Live Feeds in Marine Aquaculture

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Live Feeds in Marine Aquaculture


Live Feeds in Marine Aquaculture Edited by

Josianne G. Støttrup, PhD Danish Institute for Fisheries Research, Charlottenlund, Denmark

and Lesley A. McEvoy, PhD North Atlantic Fisheries College, Shetland Isles, UK

© 2003 by Blackwell Science Ltd, a Blackwell Publishing Company Editorial Offices: 9600 Garsington Road, Oxford OX4 2DQ, UK Tel: ⫹44 (0)1865 776868 Blackwell Publishing, Inc., 350 Main Street, Malden, MA 02148-5018, USA Tel: ⫹1 781 388 8250 Iowa State Press, a Blackwell Publishing Company, 2121 State Avenue, Ames, Iowa 50014-8300, USA Tel: ⫹1 515 292 0140 Blackwell Publishing Asia Pty Ltd, 550 Swanston Street, Carlton South, Victoria 3053, Australia Tel: ⫹61 (0)3 9347 0300 Blackwell Wissenschafts Verlag, Kurfürstendamm 57, 10707 Berlin, Germany Tel: ⫹49 (0)30 32 79 060 The right of the Author to be identified as the Author of this Work has been asserted in accordance with the Copyright, Designs and Patents Act 1988. 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, except as permitted by the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher.

First published 2003 by Blackwell Science Ltd Library of Congress Cataloging-in-Publication Data is available

ISBN 0-632-05495-6 A catalogue record for this title is available from the British Library Produced and set in Times by Gray Publishing, Tunbridge Wells, Kent Printed and bound in Great Britain by MPG Books, Bodmin, Cornwall For further information on Blackwell Science, visit our website: www.blackwell-science.com

Contents

Foreword Preface Contributors Abbreviations 1 Status of Marine Aquaculture in Relation to Live Prey: Past, Present and Future David A. Bengtson 1.1 A Historical Perspective 1.2 Marine Aquaculture Today and in the Future 1.3 The Status of Larviculture and Live Feed Usage 1.3.1 Africa 1.3.2 Asia 1.3.3 Europe 1.3.4 North America 1.3.5 Oceania 1.3.6 South America, including Central America and the Caribbean 1.4 Why is Live Feed Necessary? 1.5 Problems and Prospects with Alternatives to Live Feed 1.6 Conclusions 1.7 References 2 Production and Nutritional Value of Rotifers Esther Lubzens and Odi Zmora 2.1 Introduction 2.2 Biology and Morphological Characteristics of Rotifers 2.2.1 General biology 2.2.2 Taxonomy 2.2.2.1 The genus Brachionus 2.2.3 Morphology and physiology 2.2.3.1 Feeding 2.2.3.2 Digestion

xiii xv xvi xviii 1

1 5 7 7 8 9 10 10 11 11 12 13 13 17

17 19 19 21 21 23 23 25

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Contents

2.3

2.4

2.5

2.6

2.7 2.8

2.2.3.3 Body fluids and excretion 2.2.3.4 Movement 2.2.3.5 Nervous system and sensory organs 2.2.4 Reproduction 2.2.4.1 Asexual and sexual reproduction 2.2.4.2 Reproductive rates 2.2.4.3 Sexual reproduction and resting egg formation Culturing Rotifers 2.3.1 Selection of species and/or strain 2.3.2 Maintaining water quality in culture tanks 2.3.2.1 Organic particles 2.3.2.2 Bacteria and other organisms in the culture tanks 2.3.3 Choosing the most appropriate culture techniques 2.3.3.1 Small-scale laboratory cultures 2.3.3.2 Mass cultures Advanced Warning on State of Cultures 2.4.1 Egg ratio 2.4.2 Swimming velocity 2.4.3 Ingestion rate 2.4.4 Viscosity 2.4.5 Enzyme activity 2.4.6 Diseases Nutritional Quality of Rotifers 2.5.1 Number of rotifers consumed by larvae 2.5.2 Dry weight and caloric value 2.5.3 Biochemical composition 2.5.3.1 Protein and carbohydrate content 2.5.3.2 Lipid composition 2.5.3.3 Vitamin enrichments 2.5.4 Effects of starvation Preserved Rotifers 2.6.1 Preservation at low temperatures 2.6.2 Cryopreservation 2.6.3 Resting eggs Future Directions References

3 Biology, Tank Production and Nutritional Value of Artemia Jean Dhont and Gilbert Van Stappen 3.1 Introduction 3.2 Biology of Artemia 3.2.1 Morphology and life cycle 3.2.2 Ecology and natural distribution 3.2.3 Taxonomy

26 26 26 27 27 29 31 31 31 32 33 33 34 35 36 43 43 44 44 44 44 44 45 45 46 46 46 47 48 48 49 49 50 50 52 52 65

65 67 67 73 75

Contents

3.2.4 Strain-specific characteristics 3.2.4.1 Size and energy content 3.2.4.2 Hatching quality 3.2.4.3 Diapause characteristics 3.2.4.4 Growth rate of nauplii 3.2.4.5 Temperature and salinity tolerance 3.2.4.6 Life-history traits and reproductive capacity 3.2.4.7 Nutritional value 3.2.5 Cyst biology and diapause 3.2.5.1 Cyst morphology and physiology 3.2.5.2 Cyst metabolism and hatching 3.2.5.3 Diapause 3.3 Production Methods: Tank Production of Artemia Biomass 3.3.1 Advantages of tank production and tank-produced biomass 3.3.2 Physicochemical conditions 3.3.3 Artemia strain selection and culture density 3.3.4 Feeding 3.3.5 Infrastructure 3.3.6 Culture techniques 3.3.7 Control of infections 3.3.8 Harvest and processing of cultured Artemia 3.3.9 Production figures of intensive Artemia cultures 3.4. Biochemical composition 3.4.1 Proximate composition 3.4.1.1 Cysts and decapsulated cysts 3.4.1.2 Nauplii 3.4.1.3 Juveniles and adults 3.4.2 Lipids 3.4.2.1 Cysts and nauplii 3.4.2.2 Ongrown Artemia 3.4.3 Proteins 3.4.4 Vitamins 3.5 Applications of Artemia 3.5.1 The future use of Artemia in aquaculture 3.5.2 Hatching 3.5.3 Harvesting hatched nauplii 3.5.4 Decapsulation 3.5.5 Enrichment 3.5.5.1 Lipid enrichment 3.5.5.2 Phospholipid enrichment 3.5.5.3 Protein enrichment 3.5.5.4 Vitamin enrichment 3.5.5.5 Enrichment with prophylactics 3.5.5.6 Enrichment with other products

vii

76 77 77 77 78 78 78 79 79 79 80 81 83 83 84 86 86 88 91 92 93 93 94 94 94 94 95 96 96 97 97 98 99 99 99 102 104 105 105 107 108 109 110 110

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Contents

3.5.6 Cold storage 3.5.6.1 Survival at low temperatures 3.5.6.2 Maintenance of nutritional value 3.5.6.3 Other advantages 3.5.7 Use of juvenile and adult Artemia 3.6 References

110 110 110 111 111 112

4 Production, Harvest and Processing of Artemia from Natural Lakes Gilbert Van Stappen

122

4.1 Introduction 4.2 Pond Production of Artemia Cysts and Biomass 4.2.1 Permanent solar salt operations 4.2.2 Seasonal units 4.2.3 Site selection 4.2.3.1 Climatology 4.2.3.2 Topography 4.2.3.3 Soil conditions 4.2.4 Pond adaptation 4.2.4.1 Deepening the ponds 4.2.4.2 Dike construction 4.2.4.3 Screening 4.2.5 Preparation of ponds for Artemia cultivation 4.2.5.1 Liming 4.2.5.2 Predator control 4.2.5.3 Fertilisation 4.2.5.4 Inorganic fertilisers 4.2.5.5 Organic fertilisers 4.2.5.6 Combination of organic and inorganic fertilisers 4.2.6 Artemia inoculation 4.2.6.1 Artemia strain selection 4.2.6.2 Inoculation procedures 4.2.7 Monitoring and managing the culture system 4.2.7.1 Monitoring the Artemia population 4.2.7.2 Abiotic parameters influencing Artemia populations 4.2.7.3 Biotic factors influencing Artemia populations 4.3 Artemia Harvesting and Processing Techniques 4.3.1 Harvesting techniques 4.3.2 Processing techniques 4.4 Artemia Cyst Harvesting and Processing Techniques 4.4.1 Harvesting techniques 4.4.2 Brine processing 4.4.2.1 Brine dehydration 4.4.2.2 Size separation in brine 4.4.2.3 Density separation in brine

122 123 123 124 125 126 126 126 126 127 127 128 128 128 128 129 130 130 130 131 131 131 132 132 133 133 134 134 135 137 137 137 137 139 139

Contents

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4.4.2.4 Initial (or ‘raw’) storage 4.4.2.5 Cold storage 4.4.3 Freshwater processing 4.4.4 Drying 4.4.4.1 Layer drying in open air 4.4.4.2 Layer drying in oven 4.4.4.3 Fluidised bed drying 4.4.5 Prepackaging, packaging and storage 4.5 References

139 140 140 140 141 141 141 143 143

5 Production and Nutritional Value of Copepods Josianne G. Støttrup

145

5.1 Introduction 5.2 Biology 5.2.1 General characteristics 5.2.1.1 Calanoida 5.2.1.2 Harpacticoida 5.2.1.3 Cyclopoida 5.2.2 Copepod morphology 5.2.2.1 Digestive system 5.2.2.2 Circulatory system 5.2.2.3 Nervous system 5.2.2.4 Reproductive system 5.2.3 Reproduction 5.2.4 Resting or diapause eggs 5.2.5 Development, size and growth 5.2.5.1 Life cycle 5.2.5.2 Mortality 5.2.5.3 Size 5.2.5.4 Generation time 5.2.6 Feeding, food quality and food availability 5.2.6.1 Calanoids 5.2.6.2 Harpacticoids 5.2.6.3 Cyclopoids 5.3 Production Methods 5.3.1 Extensive and outdoor cultures 5.3.1.1 Harvest of wild zooplankton 5.3.1.2 Production in enclosed fjords or sea areas 5.3.1.3 Production in outdoor ponds or large tanks 5.3.2 Intensive culture of copepods 5.3.2.1 Calanoids 5.3.2.2 Harpacticoids 5.3.2.3 Cyclopoids

145 145 145 146 149 149 149 152 153 153 153 155 156 156 156 157 158 158 159 159 161 163 168 168 168 168 171 175 175 181 187

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Contents

5.4 Biochemical Composition 5.4.1 Carbon 5.4.2 Lipids 5.4.3 Protein 5.4.4 Free amino acids 5.4.5 Vitamin C 5.4.6 Carotenoids 5.4.7 Chitin 5.4.8 Enzymes 5.5 Nutritional Value for Fish Larvae 5.6 Application in Marine Aquaculture 5.7 References 6 The Microalgae of Aquaculture Arnaud Muller-Feuga, Jeanne Moal and Raymond Kaas 6.1 Introduction 6.2 Biology of Microalgae 6.2.1 General characteristics of microalgae 6.2.2 Growth 6.2.3 Substrates of photoautotrophy 6.2.3.1 Light 6.2.3.2 Mineral nutrients 6.2.4 Substrates of heterotrophy 6.2.5 Other factors affecting growth 6.2.5.1 Temperature 6.2.5.2 Salinity 6.2.5.3 Metabolites 6.2.5.4 pH 6.2.5.5 Mixing 6.3 Biochemical Composition of Microalgae 6.3.1 Gross biochemical composition 6.3.2 Vitamins 6.3.3 Sterols 6.3.3.1 Bacillariophyceae 6.3.3.2 Prymnesiophycaea 6.3.3.3 Prasinophyceae 6.3.3.4 Cryptophyceae 6.3.4 Fatty acids 6.3.4.1 Bacillariophyceae 6.3.4.2 Prymnesiophycaea 6.3.4.3 Prasinophyceae 6.3.4.4 Chlorophyceae 6.3.4.5 Cryptophyceae 6.3.4.6 Eustigmatophyceae

189 189 190 190 191 191 191 191 191 191 194 195 206

206 206 206 209 213 213 217 217 218 218 219 219 220 220 221 222 223 223 225 227 227 228 228 229 229 230 231 232 232

Contents

6.4 Production Methods for Aquacultural Microalgae 6.4.1 State of the art of microalgal production techniques in hatcheries 6.4.1.1 Asepsis and quality controls 6.4.1.2 Culture medium and temperature 6.4.1.3 Running the cultures 6.4.1.4 Efficiency 6.4.2 Methods of improvement 6.4.2.1 Continuous cultures 6.4.2.2 The increase in production yields 6.4.3 Heterotrophic production 6.4.4 Discussion 6.5 References 7 Uses of Microalgae in Aquaculture A. Muller-Feuga, R. Robert, C. Cahu, J. Robin and P. Divanach 7.1 Introduction 7.2 Microalgae as Food for Molluscs 7.2.1 Microalgae as a potential food source in mollusc hatcheries 7.2.1.1 Size 7.2.1.2 Digestibility 7.2.1.3 Nutritional value: biochemical composition of microalgae 7.2.1.4 Microalgae bulk production 7.2.2 Microalgal requirements in mollusc hatcheries 7.2.2.1 Feeding broodstock 7.2.2.2 Feeding larvae 7.2.2.3 Feeding spat 7.2.3 Microalgal substitutes for bivalve feeding 7.3 Microalgae as Food for Shrimp 7.3.1 Development of penaeid shrimp 7.3.2 Selection of algal species used for rearing shrimp larvae 7.3.3 Ingestion and filtration rates for shrimp larvae fed microalgae 7.3.4 Nutrient supply from algae in relation to larval shrimp requirements 7.3.5 Substitution of spray-dried algae or microparticulate compound diets for live algae 7.3.6 Other roles of algae in shrimp larval growth 7.3.7 Feeding microalgae to shrimp juveniles and adults 7.4 Microalgae as Food for Live Prey 7.4.1 Feeding live prey with live microalgae 7.4.2 Nutritional value of algae for live prey 7.4.2.1 Proteins and proximate composition 7.4.2.2 Fatty acids 7.4.2.3 Other lipid components

xi

233 233 234 236 236 236 237 238 238 240 242 243 253

253 254 255 255 256 257 257 257 258 259 261 261 263 263 263 265 266 268 269 269 270 270 271 271 272 274

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Contents

7.4.3 Vitamins 7.4.4 Minerals 7.4.5 Influence of algae on live feed and larval microbiology 7.4.6 Substitutes for live microalgae 7.5 Importance of Microalgae in Marine Finfish Larviculture 7.5.1 Range of microalgal action 7.5.2 Effects on endotrophic larval stages 7.5.3 Effects on the yolk-sac drinking stage 7.5.3.1 Drinking and ingestion of dissolved organics 7.5.3.2 Ingestion of microalgae 7.5.3.3 Digestion and assimilation of microalgae 7.5.4 Resistance to delay in first zooplanktonic feeding 7.5.5 Process and efficiency of first feeding 7.5.6 Effect on survival and growth efficiency at first feeding 7.5.7 Stimulation of digestive functions and gut flora 7.5.8 Effects on early exotrophic larvae 7.5.9 Indirect effects of microalgae on larvae 7.5.10 Future developments 7.6 References

275 275 275 276 279 279 279 280 280 281 282 283 283 284 285 286 286 288 288

Appendix I Appendix II Appendix III Appendix IV

300 304 306 307

Taxonomic Index

309

Common Names Index

312

Subject Index

313

Foreword

In the preface the editors point out that marine aquaculture has shown an important evolution from a relatively modest operation to a mature bio-industry, both in its research and development as well as for the industry. The maturation of the commercial ventures is seriously indebted to the huge progress in research and development in various disciplines relevant to aquaculture. Various indeed: if one feature can typify aquaculture research it is multidisciplinarity. Originally, the typical aquaculture researcher was a combination of a marine biologist, engineer, biochemist, physiologist, ecologist (at best), and a part-time plumber. Common knowledge on distinct topics was limited and progress was often achieved through sound, albeit empirical, experiments. Trial and error ruled, not necessarily in a one-to-one ratio. But as knowledge has broadened and deepened, aquaculture scientists became more specialized and fundamental research gradually came to underpin empirical findings. With the drive for specialization it became harder for the individual scientist to keep track of all pertinent information as well as new developments in research fields other than his or her own. This explains and justifies the multiple initiatives to provide publications, like this book, offering comprehensive updates on a selected topic. To initiate such an initiative may well turn out to be a tedious job, but fortunately strong bonds were forged between leading research groups back in the days when the aquaculture ‘who’s who’ could still be printed on a single sheet of paper. And although some degree of (mostly healthy) competition exists, it is still rather easy to find enthusiastic and dedicated authorities willing to contribute to essential reviews of the state-of-the-art of, for instance, live feed technology. As aquaculture developed, live feed has often been a bottleneck in the larviculture of many species of fish and shellfish, especially at times when upscaling from laboratory and pilot trials to large industrial units. It is therefore a pleasure and an honor to introduce Live Feed in Marine Aquaculture wherein all its relevant issues are covered by representatives of some of world’s finest aquaculture research groups. Prof. Dr Patrick Sorgeloos

Preface

The past two decades have witnessed a dramatic expansion in the culture of marine finfish and crustaceans. Marine larviculture without live feed, or crustacean cultures without microalgae, are rarities in commercial aquaculture. The development of commercial formulated feeds remains today’s upcoming challenge. In the meantime, the industry continues the struggle to produce stable quantities of high-quality live feeds. The different species used in marine aquaculture differ in their biology and culture requirements, providing ample challenges for the novice and requiring expertise in a commercial enterprise. This book includes information on the biology and culture of copepods as well as of the better-known traditional live feeds such as rotifers and Artemia. Enrichment techniques for rotifers and Artemia have greatly improved their nutritional value for marine fish species and have contributed to the expansion of the industry. Nutritional defects, however, are still evident in some species and in other cases subtle differences such as decreased tolerance to low temperatures observed during the juvenile stages in marine fish are attributed to poor nutritional diets during the larval stages. With the increasing emphasis on fish welfare and the need to produce high quality fish both for the aquaculture industry and for stocking purposes, larval nutrition will continue to be a main focus area for research within marine aquaculture. Filtering molluscs and penaeid shrimps require microalgal diets at least during some stage in their development. The development of mollusc culture is closely related to the quantity and quality of phytoplankton produced. In shrimp culture, despite the development of formulated diets, phytoplankton is still used in hatcheries to supplement the diet during the larval stages. Survival and growth in marine fish larvae can be improved by the addition of live cultures. Although their role is not fully understood, their positive effects are well documented. This book provides the reader with the compiled information on most of the live feeds used in modern marine aquaculture. Although it may not be exhaustive, it will supply the basic information needed on the biology of the species and an introduction to the relevant literature. It will also serve as a practical guide, intended to provide the reader with a good overview on culture techniques for the different species involved and with substantial reference to related literature. Three chapters deal with the hatchery production, use and nutritional value of respectively Artemia, rotifers and copepods. A further chapter deals with the production, harvest and processing of Artemia from natural lakes. Two chapters on microalgae deal with their use and production in aquaculture providing the reader with a broad insight on the importance of phytoplankton in marine aquaculture, their production and nutritional value. The book is intended for advanced undergraduates, postgraduates and researchers in the field of marine aquaculture. It may also be relevant to experimental researchers working on physiology, behaviour or energetics in these species, or to hatchery biologists who may wish to diversify or improve their culture methods.

Contributors

David A. Bengtson Professor and Graduate Program Director, Department of Fisheries, Animal and Veterinary Science, University of Rhode Island, Kingston, RI 02881, USA. Tel.: (1) 401 874 2668, fax: (1) 401 874 4017, e-mail: [email protected] Chantal Cahu Senior research scientist, Laboratoire Nutrition, IFREMER, Centre de Brest, BP 70, 29280 Plouzané, France. Tel.: (33) 2 98 22 40 40, fax: (33) 2 98 22 45 45, e-mail: [email protected] Pascal Divanach Head of Aquaculture Department, Institute of Marine Biology of Crete, PO Box 2214, Post of Poros, 71003 Iraklion, Crete, Greece. Tel.: (308) 81 24 15 43, e-mail: [email protected] Jean Dhont Researcher, Laboratory of Aquaculture & Artemia Reference Center, Ghent University, Rozier 44, 9000 Gent, Belgium. Tel.: (32) 9 264 3754, Fax: (32) 9 264 4193, e-mail: [email protected] Raymond Kaas Senior research scientist, Laboratoire Algae Biotechnology, IFREMER, Centre de Nantes, BP 21 105, 44 311 Nantes cedex 03, France. Tel.: (33) 2 40 37 41 09, fax: (33) 2 40 37 40 71, e-mail: [email protected] Esther Lubzens Department of Marine Biology and Biotechnology, Israel Oceanographic and Limnological Research, National Institute of Oceanography, PO Box 8030, Haifa 31080, Israel. Tel.: (972) 4 8515202, fax: (972) 4 8511911, e-mail: [email protected] Jeanne Moal Senior research scientist, Laboratoire Invertebrate Physiology, IFREMER, Centre de Brest, BP 70, 29280 Plouzané, France. Tel.: (33) 2 98 22 40 40, fax: (33) 2 98 22 45 45, e-mail: [email protected] Arnaud Muller-Feuga Head, Laboratoire Algae Biotechnology, IFREMER, Centre de Nantes, BP 21 105, 44 311 Nantes cedex 03, France. Tel.: (33) 2 40 37 42 20, fax: (33) 2 40 37 40 71, e-mail: [email protected]

René Robert Senior research scientist, Laboratoire Invertebrate Physiology, IFREMER, Centre de Brest, BP 70, 29280 Plouzané, France. Tel.: (33) 2 98 22 40 40, fax: (33) 2 98 22 45 45, e-mail: [email protected] Jean Robin Senior research scientist, Laboratoire Nutrition, IFREMER, Centre de Brest, BP 70, 29280 Plouzané, France. Tel.: (33) 2 98 22 40 40, fax: (33) 2 98 22 45 45, email: [email protected] Josianne G. Støttrup Senior research scientist, Danish Institute for Fisheries Research, Department for Marine Ecology and Aquaculture, Charlottenlund Castle, DK-2920 Charlottenlund, Denmark. Tel.: (45) 3396 3394, fax: (45) 3396 3333, e-mail: [email protected] Gilbert Van Stappen Researcher, Laboratory of Aquaculture & Artemia Reference Center, Ghent University, Rozier 44, B-9000 Ghent, Belgium. Tel.: (32) 9 264 37 54, fax: (32) 9 264 41 93, e-mail: [email protected] Oded Zmora National Center for Mariculture, Israel Oceanographic and Limnological Research Ltd, PO Box 1212, Eilat 88112, Israel. Tel.: (972) 7 6361442, fax: 972 7 6375761, e-mail: [email protected]

Abbreviations

AA ARA AscA AscAS ATP AWL BOD DHA DPH DPF DPPC DW EEZ EFA EPA ESD FAO FCE FW GMO GSL HUFA ILL ISA LC50 LNA L-type PAR PL PLa PUFA SCP SFB SGR SL S-type TAG WW

Amino acids Arachidonic acid; 20:4n-6 Ascorbic acid Ascorbic acid-2-sulfate Adenosine triphosphate Air-water lift system Biochemical oxygen demand Docosahexaenoic acid; 22:6n-3 Days post-hatching Days post-(first) feeding Dipalmitoyl phosphatidylcholine Dry weight Exclusive economic zones Essential fatty acids Eicosapentaenoic acid; 20:5n-3 Equivalent spherical diameter United Nations Food and Agriculture Organisation Food conversion efficiency Weight after preservation in buffered saline formaldehyde Genetically modified organisms Great Salt Lake Highly unsaturated fatty acids with 20–22 carbon atoms and more than three double bonds Incipient limiting level International Study on Artemia Lethal concentration for 50% of the sampled population Linolenic acid; 18:3n-3 Large type Photosynthetically active radiation Phospholipid Post-larvae Polyunsaturated fatty acids with more than one double bond Single cell proteins San Francisco Bay Specific growth rate Standard length Small type Triacylglycerols Wet weight

Chapter 1

Status of Marine Aquaculture in Relation to Live Prey: Past, Present and Future David A. Bengtson

1.1 A Historical Perspective It is difficult to determine exactly where and when marine aquaculture began. Milkfish culture has been conducted in Asia for centuries, based on the capture of fry from the wild (Pamplona & Mateo 1985; Liao 1991), so that modern rearing methods and live feed in the hatchery were not required. The efforts to repopulate the seas of Europe and North America in the late 1800s may provide a more useful starting point for a brief historical review of the modern methods. In response to the fishery crisis at that time, ‘hatcheries’ were constructed in several countries for the purpose of providing fertilised eggs, developing embryos and larvae for distribution back into the ocean. The hope was that these would thrive and be recruited into the commercial fisheries. Given the knowledge of freshwater fish culture in Europe and the Americas, especially of salmonid culture, which had been rapidly developing since the mid-1800s and the attendant propagation, transportation and introduction of salmonid populations (Stickney 1996), this was not an unreasonable hope for the times. By the 1890s, Britain, France, Canada and the USA all had fish hatcheries devoted to the propagation of commercially important species, such as cod (Gadas morhua), haddock (Melanogrammus aeglefinus), turbot (Scophthalmus maximus = Psetta maxima), winter flounder (Pleuronectes americanus) and lobster (Homarus sp.). The prevailing practice was to obtain gravid adults of a given species, strip them of their gametes for purposes of controlled fertilisation, sometimes on-board ship (some of the hatcheries were in fact ships), sometimes on shore, and maintain them no longer than the prolarva stage prior to release back to the ocean. The reason for the release at such an early stage of development was simple: there was no convenient live feed with which to provide them for their postlarval survival and growth. Cod larvae were raised in concrete ponds in Flødevigen, Norway, in the 1880s on a diet of natural zooplankton and in the absence of predators (Rognerud 1887), but apparently the results of this ‘experiment’ were interpreted to mean that the larvae should survive in nature, not that juveniles could be reared for release. It is only with the benefit of hindsight that we know that these ocean stocking efforts were doomed to fail, owing to the high mortality rates of fish early life-stages in the oceans. Nevertheless, many of these programmes were sustained for decades until the lack of evidence of any success from them became apparent. We will never know whether earlier discovery of easily culturable live

2


feeds would have allowed hatchery culture of these species to a later stage when they might have had better chances of oceanic survival. Indeed, the field of stock enhancement might have been advanced by several decades had convenient live feeds been available in the late 1800s. Just as many of the ocean stocking programmes of the late 1800s and early 1900s were being phased out, two developments occurred half a world apart that paved the way for much of the development of modern marine aquaculture. First, nauplii of the brine shrimp, Artemia, were found to be a good food for raising both freshwater and some marine larval fish (Seale 1933; Gross 1937; Rollefsen 1939). This allowed the culture of at least some fish species (those with mouths large enough to ingest Artemia nauplii as a first food). The use of Artemia nauplii as a convenient live feed, not only for fish, but also (and especially) for crustaceans, has perhaps done as much for the explosion of marine aquaculture in the late 1900s as any other development. Secondly, in the 1930s, Japanese researchers, beginning with Dr M. Fujinaga, began research on the culture of the kuruma prawn, Penaeus japonicus, which subsequently led to the development of the shrimp industry that we know today (Liao & Chien 1994). That research, interrupted unfortunately by World War II, continued through the 1960s, when commercial culture of P. japonicus was finally achieved. Meanwhile, techniques were developed in the 1920s and 1930s that led to the development of molluscan hatcheries. Oyster culture, which has been known since Roman times, expanded in Japan in the seventeenth century with the finding that oyster larvae would settle on bamboo stakes, and expanded further in Europe, North America and Australia in the nineteenth century based on bottom culture (Bardach et al. 1972). Similarly, clam culture has been known in Japan and mussel culture has been known in France for several hundred years (Bardach et al. 1972). However, molluscan culture always relied on the settling of larvae from the natural zooplankton (and still does in many areas). Wells (1920) used a milk clarifier to retain oyster larvae while their water was being changed. Although hatchery spawning of oysters had been demonstrated as early as 1879, no one had been able successfully to change oyster culture water, and therefore replenish the algal food, without losing the larvae (Wells 1920). Wells (1927) then went on to raise clam larvae as well. Spawning and successful larval culture of mussels was not achieved until the early 1950s (Loosanoff & Davis 1963). Investigations of algal feeds for the rearing of molluscan larvae took place in the 1930s at both the Conwy, Wales, Fisheries Experiment Station (Walne 1974) and the Milford, USA, Bureau of Commercial Fisheries Biological Laboratory (Loosanoff & Davis 1963). Fertilisation of large tanks of filtered seawater to induce mixed phytoplankton blooms as food for molluscan larvae was carried out continuously beginning in 1938 (Loosanoff & Davis 1963), despite the contention that ‘large-scale cultivation of microalgae … was probably first considered seriously in Germany during World War II’ (Becker 1994). Decades of work at the Conwy and Milford laboratories paved the way for hatchery production of molluscs for commercial aquaculture in which natural settling of larvae was either impossible or undesirable. The culture of algae seems to have its origins in the late 1800s and was enabled by the methods developed by bacteriologists (Bold 1950). Marine algal culture lagged behind its freshwater counterpart, which successfully used uncomplicated media (Pringsheim 1924; Schreiber 1927) in the early 1900s. A significant advance in marine algal culture was reported by Gross (1937), who tried to culture diatoms and dinoflagellates, but whose

Status of Marine Aquaculture in Relation to Live Prey: Past, Present and Future

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attention was drawn to ‘nannoplankton flagellates, most of them probably unknown systematically’ of about 2–10 ␮m in size. He was able to culture these and use them as feed for harpacticoid copepods over three copepod generations. He summarised his work by writing ‘All these experiments led me to the conclusion that the autotroph nannoplankton flagellates are of great importance in the food economy of the sea.’ Little did he know that they would also be of great importance in aquaculture. Methods for marine algal culture continued to advance during the middle of the twentieth century with the development of artificial media (Provasoli et al. 1957) and the development of ‘f’ medium for the enrichment of seawater (Guillard & Ryther 1962). Improved methods for monospecific algal cultures allowed expansion of hatcheries for molluscan aquaculture and enabled culture of live invertebrates as feed for larval fish and crustaceans. Another extraordinarily important advance was made in the 1960s, when Japanese researchers discovered that rotifers, Brachionus plicatilis, previously considered a pest in culture ponds, could be used as a first food for larvae of both freshwater and marine fish species (Hirata 1979). This advance clearly allowed the culture of many more species whose larvae hatched at such a small size that their mouth gapes were insufficient for the ingestion of the larger Artemia prey. In retrospect, considering the large number of commercially important marine fish species that have been brought into culture and that rely on rotifers as first food in culture facilities, the debt to those initial Japanese culturists is profound. The 1960s saw widespread interest in the culture of commercially important marine fish species, first from a research perspective. In Japan, efforts were made to culture larvae of red sea bream, flounder and puffer fish, among others. In Britain, the White Fish Authority engaged in activities particularly in the area of flatfish culture (Shelbourne 1964) that ultimately led to the first commercial production of turbot in 1976 (Person-Le Ruyet et al. 1991). In France, research conducted primarily in the 1970s led to the development of the French sea bass and turbot industries in the 1980s (Person-LeRuyet et al. 1991; Coves et al. 1991). In many countries, including the USA, interest in larval fish biology from a fisheries perspective caused many laboratories to begin rearing larval fish on fieldcollected zooplankton, sometimes supplemented with rotifers and Artemia nauplii (e.g. Houde 1972), in order to conduct fisheries research (e.g. Laurence et al. 1981). Norwegian scientists, using some pertinent results from the cod-spawning and restocking efforts 100 years earlier, began pond culture of cod larvae using natural zooplankton in the mid-1970s, and followed that with a major research programme on halibut culture beginning in the 1980s. Many of the above efforts documented the difficulty of rearing the extremely delicate marine larvae through the first-feeding stages and on to metamorphosis and subsequent grow-out (e.g. Jones 1972). Clearly, early efforts at rearing larval marine fish, whether using natural zooplankton, rotifers or Artemia, were fraught with difficulties, to the point that the famous Kyoto conference in 1976 declared larval rearing a major bottleneck in marine aquaculture (Pillay 1979). As the 1970s saw the beginning of commercial production of several marine finfish and penaeid shrimp species, this decade is also noteworthy for the discovery that live feeds vary significantly in quality. From early reports suspecting pollutants (Bookhout & Costlow 1970) to later, more definitive, studies (Watanabe et al. 1978; and several papers from the International Study on Artemia: see Persoone et al. 1980), it became very obvious that different geographical strains of brine shrimp differed in their ability to support good survival and growth of marine larvae. The finding that the differences were due primarily to

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fatty acid profiles led to productive collaborations between aquaculturists and biochemists that have resulted in literally hundreds of publications on the subject, as well as commercial products that have played no small role in the development of marine aquaculture. The lessons learned from brine shrimp were also shown to apply to rotifers (Lubzens et al. 1984), to be intimately connected to algal food supply (Léger et al. 1986) and to explain much of the high quality of natural zooplankton as a food item. The necessity for aquaculturists to understand in detail the physiology and biochemistry of the organisms that they raise has contributed much to making marine aquaculture a sophisticated industry. One event of the 1970s played a major role in the development of marine aquaculture, especially stock enhancement: the establishment of exclusive economic zones (EEZs). This event convinced the Japanese that they needed to become self-sufficient in seafood production, because they could no longer fish at will in the coastal waters of many nations and because they saw that an interruption of supplies on an international scale was a real possibility (Sproul & Tominaga 1992). The Japanese government responded by embarking on a massive research and hatchery-building campaign (Davy 1990, 1991). National and prefectural hatcheries now produce millions of fish, prawns and crabs for release into Japanese waters each year through the efforts of the national and prefectural Sea Farming Associations. As part of this effort, Japanese researchers have often led the way in marine aquaculture research and the practical applications of that research can be seen around the world. Investigations into the improvement of live feed, especially rotifers and Artemia, have certainly been a major contribution of the Japanese research programme. The last quarter of the twentieth century saw the explosion of marine aquaculture, both shrimp and fish. The aforementioned Japanese work on kuruma prawn led ultimately to the culture of numerous penaeid species around the world. Although postlarval shrimp for stocking into grow-out ponds were for years collected from the wild, the recent trend has been toward hatchery production, which is heavily dependent on microalgae and Artemia nauplii as larval feeds. Japanese research on red sea bream (Pagrus major) similarly led to the development of that species for commercial aquaculture in Japan as well as for stock enhancement. Research on other fish species in various areas of the world has led to large-scale aquaculture production of gilthead sea bream (Sparus aurata) and sea bass (Dicentrarchus labrax) in the Mediterranean region, Asian sea bass (Lates calcarifer) in the Indo-West Pacific region, turbot in western Europe and olive flounder (Paralichthys olivaceus) in east Asia, among other species, all dependent on live feed in the hatchery stage. Although commercial aquaculture of these species has become well established, a variety of species is still undergoing commercial growing pains, for example, Atlantic halibut (Hippoglossus hippoglossus), several groupers (Epinephelus spp.) and cod; again, all require live feed in the hatchery. Indeed, of all the marine fish species in production or in the research and development pipeline, it seems that only the wolfish species (Anarhichas spp.) can be routinely fed formulated diets directly upon hatching. It appears likely that live feed will be required well into the future, not only for the established and nearly established species, but also for the plethora of new bream, sciaenid, flounder and sole species currently poised to make their debut appearances on the world’s commercial aquaculture stage. One of the more interesting controversies in the live feed area is the view that natural or cultured copepods are necessary for at least some species, as opposed to the view that rotifers and Artemia nauplii are quite sufficient. The former view seems to come from the


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Nordic countries, particularly Norway and Denmark. A valuable result of this controversy has been the extensive biochemical analyses of natural zooplankton to determine whether particular nutritional ingredients are present there, but lacking in rotifers and Artemia. Van der Meeren and Naas (1997) provide an excellent review of larval fish rearing in large, enclosed systems primarily through the use of natural zooplankton. Norwegian scientists have pioneered this field of larviculture in ponds and natural inlets that have been closed off from the sea. The procedure requires either the filtration of incoming water or the use of rotenone to kill any predators. The enclosures can be fertilised to increase the phytoplankton productivity within, so that large populations of copepods are available on which the larvae can feed. Alternatively, larvae can be reared in a bag enclosure and provided with additional zooplankton that has been collected from the adjacent waters by the use of a plankton wheel (see van der Meeren & Naas 1997, p. 373). Cod have been raised successfully in Norwegian lagoons using these methods and halibut have been raised there in bags. A company in Denmark produces turbot larvae in large concrete tanks in which zooplankton ‘blooms’ are induced. In addition, at least one of the tanks is devoted exclusively to extensive copepod culture and used to feed the larval fish tanks if live prey levels therein fall too low. While larvae of all these species grow extremely well on the natural zooplankton, the production at such facilities is, almost by definition in the temperate zone, seasonal and not amenable to more intensive production methods in which juveniles must be produced year-round. However, some species produce larvae with mouths that are too small to ingest even rotifers at first feeding and an alternative live prey, such as copepod nauplii, would be necessary to culture such species. Furthermore, it is well known that the nutritional value of copepods is better than that of the convenient live feeds such as rotifers and brine shrimp. Thus, abundant rationale exists for research on the mass production of copepods. Research efforts into rearing copepods in intensive indoor systems have shown some promise, but commercial-scale production has not been achieved (see review by Støttrup 2000). Oddly, the use of live prey in hatcheries may be strongly related to one of the banes of the marine larviculturist, disease. As beneficial, indeed critical, as rotifers have been to the development of marine fish larviculture, it has been known for some time that rotifer cultures fed to a tank of fish can also carry pathogenic bacteria, such as Vibrio spp., that can lead to subsequent disease problems (Gatesoupe 1982). In a similar way, bacteria from Artemia hatching water, if the Artemia cysts have not been decapsulated or otherwise disinfected, can introduce to the fish tanks xenobiotics from wherever in the world the Artemia cysts originated. Disease has become a major consideration in hatcheries and the microbial ecology of hatchery tanks has become an area of intense research which one hopes will lead to more predictable hatchery outputs (Vadstein et al. 1993; Vadstein 1997). Live feeds thus have both good and bad aspects, and one challenge of the future is to minimize the bad aspects.

1.2 Marine Aquaculture Today and in the Future At the time of writing, the United Nations Food and Agriculture Organisation (FAO) had just released its preliminary estimates for fisheries and aquaculture statistics from 1999, which indicate that world aquaculture production was 32.9 million tonnes (19.8 million t from freshwater, 13.1 million t from marine) (FAO 2000). Thus, aquaculture makes up

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more than 35% of the total 92.6 million tonnes of fisheries products consumed by humans. Marine aquaculture has been growing by about 0.9 million tonnes per year in recent years, while the growth of freshwater aquaculture has been closer to 1.1 million tonnes per year. In 1998, the last year for which full statistics are available, aquaculture production of purely marine fish was 781,000 t, thus lagging behind crustaceans, mostly shrimp (1,564,000 t), and diadromous fish, mostly salmonids (1,909,000 t), and far behind freshwater fish, mostly carp (17,355,000 t) (FAO 2000). The major research endeavours in marine hatchery aquaculture today can be divided into three broad categories: improving reliability of production for existing species, development of culture methods for new species, and maximising the survival probability in the wild for hatchery-reared fish in stock enhancement programmes. Production reliability is being improved by several strategies. Selective breeding programmes for both fast growth and disease resistance should result in improved hatchery production in future years and those for improved flesh quality should ultimately yield a better product going to market. Improved management of microbial ecology in hatchery tanks through better husbandry, use of probiotics, etc., should also help production reliability. Development of vaccines, delivered by injection to older juveniles and by immersion to younger juveniles, should likewise aid in the minimisation of disease problems. Finally, the search for replacements for live feed proceeds apace as the world-wide availability of Artemia remains a question (see below) and the culture of algae and rotifers continues to be a labour-intensive requirement for marine hatcheries. The development of culture methods for new species tends to demonstrate the similarity of the requirements for raising different marine fish species, rather than differences between them. The research in this area generally involves the fine-tuning of widely accepted principles and procedures for application to the new species in question; if the culture of a species requires more than fine-tuning, its commercial development can be slowed or impaired (as in the case of Atlantic halibut). For example, if live feed other than rotifers and Artemia are required, the development of a new species is immediately hindered. The maximisation of survival of hatchery fish in the wild has been primarily the province of Japanese researchers (e.g. Tsukamoto et al. 1989; Yamashita et al. 1994), but their methods have more recently been adapted by others (e.g. Leber et al. 1997; Otterå et al. 1999). The basis of this area of research is the production of very high-quality juveniles from hatcheries (using only first-generation broodstock to maintain genetic integrity with the natural population), the identification of optimal release strategies (fish size, season, release site) and the use of conditioning methods, both in the hatchery and in the wild, to allow the fish to make the transition from hatchery to natural environment with maximum likelihood of survival. The fish are generally released as juveniles and therefore well adapted to formulated diets, but clearly the use of high-quality live feed is necessary earlier in the hatchery to produce the high-quality juveniles needed for release. As we proceed into the future, a few big questions dominate the landscape. The overriding one is ‘How do we make aquaculture sustainable?’ The environmental consequences of the explosion of marine aquaculture in the last quarter of the twentieth century have become a major international concern within the past decade. From shrimp farming in mangrove areas to organic enrichment from salmon net-pen culture, the ecological insults brought about by marine aquaculture are trumpeted to the world’s consumers by environmental groups. The global aquaculture industry is responding (Boyd 1999; SSFA & NAFC 2000) and there is cause for optimism that improved practices will be the norm in the future.


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A second major question is ‘What will we feed aquacultured organisms in the future?’ This question applies both to hatchery-reared fish and crustaceans and to those in grow-out operations. At the hatchery level, the industry is currently undergoing a kind of crisis in Artemia cyst availability. This is due in large part to recent poor harvests from the traditionally productive Great Salt Lake, Utah, USA, along with increased regulation of harvests from those waters. The identification of new sources of Artemia cysts for harvest, for example in Asian countries that once belonged to the Soviet Union, allow some hope that this crisis will soon fade. Recalling that the last Artemia crisis in the mid-1970s led to the discovery of new geographical strains and focused research on Artemia cyst quality, one wonders what the current crisis will yield. A renaissance in research on formulated diets to replace Artemia is already underway (Kolkovsky et al. 1997; Yúfera et al. 1999) and one hopes that the results will be more commercialisable than those from the flurry of research on microdiets that arose from the last Artemia crisis. In a manner similar to the Artemia crises, periodic shortages of fish meal world-wide (usually due to climatic conditions off western South America) bring about intensive research into fish meal replacements. Recently, however, the aquaculture industry, as well as environmental groups, have questioned whether the projected growth of the industry over the next 30 years is possible in the light of fish meal availability even in the best of times (Naylor et al. 2000). It appears that partial or complete replacement of fish meal in the formulation of diets for some species will be necessary or desirable if the industry hopes to grow to the degree necessary. While this is a question primarily for grow-out producers, the ramifications will certainly be felt all the way back to the hatchery phase of the industry (Will we no longer grow species that require fish meal? Should we select for individuals that have minimal fish meal requirements?). One final major question concerns the role that biotechnology will play in aquaculture. Clearly, the biotechnology industry is already playing a role in products for the prevention, diagnosis and treatment of disease. Genome mapping is beginning for some of the major aquaculture species (M. Gomez-Chiarri, personal communication), but the question of whether genetically modified organisms (GMO) will be allowed in the marketplace is still a question for regulators. An even greater question is whether consumers will accept such products. It is likely that the answers to those questions will become apparent with GM products from terrestrial agriculture before aquaculture will address them in a major way.

1.3 The Status of Larviculture and Live Feed Usage It may be useful in this introductory chapter to describe the status of marine finfish and crustacean larviculture and live prey usage in different regions of the world, so that the reader receives a broad overview on a global scale. The review will be presented continent by continent, in alphabetical order, based on production figures supplied by FAO for the calendar year 1997 (FAO 1999) and various articles as cited.

1.3.1 Africa Africa’s marine finfish and crustacean production comes largely from countries bordering the Mediterranean. Egypt is the leading producer, with over 16,000 t of mullet production, but

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these are grown from wild-captured fry, so no live feed is used (Wassef 2000). Egypt also produces more than 2000 t each of sea bass and seabream; the fry are mostly collected from the wild, but hatchery production is expanding, therefore requiring the use of rotifers and Artemia (Wassef 2000). Morocco and Tunisia also produce hundreds of tonnes each of sea bass and sea bream, so also require hatchery production using rotifers and Artemia (Romdhane 1992). Madagascar and the Seychelles Islands produce significant quantities of shrimp, Penaeus monodon, and South Africa has a small production of Penaeus indicus and P. japonicus, all of which require algae and Artemia as live feeds in the hatchery.

1.3.2 Asia Moving out of Africa and proceeding through Asia from west to east, one finds that Israel and Turkey, like Egypt, produce significant quantities of sea bass and sea bream, all apparently from hatchery production and requiring rotifers and Artemia. Iran and Saudi Arabia both report production of hundreds of tonnes of penaeid shrimp, requiring the use of algae and Artemia in hatcheries. A small amount of marine finfish culture is reported from Kuwait and Qatar. Penaeid shrimp culture dominates the mariculture of Pakistan (ca. 50 t), India (⬎50,000 t), Sri Lanka (ca. 5000 t), Bangladesh (⬎50,000 t), Myanmar (ca. 8 t) and Vietnam (ca. 80,000 t). Although some extensive culture using wild-caught shrimp still exists in India and Vietnam (Binh & Lin 1995; Shetty & Satyanarayana Rao 1996), the majority of the above production appears to rely on hatchery production using the normal methods with algae and Artemia (Shetty & Satyanarayana Rao 1996; Nien & Lin 1996). Thailand is the world’s largest shrimp producer (FAO 2000), based primarily on P. monodon, with production of ⬎200,000 t in 1997 (FAO 1999). Since this production is almost exclusively hatchery based, use of algae and Artemia is extremely heavy. Thailand also has significant production of Asian sea bass, L. calcarifer (⬎4000 t), grouper, Epinephelus spp. (ca. 800 t), and threadfin, Eleutheronema tetradactylum (ca. 400 t), requiring hatchery usage of rotifers and Artemia. The Philippines, Indonesia and Malaysia are somewhat similar to Thailand, having predominantly shrimp culture with P. monodon as the major species (although with substantial culture of Penaeus merguiensis and Metapenaeus spp. as well), but also exhibiting increasing production levels of finfish, L. calcarifer in the case of Indonesia and a variety of species (e.g. snappers, basses, rabbitfish, groupers) in the case of the Philippines and Malaysia. Thus, these countries also have significant hatchery production of both shrimp (using algae and Artemia) and fish (using rotifers and Artemia). It should be pointed out that both Indonesia and the Philippines are predominated by milkfish culture, but that industry still depends largely on capture of fry from the wild and therefore does not require live feed for larviculture. Singapore, Hong Kong and Taiwan are similar to each other in having their fish and crustacean mariculture activities dominated by finfish culture, with relatively little, if any, penaeid shrimp culture. They all culture Asian sea bass, groupers and snappers to greater or lesser degrees, and Hong Kong produces significant quantities (ca. 800 t) of silver bream, Rhabdosargus sarba, but Taiwan produces a wide variety of marine finfish, including over 4000 t of black sea bream, Acanthopagrus schlegeli, and ca. 400 t of red sea bream, Pagrus major, among many others. Culture of these high-value species is quite industrialised, with significant hatchery production relying on the standard formula of rotifers and Artemia.


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The People’s Republic of China is the world’s largest aquaculture producer, responsible for a remarkable two-thirds of all aquaculture production globally. Their marine finfish and crustacean production is a fairly minor component of their total production, but still dwarfs that of most other countries. Shrimp production, mostly Penaeus chinensis, still exceeds 100,000 t per year despite problems with disease epidemics in the 1990s. Cen and Zhang (1998) state that all shrimp seed for production now comes from ‘a controlled environment’, rather than being collected from the wild. It is impossible to determine from FAO statistics the production of individual marine fish species in China, but Cen and Zhang (1998) report 145,000 t of production in 1995 (which had apparently increased to ⬎250,000 t by 1997), including mullets, breams, groupers, tilapia, Asian sea bass, puffer fish and olive flounder. Japan produces far more marine finfish than shrimp. Only a little more than 2000 t of kuruma prawn, P. japonicus, the species that began the industry, is still produced commercially in Japan. Japanese hatcheries, however, produce prodigious amounts of both finfish and shrimp for stock enhancement and sea ranching efforts. Oddly, yellowtail, Seriola quinqueradiata, the fish with the largest production in commercial aquaculture (nearly 140,000 t) is still dependent on wild-caught fry. Other major finfish produced commercially include red sea bream (⬎80,000 t), olive flounder (⬎8500 t), Tetraodontidae (nearly 6000 t) and jack mackerels, Trachurus spp. (⬎5700 t). These require hatchery production using the rotifer and Artemia techniques that the Japanese largely developed. Hatchery rearing with rotifers and Artemia is also necessary for the production of fry for the stock enhancement programmes. Major species with numbers of finfish fry released in 1995 are: P. olivaceus (23 million), P. major (19 million) and A. schlegeli (6 million) (Fushimi 1998). These impressive numbers are, however, surpassed by those for kuruma prawn, 305 million, which requires algae and Artemia in the hatcheries. Overall, Japan produced seed for stock enhancement of 80 species in a total of 284 facilities (such as national, prefectural and local hatcheries), with 11 species receiving more than 10 million seed and 33 species receiving at least 1 million seed (Fushimi 1998). This production included molluscs and echinoderms as well as fish and crustaceans, but clearly Japan is a major user of live feeds such as algae, rotifers and Artemia for marine finfish and crustacean culture in both commercial aquaculture and governmental stock enhancement efforts. Finally, South Korea has been rapidly expanding its marine finfish culture, primarily P. olivaceus (⬎26,000 t), while maintaining production of a few hundred tonnes of red sea bream and yellowtail and ⬎12,000 t of various other species. The flounder culture, as in Japan, is totally dependent on hatchery production of fingerlings, with both rotifers and Artemia required as live feed.

1.3.3 Europe Although minimal production of penaeid shrimp species is reported in Albania, Cyprus, France, Greece, Italy and Spain (requiring use of algae and Artemia), the production of marine finfish far outweighs that of marine crustaceans in Europe. In the Mediterranean countries plus Portugal, the dominant species are sea bass, with over 24,000 t, and sea bream, with nearly 30,000 t, reported for 1997 (FAO 1999). Greece is by far the leader, with over 15,000 t of sea bass and over 18,000 t of sea bream. Cyprus, France, Greece,

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Italy, Portugal and Spain all produce minor to significant quantities of other finfish species as well, for an additional total of between 4500 and 5000 t. All of this production is based on hatchery-raised fry. Sea bass can feed on Artemia as a first feed, whereas sea bream also require rotifers prior to feeding on Artemia. In northern Europe, the major species are cod and halibut and there is much greater usage of natural zooplankton in addition to, or in place of, rotifers and Artemia. Although research on culture of both species has been going on since the early 1970s, the actual commercial production is still rather small, but growing. Norway is the leader in cod production, with slightly more than 300 t of commercial production in 1997, but Norwegian scientists have also been engaged for a number of years in production of cod fingerlings for stock enhancement projects. Cod larvae are produced in tanks, ponds or blocked-off sections of fjords, and are fed natural zooplankton obtained from the same or similar enclosed bodies of water which have been fertilised to bring about phytoplankton blooms (Huse 1991). Halibut are produced in a variety of enclosed systems and can eat Artemia as first food, but Norwegian producers argue that natural zooplankton is also necessary during the larval stages for production of good-quality fry. Production of halibut has more recently been effected in Iceland, which has now become the leading producer of halibut juveniles, despite the fact that the halibut larvae in Iceland are raised without natural zooplankton (K. Pittman, personal communication).

1.3.4 North America Relatively little culture of marine finfish and crustaceans is reported from North America. Culture of cold-water finfish in Canada, using rotifers and Artemia as live feed for larvae, is still in the trial phases. In the USA, commercial production is reported for red drum (Sciaenops ocellata) and summer flounder (Paralichthys dentatus), both of which require rotifers and Artemia as prey in the hatchery. Oddly, FAO includes hybrid striped bass (Morone saxatilis ⫻ Morone chrysops) as a marine fish species in its statistics, even though the fish are reared in fresh water. The larvae of those bass are mostly raised in earthen ponds, which are fertilised in spring to induce blooms of phytoplankton and zooplankton before the introduction of the fish larvae (Harrell 1997). Since the early 1980s, hatchery production of a few species has been necessary for enhancement, restoration of stocks or mitigation of environmental impacts; striped bass (M. saxatilis), red drum (S. ocellata) and spotted sea trout (Cynoscion nebulosus) are the most noteworthy of these. In addition, the USA reported production of 1200 t of Litopenaeus vannamei in 1997, originating from intensive hatcheries with heavy use of algae and Artemia. Mexico has become a large producer of shrimp (L. vannamei), with over 17,000 t of production reported in 1997.

1.3.5 Oceania The majority of production here is penaeid shrimp. Australia reported nearly 1600 t of P. monodon production in 1997 and New Caledonia over 1100 t of Penaeus spp. All are from hatchery origin, requiring algae and Artemia. Other island nations (Fiji Islands,


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French Polynesia, Guam and the Solomon Islands) all report minor production (⬍50 t each) of various penaeids. Australia also reported over 500 t of Asian sea bass production.

1.3.6 South America, including Central America and the Caribbean With the exception of Chile, the culture of marine finfish and crustaceans in this region is overwhelmingly dominated by penaeid shrimp. Ecuador is the clear leader, with 1997 production of over 120,000 t. Other countries producing between 2000 and 10,000 t include Brazil, Colombia, Costa Rica, Guatemala, Honduras, Nicaragua, Panama, Peru and Venezuela. Although wild seed is still used in some places, the trend is for increased reliance on hatchery production of postlarvae. The hatchery techniques are by now quite standard throughout the region, with algae and Artemia as the live feeds of choice, just as they are elsewhere in the world. Chile has been rapidly increasing its finfish aquaculture industry and is poised to become the world leader in salmon production, but it also is producing turbot in significant quantities for export to Europe. Hatchery rearing of these turbot depends on both rotifers and Artemia in the same way that they are used by the European turbot industry. To summarise this geographical review, hatchery production of penaeid shrimp postlarvae around the world depends on the use of live algae for the early stages and Artemia for the later stages. Usage of formulated diets to supplement and eventually replace Artemia is apparently increasing (see below), but live feed is still dominant at this point. For marine finfish, hatchery production of juveniles globally is normally accomplished just with Artemia, if the mouth gape is large enough at first feeding, or with rotifers and Artemia, if a smaller initial feed is required. It should be pointed out that algae is routinely used in marine fish culture of the so-called ‘green-water’ method, but it is still not clear to what degree the algae may be contributing directly to the nutrition of the larvae (Reitan et al. 1997). The use of natural zooplankton, or the use of cultured foods other than algae, rotifers or Artemia, is limited to a few places in the world, but it can be very important in those particular places.

1.4 Why is Live Feed Necessary? Fish biologists categorise larvae of two types: precocial and altricial. Precocial larvae are those that, when the yolk sac is exhausted, appear as mini-adults, exhibiting fully developed fins and a mature digestive system including a functional stomach. Such fish can ingest and digest formulated diets as a first food and are best exemplified by the salmon and trout raised extensively in hatcheries around the world without the benefit of live food. Altricial larvae are those that, when the yolk sac is exhausted, remain in a relatively undeveloped state. The digestive system is still rudimentary, lacking a stomach, and much of the protein digestion takes place in hindgut epithelial cells (Govoni et al. 1986). Such a digestive system seems (at this point) to be incapable of processing formulated diets in a manner that allows survival and growth of the larvae comparable to those fed on live feed. Altricial larvae therefore appear to require live feed, but there may be other reasons besides the digestibility question. Live feeds are able to swim in the water column and are thus constantly available to the

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larvae. Formulated diets tend to aggregate on the water surface or, more commonly, sink quickly to the bottom, and are thus normally less available to the larvae than are the live feeds. In addition, the movement of live feed in the water is likely to stimulate larval feeding responses, since evolutionary history has probably adapted them to attack moving prey in nature. Formulated diets are generally capable of moving only in a downward direction, towards the bottom. Finally, live prey, with a thin exoskeleton and high water content, may be more palatable to the larvae once taken into the mouth, compared with the hard, dry formulated diets. This last point is rather critical, especially when considered in light of the fish larva’s absence of feeding appendages; any foods must enter the mouth whole (i.e. the larva’s mouth gape must be of sufficient size for particle ingestion to occur) and they are quickly either accepted or rejected on the basis of palatability. Jones et al. (1997a,b) reviewed the digestive physiology and nutrition of larval crustaceans and remarked that larval penaeid shrimp and late larval Macrobrachium can successfully use artificial diets. Crustacean larvae such as shrimp are qualitatively different from fish larvae. They are filter feeders as early larvae and by the time they can feed on live zooplankton, they possess not only feeding appendages with which to manipulate the prey organisms captured, but also a gut morphology and physiology with which to digest formulated diets more effectively (Jones et al. 1997b). Larval shrimp tanks tend to have greater water movement than do larval fish tanks, so that the formulated feeds are better able to stay in the water column and the shrimp can capture the diets with their feeding appendages rather than having to ingest them in a single gulp as larval fish must do. As a result, shrimp larvae have been shown to survive and grow well on formulated feeds (Jones et al. 1997a), whereas fish larvae do not (Holt 2000).

1.5 Problems and Prospects with Alternatives to Live Feed Pelleted diets became common in the salmonid industry during the middle of the twentieth century. As marine larviculture developed in the 1960s and 1970s, aquaculturists perceived that, even though encysted crustaceans such as Artemia provided a convenient live feed, they were still not as convenient as a formulated diet would be. In addition, the aforementioned Artemia crisis of the 1970s suggested that, if marine aquaculture were to achieve its potential rate of expansion, it needed to be unimpeded by a shortage of live feed. The natural tendency was to enlist the aid of fish nutritionists who specialised in pelleted diets for grow-out and to convince them somehow to make those pelleted diets small enough for fish or crustacean larvae to eat. The results were not encouraging (Girin 1979). Formulated feeds (often referred to as artificial diets, inert diets or prepared diets) could not provide nearly as good survival and growth of marine larvae as could live prey, although some studies indicated that partial substitution of Artemia with formulated feeds did yield survival and growth of larvae equal to that of larvae fed Artemia alone (e.g. Beck & Bengtson 1979). With the advent of microencapsulation technology, primarily for the pharmaceutical industry, came the hope that this technology might be applied to the development of successful microdiets for larviculture as well. Commercial microencapsulated larval feeds were developed in the 1980s and showed much greater success with shrimp larvae than


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with fish larvae (Jones et al. 1993; Person Le Ruyet et al. 1993). In recent years, there has been a tendency to use formulated feeds in shrimp hatcheries to minimise usage of Artemia, but the usage in fish hatcheries appears to be less (although the exact numbers are not known). As argued previously (Bengtson 1993), breakthroughs in the development of microdiets for marine fish larvae require the multidisciplinary efforts of biochemists, nutritionists, chemical engineers and others all working with aquaculturists. These multidisciplinary teams have been forming and conducting research, but so far the ‘magic bullet’ remains elusive.

1.6 Conclusions Marine finfish and crustacean aquaculture has greatly expanded since the early 1980s, enabled in large part by the development of fairly standard hatchery protocols for live feed usage around the world. When new species are developed or new geographical regions opened up to aquaculture, it is a great benefit to have a standard ‘menu’ with a minimal number of well-established ‘entrees’ to use as live feed in the hatchery. The fact that development of new feed(s) is not required for the development of each new species has greatly facilitated aquaculture expansion. The major questions that remain for the future involve the availability and costs of feed for larvae: To what degree will Artemia availability and cost limit the expansion of marine aquaculture? To what degree can formulated diets replace live feeds generally, and at what cost? How can the costs of live feed culture (primarily algae, rotifers and alternative prey such as copepods) be minimised so that hatcheries can produce fingerlings/postlarvae at lower cost? The Kyoto conference identified larviculture as a bottleneck in aquaculture primarily on technical grounds. As many of the technical problems have been overcome, thanks to the tireless efforts of researchers around the world, there is now a need to reduce the economic bottleneck that larviculture still exerts on the commercial culture of many species.

1.7 References Bardach, J.E., Ryther, J.H. & McLarney, W.O. (1972) Aquaculture: The Farming and Husbandry of Freshwater and Marine Organisms. Wiley-Interscience, New York. Beck, A.D. & Bengtson, D.A. (1979) Evaluating effects of live and artificial diets on survival and growth of the marine atherinid fish Atlantic silverside, Menidia menidia. In: Finfish Nutrition and Fishfeed Technology. Vol. I (Ed. by J.E. Halver & K. Tiews), pp. 479–489. Heenemann, Berlin. Becker, E.W. (ed.) (1994) Microalgae: Biotechnology and Microbiology. Cambridge University Press, Cambridge. Bengtson, D.A. (1993) A comprehensive program for the evaluation of artificial diets. J. World Aquacult. Soc., 24, 285–293. Binh, C.T. & Lin, C.K. (1995) Shrimp culture in Vietnam. World Aquacult., 26, 27–33. Bold, H.C. (1950) Problems in the cultivation of algae. In: The Culturing of Algae (Ed. by J. Brunel, G.W. Prescott & L.H. Tiffany), pp. 11–17. Charles F. Kettering Foundation, New York. Bookhout, C.G. & Costlow, J.B., Jr (1970) Nutritional effects of Artemia from different locations on larval development of crabs. Helgoländ. Meeresunters., 20, 435–442.

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Boyd, C.E. (1999) Codes of Practice for Responsible Shrimp Farming. Global Aquaculture Alliance, St. Louis, MO. Cen, F. & Zhang, D. (1998) Development and status of the aquaculture industry in the People’s Republic of China. World Aquacult., 29, 52–56. Coves, D., Dewavrin, G., Breuil, G. & Devauchelle, N. (1991) Culture of sea bass (Dicentrarchus labrax L.). In: Handbook of Mariculture, Vol. II, Finfish Aquaculture (Ed. by J.P. McVey), pp. 3–20. CRC Press, Boca Raton, FL. Davy, F.B. (1990) Mariculture in Japan. 1. Development of an industry. World Aquacult., 21, 36–47. Davy, F.B. (1991) Mariculture in Japan: current practices. World Aquacult., 22, 30–35. FAO (1999) Aquaculture production statistics, 1988–1997. FAO Fisheries Circular No. 815, Revision 11. Food and Agriculture Organisation of the United Nations, Rome. FAO (2000) The state of world fisheries and aquaculture, 2000. http://www.fao.org/DOCREP/003/ X8002E/X8002E00.HTM Fushimi, H. (1998) Developing a stock enhancement program based on artificial seedlings: activities of the Japanese Sea-Farming Association (JASFA) in the last decade. In: Nutrition and Technical Development of Aquaculture: Proceedings of the Twenty-Sixth US–Japan Aquaculture Symposium (Ed. by W.H. Howell, B.J. Keller, J.P. Park, et al.), pp. 95–104. UJNR Technical Report No. 26. University of New Hampshire Sea Grant Program, Durham, NH. Gatesoupe, F.J. (1982) Nutritional and antibacterial treatments of live food organisms: the influence on survival, growth rate and weaning success of turbot (Scophthalmus maximus). Ann. Zootechn., 31, 353–368. Girin, M. (1979) Feeding problems and the technology of rearing marine fish larvae. In: Finfish Nutrition and Fishfeed Technology, Vol. I (Ed. by J.E. Halver & K. Tiews), pp. 359–366. Heenemann, Berlin. Govoni, J.J., Boehlert, G.W. & Watanabe, Y. (1986) The physiology of digestion in fish larvae. Environ. Biol. Fish., 16, 59–77. Gross, F. (1937) Notes on the culture of some marine plankton organisms. J. Mar. Biol. Assoc. UK, 21, 753–768. Guillard, R.R.L. & Ryther, J.H. (1962) Studies on marine planktonic diatoms. I. Cyclotella nana Hustedt and Detonula confervacae (Cleve) Gran. Can. J. Microbiol., 8, 229–239. Harrell, R.M. (ed.) (1997) Striped Bass and other Morone Culture. Elsevier, Amsterdam. Hirata, H. (1979) Rotifer culture in Japan. In: Cultivation of Fish Fry and its Live Food (Ed. by E. Styczynska-Jurewicz, T. Backiel, E. Jaspers, et al.), pp. 361–375. European Mariculture Society, Special Publication No. 4, Bredene. Holt, G.J. (2000) Symposium on Recent Advances in Larval Fish Nutrition. Aquacult. Nutr., 6, 141. Houde, E.D. (1972) Some recent advances and unsolved problems in the culture of marine fish larvae. Proc. World Maricult. Soc., 3, 83–112. Huse, I. (1991) Culturing of cod (Gadus morhua). In: Handbook of Mariculture, Vol. II, Finfish Aquaculture (Ed. by J.P. McVey), pp. 43–51. CRC Press, Boca Raton, FL. Jones, A. (1972) Studies on egg development and larval rearing of turbot, Scophthalmus maximus L., and brill, Scophthalmus rhombus L. in the laboratory. J. Mar. Biol. Assoc. UK, 52, 965–986. Jones, D.A., Kamarudin, M.S. & LeVay, L. (1993) The potential for replacement of live feeds in larval culture. J. World Aquacult. Soc., 24, 199–210. Jones, D.A., Kumlu, M., Le Vay, L. & Fletcher, D.J. (1997a) The digestive physiology of herbivorous, omnivorous and carnivorous crustacean larvae: a review. Aquaculture, 155, 285–295. Jones, D.A., Yule, A.B. & Holland, D.L. (1997b) Larval nutrition. In: Crustacean Nutrition (Ed. by L.R. D’Abramo, D.E. Conklin & D.M. Akiyama), pp. 353–389. World Aquaculture Society, Baton Rouge, LA.


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Kolkovsky, S., Koven, W. & Tandler, A. (1997) The mode of action of Artemia in enhancing utilization of microdiet by gilthead seabream Sparus aurata larvae. Aquaculture, 155, 193–205. Laurence, G.C., Smigielski, A.S., Halavik, T.A., et al. (1981) Implications of direct competition between larval cod (Gadus morhua) and haddock (Melanogrammus aeglefinus) in laboratory growth and survival studies at different food densities. Rapp. Procès-verbaux Réunions Cons. Int. Expl. Mer, 178, 304–311. Leber, K.M., Blankenship, H.L., Arce, S.M., et al. (1997) Influence of release season on size-dependent survival of cultured striped mullet, Mugil cephalus, in a Hawaiian estuary. Fish. Bull., 95, 267–279. Léger, P., Bengtson, D.A., Simpson, K.L., et al. (1986) The use and nutritional value of Artemia as a food source. Oceanogr. Mar. Biol. Annu. Rev., 24, 521–623. Liao, I.-C. (1991) Milkfish culture in Taiwan. In: Handbook of Mariculture, Vol. II, Finfish Aquaculture (Ed. by J.P. McVey), pp. 91–115. CRC Press, Boca Raton, FL. Liao, I.-C. & Chien, Y.-H. (1994) Culture of kuruma prawn in Asia. World Aquacult., 25, 18–33. Loosanoff, V.L. & Davis, H.C. (1963) Rearing of bivalve mollusks. Adv. Mar. Biol., 1, 1–136. Lubzens, E., Marko, A. & Tietz, A. (1984) Lipid synthesis in the rotifer Brachionus plicatilis. European Mariculture Society, Special Publication, 8, 201–209. Naylor, R.L., Goldburg, R.J., Primavera, J.H., et al. (2000) Effect of aquaculture on world fish supplies. Nature, 405, 1017–1024. Nien, N.M. & Lin, C.K. (1996) Penaeus monodon seed production in central Vietnam. World Aquacult., 27, 6–18. Otterå, H., Kristiansen, T.S., Svåsand, T., et al. (1999) Enhancement studies of Atlantic cod (Gadus morhua L.) in an exposed coastal area in western Norway. In: Stock Enhancement and Sea Ranching (Ed. by B.R. Howell, E. Moksness & T. Svåsand), pp. 257–276. Fishing News Books, Oxford. Pamplona, S. & Mateo, R. (1985) Milkfish farming in the Philippines. In: Reproduction and Culture of Milkfish (Ed. by C.S. Lee & I.-C. Liao), pp. 141–163. Oceanic Institute, Waimanalo, HI; Tungkang Marine Laboratory, Pingtung, Rebublic of China. Person Le Ruyet, J., Alexandre, J.C., Thébaud, L., et al. (1993) Marine fish larvae feeding: formulated diets or live prey? J. World Aquacult. Soc., 24, 211–224. Person-Le Ruyet, J., Baudin-Laurencin, F., Devauchelle, N., et al. (1991) Culture of turbot (Scophthalmus maximus). In: Handbook of Mariculture, Vol. II, Finfish Aquaculture (Ed. by J.P. McVey), pp. 21–41. CRC Press, Boca Raton, FL. Persoone, G., Sorgeloos, P., Roels, O., et al. (eds) (1980) The Brine Shrimp Artemia, Vol. 3, Ecology, Culturing, Use in Aquaculture. Universa Press, Wetteren. Pillay, T.V.R. (1979) The state of aquaculture 1976. In: Advances in Aquaculture (Ed. by T.V.R. Pillay & W.A. Dill), pp. 1–10. Food and Agriculture Organisation of the United Nations, Rome. Pringsheim, E.G. (1924) Algenkultur. Abderhalden’s Handbuch der biologische Arbeitmethoden, Vol. XI, No. 2, pp. 377–406. Provasoli, L., McLaughlin, J.J.A. & Droop, M. (1957) The development of artificial media for marine algae. Arch. Mikrobiol., 25, 392–428. Reitan, K.I., Rainuzzo, J.R., Øie, G., et al. (1997) A review of the nutritional effects of algae in marine fish larvae. Aquaculture, 155, 207–221. Rognerud, C. (1887) Hatching of cod in Norway. Bull. US Fish Commission, 7, 113–119. Rollefsen, G. (1939) Artificial rearing of fry of seawater fish – preliminary communication. Rapp. Procès-verbaux Réunions Cons. Perm. Int. Explor. Mer, 109, 133. Romdhane, M.S. (1992) Aquaculture in Tunisia. World Aquacult., 23, 27–30. Schreiber, E. (1927) Die Reinkultur von marinen Phytoplankton und deren Bedeutung für die Erforschung der Produktionsfähigkeit des Meerwassers. Meeresunters. Helgoland., N.F., 10, 1–34.

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Seale, A. (1933) Brine shrimp (Artemia) as a satisfactory food for fishes. Trans. Am. Fish. Soc., 63, 129–130. Shelbourne, J.E. (1964) The artificial propagation of marine fish. Adv. Mar. Biol., 2, 1–83. Shetland Salmon Farmer’s Association & North Atlantic Fisheries College (2000) Code of Best Practice for Shetland Salmon Farming. SSFA Code of Best Practice and Appendices, March 2000, Version 1.0. North Atlantic Fisheries College Publication. Shetty, H.P.C. & Satyanarayana Rao, G.P. (1996) Aquaculture in India. World Aquacult., 27, 20–24. Sproul, J.T. & Tominaga, O. (1992) An economic review of the Japanese flounder stock enhancement project in Ishikari Bay, Hokkaido. Bull. Mar. Sci., 50, 75–88. Stickney, R.R. (1996) Aquaculture in the United States: A Historical Survey. John Wiley & Sons, New York. Støttrup, J. (2000) The elusive copepods: their production and suitability in marine aquaculture. Aquacult. Res., 31, 703–711. Tsukamoto, K., Kuwada, H., Hirokawa, J., et al. (1989) Size-dependent mortality of red sea bream, Pagrus major, juveniles released with fluorescent otolith tags in News Bay, Japan. J. Fish Biol., 35A, 59–69. Vadstein, O. (1997) The use of immunostimulation in marine larviculture: possibilities and challenges. Aquaculture, 155, 401–417. Vadstein, O., Øie, G., Olsen, Y., et al. (1993) A strategy to obtain microbial control during larval development of marine fish. In: Fish Farming Technology (Ed. by H. Reinertsen, L.A. Dahle, L. Jørgensen, et al.), pp. 69–75. A.A. Balkema, Rotterdam. Van der Meeren, T. and Naas, K.E. (1997) Development of rearing techniques using large enclosed ecosystems in the mass production of marine fish fry. Rev. Fish. Sci., 5, 367–390. Walne, P.R. (1974) Culture of Bivalve Mollusks: 50 Years Experience at Conwy. Fishing News Books, Farnham. Wassef, E.A. (2000) Status of aquaculture in Egypt. World Aquacult., 31, 29–32, 60–61. Watanabe, T., Arakawa, T., Kitajima, C., et al. (1978) Nutritional quality of living feed from the viewpoint of essential fatty acids for fish. Bull. Jpn. Soc. Scient. Fish., 44, 1223–1227. Wells, W.F. (1920) Artificial propagation of oysters. Trans. Am. Fish. Soc., 50, 301–306. Wells, W.F. (1927) Report of the experimental shellfish station. Report NY State Conserv. Dept, 16, 1–22. Yamashita, Y., Nagahora, S., Yamada, H., et al. (1994) Effects of release size on survival and growth of Japanese flounder Paralichthys olivaceus in coastal waters off Iwate Prefecture, northeastern Japan. Mar. Ecol. Prog. Ser., 105, 269–276. Yúfera, M., Pascual, E. & Fernández, C. (1999) A highly efficient microencapsulated food for the rearing of early larvae of marine fish. Aquaculture, 177, 249–256.

Chapter 2

Production and Nutritional Value of Rotifers Esther Lubzens and Odi Zmora

2.1 Introduction For almost four decades, rotifers have been used as food organisms for cultured marine fish larvae. A continuous, stable and reliable supply of nutritionally adequate rotifers is the key to the flourishing culture of marine finfish in various parts of the world. Major fish species produced today using rotifers during the early developmental stages include yellowtail (Seriola quinqueradiata), red sea bream (Pagrus major), Asian sea bass (Lates calcarifer), turbot (Scophthalmus maximus), mullet (Mugil cephalus), pufferfish (Fugo rubripes), gilthead sea bream (Sparus aurata) and the European sea bass (Dicentrarchus labrax) (FAO 1998). Rotifers are also used as food for culturing penaeid shrimp (Samocha et al. 1989) and crabs (Keenan & Blackshaw 1999). Rotifers serve as a ‘living capsule’, providing the nutrients required by the cultured marine fish larvae for proper development. The incidental choice of rotifers (Ito 1960; see reviews by Hirata 1980; Nagata & Hirata 1986; Hagiwara et al. 2001) as food for early developmental stages of small-mouthed larvae has, therefore, been proven a success. Rotifers are not, in most cases, the natural food of marine fish larvae, which have at their disposal a wide range of food organisms in their natural habitats. In culture, fish larvae are fed on two or three organisms during the initial 10–30 days of exogenous feeding. These include rotifers of the species Brachionus rotundiformis and/or Brachionus plicatilis and brine shrimp (Artemia) nauplii. Brachionus rotundiformis is also known as the S-type (small-type) rotifer and B. plicatilis as the L-type (large-type) rotifer (Fig. 2.1). Artemia cysts are obtained mostly from natural sources, while rotifers must be cultured in hatcheries, as there is no other large supply of them. Copepods are a major component of the natural diet of marine fish larvae. Attempts to culture copepods have met with some success in recent years (Støttrup & Norsker 1997; Støttrup 2000; Hagiwara et al. 2001; Payne & Rippingale 2001). However, given the relative ease and low cost of culturing rotifers in high densities, it is unlikely that copepod cultures will replace rotifers as an economically viable alternative in the near future. Copepods do have some advantages over rotifers, including the wide range of body sizes both within and between species. In particular, the early stage nauplii and copepodites can be extremely useful as initial prey for species that have very small larvae with small mouth gape at first feeding (e.g. the grouper, Epinephelus sp.). The specific

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Fig. 2.1 Rotifers used in raising marine fish larvae: Brachionus rotundiformis (left) and B. plicatilis (right). (Photograph: Irena Pekarsky.) Table 2.1 A comparison between cultured rotifers (Brachionus plicatilis and Brachionus rotundiformis) and natural zooplankton harvested in the wild as food for early developmental stages of marine fish larvae. Characteristic

Rotifers

Natural zooplankton

Size

⬇90–350 ␮m, depending on species and developmental stage

Variable but includes organisms less than ⬃60 ␮m in size

Body shape

Round and flat, without spines

Variable with many spiny species

Distribution in water column and swimming

Usually planktonic and relatively slow moving

Variable with some benthic species and fast-moving species

Density

Tolerance of high densities

Some species (e.g. copepods) with low tolerance to high densities

Salinity

Tolerance to a wide range of salinities

Species-specific and variable tolerance

Supply

Manipulated and regulated; reliable, depending on culture facilities

Unpredictable; varies daily between locations

Nutritional quality

Can be manipulated and regulated

High but unpredictable and variable

Digestibility

Lorica and eggs not digested

Variable

Transmission of parasites and predators of fish larvae

Minimal

Realistic

Transmission of therapeutic agents and probiotics

Feasible

Doubtful or unlikely

advantages of using rotifers as food for early stages of development of marine fish larvae, as opposed to zooplankton, are summarised in Table 2.1. Rotifers (B. plicatilis) were found important in raising early developmental stages of cultured crustaceans, such as the mud crab (Scylla serrata) up to zoea 5 stage (Baylon &

Production and Nutritional Value of Rotifers

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Failama 1999; Keenan & Blackshaw 1999; Li et al. 1999; Mann et al. 1999; Quinitio et al. 1999) or the marine shrimp (Penaeus semisulcatus) (Samocha et al. 1989). Raising freshwater Brachionus calyciflorus in culture systems has been examined for freshwater, edible and ornamental fish species (De Luca et al. 1990; Awïass 1991; Awaïss et al. 1992a,b; Rico-Martinez & Dodson 1992; Ludwig 1994; Awaïss & Kestemont 1997a,b; Lim & Wong 1997; Isik et al. 1999; Hagiwara et al. 2001). The culture of freshwater rotifer species has not yet had the same impact on aquaculture as that of the marine species, although B. plicatilis rotifers were found to be helpful in culturing freshwater ornamental carp larvae (Lubzens et al. 1987). Huge numbers of rotifers, easily reaching several billions, may be required each day for raising marine fish larvae in commercial hatcheries (Lubzens et al. 1997, 2001). The amount needed ranges from 20,000 to 100,000 rotifers per fish larva during the 20–30 days of culture (Kafuku & Ikenoue 1983; Lubzens et al. 2001). The nutritional quality of the rotifers must be assured and controlled by the use of well-established and tested culture and enrichment methods. Furthermore, the rotifer culturist must be aware of and able to deal with occasional unpredicted events that can lead to low production. These may result from inadequate seawater supply, pollution of the culture water with waste products, quality and quantity of food provided to the rotifers, disease or insufficient biological information. The methods developed for providing an adequate supply of rotifers to a variety of small-mouthed fish larvae rely on extensive studies into their biology, feeding, reproductive strategies, genetics, physiology and biochemistry. This basic information provides the culturists with the tools for adapting rotifers for mass culture techniques and in solving problems arising during their culture. Several reviews have been published on rotifer morphology, biology, taxonomy and culture, and some of them have focused on production of rotifers as live food in aquaculture (e.g. Ruttner-Kolisko 1974; Nagata & Hirata 1986; Lubzens 1987; Fukusho 1989a,b; Lubzens et al. 1989, 2001; Fulks & Main 1991; Clément & Wurdak 1991; Wallace & Snell 1991; Nogrady et al. 1993; Hagiwara et al. 1997, 2001). The present review attempts to provide information on euryhaline rotifers, with a specific emphasis on the genus Brachionus (Family Brachionidae; Order Monogononta) and equip the newcomer to this field with the basic concepts and tools for using rotifers as food for cultured fish larvae. Assistance in evaluating the state of cultures is provided in Appendix I.

2.2 Biology and Morphological Characteristics of Rotifers 2.2.1 General biology The phylum Rotifera (previously known as the Rotatoria; see Ricci 1983) consists of a relatively small group of minute, unsegmented, pseudocoelomate, aquatic invertebrates with bilateral symmetry. Most rotifers are free-crawling or swimming, but sedentary and colonial forms are also known (Ruttner-Kolisko 1974; Pontin 1978; Wallace & Snell 1991; Nogrady et al. 1993). About 2000 species populate freshwater lakes and ponds, but several species are known from brackish or marine waters and from mosses and lichens in moist terrestrial habitats. Although Rotifera is a small phylum, rotifers are extremely important in the freshwater environment, contributing up to 30% of the total plankton biomass. By

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consuming bacteria and/or algae, they form the link between primary producers and secondary consumers or predators such as fish and insect larvae. Rotifers have several distinctive features (Fig. 2.2). They are characterised by an anterior, apical, ciliated corona (‘crown’), functional in swimming and feeding. The presence of the corona distinguishes rotifers from all other metazoans, and the metachronal movements of the cilia in the anterior rotating apparatus, giving the illusion of two turning wheels, is one of the distinctive characteristics of organisms belonging to this group. Their body shape ranges from saccate to cylindrical and typically four regions can be distinguished: a head bearing the corona, a neck of variable length, a body and a foot, typically possessing two toes (but the number may range from none to four) that usually retract during swimming. The pedal glands found in the foot secrete a sticky cement material for temporary attachment of rotifers to the substratum. The body of rotifers is covered by an extracellular cuticle that is a gelatinous secretion of the underlying integument and has no skeletal function (Clément & Wurdak 1991). A dense intracytoplasmic lamina located within the syncytial integument forms the peripheral skeleton that serves for muscle attachment. Species in which extensive regions of the integument are thickened are known as loricated forms (including Brachionidae) and others with a thin, more flexible integument are known as illoricated. The thickness of the body wall has little taxonomic significance, as loricated and illoricated forms may be found within one genus. Several regions with thinner intracytoplasmic lamina provide flexibility in loricated forms. These regions include the corona, foot and articulations between movable spines and the body.

Fig. 2.2 Morphology and inner organisation of a Brachionus sp. female (left) and male (right). a, dorsal antenna; b, bladder; bt, buccal tube; c, corona; e, eye; eg, egg; f, foot; fg, foot gland; g, central ganglion; la, lateral antenna; m, mastax; mu, muscle; o, oesophagous; ov, ovary; p, prostate; pe, penis; s, sensory cirri; sg, stomach gland; st, stomach; t, toe; te, testis; tr, trophy; v, vas deferens. (From Koste & Shiel 1987. Reproduced from Invertebrate Taxonomy, Volume 7 with permission of CSIRO Publishing.)


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Rotifers possess an internal fluid-filled space known as a pseudocoelom that is bound externally by the integument and internally by the epithelial cells of the various organs (digestive, protonephridial and reproductive). There are no respiratory or circulatory systems in rotifers, and the pseudocoelom internal fluid, bathing the internal organs, is equivalent to the circulatory system. Its composition is regulated by the protonephridia and it is replenished by the digestive tract. Rotifers are also characterised by the syncytial structure of their body parts. Cell membranes in tissues disappear after embryonic development, forming multinucleated or syncytial tissues. All individuals of a species have a consistent number of nuclei in each organ. This situation, known as eutely, is also found in nematodes. The total number of nuclei, ranging from 900 to 1000, is fixed for life during embryonic development, indicating a limited capacity for repairing damage.

2.2.2 Taxonomy Taxonomic classification of rotifers is under constant review (Garey et al. 1998; Melone et al. 1998; Segers 1998). Traditionally, the phylum is divided into two superclasses: the Seisona and Eurotatoria, the latter consisting of two classes: the Bdelloidea with two gonads, and the Monogononta with one gonad (Melone et al. 1998). The Monogononta contain over 90% of all rotifer species, with more than 1600 species, in about 95 genera of benthic, free-swimming and sessile forms. All are assumed to be dioecious with one gonad. Females possess one ovary with a vitellarium and males, if they occur, are structurally reduced with a vestigial gut. Males are present for brief periods (a few days or weeks). The family of Brachionidae comprises six genera of common rotifers, including Brachionus with about 25 species of littoral and planktonic rotifers. 2.2.2.1 The genus Brachionus The distinctive morphological characteristics of the genus Brachionus are an oval body that is flattened dorsoventrally, and a lorica with six spines on the dorsal anterior margin (Fig. 2.2; see Section 2.2.3). The foot is very mobile, annulated, fully retractable into the lorica and relatively long. The corona has five distinct lobes with long, sensory bristles between them. A cerebral eye is always present. The mastax (see below) is large with malleate trophi. The oesophagous is thin-walled and the cellular stomach is clearly separate from the intestine. There are two gastric glands of varying shape and the anus is terminal. There are four flame cells along the protonephridial duct on each side of the body connecting with a large urinary bladder. The vitellarium carries several cells (see below) and eggs are carried at the foot opening. One female can carry one to three amictic eggs or several male eggs, and each egg is individually attached to the female body. Usually, a female carries only one or, rarely, two resting eggs and these resting eggs are easily distinguished from other eggs by their darker orange–brown colour and a space or vacuole between the two egg membranes (see below). The family Brachionidae comprises a large number of species, most of them inhabiting freshwater. A few species are known from the marine environment and several from euryhaline waters. Two of these (B. plicatilis and B. rotundiformis; Fig. 2.1) are being used extensively in mass cultures and serve as food for early developmental stages of marine fish larvae. In the early days of devising methods for mass culture of rotifers, the cultured species was identi-

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Fig. 2.3 The lorica of Brachionus plicatilis (left) and B. rotundiformis (right) showing the anterior spines. (Photograph: Irena Pekarsky.)

fied as B. plicatilis showing two morphotypes known as L-type (large type) and S-type (small type). Accumulating data showing that these morphotypes do not differ in size and shape alone led to taxonomic identification as two species, B. plicatilis and B. rotundiformis (Segers 1995). The lorica of female B. plicatilis is significantly larger than that of B. rotundiformis and the anterior spines on the lorica of B. plicatilis are obtuse, in contrast to the pointed spines of B. rotundiformis (Hagiwara et al. 1995b; Fig. 2.3). Chromosome number (Rumengan et al. 1991), allozyme profiles (Fu et al. 1991a,b), reproductive isolation (Fu et al. 1993; Hagiwara et al. 1995b; Rico-Martinez & Snell 1995; Gomez & Serra 1995) and microsatellites (Gomez et al. 1998; Gomez & Carvalho 2000; Boehm et al. 2000) suggest genetic differences between these two species. However, recent data indicate that B. rotundiformis is composed of two genotypes, B. rotundiformis SM and B. rotundiformis SS, with no evidence of gene flow between them (Serra et al. 1998; Fig. 2.4). These studies show that B. plicatilis is a euryhaline, low-temperature group, B. rotundiformis SM is adapted to high temperatures and low salinities and B. rotundiformis SS is adapted to high-temperature and highsalinity conditions. In addition, the groups show different mictic responses to density, salinity and temperature, and mating experiments showed that most copulations occurred within a group and no hybrids could be obtained in laboratory cross-mating experiments. There are also some differences in the way rotifers carry their resting eggs. In B. plicatilis and B. rotundiformis SS, resting eggs are carried outside the female lorica in the same way as amictic or male eggs, while in B. rotundiformis SM they are retained within the body of the female (Serra et al. 1998). Production of resting eggs, their preservation and hatching are further discussed in later sections in this chapter. Caution is advised when considering older publica-


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Fig. 2.4 Differences between Brachionus plicatilis SS (SS), B. rotundiformis SM (SM) and B. plicatilis (L). Average body length (top number) and width (lower number) and their standard deviations (in parentheses) are shown below each rotifer. (From: M. Serra, A. Gomez & M.J. Carmona, 1998. Reproduced from Hydrobiologica 387/388, 373–384. With kind permission of Kluwer Academic Publishers.)

tions, as there is evidence of confusion of B. plicatilis and B. rotundiformis. It may be assumed that B. rotundiformis is the main cultured species in tropical areas.

2.2.3 Morphology and physiology The anterior end, or ‘head’, carries the corona, which is comprised in Brachionus females of two concentric ciliary crowns; the outer one is the cingulum, which is responsible for swimming and the inner one, the pseudotrochus, is composed of cilia organised into membranelles or cirri (Clément & Wurdak 1991). The pseudotrochus sweeps the food particles towards the mouth. The head also carries the ventral mouth opening and several sensory organs, including antennae, cirri and light-sensitive ocelli (Fig. 2.2). 2.2.3.1 Feeding Food is captured by the corona, enters the mouth opening and passes through the buccal tube to the pharynx. The buccal tube is equipped with a variety of sensory receptors and ends in the buccal velum, a supple myelin-like structure whose function is to prevent rejection of food that has reached the pharynx. The longitudinal and circular muscle sheaths surrounding the buccal tube can, however, evoke rejection of the ingested food at this level. The pharynx houses the muscular mastax, where the translucent hard trophi break down

24


the food, probably with the aid of enzymes produced by ‘salivary glands’. The trophi, composed of hard parts, are of extreme taxonomic importance for the characterisation of rotifer families, genera or even species. The food leaving the mastax enters the cuticular region of the oesophagus and passes through the ciliary region before entering the stomach. Large syncytial gastric glands empty into the alimentary tract at the junction between the ciliary oesophagous and the stomach and their secretion aids in the extracellular digestion occurring within the stomach lumen. The stomach leads into the intestine and then into a cloaca, which voids fluid from the bladder or paired protonephridia, and eggs from the oviduct. Sensory receptors are located all along the alimentary canal, with a prominent sensory organ between the trophi on the mastax floor and sensory receptors on the mastax ceiling. The sensory receptors that participate in feeding behaviour are connected either to the brain or to the mastax ganglion, and these centres regulate the movement of circular and longitudinal muscles surrounding the buccal cavity and mastax, respectively (Clément & Wurdak 1991). The most common method of feeding in planktonic brachionid rotifers is by filter feeding. This type of feeding, also described as ‘microphagus feeding’ (Pourroit 1977; Clément et al. 1983), is found in rotifers having a developed ciliary corona and a crushing type of mastax. The type of food consumed, particularly its size, is directly dependent on the size and form of the ciliary apparatus and of the mastax. The rotational movement of the cilia directs a water current containing food particles towards the mouth and those that are suitable are swallowed, indicating a sensory mechanism regulating food selection. Brachionus plicatilis seems to have few food preferences and can reproduce when fed with various species of algae, yeast or bacteria (Clément et al. 1983; Hansen et al. 1997). Three sites were suggested to regulate the intake of food in B. calyciflorus and these are probably also operating in B. plicatilis (Clément et al. 1983). First, the anterior ciliated apparatus composed of tactile and/or chemical receptors may regulate the muscles governing the position of the pseudotrochus cilia in such a way that they would form a screen to keep certain particles from entering the mouth. Secondly, chemoreceptors in the buccal tube may activate the longitudinal and circular muscles. The contraction of these muscles will result in rejection of food particles if they are deemed unsuitable. Finally, the ground particles come into contact with sensory receptors in the mastax, and the chemical or tactile recognition at this site will determine the continuation or cessation of mastax grinding movements. If the food is found to be unsuitable, the sensory receptors will evoke expulsion of the food from the buccal canal. Several studies indicate that food selectivity by suspension feeders such as rotifers is mainly based on prey size (Rothhaupt 1990a,b; Hansen et al. 1997). However, selectivity was reported in several species and depended on cell surface, physiological condition of algal cells and algal motility (summarised in Hansen et al. 1997). The prey size spectrum for B. plicatilis ranges from approximately 1.4 to 4838 ␮m3, or 1.4 to 21 ␮m of equivalent spherical diameter (ESD), with changes in efficiency with different particle size. Optimal grazing was reported on Tetraselmis suecica, with an ESD of 8.3 ␮m. The range covers sizes from bacteria to dinoflagellates (Hansen et al. 1997) and the largest particle size caught by rotifers is dependent on its body size (Hino & Hirano 1980). Rotifers were described as mechanical grazers as they were found to graze non-selectively when offered two algal species with different cell size. Their ingestion rates reflected the exact proportion of the volume of each species of algae (Hansen et al. 1997).


25

Three models have been proposed to describe the effect of food particle concentration on feeding rate of filter-feeding zooplankton (Rothhaupt 1990b). In the rectilinear model, clearance rates increase up to a certain food concentration that is termed the incipient limiting level (ILL). This model represents a feeding mode where numerous small particles can be collected simultaneously, without the interference of the particles with the feeding process until the ILL is reached. Maximal ingestion rates are probably determined by the gut packing and the rate of gut evacuation and, therefore, will be reached at lower food concentration with larger food items. Above the ILL, ingestion rates remain constant. The curvilinear model proposes continuously decreasing clearance rates with increasing food concentration, and is expressed by Michaelis–Menten (Michaelis & Menten 1913) or Ivlev (1960) equations. This model indicates a typical increasing interference with the feeding process and is observed with larger than optimal food particle size. Interference could be indicated by mechanical interference caused by, for example, the increased handling time necessary for processing large prey items, during which there is an interruption in collection of new particles. In this model there is a gradual increase in ingestion rates until a plateau is reached. The filtration or clearance rate is high at low food particle concentrations and declines in a curvilinear manner with the increase in food concentration. Finally, in the sigmoid model, clearance or filtration rates increase sigmoidally with increasing food particle concentration until a maximal value is reached, and then they decrease with any further increase in food particle concentration. The ingestion rate remains constant after the peak value has been reached by the clearance rates. Thus, the filtration rate is adjusted to the food concentration and maintains a full gut at low energy expenditure. Studies have indicated curvilinear (Hansen et al. 1997) or sigmoid (Navarro 1999) feeding models for B. plicatilis and a curvilinear feeding model (but not significantly different from the sigmoid model) for B. rotundiformis (Navarro 1999). In experiments using Nannochloropsis oculata as food for B. plicatilis, the ILL was twice that of B. rotundiformis (approximately 2 ⫻ 106 and 1 ⫻ 106 cells ml⫺1, respectively). Additional ILL values for B. plicatilis were obtained for other food sources; 4–8 ⫻ 106 cells ml⫺1 for feeding with Monochrysis (now Pavlova) lutheri and Saccharomyces cerevisae (Lebedeva & Orlenko 1995), 2.1 ⫻ 106 cells ml⫺1 with Chlorella sp. (probably known today as Nannochloropsis sp.; Hirayama & Ogawa 1972), 1.5 ⫻ 106 cells ml⫺1 with Chlamydomonas and 0.1 ⫻ 106 cells ml⫺1 (Chotiyaputta & Hirayama 1978). A large range (0.64–10 ␮l h⫺1 per individual) in filtration rates was found for B. plicatilis, depending on the species of algae used as food and on the food density (Hirayama 1990). When fed with N. oculata, B. plicatilis rotifers ingested between 60 and 90% of their dry weight per day, while B. rotundiformis consumed 160% of their dry weight per day (Navarro 1999), reflecting their different body size. There were no significant differences for values obtained for filtration and ingestion rates between live (N. oculata) and freeze-dried algae in these experiments (Navarro 1999). 2.2.3.2 Digestion The crushed food particles pass through the oesophagous into the stomach and intestine. The stomach is sometimes pigmented by recently ingested food and the colour depends on the type of consumed food. Gastric glands in the anterior part of the stomach may aid in extracellular digestion. Several digestive enzymes have been reported from B. plicatilis,

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including proteases (Hara et al. 1984a,b; Kuhle & Kleinow 1985; Wethmar & Kleinow 1993), ␣-amylase, laminarinase, cellulase, cellobiohydrolase, lysozyme and ␤-1,3-glucanase (Kuhle & Kleinow 1985, 1989, 1990; Hara et al. 1997). The membrane lining the stomach lumen shows invagination of vesicles that coalesce into vacuoles and large oil droplets towards the interior, probably indicating absorption of the digested food (Clément & Wurdak 1991). 2.2.3.3 Body fluids and excretion As mentioned before, rotifers do not have a respiratory system or a circulatory system and body fluids are located in the pseudocoelom. Like other pseudocoelomates, rotifers exchange gases and dispose of nitrogenous wastes by diffusion through their body surface. Muscular contraction of the body aids in circulating the body fluid. Osmoregulation, at least in freshwater species, is carried out by protonephridia. The protenephridial system comprises two lateral parallel tubules with several fan-shaped flame cells opening into them. The flame cells are equipped with cilia that pump the body fluids and pass them into the tubules, where they are drained into the urinary bladder. The bladder empties the accumulated fluids, by contraction, into the cloaca. Brachionus plicatilis was found to adjust the body fluid osmolarity to that of the external concentrations ranging from 32 to 957 mosmol l⫺1, indicating that they are essentially osmoconformers. However, at an external concentration of 32 mosmol l⫺1, the body fluid osmolarity was 59 mosmol l⫺1, demonstrating that these rotifers are unable to tolerate low external concentrations (Epp & Winston 1977). 2.2.3.4 Movement The body fluids function as a hydrostatic skeleton, interacting with the muscular system. The muscular system consists of striated and smooth muscles occurring in small longitudinal and circular bands. They are inserted onto the integument, or join the integument and internal organs. In loricate species such as the Brachionidea, the contraction of muscles inserted on the integument and the resistance of the body fluids facilitate the movement of the animal in the water, with longitudinal muscles shortening and circular bands elongating the shape of the body. Contraction of longitudinal muscles that are inserted on the corona or foot facilitate their retraction into the lorica under unfavourable conditions or during swimming. Muscles inserted in the viscera, or forming part of the viscera, facilitate the contraction of organs such as the digestive gland, vitellarium and urinary bladder. 2.2.3.5 Nervous system and sensory organs The co-ordination of all of these functions is under the regulation of a nervous system that consists of a single large cerebral ganglion (‘brain’) that is located in the anterior part of the body, dorsally below the corona. Paired, ventral neurons proceed from the brain along the length of the body into the foot, branching off to various organs. There are a few ganglia located on the mastax and foot and at the exit points for lateral nerves. The sensory organs can be divided into mechanoreceptors, chemoreceptors and photoreceptors. Mechanoreceptor bristles are located on the corona and on antennae distributed on various regions of the


27

integument, including the foot region. The corona also carries chemoreceptors that function in accepting or rejecting food particles. These also function in copulation in males, by detecting the female’s pheromone (see Section 2.2.4.1). Several species possess one or more pigmented photoreceptive eyespots. A red eyespot is very distinctive in the anterior part of Brachionus species that have been fed on algae.

2.2.4 Reproduction The reproductive organs of female Monogononts are, as the name implies, composed of a single gonad. The gonad consists of the syncytial ovary that contains the ovocytes, the yolkproducing syncytial vitellarium and the follicular layer that surrounds the ovary and vitellarium and forms an oviduct leading to the cloaca. The total number of ovocytes is present at birth. Rotifers are generally oviparous, with embryos developing outside the maternal body. 2.2.4.1 Asexual and sexual reproduction Nearly all rotifers seen in nature are females. Males occur only for short periods and in many species have never been observed. During favourable conditions, the population increases through diploid parthenogenesis, whereby diploid females produce diploid eggs known as amictic eggs. Monogonont species (e.g. B. plicatilis; Fig. 2.5) can reproduce either by parthenogenesis or through sexual reproduction. In general, rotifers of this group reproduce by cyclic parthenogenesis, meaning that asexual reproduction is prevalent, but under certain circumstances sexual reproduction may occur. Females are always diploid and males, when they appear, are haploid and very much reduced in size compared with females. Diploid females can either be amictic or mictic and morphologically they are indistinguishable. Amictic females produce parthenogenetically diploid eggs that develop mitotically into females, while mictic females produce, parthenogenetically, haploid eggs via meiosis. Thus, it is easy to obtain genetic clones from cultures originating from one amictic female. If a mictic female does not mate and is not fertilised, the haploid eggs form into males, but a mated mictic female that is fertilised will form diploid resting eggs (subitaneous eggs or cysts). Parthenogenically formed eggs (diploid or haploid eggs) will develop immediately into embryos and hatch. Resting eggs will hatch under appropriate conditions into amictic females, after a dormant period (Hagiwara 1996; Lubzens et al. 2001). Thus, amictic females produce parthenogenically diploid amictic eggs and mictic females produce parthenogenically haploid male eggs or sexually diploid resting eggs that hatch into diploid amictic females. In some rotifer genera (e.g. Asplanchna, Conchilus and Sinantherina), amphoteric females that produce diploid and haploid eggs have been observed (King & Snell 1977). Production of resting eggs or subitaneous eggs via parthenogenesis and the occurrence of diapausing amictic eggs have also been observed (see discussion in Nogrady et al. 1993; Gilbert & Schreiber 1995). In the Brachionidae, the eggs are attached to the body of the female by a thin thread, and the embryos of amictic eggs hatch and are released from the maternal body leaving the egg shells still attached to their mother. Resting eggs at the initial stages of formation cannot be distinguished from those of amictic eggs. In B. plicatilis, where the resting eggs are formed outside the female’s body (Fig. 2.5), they are also attached by a thin thread to their mother until the end of their formation and later

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Fig. 2.5 Schematic explanation of sexual and asexual cycles of reproduction in Brachionus plicatilis. (Photograph: Gidon Minkoff and Esther Lubzens.)

released and sink to the bottom of the culture vessel, pond or lake sediment. However, the resting eggs of B. rotundiformis (SM type rotifer strains) that develop within the maternal organism are not released from it. They will sink to the bottom of the culture vessel, pond or lake sediment with the death of their mother and are finally released only after the decomposition of her body. Resting eggs survive for long periods and have been hatched from sediment samples more than 60 years after their formation (Kotani et al. 2001). Males are known from a limited number of monogonont species, but it is generally assumed that all members of this group are capable of producing males, including the


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brachionid species, B. plicatilis and B. rotundiformis. Males are much smaller than females and typically very fast moving. They have a rudimentary digestive gut and a sac-like testis containing free-swimming spermatozoa. A vas deferens leads from the testis to a penis and one or two prostate glands discharge into it. Males attempt copulation with amictic or mictic females and mating occurs at the region of the corona or cloaca. Successful fertilisation occurs in newly emerged females for a very limited period (Snell & Hawkins 1983). A pheromone produced by females was identified first in B. plicatilis. It is a 29 kDa glycoprotein and is found in amictic and mictic females (Snell et al. 1988, 1995; Snell & Nacionales 1990). Its molecular structure probably differs from that of B. rotundiformis and this may serve as a species-specific barrier limiting interbreeding between these species (Kotani et al. 1997). The mode of cyclical parthenogenesis combines the advantages of rapid clonal propagation via diploid, ameiotic parthenogenesis with the advantages of sexual recombination and is found in monogonont rotifers and cladocerans and in some other animal phyla such as aphids. These groups tend to inhabit time-varying environments that may become periodically unsuitable and therefore must be recolonised (Serra & King 1999). Parthenogenic reproduction is predominant in the initial stages of colonisation as it produces rapid population growth and during this phase the genotype is copied without genetic recombination. Genetic recombination occurs during sexual reproduction that leads to the formation of resting eggs. The resting egg is the life-cycle stage having the greatest capacity to disperse in both time and space. The cues initiating meiosis in diploid mictic females are not well understood, but from laboratory studies they include nutritional, population density, salinity and genetic factors (Pourriot & Snell 1983; Serra & King 1999; Ricci 2001). In response to a mictic cue, the amictic female will produce amictic and mictic daughters. The proportion of mictic daughters of B. plicatilis is variable (18–66%), and depends on food quality and salinity (Lubzens et al. 1985; Lubzens & Minkoff 1988). Snell (1987) reported an average mictic ratio of 21.2 ⫾ 3.84% (mean ⫾ SD) for B. plicatilis and suggested 50% as the highest optimal mictic ratio in nature. Using a demographic model, Serra and King (1999) showed that the frequency of mictic females that will maximise the population long-term fitness depends on population mortality and birth rates. This means that a population may adjust its relative rates of mictic and amictic production in response to environmentally induced changes, and different mixis patterns are expected in different types of habitats or culture conditions. They concluded that intermediate mictic ratios are optimal in density-dependent growth conditions and that optimal mictic ratios are higher when habitat conditions are better. This means that optimal culture conditions will result in higher production of resting eggs. 2.2.4.2 Reproductive rates The intrinsic reproductive rates of rotifer populations (r or G values; see below) increase exponentially with increasing food concentration, and the relationship can be described by a modified Monod model, with a threshold for zero population growth and a plateau for maximal population growth values (Rothhaupt 1990c). This means that the population growth will not increase beyond a specific concentration of food and may even decrease at relatively high food concentrations. The parameters of this model depend on the food

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Table 2.2 Life tables of amictic females (AM), non-fertilised mictic females producing male eggs (M) and fertilised mictic females producing resting eggs (FM). AM

M

FM

Average life span (Σlx/n) Days Hours Range (days)

7.07 ⫾ 0.06 169.71 ⫾ 23.93 4–9

7.45 ⫾ 0.70 178.8 ⫾ 16.8 4–11

11.96 ⫾ 0.44 287.0 ⫾ 10.56 4–20

Preoviposition period Hours % of total lifespan

36.8 ⫾ 12.39 22

41.45 ⫾ 11.21 23

82.0 ⫾ 11.97 25

Oviposition period Hours % of total lifespan

108.8 ⫾ 17.84 64

61.09 ⫾ 16.5 34

107.66 ⫾ 43.7 37

Postoviposition period Hours % of total lifespan

24.08 14

76.26 43

108.00 38

No. of eggs per female Range

21.5 ⫾ 0.43 17–24

15.18 ⫾ 0.8 9–19

3.31 ⫾ 0.19 1–5

Rate of egg production Hours Eggs produced day⫺1

4.75 5.05 ⫾ 1.59

3.63 6.61 ⫾ 0.35

42.11 0.57 ⫾ 0.2

No. of replicates

14

11

35

Brachionus plicatilis rotifers were individually incubated in 1 ml seawater (at a salinity of 9 ppt), fed on Chlorella stigmatophora (3 ⫻ 106 cells ml⫺1) and incubated at 25 ⫾ 1°C under constant illumination. New offspring were separated from their mothers every 8–10 h and placed individually in new wells. The parent females were also transferred every 8–10 h to new wells containing freshly prepared medium. Values are given as means ⫾ SD. The detailed experimental set-up is described in Lubzens and Minkoff (1988, Table 1).

particle size since this determines the relative rate of consumption (or clearance rate, as discussed above) and on the nutritional quality of the food provided to the rotifers. The fecundity of rotifers depends on whether asexual or sexual reproduction takes place. An example given in Table 2.2 shows that B. plicatilis amictic females produced 17–24 eggs during their lifetime, compared with 1–5 resting eggs produced by fertilised mictic females (Lubzens & Minkoff 1988; unpublished results of Experiment 1). In addition, the production of amictic eggs is, on average, 10 times faster (about 5 eggs/day) than that of the resting eggs during a similar oviposition period (approximately 108 h) (Table 2.2). The unfertilised mictic females produced 9–19 eggs, but these did not contribute to the increase in population as they form males. The number of eggs produced by a female (R0) is dependent on the food algal species, with Synechococcus elongates and Tetraselmis tetrathele supporting the highest rates of reproduction (Hirayama et al. 1979). The optimal temperature for culturing rotifers depends strongly on the species, as the optimal temperatures for B. plicatilis (10–30°C) are lower than those for B. rotundiformis (24–35°C) (see reviews by Hirano 1987; Lubzens et al. 1987, 1989; Rumengan & Hirayama 1990; Hirayama & Rumengan 1993). Within each species, differences have been found in the reproductive rates under the same culture conditions, indicating intraspecific variability (Hino & Hirano 1977; Lubzens 1989; Lubzens et al. 1989; Hagiwara 1994).


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2.2.4.3 Sexual reproduction and resting egg formation It is widely accepted that the occurrence of sexual reproduction in B. plicatilis and B. rotundiformis is genetically determined (Hino & Hirano 1976; Snell & Hoff 1985; Hagiwara et al. 1988b; Lubzens 1989). This means that not all rotifer cultures will show sexual reproduction, even if the optimal environmental conditions are provided. The optimal conditions for encouraging the production of resting eggs also differ between B. plicatilis and B. rotundiformis. Moreover, within each species, there is a large variation between cultures in the number of resting eggs they will produce under identical culture conditions. Therefore, the selection of the appropriate strain or culture is imperative for successful production of resting eggs. As mentioned before, the stimulus for initiating sexual reproduction is still poorly understood, although several studies have been conducted to elucidate the environmental requirements for amictic and mictic production. Sexual production has more constraints than asexual reproduction. Sexual reproduction is restricted by population density (Snell & Boyer 1988) and its occurrence requires more optimal conditions of food availability, salinity and temperature than those supporting asexual reproduction (Lubzens et al. 1985, 1993; Snell 1986; Snell & Boyer 1988; Serra & King 1999). Differences have also been found between B. plicatilis and B. rotundiformis in the environmental conditions that encourage resting egg production. It has been shown that lower temperatures and lower salinities are required for B. plicatilis rotifers, while higher temperatures and relatively higher salinities will encourage higher resting egg production in B. rotundiformis (Hagiwara & Lee 1991). One of the dominating factors is the rotifer culture density. While a threshold density (approximately 10 rotifers/ml) is required to attain successful fertilisation, densities exceeding about 150 rotifers/ml result in a lower production of resting eggs (Hagiwara et al. 1997). A semicontinuous system that maintained rotifer density improved resting egg production of B. plicatilis rotifers but reduced resting egg production in B. rotundiformis cultures. Frequent renewal of culture media has been implicated in the reduced production of resting eggs, and the occurrence of a ‘density-dependent factor’ associated with resting egg production has been suggested (Hino & Hirano 1976). To date, this factor has not been identified. Fresh or preserved algae (frozen and/or concentrated) have been used in the production of resting eggs, as well as in mass production of rotifers (Hagiwara et al. 1997). Specific bacterial strains were reported to encourage resting egg production (Hagiwara et al. 1994). Preliminary experiments were reported on testing the effect of invertebrate and vertebrate hormones for encouraging mixis in B. plicatilis (Gallardo et al. 1997, 1999).

2.3 Culturing Rotifers The success of rotifer cultivation is dependent on selecting the most suitable rotifer species or strain for local culture conditions, maintaining water quality in culture tanks and choosing the most appropriate culture technique.

2.3.1 Selection of species and/or strain The selection of the strain is the most crucial step in initiating mass cultures. Size, type of reproduction (asexual versus sexual) and reproductive rates are species or strain specific.

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Culture temperatures, salinities, type of food and its quantity all modulate the type of reproduction and its rates. Mass production of rotifers is better achieved by encouraging rotifers to reproduce asexually, since sexual reproduction results in males and resting eggs. Male are nutritionally inferior to females owing to their fast swimming behaviour and lack of a digestive system, which means they cannot be enriched with essential nutrients required by fish larvae. The number of resting eggs produced by one female is significantly lower than the number of eggs produced parthenogenically, and resting eggs do not hatch immediately. Sexual reproduction can be avoided by using specific genetic strains (Hino & Hirano 1976; Lubzens 1989; Hagiwara et al. 1995b) that do not reproduce sexually or by culturing rotifers at relatively high salinities. In general, frozen or live algae support higher reproductive rates than yeast or dried algae (Lubzens et al. 2001). A careful calculation is required on the optimal trade-off between the higher cost of algae and increased rotifer production. The amount of food that has to be supplied daily to each culture tank depends on the reproductive rate of the rotifers. Usually, 1–4 g of baker’s yeast (or 30% of this weight as dry yeast) is supplied per million rotifers per day. The amount varies according to the temperature of the culture, salinity (more is needed at lower salinities and at higher culture temperatures), rotifer species and rotifer density. It has been calculated that 105–107 yeast cells need to be supplied daily for each cultured rotifer, and 1 g of yeast can produce about 80,000 B. plicatilis or 100,000 B. rotundiformis rotifers (Hirano 1987). The routine practice is to make a daily count of the number of rotifers and the number of eggs they carry in 1 ml samples, calculating the increased daily increment. The rate of reproduction of cultures is determined as r (sometimes referred to as G): r⫽

1 ln ( Nt ⫺ N0 ) T

(2.1)

where T ⫽ duration of culture in days, N0 ⫽ initial number of rotifers and their eggs, Nt ⫽ total number of rotifers and their eggs after T days of culture. The r-values for B. plicatilis strains usually range from 0.23 to 1.15, and for B. rotundiformis values from 0.54 to 1.37 have been recorded, depending on salinity and temperature (Hirayama et al. 1979; Lubzens et al. 1989, 1995a; Hagiwara et al. 1995b; Hansen et al. 1997). The type of food (wet yeast, dry yeast, live, frozen or dried algae; Lubzens et al. 2001) and its amount directly affects reproductive rates. For example, the r-values for B. plicatilis (Nagasaki strain) cultured in seawater at a salinity of 17 ppt and fed with N. oculata, were 0.90, 1.15 or 0.52 at culture temperatures of 25, 30 or 35°C, respectively. Under the same culture conditions, the r-values observed for B. rotundiformis rotifers were 0.54, 0.77 and 1.20 at 25, 30 or 35°C, respectively (Hagiwara et al. 1995b). The maximal r-values were 0.51, 0.55, 0.69, 0.74, 0.92 or 0.96 when rotifers were fed with Cyclotella cryptica, Nitzschia closterium, Monochrysis (now Pavlova) lutheri, Eutreptiella sp., Chlamydomonas sp. or Synechococcus elongates, respectively (Hirayama et al., 1979). As reproductive rates increase in a curvi-linear manner with the food concentration, comparisons between the efficacy of various food sources should be made at the Km or saturation values for each food type (Hansen et al. 1997).

2.3.2 Maintaining water quality in culture tanks The optimal range of pH for culturing rotifers is 7.5–8.5 (Hirano 1987; reviewed in Fulks & Main 1991) and the pH affects the percentage of un-ionised ammonia (NH3-N) in the


33

water. The pH of the cultures plays an important role since the toxicity of NH3-N released from ammonium (NH ⫹ 4 -N) is a function of the pH, temperature and salinity. Owing to the extreme importance of these parameters, it is highly advisable to consult the relevant tables in Bower and Bidwell (1978). The optimal level for ammonia is ⬍1 mg l⫺1 and the acceptable range for ammonia and nitrate levels is 6–10 mg l⫺1. Rotifer cultures require aeration and the dissolved oxygen level should be maintained above 4 ppm (Fulks & Main 1991). Adequate aeration should be provided using perforated polyvinyl chloride (PVC) tubes, air-stones or small-diameter open-ended tubes. Pure oxygen gas should be provided through perforated tubes in high-density rotifer cultures (see below). The use of ozone is extremely useful for high-density rotifer cultures in a recycled system (Suantika et al. 2001). Ammonia levels in this system were reduced by 67%, nitrite levels by 85% and nitrate levels by 67%. Ozone also reduced the number of particles and the number of bacteria in the culture water of B. plicatilis. In general, the water quality parameters are better in B. plicatilis cultures, because they are maintained at lower temperatures than B. rotundiformis. At lower temperatures (20–25°C), more oxygen can be dissolved in seawater (a higher saturation point), there is a lower rate of reproduction of bacteria, resulting in a lower bacterial load in the culture and several contaminant protozoans (e.g. Vorticella sp., Zoothanmium sp. or Euplotes sp.) do not proliferate as quickly as at higher temperatures (⬃25°C or even slightly higher). Moreover, as it takes longer for a culture to collapse, it is possible to take preventive measures to avoid the complete loss of B. plicatilis cultures. There is no need to illuminate rotifer cultures and in most facilities they are exposed to natural daylight. Direct illumination or exposure to sunlight may encourage uncontrolled growth in some filamentous algal species that are not consumed by rotifers, and this may lead to increased pollution of the culture media. 2.3.2.1 Organic particles Surplus food is one of the main contributors to the deterioration in water quality in culture tanks. In addition to its effect on lowering pH, thereby increasing ammonia and nitrates, it encourages bacterial growth. A careful balance must be maintained between the density and number of rotifers and the allotted food ration, to avoid the accumulation of excess organic matter in rotifer culture tanks. The limited capacity of rotifers to filter the culture medium means that this volume should contain all the nutrients required to meet metabolic needs and support reproduction. The problem is particularly acute in cultures with low rotifer density (less than approximately 50 ml⫺1) that are fed with baker’s yeast, as the volume filtered by all the rotifers is not sufficient to clear the water of the food particles. At low rotifer density, especially if the rotifers are only fed once daily, the food is not consumed quickly enough and significant amounts will either remain suspended, forming particles, or sink to the bottom. This can easily be avoided by dividing the daily food ration into four to six meals a day or by continuous feeding using a peristaltic pump. 2.3.2.2 Bacteria and other organisms in the culture tanks When considering the role of bacteria in rotifer culture tanks, two avenues have been explored: first, the success or failure of rotifer cultures in the presence of specific bacterial

34


species and, secondly, those species that may positively or negatively affect the culture of fish larvae when they are introduced to fish larvae tanks along with the rotifers. Opportunistic bacteria that are pathogenic to fish are common in seawater and may proliferate in the seawater used for live food cultures owing to the high loads of organic matter (reviewed in Skjermo & Vadstein 1999). The Vibrio spp. are among the main pathogenic bacteria infecting fish and they grow especially quickly under the low oxygen concentrations that may prevail in rotifer cultures. In these cultures, they may grow more rapidly than other bacteria and eventually dominate the bacterial assemblage. It is extremely difficult to obtain bacteria-free rotifer cultures, as sterilisation methods and the use of antibiotics are not very effective (Maeda et al. 1997; Rombaut et al. 1999a). One way of overcoming this problem is to regulate the bacterial species occurring in the rotifer cultures by introducing selected, non-pathogenic species. Several bacteria species are beneficial to rotifer culture since they produce important metabolites such as vitamin B12 (Scott 1981; Hirayama & Funamoto 1983; Yu et al. 1988, 1989), encourage sexual reproduction and resting egg formation (Hagiwara et al. 1994), and act as a direct source of food (Aoki & Hino 1996; Hino et al. 1997). However, bacteria that are toxic to rotifers have also been reported (Yu et al. 1990). It should also be noted that the culture temperature may have an important effect on both the type and rate of proliferation of bacteria, and this temperature will generally be lower for B. plicatilis rotifer cultures, possibly increasing their stability by reducing the reproductive rate of bacteria. Introducing selected, non-pathogenic bacteria may not only serve to curtail the proliferation of pathogenic bacteria attacking rotifers or fish larvae, but also be used to introduce beneficial microbial fauna (‘probiotics’) into the digestive system of the fish larvae via the rotifers. Probiotics are defined as ‘microbial cells that are administered in such a way as to enter the gastrointestinal tract and be kept alive, with the aim of improving health’ (Gatesoupe 1999). Lactic acid bacteria and Bacillus sp. spores that were introduced into the culture medium of rotifers increased rotifer production and growth of turbot larvae (Gatesoupe 1991, 1993). Alternatively, rotifers can serve as vectors for probionts that will colonise the larval gut (Fjelheim et al. 1999) and prevent disease outbreaks (Grisez et al. 1997). The bacterial microflora of rotifers from culture tanks can be artificially changed by incubation for 1 h in bacterial suspensions consisting of one or more probiotic strains. The probiont bacteria persist in the rotifers’ microflora for 4–24 h (Markridis et al. 2000) and the treated rotifers will transmit this bacterial gut flora to the fish larvae’s guts after ingestion. In addition to bacteria, the culture water may harbour many other organisms such as viruses, fungi or yeast that may be harmful to rotifers. Organisms that may compete with the rotifers for their food, such as ciliates, copepods or cladocerans, may also be present (Colorni et al. 1991; Comps et al. 1991a,b; Maeda & Hino 1991; Hagiwara et al. 1995a; Comps & Menu 1997; Jung et al. 1997).

2.3.3 Choosing the most appropriate culture techniques Rotifers can easily be maintained in all scales of culture. Two types of culture can be distinguished, the first of which are small-scale laboratory cultures for studying rotifer biology and physiology, or for maintaining species and genetic strains. These cultures are maintained as a ‘live library’ in small volumes and serve to initiate mass cultures with


35

specific important traits, such as size, adaptation to high or low culture temperatures, and salinity. The second type comprises mass cultures for supplying rotifers in the quantities required for intensive larval fish production. 2.3.3.1 Small-scale laboratory cultures The aim of these cultures is to maintain their specific genetic traits for long periods and facilitate their availability to mass culture facilities whenever they are needed. The following description demonstrates the culture procedure for rotifer species and strains in the authors’ laboratory using Mediterranean seawater (40 ppt). Rotifers in this type of culture can also be maintained in seawater prepared from dry sea salts or in artificial seawater. Seawater Natural seawater should be filtered through a 0.2 ␮m membrane filter and heat sterilised at 100°C at atmospheric pressure for 30 min to avoid the formation of insoluble precipitates. The cooled, sterile seawater can be kept for several days at room temperature or several weeks in a refrigerator, if the bottles are kept sealed. Sterile distilled water is used for dilution of seawater whenever required. Culture procedure Ehrlenmeyer flasks (100 ml in volume) or 50 ml sterile disposable tubes can be used for culture. Each heat-sterilised flask or sterile tube is filled with 10–20 ml sterile seawater and 40–60 rotifers are added. Usually, a salinity of 30 ppt is suitable for most strains and also ensures that in most cultures asexual reproduction will prevail. A drop of concentrated algae (see below) is added to each culture and the flasks or tubes are incubated at one or more of the following temperatures: 20, 25 or 35°C, depending on the strain or species of the rotifer. Brachionus plicatilis cultures are best kept at 20 and 25°C, while B. rotundiformis cultures should be kept at 25, 30 or 35°C. The cultures are fed ad libitum every 2 days with concentrated algae and the amounts will vary with the reproductive rate of the cultured rotifers. This depends on the species, strain, temperature and salinity, and empirical experience is the best way to determine the appropriate amount. Caution is needed not to provide too much food as this may reduce reproductive rates and result in the collapse of the culture. Cultures are renewed every 7–10 days. To avoid loss of cultures, two or more copies of each culture are maintained at all times, by placing each one at a different culture temperature. The old cultures are kept (even without feeding) at room temperature, as a back-up, until the cultures are renewed. While these cultures do not reproduce at optimal rates, the rotifers remain alive and can be used for replacement in case of failure of the previous renewed cultures. Culture of algae Culture of Nannochloropsis sp. was found to be the most convenient source of food for rotifer cultures. Nannochloropsis sp. originated from Japan and is cultured in 2–3 litre batches at 25°C, under constant illumination. The culture medium is prepared from natural seawater enriched with a modification of Guillard f/2 medium (Lubzens 1981). Algae are

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cultured for 4–6 days and harvested in the log phase period of growth by centrifugation at 4200 g, for 15 min at room temperature. The algal pellet is resuspended in a small volume of culture medium and kept in 1–2 ml aliquots (in Eppendorf tubes) at 4°C for 2–4 days or frozen at ⫺20°C for up to 12 months. 2.3.3.2 Mass cultures Three methods are used today for obtaining large numbers of rotifers: batch cultures, semicontinuous cultures and continuous cultures. Batch cultures This type of culture was initially developed in 1964 (Hirata 1964; for reviews, see Hirata 1979, 1980; Lubzens 1987) and has been greatly modified in recent years by many fish hatcheries (Lubzens et al. 1997, 2001). Four types can be described for this system (Table 2.3). In Type I cultures, rotifers are introduced at low density into ‘green water’ produced in fertilised tanks or ponds. Rotifers are collected (‘harvested’) after all the algae have been consumed, and are used as food for fish larvae. A small number is used to inoculate newly prepared ‘green Table 2.3 Types of batch culture method. Type I

Type II

Type III

Type IV

Place and volume of cultures

Outdoor; 20–100 t

Indoor; 1–5 t

Outdoor; 10–100 t

Indoor; 1 t

Step 1

Growth of phytoplankton (‘green water pond’)

Inoculate rotifers at high density (50–100 ml⫺1) in one-fifth volume of the culture tank

Inoculate rotifers into a large tank at a low density (10–20 ml⫺1)

Inoculate rotifers into 1000 l tank at high density (⬃5000– 10,000 ml⫺1)

Step 2

Inoculate rotifers at low density (1–5 ml⫺1)

Feed with yeast

Feed with yeast

Feed with concentrated Chlorella

Step 3

Culture rotifers until density reaches 20–50 ml⫺1

Increase the volume Culture until rotifer Culture rotifers of the culture density reaches for 2–3 days according to the 100–200 ml⫺1 reproductive rate of rotifers to maintain relative high density

Step 4

Sieve the whole culture to harvest rotifers



Sieve rotifers

Step 5

Reinoculate a new ‘green pond’ with part of the rotifers

Reinoculate a new tank and clean old tank and pipes with chlorine solution

Reinoculate a new culture tank and clean old tank and pipes with chlorine solution

Reinoculate a new culture tank and clean old tank and pipes with chlorine solution

Step 6

Enrich rotifers

Enrich rotifers

Enrich rotifers

Enrich rotifers

Duration of culture

Unpredictable

4–5 days

Depends on reproductive rate

2–3 days


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water’ ponds. A recent adaptation (Lubzens et al. 1997) for batch culture (Type II) relies on a series of tanks. The culture system depends on short (1 week) production cycles with a cleaning interval of tanks and aeration tubing. The culture in each tank is started with one-fifth of its volume with rotifers at a density of approximately 200–300 ml⫺1. Rotifers are fed mainly with baker’s yeast and the daily ration is divided into three to five meals. The volume of culture medium is increased daily with seawater depending on the reproductive rate, thus maintaining a relatively high rotifer density in the culture. This may reach 300–500 rotifers ml⫺1 at the end of the culture period. The final volume of the tank is reached after 4–5 culture days. The rotifers are then sieved and concentrated, and the culture tank and its accessories (sieves, aeration tubes) are sterilised in hypochlorite solution (10 ppm of commercial grade bleach) for 24 h, followed by thorough rinsing with seawater before reuse. The concentrated rotifers are immersed briefly (5–15 min) in dechlorinated freshwater to remove ciliates and other undesired organisms. Practical experience has shown that rotifers are not affected by this brief immersion period in freshwater. Most of the concentrated rotifers are enriched (see below) with essential fatty acids and proteins before being fed to fish or crustacean larvae and a small fraction is used to inoculate the new culture tanks. This system is based on six tanks. Each day, one tank is harvested and another is inoculated. The volumes of the tanks range between 3 and 10 m3, depending on the number of rotifers needed in the hatchery, and their number should take into account an occasional, unpredictable collapse of at least one culture per week. This safeguard can be reduced as experience is gained in the hatchery. Preliminary experiments (Ressem et al. 2001), were reported recently on a 3 day cycle of B. plicatilis batch culture system in 600 litre cylindroconical fibreglass tanks, using 1 g (dry weight) of yeast and 0.1 g (dry weight) of Algamac 2000 (Bio-Marine, Aquafauna, USA) per 106 rotifers per day, at 22–23°C and 20–25 ppt salinity. Rotifers are inoculated at a density of ⬃250 ml⫺1 and harvested after 3 days at ⬃750 rotifers ml⫺1 and the cultures seem very stable, owing to the short culture cycle. One of the advantages of using the Algamac 2000 is that the rotifers are already enriched during their culture period. This is in addition to the low cost of food for this rotifer production system. Another approach to mass culture is based on inoculating rotifers into the culture tank that contains the final volume of culture medium (Type III), at an initial low rotifer density. The rotifers are fed with yeast, other dry food or concentrated algae, and harvesting is performed when high densities (⬎200 rotifers ml⫺1) are reached. This system is less stable than the one described previously since, at the early stages of the culture, the surplus of food remaining in the tanks will encourage bacterial growth and possible reduction in the pH. As the density of rotifers increases, the amount of surplus organic material from food is reduced and the system becomes more stable. The introduction of refrigerated and condensed freshwater Chlorella enriched with vitamin B12 (Chlorella Industry Co., Japan) has facilitated the maintenance of extremely highdensity rotifer mass cultures and changed dramatically the future potential for providing rotifers for fish and crustacean hatcheries (Yoshimura et al. 1996, 1997; Balompapueng et al. 1997a). These high-density culture systems consist of 1 m3 tank units in which rotifers (B. rotundiformis) are batch-cultured at 2–3 day intervals (Type IV). Cultures are initiated at a density of 10,000 rotifers ml⫺1, and after 2–3 days with concentrated algae the culture density reaches 20,000–30,000 rotifers ml⫺1. The large number of rotifers needed for starting the first round of these cultures can originate from scaling-up a small initial culture by

38


traditional mass culture methods or, alternatively, from hatched resting eggs if they are available (see below for further discussion of this issue). Oxygen gas has to be supplied to these cultures to overcome the shortage of dissolved oxygen that results from the high amounts of food and the subsequent increase in the rotifer population. In addition, at these high densities, ammonia excreted by the rotifers becomes a significant problem. The pH of the culture also increases, presumably owing to the liberation of carbon dioxide from the water, by the oxygen gas supplied to the culture. This, in turn, results in a higher proportion of toxic un-ionised ammonia of the total ammonia, which increases with increased pH, salinity and the culture temperature. Regulation of the pH at 7.0 by hydrochloric acid minimises these effects. A special nylon filtration mat is used for removal of large amounts of suspended organic material that may also include protozoans, fungi and bacterial flocculations, which may be harmful to the fish larvae. The small space and reduced labour required are the main advantages of this ultra-high-density culture system. In hatcheries, this relatively compact, closed system permits the maintenance of separate cultures of several genetic rotifer strains, or populations of different sizes. Thus, it assists in providing a variety of rotifers for larvae of different fish species, without enlarging the hatchery facilities. Fish larvae prefer smaller rotifers immediately after hatching and larger ones as they grow, and providing them with this size variation increases their survival (Lubzens et al. 1989). Most hatcheries aim to culture a variety of fish species that may differ in their nutritional requirements during early life stages, and these can be more easily provided by the compact, high-density cultures. The relatively high culture temperatures (⬎25°C) required by small rotifers can be easily maintained in these compact systems. At the same time, fish cultured at low temperatures can benefit from this system by using rotifers cultured at relatively low temperatures. The convenience and accessibility of concentrated algae encourage its use also in intensifying traditional culture systems. Semi-continuous cultures This culture system relies on periodic (usually daily) harvesting of rotifers by removal of part of the culture medium and replacing it with new seawater (see Hirata 1980). This method has been termed the ‘thinning method’ (Fukusho 1989b). The volume removed depends on the reproductive rate of the rotifers and harvesting removes only the number of rotifers gained by reproduction from the previous harvesting period. Hirano (1987) suggested that 6–7% of the biomass can be removed daily in rotifer cultures maintained on baker’s yeast. In general, these systems often rely on large volume tanks ranging from 3000 to 300,000 litres (Hirata 1980; Fukusho 1989b; Lubzens et al. 1997). These cultures are characterised by relatively lower rotifer density (100–300 rotifers ml⫺1) and baker’s yeast is used as food. Cultures continue for several days or even weeks but, eventually, excretory products (solid wastes and nitrogenous products) that accumulate in the tank lead to their collapse. Continuous removal of the solid waste enhances rotifer reproduction and results in densities reaching 400 individuals ml⫺1 and the cultures are stable for over 30 days with daily harvesting periods. In these ‘feedback’ cultures (Hirata 1980), the solid waste is transferred to decomposing tanks and, after its breakdown to nitrogen, carbon and phosphates, it is used as fertiliser for algal cultures. However, as mentioned previously, these large-volume tanks harbour many other organisms that either compete for food with the rotifers (e.g. ciliates, copepods, cladocerans) or harm them (e.g. viruses, bacteria, fungi or yeast). Since these


39

and other pathogens can be transferred to the fish tanks (Gatesoupe 1990; Blanch et al. 1991; Verdonck et al. 1997; Munro et al. 1999), the cultivation of rotifers with selected, non-pathogenic bacterial strains has been suggested (Gatesoupe 1999; Markridis et al. 1999, 2000; Rombaut et al. 1999a,b) to curtail the growth of undesired micro-organisms and contribute to the stabilisation of the rotifer cultures (for further details see Section 2.3.2.2). High-density semi-continuous culture, using a modified commercially available formulated rotifer diet with recirculation of the culture media, was described recently (Suantika et al. 2000). This system consists of a 100 litre rotifer (B. plicatilis) culture tank, a settlement tank for suspended particulate matter, a protein skimmer and a biofilter. The amount of food provided to the rotifers in this system is adjusted to the circulation flow rate and loss of feed by the protein skimmer. Rotifer cultures are initiated at a density of approximately 250 rotifers ml⫺1, with the density reaching 3000 ml⫺1 after 8 days of culture and being maintained at about this level for over 1 month. About 20% (3–6 ⫻ 107 rotifers) of the standing stock is harvested daily. Adequate water quality is maintained by a daily 500% recirculation rate. Although it is possible to obtain densities of 8000 rotifers ml⫺1 after 8 days of culture following inoculation, the authors recommend that rotifer production should be performed at a lower density, where the system is more stable. The total rotifer production in this system is 1.7 ⫻ 109 rotifers over 32 culture days. Introducing ozone into this system (Suantika et al. 2001) improved its performance significantly by supporting a higher rotifer biomass (16,000 rotifers ml⫺1) and prolonging by 4 days the duration of rotifer production. An experimental, semi-continuous culture system using dried Nannochloropsis powder was described recently (Navarro & Yufera 1998). In this system, rotifers were cultured in 1 litre flasks, with gentle aeration in a thermoregulated chamber at 25°C, at a salinity of 18 ppt and under 2500 lux of continuous illumination. They were provided daily with 25 mg of dried N. oculata powder and the effect of daily dilution rates of 0.1, 0.2, 0.3, 0.4, 0.5 and 0.6 was investigated to establish the best production (mg rotifers day⫺1) and food conversion efficiencies (mg rotifer developed mg⫺1 microalgae consumed) for B. plicatilis or B. rotundiformis. The results showed that the optimal dilution rates were 0.2 and 0.3 for B. rotundiformis and B. plicatilis, respectively. At these dilution rates, the rotifer production reached 16.75 ⫾ 0.37 mg l⫺1 day⫺1 for B. plicatilis and 6.12 ⫾ 0.08 mg l⫺1 day⫺1 for B. rotundiformis. The average conversion efficiency at these conditions was 0.76 and 0.30 mg rotifers mg⫺1 algae for B. plicatilis and B. rotundiformis, respectively. This means that under optimal conditions, 76 and 30% of the microalgae biomass was transformed into rotifer biomass of the respective species. The lower efficiency for B. rotundiformis may be attributed to the relatively low culture temperature (25°C) used for this thermophilic species. The application of this system on a large scale awaits additional experimental results. Continuous cultures These cultures are based on the chemostat or turbidostat models of micro-organisms and are fully controlled (temperature, pH, oxygen supply and density of cultured organisms) and highly dependable (Walz 1993; Walz et al. 1997). They offer easy manipulation of rotifer physiological and nutritional quality. Log-phase produced rotifers can be harvested continuously and their nutritional quality is maintained by providing adequate food organisms (James et al. 1983, 1987; James & Abu-Rezeq 1989a,b, 1990). A more recent adaptation of this method involves high-density cultures (excess of 10,000 rotifers ml⫺1) using the

40


concentrated freshwater Chlorella (Fu et al. 1997). Part of the efficiency of rotifer production in these intensive, chemostat-like systems may be attributed to bacterial growth and consumption by the cultured rotifers. Bacterial nitrogen corresponding to approximately 20% of the algal feeding was consumed by rotifers in systems operating with unlimited food supply (Aoki & Hino 1996; Hino et al. 1997). These systems are compact (1000 litres of culture provides 1.7–3.5 billion B. rotundiformis daily, and 500 litres provides 0.13–0.27 billion B. plicatilis daily). However, their initial cost exceeds that of more conventional installations and they depend on a constant supply of concentrated algae. While more work is needed to optimise the culture system for B. plicatilis, it should be noted that their biomass (depending on their size) is at least three times higher (expressed as dry weight; Yufera et al. 1997) than that of B. rotundiformis. This means that a similar biomass per culture volume will consist of a smaller number of individuals. Moreover, the lower number of B. plicatilis produced per day may also be attributed to their lower metabolism and rate of reproduction at the optimal culture temperatures (20–24°C). Culture tanks A very large array and configuration of culture tanks is used for semi-continuous and batchculture rotifer mass production systems, in terms of both volume and shape, and these have been summarised by Fulks and Main (1991, pp. 323–326). They are round, square, cylindrical, conical or rectangular, and mass cultures are performed in volumes ranging from 150 to 300,000 litres (or 300 m3). An example of an indoor 100 m3 rotifer culture tank is shown in Fig. 2.6 and an example of a 500 litre cylindrical tank with a conical outlet is shown in Fig. 2.7. These culture tanks are made from concrete, fibreglass, polycarbonate or plastic, or nylon disposable bags hanging on metal frames. The chemostat-type, continuous culture system described by James & Abu-Rezeq (1989a,b) relies on 1000 litre culture tanks. The high rotifer density cultures relying on

Fig. 2.6 Concrete rectangular culture tank with maximum capacity of 100 m3 seawater used in culturing rotifers in Japan. (Photograph: Esther Lubzens.)


41

concentrated Chlorella cells use 100 or 1000 litre cylindrical tanks (Fu et al. 1997), and the system designed by Suantika et al. (2000) consisted of 100 litre cylindroconical tanks. The large 6–300 m3 concrete tanks are usually used for outdoor, semi-continuous cultures, where rotifer density is relatively low. In some places, these tanks have a conical bottom with an outlet that permits the removal of accumulated debris at least once a day, or is used for harvesting of rotifers. These tanks are difficult to clean, and bacteria or other organisms may accumulate in crevices that are abundant on their rough surface. For this reason, the old system of rotifer culture tanks at the National Mariculture Center (Eilat, Israel), consisting of six, 20 m3, concrete, circular tanks (Fig. 2.8) has been abandoned and replaced with an adaptation of the system devised at the Israel Salt Co. (Lubzens et al. 1997). This

Fig. 2.7 Cylindrical 500 litre fibreglass culture tanks, with conical bottom outlets for culturing rotifers at the National Mariculture Center, Eilat, Israel. (Photograph: Esther Lubzens.)

Fig. 2.8 Round 20 m3 concrete tanks for culturing rotifers at the National Mariculture Center, Eilat, Israel. Note the small tank at the lower part of the picture, near the outlet of the concrete tank, used for rotifer collection (or ‘harvesting’). (Photograph: Esther Lubzens.)

42


Fig. 2.9 Rectangular bathtub-shaped fibreglass tanks (3000 litres) for culturing rotifers (Salt. Co., Israel). Note the cylindrical tanks with conical outlets in the background used for nutrient enrichment of rotifers or Artemia. (Photograph: Esther Lubzens.)

alteration also involved changing from a semi-continuous culture system to batch culture production. Most batch culture or continuous systems are performed in relatively small volumes, ranging from 150 to 3000 or even 10,000 litres. The smaller volumes (150–2000 litres) can be cultured in nylon disposable plastic bags that are placed on a circular or rectangular metal frame and have the great advantage of being used only once for 2–6 days. The most reliable system in Israel consists of smooth-surface rectangular ‘bathtub’-shaped tanks made from fibreglass (Fig. 2.9), with volumes ranging from 3000 to 10,000 litres. The tanks are shaped with a slanting bottom, leading to an external outlet that facilitates removal of debris and rotifers. The tanks can be easily cleaned with bleach and freshwater every 5–7 days, at the end of each batch culture. One of the advantages of the smaller volume batch cultures is that they can be placed indoors (in contrast to the large-volume, concrete tanks that are usually placed outdoors), thus avoiding the colonisation of the rotifer cultures with other organisms. The decision on the choice of the type and shape of culture tanks depends greatly on the available facilities and budget. For example, the use of fibreglass rectangular or circular tanks has proved to be beneficial in the authors’ facilities. The volume of the culture tank depends on the expected production in the hatchery and the convenience of handling. Rotifers have to be counted and inspected daily from each culture tank, independent of the volume. This means that it is more cost-effective to maintain a small number of large-volume tanks than a large number of small-volume tanks. However, the accidental loss of a culture in a large-volume tank will obviously entail the loss of a larger number of rotifers. Taking into consideration the greater reliability of the batch culture system, the policy that can be adopted by a new facility is to plan on a tank containing a volume that will suffice for 1 day’s supply of rotifers in the hatchery. This will mean that the hatchery’s culture will consist of six or seven tanks, where every day one tank is harvested and one tank is inoculated with a new culture (see also Lubzens et al. 1997). New culture facilities should also maintain one extra tank, until the culture systems have become completely reliable and predictable.


43

Choosing the best system Cost of production, reliability and the practical experience of the staff are the main factors that dictate the choice of system. The cost of production of rotifers depends greatly on the total scale of production, with an estimated cost of US $0.04 per million rotifers in a system producing 4 billion per day, and US $0.15 in a 1 billion per day production system (Lubzens et al. 1997). The production rate can also be improved by using concentrated algae. Although these are more expensive than yeast, they offer several advantages: higher rotifer reproductive rates, improved stability of rotifer cultures, allowing them to be used in high-density culture systems, and higher resting egg production. The cost of concentrated 18 litre canisters containing concentrated Chlorella at a density of 20 billion cells ml⫺1 ranges from US $140 to 150 (Hagiwara et al. 2001). Recent reports (Hayashi et al. 2001) on successful incorporation of n-3 highly unsaturated fatty acids (HUFA) by Chlorella cells will eliminate the need to enrich rotifers mass cultured with this alga, prior to feeding them to the fish larvae. Several other companies have been involved in marketing concentrated algae with prices ranging from US $300 to 400 kg⫺1 of dried algae or US $50–65 for 1 litre of concentrated algae (16–20% dry matter), depending on the species, purchasing quantities and shipment destinations. In Europe, North America and Japan, the cost of labour is one of the main concerns in these calculations, whereas the cost of equipment is the main concern in developing countries. The reliability of each system dictates the number of replicate culture tanks that should be installed to meet the required production, with fewer replicate cultures needed in more reliable systems.

2.4 Advanced Warning on State of Cultures Evaluating the physiological state of the rotifer culture is extremely important in hatcheries since larval production depends on a predictable and reliable daily supply of rotifers. Six parameters have been used as early warning signals for the state of the culture and these may be strain specific.

2.4.1 Egg ratio The number of eggs predicts the state of the culture for the forthcoming 24 h. Therefore, the egg ratio (E/N), determined as the number of eggs (E) divided by the total number of females (N) in a sample, is an important indicator of the status of a culture. Usually, a sample of 30–40 ml or even larger is removed daily from each culture for determining the state of the culture (see Appendix I) and for evaluating the amount of food that should be provided to that specific culture. The total number of females and the total number of eggs in 1 ml subsamples (from the 30–40 ml sample) is counted and the ratio E/N is calculated for each sample. Three different 1 ml aliquots should be counted to obtain a more reliable estimate and more aliquots should be counted if the variation exceeds 10% between the counted rotifers in three aliquots. In general, the egg ratio depends on food quality and quantity and is affected by abiotic parameters such as oxygen level, culture temperature, salinity, pH and ammonia levels. Any deviation from optimal conditions of these parameters will be

44


reflected by low egg ratios in the rotifer cultures, except that a short duration at low temperatures will not reduce the egg number. The critical ratio is strain specific. For example, in one study an egg ratio of less than 0.13 for B. plicatilis cultures indicated their instability and possible future collapse (Snell et al. 1987). Therefore, a low or high egg ratio should be considered as one of the indicators, in addition to others, of the state of a rotifer culture.

2.4.2 Swimming velocity Swimming velocity is a quick indicator of the current state of the culture (Snell & Hoff 1988; Korstad et al. 1995). It is reduced at high un-ionised ammonia concentration and at starvation. Extremely low or high values of temperature and pH have the same effect.

2.4.3 Ingestion rate Ingestion of fluorescent labelled beads by neonates hatching from eggs was reported to be related to water quality parameters (Juchelka & Snell 1994). An adaptation of this test to testing water from rotifer mass cultures should be considered in the future.

2.4.4 Viscosity It has been demonstrated that the relative viscosity of the culture medium increases with the age of the culture, resulting in reduced swimming speed, ingestion rates, mean longevity and mean number of offspring of the rotifers. Therefore, direct measurements of viscosity could be an indicator of approaching problems in maintaining culture stability. This phenomenon is of great importance in the high-density continuous cultures, where the culture media contain high concentrations of concentrated algae and excretory products (Hagiwara et al. 1998).

2.4.5 Enzyme activity Changes in the activity of endogenous esterases, phospholipases and glucosidases, has been suggested as a monitoring tool for water quality in rotifer culture tanks (de Araujo et al. 2000, 2001; Hagiwara et al. 2001). The tests are based on reduced enzyme activity in rotifers exposed to toxins (Burbank & Snell 1994; Snell & Janssen 1995, 1998). In these tests, rotifers of similar age, size and physiological condition are provided from hatched resting eggs. It is not yet known whether rotifers removed from mass cultures are suitable candidates for these tests.

2.4.6 Diseases Occurrence of disease will eventually lead to the collapse of the culture and, therefore, early detection is important for taking appropriate measures. Rotifers in mass cultures have


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been observed to be infected by fungal, viral and yeast-like organisms (Colorni et al. 1991; Comps et al. 1991a,b, 1993; Comps & Menu 1997; Zmora 1991). As mentioned before, the culture temperature may affect the type of bacteria or other opportunistic organisms and the rate of their proliferation. Therefore, the incidence of specific infections, their type and rate of infection may vary between B. plicatilis and B. rotundiformis, as they are cultured at relatively low or high temperatures, respectively. While there are no known remedies for rotifer diseases, it has been observed (Zmora 1991) that providing Nannochloropsis to infected yeast-fed rotifer cultures will reduce the infection rate. The algae do not cure the rotifers, but the accelerated reproduction of the cultured rotifers fed algae results in new generations that show a low incidence of the disease (O. Zmora, unpublished results). These cultures remain susceptible and the diseases may recur if the cultures are exposed to additional stress. Swimming speed, egg ratio and health conditions must be inspected daily on samples removed from each culture tank.

2.5 Nutritional Quality of Rotifers After establishing mass culture techniques, it is necessary to ensure that the rotifers are nutritionally adequate for the fish larvae. This is usually performed by a step known as ‘enrichment’, where the rotifers are collected or harvested from the culture tanks into containers where they are kept at very high densities (usually more than 100,000 ml⫺1), and incubated for 8–20 h with enrichment dietary components that are specifically required by the fish larvae. In addition to enrichment with protein, lipids or carbohydrates, the rotifers can be enriched with antibiotics (Verpraet et al. 1992) or with probiotic bacteria (Markridis et al. 1999, 2000).

2.5.1 Number of rotifers consumed by larvae The nutritional value of rotifers depends on their dry weight, caloric value and chemical composition (reviewed in Lubzens et al. 1989). Amictic rotifer eggs and the loricae of rotifers are not digested by larval fish in their early developmental stages (Lubzens et al. 1989). Dynamic, physiological processes such as satiation, starvation and reproduction also affect the chemical composition of rotifers (Yufera & Pascual 1989; Olsen et al. 1993; Øie & Olsen 1997; Yufera et al. 1997; Markridis & Olsen 1999). The number of rotifers consumed by the larvae determines the quantity of food reaching their gut. In red sea bream, the number of rotifers consumed daily increases with the size or age of the larva, from 55–72 rotifers per 3.9 mm length larva to 4700 per 11.4 mm length larva (Fukusho 1989b; Fig. 2.10). There is a large variation in sizes between B. plicatilis and B. rotundiformis rotifers and between various populations within each species (Fu et al. 1990, 1991a,b; Hagiwara et al. 1995b), and this is reflected in their nutritional quality and consumption by fish larvae (Lubzens et al. 1989; Polo et al. 1992). Moreover, size changes occur during the life cycle of rotifers within each population or species, with rotifers growing in size from the time of hatching from the amictic egg until they reach sexual maturity. Size also depends on culture conditions such as salinity, temperature or diet (Snell & Carillo

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Fig. 2.10 Number of rotifers consumed daily by fish larvae: red sea bream, striped knifejaw and black sea bream. (Adapted from Fukusho 1989b.)

1984). All of these factors contribute to the difficulties in comparing results from different publications.

2.5.2 Dry weight and caloric value The dry weight of rotifers depends on their size and nutritional state (Lubzens et al. 1989). At approximately 600–800 ng, B. plicatilis rotifers are three to four times heavier than B. rotundiformis (approximately 200 ng) and this changes with their reproductive rate (Yufera et al. 1997). The caloric value was found to depend on the diet and ranged from 1.34 × 10⫺3 cal per rotifer fed on baker’s yeast to 2.00 × 10⫺3 cal per rotifer after 6 h enrichment with a formulated enrichment diet (Fernandez-Reiriz et al. 1993).

2.5.3 Biochemical composition 2.5.3.1 Protein and carbohydrate content Rotifer protein content ranges from 28 to 63% and lipid content from 9 to 28% of the dry weight (Lubzens et al. 1989; Frolov et al. 1991; Frolov & Pankov 1992; Nagata & Whyte 1992; Fernandez-Reiriz et al. 1993; Reitan et al. 1993; Rainuzzo et al. 1994; Øie & Olsen 1997; Markridis & Olsen 1999). The carbohydrate content ranges from 10.5 to 27% of the dry weight (Whyte & Nagata 1990; Frolov et al. 1991; Frolov & Pankov 1992; Nagata & Whyte 1992; Fernandez-Reiriz et al. 1993) and it is composed of 61–80% glucose (which is present mainly as glycogen), 9–18% ribose and 0.8–7.0% of galactose, mannose, deoxyglucose, fucose and xylose (Nagata & Whyte 1992). Food ration affects the reproductive rate of rotifers and their protein, lipid and carbohydrate content. The protein content of individual rotifers increases by 60–80% with increasing


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food ration, but the amino acid profiles of rotifers are unaffected by either food ration or type of food provided to rotifers (Lubzens et al. 1989; Frolov et al. 1991; Tamaru et al. 1991, 1993; Frolov & Pankov 1992; Nagata & Whyte 1992; Fernandez-Reiriz et al. 1993; Øie & Olsen 1997). 2.5.3.2 Lipid composition The lipid content of rotifers varies generally between 9 and 28% of their dry weight and has been found, unequivocally, to have the greatest influence on growth and survival of marine fish larvae. About 34–43% of the lipids in rotifers are phospholipids and 20–55% are triacylglycerols, with small amounts of monoacylglycerols, diacylglycerols, sterols, sterol esters and free fatty acids (Teshima et al. 1987; Frolov et al. 1991; Nagata & Whyte 1992; Fernandez-Reiriz et al. 1993; Rainuzzo et al. 1997). Rotifer phospholipids and triacyglycerols display similar fatty acid profiles, but these are greatly affected by the lipids provided in their diet. Eicosapentaenoic (EPA) and docosahexaenoic (DHA) acids (20:5n-3 and 22:6n-3, respectively) have been known for several decades to be essential fatty acids for the survival of marine fish larvae (Owen et al. 1975; Fujita 1979; Watanabe et al. 1983). In general, marine fish contain large amounts of EPA and DHA in the phospholipids of their cellular membranes and, since they cannot synthesise them from linolenic acid (18:3n-3), these acids are essential dietary constituents. More specifically, DHA is present in high concentrations in neural and visual membranes, and insufficiency in the larval diet may result in serious consequences for a wide range of physiological and behavioural processes. These include impaired pigmentation and vision at low light intensities, leading to low hunting capabilities of the developing larvae and impaired development of the neuroendocrine system (Bell et al. 1995; Sargent et al. 1997, 1999; Estevez et al. 1999). Similarly, fish have a limited capacity to convert linoleic acid (18:2n-6) to n-6 HUFA, including arachidonic acid (ARA, 20:4n-6), which has gained increased attention in recent years (Sargent et al. 1997, 1999) and is now also considered an essential fatty acid. It has been shown to improve stress tolerance in fish larvae (Koven et al. 2001). ARA is the main precursor fatty acid of eicosanoids that are converted to biologically active compounds including prostaglandins, thromboxanes and leukotrienes, or can function in response to hormonal stimulation. Accumulating evidence points to the importance of supplying an optimal blend of EPA, DHA and ARA in the diet of marine fish larvae. Moreover, phospholipids rather than triacylglycerols are the preferred vehicle for delivery of these polyunsaturated fatty acids (PUFA). This probably relates to the limited ability of fish larvae to synthesise phospholipids de novo. It has been suggested that the optimal ratio for DHA:EPA:ARA is 1.8:1:0.12 for turbot (Sargent et al. 1999) and that excess ARA may have a deleterious affect (Bessonart et al. 1999; Estevez et al. 1999). While it is well known that rotifers can be easily cultured on yeast, rotifers reared in this way are nutritionally inadequate for marine fish larvae as they lack adequate amounts of DHA, EPA or ARA. Rotifers have to be enriched with these fatty acids and enrichment methods include feeding rotifers with algae, lipid emulsions, microparticulates or microcapsules containing lipids, or lipids with protein and carbohydrates (reviewed in Lubzens et al. 1989; Rainuzzo et al. 1997; Sargent et al. 1997). The quantitative and qualitative lipid content of rotifers can be manipulated by short or long enrichment periods on various diets containing lipid emulsion. Rotifer phospholipids were less influenced by these

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diets than the triacylglycerol fraction (Rainuzzo et al. 1997). Commercial fish oils that consist mainly of triacylglycerols are the main source of DHA or EPA. Their content depends on the fish species, but they usually are poor in ARA, except for tuna orbital oil that may contain about 2% of ARA. These oils are usually incorporated into most artificial diets for rotifers. Lipids can also be provided by feeding algae to rotifers, e.g. EPA-rich Nannochloropsis and DHA-rich Isochrysis, both of which contain ARA. A mutant strain of Nannochloropsis that was found to be deficient in EPA (Schneider et al. 1995) contains abundant ARA (23.3 mol% in whole cell extracts of the mutant strain versus 4.2 mol% in the wild type) and can be used as an alternative source of this fatty acid. The algae can be supplied directly from cultures, as a live or frozen concentrated paste or after freeze-drying (Watanabe et al. 1983; discussed in Lubzens et al. 1989 and in Fulks & Main 1991; Frolov et al. 1991; Lubzens et al. 1995a; Takeyama et al. 1996; Navarro & Yufera 1998). More recently, bacteria isolated from Antarctica, containing high levels of EPA, were suggested as a potential alternative enrichment food for rotifers (Nichols et al. 1996). Another lipid enrichment source is the freshwater Chlorella (Hayashi et al. 2001). The lipid content of rotifers is usually lower than that of their food organism, indicating that lipids are utilised by the rotifers. Rotifers utilise more DHA in highly reproducing cultures (Øie & Olsen 1997) and lipid utilisation is temperature dependent (Olsen et al. 1993). They accumulate about three to five times more total lipids when they are kept at 10°C than at 25°C (Lubzens et al. 1995b) and these results suggest that higher enrichment levels will be obtained if this procedure is performed at relatively low temperatures (depending on the rotifer strain), where reproductive rates and utilisation rates are slowed down. 2.5.3.3 Vitamin enrichments The importance of enriching rotifers with vitamins has not been studied extensively. In addition to vitamin B12, which was mentioned previously, fat-soluble vitamins (A, D and E) were found to promote rotifer reproduction (Hirayama 1990). The content of water-soluble vitamins in rotifers increased after changing their diet from baker’s yeast and lipid emulsion to Isochrysis. The most significant increase was in the content of ascorbic acid and thiamin. Rotifers fed on algae contain sufficient amounts of these vitamins to meet the nutritional requirements of fish larvae (Lie et al. 1997). However, enhanced levels of lipid-soluble vitamins in rotifers, caused by feeding them fish oil emulsions, were quickly depleted when the rotifers were switched to a diet of Isochrysis. Nevertheless, their final content in rotifers exceeded the recommended levels needed for proper growth of fish larvae (Lie et al. 1997). Vitamin C (ascorbic acid) not only stimulates rotifer growth (Satuito & Hirayama 1991) but also contributes significantly to the survival of fish larvae (Dabrowski & Ciereszko 1993; Dabrowski & Blom 1994). Its content in rotifers was found to depend on the diet, as it is highly abundant in several species of algae. Maximal levels were achieved in rotifers enriched with ascorbyl palmitate (Merchie et al. 1997).

2.5.4 Effects of starvation One of the main problems in providing rotifers to larvae is the deterioration in their nutritional quality due to starvation that results from extended periods of residence in the fish


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tanks. About 40–50% of the rotifer body mass is lost during 4–5 days of starvation at 18–20°C and the rate of decrease is positively related to temperature (Markridis & Olsen 1999). During starvation, preferential degradation of lipids, carbohydrates and amino acid takes place, leading to an increase in the proportion of proteins in the dry weight. The chemical composition of rotifers (e.g. protein content) can be presented in either mg g⫺1 dry weight or ng rotifer⫺1, and while large differences may occur in the content per rotifer (especially during starvation), these may not be reflected in the ratio of mg g⫺1 dry weight. During the first 8 h of starvation, free amino acids are used as the main energy source by rotifers, with lipids and carbohydrates being utilised later. Lipids serve as the main source of energy and different lipid classes are mobilised at different rates during starvation. An increase in the proportion of diacylglycerols, monoacylglycerols, sterols and free fatty acids results from mobilisation of triacylglycerols, and this also leads to an increase in the proportion of polar lipids. The mobilisation of sterols and wax esters follows the hydrolysis of triacylglycerols and carbohydrates, and coincides with increased mortality of rotifers (Frolov & Pankov 1992). The loss of lipids depends on temperature and can reach 19% of total lipids per day at 18°C, and the content of n-3 fatty acids is reduced more rapidly than other lipids (Olsen et al. 1993). The protein content of each rotifer is reduced during starvation, but the amino acid composition is rather stable (Frolov & Pankov 1992). The effect of starvation can be partially overcome by supplying algae to the fish tanks (Markridis & Olsen 1999). These results suggest that newly fed and enriched rotifers should be supplied daily to cultured larvae.

2.6 Preserved Rotifers Meeting the demands of fish larvae is a continuous effort from the day of first feeding up to the time that larvae are fed on other food sources (e.g. Artemia). While the usual practice is to depend on daily harvesting of rotifers from live cultures, various methods of storing rotifers have been explored.

2.6.1 Preservation at low temperatures Frozen rotifers are not usually adequate as food because of leaching of nutrients after thawing, their lack of buoyancy and motility, and they may also cause a deterioration in the water quality if introduced into the culture tanks. Live B. plicatilis can be stored at 4°C at relatively high densities, for at least 1 month (Lubzens et al. 1990), but this period is shorter for the thermophilic B. rotundiformis strains (Assavaaree et al. 2001a,b; see below). Storage at low temperature can help to reduce the daily tasks of harvesting and enriching rotifers, which may be performed every few days. It can also be helpful in storing surplus rotifers produced on one day for later use, providing a safeguard against unpredicted culture crashes, and facilitating transport of rotifers between sites of production and culture facilities. While storage at 4°C is feasible in most places, rotifers continue to reproduce at this temperature and require periodic feeding and exchange of culture media. Rotifers can be kept at ⫺1°C without feeding or water exchange for about 2 weeks (Lubzens et al. 1995b), but storage at this temperature requires more specific equipment.

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As storage at low temperature was found to be relatively easy and successful for B. plicatilis rotifers, extending this method to B. rotundiformis has been examined recently (Assavaaree et al. 2001a,b). The results show that B. rotundiformis rotifer strains are less tolerant to 4°C than B. plicatilis rotifer strains, and the strains known as SS type were the most susceptible and showed the lowest survival. While the mean survival of B. plicatilis strains ranged from 26.9 to 63.7% after 30 days at 4°C, only one B. rotundiformis strain survived after 15 days at this temperature and only four strains survived after 7 days. A clear variation was found between S and SS type strains, with a lower survival of SS-type strains at low temperatures. At 12°C, more than 80% of B. rotundiformis rotifers (of an S-type strain and an SS-type strain) survived after 10 days. Survival of an S-type strain was higher at 35 ppt, while the SS-type strain survived best at 17 ppt. Feeding rotifers at intervals of 2 days improved their survival over those fed only at the beginning of the experiment or at intervals of 4 days, and changing the culture media every 4 days suppressed survival. An acclimation period of 24 h at 20°C, before transferring of rotifers from their usual culture temperature (28°C) to 12°C, resulted in higher survival of SS-type rotifers but had no effect on S-type rotifers. These results indicate the difficulties in attempting to keep small-type rotifers at low temperatures for extended periods, and the optimal conditions have to be empirically determined for each strain.

2.6.2 Cryopreservation Long-term preservation of genetically important strains can be achieved by cryopreservation. Amictic eggs (but not adults) are preserved in liquid nitrogen after they have been impregnated with cryoprotective agents such as dimethyl sulfoxide (DMSO) or propanediol (Toledo & Kurokura 1990; Toledo et al. 1991; Hadani et al. 1992). This method ensures full preservation of genetic traits of importance to aquaculture and is especially important for those strains that do not produce resting eggs. A small collection of cryopreserved B. plicatilis and B. rotundiformis strains is kept at the authors’ laboratory and serves as an alternative source for live cultures. Since this is a relatively expensive method, it is not suitable for preservation of large numbers of rotifers for direct use as food after thawing.

2.6.3 Resting eggs Artificially produced rotifer resting eggs have been offered as an alternative route for supplying rotifers without depending on the daily production cycle used in marine hatcheries. Resting egg production is genetically determined (Hino & Hirano 1976, 1977), with large variations between rotifers originating from eggs produced by one clone (Lubzens 1989). The production of these eggs can be manipulated by environmental factors, such as salinity, food quantity and quality, rotifer culture density, exchange of culture media and temperature, and varies between B. plicatilis and B. rotundiformis species. (Hino & Hirano 1976, 1977, 1984, 1985, 1988; Hagiwara et al. 1988a,b, 1989; Snell & Hoff 1985; Lubzens et al. 1985, 1993; Snell 1986; Lubzens 1987; Lubzens & Minkoff 1988; Hagiwara & Lee 1991; Hagiwara & Hirayama 1993; Hamada et al. 1993; Kogane et al. 1997).


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Basically, it is relatively easy to obtain rotifer resting eggs, but the main problem is the relative number produced in relation to the invested effort. The first step is choosing an appropriate strain, as their production is genetically determined. For B. plicatilis, cultures should be maintained at salinities below 30 ppt and temperatures should preferably range from 12 to 25°C. Rotifer density should be kept low, not exceeding 150 rotifers ml⫺1, and food (preferably Nannochloropsis sp.) should be provided in adequate quantities. After introducing a rotifer inoculum of 5–10 individuals ml⫺1, the rotifers will start reproducing and, depending on the culture temperature, mictic females carrying male eggs will appear after 2–3 days. This will be followed by the appearance of males and of fertilised females carrying resting eggs, after 4–6 days. These resting eggs will be released from the females at the end of their development and will sink to the bottom of the culture container. With the appearance of resting eggs, the number of females carrying male eggs will start to decline and this will be followed by a decline in the relative abundance of males and reduced production of resting eggs. Most often, cultures at this stage collapse and production of resting eggs is stopped. The production cycle lasts for 10–24 days. The resting eggs can be collected from the debris found at the bottom of the culture container, by sieving the culture water with agitation, through 200 ␮m mesh plankton net, through which the eggs will pass. The material collected on the sieve should be resuspended in clean seawater and the sieving process repeated several times, until most of the eggs have been freed from the debris. Removal of the remaining debris can be achieved by suspending the eggs in clean, diluted seawater (e.g. 10 ppt) in a separating funnel and introducing an aeration tube just above the bottom outlet. A very fine airstream will result in flotation of the debris and sinking of the eggs to the funnel outlet, where they can be collected. The collected eggs should be stored in the dark (in aluminium foil-covered glass tubes or any other container) at a low temperature. The simplest method is to suspend the eggs in capped tubes containing clean or sterile 10 ppt seawater and store them in a refrigerator (Minkoff et al. 1983). More recent results showed that eggs may also be stored in a dried form: desiccated, lyophilised or canned under pressure (Balompapueng et al. 1997b). Hatching of resting eggs is achieved by transferring them from the stored container into fresh seawater and incubation at 10–25°C, with illumination (Minkoff et al. 1983; Hagiwara & Hino 1990). Kogane et al. (1997) showed that a low-temperature treatment could improve the efficiency of resting egg production by culturing B. plicatilis for 20 days at 12°C before transferring them to 25°C. The optimal conditions for production of B. rotundiformis differ from those of B. plicatilis, with resting egg production encouraged at higher salinities in B. rotundiformis (Hagiwara et al. 1989). Mass production of rotifer (B. plicatilis) resting eggs has been reported in 50 m3 tanks (Hagiwara et al. 1993a) and improvements in culture techniques have been tested, including the use of a semi-continuous culture method to maintain relatively low rotifer density and the use of a nylon filter system for removing excess debris (Hagiwara et al. 1993a, 1997; Hagiwara & Hirayama 1993; Hagiwara 1994; Balompapueng et al. 1997a). Methods have been devised for optimal storage of large quantities of resting eggs, including removal of attached bacteria by rinsing eggs with hypochlorite (1.0 mg l⫺1) or sodium nifurstyrenate (5.0 ␮g ml⫺1) for improving hatchability (Balompapueng et al. 1997a,b). Hatching conditions are well established, facilitating the use of resting eggs at the required time (Minkoff et al. 1983; Pourriot & Snell 1983; Hagiwara et al. 1985, 1995c; Hagiwara &

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Hino 1989, 1990; Hagiwara 1996; Balompapueng et al. 1997a). These eggs can be used directly for feeding fish larvae (Hagiwara & Hirayama 1993; Hagiwara et al. 1993b) or for initiating mass cultures, but asexual reproductive rates in rotifers hatched from resting eggs may show large fluctuations from those of the parent culture (Lubzens 1989). The cost of producing resting eggs exceeds several-fold that of producing mass-cultured rotifers (Lubzens 1989; Lubzens et al. 2001) and therefore has not yet been extensively adapted as a direct source of food for fish larvae. Moreover, rotifers hatched from artificially produced resting eggs may show a high occurrence of mixis, since the strain chosen for production of resting eggs has been selected for this purpose. Therefore, caution is required in using commercially available resting eggs for initiating mass cultures, as these cultures may show a high incidence of males under specific culture conditions (e.g. low salinity for B. plicatilis).

2.7 Future Directions The prospects of replacing live rotifers as food for early developmental stages of fish larvae are far from feasible, despite considerable efforts in this direction (Tandler 1984, 1985; Lubzens 1987; Kolkovsky & Tandler 1995). Generally speaking, current methodologies of producing and enriching rotifers have succeeded in meeting the demands of the industry. However, the current pressing need for very small live food is difficult to meet, although several ‘super small’ genetic strains have been found and cultured (Hagiwara et al. 2001). Improved methods for predicting the physiological state of rotifers in mass cultures could be helpful in avoiding the collapse of cultures. Using preserved rotifers may alleviate the immediate dependence of hatcheries on the daily production of rotifers. This includes keeping live rotifers at low temperatures or, alternatively, as resting eggs. Cheaper methods for resting egg production of inbred lines will be of great advantage in reaching this goal.

Acknowledgements The financial support for rotifer species and strain cultures (1991–2000) by the Kunin Lunenfeld Foundation is greatly acknowledged. The advice, support and patience of the editor, Dr J. Støttrup, made this review possible.

2.8 References Aoki, S. & Hino, A. (1996) Nitrogen flow in a chemostat culture of the rotifer Brachionus plicatilis. Fish. Sci., 62, 8–14. Assavarree, M., Hagiwara, A., Ide, K., Maruyama, M. & Lubzens, E. (2001a) Low-temperature preservation (at 4°C) of marine rotifer Brachionus. Aquacult. Res., 32, 29–39. Assavaaree, M., Hagiwara, A. & Lubzens, E. (2001b) Factors affecting low temperature preservation of the marine rotifer Brachionus rotundiformis Tschugunoff. Hydrobiologia, 446/447, 355–361. Awaïss, A. (1991) Mass culture and nutritional quality of the freshwater rotifer (Brachionus calyciflorus P.) for grudgeon (Gobio gobio L.) and perch (Perca fluviatus L.) larvae. In: Larvi’91 – Fish


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Chapter 3

Biology, Tank Production and Nutritional Value of Artemia Jean Dhont and Gilbert Van Stappen

3.1 Introduction Artemia has probably been known and used within its natural distribution areas for centuries. However, its fame elsewhere only began to rise in the 1930s when some investigators adopted it as a convenient replacement for the natural diet of fish larvae, thus realising the first breakthrough in the culture of commercially important fish species (Sorgeloos 1980). In the 1950s, Artemia cysts were still predominantly marketed for the aquarium and pet trade at costs as low as US $10 kg⫺1. There were only two commercial sources: the coastal saltworks in the San Francisco Bay (SFB, California, USA) and the Great Salt Lake (GSL, Utah, USA). With fish and shrimp aquaculture developing from the early 1960s, new marketing opportunities were created for Artemia cysts. However, by the mid-1970s, increased demand, declining harvests from the GSL, high import taxes in some developing countries and, possibly, an artificial cyst shortage created by certain companies resulted in a severe price rise of up to US $50 or 100 kg⫺1 Artemia cysts by the end of the 1980s (Bengtson et al. 1991). The dramatic impact of the cyst shortage on the expanding aquaculture industry encouraged research on rationalising the use of Artemia and exploration of new cyst resources. During that period, the commercial exploitation of several other natural sources (Argentina, Australia, Canada, Colombia, France, PR China) and managed Artemia production sites (Brazil, Thailand) occurred. On the initiative of the Artemia Reference Center (Ghent University, Belgium) the International Study on Artemia (ISA) was established to co-ordinate a variety of different research initiatives (Sorgeloos 1979). The cyst shortage also simultaneously encouraged the search for alternatives for Artemia such as microencapsulated diets (Jones et al. 1993; Samocha et al. 1999) with the aim of eliminating live feed in fish larval nutrition, a process that continues today with slow but steady progress. With the development of improved techniques for cyst and nauplii applications (Léger et al. 1987a) and the exploitation of new natural resources, cyst prices returned to normal and annual market supply reached over 50 t by the 1980s. During the 1980s, improved techniques for harvesting from the open water and favourable hydrological and climatic conditions enabled a 10-fold increase in the yields from GSL (⬎200 t processed product; Fig. 3.1), while the hatching quality was also improved. This led to the precarious situation where, by the end of the 1980s, cyst supply was more than 90% dependent on one resource, namely

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10,000

Harvested raw product (t)

9,000 8,000 7,000 6,000 5,000 4,000 3,000 2,000 1,000 0 ’84

’86

’88

’90

’92 Year

’94

’96

’98

2000

Fig. 3.1 Cyst harvest (in tonnes of raw product) from the Great Salt Lake (Utah, USA). (Data from USGS 2001.)

GSL. Despite its size, the GSL remains a natural ecosystem subject to climatic and other influences, and this has been illustrated by unpredictable and fluctuating cyst harvests. This situation urged producers in the 1990s to explore new sites such as Lake Urmia in Iran, Aibi Lake in China, Bolshoye Yarovoye in Siberia, Kara Bogaz Gol in Turkmenistan, and several lakes in Kazakhstan (Lavens & Sorgeloos 2000). In addition, numerous managed ponds and saltworks world-wide provided small quantities (1–20 t each). Although these sites did not necessarily contribute substantially to the world supply of cysts, they provide interesting opportunities for local commercial development. New insights into hatching characteristics and nutritional essentials gave rise to the segregation of different cyst qualities. Whereas in the early 1990s, cysts with good hatching quality could be purchased for as low as US $20 kg⫺1, prices of small sized cysts with high eicosapentaenoic acid (EPA; 20:5n-3) levels could reach over US $100 kg⫺1 at times of short supply owing to their critical role as starter food for marine fish larvae. With the severe cyst shortage in the mid-1990s and at the end of the twentieth century (Fig. 3.1), cyst prices inflated to levels around US $100 kg⫺1 for GSL product and nearly US $200 kg⫺1 for the EPA-rich product. Following the superharvest of over 9000 t of raw cysts from the GSL in 2000–2001, prices started to fall again. Since the early 1990s, cyst consumption has increased exponentially as a consequence of the booming shrimp and marine fish industries. In 1997, some 6000 hatcheries required over 1500 t of cysts annually. Some 80–85% of the total sales of Artemia went to shrimp hatcheries, mainly in China and south-east Asia, as well as Ecuador and a few other Latin American countries; the remainder went to marine fish larviculture in Europe, China and Japan, as well as to the pet fish producers. However, the rationalisation of the use of Artemia in hatcheries (Sorgeloos et al. 1998, 2001) has enabled a dramatic reduction in the required amount of cysts per unit of fish or shrimp produced. For instance, formerly, a typical Mediterranean sea bass and sea bream hatchery would have been using some 150 kg cysts to produce 1 million fry, whereas nowadays the required amount of cysts is only 90 kg for bass and 70 kg for bream. Likewise, in shrimp hatcheries, the consumption of cysts dropped from 10 kg per million

Biology, Tank Production and Nutritional Value of Artemia

67

postlarvae to less than 5 kg. Although there is no doubt that Artemia will gradually be replaced by formulated diets, it is obvious that the use of nauplii will continue to be market driven for at least a few more years and that record harvests at GSL and new locations may relieve the pressure or even reverse the current trends. Comprehensive literature reviews on the use of Artemia as live food in fish and shellfish larviculture have been published by Léger et al. (1986) and Sorgeloos et al. (1998, 2001). Today, Artemia is used in the mass culture of different sea bream species, sea bass species, wolf fish, cod, turbot, halibut, flounder species and other flatfish, milkfish, sturgeon, different carp and catfish species and whitefish species. The same is true for commercially important crustaceans such as several shrimp and prawn species, crawfish, several edible crab species and lobster. Nauplii in instar I and II stages are, undoubtedly, the most widely used forms of Artemia in aquaculture. They are also the easiest and earliest live food, being obtained directly from the cysts. However, it should be borne in mind that any farmer would switch to formulated feed as soon as this proves to be more cost-effective than Artemia. This switch will be triggered not only by the constantly improving quality of formulated feed, but also by price and quality of Artemia. In general, most fish and shrimp larvae accept formulated feed more easily as they grow bigger. This is not only a matter of size of mouthparts and particle size, but also a matter of the developmental stage and efficiency of the digestive system. As a consequence, Artemia is essential only for those species that require live food in their early life stages. Brine shrimp are mostly used as freshly hatched nauplii or as ‘enriched’ nauplii (see Section 3.5.5). Hatching procedures can be simplified and improved through prior ‘decapsulation’ of cysts (see Section 3.5.4), a process that also improves the quality of poor or nonhatching cysts. Juvenile and adult Artemia, often referred to as ‘biomass’, can be obtained through culturing (see Section 3.3.1) or can be harvested from salt ponds or lakes (Baert et al. 1996). In China, thousands of tonnes are collected on an annual basis from the Bohai Bay salt ponds and are used in the local culture of Chinese white shrimp, Penaeus chinensis (Tackaert & Sorgeloos 1991). Although live biomass has a higher nutritive value, most of the 3000 t that is harvested annually is marketed in frozen form. Part of it is also flaked, dried or incorporated in compound diets.

3.2 Biology of Artemia 3.2.1 Morphology and life cycle In its natural environment under certain conditions, Artemia produces cysts that float at the water surface and are driven ashore by wind and waves. These cysts are metabolically inactive and do not develop further as long as they are kept dry. Upon immersion in seawater, the biconcave cysts hydrate and become spherical and, within the shell, the embryo resumes its interrupted metabolism. After about 20 h the outer membrane of the cyst bursts (‘breaking’) and the embryo appears, surrounded by the hatching membrane (Fig. 3.2). While the embryo hangs underneath the empty shell (the ‘umbrella’ stage) the development of the nauplius is completed and within a short period the hatching membrane is ruptured (‘hatching’), giving rise to the free-swimming nauplius (Fig. 3.3).

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Fig. 3.2 Cyst in breaking stage. (1) Nauplius eye.

Fig. 3.3 Embryo in ‘umbrella’ stage (left) and instar I nauplius (right). (1) Nauplius eye; (2) antennula; (3) antenna; (4) mandible.

The first larval stage (instar I; 400–500 ␮m in length) has a brownish-orange colour, a red nauplius eye in the head region and three pairs of appendages: the first antennae (sensorial function), the second antennae (locomotory plus filter-feeding function) and the mandibles (food uptake function). The ventral side is covered by a large labrum (food uptake: transfer of particles from the filtering setae into the mouth). The instar I larva does not take up food as its digestive system is not yet functional; it relies completely on its yolk reserves. After about 8 h, the animal moults into the second larval stage (instar II). Small food particles (e.g. algal cells, bacteria, detritus) ranging in size from 1 to 50 ␮m are filtered out by the second antennae and ingested into the now functional digestive tract. The larva grows and differentiates through a number of moults; although there has been considerable disagreement about the exact number of larval stages, generally one naupliar, four metanaupliar, seven postmetanaupliar and five postlarval stages have been described (Hentschel 1968; Schrehardt 1987). Paired lobular appendages appear in the trunk region and differentiate into thoracopods (Fig. 3.4). On both sides of the nauplius eye lateral complex eyes begin to develop


69

Fig. 3.4 Instar V larva. (1) Nauplius eye; (2) lateral complex eye; (3) antenna; (4) labrum; (5) budding of thoracopods; (6) digestive tract.

Fig. 3.5 Head and thoracic region of young male. (1) Antenna; (2) telopodite; (3) exopodite.

(Figs 3.4, 3.5). From the 10th instar stage onwards, important morphological and functional changes begin to take place, i.e. the antennae lose their locomotory function and undergo sexual differentiation. In males (Figs 3.6, 3.7) they develop into hooked graspers, while the female antennae degenerate into sensorial appendages (Fig. 3.8). The thoracopods are now differentiated into three functional parts (Fig. 3.9): the telopodites and endopodites (locomotory and filter-feeding), and the membranous exopodites (gills). Adult Artemia are typical primitive arthropods (8–12 mm in length) having an elongated segmented body with two stalked complex eyes, a linear digestive tract, sensorial antennulae, a pair of functional thoracopods on each of the 11 thoracal segments (Figs 3.7, 3.8) and a furca on the last abdominal segment. The entire body is covered with a thin, flexible exoskeleton of chitin to which muscles are attached internally. The male (Fig. 3.7) has a paired penis on the first of the eight abdominal segments (Fig. 3.10). Female Artemia can easily be recognised by the brood pouch or uterus situated in the same segment, just behind the 11th pair of thoracopods (Figs 3.8, 3.10). The female reproductive system consists of ovaries and oviducts leading into the single, median uterus, wherein several clusters of shell glands open. The ovaries are paired tubular structures extending into the abdomen

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Fig. 3.6 Head of an adult male. (1) Antenna; (2) antennula; (3) lateral complex eye; (4) mandible.

Fig. 3.7 Adult male.

Fig. 3.8 Adult female.


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Fig. 3.9 Detail of anterior thoracopods in adult Artemia. (1) Exopodite; (2) telopodite; (3) endopodite.

Fig. 3.10 Artemia couple in riding position. (1) Uterus; (2) penis.

Fig. 3.11 Uterus of ovoviviparous Artemia filled with nauplii (first larvae are being released). (1) Ovary with eggs.

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Fig. 3.12 Uterus of oviparous Artemia filled with cysts. (1) Brown shell glands.

(Fig. 3.11). Adult females ovulate approximately every 140 h, depending on rearing conditions and whether development of embryos occurs oviparously or ovoviviparously. In females, spawning is followed by a moult, after which ovulation takes place. The oviducts emerge from the ovaries near the anterior part of the third abdominal segment (Cassel 1937). Each oviduct empties into the anterolateral border of the uterus. The lateral pouches function as seminal receptacles during the time between copulation and fertilisation (within 1 h) (Benesch 1969; Criel 1980a,b). Once ripe, the eggs developing in the ovaries become spherical and migrate via two oviducts into the unpaired uterus. Fertilised eggs normally develop into free-swimming nauplii (ovoviviparous reproduction) (Fig. 3.11), which are released by the mother. In extreme conditions (e.g. high salinity, low oxygen levels) the embryos only develop up to the gastrula stage. At this point they are surrounded by a thick shell (secreted by the brown shell glands located in the uterus), enter a state of metabolic dormancy (diapause) and are then released by the female (oviparous reproduction) (Fig. 3.12). The shell glands consist of several cell clusters, and can vary from dark brown to white or even colourless, depending on reproductive strategy. In principle, both oviparity and ovoviviparity are found in all Artemia strains, and females can switch reproductive modes from one ovulation to the next. Although females may differ in their genetic tendency to reproduce either ovoviviparously or oviparously, no Artemia are known to lack completely the ability to produce ovoviviparous nauplii. The cysts usually float in the high-salinity waters and are blown ashore where they accumulate and dry. As a result of this dehydration process the diapause mechanism is generally inactivated; cysts are now in a state of quiescence and can resume their further embryonic development when hydrated in optimal hatching conditions. Under optimal conditions brine shrimp can live for several months, grow from nauplius to adult in only 8 days and reproduce at a rate of up to 300 nauplii or cysts every 4 days. However, not all encysted embryos produced by oviparous animals enter diapause, and nauplii emerge from some cysts without dehydration or other treatment (Jensen 1918; Mathias 1937; Lochhead & Lochhead 1940; Dutrieu 1960a,b; Morris & Afzelius 1967;


73

alveolar layer

outer cuticular membrane embryonic cuticle

inner cuticular membrane embryo

Fig. 3.13 Schematic diagram of the ultrastructure of an Artemia cyst. (Modified from Morris & Afzelius 1967.)

Benesch 1969; Anderson et al. 1970). These cysts are surrounded by a much thinner shell than those that enter diapause (Lochhead & Lochhead 1940). Moreover, changes in cysts post release suggest that diapause is established gradually after release (Jardel 1986). The cryptobiotic cyst shell has two important layers in addition to the hypochlorite-soluble, double-layered outer chorion secreted by the shell glands, and the hypochlorite-resistant embryonic cuticle (Morris & Afzelius 1967) (see also Section 3.2.5.1). These are the outer cuticular membrane, separating chorion from embryonic cuticle, and the inner cuticular membrane, which delineates the embryo from the fibrous layer of the embryonic cuticle (Fig. 3.13).

3.2.2 Ecology and natural distribution Artemia populations are found in about 500 natural salt lakes and artificial salterns scattered throughout the tropical, subtropical and temperate climatic zones, along coastlines as well as inland. The distribution of these sites over the continents is very uneven, mainly reflecting sampling and exploration activities (Fig. 3.14). As such, it does not give a precise picture of the actual global occurrence of Artemia. The decline of Artemia cyst harvests from the GSL in Utah, USA, since 1977 (Lavens & Sorgeloos 2000) has intensified the search for alternative resources, especially in inland lakes that are sufficiently large and productive to justify commercial exploitation. As a result, several sites, especially in continental Asia, are exploited occasionally or on a regular basis (with some local investment), and these cysts are being used world-wide in aquaculture. The identity or location of these sites has still not reached scientific literature, and attempts are seldom made to perform a systematic

74


Fig. 3.14 World distribution of Artemia.

characterisation of the respective strains. A continued survey will undoubtedly lead to the discovery of many more Artemia biotopes in different parts of the world. Two critical factors determine the population dynamics of Artemia and its biogeographical distribution: first, whether water body conditions allow the animals to survive throughout the year and, secondly, whether or not the seasonality of the environment is predictable (Lenz 1987; Amat et al. 1995). The common feature of all Artemia biotopes is their high salinity. Salinity is without doubt the predominant abiotic factor determining the presence of Artemia and consequently limiting its geographical distribution. Its physiological adaptations to high salinity provide a very efficient ecological defence against predation, as brine shrimp possess:

• • •

a very efficient osmoregulatory system the capacity to synthesise very efficient respiratory pigments to cope with the low oxygen levels at high salinities the ability to produce dormant cysts when environmental conditions endanger the survival of the species.

Other variables (temperature, light intensity, primary food production) may have an influence on the quantitative aspects of the Artemia population, or may cause only a temporary absence of brine shrimp. For physiological reasons the salinity optimum is situated towards the lower end of the salinity range, as higher ambient salinity requires higher energy costs for osmoregulation. Ambient salinity also plays a role in cyst metabolism, as Artemia cysts will only start to develop when the salinity of the medium drops below a certain threshold value. No Artemia are found in cold tundra or frost climates, as the year-round extremely low temperatures preclude Artemia development. Most strains do not seem to survive prolonged temperatures below 5°C unless in the form of cysts. The maximum temperature tolerated by


75

Artemia populations has repeatedly been reported to be close to 35°C, a temperature often attained in the shallow tropical salterns that constitute a large part of the Artemia habitats. This tolerance threshold is, however, strain dependent. Moreover, physiological adaptation of SFB Artemia to high temperatures (40°C) after a number of generations in Vietnamese salt ponds has also been reported (Clegg et al. 2001). As for salinity, temperature optima are difficult to define and are strain dependent; in general, however, the optimum for Artemia is in the range 25–30°C. The ametabolic dehydrated cysts are resistant to a wider temperature range than would ever occur in nature. Artemia is a non-selective filter feeder of organic detritus, microscopic algae and bacteria. The Artemia biotopes typically show a very simple trophic structure and low species diversity: the absence of predators and food competitors allows brine shrimp to develop into monocultures. As Artemia is incapable of active dispersion, wind and waterfowl (especially flamingos) are the most important natural dispersion vectors. The floating cysts adhere to feet and feathers of birds and, when ingested, they remain intact for at least 2 days in the digestive tract of birds. Consequently, the absence of migrating birds is probably the reason why certain areas that are suitable for Artemia (e.g. salinas along the north-east coast of Brazil) are not naturally inhabited by brine shrimp.

3.2.3 Taxonomy The brine shrimp Artemia comprises a group of zygogenetic and parthenogenetic, morphologically similar species very likely to have diverged from an ancestral form living in the Mediterranean area some 5.5 million years ago (Abreu-Grobois & Beardmore 1982; Abreu-Grobois 1987; Badaracco et al. 1987). Speciation in the genus should be regarded as a complex, multidimensional process involving a variety of environmental and genomic factors. The identification of zygogenetic Artemia species has been established by a multidisciplinary approach, including cross-breeding tests, morphological differentiation, cytogenetics, allozyme studies, and nuclear and mitochondrial DNA sequencing. With the exception of cross-mating, all of these techniques have also contributed to identifying the parthenogenetic types described as A. parthenogenetica by Barigozzi (1974), as well as to gaining insight on population structure, origin and amount of clonal diversity. In 1755 Schlosser described the brine shrimp based on material collected from the solar saltworks near Lymington, England (no longer in existence) (Kuenen & Baas-Becking 1938). Linnaeus in 1758 classified it as Cancer salinus, and Leach in 1818 renamed the brine shrimp as Artemia salina (Artom 1931). Very often authors have named all brine shrimps A. salina. While for some time the name A. tunisiana was used, Artemia salina is now only recognised as a valid name for the zygogenetic species found in the Mediterranean area (Mura 1990; Triantaphyllidis et al. 1997b). The differentiation of seven zygogenetic species, defined primarily by the criterion of laboratory reproductive isolation, and many parthenogenetic strains is currently acknowledged. Endemic to the Old World are the parthenogenetic types designated by Barigozzi (1974) as A. parthenogenetica (with different levels of ploidy, found in Europe, Africa, Asia and Australia), the zygogenetic A. salina, Leach 1819 (Mediterranean area) (Triantaphyllidis

76


et al. 1997b), A. urmiana (Günther, 1890) (Iran), A. sinica (Cai 1989) (continental China), Artemia sp. (Pilla & Beardmore 1994) (Kazakhstan) and A. tibetiana (Abatzopoulos et al. 1998) (Tibet). Endemic to the New World are A. persimilis (Piccinelli & Prosdocimi 1968) (Argentina) and A. franciscana (Kellogg 1906) (North, Central and South America), with A. (franciscana) monica being a special case of a population described for an ecologically unique habitat (Mono Lake, USA). The genus Artemia is thus a complex of sibling species and superspecies, defined largely, but not completely, by the criterion of reproductive isolation. Very rarely, it has been shown that genetically extremely distinct and allopatric species can produce laboratory hybrids (Pilla & Beardmore 1994). Coexistence of two species in the same saline habitat is possible: mixtures of parthenogenetic and zygogenetic populations have been reported in Spain, Italy, and central and northern China. Parthenogenetic types tend to predominate in more disturbed, stressful, conditions of salinity, temperature and food availability (Browne & Bowen 1991; Lenz & Browne 1991). Laboratory competition experiments where Artemia adults, belonging to different species, were raised together and reproduced for a maximum of 3 months (25°C, ⫾80 g l⫺1, fed Dunaliella) resulted in the dominance of A. franciscana over parthenogenetic populations on the one hand, and parthenogenetic populations over A. salina on the other (Browne 1980; Browne & Halanych 1989).

3.2.4 Strain-specific characteristics The world-wide distribution of the brine shrimp Artemia in isolated habitats (about 500 natural salt lakes and artificial salterns) with specific ecological conditions has resulted in numerous geographical strains, or genetically different populations within the same sibling species. The parthenogenetic Artemia with their great clonal diversity, as evident from morphology (Hontoria & Amat 1992; Triantaphyllidis et al. 1997a), and cytological and allozyme studies (Abreu-Grobois & Beardmore 1982; Abreu-Grobois 1987; Abatzopoulos et al. 1993) and different ploidy levels (diploid, triploid, tetraploid, pentaploid), display a wide genotypic variation. While the nutritional value can be manipulated through enrichment, other qualities favourable for aquaculture can be obtained by selection of strains and/or their crossbreeds. In spite of the fluctuations in the harvest, over 90% of all marketed cysts originate from the GSL, but Artemia cysts are commercially available from various production sources in America, Asia, Australia and Europe. Knowledge of the characteristics (both genotypic and phenotypic) of a particular batch of cysts can greatly increase the effectiveness of its use in a fish or shrimp hatchery. Among these strains a high degree of genetic variability as well as a unique diversity in various quantitative characteristics have been observed (Browne et al. 1991). Some of this variability is phenotypical, such as the nutritional composition of the cysts (Léger et al. 1986), and changes from batch to batch. Other characteristics such as cyst diameter and resistance to high temperature are considered strain-specific and remain relatively constant (Vanhaecke & Sorgeloos 1980a), i.e. they have become genotypical as a result of long-term adaptations of the strain to the local conditions.


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Table 3.1 Size, individual dry weight and energy content of Artemia instar I nauplii from different cyst sources hatched in standard conditions (35 g l⫺1, 25°C).

Cyst source

Length (mm)

Dry weight (␮g)

Energy content (10⫺3 J)

San Francisco Bay, USA Macau, Brazil Great Salt Lake, USA Shark Bay, Australia Chaplin Lake, Canada Tanggu, Bohai Bay, PR China Aibi Lake, PR China Yuncheng, PR China Lake Urmiah, Iran

428 447 486 458 475 515 515 460 497

1.63 1.74 2.42 2.47 2.04 3.09 4.55 2.03 —

366 392 541 576 448 681 — — —

3.2.4.1 Size and energy content The nutritional effectiveness of a food organism is primarily determined by its ingestibility and, as a consequence, by its size and form. Data on biometrics of nauplii from various Artemia strains are given in Table 3.1. In spite of small variations between batches of the same strain, possibly caused by environmental and/or processing factors, generally the cyst diameter of different production batches of the same strain remains rather constant. Other biometrical characteristics, such as cyst volume, cyst dry weight, instar I naupliar length, individual naupliar weight and naupliar volume, and energy content, show a high correlation with the cyst diameter (Vanhaecke et al. 1983). As a consequence, biometrical parameters, in particular cyst diameter, are good tools to characterise Artemia strains, and to help define the origin of unknown or even mixed cyst samples (Vanhaecke & Sorgeloos 1980a). Some general correlations can also be made between sibling species and size: parthenogenetic Artemia produce large cysts; A. salina, large cysts with a thick chorion; A. franciscana and A. persimilis, small or intermediate cysts with a thin chorion. 3.2.4.2 Hatching quality Comparative studies of hatching behaviour of cysts of different origin show a considerable variation in hatching percentage, rate and efficiency (Vanhaecke & Sorgeloos 1982, 1983). However, none of these parameters is strain-specific as they are influenced by a wide array of factors such as harvesting, processing, storage and hatching techniques, as well as production conditions affecting the parental generation. For optimal use of Artemia in aquaculture the hatching characteristics of each batch of cysts being used should be known. 3.2.4.3 Diapause characteristics As diapause can be considered as a life-cycle strategy to overcome temporarily adverse conditions, and to synchronise population developments to the variations of their specific biotype, the process of diapause and its deactivation is likely to be adapted to the population’s habitat (Lavens & Sorgeloos 1987). Adaptations to local conditions may have contributed to strain-specific differences in diapause sensitivity. In response to simple dehydration by storage in a highly saline medium or by air-drying, SFB-type cysts are gradually released

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from diapause (Versichele & Sorgeloos 1980; Vu Do Quynh & Nguyen Ngoc Lam 1987), while cysts from inland salt lakes (GSL, southern Siberia) need a period of cold storage or hibernation of several weeks to break diapause. In addition, differences in tolerance and responsiveness of different strains to hydrogen peroxide (H2O2) treatment during diapause deactivation (see Section 3.2.5.3) may be partially genetic and thus strain-specific. Although general recommendations can be formulated with regard to H2O2 concentration and exposure time, a limited screening of different combinations is therefore needed when the hatching of new batches is being optimised (Van Stappen et al. 1998). 3.2.4.4 Growth rate of nauplii Standard culture tests with brine shrimp from different geographical origins show important differences in growth rate even within the same sibling species, but not among batches of the same strain (Vanhaecke & Sorgeloos 1980b). Although the population growth of Artemia in the field (e.g. after inoculation) is determined by a variety of factors, selection of a strain with a high potential growth rate will have a positive impact on maximal production output. 3.2.4.5 Temperature and salinity tolerance Both temperature and salinity significantly affect survival and growth, the effect of temperature being more pronounced. A broad range of temperatures and salinities meets the requirements for ⬎90% survival (Vanhaecke et al. 1984). Strains from thalassohaline biotopes share a common preferred temperature range around 20–25°C, where mortalities are ⬍10%. Interaction between temperature and salinity is limited; substantial differences in tolerance have been recorded at low salinities (around 5 g l⫺1) and high temperatures (30–34°C). At elevated temperatures the survival of the GSL strain is significantly higher than for other strains. 3.2.4.6 Life-history traits and reproductive capacity Life history and reproductive characteristics of Artemia strains are important factors when an introduction of brine shrimp to a new habitat is considered, especially when competition with a local strain is to be expected. These competitive abilities are related to factors such as the length of reproductive, prereproductive and postreproductive periods, total lifespan, number of offspring per brood, broods per female, interval between broods, etc. In general, New World (zygogenetic) populations have a very large number of offspring per brood, a large number of offspring per female per day and a fast development time to sexual maturity. These are all favourable characteristics compared with those of Old World zygogenetic and parthenogenetic Artemia (Browne et al. 1984, 1991). Age at first reproduction is a key factor determining the population growth rate, and the rate of colonisation of new environments with limited nutrient resources. Consequently, if environmental preferences and nutritional factors do not interfere, New World zygogenetic species generally outcompete parthenogenetic strains, the latter in their turn predominating over Old World zygogenetic species. Inoculation experiments in natural habitats therefore require prior screening of candidate strains and of indigenous local populations, as well as the study of prevailing environmental conditions.


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3.2.4.7 Nutritional value In the late 1970s, when many fish and shrimp hatcheries started to commercialise, switching from one source of Artemia to another provoked unexpected problems. Aquaculturists even noticed highly significant differences when using different batches from the same geographical origin (Léger & Sorgeloos 1984). In particular, the lipid and fatty acid compositions, as well as the metabolisation of fatty acids in the Artemia, seemed to differ widely from strain to strain, and even from batch to batch, as a consequence of the fluctuations in biochemical composition of the primary producers (mainly unicellular algae) available to the adult population (Léger et al. 1987a). Cyst products from inland resources are more constant in fatty acid composition, albeit at suboptimal low levels. Appropriate enrichment techniques have thus been developed to improve the lipid profile of deficient Artemia strains, taking advantage of the indiscriminate filter-feeding behaviour of Artemia (Léger et al. 1987b). Applying simple methods, lipophilic compounds can easily be incorporated into Artemia before being offered as live feed (see Section 3.5.5). A number of other compounds also varies from strain to strain: nutritional components such as total amount of free amino acids, pigments (canthaxanthin), vitamin C, minerals and trace elements, as well as contamination with chemicals such as pesticides and heavy metals. In most cases these variations are not strain-specific, but correspond to different production conditions. Their effects on larviculture success are far less significant than nauplii fatty acid composition.

3.2.5 Cyst biology and diapause 3.2.5.1 Cyst morphology and physiology A schematic diagram of the ultrastructure of an Artemia cyst is given in Fig. 3.13. The cyst shell consists of three layers:

•

• •

Alveolar layer: This hard layer consists of lipoproteins impregnated with chitin and haematin. The haematin concentration determines the colour of the shell, i.e. from pale to dark brown. Its main function is to provide protection for the embryo against mechanical disruption and ultraviolet (UV) radiation. This layer can be completely removed (dissolved) by oxidation treatment with hypochlorite (cyst decapsulation, see Section 3.5.4; Bruggeman et al. 1980). Outer cuticular membrane: This protects the embryo from penetration by molecules larger than the carbon dioxide molecule (multilayer membrane with a very special filter function; acts as a permeability barrier). Embryonic cuticle: This transparent and highly elastic layer is separated from the embryo by the inner cuticular membrane (develops into the hatching membrane during hatching incubation). The embryonic cuticle is apparently impermeable to non-volatile solutes (De Chaffoy et al. 1978; Clegg & Conte 1980). The embryo is an undifferentiated gastrula, which is ametabolic when water content is below 10% and can be stored for long periods without losing its viability. The viability is affected when water levels are higher than 10% (start of metabolic activity) and when cysts are exposed to oxygen; i.e. in the presence of oxygen, cosmic radiation results in the formation of free radicals, which destroy specific enzymatic systems in the ametabolic Artemia cysts.

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3.2.5.2 Cyst metabolism and hatching Given favourable environmental conditions the metabolism and development of encysted embryos are rapidly reinitiated. When incubated in seawater the biconcave cyst swells and becomes spherical within 1–2 h. Respiration, RNA and protein synthesis begin within minutes (Clegg & Conte 1980), supporting the conclusion that encysted embryos contain all the components needed for these activities. After a period of postdiapause development, in the order of 8–24 h depending on temperature and salinity, the cyst shell (including the outer cuticular membrane) bursts (breaking stage) and the embryo surrounded by the hatching membrane becomes visible. The embryo then leaves the shell completely and hangs underneath the empty shell (the hatching membrane may still be attached to the shell). Through the transparent hatching membrane one can follow the differentiation of the pre-nauplius into the instar I nauplius which starts to move its appendages. Shortly thereafter, the hatching membrane breaks open (hatching) and the free-swimming larva emerges head-first. Dry cysts (water content from 2 to 5%; Fig. 3.15) are very resistent to extreme temperatures; i.e. hatching viability is not affected in the temperature range ⫺273°C (Skoultchi & Morowitz 1964) to 60°C; above 60°C and up to 90°C only short exposures can be tolerated. Hydrated cysts have far more specific tolerances: mortalities occur below ⫺18°C and above 40°C; a reversible interruption of the metabolism (viability not affected) occurs between ⫺18 and 4°C and between 33 and 40°C, although the upper and lower temperature limits vary slightly from strain to strain. Active cyst metabolism occurs between 4 and 33°C. Within this range, the hatching percentage remains constant but the nauplii hatch earlier as the temperature increases.

Fig. 3.15 Cellular metabolism in Artemia cysts (as a function of hydration level).


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As for other environmental conditions, optimal hatching outputs are reached in the pH range 8–8.5. As a consequence, the addition of NaHCO3 (up to 2 g l⫺1), to artificial or diluted seawater or to dense suspensions of cysts, results in improved hatching. This may be related to the optimal pH activity range for the hatching enzyme. Increased hatching has been reported with increasing oxygen level in the range 0.6–2 ppm, and maximal hatching above this concentration. To avoid oxygen gradients during hatching a good homogeneous mixing of the cysts in the incubation medium is required. As stated above, hatching in a higher salinity medium will consume more of the energy reserves of the embryo. Above a threshold salinity (varying from strain to strain, but ⬎90 g l⫺1 for most strains), the amount of water that can be taken up is insufficient to support the embryo’s metabolism. Optimal salinity for hatching is equally strain specific, but generally situated in the range 15–70 g l⫺1. Although the physiological role of light during the hatching process is poorly understood, brine shrimp cysts, when hydrated and in aerobic conditions, need a minimal light triggering for the onset of the hatching process, related to light intensity and/or exposure time. Little is known about the exact light requirements, but generally strong illumination (about 2000 lux at the water surface) is recommended, at least during the first hours after complete hydration. Finally, hatchability of cysts is largely determined by the conditions and techniques applied for harvesting, cleaning, drying and storing of the cyst material (see Section 4.4). Hatching quality in stored cysts slowly decreases when cysts contain 10–35% water (Fig. 3.15). This process may, however, be retarded when the cysts are stored at freezing temperatures. The exact optimal water level within the cyst is not known, although there are indications that too severe dehydration (down to 1–2%) results in a drop in viability. Water content of about 5% is a reasonable value. Water levels in the range 30–65% initiate metabolic activity, eventually reducing the energy content to levels insufficient to reach the state of emergence under optimal hatching conditions (Fig. 3.15). Furthermore, a depletion of the energy reserves occurs when the cysts undergo subsequent dehydration/hydration cycles. Long-term storage of such material may result in a substantial decrease in hatching success. Cysts exposed for too long a period to water levels exceeding 65% will have completed their pre-emergence embryonic development. Subsequent dehydration of these cysts will, in the worst case, result in the killing of the now differentiated embryos. Sufficiently dehydrated cysts only keep their viability when stored under vacuum or in nitrogen. The presence of oxygen results in a substantial reduction in hatching success as a result of the formation of highly detrimental free radicals. Even correctly packaged cysts are preferentially stored at low temperatures. When frozen, the cysts should be acclimated for 1 week at room temperature before hatching. 3.2.5.3 Diapause As Artemia is an inhabitant of biotopes characterised by unstable environmental conditions, its survival during periods of extreme conditions (e.g. desiccation, extreme temperatures, high salinities) is ensured by the production of dormant embryos. Artemia females can indeed easily switch from the production of live nauplii (ovoviviparity) to cyst formation (oviparity) in response to fluctuating circumstances. The basic mechanisms involved in this switch are

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not yet fully understood, but sudden fluctuations in, for example, oxygen levels and salinity seem to trigger oviparity. The phenomenon of diapause in Artemia cysts has been the subject of numerous studies and reviews (Clegg & Conte 1980; Drinkwater & Crowe 1987; Lavens & Sorgeloos 1987; Drinkwater & Clegg 1991). The study of the mechanisms of diapause induction and deactivation can help to maximise the hatching yield of the commercially important Artemia cysts. The termination of diapause is a complex process, influenced by multiple and mutually interfering genotypical and environmental factors. The triggering mechanism for the induction of the diapause state is not yet known. In principle, Artemia embryos released as cysts in the medium are, or soon become, diapausing (Clegg et al. 1996). Upon the deactivation of diapause by environmental factors (Drinkwater & Crowe 1987), cysts enter the stage of quiescence and metabolic activity can be resumed when they are exposed to favourable conditions in terms of temperature, oxygen and light, eventually resulting in hatching. In this phase the metabolic arrest is uniquely dependent on external factors. As a result, synchronous hatching occurs, resulting in a fast start and consequent development of the population shortly after the re-establishment of favourable environmental conditions. This allows effective colonisation in temporal biotopes. Moreover, diapause or a similar state can be reinduced by exposure to anoxia (Drinkwater & Clegg 1991; Clegg 1993; Clegg et al. 2000) or by a well-dosed heat shock (Abatzopoulos et al. 1994). For the user of Artemia cysts, several techniques have proven successful in terminating diapause. It is important to note here that the sensitivity of Artemia cysts to these techniques shows strain- or even batch-specificity (see Section 3.2.4), hence the difficulty in predicting the effect on hatching outcome. When working with new or relatively unknown strains, the relative success or failure of certain methods has to be found out empirically. In many cases, the removal of cyst water is an efficient way to terminate the state of diapause. This can be achieved by drying the cysts at temperatures not exceeding 35–40°C or by suspending the cysts in a saturated NaCl brine solution (300 g l⫺1). As some form of dehydration is part of most processing and/or storage procedures, diapause termination does not require any particular extra manipulation. Nevertheless, with some strains of Artemia cysts, the usual cyst processing techniques do not yield a sufficiently high hatching quality, indicating that a more specific diapause deactivation method is necessary. The following procedures have proven to be successful when applied with specific sources of Artemia cysts.

• •

Freezing: This ‘imitates’ the natural hibernation period of cysts originating from continental biotopes with low winter temperatures (GSL, Utah, USA; continental Asia). Incubation in H2O2 solution: In most cases, the sensitivity of the strain (or batch) to this product is difficult to predict, and preliminary tests are needed to provide information about the optimal dose and period to be applied, and about the maximal effect that can be obtained. Overdosing results in reduced hatching or even complete mortality as a result of toxicity of the chemical. In some cases no effect at all is observed. In general, other diapause termination techniques (cyclic dehydration/hydration, decapsulation, other chemicals) give rather erratic results and/or are not user friendly. However, one should keep in mind that the increase in hatching percentage after any procedure may (even partially) be the result of a shift in hatching rate (earlier hatching).


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3.3 Production Methods: Tank Production of Artemia Biomass 3.3.1 Advantages of tank production and tank-produced biomass Although tank-produced Artemia biomass is far more expensive than pond-produced brine shrimp its advantages may, depending on the local condition, justify its application:

• • •

year-round availability of ongrown Artemia, independent of climate or season; specific stages (juveniles, preadult, adults) or prey with uniform size can be harvested as a function of size preferences of the predator; quality of the Artemia can be better controlled (e.g. nutritional content, free from diseases).

High-density intensive culture techniques offer two main advantages compared with pond production techniques. First, there is no restriction with regard to production site or time, since the culture procedure does not require highly saline waters or specific climatological conditions. Secondly, controlled production can be performed with very high densities of brine shrimp, e.g. several thousand animals per litre versus a maximum of a few hundred animals per litre in outdoor culture ponds. As a consequence very high production yields per volume of culture medium can be obtained with tank rearing systems. Since the early 1990s, several superintensive Artemia farms have been established, e.g. in the USA, France, the UK and Australia, to supply local demand. Depending on the selected culture technology and implantation facilities, production costs are estimated at US $2.5–12 kg⫺1 live weight Artemia, with wholesale prices varying from US $25 to 100 kg⫺1. In practice, when setting up an Artemia culture one should start by listing the prevailing culture conditions and available infrastructure. The abiotic and biotic conditions relevant for Artemia culture are:

•

• • •

physicochemical conditions: – ionic composition of the culture media – temperature – salinity – pH – oxygen concentration – water quality (nitrogen metabolites, particles, etc.) Artemia: – strain selection – culture density feeding: – feeding strategy – selection of suitable diets infrastructure: – tank and aeration design – filter design – recirculation unit

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•


– heating – feeding apparatus culture techniques: – stagnant culture – open flow-through system – recirculation type.

3.3.2 Physicochemical conditions Salinity and ionic composition of the culture media Although, in the wild, Artemia only occurs in highly saline waters (mostly above 100 g l⫺1), brine shrimp do thrive in natural seawater. The best physiological performance, in terms of growth rate and food conversion efficiency, is at salinities from 32 to 65 g l⫺1, depending on the cultured strain. For Artemia culture, the use of natural seawater of 35 g l⫺1 is the most practical. Small adjustments in salinity can be made by adding brine or diluting with tap water free from high levels of chlorine. Beside natural seawater or diluted brine, several artificial media with different ionic compositions are used with success in indoor installations for brine shrimp production. Since ionic composition is so important, concentrated brine (150 g l⫺1) from salinas can also be transported to the culture facilities and diluted with freshwater before use. Temperature, pH and oxygen concentration Temperature must be maintained between the specific optimal levels of the selected Artemia strain. For most strains a common range of preference is 19–25°C (Table 3.2). In the literature, it is generally accepted that the pH tolerance of Artemia ranges from 6.5 to 8. The pH tends to decrease during culture as a result of denitrification processes. When the pH drops below 7.5 small amounts of NaHCO3 (technical grade) should be added to increase the buffer capacity of the culture water. The pH is commonly measured using a calibrated electrode or with simple analytical test kits. In the latter case, the method must be suitable for seawater. With regard to oxygen, biomass production will decrease at concentrations below 2 mg l⫺1. For optimal production, oxygen concentrations higher than 2.5 mg l⫺1 are suggested. However, continuously maintaining oxygen levels higher than 5 mg l⫺1 will result in the production of pale animals (low in the respiratory pigment haemoglobin), possibly with a lower individual dry weight, which may therefore be less perceptible to and attractive for the predators. A dark red coloration (high haemoglobin content) is easily obtained by applying regular oxygen stresses (by switching off the aeration for a few minutes several times a day) a few days before harvesting. Oxygen levels should be checked regularly as they may drop significantly, especially after feeding. Oxygen is conveniently measured in the culture tank with a portable oxygen electrode. When oxygen occasionally drops below 30% saturation (i.e. 2.5 mg O2 l⫺1 in seawater of 32 g l⫺1 salinity), aeration intensity should be increased temporarily or air stones added. If oxygen levels remain low, the aeration capacity should be increased. It is important to remember that for a given air flow, the oxygen level is increased more effectively by small air bubbles than by large ones. However, very small air bubbles can become trapped between the thoracopods, causing the animals to float and congregate at the surface.


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Table 3.2 Effect of temperature on different production parameters for various geographical strains of Artemia. Temperature (°C) Geographical strain

20.0

22.5

25.0

27.5

30.0

32.5

San Francisco Bay, California, USA Survival (%) 97 Biomass production (%)a 75 Specific growth rateb 0.431g Food conversionc 3.89e

97 101 0.464d 3.35d

94 100 0.463d,e 3.64e

91 94 0.456e 3.87e

66 88 0.448f 4.15f

na na na na

Great Salt Lake, Utah, USA Survival (%) Biomass production (%)a Specific growth rateb Food conversionc

85 104 0.437f 2.90e

89 122 0.454e 2.65d,e

89 128 0.460d,e 2.62d

87 135 0.465d 2.40d

88 78 0.406g 4.14g

Chaplin Lake, Saskatchewan, Canada Survival (%) 72 Biomass production (%)a 78 0.422f Specific growth rateb c Food conversion 3.42e

75 102 0.452d,e 3.00d

77 108 0.459d 3.03d

65 106 0.456d 3.11d

50 90 0.437e,f 3.72d

na na na na

Tanggu, PR China Survival (%) Biomass production (%)a Specific growth rateb Food conversionc

94 61 0.343e 5.42e

91 80 0.371d 4.46d,e

93 92 0.387d 3.84d

84 85 0.378d 4.22d,e

54 16 0.208g 22.04g

77 69 0.392g 3.79h

95 41 0.299f 7.22f

Data compiled from Vanhaecke and Sorgeloos (1989). a Expressed as % recorded for the Artemia reference strain (San Francisco Bay, batch 288–2596) at 25°C after 9 days’ culturing on a diet of Dunalliella cells; bSpecific growth rate k ⫽ ln(Wt ⫺ W0) T⫺1, where T ⫽ duration of experiment in days (⫽9); cFood conversion ⫽ F(Wt ⫺ W0), where F ⫽ ␮g dry weight Dunalliella offered as food, Wt ⫽ ␮g dry weight Artemia biomass after 9 days’ culturing, and W0 ⫽ ␮g dry weight Artemia biomass at start of experiment; d–gMeans with the same superscript letter are not significantly different at the p ⬍ 0.05 level. na, not analysed.

Water quality The quality of the culture medium is primarily affected by excess particles as well as by soluble waste products such as nitrogenous compounds. High levels of suspended solids will affect production characteristics, either by their interference with uptake of food particles and propulsion by the Artemia, or by enhancing bacterial growth that will compete for oxygen and eventually infest the culture tank. Soluble waste products give rise to toxic nitrogenous compounds. The tolerance levels in Artemia for ammonia, nitrite and nitrate in acute and chronic toxicity tests with, for instance, GSL brine shrimp larvae showed no significant effect on survival [median lethal concentration ⫺1 (LC50)] or growth for concentrations up to 1000 mg l⫺1 NH⫹ NO⫺2 (Chen et al. 4 , 320 mg l 1989). For nitrate, no effects were observed at 1000 mg l⫺1 and it is therefore considered non-toxic. It is therefore unlikely that N-components will interfere directly with Artemia cultures. Nevertheless, the presence of soluble substances should be restricted as much as possible since they are an ideal substrate for bacteria. Excess soluble waste products can only be eliminated by diluting the culture water with clean water, be it new or recycled.

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3.3.3 Artemia strain selection and culture density Strain selection Based on laboratory results (Table 3.2), guidelines are provided for strain selection as a function of optimal temperature and culture performance. The optimal strain should be selected according to specific culture conditions. Culture density of Artemia Unlike other crustaceans, Artemia can be cultured at high to very high densities without affecting survival. Depending on the applied culture technique, inoculation densities up to 5000 larvae per litre for batch culture, 10,000 for closed flow-through culture and 18,000 for open flow-through culture can be maintained without influencing survival and growth. Above these densities, culture conditions become suboptimal (water quality deterioration, lower individual food availability), and growth and survival decrease (Table 3.3). Crowding seems to affect ingestion rate and thus growth. In stagnant systems, a clear decrease in growth rate with increasing animal density has been observed (Dhont et al. 1993). The costeffectiveness of a culture increases with increasing Artemia density. In an open flow-through system, maximal densities will be limited by feeding rate, while in recirculating and stagnant system the preservation of water quality will determine a safe feeding level, which in turn determines the animal density at which the individual feed amount still allows a satisfactory growth rate. After some culture trials with increasing animal densities, the maximal density can be identified as the highest possible density where no growth inhibition occurs.

3.3.4 Feeding Artemia is a continuous, non-selective, particle-filtering organism. Various factors may influence the feeding behaviour of Artemia by affecting the filtration rate, ingestion rate and/or assimilation: the quality and quantity of the food offered, the developmental stage of the larvae and the culture conditions. More detailed information about these processes is given in Coutteau and Sorgeloos (1989). Selection of a suitable diet Artemia can take up and digest exogenous microflora as part of the diet. Bacteria and protozoans, which develop easily in the Artemia cultures, are able to biosynthesise essential nutrients as they use the supplied brine shrimp food as a substrate. In this way they compensate for possible deficiencies in the diet composition. The interactions with bacteria make it a hard task to identify nutritionally adequate diets per se, and growth tests are difTable 3.3 Directive animal densities under different culture conditions. Culture system Open flow-through Closed flow-through Stagnant

Animals/litre 18,000 ⬎10,000 5000–10,000 5000 20,000

Culture period

Growth

Reference

To adult To adult To adult 7 days 7 days

High Moderate High High Low

Tobias et al. (1979) Lavens et al. (1986) Dhont et al. (1991)


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ficult to run under axenic conditions. As a consequence, nutritional composition of the diet does not play the most critical role in the selection of diets suitable for high-density culture of brine shrimp. The criteria used generally include:

• • • • • • •

availability and cost particle size composition (preferentially ⬍50 ␮m) digestibility consistency in composition among different batches and storage capacity solubility (minimal) food conversion efficiency (FCE) buoyancy.

Commonly used food sources are listed below.

•

•

•

•

Microalgae: These undoubtedly yield best culture results but it is rare that sufficient algae are available at a reasonable cost. Mass culturing of suitable algae for Artemia is most often economically unrealistic, so their use can only be considered in locations where algal production is an additional feature of the main activity. Furthermore, not all species of unicellular algae are considered capable of sustaining Artemia growth (D’Agostino 1980); For example, Chlorella and Stichococcus have a thick cell wall that cannot be digested by Artemia, Cocochloris produces gelatinous substances that interfere with food uptake and some dinoflagellates produce toxic substances. Dried algae: In most cases algal meals give satisfactory growth performance, especially when water quality conditions are kept optimal. Drawbacks in the use of these feeds are their high cost (⬎US $12 kg⫺1), as well as their high fraction of water-soluble components, which cannot be ingested by the brine shrimp, but will interfere with the water quality of the culture medium. Bacteria and yeasts: Single-cell proteins (SCP) have several characteristics that make them an interesting alternative to microalgae: – the cell diameter is mostly smaller than 20 ␮m – the nutritional composition is fairly complete – the rigid cell walls prevent the leakage of water-soluble nutrients in the culture medium – products are commercially available at acceptable cost (e.g. commonly used in cattle feeds). The highly variable production yields that often occur when feeding a yeast mono-diet are assigned to nutritional deficiencies of the yeast diet and should therefore be met by supplementation with other diets. For some SCP, digestibility by the Artemia can be a problem. Complete removal of the complex and thick yeast cell wall by enzymatic treatment and/or supplementation of the diet with live algae significantly improve the assimilation rate and growth rate of the brine shrimp (Coutteau et al. 1992). By-products from the food industry: Non-soluble by-products from agricultural crops or from the food-processing industry, such as rice bran, corn bran, soyabean pellets and lactoserum, appear to be a very suitable feed source for high-density culturing of Artemia (Dobbeleir et al. 1980). Their main advantages are their low cost and world-wide availability. Equally important in the evaluation of dry food is the consistency of the food quality and supply, and the possibility for storage without loss of quality. Bulk products must be stored in a dry and, preferentially, cool place.

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1

3 2

T

Fig. 3.16 Feeding strategy with cultured Artemia. (1) Look through looking glass to turbidistick (or submerge stick in cylinder with appropriate mesh). (2) Submerge turbidistick until contrast between black and white disappears. (3) Read depth of submergence in cm ⫽ T. During first week: T ⬍ 15 cm: stop feeding and/or increase water renewal; 15 cm ⬍ T ⬍ 20 cm: maintain actual feeding ratio; T ⬎ 20 cm: increase feeding ratio and/or add food manually. During next week: T ⬍ 20 cm: stop feeding and/or increase water renewal; 20 cm ⬍ T ⬍ 25 cm: maintain actual feeding ratio; T ⬎ 25 cm: increase feeding ratio and/or add food manually.

Feeding strategy Since Artemia is a continuous filter-feeding organism, highest growth and minimal deposition of unconsumed food is achieved when food is distributed as frequently as possible. When feeding SCP, algae or yeast, concentrations should be maintained above the critical minimum uptake concentration, which is specific for the algal (or other) species and the developmental stage of Artemia (Abreu-Grobois et al. 1991). Levels of dry feeds, consisting of fragments and irregular particles, cannot be counted in the culture tank. Therefore, a correlation between optimal feed level and turbidity of the culture water has been developed, whereby the feed concentration in a culture tank is determined by measuring the turbidity of the water with a simplified Secchi-disc (Fig. 3.16).

3.3.5 Infrastructure Tank and aeration design Artemia can be reared in containers of any possible shape as long as the installed aeration ensures proper oxygenation and adequate mixing of feed and animals throughout the total culture volume. However, aeration should not be too strong. Thus, aeration and tank design must be considered together as the circulation pattern is determined by the combination of both. A wide variety of culture tanks has proven to be suitable (Dhont & Lavens 1996). For cultures up to 1 m3, rectangular tanks are most convenient. They can be aerated either with an air–water lift (AWL) system (Fig. 3.17), by an aeration collar mounted around a central standpipe or by perforated polyvinyl chloride (PVC) tubes fixed to the bottom of the tank.


89

Fig. 3.17 Air–water lift.

For large volumes (⬎1 m3), it is advantageous to switch to cement tanks covered on the inside with impermeable plastic sheets or coated with special paint. These large tanks are traditionally operated as raceway systems. They are oblong, approximately 1.5 m wide and with a height:width ratio kept close to 1:2. The length is then chosen according to the desired volume. The corners of the tank may be curved to prevent dead zones where sedimentation can take place. A partition, to which AWLs are fixed, is installed in the middle of the tank and ensures a combined horizontal and vertical movement of the water, which results in a screw-like flow pattern (Bossuyt & Sorgeloos 1980). If axial blowers are used for aeration, the water depth should not exceed 1.2 m to ensure optimal water circulation. Filter design The most important and critical equipment in flow-through culturing is the filter used for efficient evacuation of excess culture water and metabolites without losing the brine shrimp from the culture tank. These filter units should be able to operate without clogging for at least 24 h, to reduce risks of overflowing. Traditionally, filters are constructed as a PVC frame around which an interchangeable nylon screen is fixed. The aeration is positioned at the bottom of the filter, ensuring a continuous friction of air bubbles against the sides of the filter screen and thus reducing filter-mesh clogging (Fig. 3.18). A more sophisticated type of cylindrical filter system consists of a welded-wedge screen cylinder, made of stainless steel. This welded-wedge system has several advantages with respect to the filter-screen types:

• •

Larger particles with an elongated shape can still be evacuated through the slit openings. The specially designed V-shape of the slit openings creates specific hydrodynamic suction effects, as a result of which filter particles that are only slightly smaller than the slit opening are sucked through.

90


65

filter bag

70

20

PVC frame

60

air

55

65

60

20

15

aeration collar 25 4

55

Fig. 3.18 Construction of filters used in Artemia culture (dimensions in cm).

•

Possible contact points between particles and filter are reduced to two instead of four mesh borders, which consequently reduces the chances of clogging.

This filter can be operated autonomously for much longer periods than traditional mesh filters. Therefore, proportionally smaller welded-wedge filters can be used, leaving more volume for the animals in the culture tank. As brine shrimp grow, the filter is regularly switched for one with a larger mesh or slit opening to achieve maximal evacuation of moults, faeces and other waste particles from the culture tanks. A set of filters covering a 14 day culture period should consist of approximately six different slit or mesh openings ranging from 120 to 350 ␮m. Heating When ambient temperature is below the culture optimum range (25–28°C), heating is imperative. Small volumes (⬍1 m3) are most conveniently heated using electric thermoregulated resistors. Depending on the ambient temperature, a capacity up to 1000 W m⫺3 must be provided. For larger volumes, a heat exchanger consisting of a thermostatically controlled boiler with copper tubing under the bottom of the culture tank is recommended. Heat losses can be avoided by insulating the tanks with Styrofoam and covering the surface with plastic sheets. Feed distribution apparatus Dry feed cannot be distributed directly to the culture tank, but should be suspended by mixing it in tap water or seawater beforehand. The feed suspension is distributed to the culture tanks via a timer-controlled pump. The volume of the food tank should be big enough to hold the highest daily food ration at a concentration maximum of 200 g food l⫺1. Even at those concentrations, the food suspension is so thick that there is a high risk of blocking of the food lines.


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3.3.6 Culture techniques Depending on the objectives and the opportunities, different culture procedures for highdensity intensive Artemia production may be applied. The final selection of the type of culture installation will be subject to local conditions, production needs and investment possibilities. Decisions need to be made as to: (a) whether or not the water should be renewed (open flow-through); and (b) in the latter case, whether a particular water treatment should be applied (closed flow-through or stagnant or batch system). There are many kinds of transition types, ranging from open flow-through with 0% recirculation to closed flow-through with 100% recirculation. In reality, even at complete recirculation, a small part of the culture water must be regularly renewed. The culture system should be designed in such a way that the water quality can be maintained as close to optimal as possible. This means that the concentration of particles and soluble metabolites should remain minimal to prevent toxicity problems, proliferation of micro-organisms and interferences with the filter-feeding apparatus of the brine shrimp. Stagnant systems Stagnant systems are the simplest concept for intensive Artemia culture: no wastewater evacuation, filter systems or water treatment are involved. The culture is started in an aerated tank and biomass is harvested after a reduced culture period. The main disadvantage is that high animal densities do not allow for extended culture periods because of the degradation of the water quality. Successful trials with 10 animals l⫺1 on micronised soya pellets yielded Artemia juveniles of 3 mm in length and over 75% survival in 7 days (Dhont et al. 1993). Open flow-through A discontinuous or continuous renewal of culture water by clean seawater, with consequent dilution of particulate and dissolved metabolites, will result in the best possible culture conditions and highest production capacities. Application of an open flow-through culture technique, however, is limited to those situations where large volumes of sufficiently warm seawater (or brine) are available at relatively low cost, or where large quantities of algal food are available, such as from effluents from artificial upwelling projects, tertiary treatment systems or intensive grow-out ponds of shrimp. The water retention time is chosen so as to reach an optimal compromise between efficient evacuation of wastewater and minimal food losses. A very simple semi flow-through system has been developed by Dhert et al. (1992). The system does not require the use of feeding pumps and involves minimal care. The pilot system consists of six oval raceway tanks of 1 m3 and six reservoir tanks of the same capacity placed above each culture tank. These reservoir tanks hold seawater and food (squeezed rice bran suspension), and need manual refilling only once or twice a day. They are slowly drained to the culture tanks, and flow rate is easily adjusted by means of a siphon of a selected diameter. Retention time is at least 12 h. The culture effluent is drained using weldedwedge filters as described above. This technique involves minimal sophistication and appears to be very predictable in production yields, which are between those obtained in batch and flow-through systems (see Section 3.3.9 for production figures).

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Closed flow-through (recirculation) systems When only limited quantities of warm seawater are available, open flow-through systems cannot be considered. High-density flow-through culturing of Artemia can only be sustained by recirculating the culture water via a water treatment unit. This unit should be designed to remove particles and decrease levels of harmful nitrogen components. Several suitable recirculation systems have been designed, proving that more than one solution exists to treat effluents from intensive Artemia cultures. However, at least one option will be most appropriate to a given situation when local conditions are taken into account. In general, a recirculation unit consists of a mechanical treatment that removes flocculations, particles, debris, etc., and a biological treatment that breaks down ammonia to nitrite and nitrate, and lowers the biological oxygen demand (BOD). A unit that removes the soluble fraction is also included. An overview of the most frequently used treatment components is presented in Table 3.4.

3.3.7 Control of infections Heavy losses of pre-adults may be due to infections with the filamentous bacterium Leucothrix, which particularly occur in nutrient-rich media (Solangi et al. 1979). The Leucothrix colonies fix on to the exoskeleton, by preference on the thoracopods, and become visible only from instar V/VI stage onwards. The brine shrimp suffer physically, as the movements of their thoracopods become affected and, consequently, filtration rates are reduced. Ultimately, growth and moulting are arrested, and overfeeding of the tanks occurs, eventually resulting in a collapse of the Artemia culture. One cure may be the application of tetramycin; however, antibiotics cannot be used in recirculation systems as they will affect the biological treatment unit. Raising salinity from 35 to 50–60 g l⫺1 appears to be the most practical solution, coupled with a higher water renewal rate of 25% instead of 10% on a weekly basis (Lavens et al. 1986). A second observed disease in Artemia cultures is the ‘black disease’. Black spots appear primarily on the extremities, such as on the thoracopods and antennae. This disease consists of the detachment of the epidermis from the cuticula, and is caused by a dietary deficiency, which interferes with lipid metabolism (Hernandorena 1987). In high-density cultures of Artemia using agricultural by-products as a food source, the black disease is observed when water quality deteriorates (probably interfering with the composition of the bacterial population and consequently the diet composition) and/or when feeding rates are not optimal. Improving these conditions does not save the affected animals, but appears to avoid further losses.

Table 3.4 Water treatment systems used in intensive Artemia culture. Treatment

Type

Reference

Mechanical treatment

Plate separator Cross flow sieve Foam fractioner or protein skimmer

Lavens et al. (1986) Brisset et al. (1982) Lim et al. (2001)

Biological treatment

Rotating disc contactor

Bossuyt & Sorgeloos (1980)


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3.3.8 Harvest and processing of cultured Artemia Harvesting of high-density cultures of Artemia can be facilitated by taking advantage of the surface respiration behaviour of the animals. When the aeration in the culture tank, together with the flow-through and the automatic feeding are interrupted, oxygen levels in the water drop very quickly and all waste particles sink to the bottom; after about 30 min the Artemia respond to the oxygen depletion by concentrating at the water surface, where they perform surface respiration. The concentrated population, free from suspended solids, can easily be scooped out with a net of an appropriate mesh size. When the culture water is not loaded with particles, brine shrimp can be harvested by draining the complete culture over a sieve, which should be partially submerged. The Artemia should be washed thoroughly in freshwater or seawater. The harvested Artemia can then be offered as live food for freshwater as well as marine predators. The salinity of the predator culture water is of no concern, as Artemia is a hypo-osmoregulator, and its body fluids have a constant and low salt content of about 9 g l⫺1. In seawater, they remain alive without feeding for several days. When transferred into freshwater, Artemia will continue to swim for another 5 h, after which time they eventually die as a result of osmoregulatory stress. Live brine shrimp can be transported in plastic bags containing cooled seawater under oxygen. Harvested Artemia that are not for immediate consumption can be frozen or dried in flakes. To ensure optimal product quality, brine shrimp biomass must be frozen immediately after thorough washing with freshwater when still alive. The biomass should be spread out in thin layers (1 cm) in plastic bags or on ice trays, and transferred to a quick freezer (at least ⫺15°C). The adult exoskeleton is not damaged when the biomass is frozen properly. Upon thawing, the Artemia cubes yield intact animals, which do not pollute the water by leaching of body fluids.

3.3.9 Production figures of intensive Artemia cultures Figure 3.19 provides a summary of average production data expressed as Artemia survival and length, obtained in the different culture systems described in this chapter. After 8

100

7 6 growth (mm)

survival (%)

80 60 40

5 4 3 2

20

1 0

0 0

2

4

6

8

10

time (days)

12

14

0

2

4

6

8

10

12

14

time (days)

Fig. 3.19 Production figures of various intensive Artemia cultures. (ⵧ) 1000 litre open flow-through culture on rice bran diet (Dhert et al. 1992); (䉫) 200 litre open flow-through culture on Chaetoceros diet (modified from Lavens 1989); (⌬) 300 litre closed flow-through culture on a mixture of soyabean waste and corn bran (modified from Lavens 1989); (䊊) 3 m3 closed flow-through raceway culture on rice bran diet (modified from Lavens 1989); (⫻) 500 litre batch culture on a mixture of pea and corn bran (Dhont, unpublished data).

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2 weeks of culturing, pre-adult or adult Artemia with an average length of 5 mm or more can be harvested. In a flow-through culture there is a slight but continuous mortality during the whole of the culture period. No significant differences in survival are observed between open flow-through and recirculating cultures (Fig. 3.19). In stagnant cultures, there is a significantly higher mortality during the end of the first week of culture, which can be explained by the deterioration of the water quality, probably because the early naupliar stages are more sensitive than the juvenile or pre-adult stages. Average production yields harvested after 2 weeks (live wet weight Artemia biomass relative to tank volume) amount to 5, 15 and 25 kg m⫺3 for batch production, and flow-through systems using micronised feeds and live algae, respectively. These differences in production figures are mainly the result of differences in maximum stocking density at the start and survival at the end of the culture trial.

3.4 Biochemical Composition 3.4.1 Proximate composition Dealing with the biochemical composition of any living organisms usually involves a great deal of generalisation. Presented figures inevitably consist of average values that may conceal subtle changes or intriguing differences. In the case of Artemia, distinctions between species, sources and life-stage must be made, owing to its specific nature and the range of its applications. Unlike algae or rotifers, different life stages of Artemia are used as larval food: from the embryonic form (as decapsulated cysts), through non-feeding nauplii and enriched nauplii, to adult biomass. These life stages may show important biochemical differences. Different Artemia strains or even Artemia from the same strain but from another batch or a different seasonal harvest may show variation in their composition. However, the most important differences in composition can be observed with the exogenous feeding stages (instar II and later). Since Artemia is a non-selective filter-feeder with a relatively high ratio of gut content to body volume, its composition is highly dependent on its diet. This section aims to give a comprehensive picture of the composition of various Artemia forms (see Table 3.5). 3.4.1.1 Cysts and decapsulated cysts Since the chorion of Artemia cysts is completely indigestible by all known cultured species, the biochemical composition of intact cysts could be considered irrelevant, since they cannot be used as a food source. However, once this chorion has been removed by chemical decapsulation (see Section 3.5.4) the remaining embryo is digestible, and has been used successfully in the larviculture of carp, catfish, milkfish and some marine shrimp. 3.4.1.2 Nauplii There is a crucial distinction between the first larval stage or instar I, which is nonfeeding, and subsequent stages (instar II and further), which have a functional digestive


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Table 3.5 Proximate composition of different developmental stages of Artemia (% on a dry weight basis). Artemia stage Source (refs)

Protein (%)

Lipid (%)

Carbohydrate Ash (%) (%)

Fibre (%)

DW (␮g)

Cysts GSL, USA (a) SFB, USA (e) Mexico (e)

55.8 45.2 41.4–50.2

11.2 3.9 0.3–1.0

6.9 36.3 36.4

— — —

4.83 — —

Decapsulated cysts GSL, USA (a) SFB, USA (d)

50.6 67.4

14.7 15.7

6.6 —

10.6 —

— —

3.42 —

Nauplii GSL, USA (a) GSL, USA (b) GSL, USA (c) PR China (b) France (b) SFB, USA (b,d)

56.2 41.6–47.2 61.9 47.3 55.7 41.9–59.2

17.0 20.9–23.1 14.4 12.0 12.4 15.9–27.2

3.6 10.5 10.6 — — 11.2

7.6 9.5 7.1 21.4 15.4 8.17

— — 5.9 — — —

2.31 1.65–2.70 — 3.09 2.7–3.1 1.45–2.87

Adults: wild population San Diego, USA (b) SFB, USA (b) Italy (g)

64.0 50.2–58.0 41.9

12.0 2.4–19.3 3.5

— 17.2 —

20.6 29.2 —

— — —

— — —

Adults: cultured GSL, USA (b,c,f) France (b) SFB, USA (d) Italy (g)

50.8–67.4 53.7 39.4–64.0 55.4

10.8–30.6 9.4 4.5–12.1 4.0

4.0–12.3 — — 20.0

5.2–13.6 21.6 — 20.6

4.2 — — —

— — — —

5.9 5.2 5.8–12.6

References: a, García-Ortega et al. (1998); b, Léger et al. (1986); c, Lim et al. (2001); d, Dendrinos and Thorpe (1987); e, Correa Sandoval et al. (1993); f, Correa Sandoval et al. (1994); g, Trotta et al. (1987). DW, dry weight; GSL, Great Salt Lake; SFB, San Francisco Bay; —, not mentioned.

system. Instar I nauplii survive by depleting their yolk reserves, and gradually decrease in nutritional value and in energetic content. Their initial composition reflects the parental characteristics, both genetic and phenotypic, while from instar II nauplii onwards, the composition will also be influenced by the diet. Several publications offer data on naupliar composition, but few authors make a clear distinction between instar I and instar II stages. This is understandable given the fact that asynchronous hatching produces batches of nauplii consisting of different stages. 3.4.1.3 Juveniles and adults Ongrown Artemia are used much less frequently in aquaculture than nauplii, for the reason that, while nauplii can easily be obtained through simple hatching of widely available and storable cysts, culturing Artemia requires a considerable amount of labour and infrastructure. Even though various culture techniques have been developed to suit all kinds of local conditions (Section 3.3; Dhont & Lavens 1996), they often remain a significant extra investment. However, the nutritional value of ongrown Artemia compared with freshly hatched nauplii is superior, at least with respect to its protein quality and individual energetic content.

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3.4.2 Lipids The lipid fraction has undoubtedly received most attention in marine larviculture, yielding a wealth of published lipid analysis on all kind of Artemia strains and live stages. Published values seem to exhibit important differences (Table 3.6). Besides natural fluctuations, these differences may also arise from different analytical methodologies or inaccurate definitions of what exactly was analysed (e.g. exact strain or larval stage). 3.4.2.1 Cysts and nauplii The lipid content and profile of cysts and instar I nauplii are not affected by diet or environmental conditions. The differences in lipid profile that are observed between strains (Triantaphyllidis et al. 1995; Han et al. 2000b) can reflect either genetic characteristics or the lipid profile of the food of the parental population. However, the lipid profile is considered to be environmentally rather than genetically determined, as several authors have demonstrated that the fatty acid profile of Artemia adults and their offspring clearly reflects the composition of the parental diet, regardless of the strain (Vos et al. 1984; Millamena et al. 1988; Lavens et al. 1989; Navarro & Amat 1992). Many publications include values for 15 or more different fatty acids in Artemia nauplii, but according to Léger et al. (1986) only six fatty acids (16:0, 16:1n-7, 18:1n-9, 18:2n-6, 18:3n-3 and 20:5n-3) actually make up Table 3.6 Fatty acid composition of different developmental stages of Artemia (mg g⫺1 dry weight).

Artemia stage Source (refs)

Palmitic acid (16:0)

Cysts GSL, USA (a)

12.7

3.9

19.1

5.5

Decapsulated cysts GSL, USA (a,b) France (b)

16.1–25.7 16.5

5.0–8.1 9.8

24.2 20.9

13.2–19.4 3.6 12.6

4.1–7.4 0.1 6.7

15.7 18.6 14.4 23.6

Nauplii GSL, USA (a,c,d) SFB, USA (e) PR China (f,g,h) Urmia, Iran (g) A. parthenogenetica (g,h) Madagascar (i) A. persimilis, Arg (h) A. tibetiana, China (j) Adults GSL, USA (k) GSL, USA (l) SFB, USA (c)

3.9 9.1 4.5–14.5

Palmitoleic acid (16:1n-7)

Linolenic acid (18:3n-3)

EPA (20:5n-3)

DHA (22:6n-3)

27.2

3.2

0.1

6.9–11.1 34.2–49.4 7.3 22.7

3.9–4.7 6.2

0.1

20.3–34.8 6.1 17.8

5.7–10.1 28.6–40.0 1.8 8.2 11.0 3.6–39.3

tr

1.6

23.7

12.2

6.8

3.5–8.9 0.6 1.4–7.5 2.7 3.5–14.7

0.0–0.4 0.4 0.0–0.4

20.2 9.5 2.4

21.2 17.9 40.4

7.4 8.0 6.2

6.25 16.7 tr

24.5 0.0 44.7

0.0 0.3 0.2

1.7 0.0 3.5–13.5

1.7 2.8 2.5–3.9

0.8 0.0 0.1–0.4

1.7 4.3 0.2–1.0

Oleic acid (18:1n-9)

7.4 18.3 9.0–17.1

Linoleic acid (18:2n-6)

15.4 15.9 1.7–30.5

References: a, García-Ortega et al. (1998); b, Lavens et al. (1989); c, Estevez et al. (1998); d, Han et al. (2000a); e, Dendrinos and Thorpe (1987) (adults fed on yeast); f, Dhert et al. (1993); g, Triantaphyllidis et al. (1995); h, Han et al. (2000b); i, Triantaphyllidis et al. (1996); j, Han et al. (1999); k, Lim et al. (2001) (adults fed on rice bran); l, Lavens (1989) (adults fed on soya pellets). EPA, eicosapentaenoic acid; DHA, docosahexaenoic acid; GSL, Great Salt Lake; SFB, San Francisco Bay; tr, trace.


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about 80% of the total fatty acid pool in an Artemia sample. In later publications, some authors also detected significant levels of 18:1n-7, which was not reported earlier, probably reflecting increased chromatographic resolution (Estevez et al. 1998; Han et al. 2000b). Léger et al. (1986) compiled data on almost 150 fatty acid analyses of nauplii from about 20 different Artemia sources and came to the following conclusions. Oleic acid (18:1n-9) is often the most abundant fatty acid. It also has the most stable occurrence (lowest coefficient of variation). Together with palmitic (16:0) and palmitoleic acids (16:1n-7), it accounts for 40–60% of the total fatty acids in Artemia. The level of 16:0 is fairly constant over different strains, but levels of 16:1n-7 are more variable. Léger et al. (1986) concluded that Artemia nauplii contain between 0.4 and 33.6% linolenic acid (18:3n-3), but that the distribution of these levels is actually bimodal: 36% of the samples contain more than 20% of their total fatty acids as linolenic acid, while 43% of the samples contain less than 10% linolenic acid. The levels of EPA (20:5n-3) seem to be inversely related to linolenic acid levels. Most of these patterns can be recognised in the more recent compilation presented in Table 3.6. Only data expressed in mg g⫺1 dry weight have been listed here, as these not only reflect the relative proportions of the various fatty acids, but also indicate their quantities with reference to Artemia body weight. 3.4.2.2 Ongrown Artemia As soon as brine shrimp start feeding (at instar II), their lipid profile will quickly reflect the profile of their diet. This is the basis of the enrichment technique that is discussed in detail in Section 3.5.5. Just as for nauplii, the lipid composition of natural Artemia biomass will reflect the composition of its diet. The metabolic pathways and abilities of Artemia to convert one fatty acid to another are not yet entirely elucidated. Indications that Artemia converts docosahexaenoic acid (DHA) into EPA (McEvoy et al. 1995) were proven to be correct by Navarro et al. (1999). The conversion rate seems to vary according to the strain (Evjemo et al. 1997). Thus, it should be borne in mind that data on lipid composition of Artemia biomass reflects a variety of influences such as diet, season, geography, strain, life stage and physiological stage, and as such the figures presented only offer an indication of possible levels.

3.4.3 Proteins Some confusion appears when reviewing amino acid profiles of Artemia because of different methods of analysis or reporting the data. Nevertheless, it seems correct to state that protein levels and amino acid profiles show much less fluctuation between strains or life stages than, for example, the lipid fraction (Table 3.7). Adult Artemia have a slightly higher protein content than nauplii and contain slightly more essential amino acids than nauplii. The higher protein content of cysts compared with decapsulated cysts or nauplii is due to the presence of the chorion, which is composed of lipoprotein impregnated with chitin and haematin (García-Ortega et al. 1998). Nauplii contain markedly lower levels of free amino acids compared with wild copepods (Tonheim et al. 2000) but, generally, Artemia nauplii as well as adults contain sufficient levels of the 10 amino acids that are considered essential for fish larvae. Even so, methionine seems to be the first

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Live Feeds in Marine Aquaculture Table 3.7 Amino acid composition of different developmental stages of Artemia (g 100 g⫺1 protein). Stage (refs) Cysts(b)

Decaps. cysts (a,c)

Nauplii (a,c)

Juveniles and adults (b,c,e)

Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Cystine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Histidine Lysine Arginine Tryptophan

8.6 4.1 6.3 10.3 5.0 4.0 5.1 1.3 4.9 2.1 4.4 6.0 5.2 3.4 2.7 7.1 6.4 0.9

7.7–11.6 4.3–5.8 5.7–7.7 11.0–13.9 3.6 3.9–4.1 4.1–4.8 1.1–1.3 4.9–6.2 2.5–3.0 4.8–9.9 6.6 3.3–4.4 3.9–5.2 2.7–3.2 7.1–9.7 7.0–9.3 1.3

7.6–9.4 4.3–5.2 5.2–5.5 11.2–12.3 4.2 4.0–4.9 4.4–5.1 1.1–1.6 4.9–5.9 2.3–1.9 4.7–5.3 6.5–7.4 3.4–3.7 3.9–4.4 2.5–3.5 7.3–8.1 6.8–16.1 1.2

5.8–10.5 2.4–6.3 2.6–6.2 7.5–13.9 3.3–10.9 2.7–11.0 3.6–6.7 0.1–2.3 3.2–7.4 0.7–2.3 3.0–6.3 4.5–8.2 2.2–5.0 2.3–6.4 1.3–3.6 4.2–9.9 2.7–8.2 —

Total amino acid

87.8

85.5–111.6

85.5–105.7

52.1–125.1

References: a, García-Ortega et al. (1998); b, Lim et al. (2001) (adults fed on rice bran); c, Dendrinos and Thorpe (1987) (adults fed on yeast); e, Trotta et al. (1987). Decaps., decapsulated.

limiting amino acid when feeding nauplii to fish larvae (Fyhn et al. 1993; Conceição et al. 1997; Helland et al. 1999). Most protein in Artemia nauplii consists of small size proteins with a molecular weight between 7.4 and 49.2 kDa (García-Ortega 1999). The presence of these low molecular weight peptides and free amino acids in nauplii, together with their autolytic capacity and high solubility, accounts for the easy digestion of the proteins by fish larvae.

3.4.4 Vitamins Most data on vitamins relate to Artemia franciscana (Table 3.8). An account of differences between strains could only be found for ascorbic acid (AscA) forms in cysts in Merchie et al. (1995). They observed considerable differences in ascorbic acid 2 sulfate (AscAS) concentrations (296–517 ␮g g⫺1 dry weight expressed as AscA) when comparing 10 different strains. Furthermore, they provided further evidence for the complete conversion of the ascorbic acid 2 sulfate form to free ascorbic acid. Whether AscAS serves as a storage form of AscA to satisfy the brine shrimp’s larval requirements after hatching (Mead & Finamore 1969), or acts as a sulfating agent during embryonic development (Mead & Finamore 1969; Bond et al. 1972) is unclear. With the exception of AscA and thiamine, Artemia nauplii contain higher vitamin levels than natural marine zooplankton (Mæland et al. 2000) and according to the NRC standards (NRC 1993; Kaushik et al. 1998) Artemia seems to cover the minimal dietary requirements of


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Table 3.8 Vitamin levels in different developmental stages of Artemia (␮g g⫺1 dry weight). Cysts Ascorbic acid Ascorbic acid 2 sulfate Thiamin Riboflavin Niacin Pantothenic acid Vitamin B6 Biotin Folate Vitamin B12 Choline chloride Inositol Piridoxine HCl

861 ⫾ 16c 7.13d 23.2d 108.7d 72.6d 10.5d

Nauplii

Adults

692 ⫾ 89a

49b

7.5 ⫾ 1.1a 47.3 ⫾ 6.0a 187 ⫾ 8a 86 ⫾ 19a 9.0 ⫾ 5.0a 3.5 ⫾ 0.6a 18.4 ⫾ 3.4a 3.5 ⫾ 0.8a

27b 17b 130b 68b 1b 1b 3b 6100b 1200b 8b

Mæland et al. (2000); bSimpson et al. (1983); cMerchie et al. (1995); Léger et al. (1986).

a

d

fish larvae. However, exact dietary vitamin requirements have not been established for many marine fish larvae.

3.5 Applications of Artemia 3.5.1 The future use of Artemia in aquaculture Although there is no doubt that Artemia will gradually be replaced by formulated diets, the use of nauplii will continue to be market driven for at least a few more years (see also Section 3.1). Increased harvests at GSL and new locations may relieve the pressure or even reverse the current trends, but it seems far from redundant to improve and promote the existing applications of Artemia in live food production.

3.5.2 Hatching Although hatching Artemia cysts appears to be simple, several factors are critical for the successful hatching of the large quantities needed in larval fish production. Optimal hatching conditions are (Lavens & Sorgeloos 1996):

• • • • • •

constant temperature of 25–28°C 15–35 g l⫺1 salinity pH around 8.0 minimum oxygen levels of 2 mg l⫺1, preferably 5 mg l⫺1(see below) maximum cyst densities of 2 g l⫺1 strong illumination of 2000 lux.

All of these factors will affect the hatching rate and maximum output, and hence the production cost of the harvested Artemia nauplii. Best hatching results are achieved in containers

100


Fig. 3.20 Large-scale hatching set-up.

with a conical bottom, aerated from the bottom with air-lines (Fig. 3.20). Cylindrical or squarebottomed tanks will have ‘dead spots’ in which Artemia cysts and nauplii accumulate and suffer from oxygen depletion. Transparent or translucent containers will facilitate inspection of the hatching suspension, especially when harvesting. The underlying physiological processes of hatching are described in detail in Section 3.2.5.2, but practical implications are reiterated here. The aeration intensity must be sufficient to maintain oxygen levels above 2 mg l⫺1, preferably 5 mg l⫺1. The optimal aeration rate is a function of the tank size and the density of cysts incubated. Excessive foaming can be reduced by disinfection of the cysts before incubation and/or by the addition of a few drops of a non-toxic antifoam agent (e.g. silicone antifoam). The temperature of the seawater should be kept in the range of 25–28°C; below 25°C cysts hatch more slowly and above 33°C cyst metabolism is irreversibly stopped. The effect of more extreme temperatures on the hatching output is largely strain specific. Quantitative effects of the incubation salinity on cyst hatching are related primarily to the hydration level that can be reached in the cysts. Above threshold salinity, the cysts do not absorb sufficient quantities of water. This threshold value varies from strain to strain, but is approximately 85–90 g l⫺1 for most Artemia strains. Secondly, the incubation salinity will interfere with the amount of glycerol that needs to be built up to reach the critical osmotic pressure within the outer cuticular membrane of the cysts. The fastest hatching rates will thus be noted at the lowest salinity levels, since it will take less time to reach breaking. Optimal hatching can be obtained in the range 15–70 g l⫺1. For practical convenience, natural seawater is most often used to hatch cysts. For some sources of cysts, hatching at low salinity results in higher hatching efficiencies, and the nauplii have a higher energy content (Table 3.9). The pH must remain above 8 during the hatching process for optimal functioning of the hatching enzyme. If necessary, for instance when low-salinity water is used, the buffer


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Table 3.9 Effect of incubation at low salinity on hatching percentage, individual nauplius weight and hatching output for Artemia cysts from different geographical origins. Salinity Cyst source

35 g l⫺1

5 g l⫺1

% diff.

Hatching percentage San Francisco Bay, USA Macau, Brazil Great Salt Lake, USA Shark Bay, Australia Chaplin Lake, Canada Bohai Bay, PR China

71.4 82.0 43.9 87.5 19.5 73.5

68.0 86.4 45.3 85.8 52.2 75.0

⫺4.8 ⫹5.3 ⫹3.1 ⫺1.9 ⫹167.6 ⫹2.0

Naupliar dry weight (␮g) San Francisco Bay, USA Macau, Brazil Great Salt Lake, USA Shark Bay, Australia Chaplin Lake, Canada Bohai Bay, PR China

1.63 1.74 2.42 2.47 2.04 3.09

1.73 1.76 2.35 2.64 2.28 3.07

⫹6.1 ⫹1.1 ⫺2.5 ⫹6.9 ⫹11.8 ⫺0.6

Hatching output (mg nauplii g⫺1 cysts) San Francisco Bay, USA Macau, Brazil Great Salt Lake, USA Shark Bay, Australia Chaplin Lake, Canada Bohai Bay, PR China

435.5 529.0 256.5 537.5 133.8 400.5

440.2 563.7 257.0 563.3 400.4 406.0

⫹1.1 ⫹6.6 ⫹0.2 ⫹4.8 ⫹199.3 ⫹1.4

Modified from Vanhaecke et al. (1984).

capacity of the water should be increased by adding up to 1 g NaHCO3 l⫺1. Increased buffer capacity is also essential when high densities of cysts are hatched (because of high carbon dioxide production). Cyst density interferes with other abiotic factors that are essential for hatching, such as pH, oxygen and illumination. The density may be as high as 5 g l⫺1 for small volumes (⬍20 litres) but should be decreased to maximum 2 g l⫺1 for larger volumes, to minimise mechanical injury to the nauplii and to avoid suboptimal water conditions. Strong illumination (about 2000 lux at the water surface) is essential, at least during the first few hours after complete hydration, to trigger the start of embryonic development. Although this level of illumination can generally be attained in daytime by using transparent tanks set up outdoors in the shade, it is advisable to keep the hatching tanks indoors and to provide artificial illumination, so as to ensure good standardisation of the hatching process. When hatching large quantities or high densities of cysts, an impressive bacterial load rapidly develops (Dehasque et al. 1993). This is a potential source of pathogens, a competitor for oxygen and a general threat to hatchery hygiene. Reducing bacterial development during hatching will improve the hygienic status of nauplii and may result in better hatching yields. It can be achieved through simple disinfection of the cysts using liquid bleach solution, through decapsulation (see Section 3.5.4) or through the use of recently

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developed cyst and enrichment products that achieve disinfection during the course of the hatching process (Sorgeloos et al. 2001). Attention should be paid to the selection of Artemia cyst batches with good hatching synchrony (less than 7 h between hatching of first and last nauplii) and high hatching efficiency (more than 200,000 nauplii per gram product), as considerable variation has been demonstrated for cysts from different sources, and even among batches from the same strain (Vanhaecke & Sorgeloos 1982).

3.5.3 Harvesting hatched nauplii After hatching and before feeding to fish/crustacean larvae, the nauplii should be separated from the hatching wastes (empty cyst shells, unhatched cysts, debris, micro-organisms and hatching metabolites). Five to ten minutes after switching off the aeration, cyst shells will float and can be removed from the surface, while nauplii and unhatched cysts will concentrate at the bottom (Fig. 3.21). Since nauplii are positively phototactic, their concentration can be improved by shading the upper part of the hatching tank (use of cover) and by focusing light on the bottom part of the conical tank. Nauplii should not be allowed to settle for too long in the bottom of the conical container, as they will quickly suffer from oxygen depletion. First, unhatched cysts and other debris that have accumulated underneath the nauplii are siphoned or drained when necessary (i.e. when using cysts of a lower hatching quality). Then the nauplii are collected on a filter with a fine mesh screen (⬍150 ␮m), which should be submerged at all times to prevent physical damage to the nauplii. They are rinsed thoroughly with water to remove possible contaminants and hatching metabolites such as glycerol. In commercial operations the use of a concentrator/rinser (Fig. 3.22) allows fast harvesting of large volumes of Artemia nauplii and complete removal of debris from the hatching medium. Since instar I nauplii rely solely on their endogenous yolk reserves they should be harvested and fed to the fish or crustacean larvae in their most energy-rich form, i.e. as soon as

Fig. 3.21 Hatching tank after switching off aeration.


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possible after hatching. Farmers often overlook the fact that an Artemia nauplius in its first stage of development cannot take up food and thus consumes its own energy reserves. At the high temperatures that occur during cyst incubation, the freshly hatched Artemia nauplii develop into the second larval stage within a matter of hours. It is important to feed first instar nauplii to the predator rather than starved second instar meta-nauplii which will already have consumed 25–30% of their energy reserves within 24 h after hatching (Fig. 3.23). Moreover, instar II Artemia are less visible as they are transparent. They are also larger and swim more rapidly than first instar larvae. As a result they are less accessible as prey. Furthermore, they contain lower amounts of free amino acids, and their lower individual organic dry weight and energy content will reduce the energy uptake by the predator per hunting effort. All this may be reflected in a reduced growth of the larvae, and increased Artemia cyst usage and cost, as about 20–30% more cysts need to be hatched to feed the same weight of starved meta-nauplii to the predator (Léger et al. 1986).

Fig. 3.22 Concentrator/rinser used for an efficient harvest of large amounts of hatched Artemia.

Fig. 3.23 Energy content and dry weight of instar I, instar II, cold stored nauplii and decapsulated cysts. (Modified from Léger et al. 1987a.)

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3.5.4 Decapsulation Decapsulation is the process whereby the chorion that encysts the Artemia embryo is completely removed by a short exposure to a hypochlorite solution (Bruggeman et al. 1980). The use of decapsulated cysts as a food source is much more limited than the use of Artemia nauplii. Nevertheless, dried decapsulated Artemia cysts have proven to be an appropriate feed for larval rearing of various species such as the freshwater catfish (Clarias gariepinus), the common carp (Cyprinus carpio), and marine shrimp and milkfish larvae (Verreth et al. 1987; Vanhaecke et al. 1990; Stael et al. 1995; Ribeiro & Jones 1998; Sui 2000). Using decapsulated cysts in larval production offers a number of advantages over nauplii and non-decapsulated cysts:

• • •

• •

The daily production of nauplii, a labour-intensive job that requires additional facilities, is avoided. Cyst shells are not introduced into the culture tanks. When hatching normal cysts, the complete separation of Artemia nauplii from their shells is not always possible. Unhatched cysts and empty shells cannot be digested by fish or shrimp larvae and may obstruct the gut when ingested. Nauplii that are hatched out of decapsulated cysts have a higher energy content and individual weight (30–55% depending on strain) than ‘regular’ instar I nauplii from nondecapsulated cysts, because they do not expend energy breaking out of the shell. In some cases, where cysts have a relatively low energy content, the hatchability may be improved by decapsulation, because of the lower energy requirement to break out of a decapsulated cyst (Table 3.10). Decapsulation results in complete disinfection of the cyst material. Cysts with poor hatching quality or even non-hatching cysts can still be used as a food source.

Decapsulated cysts, however, have the disadvantage that they are non-motile and thus less visually attractive to the predator. Moreover, decapsulated cysts dehydrated in brine sink rapidly to the bottom, thus reducing their availability for fish larvae feeding in the water column unless adequate mixing of the culture water is applied. Older penaeid larvae, however, are mainly bottom feeders and do not find this a problem. From the nutritional point of view, the gross biochemical composition of decapsulated cysts is comparable to that of freshly hatched nauplii (García-Ortega et al. 1998) (Tables 3.5–3.7). Table 3.10 Improved hatching characteristics (%) of Artemia from different geographical origins as a result of decapsulation.

Cyst source

Hatchability

Naupliar dry weight

Hatching output

San Francisco Bay, USA Macau, Brazil Great Salt Lake, USA Shark Bay, Australia Chaplin Lake, Canada Bohai Bay, PR China

⫹15 ⫹12 ⫹24 ⫹4 ⫹132 ⫹4

⫹7 ⫹2 ⫺2 ⫹6 ⫹5 ⫹6

⫹23 ⫹14 ⫹21 ⫹10 ⫹144 ⫹10

Modified from Bruggeman et al. (1980).


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In addition, their individual dry weight and energy content is on average 30–40% higher than for instar I nauplii (Fig. 3.23). The decapsulation procedure (Appendix II) involves the hydration of the cysts (as complete removal of the envelope can only be performed when the cysts are spherical), removal of the brown shell in a hypochlorite solution and deactivation of the remaining hypochlorite by washing. These decapsulated cysts can be directly hatched into nauplii, or dehydrated in saturated brine and stored for later hatching or for direct feeding. They can be stored for a few days in the refrigerator at 0–4°C without a reduction in hatchability. If storage for prolonged periods is needed (weeks or few months), the decapsulated cysts can be transferred into a saturated brine solution. During overnight dehydration (with aeration to maintain a homogeneous suspension) cysts have released over 80% of their cellular water, and upon interruption of the aeration, the now coffee-bean-shaped decapsulated cysts settle out. After harvesting of these cysts on a mesh screen they should be stored cooled in fresh brine. Since they lose their hatchability when exposed to UV light it is advisable to store them protected from direct sunlight.

3.5.5 Enrichment The nutritional value of Artemia nauplii is easy to manipulate thanks to their primitive feeding characteristics. After about 8 h posthatch, brine shrimp moult to the second naupliar stage, instar II, and start filtering particles smaller than 25 ␮m irrespective of their nature (Makridis & Vadstein 1999; Gelabert Hernandez 2001). Taking advantage of this non-selective filter feeding, simple methods were developed to incorporate various kinds of products into nauplii before feeding them to predatory larvae (Fig. 3.24). This ‘bioencapsulation’ or ‘enrichment’ is a now very common practice in fish and crustacean hatcheries for enhancing the nutritional value of this live feed or for delivering specific ingredients to cultured larvae. 3.5.5.1 Lipid enrichment In the early 1970s, several authors reported problems with the larviculture of shrimp, crab, prawn, lobster and marine fish larvae when using Artemia sources other than SFB Artemia. After considerable efforts by diverse research groups, these striking differences in culture results could be related to the origin of the Artemia strain used (Bengtson et al. 1991).

Fig. 3.24 Principle of bioencapsulation or enrichment.

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Comparative studies with different strains of Artemia (Kanazawa et al. 1979) and with other zooplankton (Nellen et al. 1981; Watanabe et al. 1983a) revealed differences in levels of specific polyunsaturated fatty acids. Initially, most attention was focused on the presence of EPA (20:5n-3) in Artemia and its importance in the successful production of marine fish and crustacean larvae (Watanabe et al. 1983b; Léger et al. 1985). In the late 1980s and early 1990s, attention shifted to DHA when several authors documented the importance of DHA, more particularly the requirement for high DHA:EPA ratios (Lavens et al. 1995; Kraul 1993; Reitan et al. 1994; Mourente et al. 1993). A myriad of studies has been carried out on the essentiality of long-chain, highly unsaturated fatty acids (HUFAs) in several fish and shrimp species to gain a better understanding of their true requirements, with the ultimate goal of optimising larval feed and/or enrichment products. For the latest developments and insights into larval fish lipid nutrition, the reader is referred to the reviews by Sargent et al. (1999a,b). To provide fish larvae with adequately enriched Artemia, the following points should be considered.

• • •

• • • •

Most marine fish larvae cannot synthesise DHA, EPA or arachidonic acid (ARA, 20:4n-6) from shorter chain precursors and they must be provided preformed in the larval diet (see review by Sargent et al. 1997). ARA (20:4n-6) is the major precursor for eicosanoids in fish, as in mammals (Castell et al. 1994; Bell et al. 1995). EPA is present in large amounts in the cellular membranes of marine fish larvae and is also a precursor of eicosanoids (Sargent et al. 1993). Eicosanoids formed from EPA are less biologically active than ARA-derived eicosanoids, and EPA is important in modulating eicosanoid production by competing for the same enzyme systems that convert ARA to eicosanoids (Sargent 1995). DHA is a major constituent of neural and visual cell membranes and, thus, is essential for a range of physiological processes that are crucial to fast-growing marine fish larvae (Sargent et al. 1999b). The requirements for these essential fatty acids cannot be considered separately. Altering the dietary dose of one of them will influence the ARA:EPA:DHA balance owing to competitive interactions and metabolic conversions (Sargent et al. 1999b). Besides absolute HUFA requirements, the importance of polar lipids and the distribution of HUFA between dietary phospholipids and triacylglycerols (TAG) should not be overlooked (see Section 3.5.5.2). Dietary HUFA requirements seem to be, at least to some extent, species specific.

These facts have various consequences on the usefulness of enriched Artemia as larval food and the modalities of enrichment procedures.

•

It has been proven that optimised DHA levels and high DHA:EPA ratios improve growth, stress resistance and proper pigmentation, especially in marine flatfish (Watanabe et al. 1983b; Kraul 1993; Mourente et al. 1993; Reitan et al. 1994; Copeman et al. 1999). While the DHA:EPA ratio in enriched Artemia rarely exceeds 2:1, marine zooplankton generally have ratios substantionally higher than 1:1, often 4:1 and higher (Shields et al. 1999; see also Chapter 5). Consequently, zooplankton-fed halibut larvae have a much higher DHA level and DHA:EPA ratio than enriched Artemia-fed larvae (McEvoy et al. 1998b).


•

•

107

Although ARA is an essential precursor of eicosanoids, the dietary requirements for ARA are relatively low. Moreover, dietary levels must be carefully chosen, as excessive ARA:EPA ratios tend to exert negative effects on pigmentation (McEvoy et al. 1998a; Estevez et al. 1999; Copeman et al. 1999). Although the exact requirements and effects of ARA in relation to EPA and DHA are not fully understood, they are likely to be species specific. It is difficult to maintain high DHA levels in enriched A. franciscana because of the rapid retroconversion of DHA to EPA, resulting in decreased DHA:EPA ratios as soon as enrichment is interrupted (Navarro et al. 1999). An interesting solution to this problem may stem from the capacity of some Artemia strains to reach high DHA levels during enrichment (Dhert et al. 1993; Velazquez 1996) and to maintain them during subsequent starvation (Evjemo et al. 1997; Han et al. 2000a). Although retroconversion of DHA to EPA may also occur in rotifers, it seems to occur at a lower rate than in Artemia and it is easier to maintain high DHA levels in rotifers.

Although Artemia is often an inferior food source for fish larvae compared with wild zooplankton, the ability to produce any amount of biomass within 24 h, in contrast to zooplankton, and the constant improvement of enrichment products ensure its continued use in marine fish larviculture. In parallel to this relentless unravelling of the biochemical pathways, physiological functions and dietary requirements, numerous enrichment products and procedures were developed, using selected microalgae and/or microencapsulated products, yeast and/or emulsified preparations, self-emulsifying concentrates and/or microparticulate products (reviewed by McEvoy & Sargent 1998). Although initially the composition of enrichment products was often based on empirical trials of variable components, they are increasingly supported by sound insight into the true dietary requirements of marine fish larvae. Nevertheless, we are still far from understanding every species’ requirements and it therefore seems a logical approach to tune the composition of the larval food and associated enrichment products to reflect the natural diets of the larvae; namely, yolk and zooplankton (Sargent et al. 1999b). Currently, the highest enrichment levels are obtained using emulsified concentrates (Table 3.11). This procedure, developed by Léger et al. (1987b), involves the incubation of freshly hatched nauplii in an enrichment emulsion for a period up to 24 h (for detailed procedure, see Appendix III). 3.5.5.2 Phospholipid enrichment Several marine fish and shrimp larvae seem to have a requirement for phospholipids (see review by Coutteau et al. 1996). Geurden et al. (1997) demonstrated that the observed positive effect of phospholipid supplementation was not connected to the role of phospholipids as an additional HUFA source, which suggested that dietary phospholipids were necessary to compensate for a limited ability for de novo biosynthesis by the fish larvae. However, enriching Artemia with traditional products seems to increase the fraction of TAG at the expense of the phospholipid fraction (McEvoy et al. 1996). Although fish larvae appear to have a certain ability to convert fatty acids between phospholipids and TAG, the relatively high proportion of linolenic acid (LNA, 18:3n-3) in the phospholipids of enriched Artemia will hinder fish larvae in assimilating sufficient EPA and DHA (mainly

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Table 3.11 Overview of lipid levels obtained through enrichment of Artemia franciscana by various authors (mg g⫺1 dry weight). Stage Reference

LNA (18:3n-3)

ARA (20:4n-6)

EPA (20:5n-3)

DHA (22:6n-3)

DHA:EPA

Nauplii Dhert et al. (1993) Dhert et al. (1993) Triantaphyllidis et al. (1995) Evjemo et al. (1997) Harel et al. (1999) Estevez et al. (1998) Estevez et al. (1999) Narciso et al. (1999) Han et al. (2000b)

39.8 32.7–40.3 — — — 20.6–21.7 — — 25.6–45.0

1.0 2.8–4.2 — — 2.4–2.7 2.1–14.3 1.0–7.9 — 2.4–4.9

10.7–17.8 4.4–19.5 17.7 17.4 9.5–11.9 10.0–25.1 5.5–33.2 4.1–20.2 29.6–53.2

27.5–45.7 0.8–9.5 25.1 36.6 21.1–39.1 5.2–16.6 3.0–10.0 1.4–11.1 11.2–28.9

1.2–4.9 1.4–0.5 1.4 2.1 1.7–3.9 0.3–1.3 0.1–1.3 0.0–0.7 0.3–0.5

Adults Han (2001) Lim et al. (2001) Dhont (unpubl.)

14.9–26.0 1.8 1.7

13.4 — 3.8

23.1 5.6 9.5

4.6 1.9 4.0

0.2 0.3 0.4

LNA, linolenic acid; ARA, arachidonic acid; EPA, eicosapentaenoic acid; DHA, docosahexaenoic acid; —, not determined.

present in the TAG of the ingested Artemia) to replace the LNA in the ingested phospholipids (Bell et al. 2001). Increasing the polar lipid level in Artemia with emulsions is difficult but, using liposomes, McEvoy et al. (1996) obtained significantly higher levels. However, as these authors point out, this increase in polar lipid may have represented solidified, indigestible dipalmitoyl phosphatidylcholine (DPPC, incorporated to stabilise the liposomes) remaining in the guts of the Artemia and, if so, it is questionable whether this DPPC could be digested by fish larvae. In an effort to determine the most effective molecular carrier of DHA for Artemia, DHAethyl esters were compared with DHA-containing phospholipids. Harel et al. (1998) found significantly higher absorption of DHA using 10% dietary phospholipids compared with 5%, while no further improvement in absorption was obtained at higher phospholipid percentages. In further studies, it was observed that mixtures of phospholipids with DHA sodium salts resulted in maximal absorption of DHA phospholipids in Artemia (Harel et al. 1999) and may be used to increase the polar lipid content in larval live food. 3.5.5.3 Protein enrichment Compared with lipids, much less research has been carried out on the role and requirements for protein in larval nutrition, despite the fact that amino acid catabolism is a major source for energy in fish larvae (Dabrowski 1983) and amino acids are essential for the synthesis of proteins and enzymes (Conceição 1997). Moreover, larvae seem to have higher amino acid requirements than juvenile or adult fish (Dabrowski 1986; Fiogbé & Kestemont 1995). Although it is believed that Artemia contain adequate levels of most amino acids, the fraction of free amino acids is low compared with levels in wild copepods (Tonheim et al. 2000), especially in methionine. Conceição et al. (1997) observed growth retardation in turbot larvae fed on Artemia and suspected it to be related to methionine deficiency.


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Table 3.12 Vitamin levels obtained by enrichment (␮g g⫺1 dry weight).

Vitamin

(Ref.)

Ascorbic acid

(a) (c) (b) (c) (b) (b) (b) (b) (b) (b) (b)

Thiamin Riboflavin Niacin Pantothenic acid Vitamin B6 Biotin Folate Vitamin B12

Unenriched nauplii

24 h enriched nauplii

692

3100 1000–12,000 8.8 20–40 38.0 202 81 6.7 4.0 12.2 3.9

7.5 47.3 187 86 9.0 3.5 18.4 3.5

References: a, Merchie et al. (1995); b, Mæland et al. (2000); c, Olsen et al. (1999).

Dendrinos and Thorpe (1987) cultured Artemia on different types of protein and observed an increase in total protein content when feeding Candida utilis and Saccharomyces cerevisiae. The pattern for levels of essential amino acids was not very clear: some levels clearly increased (leucine, isoleucine, tyrosine, lysine) but others remained unchanged or decreased slightly (phenylalanine, histidine, arginine). Tonheim et al. (2000) demonstrated that methionine levels in nauplii could be boosted 20–30-fold in 16 h by simply adding dissolved methionine in the water. This increase could be doubled (to 60-fold of the unenriched control) when incorporating methionine in liposome droplets. 3.5.5.4 Vitamin enrichment Enrichment of Artemia for 48 h with DHA-Selco (Inve, Belgium) led to increased levels of thiamin, niacin and pantothenic acid, but no changes in the content of AscA, riboflavin, or vitamin B6 or B12 (Mæland et al. 2000). Better results for selected vitamins are obtained using specific enrichment preparations. Tests have been conducted to incorporate extra AscA into Artemia nauplii in a stable and bioavailable form (see Table 3.12). In a 24 h enrichment with self-emulsifying concentrates containing 10–20% ascorbyl palmitate (AscP), levels up to 2.5 mg free AscA g⫺1 dry weight were achieved in brine shrimp nauplii (Merchie et al. 1995). When these vitamin C-enriched Artemia were fed to turbot larvae, no differences in growth or overall survival could be detected compared with those fish fed the non-enriched live food containing 500 ␮g AscA g⫺1 dry weight. However, the larvae given the high AscA treatment showed a better pigmentation rate compared with the control group. Evaluation of the physiological condition through a salinity stress test also revealed an improvement. Cumulative mortalities after challenge with Vibrio anguillarum amounted to 50% for the control versus 40% for the AscA-supplemented fish, with a slower onset of mortality for the AscA-fed fish (Merchie et al. 1995). High levels of ␣-tocopherol can be bioaccumulated and maintained in Artemia nauplii, making this live food delivery system useful for studying dietary requirements as well as antioxidative effects of vitamin E (Huo et al. 1996).

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Vitamin A levels in Artemia nauplii could be raised from 1.3 to 1283 IU g⫺1 dry weight over an 18 h period through the addition of vitamin A palmitate to an egg-yolk-based emulsion (Dedi et al. 1995). However, Rønnestad et al. (1998) found striking differences in vitamin A and carotenoid composition between halibut larvae fed SuperSelco-enriched Artemia and a species of copepod (Temora). Although the pigmentation status was not systematically recorded, the authors observed a higher incidence of malpigmentation with Artemia-fed larvae. 3.5.5.5 Enrichment with prophylactics Although the use of antibiotics in larviculture is rightfully questioned, disapproved of or even banned, it may be of interest to mention that techniques have been developed for oral biomedication, rather than administration via the culture water (‘bath treatments’). Doses ranging from 20 to 100 ppm sulfadrugs can be incorporated in sea bass and turbot larvae tissue, respectively, within less then 4 h by feeding them with specifically enriched Artemia (Chair et al. 1996; Gapasin et al. 1996). 3.5.5.6 Enrichment with other products The effectiveness of Artemia nauplii as a dietary carrier system could be tested for various other nutritional components, e.g. liposoluble products administered via an emulsion, watersoluble compounds via liposomes (Hontoria et al. 1994) and microcapsule delivery (Sakamoto et al. 1982). However, for each nutrient, the usefulness of the Artemia bioencapsulation method remains to be verified by chemical analysis.

3.5.6 Cold storage 3.5.6.1 Survival at low temperatures Moulting of the Artemia nauplii to the second instar stage can be delayed and their energy metabolism greatly reduced by storage of the freshly hatched nauplii at a temperature below 10°C at densities of up to 8 million/l (Léger et al. 1983). Nauplii can be stored for more than 24 h without significant mortalities and a reduction in energy of less than 5%. Only slight aeration is needed to prevent the nauplii from accumulating at the bottom of the tank where they might suffocate. Nauplii stored at 20 million/l showed good survival (⬎70%) even after 72 h when kept at 12°C with a slight injection of pure oxygen (Anbaya Almalul 2000). Evjemo et al. (2001) kept nauplii after enrichment at moderate densities (⬍100,000 l⫺1) and recorded survival above 70% for temperatures between 8 and 19°C. At 5°C and above 19°C survival decreased rapidly after 48 h. 3.5.6.2 Maintenance of nutritional value Initially, it was generally accepted that nauplii stored at low temperatures maintained their biochemical composition (Fig. 3.23), especially the lipid content (Léger et al. 1983). More detailed analysis revealed that protein is fairly well conserved even after 96 h (Evjemo et al. 2001), but fatty acid levels, mainly DHA, decrease significantly. Estevez et al. (1998) demonstrated that up to 70% of the DHA obtained through enrichment was catabolised, while


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losses of EPA were more moderate and depended on the type of enrichment received. The fact that Artemia retroconvert DHA to EPA (Navarro et al. 1999) may also account for the reduced losses of EPA compared to DHA. An interesting observation was made by Evjemo et al. (1997): they confirmed that, after enrichment, A. franciscana catabolised DHA at a rate that increased with temperature but demonstrated that, under similar circumstances, DHA levels in A. sinica remained at almost constant high levels after enrichment and at temperatures ranging from 6 to 22°C. 3.5.6.3 Other advantages Cold storage enables the farmer to reduce hatching efforts (less frequent hatching and harvesting, fewer tanks, larger volumes). Using cold storage also allows for more frequent and automated distribution of nauplii to larvae. This appears to be beneficial for fish and shrimp larvae as food retention times in the larviculture tanks can be reduced and hence growth of the Artemia in the culture tank can be minimised. For example, applying one or two feedings per day, farmers often experienced juvenile Artemia in their larviculture tanks. With poor hunters such as the larvae of turbot, using cold-stored, less active Artemia as live prey resulted in a much more efficient food uptake (Léger et al. 1986).

3.5.7 Use of juvenile and adult Artemia Besides nutritional and energetic advantages, the use of Artemia biomass for feeding postlarval shrimp also results in improved economics, as expenses for cysts and weaning diets can be reduced. Dhert et al. (1993) developed a simple culture system for juvenile and adult Artemia as food for postlarval (PLa) Penaeus monodon. The growth performance of shrimp reared from PLa-4 to PLa-25 on juvenile Artemia live prey is identical to the growth obtained when feeding newly hatched Artemia, but the PLa-25 reared with juvenile brine shrimp display significantly better resistance in salinity stress tests, i.e. the stress sensitivity index dropped from 138 with freshly hatched nauplii to 36 when feeding juvenile Artemia. Similarly, Olsen et al. (1999) proved that halibut larvae fed with gradually increasing sizes of nauplii showed the same satisfactory growth and survival as larvae fed short-term enriched nauplii, but the quality of the halibut larvae (proper pigmentation, eye migration and lack of deformities) was significantly higher. Lim et al. (2001) developed a pilot-scale culture unit for ongrown Artemia for use in ornamental fish in Singapore and proved it to be cost-effective, with a payback period of less than 18 months. It offers local fish breeders a cheaper, less labour-intensive and nutritionally suitable alternative to the traditional Moina culture. Although the fresh, live form has the highest nutritive value, harvested Artemia can also be frozen, freeze-dried or acid-preserved for later use (Abelin et al. 1991; Naessens et al. 1995), or made into flakes or other forms of formulated feed (Sui 2000). Artemia biomass is apparently a good food for the maturation of several species of penaeid shrimp. Recent culture tests in Ecuador and the USA have shown that polychaetes, which have been identified as a critical fresh-food component in the maturation diet of Litopenaeus vannamei (Bray & Lawrence 1991), can be successfully replaced by frozen Artemia biomass (Naessens et al. 1997).

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Chapter 4

Production, Harvest and Processing of Artemia from Natural Lakes Gilbert Van Stappen

4.1 Introduction Highly saline lakes with natural Artemia populations can vary in size from a few hectares to large inland lakes like Great Salt Lake (GSL, Utah, USA; Fig. 4.1) and Lake Urmiah (Iran), both 4000–6000 km2 in size. In these inland lakes, population densities are usually low and fluctuate mainly as a function of food availability, temperature and salinity. The size and lack of suitable infrastructure make management of such lakes very difficult, restricting the main activity to extensive harvesting of Artemia biomass and/or cysts. Until recently, this harvesting was performed at most sites without too much concern for sustainable exploitation or the carrying capacity of the available Artemia population. Only the sharp decrease in the harvests in GSL at the end of the 1990s has urged both the Artemia industry and wildlife authorities, at GSL and other sites, to perform population assessment studies. Since the low harvests at GSL in the late 1990s (see Chapter 3), several population monitoring and modelling studies have been realised, and research-based management of the

Fig. 4.1 Harvesting of Artemia cysts from Great Salt Lake, Utah, USA.

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lake started in 1996. A co-operative research programme was launched by the Utah State Department of Natural Resources, the Utah Division of Wildlife Resources, the Utah State University and the United States Geological Survey. This research programme focuses on the study of the lake’s ecosystem and the sustainable exploitation of its brine shrimp resources, by studying its population dynamics and formulating quantitative population predictions. It includes the study of the abiotic parameters and the phytoplankton population, and links are made with the population’s age structure and reproduction characteristics (Stephens 2000; Belovsky et al. 2000). Through the identification of a minimal viable population size, the concentration of cysts found is used as a criterion to determine the harvestable amounts, and to open and close the harvesting season. Harvesting cysts is sometimes undertaken by collecting cysts from the shore, when boats are not available. However, best practice is to collect the cyst accumulations from the water surface, as this product contains fewer impurities and has a higher viability. On an industrial scale (e.g. at GSL), floating cyst accumulations are spotted by aircraft, then concentrated (Fig. 4.1) and pumped into vessels.

4.2 Pond Production of Artemia Cysts and Biomass 4.2.1 Permanent solar salt operations Mechanised salt production operations consist of several interconnected evaporation ponds and crystallisers. In these salt operations, pond size can vary from a few to several hundred hectares, each with depths of 0.5–1.5 m (Fig. 4.2). Seawater is pumped into the first pond and flows by gravity through consecutive evaporation ponds. While passing through the pond system, salinity gradually increases as a result of evaporation, and salts with low solubility precipitate as carbonates and sulfates. Once the seawater has evaporated to about one-tenth of its original volume (about 260 g l⫺1), this ‘mother brine’ is pumped into the crystallisers, where NaCl precipitates. Before all NaCl has crystallised, the mother liquor, now called bittern, must be drained off. Otherwise, the NaCl deposits will be contaminated with MgCl2, MgSO4 and KCl, which start precipitating at this elevated salinity. The technique of salt production thus involves fractional crystallisation of the salts in different ponds. To ensure that the different salts precipitate in the correct pond, salinity in each pond is strictly controlled and during most of the year kept at a constant level. Within such salt production facilities, brine shrimp are mainly found in ponds at intermediate salinity levels. As Artemia have no defence mechanisms against predators, the lowest salinity at which animals are found is also the upper salinity tolerance level of possible predators (i.e. 80–140 g l⫺1). Above 250 g l⫺1, animal density decreases. Although live animals can be found at higher salinity, the need for increased osmoregulatory activity, requiring higher energy inputs, negatively influences growth and reproduction, eventually leading to starvation and death. Cysts are produced in ponds having intermediate and high salinity (80–250 g l⫺1). The population density depends on food availability, temperature and salinity, but generally numbers of animals and thus cyst yields are low. Moreover, the stable conditions prevailing in the ponds of these saltworks often result in stable populations in which the ovoviviparous reproduction mode dominates.

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Fourth evaporator

Crystallisers

Third evaporator

Second evaporator

First evaporator

Pump1

Dike

Pump2 Large salt operation (Port Saïd, Egypt) : First evaporator = 600ha; Salinity : 35ppt – 80ppt Second evaporator = 100ha; Salinity : 80ppt – 140ppt Third evaporator = 75ha; Salinity : 140ppt – 180ppt Fourth evaporator = 70ha; Salinity : 180ppt – 260ppt Crystallisers = 125ha; Salinity > 260ppt Fig. 4.2 Schematic outline of a typical permanent saltwork, Port Saïd, Egypt.

4.2.2 Seasonal units In the tropics and subtropics, small-scale artisanal saltworks (saltstreets) operate during the dry season, when evaporation of water from the ponds exceeds precipitation. The ponds are just a few hundred square metres in size and are 0.1–0.6 m deep (Fig. 4.3). Salt production is abandoned during the rainy season, when evaporation ponds are often turned into fish/ shrimp ponds. Although salt production in these saltstreets is based on the same chemical and biological principles as in the large saltfarms, production methods differ (Vu Do Quynh & Nguyen Ngoc Lam 1987). At the beginning of the production season, all ponds are filled with seawater. For the remainder of the season water is kept in each pond until the salinity reaches a predetermined level and is then allowed to flow into the next pond, holding water of a higher salinity. The salinity in the different ponds is not kept constant as in continually operated saltworks. Sometimes, to increase evaporation further, ponds are not refilled immediately but left dry for 1 or 2 days. During that time the bottom heats up, which further enhances evaporation. Once the salinity reaches 260 g l⫺1, water is pumped to the crystallisers, where NaCl precipitates. Artemia thrive in ponds where salinity is high enough to exclude predators (between 80 and 140 g l⫺1). As seasonal systems are often small, they are fairly easy to manipulate. Hence, higher food levels and thus higher animal densities can be maintained. In addition, the shallow ponds, high algal density, use of organic manure and discontinuous pumping cause abiotic factors such as temperature, oxygen and salinity to fluctuate, creating an unstable environ-


Storage and evaporation area

Evaporation pond (first)

Evaporation pond (second)

Crystalliser



Crystalliser





Canal Storage and evaporation area

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Crystalliser

Crystalliser

Small salt operation (Vinh Tien, Vietnam) Storage and evaporation area = 0.4ha – 1ha; Salinity: 10ppt – 50ppt Evaporation pond (first series) = 0.2ha – 0.5ha; Salinity: 40ppt – 120ppt Evaporation pond (second series) = 0.2ha – 0.5ha; Salinity: 80ppt – 160ppt Crystallisers = 0.2ha – 0.5ha; 120ppt – all water evaporated. Fig. 4.3 Layout of a typical artisanal saltfarm, Vietnam.

ment. This, together with the fact that population cycles are seasonally interrupted, seems to favour oviparous reproduction. This culture system is referred to as semi-intensive (Tackaert & Sorgeloos 1991) and static (Vu Do Quynh & Nguyen Ngoc Lam 1987), as ponds are managed intensively (i.e. inoculation of selected strains, manipulation of primary and secondary production, predator control) and are not interconnected. Once stocked, ponds are managed as separate units, and green water is pumped from a common fertilisation pond and, if necessary, mixed with brine to maintain high salinity levels in the culture ponds (Baert et al. 1996). As the Artemia population is maintained throughout the culture season, and animals are introduced only once per season, this system can be described as ‘one-cycle’ (Baert et al. 1997). More stable cyst yields can be obtained by the multicycle system, where ponds are drained and restocked several times per season (Baert et al. 1997). Food levels in this system, where only the first generation is allowed to develop and release cysts, allow higher numbers of reproducing adults and larger brood sizes. In the one-cycle system, in contrast, the food levels can only be increased to a limited extent, as the pumping rates (limited by salinity constraints) and the use of organic manure are already maximised.

4.2.3 Site selection Integrating Artemia production in an operational solar saltwork or shrimp/fish farm improves the cost-effectiveness. Ponds can be constructed close to evaporation ponds with

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the required salinity, or low-salinity ponds already existing in the salt operation can be modified (Tackaert & Sorgeloos 1993). Several criteria have to be taken into account when selecting a site for Artemia production. 4.2.3.1 Climatology The presence of sufficient amounts of highly saline water is imperative. Therefore, Artemia culture is mostly found in areas where evaporation rates are higher than precipitation rates during extended periods of the year (e.g. a dry season of more than 4 months in the tropical–subtropical belt). Evaporation rates depend on temperature, wind velocity and relative humidity. The presence of solar saltfarms in the neighbourhood is a clear indication that Artemia pond culture is possible during at least part of the year. As temperature also influences population dynamics directly, this climatological factor should receive special attention. Temperatures that are too low will result in slow growth and reproduction, whereas high temperatures can be lethal. Optimal culture temperatures are strain-dependent. 4.2.3.2 Topography Flat land allows easy construction of ponds with regular shapes. A gradual slope can eventually facilitate gravity flow in the pond complex. The choice between dug-out (entirely excavated) and level ponds (bottom at practically the same depth as the surrounding land and water retained by dikes or levees) will depend on the type of ponds already in use. Locating the Artemia ponds lower than all other ponds is good practice, as the water flow into the ponds is much higher than the outflow (usually ponds are only drained at the end of the culture season). Making use of gravity or tidal currents to fill the ponds, even if only partially, will reduce pumping costs. 4.2.3.3 Soil conditions Because long evaporation times are needed to produce high-salinity water, leakage and/or infiltration rates should be minimal. Heavy clay soils with minimal contents of sand are the ideal substrate. An additional problem may be the presence of acid sulfate soils, often found in mangrove or swamp areas. When exposed to air such soils form sulfuric acid, resulting in a pH reduction in the water, which may have a negative effect on the phytoplankton population, the most important food source for Artemia. The presence of high levels of organic material in the pond bottom may also cause problems; especially when used for dike construction, such earth tends to shrink, thus lowering the dike height considerably. Moreover, problems can arise with oxygen depletion at the pond bottom, where organic material is decomposing, but these problems will gradually disappear when the same pond is used for several years.

4.2.4 Pond adaptation In large salt operations, adaptation of the existing ponds is normally not possible and generally not needed. The only adaptation needed is the installation of screens to reduce


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the number of predators entering the evaporators (see below). This is especially important in regions where predators are found at high salinity (e.g. the Cyprinodont fish Aphanius). In artisanal saltworks, ponds are often very shallow, sometimes resulting in water temperatures that are too high for Artemia (⬎40°C) and promoting phytobenthos rather than the required phytoplankton. Integration of Artemia production with such systems requires that ponds should be deepened, dikes heightened and screens installed to prevent predators from entering the culture ponds. Under windy conditions, high wave action will enhance evaporation. However, to reduce foam formation (in which cysts get trapped) at the downwind side of the pond, wave breakers should be installed (Fig. 4.4), which will also act as cyst barriers and facilitate harvesting. 4.2.4.1 Deepening the ponds In regions with high air temperatures, deepening the ponds is crucial. Depths of 40–50 cm are to be recommended. High water levels are needed not only to prevent lethal water temperatures but, at the same time, to reduce growth of benthic algae. Ponds are usually deepened by digging a peripheral ditch and using the excavated earth to heighten the dikes. 4.2.4.2 Dike construction To prevent leakage, newly constructed dikes need to be well compacted (Kungvankij & Chua 1986). When heightening old dikes, leaks will occur most frequently at the interface of old and new soil. To prevent such leaks from occurring, the old dike should first be wetted and ripped before new soil is added. Dikes are often inhabited by burrowing crabs that cause leakage and destabilisation. Filling nests with CaO and clay will reduce leaks caused by these crabs. Dike slopes should have a 1:1 ratio (height:width) to prevent excessive erosion.

Fig. 4.4 Floating bamboo poles used as wave breakers for the harvesting of Artemia cysts.

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4.2.4.3 Screening Intake waters should be screened to prevent predators from entering the culture ponds. Two types of filter can be used: filter bags (in plastic mosquito-screen, polyurethane or nylon) or stainless-steel screens. As intake water is often heavily loaded with particles, sequential screening is recommended, where an array of different screens, each with a smaller mesh size than the previous one, is used. Both stainless-steel screens and filter bags should be cleaned regularly. For small ponds, the use of filter boxes can be considered. In such boxes a stainlesssteel, welded-wedge filter is installed under an adjustable angle. Water is lifted by a pump into an overhead compartment from where the water is drained over the filter screen. When using such filters, even small competitors such as copepods can be removed (up to 90%). Results are especially good when Artemia culture periods are relatively short (6–8 weeks). However, the high cost of these units restricts their use to regions where highly saline water is not abundant and/or where the presence of (small) predators seriously hampers Artemia culture.

4.2.5 Preparation of ponds for Artemia cultivation 4.2.5.1 Liming Normally, ponds used to culture Artemia do not need liming. The highly saline water often has a hardness of more than 50 mg CaCO3 l⫺1 (owing to the presence of carbonates). Liming ponds with such hardness will not further improve yields. Liming should be considered when culture water has a pH of less than 7.5 and stimulating an algal bloom proves difficult. The liming substances most often used in aquaculture are agricultural lime, CaO, quicklime and Ca(OH)2 or hydrated lime (Boyd 1990). Using CaO and Ca(OH)2 will result in a quick pH rise to about 10, which will kill possible pathogens and predators. CaO and Ca(OH)2 are therefore often used to disinfect the pond bottom. After 2–3 days, the pH drops to 7.5, after which normal mineralisation takes place. The lime requirement is highest for clay bottoms and acid bottoms, and when the pond water has low concentrations of Ca2⫹ and Mg2⫹. Whereas drying can be beneficial for most soils, this is not true for acid sulfate soils, often found in mangrove areas. When exposed to the air, the pyrite in these soils oxidises to form sulfuric acid. Liming of these soils requires large amounts of lime. A simpler method to reduce acidity is flushing the ponds repeatedly after oxidation (exposing the soil to the air) but, in general, culturing brine shrimp in regions with acid sulfate soils should be avoided. 4.2.5.2 Predator control Removal of predators in large salt operations is very difficult. Careful screening of intake water (see Section 4.2.4) and restricting the culture of Artemia to high-salinity ponds are of the utmost importance. If large numbers of predators are found in the culture ponds, manual removal (i.e. by trawl nets) and killing fish/shrimp accumulating at the gates using a mixture of urea and bleach (0.01–0.015 kg urea m⫺3 and 0.007–0.01 kg 75% bleaching powder m⫺3) will be necessary. In small artisanal systems, ponds should initially be filled only to a level of 10–15 cm, to ensure maximum evaporation. Thus, a salinity lethal to predators will rapidly be obtained.


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Screening of the intake water will further reduce the number of predators in the pond. As ponds often cannot be drained completely, fish, crabs and shrimp left in puddles may be killed using rotenone (0.05–2.0 mg l⫺1), tea-seed cake (15 mg 1⫺1), a combination of urea and hypochlorite (5 mg l⫺1 urea and 24 h later 5 mg l⫺1 hypochlorite), CaO or derris root (1 kg 150 m⫺3). Dipterex (2 mg l⫺1) will kill smaller predators such as copepods and is also very toxic to shrimp. The degradation rate of rotenone, chlorine and CaO to non-toxic forms is fairly rapid (24–48 h), whereas if tea-seed cake or dipterex is used, ponds should be flushed prior to the stocking of animals. 4.2.5.3 Fertilisation Fertilisers are added to the culture ponds to increase primary production. Numerous factors influence the chemistry of the fertilisers, such as the ion composition of seawater, pH and pond bottom, and algal growth (temperature, salinity, sunlight) and species composition [nitrogen:phosphorus (N:P) ratio, selective grazing pressure] (Tackaert & Sorgeloos 1991). The inorganic nutrients carbon, nitrogen and phosphorus, used by photosynthesising algae, enter the system via the photo-autotrophic pathway, whereas organic nutrients are processed through the heterotrophic pathways of heterotrophic bacteria, or are consumed directly by the Artemia. As some algae are better suited as food for Artemia than others, a high N:P ratio is usually recommended. An N:P ratio of 10 is desirable for the growth of green algae (Tetraselmis, Dunaliella) and diatoms (Chaetoceros, Navicula, Nitschzia). However, as phosphorus has poor solubility in saltwater and is quickly absorbed at the pond bottom, N:P ratios of 3–5 may be more appropriate. If too much phosphorus is added, especially at high temperatures (⬎28°C) and in the case of low turbidity (bottom visible), growth of benthic algae is promoted. Likewise, high phosphorus concentrations combined with low salinity seem to induce the growth of filamentous blue–green algae (e.g. Lyngbya, Oscillatoria). Both algae are often too large in size for ingestion by Artemia. Besides the N:P ratio, temperature, salinity, light intensity and pumping rates (input of new nutrients and carbon dioxide) also play an important role. At lower salinity and higher light intensities, high N:P ratios stimulate green algae more than diatoms. Some green algae (Nannochloropsis, Chlamydomonas) are poorly digested by Artemia. Finally, manipulation of algal populations also depends on the composition of the local algal community. The most dominant algae in the intake water will often also be the dominant ones after fertilisation. In large salt operations, costs may limit the use of fertilisers. Regular pumping is often more effective in controlling the Artemia standing crop. When pumping, new nutrients and carbon dioxide enter the culture ponds that stimulate algal growth. In areas where intake water is nutrient rich (turbidity readings, i.e. Secchi depth, ⬍40 cm), no additional fertilisation should be used. As pumping influences the retention time of the nutrients in the ponds (i.e. at high pumping rates algae will not have time to take up nutrients), fertilisation should be combined with lower pumping rates in systems with short retention times. Ideally, algal turbidity readings should be kept between 20 and 40 cm in the Artemia culture ponds by regular water intake from the fertilisation ponds. Turbidity readings of less than 20 cm may result in oxygen stress at night, especially when temperatures are high.

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4.2.5.4 Inorganic fertilisers Nitrogen fertilisation As the nitrogen influx to the system depends solely on biochemical processes (degradation of organic matter by bacteria) and the nutrient level in the intake water, nitrogen often limits algal growth. The use of nitrogen fertilisers is therefore widespread. Among the most common inorganic nitrogen fertilisers are ammonium salts [e.g. (NH4)2SO4], nitrates [e.g. Ca(NO3)2], amides and urea. The need for nitrogen fertilisation varies considerably and should be determined experimentally at each site. Usually, adding between 1 mg l⫺1 (eutrophic intake water) and 10 mg l⫺1 (oligotrophic water) nitrogen will induce an algal bloom. It is best to fertilise only the low-salinity ponds in a flow-through system. Initiating an algal bloom in high-salinity ponds is difficult and can take more than 1 month. The algae and organic matter created in the low-salinity ponds are drained to the high-salinity ponds, where they are available as food. The use of inorganic fertilisers in Artemia culture ponds is not recommended (except before introducing the nauplii), as algal densities here are limited not by the nutrient concentrations but rather by the grazing pressure imposed by the brine shrimp. Phosphorus fertilisation As with nitrogen, phosphorus enters the culture ponds with the intake water in the form of organic material that only becomes available through bacterial decomposition. Phosphorus is also found in the soil, where it is bound in the form of AlPO4 ⭈2H2O or FePO4 ⭈2H2O. Most phosphorus fertilisers precipitate, especially in saltwater ponds, i.e. by reacting with Ca2⫹. Phosphorus is also quickly absorbed at the pond bottom. In cases where the use of phosphorus fertilisers is desirable, fertilisers with a small grain size, which dissolve easily in water, should be selected. Predissolving the fertiliser in freshwater will improve its availability. 4.2.5.5 Organic fertilisers Organic fertilisers present some distinctive advantages as they contain, besides nitrogen and phosphorus, other minerals beneficial for phytoplankton growth. Fertiliser particles coated with bacteria enhance the microflora and can be used directly as food by Artemia. Finally, the use of organic fertilisers implies the recycling of a waste product. However, the use of organic fertilisers is difficult to standardise in view of their variable composition. Their use results in a higher biological oxygen demand (BOD) in the pond and they may carry pathogens. The organic fertilisers most often used in aquaculture are chicken, quail and duck manure. Cow, pig and goat dung have also been used, but seem to stimulate phytobenthos (Tackaert & Sorgeloos 1991). 4.2.5.6 Combination of organic and inorganic fertilisers A common practice is to use a combination of inorganic and organic fertilisers. Inorganic fertilisers stimulate algal growth and mineralisation of the organic fertiliser by lowering the carbon:nitrogen (C:N) ratio. However, organic fertilisers are used directly as food for the Artemia and, via slow release of nutrients, especially phosphorus, further stimulate algal


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growth. Normally, inorganic fertilisers are added to the fertilisation ponds or canals, while manure can be added directly to the Artemia culture ponds or to the fertilisation ponds.

4.2.6 Artemia inoculation 4.2.6.1 Artemia strain selection While Artemia introduction frequently ensures social and economic benefits, particularly in developing countries, it also bears certain risks (Beardmore 1987). Competition with local (or nearby) strains or species of Artemia may occur (Geddes & Williams 1987), which may lead to the extinction of some genotypes or, at worst, of one or both competitors. Competition experiments suggest that Artemia franciscana may outcompete others (Browne 1980; Browne & Halanych 1989). The effect of one introduction will not remain local but may have consequences over large areas: many saltworks in north-east Brazil are now populated with Artemia since the human intervention in Macau in 1977, followed by dispersion by wind and local waterbirds over a distance of more than 1000 km. The resolution put forward at the 2nd International Symposium on Artemia, in Antwerp, Belgium, in September 1985, is still valid: ‘… the 2nd International Symposium resolves that all possible measures be taken to ensure that the genetic resources of natural Artemia populations are conserved; such measures include the establishment of gene banks (cysts), close monitoring of inoculation policies, and where possible the use of indigenous Artemia for inoculating Artemia-free waters’ (Beardmore 1987). When the idea is thus to replace a poorly performing strain, in terms of its limited effect on algae removal in the salt production process, or its unsuitable characteristics for use in aquaculture (e.g. large cysts, particular diapause or hatching characteristics), all possible efforts should be made to collect, process and store a sufficient quantity of good quality cysts of the autochthonous gene pool that have a proven high hatch rate. Strain selection can be based on the literature data for growth, reproductive characteristics and especially temperature/salinity tolerance. In summary, a strain exhibiting maximal growth and having a high reproductive output at the prevailing temperature/salinity regime in the ponds should be selected. 4.2.6.2 Inoculation procedures Standard hatching procedures (described in Section 3.5.2) should be followed wherever possible. It is essential to harvest the nauplii in the first instar stage. Older instar stages will not survive the salinity shock when transferred from the hatching vessel (20–35 g l⫺1) to the culture ponds (80 g l⫺1 upwards). Stocking density is determined by the nutrient level and temperature found in the culture ponds. For large salt operations, depending on the size of the ponds, a stocking density of 5–10 nauplii l⫺1 should be considered. However, in large operations practical considerations, such as facilities to hatch out the required amount of cysts, may further limit the stocking density. Animals should be stocked as early as possible in the brine circuit where no predators are found. In small pond systems, the initial stocking density can be as high as 100 nauplii l⫺1 in ponds with turbidity between 15 and 25 cm. However, at such high stocking densities oxygen

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may become limiting, especially when water temperatures are high. When food for the nauplii is less abundant (turbidity readings of more than 25 cm), stocking density should be decreased to 50–70 nauplii l⫺1. Stocking at high density is thought to stimulate oviparous reproduction, whereas stocking at lower density may increase the proportion of ovoviviparity. However, since more food is available per individual, animals grow more rapidly and females have larger broods. As a result, final cyst yields do not necessarily decrease when lower stocking densities are applied.

4.2.7 Monitoring and managing the culture system Very regular monitoring of the ponds is necessary to allow correct management. The type of sampling programme depends largely on the goals. If production is the main objective, only those variables necessary to provide essential decision-making information should be monitored (temperature, salinity, turbidity, number of females and brood size). More extensive sampling programmes are required when research programmes are carried out in the culture ponds, allowing for relative estimates of population numbers, at least (Eliot 1977; Krebs 1989). The most important rule when collecting data is standardisation. This includes selection and marking of fixed sampling stations at every site, the use of fixed well-maintained and operational equipment, appropriate techniques when measuring parameters or when analysing samples, and keeping accurate data records. 4.2.7.1 Monitoring the Artemia population Samples should be collected at fixed sampling stations located in as many different strata as possible, by filtering, for instance, 5–10 litres of water over a 100 ␮m sieve, or by dragging a conical net over a certain distance through the water. Drags can be horizontal or vertical. However, the mesh size and diameter of the sampling net depend on the volume of water sampled, which in turn depends on the population density in the pond. To prevent clogging, only the distal part of the net has a small mesh size (100 ␮m), while the remainder of the net can have a mesh size of 300–500 ␮m. Samples are fixed with formalin and carefully examined, dividing animals into age or size classes, such as nauplii (no thoracopods), juveniles (developing thoracopods clearly visible) and adults (sexual differentiation apparent). The scores for each life stage of all samples taken in one pond are summed and plotted in time. Although such estimates do not give the exact number of animals per litre, they correctly reflect the variations in abundance and allow for adaptation of the management procedures. The reproductive status of the females can also be used as an indicator for the health status of the Artemia population. Large broods and short interbrood intervals show that pond conditions are good. A quick way to estimate standing crop is to use sample volume as an estimate. The sample is fixed with lugol or formalin, then biomass is transferred to a measuring cylinder, where it is allowed to settle for 10 min, after which the volume is read. Using dry weight as an estimator is only possible if samples can be cleaned properly. Wet weight should not be used, as it is very imprecise and inaccurate.


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4.2.7.2 Abiotic parameters influencing Artemia populations Temperature In deeper ponds, the water may be stratified and temperatures at the surface and bottom may differ considerably. In extreme situations this can lead to lethally high temperatures and low oxygen concentrations at the pond bottom, especially in situations of salinity stratification (i.e. a greenhouse effect resulting from the low-salinity top layer). Such conditions, indicated by large clouds of deep red Artemia at the water surface, have a negative influence on growth and survival. Regular pumping or raking of the pond bottom will prevent stratification. Salinity Salinity is an important factor in determining the success of Artemia populations (Triantaphyllidis et al. 1995). As mentioned previously, the upper salinity tolerance level of predators (e.g. fish, Corixidae) determines the threshold beyond which reasonable numbers of Artemia can be found. Water at greater salinity than 250 g l⫺1 is toxic for Artemia. Under field conditions, oviparous reproduction is often found at high salinity, probably as a result of oxygen and food stress. Oxygen Often oxygen levels will be higher at the surface than at the bottom, especially when ponds are stratified. Oxygen levels also exhibit daily cycles. Concentrations are lowest at dawn (algal respiration) and highest in the afternoon (algal photosynthesis). When exposed to oxygen stress, Artemia turn red, swim slowly and start surfacing, and growth is retarded. Extra pumping, lowering the algal concentration or circulating the water in the pond will increase oxygen levels. pH In their natural habitat Artemia are mostly found in a pH range between 7.8 and 8.2, which is often given as the optimal range. However, effects of pH on growth and reproduction have not been studied so far. Algal blooms can affect the pH (consumption of carbon dioxide), but as seawater is usually well buffered, problems with pH are rare, except in areas with acid sulfate soils. Water depth Depth is best measured using calibrated sticks placed in the pond. The water depth influences several of the other measurements (e.g. temperature, oxygen) and therefore should also be recorded. Furthermore, fluctuations in pond depth give information on pumping rates, evaporation, precipitation and leakage. 4.2.7.3 Biotic factors influencing Artemia populations Algae The easiest way to estimate algal abundance is by the measurement of turbidity with a Secchi disk. Turbidity fluctuates during the day, and wind (concentration of algae in the downwind corners) as well as suspended solids (e.g. clay) can affect turbidity readings. Turbidity readings between 25 and 35 cm are optimal. At lower turbidity levels, extra pumping of nutrientrich water is needed. At higher turbidity, the risk of oxygen depletion at dawn increases.

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Fig. 4.5 Raking the Artemia pond to remove the benthic algae.

If time and equipment are available, the algal density can be estimated by analysis of the chlorophyll concentration, if combined with a proper sampling programme. The colour of the water can give useful indications concerning the type of organisms present in the culture ponds. If problems are encountered, more thorough analysis of the algal samples is recommended. Algal composition not only influences growth and reproduction of the Artemia, but also has an effect on the nutritional value of the biomass and the cysts (e.g. fatty acid composition). Algal numbers can be increased through fertilisation. A problem often encountered in Artemia ponds is the presence of benthic and/or filamentous algae. Both types of alga are unsuitable as food for the Artemia. Development of these algae can be prevented by keeping pond water turbid and deepening the ponds. In Vietnam, pond bottoms are raked daily to remove benthic algae (Fig. 4.5). Moreover, raking brings detritus (extra food for Artemia) as well as inorganic nutrients back into suspension. Predators and competitors Possible predators of Artemia include fish (Aphanius, Tilapia), various species of insect (Corixidae) and some copepods. Rotifers and ciliates (Fabrea) are possible food competitors. Careful screening of the intake waters and increasing salinity will keep their numbers within acceptable limits. As wading waterbirds (e.g. avocets, Recurvirostra, and herons, Ardea) also take adult Artemia, bird scarers and wires stretched above the water near shallow places may be used to control these predators.

4.3 Artemia Harvesting and Processing Techniques 4.3.1 Harvesting techniques Adult Artemia biomass can be collected from large, shallow ponds with conical nets mounted in front of a motorboat or pulled by people (Figs 4.6, 4.7). In small ponds,


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Fig. 4.6 Raft with conical net used for Artemia biomass harvesting.

Fig. 4.7 Small net used for Artemia biomass harvesting.

dip-nets can be used. Alternatively, nets can be installed (temporarily) at the pond outlet and biomass is then collected automatically when water flows (by pumping or gravity) to the next pond (Fig. 4.8). After collection, biomass should be prepared for transport and further use or treatment.

4.3.2 Processing techniques Artemia biomass should be used immediately after harvesting as live food, or should be frozen or dried. Harvested biomass can be stored temporarily in nets installed in the pond (Fig. 4.9). Live transportation for marketing as a live product (⬎90% survival after 24 h) is undertaken using similar techniques to those for the transportation of live fish and shrimp

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Fig. 4.8 Installation of filter nets at the sluice gate for Artemia biomass harvesting in solar saltworks.

Fig. 4.9 Storage net for Artemia biomass harvested from seasonal salt ponds integrated for Artemia production.


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larvae. They are transported in plastic bags, filled to one-third of their capacity with seawater, and stocked with Artemia at a density of 100 g live wet weight biomass per litre. The rest of the bag is inflated with oxygen and closed off with a rubber band. The bags are packed in polystyrene boxes filled with ice. Artemia biomass can be frozen for subsequent use as a food source in fish/shrimp hatcheries or for the pet market. Alternatively, biomass can be dried and used as an ingredient for larval feeds (flakes or particulate diets). Since Artemia is rich in proteolytic enzymes it is essential to process the biomass alive. Freezing should be done as quickly as possible (thin layers, low temperature), as slow freezing will result in proteolytic activity and leaching of essential nutrients when used subsequently. The best quality biomass meal is obtained with freeze-drying or spray-drying.

4.4 Artemia Cyst Harvesting and Processing Techniques Once the cysts have been harvested, a number of processing steps should be carried out to obtain a clean, marketable product with acceptable hatching properties and shelf-life. The processing can be classified into a number of successive processing stages (Fig. 4.10): harvesting, brine processing, freshwater processing, drying, prepackaging, packaging and dry storage. The processing activities within a processing stage can vary, or they can be applied simultaneously. According to specific requirements, some processing activities may be omitted. Basically, the processor will choose a combination of processing stages and diapause deactivation techniques depending on a number of factors such as trade-off between required final quality and economic viability, strain/batch specific characteristics and local conditions (e.g. site location, storage facilities, locally available equipment and scale of operation).

4.4.1 Harvesting techniques After being released, cysts float on the water surface and are washed ashore by winds and waves. In places with changing wind direction, cysts may be carried around for a long period before they are thrown ashore. If these cysts are produced in low-salinity ponds (⬍100 ppt) or when salinity stratification takes place after rainfall, quiescent cysts may hatch. When water is very agitated and foam develops, cysts can become trapped in airborne foam. Cysts washed ashore may be exposed to high temperatures, ultraviolet (UV) irradiation and repeated hydration/dehydration cycles, which in many cases decrease the viability of the final product. When dry, these cysts may also become airborne. The production of good quality cysts with reduced contamination by impurities is maximised when they are harvested from the water surface on a regular basis.

4.4.2 Brine processing 4.4.2.1 Brine dehydration To improve storage conditions and/or to deactivate diapause, cysts are usually dehydrated (to a water content of 20–25%) in saturated brine immediately after harvesting. When size

DIAPAUSE DEACTIVATION ACTIVITIES

dehydration dehydration dehydration aging in brine/hibernation

hydration/dehydration cycles peroxide treatment

DESCRIPTION OF CYST PROCESSING STEPS

AIM OF PROCESSING STEPS AND/OR PROCESSING

HARVESTING

apply proper harvesting procedures to ensure better quality of the cysts

BRINE PROCESSING dehydration size separation density separation raw storage

FRESH WATER PROCESSING removal excess brine density separation disinfection rinsing removal excess water

to prevent quality decrease during storage and/or to deactivate diapause removal of light and heavy debris in different size range of cysts removal of heavy debris (e.g. sand, stones, high dens. organic matter) temporary storage before or in-between brine processing steps storage for diapause deactivation storage before use as wet dry product (within 2 to 3 months)

to avoid salinity increase of freshwater and sub-optimal separation to separate high sinking fraction (mainly cysts) from low density floating fraction (mainly empty/cracked shells, light organic matter) reduce bacteria load remove salt just before drying to improve drying efficiency

BRINE PROCESSING brine dehydration raw storage

storage before use as a clean wet-dry product (within 2 to 3 months)

dehydration aging in brine/hibernation dehydration

DRYING

for long time storage

PRE-PACKAGING size separation air classification temporary packaging mixing

remove cyst aggregates remove remaining empty shells and non-cyst material optimal storage prior to final packaging obtain constant hatching quality

PACKAGING

oxygen free conditions to permit long time storage (>1 year)

DRY STORAGE

special storage conditions to increase shelf life

Fig. 4.10 Overview of cyst processing. (Modified from Lavens & Sorgeloos 1987.)


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and density separation equipment is located near the collection sites, brine dehydration is either combined with or performed immediately after density separation and size separation (see below). However, when there is a long period (up to several weeks) between collection and further processing, it is advisable to perform brine dehydration before size and density separation so as to avoid any loss of cyst quality. 4.4.2.2 Size separation in brine This involves the separation of debris from the cysts (e.g. feathers, sand, wood, stones) by screening the harvested product over different mesh sizes (e.g. 1 mm, 0.5 mm, 0.15 mm). For cyst material containing a lot of heavy debris (e.g. when collected from the shore), it is more efficient to perform a density separation in brine (see below) before size separation. 4.4.2.3 Density separation in brine Removal of heavy debris in the same size range as the cysts, when performed subsequent to size separation, is done through density separation in brine. Cysts immersed in brine float, while heavy debris (e.g. sand, small stones, heavy organic matter) sinks. Density separation is often performed soon after harvesting near production sites because of the availability of saturated brine. It can be combined with brine dehydration or cysts can be transferred to a special brine dehydration tank or pond following density separation. 4.4.2.4 Initial (or ‘raw’) storage Cysts are usually stored in a raw condition as follows:

• • • •

temporary storage (days or weeks) before the next brine processing stage temporary storage before the freshwater processing stage combination of raw storage and specific diapause deactivation methods raw storage for use as a wet–dry product (within 2–3 months).

Raw storage in low-saline brine (e.g. pond brine) Many strains can be stored in pond brine with salinity as low as 100 g l⫺1 for several days at ambient temperature without a decrease in the viability. It is essential that the cysts remain under hypoxic conditions to prevent initiation of the hatching metabolism, by storing them at a relatively high ratio of cysts to brine (20–80% volume:volume) without aeration. Raw storage in saturated brine After brine dehydration cysts can be stored safely for up to 1 month at ambient temperature. The cysts can be stored in containers submerged in brine or, alternatively, excess brine can be removed (e.g. hand squeezing) and the wet–dry product can be stored in bags made of cotton or jute. When stored as a wet–dry product over longer periods (⬎1 week), in areas of high relative humidity, crude salt should be mixed with the cysts to prevent rehydration of the highly hygroscopic cysts. Besides diapause deactivation as a result of the dehydration process itself, the storage (ageing) in brine may further deactivate the diapause in certain strains and batches.

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4.4.2.5 Cold storage The cysts of many strains and batches may be stored for several months to a year at temperatures between ⫺20 and 4°C. For certain species and strains, cold storage for several months is an adequate diapause deactivation method. Although many strains and batches have been stored safely without proper dehydration, cysts are usually dehydrated in saturated brine and packed as a wet–dry product before cold storage.

4.4.3 Freshwater processing During the freshwater processing stage, the cysts are further cleaned by density separation and prepared for subsequent drying. In freshwater, the cysts will partially hydrate and, if they remain hydrated for too long a period under aerated conditions, the embryos will eventually reach a stage of initiation of the hatching process that is irreversible and they cannot then be dehydrated without affecting the viability of the embryos (Clegg & Cavagnaro 1976). Moreover, their energy reserves may have been depleted to levels that result in a decrease in hatchability. Hence, freshwater processing should be limited to a maximum of 30 min. Before density separation in freshwater, excess brine must be removed to prevent a salinity increase in the water and, consequently, suboptimal separation. Cyst material immersed in freshwater will separate into a high-density (sinking) fraction and a low-density (floating) fraction. The sinking fraction contains mainly full cysts and some non-cyst material of similar density and similar size as the full cysts. The floating fraction contains mainly empty and cracked cyst shells and light non-cyst material of a similar size range. To reduce the bacterial load of the final cyst product the cysts can be disinfected during the freshwater treatment, e.g. by adding hypochlorite (liquid bleach) to the freshwater separation tanks before adding the cyst material. The concentration of active chlorine in the freshwater of the separation tanks should be less than 200 g l⫺1. If cysts are to be dried, they must always be rinsed thoroughly with freshwater to avoid crystallisation of any remaining salts during drying and consequent damage to the cyst shells. Following separation, rinsing and collection in bags, the bulk of the freshwater can be removed by firm squeezing of the cysts. The cysts can be dehydrated in saturated brine for raw storage and used, within 1–3 months, as a clean wet–dry product. Alternatively, the cysts can be dried immediately for long-term storage, to prevent further metabolism and decreased hatchability. Further removal of excess water can be achieved by centrifugation.

4.4.4 Drying The type of drying procedure used can affect the quality of the cysts in terms of hatching percentage and rate. Drying procedures differ in a number of factors. After the freshwater treatment, the water content of the cysts should be reduced as soon as possible below the critical level of 10% to arrest metabolic activity and, consequently, to ensure a long shelflife. Below a water content of 10%, little is known about the actual relation between water content and subsequent quality and shelf-life (Clegg & Cavagnaro 1976). Usually, a final water content between 3 and 8% is the objective. The best results are obtained when a water content of 10% is reached within 8 h or less. Few data are available on possible quality improvements when the drying time is very


141

short (⬍3 h). Prolonged drying (e.g. ⬎24 h) results in a decreased hatching percentage, possibly caused by a decrease in energy reserves. The maximal drying temperature is both strain specific and dependent on the degree of dehydration of the cysts. For fully hydrated cysts, temperatures below 35°C are usually safe. As the drying proceeds, water content decreases and cysts tend to be resistant to higher temperatures. If the freshwater processing cycle is limited in time and if excess water is properly removed, cysts are only partially hydrated (water content between 40 and 45%). Consequently, they may withstand higher temperatures (for some strains/batches, up to 60°C). Finally, heterogeneous drying will result in some cysts drying very slowly, eventually not reaching a water content of 10%. This may cause both a decrease in hatching percentage and hatching rate, and a reduced shelf-life. In summary, optimal results are obtained when ensuring a fast (⬍8 h) and homogeneous drying stage to a water content below 10%, without exposing the cysts to critical temperatures. Depending on the available equipment and financial considerations, different drying techniques can be applied. 4.4.4.1 Layer drying in open air Cysts are spread in thin layers of uniform thickness (a few millimetres only) on a drying rack (trays made with 120 ␮m screen), sheltered from direct sun irradiation and provided with good air exchange. Exposure to direct sunlight may result in critical temperature increases within the cysts (through heat absorption by the dark shell) or in UV damage to the embryos. A final equilibrium will be reached, if dried for long enough, between the water content in the cysts and the relative humidity of the air; for example, at a relative humidity of 70–75% cysts may reach a water content of about 10–15% after a maximum of 48 h. In areas with variable air humidity, poor standardisation and slow drying, inherent to this method, may result in fluctuating cyst quality. Moreover, owing to poor mixing, small aggregations of cysts may form which may affect the overall quality of the final product. 4.4.4.2 Layer drying in oven Drying racks are placed in a temperature-controlled room or oven with good air exchange. Heating incoming air significantly decreases the relative humidity, thus improving the drying. The relationship between temperature tolerance and water content of the cysts should be checked to determine the most efficient temperature cycle and so avoid overheating. Although a better standardization is possible, drying may still be quite slow and the problem of cyst aggregation remains. 4.4.4.3 Fluidised bed drying The most efficient and most versatile drying is obtained by means of a fluidised bed dryer (Figs 4.11, 4.12), which consists of a conical drying chamber, a blower and a heating unit with temperature control device (Bosteels et al. 1996). The blower forces air over the heating unit into the drying chamber. Sieves at the inlet and outlet of the drying chamber allow free airflow without loss of air-suspended (fluidised) cysts. The conical shape of the drying

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100 µm screen

Removable part of cone for addition or removal of cysts Temperature sensor (control of temperature in cone)

Temperature control device

Temperature sensor (control of inflowing air) Blower unit 100 µm screen

Heating unit

Fig. 4.11 Schematic drawing of a fluidised bed dryer for Artemia cysts.

Fig. 4.12 Fluidised bed dryer for Artemia cysts.


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chamber ensures optimal mixing of the cysts throughout the drying process, which results in homogeneous drying without excessive formation of cyst aggregations. Improved drying efficiency is further obtained by the heating unit. If the inlet and cone temperature are limited to 80 and 40°C, respectively, a unit as described above will dry approximately 35 kg of wet cysts in less than 3 h to a water content below 10%. Some species and strains of cysts seem unsuited to the intense turbulence inherent to this type of drying, and give lower hatching values postdrying. To cope with these cysts, the fluidised bed dryer may include cylindrical drying chambers, into which hot air is introduced from below at a moderate rate. The air inflow is insufficient to blow the cysts high into the air, and so homogeneous mixing of the cysts is achieved by a series of blades rotating about a central vertical axis within the drying chamber.

4.4.5 Prepackaging, packaging and storage During drying (especially layer drying), small aggregations of cysts are formed, which can be removed by dry sieving to improve the visual appearance of the final product. Air separation of cysts is often applied to separate any remaining empty and cracked shells that were not removed during freshwater separation. This can be effected in a horizontal air stream in which heavy particles tend to fall down more quickly than light particles, but it is often combined with the fluidised bed drying process itself: if a mesh is installed at the top outlet of the air chamber, the air inflow can be regulated to allow empty cysts to be blown through the mesh. Variations in hatching quality of the dry cysts may require mixing of different cyst batches to ensure a marketable product of constant quality. Any type of mixing equipment may be used, provided cysts are not exposed to high humidity (to avoid rehydration). Dry cysts should be packed in oxygen-free conditions to prevent formation of free radicals resulting in the irreversible interruption of the hatching metabolism. This can be achieved by vacuum or nitrogen packing. Properly packed cysts may be stored for months or even years without any significant decrease in hatching success. Furthermore, the shelflife of dry cysts is strain- and batch-specific. Although some strains may be stored at room temperature, storage temperatures below 10°C are usually recommended.

4.5 References Baert, P., Bosteels, T. & Sorgeloos, P. (1996) Pond production of Artemia. In: Manual on the Production and Use of Live Food for Aquaculture (Ed. by P. Lavens & P. Sorgeloos), pp. 196–251. FAO Fisheries Technical Paper 361, Food and Agriculture Organisation of the United Nations, Rome. Baert, P., Anh, N.T.N., Vu Do Quynh & Hoa, N.V. (1997) Increasing cyst yields in Artemia culture ponds in Vietnam: the multi-cycle system. Aquacult. Res., 28, 809–814. Beardmore, J.A. (1987) Concluding remarks for Symposium Session I: morphology, ecotoxicology, radiobiology, genetics. In: Artemia Research and its Applications, Vol. 1, Morphology, Genetics, Strain Characterization, Toxicology (Ed. by P. Sorgeloos, D.A. Bengtson, W. Decleir & E. Jaspers), pp. 345–346. Universa Press, Wetteren.

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Belovsky, G.E., Larson, C. & Mellison, C. (2000) Brine shrimp population dynamics and sustainable harvesting in the Great Salt Lake, Utah. Progress Report to the Utah Division of Wildlife Resources, Salt Lake City, Utah, USA. Bosteels, T., Tackaert, W., Van Stappen, G. & Sorgeloos, P. (1996) Improved use of the fluidized bed dryer for Artemia cysts. Aquacult. Eng., 15, 169–179. Boyd, C.E. (1990) Water Quality in Ponds for Aquaculture. Birmingham Publishing, Birmingham, AL. Browne, R.A. (1980) Competition experiments between parthenogenetic and sexual strains of the brine shrimp, Artemia salina. Ecology, 61, 466–470. Browne, R.A. & Halanych, K.M. (1989) Competition between sexual and parthenogenetic Artemia: a re-evaluation (Branchiopoda, Anostraca). Crustaceana, 57, 57–71. Clegg, J.S. & Cavagnaro, J. (1976) Interrelationships between water and cellular metabolism in Artemia cysts. IV. ATP and cyst hydration. J. Biophys. Biochem. Cytol., 88, 159–166. Eliot, J.M. (1977) Statistical Analysis of Samples of Benthic Invertebrates. Freshwater Biology Association, Scientific Publication No. 25. Geddes, M.C. & Williams, W.D. (1987) Comments on Artemia introductions and the need for conservation. In: Artemia Research and its Applications, Vol. 3, Ecology, Culturing, Use in Aquaculture (Ed. by P. Sorgeloos, D.A. Bengtson, W. Decleir & E. Jaspers), pp. 19–26. Universa Press, Wetteren. Krebs, J.C. (1989) Ecological Methodology. Harper & Row, New York. Kungvankij, P. & Chua, T.E. (1986) Shrimp culture: pond design, operation and management. NACA Training Manual Series No. 2, 68 pp. S. Sirikarnpimp, Bangkok. Lavens, P. & Sorgeloos, P. (1987) The cryptobiotic state of Artemia cysts, its diapause deactivation and hatching: a review. In: Artemia Research and its Applications. Vol. 3, Ecology, Culturing, Use in Aquaculture (Ed. by P. Sorgeloos, D.A. Bengtson, W. Decleir & E. Jaspers), pp. 27–63. Universa Press, Wetteren. Stephens, D.W. (2000) Brine shrimp ecology in the Great Salt Lake, Utah, July (1998) through June (1999). Progress report, prepared in cooperation with Utah Division of Wildlife Resources, Great Salt Lake Ecosystem Project, Salt Lake City, Utah, USA. Tackaert, W. & Sorgeloos, P. (1991) Semi-intensive culturing in fertilized ponds. In: Artemia Biology (Ed. by R.A. Browne, P. Sorgeloos & C.N.A. Trotman), pp. 287–315. CRC Press, Boca Raton, FL. Tackaert, W. & Sorgeloos, P. (1993) The use of brine shrimp Artemia in biological management of solar saltworks. In: Proceedings of the 7th International Symposium on Salt, Vol. 1 (Ed. by H. Kakihana, H.R., Hardy, Jr, T. Hoshi & K. Tokyokura), pp. 617–622. Elsevier, Amsterdam. Triantaphyllidis, G.V., Poulopoulou, K., Abatzopoulos, T.J., Pinto Perez, C.A. & Sorgeloos, P. (1995) International study on Artemia. XLIX. Salinity effects on survival, maturity, growth, biometrics, reproductive and lifespan characteristics of a bisexual and a parthenogenetic population of Artemia. Hydrobiologia, 302, 215–227. Vu Do Quynh & Nguyen Ngoc Lam (1987) Inoculation of Artemia in experimental ponds in Central Vietnam: an ecological approach and a comparison of three geographical strains. In: Artemia Research and its Applications, Vol. 3 (Ed. by P. Sorgeloos, D.A. Bengtson, W. Decleir & E. Jaspers), pp. 253–269. Universa Press, Wetteren.

Chapter 5

Production and Nutritional Value of Copepods Josianne G. Støttrup

5.1 Introduction In nature, copepods constitute a first vital link in the marine food chain leading from primary producers to fish. In the open water marine environment, calanoids dominate the herbivorous zooplankton and provide the food-chain base for practically all marine fish larvae and planktivorous fish (Pauly & Christensen 1995). In estuaries and coastal areas, harpacticoids are an important constituent in the diet of larval and juvenile fish, flatfish and salmonids (Hicks & Coull 1983; Huys & Boxhall 1991). Thus, copepods play a central role in the global production of fish, which by 1999 supported an estimated total capture fishery production of around 92 million tonnes (FAO 2000).

5.2 Biology 5.2.1 General characteristics Copepods are aquatic animals, mostly marine, although many species occupy freshwater or estuarine habitats. The name copepod is derived from the Greek kope meaning ‘oar’ and podos meaning ‘foot’, and refers to the flat, paddle-like swimming legs. Around 200 families with some 1650 genera and 11,500 species were classified by 1993 (Humes 1994). Freeliving copepods inhabit a variety of habitats, ranging from the planktonic copepods that inhabit the world’s oceans, through benthic species that live on the surface of macroalgae or inhabit microscopic spaces in marine sediments, to subterranean species living in groundwater or in deep-sea hydrothermal vents. Almost one-third of marine copepod species are parasites or live in symbiotic relationship with other organisms. Aquaculturists may involuntarily be better acquainted with fish parasites than with the more numerous, less harmful, free-living copepod species. This chapter deals with the biology, culture and use of free-living copepod species as live feed in aquaculture. The most commonly used species in aquaculture are free-living copepods belonging to three of the ten orders of copepod (reviewed by Huys & Boxshall 1991): Calanoida,

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Harpacticoida and Cyclopoida. Figure 5.1 depicts an adult and a nauplius representative from each order. 5.2.1.1 Calanoida The calanoids are predominantly pelagic, occurring at all depths, with some near-bottom and benthic species. They are selective feeders feeding on small phytoplankton cells by filtration, or predators feeding on a variety of animal prey including copepod eggs. They are distinguished by their long antennules, as long as the body itself or even longer, with up to 27 segments and biramous antennae used as accessory locomotory appendages (Huys & Boxshall 1991; Dussart & Defaye 2001). The position of the prosome–urosome articulation is between the fifth and sixth postcephalosome somite (Mauchline 1998; Dussart & Defaye 2001). For a comprehensive review on the biology of calanoid species consult Mauchline (1998). Calanoid species used in aquaculture as live prey for marine fish larvae are listed in Table 5.1.

Fig. 5.1 Schematic diagrams of an adult and nauplius representative from each of the three orders: Calanoida, Harpacticoida and Cyclopoida. Calanoida: (a) stage CV Calanus finmarchicus; (b) stage I nauplius C. finmarchicus. Harpacticoida: (c) male CV Tisbe gracilis; (d) N1 T. gracilis. Cyclopoida: (e) C5 Cyclops strenuous; (f) N1 Eucyclops serrulatus. (From Lebour 1916–1918; Dahms & Bergmans 1988; Dussart & Defaye 2001.) [Parts (e) and (f) reproduced with kind permission of Backhuys Publishers, Leiden.]

Table 5.1 Species of calanoids used in aquaculture as live prey for marine fish species. Calanoid species

Origin or application

Marine fish species

References

Acartia longiremis

Collected from wild

Wolf fish, Anarhichas lupus

Ringø et al. (1987)

Acartia pacifica, A. plumose

Collected from wild

Asian sea bass, Lates calcarifer; grouper, Epinephelus fuscoguttatus

Sunyoto et al. (1995)

Red snapper, Lutjanus argentimaculatus

Doi et al. (1997b) Jung & Clemmesen (1997)a Turk et al. (1982)

Acartia sinjiensis Acartia tonsa Acartia tonsa

Laboratory culture

Acartia spp.

Cod, Gadus morhua Fundulus spp., Elops saurus and squid, Loligo pealie Golden snapper, Lutjanus johnii; mangrove jack, Lutjanus argentimaculatus

Schipp et al. (1999)

Eurytemora affinis

Striped bass, Morone saxatilis

Chesney (1989)

Eurytemora affinis

Striped bass, Morone saxatilis

Tsai (1991)

Turbot, Scophthalmus maximus

Kuhlmann et al. (1981)

Dolphin fish, Coryphaena hippurus

Rippingale & MacShane (1991)

Sea horse, Hippocampus angustus


West Australian dhufish, Glaucosoma hebraicum Pink snapper, Pagrus aurata Wolf fish, Anarhichas lupus

Payne et al. (2001)a Payne et al. (2001)a Ringø et al. (1987)

Eurytemora affinis, Acartia tonsa and others Gladioferens imparipes

Outdoor tanks

Gladioferens imparipes Gladioferens imparipes Gladioferens imparipes Metridia longa

Co-fed with rotifers Co-fed with rotifers Collected from wild

Halibut, Hippoglossus hippoglossus

Rønnestad et al. (1998)

Outdoor and indoor systems

Cod, Gadus morhua

Gamble & Houde (1984)

Collected from wild

Grouper, Epinephelus coioides

Toledo et al. (1999)a

Temora longicornis Mixed zooplankton/ Pseudocalanus elongates Mixed copepods: Acartia tsuensis, Pseudodiaptomus spp., Oithona sp., some harpacticoids

(continued)

Table 5.1 (continued) Calanoid species



References

Mixed zooplankton

Outdoor enclosures, wild zooplankton Wild zooplankton

Cod, Gadus morhua

Kvenseth & Øiestad (1984)

Barramundi, Lates calcarifer; Australian bass, Macquaria novemaculeata; mulloway, Argyrosomus hololepidotus; sand whiting, Sillago ciliata

Battaglene & Fielder (1997)

Collected from wild/culture ponds


McEvoy et al. (1998)a

Collected from wild or outdoor pond cultures. Supplement over a short period Collected from wild and cultured for three generations


Næss & Lie (1998)a

Flounder, Platichthys flesus

Engell-Sørensen (1997)

Added on days 35–55.

Gilthead sea bream Sparus aurata

Bedier et al. (1984)

Mixed zooplankton

Mixed species: Eurytemora affinis, Acartia teclae, Centropages hamatus Mixed species dominated by Eurytemora affinis or Centropages hamatus Mixed copepods: Temora longicornis, Centropages typicus, C. hamatus, Acartia spp., Eurytemora hirundoides Frozen copepods a

Showed improved growth or survival, or higher rates of normal pigmentation in fish compared with diets comprising rotifers or Artemia nauplii.

Production and Nutritional Value of Copepods

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5.2.1.2 Harpacticoida The harpacticoids, which include over 50% of copepod species, are primarily marine, freeliving, benthic organisms. They inhabit sediments occupying spaces between sand particles (interstitial), burrowing into sediment (burrowers), or living on sediment or plant surfaces (epibenthic). They are distinguished by their short antennules, fewer than 10 segments, and biramous antennae. The position of the prosome–urosome articulation is between the fourth and fifth postcephalosome segment (Dussart & Defaye 2001). Table 5.2 lists the harpacticoid species used in aquaculture as live food for marine fish larvae.

5.2.1.3 Cyclopoida The cyclopoids include pelagic, epibenthic, benthic and parasitic species, and inhabit both freshwater and marine environments, although they are far more abundant in freshwater. In the marine environment, cyclopoids belonging to the family Cyclopinidae are predominantly benthic and those of the Oithonidae planktonic (Huys & Boxshall 1991). The third family contributing to the free-living cyclopoids, which inhabit saline waters, is the Cyclopidae, although the majority of species in this family are freshwater species. Several of the freeliving forms are predatory, attacking predominantly smaller, less robust fish larvae (Cooper 1996). In contrast to calanoids and harpacticoids, cyclopoids have uniramous antennae used to help catching food. The antennules in cyclopoids are shorter than in calanoids, rarely reaching beyond the cephalothorax, and have six to 17 segments (Huys & Boxshall 1991; Dussart & Defaye 2001). As in harpacticoids, the position of the prosome–urosome articulation is between the forth and fifth postcephalosome segment (Dussart & Defaye 2001). Cyclopoid species are not frequently used in the larviculture of marine species, as evident in Table 5.3.

5.2.2 Copepod morphology Free-living copepods have generally cylindrical bodies with a narrow abdomen (planktonic forms) or, in the case of benthic or surface-living forms, broader bodies and/or dorsoventrally compressed forms. The trunk is composed of a thorax (metasome) and an abdomen (urosome) (Fig. 5.2). The head (cephalosome) is fused with the thorax and bears anteriorly a typical median naupliar eye, a conspicuous set of antennae and the various appendages used for feeding and swimming. This anterior part has been designated as the prosome. The abdomen is generally narrower than the thorax, with no appendages except for the caudal rami. The genital opening is usually located in the first abdominal segment, whereas the last segment bears the anal opening (anal somite). The urosome ends in a furca formed of two symmetrical rami ornamented with setae. The antennules vary in size and may be used to distinguish among the three genera, being shortest in harpacticoids and longest in calanoids. This appendage bears different setae with chemosensory or mechanosensory functions, and the form of these antennules is closely related to the lifestyle of the copepod: pelagic, benthic, littoral, etc. (Dussart & Defaye 2001). When held out laterally from the body, the antennules help to slow down the sinking rate in calanoids (Mauchline 1998). This is rarely the case in harpacticoids and

Table 5.2 Species of harpacticoids used in aquaculture as live prey for marine fish species. Harpacticoid species


Amphiascoides atopus

Marine fish species/crustaceans

References

Grass shrimp; Pacific white shrimp, Penaeus vannamie; darter goby, Gobionellus boleosoma

Sun & Fleeger (1995)

Nanton & Castell (1999)

Amonardia sp. Euterpina acutifrons

Grey mullet, Mugil cephalus

Kraul (1983)a

Euterpina acutifrons

Mahimahi, Coryphaena hippurus

Kraul et al. (1991, 1992)a

Tigriopus japonicus

Black sea bream, Mylio macrocephalus

Lee et al. (1981)a

Sand borer, Sillago sihama

Lee & Hirano (1979) in Lee et al. (1981)

Tigriopus japonicus

As a supplement

Nassau grouper, Epinephelus striatus

Tucker & Woodward (1996)

Tisbe furcata

Co-fed with rotifers

Northern anchovy, Engraulis mordax

Hunter (1976)

Tisbe holothuriae

Sole and co-fed with rotifers

Turbot, Scophthalmus maximus

Støttrup & Norsker (1997)

Tisbe holothuriae

Sole diet and co-fed with Artemia

Dover sole, Solea solea

Heath & Moore (1997)a

Tisbe spp.(dominant) Temora longicornis, Centropages hamatus, Acartia clausi, Paracalanus sp.

Outdoor tanks with multiple copepod species

Dover sole, Solea solea

Jinadasa et al. (1991)

Tigriopus japonicus

Mud dab, Limanda yokohamae

Fukusho (1980)

Tisbe holothuriae, Tisbe sp., Schizopera elatensis

Sea bream, Sparus aurata

Kahan et al. (1982)

Tigriopus japonicus

Yellow-fin sea bream, Acanthopagrus latus

Tseng & Hsu (1984)

Tigriopus japonicus

a



151

Table 5.3 Species of cyclopoids used in aquaculture as live prey for marine fish species. Calanoid species



References

Apocyclops borneoensis

Sole diet or co-fed (1:1) with Artemia nauplii

Acanthopagrus cuvieri

James & Al-Kars (1986)a

Apocyclops royi Oithona sp.

Co-fed with rotifers and Artemia

Grouper, Epinephelus spp.

Su et al. (1997)

Striped patao, Eugerres brasilianus

Alvarez-Lajonchère et al. (1996)

a


Fig. 5.2 Diagrammatic illustration of the external morphology and relevant terminology of a female calanoid.

cyclopoids, but exceptions do exist such as the planktonic species Oithona. In benthic species the antennules help in anchoring the copepod to the substratum (Björnberg 1986). In some species of all three genera one or both antennules are used to grasp the female during copulation. The antennae are used as accessory locomotory appendages in calanoids and harpacticioids, whereas the uniramous form in cyclopoids is used to help catch and handle the food (Dussart & Defaye 2001). The structure of the oral aperture and surrounding appendages differs with species according to their feeding mode. Rudiments appear during the naupliar stages but are first fully functional during the copepodite stages. The oral aperture is situated ventrally and is surrounded by mandibles, which help to macerate the food, maxillules adapted to grasp and break up the food, and maxillae, which adopt different forms according to the feeding mode of the species and help in capturing the food. In filter-feeders the maxillae bear setae where

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particles are retained, accumulated and transferred to the mouth (Mauchline 1998). Price et al. (1983) describe the maxillae periodically combing the feeding appendages, collecting particles on the setae. In cyclopoids the labrum, which protects the oral aperture anteriorly, is armed with teeth whose number and shape are used in taxonomy. The maxillipeds generally appear during the sixth naupliar stage and are situated between the maxillules and aid in scraping the substratum or in cleaning the other appendages. This appendage is very developed in Oithona species where it is used for catching food (Björnberg 1986). Attached to the thorax are four or five pairs of swimming legs, essential for locomotion. They are also useful in taxonomy because the distribution of spines and setae on the legs is species specific. In calanoids and some harpacticoids, the last pair is often modified, being often reduced in females and enlarged and asymmetrical in males and used to grasp the female during mating. In cyclopoids this pair of legs is similar in both species, but considerably reduced or even absent (Dussart & Defaye 2001). In free spawners the fifth pair of swimming legs is reduced or absent in order to facilitate swimming movements (Björnberg 1986). In male calanoids this appendage may be modified so as to enable it to grasp the female with one leg, while the other is used to transfer the spermatophore to the female. In females, which produce egg sacs, this appendage may also be modified. The body of newly hatched nauplii has an oval shape, is dorsoventrally compressed and unsegmented, with relatively few appendages and a single naupliar eye. Cyclopoid nauplii have a pear-shaped body (Dussart & Defaye 2001). In many species the first naupliar stage, abbreviated to NI, has no oral aperture and lives on its vitelline reserves, barely moving. Its body is unsegmented and has three pairs of appendages: antennules, antennae and mandibles, during the first three naupliar stages (Hicks & Coull 1983; Dahms 1993; Dussart & Defaye 2001). The musculature is most primitive in cyclopoids, resulting in simple, erratic jerks forwards or sideways, whereas calanoids exhibit circular, somersaulting or helical movements (Björnberg 1986). During the remaining three naupliar stages there is a progressive development of setae and of appendages in the posterior end of the body. The external skeleton or integument consists of several layers arranged in three distinct structures: an outer cuticle composed of a thin epicuticle and a thicker procuticle, the epidermis and the basal lamina (Dussart & Defaye 2001). The procuticle consists of chitin in a protein/lipid matrix arranged in two layers in different orientation to each other. The cuticle is not entirely impermeable but is perforated by many pores. At the articulation points the cuticle is not chitinised to allow flexibility. 5.2.2.1 Digestive system The oral aperture is formed by the anterior labrum and the posterior labium. The labrum contains the labral glands, which produce secretions that may bind the food together and initiate digestion (Dussart & Defaye 2001). The precise composition of the secretions is not known, but they contain glucids, glycoproteins and mucopolysaccharides. From the buccal cavity a dorsal oesophagus leads to the midgut. The oesophagus is lined with chitin and its wall has longitudinal folds and strong muscular tissue. In several species of calanoids and harpacticoids an extension from the midgut leading dorsally and anteriorly, called the midgut diverticulum, functions as a hepatopancreas. This diverticulum forms part of the first section of the midgut which, together with the middle section, is involved in digestion. Absorption and the


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formation of faecal pellets take place in the posterior section of the midgut, which also functions as a primitive kidney. In this region of the midgut, lipid reserves are accumulated and distributed throughout the body in small, coloured droplets or concentrated in an oil sac, which extends dorsally along the digestive tract. From here the pellets move through the hindgut, also lined with chitin and with longitudinal folds, to be expelled via the anus placed dorsally on the anal somite (Mauchline 1998; Dussart & Defaye 2001). The faecal pellets are generally ovoid. The main nitrogenous compound excreted by copepods is ammonia (Le Borgne 1986). In the harpacticoid Tisbe holothuriae, ammonia production was shown to reflect feeding activity and suggested as an ideal feedback parameter for feeding control in a continuous production system (Støttrup & Norsker 1997). Phosphate, urea and dissolved primary amines are also excreted (Dussart & Defaye 2001). 5.2.2.2 Circulatory system A heart is only found in calanoids. It is located dorsally between the second and third thoracic segments and consists of one ventricle. The colourless blood is circulated through an anterior aorta and a system of sinuses. In harpacticoids and cyclopoids with no distinct heart, peristaltic movements of the muscles of the alimentary tract ensure blood circulation (Dussart & Defaye 2001). 5.2.2.3 Nervous system The central nervous system is simple and relatively uniform in copepods, and consists of a brain situated dorsally and anterior to the oesophagus. From the brain extends a ventral nerve chord that extends to the posterior end and comprises both sensory and motor nerves. A sympathetic nervous system innervates the digestive tract. It arises posteriorly from the brain and extends ventrally to the base of the labrum at the oral aperture (Dussart & Defaye 2001). A giant fibre system formed by two pairs of interneurons and numerous giant motor fibres is involved in the rapid escape movements in copepods. The giant fibres run along the nerve chord, branching off to segmental and intersegmental nerves, innervating longitudinal body muscles and flexor muscles of the swimming legs. Anteriorly, these fibres leave the brain to innervate the antennules. The naupliar eye is characteristic in all stages of copepods and is generally red and located anterodorsally (Fig. 5.3). The pigments are carotenoids associated with proteins or pro-melanins (Dussart & Defaye 2001). 5.2.2.4 Reproductive system The genital system is located dorsally in the prosome and lies along the digestive tract. In general, it consists of paired glands, ducts and a ventrally placed genital aperture. The male genital system is unpaired in calanoids and most harpacticoids and paired in cyclopoids (Fig. 5.4) and some harpacticoids. In most copepod females the genital system consists of a single ovary. The posterior end, the germinal site of the ovary, is where the oogonia multiply. The new oocytes are located anteriorly, where they grow in size and undergo

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Eye

Fig. 5.3 Tisbe holothuriae nauplius with a characteristic nauplius eye. (Photograph: J.G. Støttrup.)

Fig. 5.4 Picture of the cyclopoid Apocyclops panamensis with paired egg sacs. (Photograph: E.E. Lipman.)


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differentiation. The ripe oocytes occupy the oviducts, which run laterally along the thorax and enter the genital somite, where they merge into a genital cavity (Mauchline 1998; Dussart & Defaye 2001). Both sexes of cyclopoids and harpacticoids have paired genital apertures located ventrally (Huys & Boxshall 1991).

5.2.3 Reproduction Most copepods reproduce sexually. The male deposits a sac containing viable sperm called a spermatophore near the genital aperture of the female. In cyclopoids, the male transfers paired spermatophores from its genital aperture to the ventral surface of the female genital somite. Most calanoids are broadcasters, shedding eggs singly into the water. The number of eggs spawned in a single event may vary from a few eggs to 50 or more eggs, and each spawning event may occur about once every 24 h for extended periods. Free-spawning species such as various Acartia species may produce between 11 and 50 eggs female⫺1 day⫺1, producing a total of up ⬎1200 from one single spawning, and Calanus species between 15 and 230 eggs female⫺1 day⫺1 to a total of up to 3800 eggs female⫺1 (Mauchline 1998). Copepods can produce eggs that are non-viable or unfertilised, depending on food conditions (Poulet et al. 1994). Several calanoids, such as Centropages furcatus, some species of Acartia and several Calanus species, are known to spawn at night. In most cases a new mating is necessary for the female to produce eggs again. Other copepods, including cyclopoids and harpacticoids, have their eggs contained within one or two egg sacs (ovisac), which remain attached to the female genital segment until they hatch. In cyclopoids the eggs are contained within paired egg sacs (Fig. 5.4) (Huys & Boxshall 1991). In calanoids the eggs are not contained in a membrane but adhere to each other as an egg mass and remain attached to the female (Mauchline 1998). Each egg sac or egg mass may contain a few to 50 or more eggs and may be produced at frequent intervals of a few days. Females from different Tisbe species were capable of fertilising up to 12 broods from a single fertilisation and up to around 21 in other harpacticoid species (Hicks & Coull 1983). Average brood sizes for Tisbe holothuriae reared on three different artificial diets ranged from 58 to 86 eggs sac⫺1 (Gaudy & Guerin 1977). A review covering 27 different harpacticoid species showed a range of 3–229 eggs sac⫺1 (Hicks & Coull 1983). There are very few reported cases of repeated mating in harpacticoids and it is assumed that the female dies after having produced one or a number of batches of eggs from a single spermatophore. Clutch size in marine cyclopoid species varies from a few eggs up to around 100, produced at intervals from ⬍1 day to 4 days (Hopcroft & Roff 1996). Egg production is measured as number of eggs female⫺1 day⫺1, independently of the spawning method. Tisbe battagliai produces up to nine egg sacs, and an average of 10 offspring O+ ⫺1 day⫺1 (Williams & Jones, 1999). Since a new egg sac cannot be produced before the nauplii have hatched from the previous one and because of their larger size, fecundity in calanoids with egg sacs is about 7.5 times lower than in free spawners (Kiørboe & Sabatini 1995). Fecundity in species with egg masses is within the range of 10–200 eggs, and that of free spawners between 30 and 700 eggs (Mauchline 1998). Higher fecundity has been shown for some harpacticoid species. In laboratory experiments, the harpacticoid T. holothuraie fed artificial diets produced 258–416 nauplii per female (Gaudy & Guerin 1977).

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Parthenogenesis, asexual reproduction through generations of females, has been demonstrated for a few harpacticoid species under laboratory conditions, but there is little information on this phenomenon or its occurrence in nature (Dussart & Defaye 2001). Normal eggs have been stored under cool conditions in aquaculture to ensure sufficient quantities for rearing fish. Støttrup et al. (1999) stored normal eggs under anoxic conditions at 4°C for up to 12 weeks to examine the effect of cold storage on lipid composition. Egg viability was not registered, although hatching rates were lower than when hatching eggs directly. Close to 100% survival was achieved when nauplii and adults of Gladioferens imparipes were stored at 8°C for up to 12 days (Payne & Rippingale 2000d).

5.2.4 Resting or diapause eggs Resting eggs are produced by several species of copepod and are the primary mode of dormant state in calanoids. These copepod eggs are laid during development, whereby development is arrested, and possess an additional external envelope of variable thickness. Resting eggs are able to withstand long periods of desiccation, heat or cold (Dussart & Defaye 2001). They may occur in high quantities in the sediments up to 3.2 million eggs m⫺2 in the Inland Sea of Japan (Kasahara et al. 1975) and up to 5.5 million eggs m⫺2 in a Norwegian enclosed pond system (Næss 1996). Most of the resting eggs in the top 8 cm of the sediment remained viable for several years (Katajisto 1996). In the laboratory, they were observed to withstand disinfection of the egg surface (Næss & Bergh 1994: Acartia clausi and Eurytemora affinis). In culture, diapause egg production can be stimulated by the culture conditions such as short daylength and enhanced by low temperature (Ban 1992) or by subjecting the culture to high densities (Ban & Minoda 1994). In the latter case, the induction was chemically mediated, possibly through metabolic products in the culture medium. Further, rearing conditions during the naupliar stages may determine whether the resultant adults produce diapause eggs (Ban 1992). Resting eggs have only been reported for calanoids (Grice & Marcus 1981; Mauchline 1998), but dormant nauplii or copepodites are known from free-living representatives of all three copepod taxa discussed in this chapter, namely the Calanoida, Harpacticoida and Cyclopoida, although in the latter no dormancy is reported in marine species (Dahms 1995). Dormancy occurs more frequently in winter in species from temperate regions and higher latititudes, although summer encystment of adult harpacticoids and in cyclopoids has been reported. Dormancy in fertilised females was reported to occur exclusively in the freshwater cyclopoid copepod Cyclops strenuous (Næss & Nilssen 1991). More recently, reproductive-resting females were reported for Coullana canadensis (i.e. Scottolana canadensis), a brackish-water harpacticoid copepod, and may also occur in other species (Lonsdale et al. 1993). Daylength and temperature were the principal environmental cues that induce females to switch from active reproductive to a resting reproductive state.

5.2.5 Development, size and growth 5.2.5.1 Life cycle Copepod species belonging to these three orders considered for aquaculture have similar life stages. Once fertilised, the eggs pass into the water or into an egg sac. The egg is spherical and


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protected by a chitinised envelope. The larva that hatches from copepod eggs, the nauplius (NI), develops through five (a few calanoid species) or six moults before passing onto the copepodite stage where they display the general adult features. Most species commence feeding during the third or fourth naupliar stages, although a few commence feeding during the second (e.g. Acartia tonsa; Berggreen et al. 1988) and even fewer during the first naupliar stage (Mauchline 1998). Harpacticoid nauplii are believed to be able to feed from the first stage and undergo six moults, the final one resulting in the first copepodite stage (CI) (Hicks & Coull 1983; Williams & Jones 1994). In cyclopoids, the first nauplii of the cyclopoid Oithona ovalis have been shown to have a functioning gut (Fanta 1976), whereas in Apocyclops royi the digestive tract is incomplete during the first stage, which is of a very short duration, lasting for a few minutes (Chang & Lei 1993). The rate of development is dependent primarily on temperature, but within the optimal range other factors such as food quantity and quality may influence development (Williams & Jones 1994; Mauchline 1998). Various species of calanoids of the genera Acartia, Calanus and Pseudocalanus and the pelagic harpacticoid Euterpina acutifrons are reported to exhibit isochronal development where the different stages have virtually the same duration (Neunes & Pongolini 1965; Klein Breteler et al. 1982; Corkett et al. 1986). In other species such as Calanus finmarchicus the stage durations differ (Tande 1988). Several harpacticoid species tend towards progressively longer stage durations (Bergmans 1981). The harpacticoid Euterpina acutifrons is pelagic and some harpacticoid species have planktonic immature stages (Neunes & Pongolini 1965). Nauplii of the epibenthic harpacticoids S. canadensis and Tisbe sp. are pelagic (Hicks & Coull 1983). In an extensive cultivation system for sole, the principal (46%) copepod species to be found in the water column were young stages of Tisbe spp. (Jinadasa et al. 1991). 5.2.5.2 Mortality Mortality rates, in terms of naupliar survival to adulthood, for Tisbe species measured in the laboratory range from 2 to 23% (Bergmans 1981), and more recent reports on mortality rates for different species of Tisbe lie within this range (e.g. Guérin et al. 2001, T. holothuriae, 14%; Volkmann-Rocco & Battaglia 1972, T. clodiensis, 16%). Thus, in cultures with Tisbe species and under appropriate conditions a survival rate of 75% to adults could be expected. Klein Breteler (1980) estimated 46–75% survivial during one generation for three calanoid species: Acartia clausi, Temora longicornis and Centropages hamatus. 5.2.5.3 Size Calanoid eggs produced in egg sacs range in diameter from 70 to around 800 ␮m and are generally larger than freely spawned eggs, which are rarely larger than 200 ␮m (Mauchline 1998). Size ranges of newly hatched nauplii also vary. Newly hatched nauplii such as A. tonsa measure less than 100 ␮m in body length (Klein Breteler et al. 1982), whereas the larger Calanus pacificus nauplii measure around 220 ␮m (Greene & Landry 1985). Schipp et al. (1999) found Acartia spp. measuring 65 ␮m in width an ideal size for first-feeing Lutjanus johnii. The nauplii of Gladioferens imparipes measuring 126 ⫻ 67 ␮m was an ideal size for first-feeding seahorse Hippocampus subelongatus (Payne & Rippingale 2000b).

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1,2

Length in mm

1 0,8 0,6 0,4 0,2 0

NI

NII

NIII

NIV

NV NVI CI Copepod stages

CII

CIII CIV F CV F CVI F

Tisbe cucumariae

T. gracilis

T. furcata

Apocyclops royi

Fig. 5.5 Sizes of naupliar (N) of three Tisbe species and copepodite (C) stages of T. gracilis and a cyclopoid species, Apocyclops royi. (Drawn from Johnson & Olsen 1948; Dahms & Bergmans 1988; Dahms et al. 1991; Chang & Lei 1993.)

Marine free-living copepods may measure up to 10 mm in length. The carnivorous adult Euchaeta elongata measures 6.3–7.4 mm in total length (Greene & Landry 1985). The harpacticoids are smaller and range in size from 0.2 to 2.5 mm (Hicks & Coull 1983). Adult sizes are generally around 0.7–1.5 mm in body length (Ikeda 1973). The small nauplii of three different harpacticoid species commonly used for cultures are similar in size, the adults measuring less than 1 mm (Fig. 5.5). Although shorter in length, their width is similar to that of small calanoids. For example, the width of the first naupliar stage of Tisbe cucumariae is 68 ␮m (Dahms et al. 1991). Newly hatched nauplii of a Tisbe sp. collected near Halifax, Canada, measured 90 ␮m in length and females grew to around 2 mm in total length (Nanton & Castell 1998). The adult females are generally larger than the males. The larger forms are epibenthic and have the ability to swim or are among the few truly planktonic harpacticoids. The first naupliar stage of the cyclopoid Apocyclops royi is the larger specimen, whereas the adults are relatively small, measuring around 1 mm in length (Fig. 5.5). Cyclopoids are generally smaller than calanoids. A study of the female size of 11 species of the family Oithonidae from the central Red Sea showed a range in body length from 0.4 to 1.4 mm (Böttger-Schnack 1988). Apart from growth in length (total length or prosome length), growth can be expressed in dry weight, which increases almost exponentially with development (Mauchline 1998) in most species. Once they reach maturity, somatic growth ceases in adult females and egg production is thus often used as an expression of growth (Kiørboe et al. 1985). 5.2.5.4 Generation time The generation time, defined as the time interval between hatching of an individual and the hatching of its progeny, differs from species to species and is positively correlated with


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increasing temperature. In calanoids reared at different temperatures, generation times varied from around 1 week in Acartia spp. (A. sinjiensis 5–6 days at 28–30°C; A. tonsa 7 days at 25°C) to months (Mauchline 1998, Table 47). Males are often smaller than females, may develop more rapidly (shorter generation time) and have a shorter longevity. Similar population characteristics are found in harpacticoids (Bergmans 1981; Hicks & Coull 1983; Dahms 1987). The generation time in 34 different harpacticoid species ranged from 7 days in Tisbe holothuriae reared at 22°C to 42 days in Robertgurnenya sp. reared at 17°C. One outlier, Huntemannia jadensis, had a generation time of 167 days, but the culture temperature was only 8°C (Hicks & Coull 1983). Food supply and salinity may also influence development rates. The duration of development from the first naupliar stage to adult in Tigriopus japonicus was shorter with increasing salinity (Hagiwara et al. 1995).

5.2.6 Feeding, food quality and food availability 5.2.6.1 Calanoids Feeding Calanoids have a non-visual, active raptorial mode of feeding, capturing and ingesting a variety of animal prey (Tiselius & Jonsson 1990), including yolk-sac cod larvae as was observed for the carnivorous Euchaeta norvegica (Yen 1987). Acartia tonsa was shown to be able to switch between particle feeding and a predatory mode (Kiørboe et al. 1996). Copepod species may also create feeding currents that entrap non-evasive prey such as copepod eggs and nauplii (Yen & Fields 1992). In laboratory experiments, Artemia nauplii were used to investigate predation rates in adult Aetideus divergens, a copepod adapted to living below the euphotic zone and to feeding on larger particles such as faecal pellets or large phytoplankton cells (Roberston & Frost 1977). However, Artemia nauplii were suggested to be easier prey than copepod nauplii (Mullin & Brooks 1967). Predation rates of different surface-dwelling calanoid species of the genera Acartia, Centropages and Temora feeding on nauplii of their own or other species are reviewed by Daan et al. (1988) and vary from 0.11 to 11 prey adult⫺1 day⫺1. However, predation rates on nauplii decreased in the presence of sufficient alternative (phytoplankton) food. Calanoids are generally herbivorous filter-feeders, able to distinguish between particles and selecting between different food particles based on size or taste (Donaghay & Small 1979; Huntley et al. 1986). Unpalatable feed, such as inert beads, was regurgitated by Eurytemora affinis (Powell & Berry 1990) and the toxic dinoflagellate Gonyaulax grindleyi by Calanus finmarchicus (Sykes & Huntley 1987). The functional response (rate of consumption of algal particles) is influenced by the size, quantity and quality of the food (Paffenhöfer 1976; Kiørboe et al. 1985; Støttrup & Jensen 1990; Kiørboe et al. 1996). Size is important relative to the structure of the oral appendages, and copepods may be less efficient in retaining small cells, such as demonstrated for A. tonsa feeding on Isochrysis galbana, requiring higher concentrations relative to other algal species to reach maximum ingestion rates (Støttrup & Jensen 1990) (Fig. 5.6). The lower size limit for particle capture in A. tonsa was estimated at 2–4 ␮m (Berggreen et al. 1988). Small size was suggested to be one of the probable causes for poor survival, development and fecundity in Gladioferens imparipes fed Nannochloropsis oculata (Payne & Rippingdale 2000a), a phytoplankton species widely used in aquaculture (see Chapters 2 and 7) as a diet for rotifers.

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Cell numbers per ml

140 120 100 80 60 40 20 0

0

5

10

15 20 Cell size (µm)

25

30

Fig. 5.6 Relationship between maximum ingestion rate and algal cell size in the adult female calanoid Acartia tonsa. (Drawn from ⌬: Kiørboe et al. 1985; ■: Cowles et al. 1988; ◆: Støttrup & Jensen 1990.)

Ingestion rates Ingestion rates are calculated in laboratory experiments from the number of cells removed from a volume of water per unit time and expressed in different units such as numbers, volume or weight copepod⫺1 day⫺1 and percentage copepod body weight ingested day⫺1. Ingestion rates in terms of carbon, nitrogen or protein are also available in the literature. The mean daily ration of adult calanoids feeding on phytoplankton and expressed as a percentage of copepod body weight ingested day⫺1 ranged from 2 to 360% (Mauchline 1998, p. 187). In A. tonsa, values from 6 to 360% are reported. The different algal species used for different copepods in culture are given in Tables 5.4–5.6. Egg production The relationship between the rate of egg production and phytoplankton concentration in copepod species for which experimental data are available indicates that the food concentrations at which egg production commences and at which it attains a maximum level differ between species. Egg production per female per day increases with increasing food concentrations to an asymptotic level (Kiørboe et al. 1985; Dam et al. 1994) (Fig. 5.7). Food quality also influences growth and reproduction. Acartia tonsa produced more eggs when feeding on I. galbana than on Thalassiosira weissflogii, despite similar ingestion rates in terms of carbon and nitrogen (Støttrup & Jensen 1990). Some species of diatom used as food for copepods have been reported to reduce fecundity (Kleppel et al. 1991; Kleppel & Burkart 1995; Ban 1999) or be deleterious to copepod embryogenesis, resulting in variable hatching success due to the presence of toxic aldehydes (Ianora et al. 1995; Kleppel & Burkart 1995; Ban 1999; Miralto et al. 1999). A diet of Skeletonema costatum resulted in sterility or death in Temora stylifera (Ianora et al. 1995). Poulet et al. (1995) also showed low hatching success in Calanus helgolandicus feeding on dense cultures of Phaeodactylum tricornutum. Jónasdóttir (1994) found that the age of the diatom culture influenced the ability of Acartia sp. eggs to hatch. Ban (1999) reviewed the literature on diatom effects and found 31 combinations involving 12 estuarine or coastal ocean copepod species and 13 diatom species where either reduced fecundity or reduced hatching, or both, was reported. Earlier findings have long established the relationship between copepod egg production and lipid levels in the diet in C. helgolandicus (Gatten et al. 1980), and it has been

Egg production, eggs per female per day


161

45 40 35 30 25 20 15 10 5 0 0

5

10

15

20

25

30

35

40

45

50

Food concentration, cells per ml Fig. 5.7 Rate of egg production by Acartia tonsa as a function of food concentration of Rhodomonas baltica. (Drawn from Kiørboe et al. 1985.)

suggested that low levels of docosahexaenoic acid (DHA, 22:6n-3) and eicosapentaenoic acid (EPA, 20:5n-3) in the diet may reduce fecundity (Lacoste et al. 2001). Although no clear relationship could be established between copepod fecundity and EPA or DHA concentration in diets (Støttrup & Jensen 1990; Koski et al. 1998; Lee et al. 1999), the highest rate of egg production coincided with the highest DHA:EPA ratio in both A. tonsa (Støttrup & Jensen 1990; Jónasdóttir 1994; Jónasdóttir & Kiørboe 1996) and G. imparipes (Payne & Rippingdale 2000a). The importance of n-3 fatty acids compared with n-6 fatty acids was demonstrated for A. tonsa in laboratory experiments (Jónasdóttir 1994). When fed Dunaliella tertiolecta, a chlorophyte deficient in n-3 polyunsaturates, fecundity was irreversibly blocked in A. tonsa (Støttrup & Jensen 1990), in Pseudocalanus elongatus (Koski et al. 1998) and in C. helgolandicus (Lacoste et al. 2001). The role of these n-3 polyunsaturated fatty acids in copepods is not clear, but they are reported to be important in maintaining membrane fluidity in cold environments (Benson & Lee 1975) and facilitating catabolism of long-chain monoenoic fatty acids (Sargent & Henderson 1986). Thus, to ensure high fecundity in calanoids, ample food of the right size and nutritional quality must be supplied. Components other than lipids may also be important for copepod fecundity. Kleppel et al. (1998) showed the dietary content of amino acids to be correlated to egg production in A. tonsa. Amino acids are important sources of organic carbon and total nitrogen in copepods (Cowie & Hedges 1996). Protein and nitrogen content in food was shown to be positively correlated with fecundity in two species of Acartia, although lipids were shown to exert a superior influence on fecundity in A. tonsa (Jónasdóttir 1994). 5.2.6.2 Harpacticoids Feeding Harpacticoids are primarily detritivorous, benthic grazers, efficiently utilising various food sources such as bacteria (Rieper 1978), macroalgae, marsh grass, algal biofilm, diatoms,

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polychaete meat, mixed cereals (Guidi 1984), shrimp solids (Norsker & Støttrup 1994) and artificial diets such as Tetramin (Guérin & Gaudy 1976). Cannibalistic behaviour towards non-related nauplii has also been shown in a harpacticoid (Lazzaretto & Salvato 1992). Although harpacticoids eat practically anything, this does not imply that they are nonselective feeders and that their offspring production is independent of the type of diet offered (i.e. food quality). Egg production Food supply in terms of quality and quantity affects feeding, development and reproduction in harpacticoids. In a study on a harpacticoid, Tisbe battagliai, Williams and Jones (1999) showed that low algal concentrations decrease fecundity. Egg production ceased in the harpacticoid Scottolana canadensis at concentrations below 2.5 ⫻ 104 cells ml⫺1 but increased exponentially with increasing concentrations to a symptotic level (Harris 1977). Fecundity, longevity, survival and population densities obtained were highest for Tisbe furcata fed the diatom Skeletonema costatum at densities of 80 cells ␮l⫺1, whereas a diet of either Rhinomonas reticulata or Pavlova lutheri gave poorer results at maximum ingestion densities (Abu-Rezq et al. 1997). Tisbe furcata exhibited a preference for dead bacteria to dead Dunaliella tertiolecta, whereas its sibling species T. holothuriae and T. battagliai were indiscriminate in their feeding (Vanden Berghe & Bergmans 1981). Further, the rate of daily naupliar production was higher when fed a mixture of Isochrysis galbana and Thalassiosira pseudonana (5.6 nauplii female⫺1 day⫺1) compared with a single algal diet (3.2 on I. galbana alone and 4.2 on T. pseudonana alone). Tisbe holothuriae females produced approximately 8.4 nauplii daily when fed a diet of Rhodomonas baltica alone, compared with 2.5 on a diet of Dunaliella tertiolecta (Norsker & Støttrup 1994). The number of offspring per female in T. holothuriae was highest when fed a mixture of Ulva sp. and Fryfood® and lowest on Spirulina, a phytoplankton very high in protein (68%) (Miliou & Moraïtou-Apostolopoulou 1991). Detritus and associated micro-organisms have been suggested as important nutritional elements in the harpacticoid diet. Carbon derived heterotrophically was assimilated at a rate eight to 10 times higher than autotrophically derived carbon in certain harpacticoids (Brown & Sibert 1977), and Guérin et al. (2001) showed that the addition of small amounts of bacteria and vitamin D2 to a commercial pet food, Tetra Min, resulted in higher percentages of fecund females and higher total egg production of the culture. Because of the difficulty in assessing the nutritional contribution from bacteria, it may be hard to establish harpacticoid production on the merit of various feeds unless the experiments are carried out axenically. Harpacticoids are not known to store lipids as an energy source during or prior to reproduction, as in many calanoids. Ingested food is thus assumed to be invested directly into energy resources of the yolk (Hicks & Coull 1983). Indeed, lipid levels in harpacticoids are generally low, or lie within the lower range of that in calanoids. Whereas calanoids obtain copious amounts of n-3 polyunsaturated fatty acids (PUFA) from their phytoplankton diet, harpacticoids feed on other sources lacking or containing low levels of these fatty acids. Nanton (1997) fed two harpacticoid species yeast with a dietary lipid content of 2.2%; yet, despite the low PUFA content in the yeast, PUFA content in both copepod species was high.


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Egg production in Tisbe holothuriae was not reduced significantly when fed Dunaliella tertiolecta, a chlorophyte with trace amounts of n-3 PUFAs, including EPA and DHA, compared with Rhodomonas baltica, an alga with a relatively high content of long-chain polyunsaturates (Norsker & Støttrup 1994). In another harpacticoid, Tigriopus japonicus, high levels of DHA (12%) and EPA (7%) were observed when fed exclusively on baker’s yeast (Watanabe et al. 1978). In the absence of dietary n-3 PUFA, these results suggested that this harpacticoid was able to synthesise significant amounts of EPA and DHA to maintain a high production level and to incorporate quantities of these fatty acids in its offspring. Similarly, Tisbe sp. and Amonardia sp. were shown to synthesise significant amounts of EPA and DHA when fed a PUFA-poor diet (Nanton 1997). When fed an EPA-rich diet, EPA was converted to DHA. Because of the ability of the copepod to manipulate its relative contents of various fatty acids, the DHA:EPA ratio remained high (⬎2) in both copepod species despite their dietary fatty acid content (Nanton 1997). Levels of arachidonic acid (20:4 n-6) in T. holothuriae and in a Canadian Tisbe sp. were high (⬎1%) even when only trace amounts were present in the diet (Norsker & Støttrup 1994; Nanton 1997). Although HUFA content may be less important for production in harpacticoids, the content of protein seems to play an important role. Guidi (1984) showed that even though food composition did not influence ingestion rate, the content of nitrogen (protein) in the diet played an important role in egg production in Tisbe cucumariae. The highest number of ovigerous females was registered on diets containing 49–52% protein (% dry weight). Although there is evidence of the ability of harpacticoids to synthesise PUFA from HUFA precursors such as 18:3n-3, this has not been tested directly. Another explanation for the high PUFA levels in harpacticoids fed low PUFA diets may be their ability to utilise marine bacteria in the culture. Harpacticoids utilise bacteria as a food source (Rieper 1978), but although bacteria containing EPA have been found in deep-sea or low-temperature environments and to a lesser extent in warm-water environments, DHA in bacteria has so far only been found in deep-sea samples (Yano et al. 1997), making bacteria an unlikely source of DHA. 5.2.6.3 Cyclopoids Feeding Cyclopoids are considered omnivorous, feeding on planktonic and benthic prey and even on their own progeny, attacking nauplii or copepodites. Chang and Lei (1993) found that Apocyclops royi is herbivorous in the early stages, switching to predominantly carnivorous feeding during the later copepodite stages. These workers observed that this species attacks and ingests Artemia nauplii (⬍1 mm). Gyllenberg and Lundqvist (1978) showed that two cyclopoid species, Cyclops oithonoides and Halectinosoma curticorne, ingest and assimilate dissolved glucose. Like harpacticoids, both species ingested bacterial matter but only H. curticorne assimilated living bacteria. For many cyclopoids, turbulence is important for encounter rates with prey (Kiørboe & Saiz 1995). In tropical waters, diel periodicity was demonstrated for Oithona plumifera, with egg sacs being laid down at dawn and hatching at dusk on the following day (Hopcroft & Roff 1996). Thus, a light regimen would be important in the culture of these species. In nature, dominance of single species among planktonic cyclopoids is not as pronounced as in calanoids (Böttger-Schnack 1988).

Table 5.4 Cultivation techniques for different calanoid species.

Species cultured

Duration of culture

Food organisms

T/S

Culture volume/ system

Densities obtained ⫺1

Harvest

References

—

Zillioux (1969)

Acartia clausi

14 months

Twice weekly: Rhodomonas baltica (up to 50,000 cells ml⫺1) Isochrysis galbana (up to 50,000 cells ml⫺1)

15°C

100 litres recirculation

ⱕ40 l

Acartia clausi

1 year

Tetraselmis suecica

20°C

20 litres

300–350 adults l⫺1

Acartia clausi

Multiple generations

Equal density of Isochrysis galbana, Monochrysis (now Pavlova) lutheri (1 ⫻ 106 cells ml⫺1)

15°C

—

—

16 eggs adult⫺1 day⫺1

Iwasaki et al. (1977)

Acartia clausi


Equal density of Isochrysis 20°C galbana, Monochrysis (now Pavlova) lutheri (1.5 ⫻ 106 cells ml⫺1)

—

—


Iwasaki et al. (1977)

Acartia clausi

55 generations

Rhodomonas sp., Isochrysis galbana, Oxyrrhis marina

15°C

22 litres, 88 litres

Acartia clausi

⬎20 generations

Dinoflagellates, concentrations at 3–10 ␮g ml⫺1, checked 1–2 times daily

20–24°C

10 day batch cycle

Acartia clausi ⫹ Tisbe furcata


Tetraselmis suecica

15°C

40 litres

1000 ind l⫺1 mixed species

Acartia tonsa

10 months

Twice weekly: Rhodomonas baltica (up to 50,000 cells ml⫺1), Isochrysis galbana (up to 50,000 cells ml⫺1)

15°C

100 litres, recirculation

ⱕ40 l⫺1

Acartia tonsa

Several generations

Thalassiosira pseudonana, Isochrysis galbana, Chroomonas salina, equal volume 3 times weekly

18°C

1500 ml

Acartia tonsa

6 months

Natural phytoplankton blooms, seawater filtered through 5 ␮m

Ambient 6–28°C/ 1–26 psu

1890 litres circular outdoor tanks, water exchange 3 times a week

Person-Le Ruyet (1975)

Klein Breteler & Gonzalez (1982) 70 eggs F⫺1 day⫺1

Khanaichenko (1998)

Person-Le Ruyet (1975) —

Zillioux (1969)

Parrish & Wilson (1978)

11–95 l⫺1


Ogle (1979)

Acartia tonsa


Acartia tonsa

12 generations

Acartia tonsa

4 months

Acartia tonsa

⬎70 generations

Acartia tonsa

⬎20 generations

Acartia spp.

⬎6 months

Centropages hamatus Centropages hamatus Centropages hamatus

1 year 1 year Multiple generations

Centropages hamatus

55 generations

Centropages typicus

1 year

Centropages typicus Eurytemora affinis

1 year Multiple generations

Daily: Isochrysis galbana (3–18 ⫻ 104 cells ml⫺1), Rhodomonas baltica (3–11 ⫻ 104 cells ml⫺1) Isochrysis galbana, Rhodomonas sp. and a diatom Defatted rice bran, 1–3 g l⫺1 culture fed twice daily

ⱕ40 l⫺1

15°C/32 psu

100 litres, aeration

17°C

1500 ml

20–25°C/ 15–25 psu

170 litres

Rhodomonas baltica, Isochrysis galbana Dinoflagellates, concentrations at 3–10 ␮g ml⫺1, checked 1–2 times daily Rhodomonas sp., Tetraselmis sp., Isochrysis sp. (total 20,000 cells ml⫺1 ratio of 2:1:1)

16–18°C/ 35 psu 20–24°C

200 litres

28–32°C/ 30–34 psu

100 litres or 1000 litres batch ⫹ aeration

Tetraselmis suecica Tetraselmis suecica Daily: Isochrysis galbana (3–8 ⫻ 104 cells ml⫺1) Rhodomonas baltica (2–5⫻104 cells ml⫺1) Rhodomonas sp., Isochrysis galbana, Oxyrrhis marina Tetraselmis suecica

15°C 20°C 15°C/28 psu

40 litres 80 adults l⫺1 20 litres 110 adults l⫺1 100 litres, aeration ⱕ40 l⫺1

15°C


15°C

40 litres

87 adults l⫺1

20°C 15–20°C

20 litres 23 litres water exchange every 2–3 weeks

92 adults l⫺1 435 ind l⫺1

Tetraselmis suecica Isochrysis galbana, Cyclotella nana, Platymonas sp., Skeletonema costatum every 2–3 weeks

10 day batch cycle

—

Klein Breteler (1980)

Zillioux & Wilson (1966) 870–1680 N l⫺1 ⫹ 170–1520 adults l⫺1 100 adults l⫺1 23–27 eggs F⫺1 day⫺1 70 eggs F⫺1 day⫺1 2000 N l⫺1 harvested after 8 days

—

Turk et al. (1982)

Støttrup et al. (1986) Khanaichenko (1998)

Schipp et al. (1999)

Person-Le Ruyet (1975) Person-Le Ruyet (1975) Klein Breteler (1980)

Klein Breteler & Gonzalez (1982) Person-Le Ruyet (1975) Person-Le Ruyet (1975) Katona (1970)

(continued)

Table 5.4 (continued)

Species cultured Eurytemora affinis co-cultured with other species

Duration of culture

Food organisms

T/S

45 days

Nannochloris

15–20°C/ 16 psu

27 m3 in outdoor tanks, 7.5% harvested daily

Nannochloris sp.

15°C/ 12 psu

150 litres

Eurytemora affinis


2000 litres

Eurytemora affinis Eurytemora affinis

Isochrysis galbana, Thalassiosira pseudonana (ratio 1:1; 5 ⫻ 104 to 5 ⫻ 105)

19°C

50 litres

Densities obtained

Harvest

Reference

1–2000 N l⫺1 day⫺1

Nellen et al. (1980)

Barthel (1983) 3000 litres⫺1

Chesney (1989) Tsai (1991)

20,000 N day⫺1

Gladioferens imparipes

28 generations

Isochrysis sp., Pavlova sp., Phaeodactylum tricornutum

12–28°C/ 2–38 psu

15 litres

Pseudocalanus sp.

55 generations


15°C


Klein Breteler & Gonzalez (1982)

Pseudocalanus sp.

70 days

Isochrysis galbana clone Iso, Thalassiosira pseudonana clone 3H, Skeletonema costatum clone Skel, Thalassiosira weissflogii clone Actin

5°C

2 litres

Davis (1983)

Rhincalanus nasutus

7 generations

Diatoms Cyclotella nana, Thalassiorsira fluviatilis, Ditylum brightwellii, Coscinodiscus wailesii ⫹ naupliar Artemia salina

12°C

19 litres, stirred ⫹ aeration

Temora longicornis

2 months

Tetraselmis suecica

20°C

40 litres

Temora longicornis


Daily: Isochrysis galbana (3–8 ⫻ 104 cells ml⫺1) Rhodomonas baltica (2–5 ⫻ 104 cells ml⫺1)

15°C/ 28 psu

100 litres, aeration

Temora lonicornis

55 generations


15°C


45 eggs F⫺1

100 adults l⫺1 ⫺1

ⱕ40 l


Mullin & Brooks (1967)

Person-Le Ruyet (1975) —

Klein Breteler (1980)

Klein Breteler & Gonzalez (1982)

Mixed copepods

min. 3 generations

Natural phytoplankton

Mixed copepods: Acartia sp., Tisbe sp., Oithona sp.

6 weeks

Approx. 1/6 volume added daily with enriched seawater phytoplankton bloom

Mixed copepods: Temora longicornis, Centropages hamatus, Acartia clausi, Paracalanus sp., Tisbe sp.

15 weeks

1/3 water exchange first month, once weekly second month, twice weekly thereafter

Ambient

21–238 N l⫺1 day⫺1

1000 m3 30 m3, outdoor

10–20°C/ 28–33 psu

Engell-Sørensen (1999b) Gaudy (1978)

85 m3, outdoor ⫹ aeration

25 ind l⫺1

Jinadasa et al. (1991)

Selected copepod species (starter cultures)

Algae from aerobic waste stabilisation ponds, mixed with seawater

20 m3

10,000 l⫺1

Harvested to inoculate nursery ponds

Quin (1993)

Selected copepod species (nursery ponds)

Algae from aerobic waste stabilisation ponds, mixed with seawater

100 m3, flowthrough, water exchange every 20–30 days

1000–6000 ind l⫺1

Batch harvest used to inoculate grow-out ponds

Quin (1993)

Selected copepod species (grow-out ponds)

Eutrophic ponds

5–200 ha

100 ind l⫺1

Up to 8 ⫻ 107 (⬍200 ␮m) ⫹ 6 ⫻ 107 (200– 500 ␮m) h⫺1 harvest

Quin (1993)

T, temperature; S, salinity; ind, individuals; N, nauplii; F, Female.

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5.3 Production Methods 5.3.1 Extensive and outdoor cultures 5.3.1.1 Harvest of wild zooplankton Copepods have been used for culturing marine species for several decades. They are collected directly from nature, often from fjords or inlets where natural densities are high, and used directly as live prey, inoculated into outdoor tanks on land to produce live zooplankton for fish larval rearing, or harvested and frozen, dried or freeze-dried for later use as an inert diet. Several types of filtering device have been developed for this purpose. Barnabé (1980) described a floating propeller-induced device that directed the water current through an elongated plankton net at around 146 m3 h⫺1. Mesh sizes below 70 ␮m quickly became clogged, whereas sizes between 70 and 100 ␮m retained rotifers almost exclusively. Above 120 ␮m few rotifers were retained, whereas numerous copepod nauplii were retained, and 250 ␮m was sufficient to retain adult copepods. The UNIK filter system described by van der Meeren and Naas (1997) represents a more sophisticated plankton-collecting device, capable of concentrating a defined size fraction of copepods at a rate of 1–5 m3 min⫺1. It consists of a flow-through system equipped with rotating discs mounted with plankton gauze of different sizes (Fig. 5.8). Huse (1994) used this system to collect wild copepods, using mesh sizes 80 and 250 ␮m to obtain equal quantities of Acartia teclae and Centropages hamatus. This size range collects almost exclusively nauplii. By increasing the mesh size of the larger gauze, the size fraction can be broadened to include copepodites (350 ␮m) and adult copepods (600 ␮m) (van der Meeren & Naas 1997). When operated at 1 m3 min⫺1 in a eutrophic (artificially fertilised) saltwater lake with copepod densities of 20–50 l⫺1, enough food could be collected theoretically to feed 40,000–100,000 turbot fry. This calculation was based on a bioenergetic food consumption model developed by van der Meeren (1991). Even larger collection devices have since been developed which enable the filtration of larger volumes of water per hour. The Baleen filter described by Quin (1993) is mounted on a craft and is capable of filtering 200 l s⫺1 on a 63 ␮m screen without damaging the zooplankton, and subsequently sorting these into size fractions and concentrating these in separate containers. A further development of the UNIK filter is used in the lagoon Parisvatnet in Norway, and consists of a floating device with a large rotor at the bottom driving a water flow upwards and through the rotating filters, with a filtering capacity of 35 m3 min⫺1 (Fig. 5.9). 5.3.1.2 Production in enclosed fjords or sea areas Copepods occurring naturally in enclosed or semi-enclosed ‘polls’ with volumes of 37,000–510,000 m3 have been utilised directly for rearing marine fish in Norway (Svåsand et al. 1998) (Fig. 5.10). Potential predators in the enclosed system were initially killed off with rotenone (Naas 1990). The phytoplankton production is enhanced by adding agricultural fertilisers and where possible water-flow to maintain a high and stable production of zooplankton. In some systems, the copepod starting culture is derived from resting eggs in the sediment. In others, wild zooplankton are collected from the sea and transferred to the


169

Fig. 5.8 UNIK filter. Top: a UNIK filter viewed from the collection side. Bottom: inside the UNIK filter. The gauze on the rotating wheels (bottom) entraps copepods, which are then flushed by water jets (visible in the bottom picture) onto a collection pipe. The two wheels have different mesh sizes and the copepods entrapped on each of these wheels are collected in two separate containers (top). (Photographs: J.G. Støttrup; see text for more detail.)

170


Fig. 5.9 Free-floating zooplankton filtering unit used in the extensive rearing of cod in Norway to concentrate zooplankton from a fjord at a capacity of 35 m3 min⫺1. (Photograph: J.G. Støttrup.)

Fig 5.10 Parisvatnet (top): a natural semi-enclosed ‘poll’ used for rearing cod juveniles in Norway. Juveniles are maintained within the fjord (bottom). (Photographs: J.G. Støttrup.)


171

enclosures. The larvae are then transferred to these enclosures at densities of 0.01–0.32 l⫺1. Additional prey may be added during the larval rearing when necessary to maintain prey densities in the range of 200–500 l⫺1. In Norway, cod production from such systems varies from 0.02 to 1.6 cod juveniles m⫺3. From 1986 to 1994 a total of around 2 million juvenile cod was produced (Svåsand et al. 1998). Disadvantages with this type of system include the inability to control production and thus food levels and predators. Lack of food results in differential growth in fish larvae, which in cod leads to higher rates of cannibalism. Blooms of toxic algae can result in total mortalities for a year class, but experience has enabled the Norwegian culturists to prevent such blooms by nutrient manipulation to encourage diatom blooms rather than flagellate blooms, and mixing of the water layers by generating flow and adding filtered seawater. Another disadvantage is that it is difficult to assess the number of surviving juveniles and thus difficult to ration food at the time of weaning to dry diets. Collecting preweaning juveniles has been tried without much success (Svåsand et al. 1998). Despite improvements and experience in managing these systems, the restrictions and unpredictability of such systems are well recognised. 5.3.1.3 Production in outdoor ponds or large tanks Outdoor production in 350–5000 m3 ponds and tanks is carried out in Europe and Asia for the culture of round fish species, such as cod and grouper, and flatfish, such as turbot. Filtered seawater is generally used in these systems. By using filters of around 20–40 ␮m, natural phytoplankton can be transferred to the ponds without the accompanying zooplankton or potential predators. The phytoplankton can be monitored, and adding nutrients, generally commercial fertilisers (e.g. NPK complex) in small quantities can induce a bloom. Regarding the amounts to add, a good rule of thumb is to identify local fjords or inshore waters where there are regular algal blooms, and to examine nitrogen and phosphorus levels before the time of these occurrences and add fertilisers to achieve similar levels. Low nitrate concentrations favour algae with low Ks (concentration at which the uptake rate of nitrate is at half the maximum rate) values of nitrate, such as small flagellates (Parsons et al. 1978; Takahashi et al. 1982). In the experiment carried out by Naas et al. (1991), small flagellates dominated during the initial period before fertilisers were added and where the nitrate levels were registered at ⬍0.5 ␮M. In conditions of non-limiting nitrate levels (⬎5 ␮M) and high oxygen concentrations, larger diatoms were favoured, giving rise to a prolific copepod production. Silicate is sometimes added and this encourages the development of diatoms, which are preferable to the development of the smaller flagellates. In a Norwegian manipulated seawater enclosure, the increase in diatom production and improved oxygen conditions resulted in an increased production of calanoid nauplii (Naas et al. 1991). For this reason the nutrients in the water intake and the type of phytoplankton found in the rearing tanks are closely monitored in the Danish extensive system for rearing marine fish larvae (Engell-Sørensen 1999a,b). Filtering devices that allow for selective sieving are used to collect primarily nauplii (80–250 ␮m), copepodite stages (80–350 ␮m) or primarily adult stages (250–600 ␮m) to inoculate the rearing tanks. In Asia a mesh size of 400–600 ␮m was used to inoculate outdoor tanks for grouper rearing with copepodite and adult stages 3 days before stocking the newly hatched fish larvae at densities of 5 m⫺3 (Toledo et al. 1999). In this system, using wild-harvested copepods (chiefly Acartia tsuensis

172


with smaller amounts of Pseudodiaptomus spp., Oithona sp. and a very few harpacticoids), an average survival of 3.4% at harvest corresponded to an average production of 0.17 grouper Epinephelus coioides juveniles m⫺3. In the Norwegian systems, prey densities range from 10 to 300 l⫺1 and around 1.4–2.8 fish larvae l⫺1 are added, resulting in the production of 3.8–40 cod juveniles m⫺3 (Svåsand et al. 1998). The plankton found in these systems consists of primarily calanoid species: Eurytemora affinis, Temora longicornis, Centropages hamatus, Acartia spp., Calanus finmarchicus, Paracalanus parvus and Pseudocalanus elongatus, but other copepods such as Oithona similis and Tisbe sp. have been reported (van der Meeren & Naas 1997). In Norway between 1989 and 1993, around 140,000 cod juveniles were produced in such systems. In Denmark, extensive systems use either outdoor concrete tanks or tarpaulin-lined earth-ponds (Fig. 5.11). Around 0.05 yolk-sac turbot larvae l⫺1 are added to each pond and during successful runs a production of around 20 juvenile turbot m⫺3 can easily be achieved. The Danish turbot-producing hatchery described by van der Meeren and Naas (1997) and Støttrup (2000) has, within a couple of years of its establishment, produced around half a million turbot juveniles each year. At least one of the concrete tanks is used entirely for the zooplankton cultures, from which water can be filtered and zooplankton added to the fish rearing tanks. Good techniques for filtering zooplankton are essential as this system relies heavily on the addition of zooplankton during the fish larval rearing. The zooplankton most commonly reported in these systems is calanoids of the genera Acartia, Centropages and Temora. Because of the relatively high energetic demands of the fast-growing fish larvae, Artemia nauplii are sometimes added as a supplement when copepod densities are low (Naas 1990; van der Meeren & Naas 1997). In a similar system in France, the dominant copepod species developing in the system were Acartia sp., Tisbe sp. and Oithona sp. (Gaudy 1978). During the initial period a bloom of rotifers reaching well over 10,000 individuals m⫺3 developed, lasting for around 20 days. A bloom of nauplii and other zooplankton then replaced the rotifers. A similar sequence has been observed in some of the productions in a tarpaulin-lined pond system used to culture flounder, Platicththys flesus (Engell-Sørensen 1999b). In this system, in two out of five successful productions over a 3 year period, rotifers dominated at the outset, peaking at approximately 350–450 rotifers l⫺1. The duration of rotifer dominance was shorter in this system, lasting for around 8 days, whereupon they were replaced by copepods and especially nauplii. Newly hatched flounder larvae were stocked at around 0.08 l⫺1 and 11–47 flounder juveniles m⫺3 were produced. Copepods identified in this system were Temora longicornis, Centropages typicus, C. hamatus, Pseudocalanus elongates, Acartia spp., Eurytemora hirundoides and occasionally Oithona similis (Engell-Sørensen 1997, 1998, 1999a). Typically, one species dominated and between two and five other calanoid species were identified. During 1997, the dominant species in two productions was T. longicornis and the concurrent dominant algal species was Thalassiosira nordenskjoldii. The naupliar concentrations around first feeding were generally in the lower range compared with those reported in the Norwegian systems, 10–50 l⫺1. The rearing system for flounder consists of three outdoor ponds (1200 m3), two used for fish larval rearing and one for copepod culture, and has produced up to 82,000 flounder juveniles each year (one season) (EngellSørensen 1999b). Regular monitoring of densities of the live prey in these outdoor systems is important for the successful rearing of marine fish larvae. Obtaining reliable density estimates of the


173

Fig. 5.11 Harvesting juvenile cod reared in earth-ponds lined with black tarpaulin in Denmark. (Photograph: J.G. Støttrup.)

copepod populations within the larger outdoor systems is not an easy task owing to their patchy distribution and changes in their distribution throughout the day. In general, a large volume (10–25 litres) from various positions in the tanks is sampled and filtered, and the copepods are identified and counted (Jinadasa et al. 1991; Toledo et al. 1999). Van der Meeren

174


(1991) used a long tube, which sampled 11.5 litres from a 3 m water column. Dussart and Defaye (2001) recommend the plankton trap of Schindler-Patalas (Schindler 1969, in Dussart & Defaye 2001) for quantitative sampling in lakes. Regardless of the sampling technique, it is advisable to take the samples at the same time each day. The zooplankton samples can be preserved in 4% formalin buffered with borax (sodium tetraborate). The method described by Mauchline (1998) is very practical and convenient. This consists of adding 30 g of borax to 1 litre of analytical reagent-grade formalin (40%). The plankton sample should be concentrated and added to a bottle of known volume; glass vials with screw-on caps with easily identifiable millilitre marks are ideal for formalin storage. The volume of the settled copepod sample should not exceed 20–25%, and the total sample (copepods ⫹ seawater) not exceed 90% of the volume of the bottle. The final 10% is filled with the buffered formalin and should be stored in the dark for at least 10 days. Since formalin is detrimental to health, care should be taken when working with this preservative. After 10 days, the sample can be filtered onto a sieve, washed in seawater and transferred to another preservative fluid, less harmful to health. The formula for such a preservative fluid is given in Mauchline (1998, p. 10). An advantage of outdoor ponds over the extensive systems that rely on the local production of zooplankton is the possibility of ‘culturing’ the zooplankton over one generation before using them as food. Trematodes and cestodes, which infect marine fish, have been identified in copepods (Bristow 1990; Robert & Gabrion 1991; Marcogliese 1995). Thus, feeding wild zooplankton directly to the fish increases the risk of infection. Since many of these parasites use copepods as intermediate hosts between compulsory hosts, the use of the first generation nauplii in the system is sufficient to reduce the risk of parasite transfer for those parasites that use copepodites or adult copepods as intermediate hosts. The juveniles are usually metamorphosed when they are collected from the ponds or tanks and transferred to indoor tanks for weaning to a dry diet and ongrowing. Both the transfer and weaning may be very stressful to the fish and heavy losses may occur during this period. Vaccination against Vibrio shortly after the transfer has been shown to mitigate these losses. At the end of the season, the tanks are emptied and left dry during the winter. In some cases around half the water is left in the outdoor tanks, enough to ensure that the whole water column does not freeze. With the falling temperatures, the copepods produce resting eggs, which survive in the sediment until the following year and are used as starter zooplankton cultures in the following year when the larval rearing season recommences (Næss 1996; K. Engell-Sørensen, personal communication). Tarpaulin bags were placed in the polls or fjords in an attempt to increase control and allow manipulation of the biotic and abiotic parameters (Svåsand et al. 1998) (Fig. 5.12). The advantage here is that the number of bags used to rear fish can be regulated according to the zooplankton supply. It is easier in these systems to estimate the numbers of surviving juveniles and thus to ensure an adequate food supply. The bags used in Norway are cylindrical, 2–8 m in diameter and 4–6 m deep, with a flow-through system with a bottom outflow (van der Meeren 1991; van der Meeren & Naas 1997). It is important to ensure that the material used is non-toxic to the fish larvae, live prey and phytoplankton. The bags are filled with filtered (80 ␮m) seawater and copepod nauplii added to densities of 100–500 l⫺1. In Norway almost half a million cod juveniles were produced in such systems between 1988 and 1994 (Svåsand et al. 1998). This production system, like many of the other extensive or semiextensive systems, needs to be developed further to ensure predictable production cycles.


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Fig. 5.12 Experimental floating tarpaulin bag (foremost) placed in an enclosed fjord for rearing cod juveniles in Norway. (Photograph: J.G. Støttrup.)

5.3.2 Intensive culture of copepods Since the early 1990s there has been a revival of attempts at intensive cultures of specific species of copepods, partly in response to a global shortage of Artemia eggs (see Chapters 3 and 4) and partly to diversification to new culture species with very small larvae, such as marine ornamental fish species (Payne & Rippingale 2000b) or species difficult to rear on the traditional live prey, rotifers or Artemia nauplii, or with a small mouth size, such as grouper (Epinephelus sp.) (Toledo et al. 1999), dhufish (Glaucosoma sp.) (Payne et al. 2001) and red snapper (Lutjanus argentimaculatus) (Doi et al. 1997a). One calanoid or harpacticoid species often dominates, the species possibly taking advantage of some environmental parameter to outcompete other species. Although the mechanisms for competition have not been fully explored, production of toxic metabolites and genetic adaptation to particular environmental conditions (niches) have been proposed (Bergmans & Janssens 1988). These dominant species are ideal candidates for intensive rearing. Several attempts to mass-culture copepods in intensive systems have been undertaken with varying success and have resulted in the development of different systems for particular species of copepods (Tables 5.4–5.6). Rearing in larger volumes (⬎10 litres for calanoids and ⬎2 litres for harpacticoids) may be more representative of the conditions required for mass rearing. For further references including small-scale (millilitre to litre) cultures, consult Ikeda (1973) and Paffenhöfer and Harris (1979). Species with relatively short generation times at ambient temperatures are best suited for aquaculture purposes. Species inhabiting coastal environments are normally more tolerant to variations in salinity and temperature and have a wider thermal and salinity tolerance. 5.3.2.1 Calanoids The most frequently cultured calanoid species belong to the genera found in coastal waters, such as those of the genera Acartia, Centropages, Eurytemora and Temora (Table 5.4). These copepods are small, with relatively short generation times and a wide thermal and

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salinity tolerance, and are easily adaptable to laboratory conditions. When reared in outdoor multispecies cultures, they tend to dominate with time. Food and feeding Most calanoids require the provision of phytoplankton, which can be filtered from the water, although Turk et al. (1982) demonstrated that it was possible to culture Acartia tonsa on rice bran. In many cases, the copepods are reared on monoalgal diets, which may not comply with all the requirements for maximum egg production. In culture systems where the culture medium is not exchanged daily, waste products and superfluous feed may accumulate and generate problems with ciliates and other contaminants, which may cause the culture to collapse. Somatic growth ceases in adult copepods and growth rate is more or less equivalent to the rate of egg production. Similar to ingestion rates, the rate of egg production in copepods is dependent on the size, quantity and quality of the algae provided. The algal concentration at which egg production commences or is at its maximum rate differs in copepod species depending on the diet, as discussed earlier. As a general rule, to reach food saturation, high ingestion rates and high egg production rates, cell concentrations of around 103 cells ml⫺1 would be sufficient using larger cells such as Thalassiosira weissflogii and Ditylum brightwellii (⬎12 ␮m in mean diameter), around 104 cells ml⫺1 using smaller cells such as Rhodomonas baltica (around 5–12 ␮m) and 105 cells ml⫺1 using yet smaller cells such as Isochrysis galbana or Pavlova lutherii (Støttrup & Jensen 1990; Payne & Rippingale 2000c; Lacoste et al. 2001). This is also illustrated in Fig. 5.6 for A. tonsa. Larger calanoid species may be less efficient in feeding on the smaller algal sizes. Food is provided at regular intervals (daily to two or three times weekly), and food density is either counted to ensure specific densities or regulated according to the water turbidity in the culture tanks. A combination of at least two algal species with high n-3 polyunsaturated lipid content, and of a size that can be utilised by both the feeding naupliar stages and the copepodite and adult stages, probably comprises an adequate diet for culture. Since the fatty acid distribution in adults and their non-feeding offspring reflects that of the adult diet, this would also ensure a suitable fatty acid distribution in the copepods used as live feed. A combination of I. galbana, high in DHA (DHA:EPA ⫽ 29.3), and R. baltica, high in EPA (DHA:EPA ⫽ 0.6), was used for the culture of A. tonsa (Støttrup et al. 1986) and G. imparipes (Payne & Rippingale 2000b; 3:1 by cell numbers). The food ration was adjusted according to water turbidity, ranging from 6–8 ⫻ 104 to 1.2–1.4 ⫻ 105 cells ml⫺1 day⫺1. In the culture system described by Støttrup et al. (1986), the eggs sedimented to the bottom from where they were siphoned daily, simultaneously siphoning out debris, faecal matter and associated ciliates (Fig. 5.13). During the siphoning, the eggs were concentrated on a 45 ␮m sieve, allowing most of the debris and ciliates to pass through and be removed from the culture. The daily removal of eggs eliminates the potential loss of nauplii through cannibalism by the adult population. The removed debris was checked daily, together with the egg count; and the presence of a particular protozoan, the ciliate Euplotes sp., heralded the deterioration of water quality and a thorough water exchange was necessary. Failing to do this would result in a culture crash. The culture was filtered through a 180 ␮m sieve submerged in seawater to retain the adult population and wash out the ciliates. A smaller sized gauze (80–120 ␮m) would also have been sufficient. The adults were then used to inoculate


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Fig. 5.13 System for culturing the calanoid Acartia tonsa at the North Sea Centre, Hirtshals, Denmark. The eggs were siphoned daily off the bottom of the 180 litre black tanks (left), concentrated on a 45 ␮m filter and transferred to 100 litre containers (right).

a new tank filled with filtered (1 ␮m) seawater. The frequency of this water exchange varied, but was generally done around every 2–4 months. A reliable batch culture system for rearing Acartia sp. is described in Schipp et al. (1999), showing consistent production results over an 8 day cycle in three 1000 litre tanks run concurrently over a period of 7 weeks. Starting with an inoculum of around 50–100 adults and 150–250 copepodites per litre, the culture contained after 7 days around 2000 nauplii, 750 copepodites and 300 adults per litre. The culture of Acartia tonsa without cultured algae has been achieved using rice bran over a 4 month experimental period (Turk et al. 1982). These authors used defatted rice bran sieved through a 73 ␮m mesh and mixed in deionised water before adding twice daily to the copepod culture at concentrations of 1–3 g l⫺1 culture. If the culture water did not clear between feedings, the feeding level was reduced. Two or three times weekly the bottom was siphoned, but because many copepods would be lost with each siphoning, this was done as infrequently as possible. As discussed later (culture tank size and shape), a deeper tank with a smaller diameter, or in this case a smaller surface area, may have alleviated this problem. The greatest advantage of this culture method is that it does not rely on algal cultures. A further advantage of this system and that of Støttrup et al. (1986) is that known age (and size) cohorts of A. tonsa can be obtained by separating out eggs or nauplii and maintaining each day’s production in separate tanks. By growing the nauplii for a set number of days, one could ensure the availability of nauplii of the required size as live feed for larvae of herring, Clupea harengus (Kiørboe et al. 1985), plaice, Pleuronectes platessa (Støttrup et al. 1986), cod, Gadus morhua (Grønkjær et al. 1995), Fundulus spp. and Elops saurus (Turk et al. 1982). Light Light levels and periodicity are rarely reported in the literature. In many cases no particular light is provided and ambient light levels are not recorded. In general, low light levels are applied. Davis (1983) used a light/dark (L/D) cycle of 10/14 h at an intensity of

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18 ␮E m⫺2 s⫺1 for rearing Pseudocalanus sp. In the system of Støttrup et al. (1986) the light intensity was 25 ␮E m⫺2 s⫺1 and lids were placed over the production tanks during the night. Photoperiod (8L/16D) and constant illumination at an intensity of 1000–1200 lux at the water surface was used for rearing four calanoid species (Person-Le Ruyet 1975). High density fluctuations were observed in the cultures independent of light regime. Several calanoids are known to spawn at night (Mauchline 1998). Egg production in the calanoid Labidocera aestiva was examined under cycling light/dark conditions and compared with constant light or darkness (Marcus 1985). In this study, maximum egg release occurred during the dark phase and a rhythmic pattern was evident under constant dark conditions but not under constant light conditions. It is also important to bear in mind that in some species a short photoperiod is the primary cue for the production of diapause eggs, as was shown for Eurytemora affinis exposed to 10 h light (60 lux)/14 dark (Ban 1992). Hence, in culture, the most favourable regimen seems to be a photoperiod of at least 12 h of light. In nature, high solar radiation is harmful to copepods; hence, adults show negative phototaxis during the day and positive phototaxis during the night (Dussart & Defaye 2001). Positive phototaxis in Gladioferens imparipes nauplii was used advantageously to concentrate nauplii before harvest from 500 litre culture units (Payne & Rippingale 2000c). In a continuous recirculation system provided with constant illumination at 1500 lux, Zillioux (1969) reported that the copepods gathered in the corners, thus facilitating harvest. Aeration and oxygen Aeration is required to help to maintain phytoplankton in suspension and to create small turbulence, which helps to distribute the copepods within the culture tanks and prevent ambient low algal concentrations due to patchy distribution of predator and prey. Aeration also helps to prevent anoxic conditions. Turbulence is less important for suspension feeders, which establish their own feeding currents, but it has no negative effect on these species. In contrast, turbulence is important in ambush-feeding copepods such as Acartidae, in that it increases the encounter rate (Kiørboe & Saiz 1995). Gentle air-lifts, air-stones or air bubbles may be used to support a gentle upward and circulating flow. Too vigorous aeration should be avoided and is unnecessary, and air bubbles that are too small may become entrapped in the copepod appendages and should be avoided. In nature, copepods live in well-oxygenated environments. Coastal species produce eggs that sink to the bottom sediment and are able to survive anoxic conditions. In outdoor ponds used for rearing marine fish larvae, oxygen supersaturation of up to around 160% has been registered (Engel-Sørensen 1998), with no evidence of adverse effects on the copepod population. Culture tank size and shape Most calanoids require large volumes and the adult density rarely exceeds 100 per litre. Higher densities have been achieved in Acartia clausi, Eurytemora longicornis and E. affinis (Katona 1970; Person-Le Ruyet 1975; Chesney 1989). Tanks used by Støttrup et al. (1986) for culturing Acartia tonsa through several hundred generations are cylindrical with a capacity of 200 litres and a centrally placed air-stone with very gentle aeration (Fig. 5.13). The bottom is flat to enable siphoning of the eggs from the bottom. A high tank height to


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tank diameter (tank bottom area) is considered advantageous in reducing the surface area to be siphoned and thus the loss of copepods. Furthermore, a central air-stone may be sufficient to ensure proper circulation within the tank column without too vigorous aeration. Turk et al. (1982) used square tanks 100 ⫻ 50 ⫻ 50 cm with air-lifts in two opposite corners. The bottom was flat to enable debris to be siphoned off the bottom, but they suggested doing this as rarely as possible to prevent loss of copepods. The low tank height to bottom surface area may result in high numbers of adult copepods being siphoned together with the bottom debris. The successful batch culture of the calanoid Acartia sp. was achieved in 1000 litre polyethylene tanks, 1.3 m in diameter with a conical base. These tanks were emptied after the 8 day batch cycle and cleaned, and a new batch culture was started (Schipp et al. 1999). Temperature and salinity Temperature plays a fundamental role in the life of the copepods, but their ability to adapt to temperatures even beyond their natural range is remarkable. Geographical populations are genetically adapted to the conditions of their natural habitat. Estuarine species are more tolerant to lower salinities and temperate species to temperature changes. Coastal species have wider thermal and salinity tolerances than oceanic species (Bradley 1986; Mauchline 1998). In culture, it is preferable to choose species with similar thermal–salinity optima to those present in the rearing facility, although it is possible to obtain species from one temperature–salinity regimen and adapt them to another, as in the case of A. tonsa (Støttrup 2000). pH Extreme pH values are often observed in eutrophic ponds, outdoor rearing tanks or bag enclosures used in extensive rearing of marine fish larvae. However, few studies have dealt with the influence of different pH levels on copepod development or production. Contamination Contamination of copepod cultures by bacterial blooms, ciliate infections, other copepods or rotifers may pose a problem. In cultures where the bottom is siphoned regularly, use of the same siphon for all copepod tanks should be avoided at all costs, and separate siphons used for each tank to avoid contamination. The same rule stands for other devices (e.g. sampling devices) that may be used in each tank. In commercial facilities, contamination by rotifers is the most likely cause of the collapse of a copepod culture, since the rotifers with their higher reproductive rate would quickly outcompete the copepods. It is therefore important to keep these cultures strictly apart. The presence of other copepods may pose a problem, although the presence of the harpacticoid Tisbe furcata in a culture of Acartia clausi helped to increase the calanoid production, ensuring a natural cleaning of the tank (Person-Le Ruyet 1975). From experience gained from extensive copepod production, it is clear that certain species of calanoid often become dominant. These species are probably the most suitable candidates for culture, such as species belonging to the genera Acartia, Eurytemora, Centropages or Temora. Ciliates are utilised by copepods and may in periods of low phytoplankton concentrations constitute the major dietary source (Poulet 1983; Mauchline 1998). In laboratory

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studies, ciliates were demonstrated to be nutritionally adequate, although the clearance rates on ciliates decreased with increasing phytoplankton concentrations (Stoecker & Egloff 1987). In intensive cultures, the presence of certain ciliates, such as Euplotes species, is often an indication of overfeeding and should be avoided. This hypotrichid ciliate is a common contaminant of live prey cultures and is easily identified under a light microcope. Should ciliates be observed, it is advisable to empty the culture using a 60 or 80 ␮m mesh size gauze, which retains the adult copepods, but allows the ciliates to be washed out. The culture can then be started afresh. Cultures may succumb to uncontrolled proliferation of bacteria, even though bacteria often constitute a part of the diet of copepods. Bacteria-colonised faecal pellets or detritus have been found in the guts of calanoids and some of the ingested bacteria were absorbed and nutritionally utilised (Mauchline 1998). It was suggested that bacteria colonising the rice bran might also have served as food in the culture of A. tonsa on this inert diet (Turk et al. 1982). Takano (1971) also speculated that the bacteria on the cereal fed to G. imparipes might have been the major nutritional source. Some bacteria, such as Vibrio sp., are known to infect copepods in eutrophic coastal waters, resulting in lower survival rates (Nagasawa & Nemoto 1986). Calanoids are sensitive to high ammonia concentrations. Ammonia concentrations of 0.12 ppm resulted in an increase in egg production but negatively affected egg viability in the calanoid A. clausi (Buttino 1994). Payne and Rippingale (2000c) found higher concentrations of ammonia and nitrate in cultures of G. imparipes, but the extent to which this effected fecundity was not examined. Submerged air-lift foam filters maintained good water quality in 500 and 1000 litre cultures of G. imparipes through rapid removal of faecal pellets (Payne & Rippingale 2000c). Water renewal in batch systems or water flow-through in continuous systems (Støttrup & Norsker 1997), or continuous water recirculation (Sun & Fleeger 1995) also helps to reduce accumulation of contaminants. Care should be taken concerning materials used to rear copepods in the laboratory. Because of their high sensitivity to a great variety of chemicals, natural or anthropogenic, copepods are used in toxicity tests to determine acute and sublethal effects of water-soluble chemicals and effluents, and in bioaccumulation studies (Toudal & Riisgård 1987; Kusk & Wollenberger 1999; Hook & Fisher 2001). Copepods are sensitive to pesticides, heavy metals and chlorinated compounds. Copepod populations are drastically reduced when exposed to rotenone, a pesticide commonly used to kill parasites or predators before rearing marine fish larvae (e.g. cod) in extensive systems (Næss 1991). Heavy metals such as cadmium or copper and chlorinated compounds are toxic to copepods (Toudal & Riisgård 1987; Mauchline 1998). Fungicide-free silicone produced for aquarium purposes should be used and corrosive materials avoided in a saltwater laboratory. Harvest, storage and transport The zooplankton are hardy enough to withstand repeated filtering, although submerged filters should be used at all times. Even when assorted sizes are used simultaneously, the filters should be constructed such that all are submerged during the filtering process. Copepods can survive for short periods on the gauze as they are transferred from one tank to another or when refilling a tank. They can also survive for an extended time (a few hours) at very high densities, providing there is sufficient oxygen. Freely spawned calanoid eggs sink to the


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bottom and can be harvested by siphoning the bottom once daily. The day’s production can be transferred to individual hatching tanks and grown to the required size to feed as live feed for fish larvae. Although the first naupliar stage does not feed in most calanoids, it is usually of so short a duration that phytoplankton can be added to the hatching tank once hatching has commenced. The method for harvest is different for copepods with egg-bearing females, since only the nauplii can be separated. Methods for harvesting nauplii are described by Payne and Rippingale (2000c), and include manual collection of nauplii using buckets with meshed sides or automatic collection of nauplii using light to concentrate the nauplii. Eggs from the daily collections can be concentrated, transferred to small vials, deoxygenated and sealed before being stored in a refrigerator for weeks to months. The eggs can be transferred directly from storage at 4°C to the hatching tanks at 16–18°C. The viability of the eggs decreases with storage time, although no attempts were made to quantify this (J.G. Støttrup, unpublished data). Kept cool, these eggs can be transported to another site or stored for later use. Newly hatched nauplii that had been cold-stored as eggs for a period of 12 weeks had lower DHA levels than and similar EPA levels to nauplii from newly spawned eggs (Støttrup et al. 1999). The DHA:EPA ratio had decreased from 2.3 to 1.7, which is still within the range found in wild zooplankton. Khanaichenko (1998) registered hatching rates down to 55% in batches stored at 4°C, compared with 96% for newly spawned eggs of Acartia sp. Eggs were also transferred directly from the storage temperature (4°C) to 20–24°C. 5.3.2.2 Harpacticoids Harpacticoids have been cultured in batch and continuous systems to provide food for marine fish larvae, and several studies have demonstrated improvements in growth and survival when using harpacticoids either as the only food source or as a supplement to traditional feeds (Lee et al. 1981; Heath & Moore 1997). Several workers have declared harpacticoids, in particular species of the genera Tisbe or Tigriopus, to be ideal candidates for cultivating in large cultures (Rothbard 1976; Fukusho 1980; Uhlig 1984). Fukusho (1991) found that the harpacticoid Tigriopus japonicus was the only species to be successfully mass cultured from a list of 11 different species examined, including species of the genera Acartia, Eurytemora and Pseudodiaptomus (Calanoida) and species of the genera Oithona (Cyclopoida). The aim was to produce high-density cultures and highest densities were obtained in extensive co-cultures of T. japonicus and rotifers. There are several advantages in using harpacticoids in culture:

• • • • • • • •

high tolerance to a wide range of environmental conditions (Lee & Hu 1981; Uhlig 1984) ability to feed on a wide range of live or inert diets (Uhlig 1984) high reproductive capacity (Uhlig 1984) relatively short life cycles (Tisbe spp. 7–29 days, Tigriopus spp. 12–21 days, Schizopera elatensis 8 days) (Hicks & Coull 1983; Uhlig 1984) ability to be cultured in high densities (Uhlig 1984) requirement for surface area rather than volume (Uhlig 1981; Støttrup & Norsker 1997) planktonic naupliar stages (Hicks & Coull 1983; Jinadasa et al. 1991) can be used as tank cleaners in rotifer cultures, other copepod cultures or larval tanks (Person-Le Ruyet 1975; Støttrup & Norsker 1997).

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Many of the techniques described for calanoids can be used to culture harpacticoids and cyclopoids. Only the points where they may differ, or peculiarities to these genera are dealt with in the following sections. Criteria for culture conditions for harpacticoids are less demanding than those for calanoids. Filtered, artificial or non-treated seawater may be used, and a whole range of inert food is acceptable to many harpacticoid species (see Table 5.5). This simplifies the culture method and eliminates the need for cultures of phytoplankton. Tisbe species, with their small nauplii (T. gracialis: NI 73 ␮m, NVI 207 ␮m, Dahms & Bergmans 1988; T. cucumariae: NI 72 ␮m, NVI 186 ␮m, Dahms et al. 1991), are adequate as a starter feed for small marine fish larvae. Crowding affects both female productivity and egg viability in T. holothuriae (Zhang & Uhlig 1993). Daily naupliar yield was highest when females were kept at a density of 40 cm⫺2. The best density for naupliar production was 180 individuals cm⫺2, resulting in around 30% larval mortality (Zhang & Uhlig 1993). Food and feeding If algae are readily available, a mixture of two algal species would be the preferred choice. Algae, which quickly sediment, are very appropriate for benthic copepods, possibly because bacteria colonise these cells, and the mixture of algae and bacteria may be a superior dietary combination for harpacticoids. Algae such as Skeletonema costatum, Rhodomonas baltica and Tetraselmis suecica quickly sediment, whereas species such as Isochrysis galbana remain in suspension and may be less available to the harpacticoids. Harpacticoids can be reared on a variety of inert feed. However, as discussed earlier, food quality affects development and fecundity, and should be considered carefully. The use of inert feed may cause hygiene problems in the culture tank. A mixed diet may provide the best nutritional value (Zhang & Uhlig 1993). Sun and Fleeger (1995) fed Amphiascoides atopus either the phytoplankton Chaetoceros muelleri, commercial fish flakes or a mixture of algae and commercial fish flakes, with equal success. The protein content of the diet affected both the development time and fecundity in Tisbe cucumariae (Guidi 1984). This may not be valid for all harpacticoids, but it may be appropriate to ensure a diet containing around 50% protein for most harpacticoids. Although HUFA do not seem to affect fecundity or the content of HUFA in the offspring, a diet containing HUFA is recommended until conclusive evidence on the role of HUFA in harpacticoid diets has been provided. The addition of vitamins (vitamin D2) to inert food was shown to improve fecundity (Guérin et al. 2001). Light Photoperiod influences both offspring production and their sex ratio. This was demonstrated in experiments comprising three successive generations of T. holothuriae (MoraitouApostolopoulou et al. 1982). A photoperiod of 12 L/12 D was shown to be most favourable for offspring production, and both photoperiod and continuous dark conditions resulted in 40–48% females among the offspring. Continuous light was the least favourable, producing the least number of offspring and lowest percentage of females (20–23%). A similar reliance on photoperiod for the synchronisation of egg production was found in Euterpina acutifrons in tropical waters (Hopcroft & Roff 1996).

Table 5.5 Cultivation techniques for different harpacticoid species.

Species cultured

Duration of culture

Amphiascoides atopus

17 weeks

Euterpina acutifrons (pelagic)

⬎1 year


⬎90 days

Euterpina acutifrons Euterpina acutifrons

Several generations


Food organisms

T/S


Densities obtained

Chaetoceros muelleri ⬎1 ⫻ 106 cells ml⫺1 2 day⫺1, or mixture of algae ⫹ 20 g commercial fish flakes 2 day⫺1

23–26°C/ 30–34 psu

1440 litres recirculation

18°C

3 litres

500 adults l⫺1

Neunes & Pongolini (1965)

Dunaliella salina, Platymonas suecica, Chroomonas fragarioides, Phaeodactilum tricornutum, Chlorella ovalis, Glenodinium sp.

21–24°C

600 litre batch 15 day cycle inoculated with 50 litres

8900 ind l⫺1

Alessio (1974)

Flagellates, diatoms, Peridineae

18°C/37– 38 psu

Algae or yeast

22°C

Chaetoceros gracilis

22–23°C/ 34 psu

50 litres

Harvest

Reference

0.5 ⫻ 106 ind day⫺1 for 10 weeks, then 2–4 ⫻ 106 day⫺1

Sun & Fleeger (1995)

356 eggs F⫺1

Zurlini et al. (1978) Ben-Amotz et al. (1987) Szyper (1989)

Nitocra spinipes

56 days

Chlorella 1–3 ⫻ 106 cells ml⫺1 or shrimp head meal

28–32°C/ 25 psu

10 litres ⫹ aeration

11 ind ml⫺1

Gopalan (1977)

Schizopera elatensis

⬎55 days

Lettuce leaves

25°C/ 35 psu

0.5–6 litres, static

412 ind ml⫺1

Kahan (1981)

Schizopera elatensis

13–21 days

Mytilus powder 5–150 mg, lettuce leaves

21–22°C/ 40 psu

1.5 litre baskets in 200 litres

29 ind ml⫺1

Kahan et al. (1982) (continued)

Table 5.5 (continued) Duration of culture

Food organisms

T/S


Scottolana canadensis


Isochrysis galbana, Thalassiosira pseudonana weekly

20°C, 25°C

2 litres, batch

Tigriopus japonicus

1 month

Chlorella and soya cake

22–25°C

Tigriopus japonicus

2 months

Artificial fish feed

Species cultured

Densities obtained

Harvest

Reference

10 eggs F⫺1 day⫺1

Harris (1977)

150 F with eggs l⫺1 ⫹aeration

Tigriopus japonicus

26 days

3 g crushed seaweed, Ulva petrusa, once weekly

22°C

2 litres semicontinuous ⫹ aeration

Tigriopus japonicus

Several generations

Green algae or artificial foods

15.6°C/ 36 psu

Tigriopus japonicus co-cultured with rotifers

89 days

Chlorella minutissima, baker’s yeast, ␻-yeast

Ambient, 5–25°C

200 m3 in outdoor ponds

Tisbe furcata

1–3 generations

Seaweed, dried mussels and scallops, faecal pellets of Nereis

17–21°C

50 ml

Tisbe furcata

⬎90 days

Dunaliella salina, Platymonas suecica, Chroomonas fragarioides, Phaeodactilum tricornutum, Chlorella ovalis, Glenodinium sp.

21–24°C

600 litre batch 15 day cycle inoculated with 50 litres

Kitajima (1973)

3500 ind l⫺1 ⫺1

⬃1000 F l

Kitajima (1973) ⫺1

26,000 N l

Rothbard (1976)

Takano (1968) in Lee & Hu (1981) —

1.9 kg day⫺1

Fukusho (1980)

513 eggs F⫺1

Johnson & Olson (1948)

6700 ind l⫺1

Alessio (1974)

Tisbe holothuriae

Tetramin 1 g day⫺1 for 30 days, then 2 g day⫺1

100 litres recirculation 12 l h⫺1

20 ind l⫺1 day⫺1

Gaudy & Guerin (1978)

Tisbe holothuriae

Tetramin 1 g day⫺1 for 30 days, then 2 g day⫺1

100 litres ⫹ 10 m2 surface area, recirculation 12 l h⫺1

70 ind l⫺1 day⫺1

Gaudy & Guerin (1978)

Kahan et al. (1982)

Tisbe holothuriae

13–21 days


21–22°C/ 40 psu


Tisbe holothuriae


Rhodomonas baltica, 400 ml added daily with water renewal

18°C/ 34 psu

3 litres in flat trays 40 ⫻ 60 cm, batch

100,000 N l⫺1 day⫺1


Tisbe holothuriae


Rhodomonas baltica 1 ⫻ 106 cells ml⫺1, continuous system, 20 l day⫺1

18°C/ 34 psu

150 litres filled with balls, continuous

1533 N and 1800 C l⫺1 day⫺1


Ulva fragments ⫹ Dunaliella, Pheodactylum or Nitzschia and boiled wheat grain fragments

Tisbe reticulata

Tisbe sp.

13–21 days

Tisbe sp. Tisbe sp. Tisbe sp.

16 days

20 cm3

Battaglia (1970)


21–22°C/ 40 psu


115 ind l⫺1

Granulated Mytilus edulis

28 psu

Floating sieves

25 F cm⫺2

0.5 g Microfeast L-10® twice per week

20°C

1 adult ⫹ 31 N ⫹ C ml⫺1

Tetraselmis suecica, Isochrysis galbana tahiti, Dunaliella tertiolecta, Rhodomonas baltica and two algal pastes

32 litres, airlift pump 70 litres ⫹

Bioballs©

T, temperature; S, salinity, ind, individuals; F, females; N, nauplii; C, copepodites.

Kahan et al. (1982) 10 N F⫺1 day⫺1

Uhlig (1984) Nanton & Castell (1997) Coli (2000)

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Light was used to harvest selectively harpacticoid nauplii from an automated continuous culture system, which was allowed to function for approximately 1 year (Støttrup & Norsker 1997). The copepodite and adult harpacticoids were negatively phototactic and were observed to swim energetically away from a sudden light source, whereas the nauplii did not seem to be affected (or were poorer swimmers) and remained in the water column close to the outlet where the harvest took place. A strong light source was thus switched on for a few minutes before the daily harvest. This form of harvest was not totally effective. Both copepodites and nauplii were harvested daily and sufficient nauplii would remain in the culture unit to supplement continuously the breeding population. Aeration Batch or continuous systems used for cultures of harpacticoids generally lack mechanical aeration. Nanton (1997) used an air-lift pump, a very effective yet gentle mode of aeration and circulation within the culture system. Thus, some aeration may be applied to maintain an even distribution of food and zooplankton, but may be less important in recirculation systems where the water volume is replaced more than once daily, or in batch systems where the whole water volume is filtered and replaced daily or every 2 days. Culture tank size and shape The mass culture of the benthic harpacticoid Tisbe spp. is dependent on the available surface area rather than culture volume (Uhlig 1981). In the batch system described by Støttrup and Norsker (1997), an average of 300,000 nauplii day⫺1 was produced from four trays measuring 40 ⫻ 60 cm and filled with around 3 litres of filtered (0.22 ␮m) seawater. This corresponds to a daily volume output of 100,000 nauplii per litre. In continuous or recirculation systems, the units are filled with small balls, or other material, which provide surface area for the copepods (Gaudy & Guerin 1978; Støttrup & Norsker 1997; Coli 2000). Very successful mass culture of the harpacticoid Amphiascoides atopus was achieved in a continuous system with a relatively small basal surface area (4 m2) and a water volume of 1440 litres (Sun & Fleeger 1995). During a 5 month harvest period this system averaged a production of 0.5 million individuals day⫺1. Temperature and salinity Most harpacticoids have wide thermal and salinity tolerances, reflective of the variable environment they inhabit. The harpacticoid Tigriopus japonicus is very tolerant to salinity changes and can survive a change from 36 to 1.8 psu, although it is doubtful that this species could be reared at this low salinity (Lee & Hu 1981). Amphiascoides atopus also possesses a wide thermal and salinity tolerance range and survived salinities from 10 to 60 psu (Sun & Fleeger 1995). Contaminants The ammonia concentration in high-density cultures of T. holothuriae varied from 1.2 to 1.8 ppm after a feeding event (Støttrup & Norsker 1997). However, the effect of ammonia levels on fecundity was not examined. Harpacticoid cultures contaminated with rotifers tend to be outcompeted by the rotifers. Ciliates may also compete for food and be detrimental


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to the culture performance. Ciliate and rotifer free cultures were obtained by exposing fecund females to weak doses (⬍1%) of chlorine for a few minutes (N.H. Norsker, personal communication). The egg sacs would separate from the females but remain viable, and would hatch upon being washed and transferred to normal medium. Adult copepods, ciliates and rotifers would not survive. As in the case of calanoids, harpacticoids are sensitive to a variety of chemicals and are used in bioassays, for example to examine toxic substances in sediments (Chandler & Green 1996). Harvest, storage and transport Since harpacticoids are not free spawners, harvest methods for collecting nauplii need to be developed. A description of the manual harvest of harpacticoid nauplii is given by Støttrup and Norsker (1997), and is similar to that used by Payne and Rippingale (2000c) for the calanoid G. imparipes. Although the Tisbe sp. used in the laboratory was robust, several species may lose their egg sacs during collection, and possibly depending on when during development the egg sac is lost, embryonic development may be arrested. An automated system, using light to concentrate the nauplii, was used for a continuous culture system for T. holothuriae (Støttrup & Norsker 1997). Although not a very efficient collecting system, since many nauplii would mature within the system and several copepodites were harvested daily, it was a relatively reliable system, needing little labour. Kahan et al. (1982) solved the problem of harvesting nauplii for feeding fish by growing the harpacticoids in floating baskets within the fish larval tanks. The basket had bottom sieves through which the nauplii could pass, but the adults were retained. Harpacticoids are relatively tolerant to high stocking densities and can be transported for a period of up to 2–3 days, kept cool in blood-transfusion bags (2 litres) at densities of up to 200,000 individuals per litre (N.H. Norsker, personal communication). Excess nauplii can also be stored at 4°C for up to 1 week and used on days when the production output is below the required amount. 5.3.2.3 Cyclopoids Very few cyclopoid species have been reared in the laboratory (Table 5.6). From the available information, Oithona spp. or Apocyclops spp. appear to be the best candidates and they are relatively easy to culture over several generations in the laboratory (Chang & Lei 1993). Oithona spp. are an ideal supplement to the traditional live feed for striped patao, Eugerres brasilianus (Alvarez-Lajonchère et al. 1996). A private hatchery in Taiwan uses the copepod Apocyclops royi as live feed for grouper larvae (Su et al. 1997), and A. borneoensis was a suitable replacement for Artemia in rearing Acanthopagrus cuvieri (James & Al-Kars 1986). Photoperiod As in the two other genera, photoperiod affects egg production in cyclopoids. This was demonstrated by Hopcroft and Roff (1996) in four species of cyclopoid copepods sampled from tropical waters around Jamaica, West Indies. The egg sacs were laid down at dawn and the nauplii hatched at dusk on the following day. Chang and Lei (1993) reported great ease in culturing A. royi in the laboratory using 600 lux and a 16 L/8 D photoperiod.

Table 5.6 Cultivation techniques for different cyclopoid species.

Species cultured

Duration of culture

Food organisms

T/S


Apocyclops borneoensis

60 days

Marine yeast, Candida sp.

28°C/20 psu

15 m3

Apocyclops dengizicus

120 days

20–25°C/ 0.5–68 psu

200 ml

Apocyclops panamensis

77 days

Apocyclops royi

Several generations Tetraselmis chuii ⬎105 cells ml⫺1

Isochrysis galbana 5 ⫻ 105 cells ml⫺1 30 psu

T, temperature; S, salinity; ind, individuals; N, nauplii; C, copepodites; F, female.

25°C/30 psu

100 litres 400 ml

Densities obtained 2300 adults l⫺1; 4400 ind l⫺1

Harvest

Reference

2.75 ⫻ 106 day⫺1

James & Al-Khars (1986)

Dexter (1993) 10.7 N ⫹ C F⫺1 4–9 days⫺1

Lipman (2001) Chang & Lei (1993)


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Aeration and oxygen Gentle turbulence is important for most cyclopoids and increases food encounter rates (Kiørboe & Saiz 1995), but as for most copepods vigorous aeration should be avoided. pH The cyclopoid Cyclops vernalis is very tolerant to low pH and can survive for several hours at pH 3.6 (Bulkowski et al. 1985). Contamination Cultures of C. vernalis could only be maintained in a Daphnia-free environment, since Daphnia easily outcompetes C. vernalis (Bulkowski et al. 1985). However, exposing the culture to low pH (3.8) for up to 7 h selectively killed off the Daphnia, whereas around 30% of the cyclopoid survived. Harvest Some cyclopoids are sensitive to handling and fecund females may lose their egg sac, as shown in Oithona sp. (Hopcroft & Roff 1996).

5.4 Biochemical Composition Båmstedt (1986) provides a comprehensive review on the chemical and energy content in pelagic copepods. Water constitutes about 82–84% (modal value; range 67–92%) of the copepod wet weight, and total organic matter 70–98% or more of the dry weight (Båmstedt 1986). The energy content ranges from 9 to 31 J mg⫺1 dry weight, being generally lowest in species from low and medium latitude, and higher in those from high latitudes.

5.4.1 Carbon Carbon content in calanoids varies from around 28 to 68%, with modal values of 40–46% dry weight (Båmstedt 1986). Species from colder regions generally contain higher carbon levels than temperate, subtropical and tropical species (Ikeda 1973). Carbon:Nitrogen (C:N) ratios are generally between 3 and 4, especially in low and medium latitudes. Hydrogen content is low, ranging from 3 to 10% of body dry weight, mostly associated with lipid content. The content of phosphorus rarely exceeds 1% (Båmstedt 1986). In the harpacticoid T. holothuriae, carbon values for females are around 40% of body dry weight, with lower values in males and higher content in egg sacs (Guérin & Gaudy 1977; Zhang & Uhlig 1993). Hydrogen content is also less than 10% (Zhang & Uhlig 1993) and C:N ratios vary between 5 and 12, depending on the diet provided; the variation is due primarily to changes in carbon content (Guérin & Gaudy 1977). When fed an algal diet, Nannochloris sp., this ratio was around 5. Crowding affected dry weight and contents of carbon, nitrogen and hydrogen in the adults and offspring (Zhang & Uhlig 1993).

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5.4.2 Lipids Lipid content in marine pelagic copepods varies with latitude, season and food availability, with a range of 2–61% in low- and medium-latitude species, and 8–73% in high-latitude species (Båmstedt 1986). Most of the low- and medium-latitude species have a lipid content of 8–12%. This author attributes the need to store lipids to maintain fecundity in times of food shortage in high-latitude species as an explanation why these copepods are generally more energy rich than those in low latitudes. The lipid stores also serve as a buoyancy aid (Sargent & Henderson 1986; Hagen 1988). Lipid storage, utilisation and transformation in marine food chains have received particular attention over several decades, and a review of lipid biochemistry in copepods is provided by Sargent and Henderson (1986). Lipid levels are often high in newly hatched nauplii owing to residual lipid stores, but these are soon used and lipid levels fall in the later nauplii and early copepodite stages. The predominant lipids during these stages are structural phospholipids (Sargent & Henderson 1986; Sargent & Falk-Petersen 1988). These phospholipids have negligible levels of 14:0, 20:1 and 22:1 fatty acids, but are rich in n-3 polyunsaturates. In the late copepodite stages lipids are accumulated in the form of wax esters or triacylglycerol (Kattner & Krause 1987; Hagen 1988), which fuel reproduction, especially in situations of food limitation (Sargent & Henderson 1986). Similarly, the classes of lipid differ in eggs from different species. The major neutral lipid in Calanus sp. eggs is triacylglycerols (Gatten et al. 1980), whereas that in Euchaeta norvegica eggs is wax esters (Lee et al. 1974). The classes of stored lipid can also change seasonally within a species. Lipids are stored during the autumn in the later copepodite stages and adults of many species. They are stored in the form of wax esters in Pseudocalanus acuspes or in the form of triacylglycerols in Acartia longiremis (Norrbin et al. 1990). Calanoid wax esters are characterised by 20:1n-9 and 22:1n-11 fatty alcohols and high levels of 18:4n-3, EPA (20:5n-3) and DHA (22:6n-3) fatty acids (Sargent & Henderson 1986; Sargent & Falk-Petersen 1988). These n-3 polyunsaturates may constitute up to 40% of the total fatty acids in the wax esters. Wax esters may make up to 90% of the total lipid in calanoids and are contained in an oil sac, which runs parallel to the gut. Herbivorous calanoids were observed to have higher lipid content and higher content of wax esters (Sargent & Henderson 1986). In non-calanoid zooplankton such as euphasiids, shorter chain fatty alcohols such as 16:0 and 14:0 dominate (Sargent & Falk-Petersen 1988). The protein and lipid content in female T. holothuriae (Mediterranean) examined by Miliou et al. (1992) was 71% and 10% of the dry weight, respectively, with little difference relating to whether or not they were carrying egg sacs. Lipid content is therefore within the range normally found in pelagic species occurring at low and medium latitudes (Båmstedt 1986). A similar pattern of lipid content was observed in calanoids, with a decrease towards early copepodite stages and an increase in late copepodite and adult stages (Miliou et al. 1992).

5.4.3 Protein Protein contents in marine pelagic copepods range from 24 to 82% dry weight and are highest in species from medium latitudes (Båmstedt 1986). In the harpacticoid T. holothuriae, protein content was 71% of the dry weight (Miliou et al. 1992).


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5.4.4 Free amino acids Free amino acids, primarily used in osmotic regulation, generally increase in level with increasing environmental salinity (Båmstedt 1986). Glycine, alanine, arginine, lysine, proline and taurine are quantitatively the most important.

5.4.5 Vitamin C Copepods, in particular omnivorous and herbivorous species, contain high levels of vitamin C, and levels ranging from 201 to 235 ␮g g⫺1 were reported in nauplii of Acartia clausi and Temora longicornis (Hapette & Poulet 1990). Vitamin C is known to stimulate reproduction in crustaceans and is suggested to induce reproduction in copepods (Hapette & Poulet 1990). Thus, copepods may be an important source of vitamin C in fish.

5.4.6 Carotenoids In a study involving over 80 species of calanoid, Fisher et al. (1964) found the predominant and possibly the only carotenoid in most species to be astaxanthin or its esters, with concentrations ranging from trace levels to 1133 ␮g g⫺1 wet weight. In a more recent study on T. longicornis, two carotenoids were evident in high quantities, with lutein present at almost four times the level of astaxanthin (Rønnestad et al. 1998). These carotenoids were not detected in Artemia, which contained primarily cryptoxanthin/canthaxanthin and an unknown retinoid component.

5.4.7 Chitin Chitin content in marine copepods ranges from 2.1 to 9.3% dry weight (Båmstedt 1986). There is an increasing interest in chitin production, in keeping with the increasing industrial utilisation of chitin and its derivative, chitosan.

5.4.8 Enzymes Levels of endoproteases, exoproteases, amylase, esterase and phosphodiesterase were high in copepods (adult Eurytemora hirundoides) (Munilla-Moran et al. 1990).

5.5 Nutritional Value for Fish Larvae Improved growth, survival and/or rates of normal pigmentation have been documented for several marine fish species fed copepods alone or as a supplement to the traditional diets of rotifers or Artemia nauplii compared with traditional diets alone (Kraul 1983; Heath & Moore 1997; McEvoy et al. 1998; Næss & Lie 1998; Nanton & Castell 1999). In many hatcheries, malpigmentation of the reared juveniles constitutes a major problem. Larval nutrition is suggested to be the major factor determining pigmentation patterns,

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possibly enhanced by suboptimal or stressful rearing conditions (McEvoy et al. 1998). Flatfish larvae fed natural or laboratory-reared zooplankton generally exhibit higher rates of normal pigmentation than do larvae fed Artemia nauplii (e.g. Seikai et al. 1987, Japanese flounder; Næss et al. 1995, Atlantic halibut; McEvoy et al. 1998, Atlantic halibut). Growth and survival was higher in seahorses, Hippocampus subelongatus, fed copepods compared with Artemia nauplii (Payne & Rippingale 2000b). A combination of rotifers and copepod nauplii increased growth and survival in larvae of turbot, Psetta maxima (Støttrup & Norsker 1997), dhufish, Glaucosoma hebraicum, and pink snapper, Pagrus auratus, compared with rotifers only, although the results for the latter species were not statistically significant (Payne et al. 2001). The documented improvements in larval growth, survival and rates of normal pigmentation are generally attributed to levels of DHA, EPA and/or arachidonic acid (ARA) in the diet (Castell et al. 1994; Reitan et al. 1994; Zheng et al. 1996; Sargent et al. 1997), and in particular to the DHA:EPA ratio in the diet (Bell et al. 1995b; Sargent et al. 1997; Nanton & Castell 1998) and EPA:ARA ratio (Bell et al. 1995a; Sargent et al. 1997; Estevez et al. 1999). DHA can be synthesised from shorter chain precursors in some marine fish larvae, but at rates insufficient to meet requirements for their normal growth and survival (Sargent et al. 1989; Bell et al. 1995b). A minimum of 0.5–1.0% of the dry weight as n-3 HUFA is required for juvenile marine fish larvae and higher amounts are probably required for the rapidly growing fish larvae (Sargent et al. 1997). Le Milinaire et al. (1983) estimated a minimum n-3 HUFA requirement of 1–3% dry weight for turbot larvae. However, the proportion of different lipids may be an important factor, since n-3 HUFA levels of 1.2% (Gatesoupe & Le Milinaire 1984), 1.5% (Støttrup & Attramadal 1992) or 2.2% (Støttrup 1992) did not improve growth in turbot larvae. Marine copepods, the principal diet for most marine fish larvae in nature, contain high levels of DHA and other PUFA, either obtained through their phytoplankton diet or accumulated despite low PUFA levels in the diet (see Section 5.2.6.2). DHA levels in wild copepods can be more than 10 times higher than in enriched Artemia (McEvoy et al. 1998). Using special emulsions, it is possible to boost DHA levels in both Artemia nauplii and rotifers to levels equivalent to zooplankton (Reitan et al. 1994), but by the time these prey are consumed by the fish larvae n-3 PUFA levels may be reduced. Rotifer residence time in larval fish tanks is crucial since n-3 HUFA levels decrease with time (Reitan et al. 1994) and retroconversion of DHA to EPA takes place in Artemia nauplii (Navarro et al. 1999) and probably in rotifers (Barclay & Zeller 1996). The ratio of DHA to EPA in calanoids and harpacticoids is generally ⬎2 (Nanton & Castell 1998, Tisbe sp.; McEvoy et al. 1998, wild calanoid zooplankton; Støttrup et al. 1999, A. tonsa nauplii from adults fed Rhodomonas baltica, Heterocapsa triquetra or Isochrysis galbana), and approximately 2 in the yolks of wild halibut eggs (Parrish et al. 1994). Reitan et al. (1994) found a positive correlation between DHA:EPA ratios in 12- and 22-day-old turbot larvae and the rate of normal pigmentation at 27 days. DHA is important in maintaining structural and functional integrity in fish cell membranes, in neural development and function, and especially in retinal development and vision (Bell & Tocher 1989; Bell & Sargent 1996). Dietary deficiency of DHA was shown to impair vision at low light intensities in juvenile herring, Clupea harengus (Bell et al. 1995b). It is suggested to play an important role in the development of normal pigmentation when provided in


193

sufficient quantities at particular times during the larval stage (Reitan et al. 1994). Næss and Lie (1998) showed that malpigmentation could be avoided by providing copepods for a short period to halibut larvae before reaching 2.5 mm mytome height. Like DHA, EPA cannot be synthesised by most marine fish – or not at a rate to satisfy demands, especially during early development – and it is therefore essential in the diet of the fish. EPA gives rise to less biologically active eicosanoids than those produced from ARA. Since it competes metabolically for the same enzyme systems required for ARA-derived eicosanoid production, EPA is very important in modulating the production of these highly biologically active eicosanoids. This metabolic interaction necessitates an optimal EPA:ARA ratio in the diet (Bell et al. 1995a). Eicosanoids of n-6 origin are important for the normal function of vital organs such as the kidney, gill, intestine and ovaries of marine fish. Levels of ARA in copepods are high (⬎1%) in both calanoids and harpacticoids, and in the latter are independent of the dietary content. Nutritional benefits of feeding copepods (in terms of improved larval fish growth, survival and frequency of normal pigmentation) seem to lessen with improved knowledge on husbandry and optimal rearing conditions for a particular species. Both ARA- and EPAderived eicosanoids are involved in the physiological reaction to stress and it may be that the optimal EPA:ARA ratios found in copepods allow the larval fish to cope better with stressful conditions (McEvoy & Sargent 1998). When improvements in husbandry remove these stressful conditions, the benefits of the optimal EPA:ARA ratio may become less apparent. Apart from the superior fatty acid composition in copepods compared with Artemia nauplii or rotifers, even after enrichment, copepods contain high amounts of polar lipids (Fraser et al. 1989). Levels of total polar lipids in wild calanoids were almost twice those in enriched Artemia (McEvoy et al. 1998). In A. tonsa adults fed different monoalgal diets and their newly hatched nauplii, total polar lipids averaged 46% and 47% of total lipids, respectively (Støttrup et al. 1999). Polar lipids are more readily digested by larvae and may also facilitate digestion of other lipids in the undeveloped gut of marine fish larvae (Koven et al. 1993). Thus, larval fish may more easily assimilate DHA and other essential fatty acids in copepods than in Artemia, where these essential fatty acids are present mainly as neutral lipids (McEvoy et al. 1998). The assimilation of wax esters is a highly efficient process in fish, even though not all of the dietary wax esters are utilised (Sargent & Henderson 1986). Long-chain monoenoic fatty acids can be exploited as metabolic fuel (catabolised) in fish, facilitated by the presence of n-3 PUFA. Varying concentrations of the carotenoid astaxanthin were found in various copepods and it was suggested that its possible value for fish is as a precursor to vitamin A (Fisher et al. 1964). The difference in carotenoid pigments between copepods and Artemia nauplii was suggested to be a possible explanation for the poor development of normal pigmentation in marine fish larvae, since canthaxanthin is not frequently encountered in the natural environment and possibly may not be easily assimilated by marine fish larvae (McEvoy et al. 1998). This was confirmed by Rønnestad et al. (1998). After 14 days of feeding on Artemia, halibut larvae contained 50–80% lower vitamin A (retinol and retinal) than those fed zooplankton, suggesting that these fish larvae were unable to convert the available carotenoids (canthaxanthin) present in Artemia. These authors cautioned against adding vitamin A to the enrichment emulsion of Artemia, since excess amounts are toxic to fish larvae. PUFA,

194


particularly DHA, are highly prone to auto-oxidation. The high levels of natural antioxidants in copepods protect these PUFA, supplying fish with both PUFA and high levels of natural antioxidants (Sargent et al. 1997). In contrast, antioxidants have to be added to enrichment emulsions for both rotifers and Artemia to prevent auto-oxidation. Copepods are also an important source of exogenous digestive enzymes and are thought to play an important role in fish larval digestion (Munilla-Moran et al. 1990). Marine zooplankton are a valuable source of lipids, essential fatty acids, protein, amino acids, easily assimilated carotenoids, minerals and enzymes. They could be an inexpensive ingredient to replace expensive fishmeal, and an alternative or supplement to Artemia nauplii or rotifers. A strong selection for copepod nauplii was observed in turbot, suggesting that this prey may be more visible, attractive or better suited in size than traditional live prey, in this case rotifers (van der Meeren 1991). In general, all stages of copepods are suitable as food, although there is an indication that copepodite stages of Acartia spp. may be less suitable due to their high escape ability (van der Meeren 1991). Harpacticoid nauplii are suitable prey for fish larvae. They have been reported to predominate within the nauplii fraction of zooplankton samples (van der Meeren 1991; Jinadasa et al. 1991) are readily consumed by, and are nutritious prey for, marine fish larvae (Kraul 1983; van der Meeren 1991; Heath & Moore 1997; Nanton & Castell 1999). In freshwater systems, methods for the production and harvest of zooplankton in large ponds utilising sewage waste have been developed and the zooplankton used as food for marine fish species (Quin 1993). The species composition was not given. However, care should be taken using products derived solely from freshwater zooplankton, since some freshwater zooplankton species may lack essential fatty acids. Watanabe et al. (1978) showed that although Daphnia sp. contained high levels of EPA it was deficient in DHA, and Moina sp. was deficient in both DHA and EPA. The temperature within the systems where the zooplankton is harvested is also important, since PUFA levels in the phospholipids in freshwater copepods increase with decreasing temperature (Farkas 1979).

5.6 Application in Marine Aquaculture The ready availability of Artemia nauplii, through the purchase of cysts and subsequent hatching of their nauplii (see Chapter 3), and the short generation time and relatively uncomplicated culture of rotifers (see Chapter 2) make these live feeds more accessible for marine hatcheries. Indeed, cultures of these organisms have been responsible for the rapid progress over recent decades in the marine aquaculture sector. Because of the rapid expansion and the increasing demand for reliable large-scale production of live feeds, research and development has been focused on enhancing culture techniques or enrichment methods to improve the availability and nutritional value of rotifers and Artemia nauplii, respectively. Little effort has been directed towards developing an adequate, large-scale simple and reliable culture technique for copepods, despite their proven superior nutritional value. With the rapid expansion of the sector and increasing interest in new species and the culture of ornamental species to replace wild fisheries, requirements arise that cannot be met by conventional species. Thus, interest in copepods has been regenerated and the use of


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copepods as live feed in aquaculture is rapidly gaining impetus. They are used as a supplement to traditional live feed, or because they are the only acceptably sized prey for small larvae of ornamental fish species or the only type of live feed that will support the rearing of particular, often new, marine fish species (Doi et al. 1997b; Næss & Lie 1998; Toledo et al. 1999; Payne & Rippingale 2000b; Payne et al. 2001). A shortage in the supply of Artemia cysts and increasing prices (Bengtson et al. 1991) have further encouraged the search for alternative live prey and the renewed interest in using copepod species. A few of the culture methods developed to date are adaptable to mass-culture techniques in commercial hatcheries, or are already in use in commercial hatcheries, although these hatcheries are generally small producers and produce fish for a limited period of the year. For example, the Danish hatchery Maximus A/S produces around half a million turbot juveniles each year in extensive systems based on copepods with supplement of Artemia to cover the energetic demands of the rapidly growing late larval stages. The productive season is limited to around 6–7 months depending on the temperature. Expansion to year-round production would require the inclusion of intensive culture techniques for live prey. Thus, the use of copepods in marine aquaculture is likely to be maintained for low-technology (extensive or semi-extensive) rearing systems and smaller hatcheries operating for a limited period each year in temperate climates, but which may be expanded to year-round production in regions with more favourable climatic conditions and for rearing ornamental fish or fish with a very small mouth size. Unless inert feeds are introduced as a viable alternative to Artemia, the future expansion of marine aquaculture may further encourage work on copepods towards the development of reliable production systems or, alternatively, the production of resting or diapause eggs for sale on a commercial scale.

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Chapter 6

The Microalgae of Aquaculture Arnaud Muller-Feuga, Jeanne Moal and Raymond Kaas

6.1 Introduction Ocean phytoplankton, with a production of several hundred billion tonnes of dry weight per year (Pauly & Christensen 1995), forms the base of the aquatic food chain, contributing to the production of some 100 million tonnes of renewable resources per year from fishing. Hence, it is hardly surprising that the microalgae composing phytoplankton play a crucial nutritional role in marine animal aquaculture, especially for molluscs, shrimp and fish. Aquaculture hatcheries often need to include a microalgal production system and, in the case of marine fish larvae, a live prey production system. Microalgae and cyanobacteria are a major component of the plant kingdom and play a major role in building and maintaining the Earth’s atmosphere by producing oxygen and consuming carbon dioxide. Several tens of thousands of species are ranked in 15 major classes. Although class membership and interrelationships are still heavily debated, a provisional structure based essentially on cytomorphological analysis is presented for this polyphyletic part of the plant kingdom (Fig. 6.1). Further developments in classification are expected in the near future related to the implementation of molecular techniques to distinguish between species. This chapter considers the biological features of microalgae most commonly used in hatcheries as well as their biochemical composition, paying special attention to those compounds beneficial to marine animal nutrition. Finally, microalgal production systems are described and compared with those implemented in hatcheries, particularly in economic terms.

6.2 Biology of Microalgae 6.2.1 General characteristics of microalgae Because of their small size and aquatic existence, microalgae do not require a rigid skeleton for countering the force of gravity in the manner of higher plants. Nevertheless, some classes, such as peridinians and diatoms, exhibit external theca.

The Microalgae of Aquaculture

207

Green algal line

Higher plants Chlorophyceae Charophyceae Prasinophyceae Euglenophyceae Cyanobacteria

Rhodophyceae

Prokaryotes

Eukaryotes

Cryptophyceae Dinophyceae

Brown algal line

Eustigmatophyceae Xanthophyceae Chloromonadophyceae Thraustochytriidae Prymnesiophyceae Phaeophyceae Chrysophyceae Bacillariophyceae Fig. 6.1 Provisional phyletic relationships between algal classes, with an indication of those of importance to aquaculture (shaded boxes).

As for most micro-organisms, reproduction is primarily asexual, although gamete-like forms have been described for certain species. Vegetative reproduction consists of the equal division of the mother cell into two daughter cells, each exactly half the size of the mother cell. The daughter cells sometimes remain within the cell wall of the mother cell until two or three divisions have occurred. Thus, the metabolism of microalgae is directed towards cell division and increase in size, using light as the sole energy source. However, under certain conditions, some algae can utilise organic substrates, in a manner similar to bacteria and fungi. Microalgae have colonised a great variety of environments, from hot-water springs to polar ices, and some species show great resistance to dryness, high salinity, low light, etc. Microalgal metabolism involves numerous biochemical compounds of interest in nutrition, cosmetics and the pharmaceutical industry. They probably exhibit a greater variety of these products than terrestrial plants that have developed from only one of the two lines of the algal evolutionary ‘tree’. Their metabolic plasticity allows species adaptation to a wide range of environmental conditions. For example, when unavailability of nitrogen prevents algae from synthesising the structural proteins required for growth and division, metabolism is directed towards the synthesis of carbohydrate compounds not requiring nitrogen (polymers such as starch and polysaccharides) and of valuable lipids such as carotenoids

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and fatty acids (Sukenik & Wahnon 1991). The metabolic plasticity of microalgae has been extensively studied, especially their ability to synthesise polyunsaturated fatty acids (PUFA) beneficial for animal nutrition. Microalgae have also developed anti-radical mechanisms against activated oxygen species. Like other micro-organisms, they show a capacity to resist competition and predation by synthesising bioactive substances. Some species develop allelopathic activities and should be more suitable for high-density production. Dunaliella tertiolecta and Pyramimonas grossii were clearly found to exhibit allelopathy (Smith 1994). Excretion of high molecular weight carbohydrates and their inhibitory action upon copepod growth has been reported by Malej and Harris (1993). As indicated below, all of these features impact to some extent on the nutrition of marine animals. The first microalgae species used in aquaculture have been selected from those that developed naturally in the marine environment of pioneering aquaculture farms and were probably the easiest ones to cultivate. Subsequently, other collected species were investigated and the most nutritionally efficient were retained. Among the numerous species tested, only a handful is widely used today. Table 6.1 summarises the 16 genera of microalgae most commonly grown for aquacultural purposes. Most of the major microalgae classes are represented in this list. The majority are eukaryotic, with plastidial membranes separating their organelles. However, cyanobacteria of the genus Arthrospira are also included because of the numerous examples of their utilisation as food in aquaculture for both larviculture and ongrowing. Those prokaryotes have photochemical systems I and II as in eukaryotes, but only chlorophyll a instead of a and b (with the exception of eustigmatophytes). Although toxic metabolites of dinoflagellates are frequently harmful for marine life, a representative of this class has recently been proposed as potentially useful in aquaculture. Thraustochytriidae are mentioned here because they are considered as heterokont algae by some authors Table 6.1 Classes, genera and species of major currently named microalgae grown for food in aquaculture, and their main utilisation (synonymous names are in parentheses). Class

Genus

Species

Main utilisation

Cyanophyceae (blue–green algae)

Arthrospira (Spirulina)

platensis, maxima

LPF

Bacillariophyceae (diatoms)

Skeletonema Phaeodactylum Chaetoceros Thalassiosira

costatum, pseudocostatum tricornutum, calcitrans, gracilis, pumilum pseudonana

BM, PS BM, PS BM, PS BM

Chlorophyceae (green algae)

Chlorella Dunaliella Nannochloris

minutissima, virginica, grossii tertiolecta, salina atomus

LPF LPF, PS BM

Prasinophyceae (scaled green algae)

Tetraselmis (Platymonas) Pyramimonas

suecica, striata, chuii virginica

LPF, BM, PS BM

Cryptophyceae

Rhodomonas

salina, baltica, reticulata

BM, PS

Eustigmatophyceae

Nannochloropsis

oculata

LPF

Prymnesiophyceae (haptophyceae)

Isochrysis Pavlova (Monochrysis)

galbana, aff. galbana ‘Tahiti’ (T-iso) lutheri, salina

BM, PS BM, PS

Dinophyceae (dinoflagellates)

Crypthecodinium

cohnii

LPF

Thraustochytriidae

Schizochytrium

sp.

LPF

LPF, food for live preys of fish larvae; BM, food for bivalve mollusc larvae; PS, food for penaeid shrimp larvae.


209

(e.g. Barclay et al. 1994; Lewis et al. 1999). They have also been classified as fungi (e.g. Gaertner 1977; Porter 1990). Depending on salinity, light and nutrient concentrations, the size and appearance of microalgae can vary markedly (Hiramatsu et al. 2000), which makes it difficult to characterise any particular species. Thus, there are some discussions about the classification of some genera, e.g. Spirulina versus Arthrospira (Guglielmi et al. 1993), Chlorella versus Nannochloropsis (Gladu et al. 1995), and of some species, e.g. Nitzschia closterium (Ehrenberg) Wm. Smith, forma minutissima versus Phaeodactylum tricornutum (Wilson 1946) and Skeletonema costatum versus S. pseudocostatum (Medlin et al. 1991). More recently, with the progress in molecular taxonomy, new classes have been proposed for almost all genera in Table 6.1, especially within Chlorophyceae (Marin & Melkonian 1999) and Prymnesiophyceae (Edvardsen et al. 2000). The microalgae of aquaculture have been well described and documented (e.g. Chrétiennot-Dinet et al. 1986). Some web sites provide extensive information on algae and can supply strains (see Appendix IV). Most of the sites providing information on microalgae are listed by the Phycological Society of America (www.psaalgae.org). Microalgae species selected for aquaculture are generally free-living (Table 6.2). All are pelagic and in the nannoplankton range (2–20 ␮m). Those with benthic behaviour have been excluded because they tend to settle on vessel walls and make cleaning difficult. Both diatoms and cyanobacteria cells often remain associated after division, forming chains of cells, so that counting is difficult and cell weight is generally not determinable. This section considers the practical aspects of the biology of microalgal species, especially those relevant to aquaculture.

6.2.2 Growth The growth of a population of micro-organisms, which results from both the increase in size and the division of the cells composing it, is currently indicated by the specific growth rate ␮, defined as dX/(Xdt), where X is the algal biomass concentration and t the time. The growth rate is generally expressed in h⫺1 for heterotrophs, while day⫺1 is more suitable for photoautotrophs, with the relation ␮(day⫺1) ⫽ 24 ␮(h⫺1). A more practical expression of ␮ over a period of time (t ⫺ t0) results from integration: ␮ ⫽ [log(C) ⫺ log(C0)]/(t ⫺ t0), where C0 is the initial concentration at time t0, and C the final concentration at time t. The doubling time ␶ of the biomass of a population is also used to characterise growth in microorganisms and can be expressed as ␶ ⫽ log(2)/␮. Table 6.3 summarises some specific growth rate values obtained for the microalgae species most often used in aquaculture. As no standard is available to define the conditions of measurement of the growth, the data should be considered with due caution. Results are very heterogeneous, even for a given species and a given irradiance. For example, the growth rates of S. costatum vary between 0.84 and 2.88 day⫺1 at 135 ␮Einstein m⫺2 s⫺1 (Curl & McLeod 1961). As shown later, they depend on various culture conditions (e.g. vessel shape and volume, concentration, mixing) and on the moment of measurement. However, an average value of 1 day⫺1 can be maintained for most species, corresponding to a doubling time of about 17 h, which is a common feature of microalgae in photoautotrophy. In rare cases, for example the heterotrophs Crypthecodinium and Schizochytrium and some species of Chlorella, the specific growth rate can reach values as high as 6.24 day⫺1 (2.7 h doubling time).

Table 6.2 Features, size and weight of the microalgae currently used in aquaculture. Genera and species

Features

Length ⫻ width (␮m)

Arthrospira

Cylindrical cells forming helicoidal trichomes

Chaetoceros calcitrans

Free-livinga, brown, four long diagonal setae

50–300 ⫻ 10 (trichome) 1–12 (cell) 5–16 –

C. calcitrans forma pumilum

Smaller than C. calcitrans, proportionally shorter setae a

Chlorella

Free-living , green, spherical, cell wall, 2–16 autospores

Crypthecodinium

Free-livinga, mobile, spherical or elliptic, incomplete cingulum, forming cyst Free-livinga, mobile, yellow to golden brown, short subapical haptonema with two flagella, no cell wall

Isochrysis galbana

I. affinis galbana (T-iso)

Identical to I. galbana

Nannochloropsis oculata

Free-livinga or aggregated, ovoid, cell wall, no chlorophyll b

Pavlova lutheri

Free-livinga, mobile, gold–brown, ovoid, short haptonema with two unequal flagella, no cell wall

3–8 3–7 1.5–10 3–7 (vulgaris) 1–11 10–30 30 5–6 ⫻ 2–4 ⫻ 2.5–3

6.4 4.6 3–5 2–4

Cell weight (pg)

11 31–61 8–22 (owb)

23–47 (owb) 31 41–83 15–19 30–44 10

3–6 7–9 ⫻ 5–7 ⫻ 3–4 3–6

30–40 (owb) 102 57 (log phase)

References Vonshak et al. (1982) Borowitzka & Borowitzka (1988) Chrétiennot-Dinet et al. (1986) Brown (1991) Lopez-Elias & Voltolina (1993) Chrétiennot-Dinet et al. (1986) Fernandez-Reiriz et al. (1989) Chrétiennot-Dinet (1990) Gladue & Maxey (1994) Hirata et al. (1997) Sournia (1986) Gladue & Maxey (1994) Chrétiennot-Dinet et al. (1986) Fernandez-Reiriz et al. (1989) Brown (1991) Fidalgo et al. (1998) Wikfors et al. (1996) Brown (1991), Brown et al. (1998) Chrétiennot-Dinet et al. (1986) Brown et al. (1993b) Gladue & Maxey (1994) Chrétiennot-Dinet et al. (1986) Fernandez-Reiriz et al. (1989) Brown (1991) Brown et al. (1993b)

Phaeodactylum tricornutum

Rhodomonas

Chained, oval, Y or spindle-shaped, single valve

3–4 ⫻ 8 to 25–35 5–10 (owb) 77 81–140 (owb)

Free-livinga, mobile, two subapical flagella ⬎10 5–12 (salina)

Schizochytrium sp. Skeletonema costatum

Tetraselmis

Thallum attached to substrates by rhizoidal network, Heterokont biflagellate spherical to ovoid zoospores Chained, spherical to cylindrical cells attached together by a ring of external gutter-shaped processes

Free-livinga, mobile, spindle-shaped cells, four polar flagella, cell wall, cyst form

13 ⫻ 8 4.5 5.3 3–10 10 ⫻ 5

b

Free-living: not colonial and not benthic cells. ow, organic weight. Total weight when not specified.

52 18

10 ⫻ 4 9–11 ⫻ 7–8 ⫻ 4–6 160–227 (owb) 7–22 10–12 ⫻ 8–10 (chui) 9–11 (striata)

a

100 (reticulata) 79

104–135 (chuii) 64–158 (striata) 200 (suecica)

Chrétiennot-Dinet et al. (1986) Fernandez-Reiriz et al. (1989) Brown (1991) Fernandez-Reiriz et al. (1989) Chrétiennot-Dinet (1990) Brown et al. (1997) Laing & Psimopoulous (1998) Renaud et al. (1999) Azevedo & Coral (1997) Azevedo & Coral (1997) Chrétiennot-Dinet et al. (1986) Brown (1991) Cognie & Barille (1999) Renaud et al. (1999) Chrétiennot-Dinet et al. (1986) Fernandez-Reiriz et al. (1989) Gladue & Maxey (1994) Wikfors et al. (1996) Wikfors et al. (1996) Laing & Psimopoulous (1998)

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Table 6.3 Growth rates of the microalgae species commonly used in aquaculture.

Genus Arthrospira Chaetoceros

Chlorella

Crypthecodinium Isochrysis

Nannochloropsis Pavlova

Phaeodactylum Rhodomonas Schizochytrium Skeletonema

Tetraselmis

Specific growth rate (day⫺1)

Culture conditionsa

References

2.47 1.68 0.87 1.37 1.47 2.03 0.3 0.16 4.75 1.98 1.4 0.75 1.49 1.56 0.48 0.28 0.92 0.9–1.2 0.25 1.5 1.6 0.37 0.30 6.24 0.84–2.88 2.2 2.0 1.35 1.68 0.74–1.14 1.4 0.11 0.36–0.41

maxima, 550, 12/12, 33°, B platensis, 273, 24/0, 35°, C calcitrans, 60, 24/0, 15°, C, 35 g l−1 gracilis, 165, 24/0, 18°, B gracilis, 332, 12/12, 18°, B vulgaris, heterotrophy, B vulgaris, axenic, B vulgaris, open condition, B pyrenoidosa, heterotrophy, B cohnii, heterotrophy, B galbana, 36, 24/0, 25°, B galbana, various nitrogen sources, B T-iso, 165, 24/0, 18°, B T-iso, 332, 12/12, 18°, B T-iso, 40 W fluorescent, 24/0, 20°, B sp., 75, 12/12, 20°, B lutheri, 20°, C lutheri, 15–23°, B lutheri, 60, 20°, B tricornutum, 23°, B tricornutum, 16°, B salina, 100, 12/12, 22°, C, 35 g l−1 sp., 80, 12/12, 25°, B, 25 g l−1 sp., heterotrophy, B costatum, 135, 24/0, 20°, B costatum, 130, 24/0, 20°, B costatum, 120, 24/0, 18°, B costatum, 100, 24/0, 20°, B, 25 g l−1 sp., N/P 2.5–80, 25°, B suecica, through strain selection, B sp., 360, B sp., 70–80, 12/12, 20°, B sp., 80, 12/12, 25°, B, 25 g l−1

Zarrouk (1966) Aiba & Ogawa (1977) Fernandez-Reiriz et al. (1989) Toro (1989) Toro (1989) Mayo & Noike (1994) Lau et al. (1994) Lau et al. (1994) Running et al. (1994) De Swaaf et al. (1999) Ryu & Tokuda (1984) Ryu & Tokuda (1984) Toro (1989) Toro (1989) Brown et al. (1998) Brown et al. (1998) Droop (1974) Goldman (1979) Hiramatsu et al. (2000) Ben-Amotz & Gilboa (1980) Sigaud & Aidar (1993) Brown et al. (1998) Renaud et al. (1999) Barclay et al. (1994) Curl & McLeod (1961) Hitchcock (1980) Mortain-Bertrand et al. (1988) Blanchemain (1993) Molina-Grima et al. (1991) Gonzalez-Chabarri et al. (1992) Molina-Grima et al. (1994a) Brown et al. (1998) Renaud et al. (1999)

a Principally refers to species, energy (in ␮Einstein m⫺2 s⫺1, if unit is not specified), light regimen (light/dark hours day⫺1), temperature (°C), renewal status (B, batch; C, continuous or semi-continuous culture), salinity.

Biomass production is the major criterion for assessing cultures of microalgae. The production P expresses the increase of biomass per unit of time. When related to the surface of illumination (g m⫺2 day⫺1) or to the volume of culture (g l⫺1 day⫺1), it is called productivity. Its calculation depends on the manner in which the culture is operated. Two situations are considered, depending on whether or not the illuminated reaction medium is renewed after inoculation. The first situation, by far the more frequent, is called batch culture and consists of no media renewal other than supplying air enriched with carbon dioxide and illumination during cultivation. Figure 6.2 presents a typical kinetic of growth for batchcultured microalgae, where five phases can be identified. The production P of biomass in this case is P ⫽ (C ⫺ C0)V/(t ⫺ t0), where V is the volume of culture produced. A culture can also be run continuously. In this case, the reaction medium is renewed and an equivalent flow Q of culture is harvested. The equation that simulates this situation


Biomass of the population

dB/dt = k2

4

213

5

3

dB/dt = k1 B

2 1 Time

Fig. 6.2 Typical growth kinetic of microalgae cultivated in a batch, and phases. 1: Lag or acclimation phase; 2: exponential phase: cells divide actively. The low concentration and abundant nutriments result in no limitation (␮ ⫽ ␮max); 3: limitation phase: exponential growth could not be sustained because of light and substrate restrictions; 4: plateau phase: cessation of growth; 5: senescence phase: death and lysis of cells.

expresses that the variation of concentration in the reactor is the difference between the growth of the biomass and the population reduction following dilution/harvesting: dC/dt ⫽ (␮ ⫺ D) C, where D is the dilution rate Q/V. Three conditions may develop:

• • •

D ⬎ ␮: Growth cannot compensate for dilution and the concentration C diminishes; the culture washes out. D ⬍ ␮: The culture has not reached its steady state and concentration still increases. D ⫽ ␮: The concentration stabilises around a mean value C.

In a stabilised continuous culture (␮ ⫽ D), the production of biomass per unit of time p is the product of the harvesting flow rate DV by the concentration C ( p ⫽ DVC). Although a continuous culture could theoretically last indefinitely, deposits on the light wall and contaminations may result in a drop in production. The longest runs reported in the literature have lasted for several months. Growth of algae, like that of all plants, is governed by environmental factors, among which the most influential are the source of energy (light or organic substrates), temperature (affecting biochemical reaction velocity), nutrients (involved in the organic growth), and other medium factors such as salinity, pH, mixing and toxins. In addition to the production of biomass, its biochemical composition is affected by these factors, which are examined below.

6.2.3 Substrates of photoautotrophy 6.2.3.1 Light Light, through photosynthesis, plays a fundamental role in the development of cultivated microalgae. It is highly variable in intensity and distribution, and both aspects have to be considered for photosynthesis.

214


Light is an energy flux, which propagates in a straight line in a homogeneous medium. The behaviour of light can be simulated either as an electromagnetic wave or as particles. In the first case, it is characterised by its frequency ␯ and its energy ε which are related by ε ⫽ h␯, where h is the Plank constant. In the second case, the particles composing light are called quanta or photons. Light is then quantified through its photon-flux density, or light irradiance, and expressed in ␮Einstein m⫺2 s⫺1 (or ␮mol-quanta m⫺2 s⫺1, or ␮mol-photons m⫺2 s⫺1). The wavelength ␭ ⫽ c/␯ (where c is the light celerity) is more commonly used to characterise the distribution of the radiations (spectrum) of a light. As a consequence, the physical energy content of ‘blue’ photons (440 nm) is higher than that of ‘red’ ones (650 nm). Light of different wavelengths is utilised differently by the photosynthetic apparatus of a plant. The photosynthetically active radiation (PAR) is found between 400 and 700 nm. Taking sunlight as an example, only about 45% (Kirk 1986) of the energy is useful for plants (found in the PAR spectrum), the remainder being either ultraviolet (shorter than 400 nm) or infrared (longer than 700 nm). Light sources specialised for plant production generally produce reinforced reddish radiation, while green radiation is minimised. Light irradiance can be measured by means of several types of quantum meter, some of which present the PAR directly in ␮Einstein m⫺2 s⫺1. Some units based on light perception by the human eye are also used to quantify light, giving readings in either lux or lumen m⫺2. The PAR corresponds roughly to the visible range of the light, but human eye sensibility is centred on yellow light (550 nm), whereas plants utilise blue light (440 nm) and the yellow to red range of wavelengths (620–680 nm). This makes it necessary to consider the nature of the light source in order to convert lux to ␮Einstein m⫺2 s⫺1. For example, for cool white fluorescent tubes, 1 ␮Einstein m⫺2 s⫺1 corresponds to 74 lux (Thimijan & Royal 1982). The PAR energy is dissipated in the culture, in the cell and in the chloroplasts as fluorescence and heat, so that only a fraction of the light is transformed into chemical energy. According to Oswald (1978), the maximum practical energy conversion efficiency for outdoor continuously mixed cultures is near 5% of total solar energy, which is one of the highest values encountered in the entire plant kingdom. Hence, the photosynthetic response in terms of accumulation of organic matter by a given plant will vary according to both the light spectrum and intensity of the source, and the nature of its photochemical apparatus. They condition the biomass production and the overall biochemical content of the cells. Protein level is thought to increase more with blue than with white light, and glycogenesis is reported to be favoured by red light (SanchezSaavedra & Voltolina 1996). The reactions of biosynthesis occur according to two successive phases, each of them under the control of photosystems I and II located in the membrane of the chloroplast. In the first phase, the ‘light reaction’, the photons are absorbed by the chlorophyll, which is activated and then returns to its previous state while producing energy, which is used to split water molecules into H⫹, O2 and electrons. The protons accumulate in the thylakoid lumen of the chloroplast, resulting in pH gradients of up to 3 units. The electrons accumulate as reduced nucleotides (NADPH) in the chloroplasts. The proton gradient is used to produce directly usable energy under the form of adenosine triphosphate (ATP). The ATP and NADPH generated are then used to produce reduced organic molecules supplying chemical energy for further biosynthesis, especially of glucids.


215

Specific growth rate (day⫺1)

To determine the influence of intensity, spectrum and regimen of delivery, the relation between photosynthesis and irradiance has been studied in wild phytoplankton by oceanographers, and in several microalgae species by plant physiologists. This became possible thanks to two recent techniques: the use of radioactive isotopes of carbon for measuring the rate of organic build-up of the cells (primary production), and the use of polarographic Clark electrodes for the study of the rate of photosynthetic oxygen production. During photosynthesis in microalgae, chlorophyll a is observed to increase almost linearly with increasing irradiance until a maximum plateau is reached, and may even diminish at higher irradiance. This loss of efficiency with high energies, also called photoinhibition, has been attributed to an overdestruction rate of the high-turnover D1 protein of photosystem II (Aro et al. 1993). In an attempt to describe the influence of light on photosynthesis, the curves of variation of the oxygen evolution rate with respect to chlororophyll a content, also called P/I curves, were fitted to mathematical equations. Several fitting parameters were determined, especially the initial slope, the maximum production and the optimal irradiance corresponding to it. The results showed variations in relation to time, species and the physiological state of the cells, and revealed a highly unstable and adaptive capacity (e.g. Macedo et al. 1998). Other works were devoted to the influence of irradiance on the specific growth rate ␮, an example of which is given in Fig. 6.3 for three microalgae. The simplifying assumption that growth is nil at dark was frequently made. As photosynthesis/irradiance and growth/irradiance curves showed similar shapes, they were often fitted using the same equations. Most of those proposed in the literature (Table 6.4) do not simultaneously take into consideration the three key criteria of consistency of this energy transformation (Muller-Feuga 1999): (a) loss of weight with respiration in the dark, (b) maximum growth rate ␮, and (c) maximum yield ␮/I. The parameter values necessary to adjust these models are not yet available for all the microalgae of aquaculture. However, saturation light energy Is is generally in the 300–600 ␮Einstein m⫺2 s⫺1 range for most species. As indicated in Table 6.3, the highest values of the specific growth rate (␮s) are between 0.5 and 1.5 day⫺1. Compensation energy Ic, though a concept familiar to oceanographers, is not available for most aquaculture species. 2.0

1.5

1.0

0.5

0.0 0

200 P. micans

400 600 Irradiance (µEinstein m⫺2 s⫺1) I. galbana T. weisflogii

Fig. 6.3 Growth versus irradiance of three species of microalgae at 18°C: Prorocentrum micans, Isochrysis galbana and Thalassiosira weisflogii. (Reprinted from Falkowski, P.G., Dubinsky, Z. & Wyman, K. (1985) Growth irradiance relationship in phytoplankton. Limnol. Oceanogr., 30, 311–321. Copyright 1985, with permission from Elsevier Science.)

216


Table 6.4 Major models of energy transformation available in literature, the number of parameters required for their fitting, and their consistency. Consistency

References Michaelis & Menten (1913) Monod (1942) Tamiya et al. (1953) Droop (1983) Kiefer & Mitchel (1983) Haldane (1930) Aiba (1982)

Number of Weight loss Maximum Maximum parameters at dark growth yield

Model ␮⫽

␮s I ␮s ⫹ I

Same model with translatory motion along y-axis ␮s I

␮⫽

⌲s ⫹ I ⫹

1

⫺

⫺

⫺

2

⫹

⫺

⫺

3

⫺

⫹

⫺

I2 K1

Tessier (1942) van Oorschot (1955)

␮ ⫽ ␮s(1⫺e⫺i)

2

⫺

⫺

⫺

Steele (1977)

␮ ⫽ ␮si e(1⫺i)

2

⫺

⫹

⫺

Peeters & Eilers (1978)

i ␮ ⫽ 2␮s (1 ⫹ ␤) 2 i ⫹2 ␤i⫹1

3

⫺

⫹

⫹

4

⫹

⫺

⫹

3

⫺

⫺

⫹

4

⫹

⫺

⫹

2

⫺

⫹

⫺

3

⫹

⫹

⫹

Bannister (1979)

Moser (1985) Molina Grima et al. (1994b) Lee et al. (1987) Muller-Feuga (1999)

␮⫽

␮⫽

␮si 1 ( K1n⫹i n ) n

⫺␮ 0

␮si n n ik ⫹i n

Same model with translatory motion along y-axis i ␮⫽ Ki ⫹ K 2 i 2 ␮⫽

2␮s (1⫺ic )(i⫺ic ) (1⫺ic )2 ⫹(i⫺ic )2

The notations are homogenised to facilitate the comparison: ␮s ⫽ growth rate at saturation; Is ⫽ saturation energy; I ⫽ energy; i ⫽ normalised energy I/Is; ic ⫽ normalised compensation energy Ic/Is. (Reprinted from Muller-Fenga (1999), Growth as a function of rationing: a model applicable to fish and microalgae. J. Exp. Mar. Biol. Ecol., 236, 1–13. Copyright 1999, with permission from Elsevier Science.)

In nature, sunlight is delivered according to nycthemeral light/dark (L/D) cycles varying with location and season. The question then arises of the necessity of simulating such cycles in controlled cultures. Toro (1989) observed no significant differences in growth rate and final concentrations between 165 ␮Einstein m⫺2 s⫺1 under continuous illumination and 332 ␮Einstein m⫺2 s⫺1 under 12/12 h L/D cycles, for Chaetoceros gracilis and Isochrysis affinis galbana ‘Tahiti’ (T-iso) cultured in batch and after 14 days (Table 6.3). Continuous illumination of indoor cultures was preferred for practical reasons, and this has proved suitable for most species cultured. The gradual extinction of light penetrating into the depth of the culture, also called the self-shading effect, creates a light gradient which makes the relationship between incident energy and growth much more complicated than for a suspension with low cell concentra-


217

tion. In intensive cultures, the increasing cell concentration diminishes the amount of energy I available for photosynthesis according to the Beer–Lambert law. Written I ⫽ I0 e⫺␣C h, this law expresses that the relative variation in intensity dI/I along the path length h of an absorbing medium is inversely proportional to the distance dh and the concentration C. This law is strictly valid for monochromatic lights as the coefficient of proportionality ␣ (sometimes called the coefficient of extinction) depends on the wavelength. Self-shading is generally the limiting growth factor in intensive culture. Some authors (Gladue & Maxey 1994) consider that this is the main disadvantage of photoautotrophic microalgae production. Because of the low concentrations attained, very large amounts of water must be handled, resulting in increasing requirements for space, labour and energy, and highly variable quality. These aspects will be considered in more detail in Section 6.4. 6.2.3.2 Mineral nutrients As for all plants, the major nutrients for microalgae are carbon, nitrogen and phosphorus in mineral form. Diatoms also require silicon. Microalgal organic matter contains about 50% carbon and 10% nitrogen. In nature, carbon is available as carbon dioxide present in the atmosphere at a concentration of 0.03%. However, in intensive cultures of microalgae, carbon dioxide is generally mixed with the air injected at the bottom of vessels. Other nutrients are supplied in the liquid culture medium according to different recipes, the main ones being Watanabe (Watanabe 1960), f series (Guillard & Ryther 1962), Zarrouk (Zarrouk 1966), Conway (Walne 1966), Kuhl (Wong 1977), AM (Fabregas et al. 1984) and Knop (Bajguz & Czerpak 1996). As indicated in Table 6.5, f/2 and Conway media provide satisfactory growth for most of the species of microalgae used in aquaculture. This simplifies their culture to some extent, as hatcheries generally cultivate more than one species. However, all other factors remaining constant, the composition of the culture media determines to a large extent the biochemical composition of the biomass. For example, comparing five media, Sánchez et al. (2000) showed that they greatly influence the fatty acid composition and the protein content of Isochrysis galbana, concluding that Ukeles medium modified by Fabregas and Herrero (1985) provides the best compromise between optimising the biochemical profiles and the growth kinetics for this species.

6.2.4 Substrates of heterotrophy Some microalgae have the ability to grow in the dark, using organic substrates in place of light energy. The addition of a small range of reduced organic compounds, such as glucose, some amino acids and some carboxylic acids in different concentrations, depending on the species cultured, can accelerate algal growth. After 7 days, the cell concentrations of C. gracilis and Pavlova lutheri cultured in light increased perceptibly after the addition of 30 ␮g l⫺1 of citric acid (Ohgai et al. 1993). Working on Tetraselmis suecica, Cid et al. (1992) showed that the supply of both light and organic substrates could affect the biochemical composition in comparison to a control without any organic compound, with much higher carbohydrates and protein content, whereas lipid content changed little. Heterotrophic cultivation was assessed with Chlorella in Asia in the early 1980s and later on with Tetraselmis (Day & Stavalos 1996). Among the 10 genera investigated,

218


Table 6.5 Liquid and gaseous substrates supply conditions for the photoautotrophic cultivation of the main species of microalgae used in aquaculture.

Species

Liquid medium

Vitamin

Arthrospira Chaetoceros

Zarrouk Conway ⫹ Si f/2 ⫹ Si f/8 ⫹ Si Knop

Autotrophic Autotrophic

Chlorella

Air supply CO2 (v/v/ha) (% in air)

20

0.5–1.5

Thiamine AM M4N Watanabe, f/2 Bristol Isochrysis galbana I. affinis galbana Tahiti (T-iso) Nannochloropsis oculata Pavlova lutheri

Phaeodactylum tricornutum Rhodomonas Skeletonema costatum Tetraselmis

f/2 f/2 ES Conway

10

Thiamine B12 Autotrophic

150 60 17

0.5

17

0.5

Thiamine f/2, h/2 B12, thiamine Conway f/2 f/2 ⫹ Si

0.5 Autotrophic

f/2 f/2 ⫹ Si AM f/2 Conway

17

0.5

17

0.5

B12 B12 B12

References Vonshak (1997) Walne (1966) Brown et al. (1993b) Soudant et al. (1998) Wong (1977) Turner (1979) Fábregas et al. (1984) Mandalam & Palsson (1998) Sung et al. (1999) Illman et al. (2000) Turner (1979) Fernandez-Reiriz et al. (1989) Sánchez et al. (2000) Brown et al. (1993a) Price et al. (1998) Albentosa et al. (1999) Turner (1979) Brown et al. (1998) Bonin et al. (1986) Fernandez-Reiriz et al. (1989) Brown et al. (1993b) Burkhardt et al. (1999) Brown et al. (1998) Bonin et al. (1986) McCausland et al. (1999) Fábregas et al. (1984) Brown et al. (1998) Renaud et al. (1999)

a

v/v/h, volume of gas per volume of culture and per hour.

Gladue and Maxey (1994) showed that Cyclotella, Ochromonas, Tetraselmis and Nitzschia have moderate to rapid growth in heterotrophy and are of potential interest for aquaculture, whereas Nannochloropsis exhibit slow growth. Thus, the choice of species and genera will probably differ depending on whether growth is by photoautotrophy or heterotrophy. When the latter condition is well accepted, doubling times are within 3.5–4.8 h (instead of 16–24 h in photoautotrophy) and maximum cell densities can reach 40–100 g l⫺1.

6.2.5 Other factors affecting growth 6.2.5.1 Temperature Raising the temperature can increase microalgal growth up to an optimum point, after which it is reduced. Table 6.6 summarises the growth temperatures of species used in aquaculture. As no standard is available to define minimum, maximum and optimum temperatures, the data should be considered with due caution. However, with respect to the


219

Table 6.6 Temperature tolerance of the main species of microalgae used in aquaculture. Temperature (°C) Species

Minimum

Maximum

Optimum

References

Arthrospira sp. Chaetoceros sp.

15 10

45 40

Chlorella sp. C. saccharophila C. sorokiniana C. fusca var. vacuolata Isochrysis galbana

22

35 26–30 38–42 34 28

35–38 16 20–30 26

I. affinis galbana ‘Tahiti’ (T-iso)

15

36

2

29

vws Flassch (1978) Wei et al. (1986) vws Kessler (1980) Kessler (1980) Kessler (1980) vws Ryu & Tokuda (1984) vws Srisudha & Nair (1996) vws Abu-Rezq et al. (1999) vws Fanuko (1981) Sigaud & Aidar (1993) vws Blanchemain (1993) Lopez-Munoz et al. (1992)

Nannochloropsis oculata Pavlova lutheri Phaeodactylum tricornutum Rhodomonas sp. Skeletonema costatum Tetraselmis sp.

3

11 11

32

2 15

30 32

11–16 22–25 30 20–25 24–25 22–26 20 16–26 20–25 20 20

vws, various web sites.

optimum temperature for culture conditions, 25°C on average is suitable for nearly all the species except for Arthrospira, S. costatum and Tetraselmis. 6.2.5.2 Salinity Most of the microalgae used in aquaculture are euryhaline. For example, Dunaliella salina is known to grow in supersaturated waters, where it develops the characteristic reddish colour of ␤-carotene. Adaptative mechanisms producing variations in the inner concentration of small molecules such as glycerol and sorbitol compensate for ambient osmotic pressure. However, Table 6.7 shows that preferred salinity for most species is below that of seawater. Although the use of seawater can be advantageous for medium preparation in intensive culture, salinity should generally be reduced to 25–27 g l⫺1 for improved growth. 6.2.5.3 Metabolites As for most micro-organisms, microalgae development is generally accompanied by the excretion of metabolites that accumulate in the reaction medium and could become inhibitory for growth. However, because of light limitation, the cell densities in photoautotrophic cultures remain small compared with those of fermentation, and such inhibitions seldom occur. In an attempt to determine the optimum cell density of outdoor Nannochloropsis cultures, Richmond and Zou (1999) removed the inhibitory activity of high-density suspensions by replacing the culture medium after separation. The concentration threshold at which inhibitory activity began to reduce the growth was in the range of 6–7 g dry weight l⫺1. According to Javanmardian and Palsson (1991), who studied

220


Table 6.7 Salinity tolerance of the main species of microalgae used in aquaculture. Salinity (g l⫺1) Species

Minimum

Maximum

Arthrospira sp.

2.5

80

A. platensis Chaetoceros sp.

1.6

88

Chlorella sp. Isochrysis galbana

20 15 10

I. affinis galbana ‘Tahiti’ (T-iso) Nannochloropsis oculata

35 36 36 35

Optimum

13–16 28.5–30.5 20–30 15 15–25 25–35

35 15–23 20–30

Pavlova lutheri

0.1

60 2–10 5–35

Phaeodactylum tricornutum Rhodomonas sp. Skeletonema costatum Tetraselmis sp.

2 0 10 10

63 40 33 45 36

25 25–30

References Borowitzka & Borowitzka (1988) Nana-Kebou-Hako (1999) Baticados & Gacutan (1977) Flassch (1978) Alias (1988) Laing & Utting (1980) Renaud & Parry (1994) Abu-Rezq et al. (1999) Renaud & Parry (1994) Wood et al. (1999) Chini Zitelli et al. (1999) Bonin et al. (1986) Brand (1984) Droop (1958) Fanuko (1981) Sigaud & Aidar (1993) Hill (1992) Blanchemain (1993) Laing & Utting (1980) Fabregas et al. (1984)

Chlorella, the inhibitory compounds could be analogous to pheromonal mating factors and could exhibit bacteriocidal activity. The accumulation of dissolved oxygen produced during cultivation may affect growth at high concentrations, inhibiting photosynthesis and even causing photo-oxidative cell destruction. This situation could occur when degassing is insufficient, and especially in closed systems (Richmond et al. 1993). The literature mentions the adverse effect of high oxygen contents on microalgae growth (e.g. Tredici et al. 1991). Ogawa et al. (1980) found that Chlorella vulgaris growth is reduced by some 20% when the partial pressure of oxygen varies from 21 to 65 kPa O2, i.e. from air saturation to a concentration of 29 g l⫺1. 6.2.5.4 pH With the exception of Arthrospira, which tolerates pH as high as 11, most species used in aquaculture require a pH between 6 and 9, with an optimum close to neutrality. Growth usually increases pH because carbon dioxide in the medium is consumed by the microalgae. For this reason, and because precipitates could appear when adding the enrichment medium, initial batch pH should be as low as growth compatibility allows. Seawater pH (around 8.3) is currently adjusted to 7.0–7.5 by addition of acid prior to culture inoculation. The carbon dioxide injected to supply photosynthetic carbon requirements also reduces the pH and helps to maintain this within the growth range for the algae. 6.2.5.5 Mixing In intensive photoautotrophic cultures, mixing is an important factor affecting growth as it allows alternate exposure of the entire microalgal population to light. Using L/D cycles,


221

it has been shown that microalgae continue to grow for a while in darkness, provided that previous light exposure has been long enough (e.g. Merchuk et al. 1998). With increasing cell density, dark zones may develop in the culture vessel. Microalgae have a limited and probably species-specific ability to ‘average’ exposure to strong light and darkness given that adequate mixing is provided (‘light integration’), but the growth will eventually decrease with increasing extension of the dark zones. Mixing is also necessary to provide gas exchange and possibly to break up nutrient and inhibiting metabolite gradients. However, because of the possible detrimental effects of mechanical forces on the integrity of the algal cells (Gudin & Chaumont 1990), mixing must not be too vigorous. The microalgae species used in aquaculture have not been studied for their tolerance to mechanical stress, especially shear forces. As most of them have flagella or appendices, they are usually fragile. For example, T. suecica loses its flagella and stops growing with strong mixing. Those microalgae with a thick cell wall, such as Chlorella and Nannochloropsis, are more tolerant.

6.3 Biochemical Composition of Microalgae The main zoological groups produced by aquaculture, marine fish, shrimps and molluscs, exhibit reduced ability to synthesise highly unsaturated fatty acids (HUFA, with 20–22 carbon atoms and more than three double bonds) by desaturation and chain elongation, low sterol synthesis and poor bioconversion ability (Teshima & Kanazawa 1974; Trider & Castell 1980; Kanazawa et al. 1985; Enright et al. 1986; Watanabe et al. 1989; Soudant et al. 2000). HUFA and cholesterol are thus essential substances that must be supplied by food sources. These substances are abundant in microalgae (Lin et al. 1982; Volkman et al. 1989), which possess ⌬4, 5 and 6 desaturases, allowing synthesis of essential PUFA (with more than one double bond) as well as a large variety of phytosterols, including cholesterol. Microalgae can also provide a large variety of vitamins to satisfy marine animal requirements (Seguineau et al. 1996; Brown et al. 1999). The data on biochemical composition presented here refer mainly to vitamins, sterols and PUFA of phytoplankton species commonly used for bivalve aquaculture. Algae constitute the basic diet of these species that cannot easily be shifted to artificial diets, as can fish and crustaceans. The data presented refer mainly to photoautotrophically produced algae, although some mixotrophic and hetrotrophic marine algae have also been described (Wood et al. 1999). Further details can be found in the well-documented reviews of Brown et al. (1997, 1999) and Volkman et al. (1981, 1989). The composition of microalgae is clearly related to their growth phase (exponential or stationary) and to culture conditions such as light frequency (Brown et al. 1993a), light intensity (Thompson et al. 1993), temperature (Thompson et al. 1992) or culture media (Wikfors et al. 1996). Results in the literature have been obtained mainly with small experimental volumes under various controlled, but highly variable, conditions (batch/continuous, L/D cycle, period of harvesting). These results were supplemented with production data from IFREMER obtained over several years and using larger volumes (300 litre tanks), but in less controlled conditions (non-axenic cultures, variable greenhouse light and temperature), that are more representative of the quality of algae distributed to bivalves in commercial hatcheries.

222


6.3.1 Gross biochemical composition Brown et al. (1997) reported the overall biochemical composition of 40 algal species grown under standard conditions (Fig. 6.4). Protein was the major organic component (15–52% of dry weight), followed by lipid (5–20%) and carbohydrates (5–12%). Lipid content was higher for diatoms, being 18% on average. Figure 6.5 shows results obtained in the IFREMER experimental hatchery with carbon dioxide-enriched aeration. The data are in the range of those reported in the literature, showing a very consistent pattern. Proteins range from 26 to 40%, and carbohydrates and lipids are stabilised around 15–17% for most species, except for diatoms whose storage product is lipid, in accordance with previous reports. Gross composition does not always correlate directly with nutritional value owing to possible deficiency in some essential nutrients. However, when specific essential nutrients are in adequate proportion, the gross composition may be important.

Fig. 6.4 Microalgal lipid, carbohydrate and protein in per cent of dry weight (aeration with 1% carbon dioxide). (From Brown et al. 1997.)

Fig. 6.5 Lipid, carbohydrate and protein in per cent of dry weight in microalgae grown in the IFREMER-Brest hatchery. Microalgae are cultured in 300 litre tanks under the following conditions: artificial ⫹ sunlight, aeration with 1% carbon dioxide, filtered (1 ␮m) seawater, acid-treated and enriched with Conway medium. The number of replications is shown in parentheses.


223

The phase of harvesting and the culture conditions (nutrients, temperature, light intensity) influence the biochemical composition. It is generally accepted that, depending on the algal species, lipids and/or carbohydrates tend to accumulate in late senescence phase (Wikfors 1986; Whyte 1987; Moal et al. 1987), whatever the conditions leading to this phase. Few studies are devoted to microalgal amino acid composition. The data of Brown et al. (1997) for 40 species show a consistent pattern, regardless of algal class. These authors considered the nutritional value of proteins to be due to their composition of essential amino acids being similar to that of the animals to which the algae are offered as feed. Aspartate and glutamate occurred in the highest concentrations (7.1–12.9%), and cysteine, methionine, tryptophan and histidine in the lowest (0.4–3.2%), with other amino acids ranging from 3.2 to 13.5%. In terms of carbohydrate composition, glucose appears as the predominant sugar in all species. There are few significant differences between algal classes, but prymnesiophytes contain more arabinose (2–12%) than other classes (0–2%). Phaeodactylum tricornutum is unique in its rich concentration of mannose (46%). Brown et al. (1997) considered this feature in relation to the low nutritional value reported by Enright et al. (1986). Brown et al. (1993a) also showed that arachidonic acid (ARA) and sugar composition in T-iso did not differ for the range of irradiances tested (50–1000 ␮Einstein m⫺2 s⫺1).

6.3.2 Vitamins Most vitamins are found in algae used in aquaculture (Bayanova & Trubachev 1981; De Roeck-Holtzhauer et al. 1991; Brown & Miller 1992; Brown & Farmer 1994; Seguineau et al. 1996; Brown et al. 1999). The greatest differences between the species concern retinol and pyridoxine in relation to the growth phase. With the onset of the stationary phase, riboflavin, thiamine and vitamin E increase; vitamin C may decrease or increase depending on species (Brown & Miller 1992; Brown & Farmer 1994; Brown et al. 1999); and vitamin A decreases (Seguineau et al. 1993). Major vitamin contents are similar for all species, showing high levels of vitamin C, vitamin E, ␣-tocopherol and niacin (vitamin PP = pellagra preventing factor. Quantitative discrepancies found in the literature are probably related to the analytical and extraction methods, sampling conservation (Brown et al. 1999) and the harvesting phase. Despite the variability in quantitative data, the concentrations found in algae are high compared with both those in human food (De Roeck-Holtzhauer et al. 1991) and recommended levels in a fish diet (Seguineau et al. 1996). The data from Brown et al. (1999) are summarised in Table 6.8.

6.3.3 Sterols Unlike human sterol, which consists exclusively of cholesterol, algal sterols are very complex and show species specificity. Phytoplankton sterols are found in free form in neutral lipids (Ballantine et al. 1979; Volkman et al. 1981). Polar sterols have also been reported, which may in fact be glycosylated forms of sterols (Véron et al. 1998). Variations in sterol levels can depend on the growth phase, nutrients and light conditions of the culture (Ballantine et al. 1979; Gordillo et al. 1998). The 14 sterols listed below can be easily recognised in gas chromatography.

Table 6.8 Reported vitamin content of microalgae. Animal requirements

Content in microalgae

Vitamin

Units

Brown et al. (1999)a

Seguineau et al. (1993)b

Seguineau et al. (1996)c

De Roeck Holtzhauer et al. (1991)d

Bayanova & Trubachev (1981)e

Ascorbic acid (C) ␤-Carotene Niacin ␣-Tocopherol (E) Thiamine (B1) Riboflavin (B2) Pantothenic acid Folates Pyridoxine (B6) Phylloquinone (K1) Cobalamin (B12) Biotin Retinol Ergocalciferol (D2) Cholecalciferol

mg g⫺1 mg g⫺1 mg g⫺1 mg g⫺1 ␮g g⫺1 ␮g g⫺1 ␮g g⫺1 ␮g g⫺1 ␮g g⫺1 ␮g g⫺1 ␮g g⫺1 ␮g g⫺1 ␮g g⫺1 ␮g g⫺1 ␮g g⫺1

2.3 (1.3–3.0) 0.6 (0.5–1.1) na 0.17 (0.07–0.29) 61 (29–109) 32 (25–50) na 21 (17–24) 8.7 (3.6–17) na 1.8 (1.7–1.95) 1.4 (1.1–1.9) ⬍0.7 (⬍0.25–2.2) ⬍0.45 ⬍0.35

1.4 (0.7–1.8) na 0.24 (0.11–0.47) 0.27 (0.18–0.33) 81 (66–86) 0.6 (0.4–0.7) na 12 (7–15) 8.3 (6.7–10) na 4.4 (1.8–7.4) 0.81 (0.7–1.0) na na na

1.3 (0.6–1.9) 0.7 (0.4–1.2) 0.24 (0.11–0.47) 0.23 (0.16–0.35) 65 (40–110) 32 (28–38) 26 (14–38) 12 (7–15) 8.3 (6.7–10) na 4.4 (1.8–7.4) 0.81 (0.7–1.0) na na na

0.5 (0.06–0.84) 1.2 (0.14–4.3) 1.1 (0.03–2.7) 1.7 (0.12–6.32) 550 (290–710) 22 (6–42) na na 96 (4–180) 10 (0–28) 277 (8–1200) na na 14 (0–39) na

1.6 (0.5–2.4) 1.4 (0.3–2.3) 0.33 (0.06–0.58) na 26 (7–48) 43 (15–66) na 26 (2–67) na na na na na na na

Data are average values for three to seven strains in each study (range in parentheses). Culture conditions and the method for harvesting and/or processing are indicated (from Brown et al. 1999). na, not analysed; values for retinol were calculated using 3100 IU ⫽ 1 mg, and those for ergocalciferol and cholecalciferol using 40,000 IU ⫽ 1 mg. a 12/12 h L/D, log phase, centrifuged and/or filtered, freeze-dried and stored at ⫺70°C. b 24/0 h L/D, late log phase, centrifuged and frozen at ⫺20°C. c 24/0 h L/D, late log phase, centrifuged and frozen at ⫺20°C. d 24/0 h L/D, harvest stage not specified, centrifuged (?) and frozen at ⫺30°C. e 24/0 h L/D, late and log phase, centrifuged (?) and analysed immediately. f New (1986), cited by Tacon (1991), for yellowtail, sea bass, sea bream and grouper. g From Conklin (1997).

Marine fishf

Prawng

0.2

0.2

0.15 0.2 20 20 50 5 20 10 0.02 1 1.9 0.025

0.04 0.1 60 25 75 10 50 5 0.2 1 1.6 0.1


225

Sterol nomenclature (trivial and systematic names):

• • • • • • • • • • • • • •

norcholesterol ⫽ 22-trans-24-5,22-dien-3␤-ol cis-dehydrocholesterol ⫽ cholesta-5,22-dien-3␤-ol trans-dehydrocholesterol ⫽ cholesta-5,22-dien-3␤-ol dihydrocholesterol ⫽ cholesta-3␤-ol cholesterol ⫽ cholesta-5-en-3␤-ol brassicasterol ⫽ 24␤-methylcholesta-5,22-dien-3␤-ol 24-methylenecholesterol ⫽ 24␤-methylenecholesta-5,24(28)-dien-3␤-ol campesterol ⫽ 24␣-methylcholesta-5-en-3␤-ol stigmasterol ⫽ 24␤-ethylcholesta-5,22-dien-3␤-ol 4␣-methylporiferasterol ⫽ 4-␣-methyl-24-␣-ethylcholesta-22-en-3␤-ol ␤-sitosterol ⫽ 24␤-ethylcholesta-5-en-3␤-ol fucosterol ⫽ 24␤-ethylcholesta-5,24(28)-dien-3␤-ol methylpavlovol ⫽ 4␣,24␤-dimethylcholestan-3␤,4␤-diol ethylpavlovol ⫽ 4␣-methyl-24␤-ethylcholestan-3␤,4␤-diol.

6.3.3.1 Bacillariophyceae Among diatoms, Chaetoceros exhibits the least diversified sterol spectrum: essentially cholesterol, 24 methylenecholesterol and fucosterol, with no differences between the species mulleri, calcitrans and gracilis (Gladu et al. 1991). Studies show considerable variability in composition owing to confusion between fucosterol and ␤-sitosterol in the earliest works. The mean values for C. calcitrans cultures grown in the IFREMER-Brest laboratory or in commercial hatcheries show the same distribution as those described in the literature (Fig. 6.6).

100

weight % of total sterols

90

Chaetoceros sp. (literature) Chaetoceros calcitrans (hatchery)

80 70 60 50 40 30 20 10

C

es D

D

eh

yd r

oc ho

m os l ho tero l le D ste ih yd rol Br ro c a 24 ssic hol -m as t et hy ero l le C ne am c h p o St este l ig r ol m a β- ste Si ro to l s Fu tero co l M s et hy tero l l M et por hy ife l r Et pav a hy lo l p vo av l lo vo l

0

Fig. 6.6 Sterol profile of the diatom Chaetoceros sp. Comparison of data reported in the literature (mean and standard deviation, n ⫽ 8) with results from cultures in the IFREMER-Brest hatchery (mean and standard deviation, n ⫽ 10) for the strain C. calcitrans.

226


The diatom Skeletonema costatum has a larger sterol spectrum, including cholesterol, campesterol, 24-methylenecholesterol, ␤-sitosterol and fucosterol (Tsitsa-Tzardis et al. 1993; Bergé et al. 1995). The cholesterol content of the S. costatum strain cultured in the authors’ laboratory was lower than that reported in the literature (Fig. 6.7). The variability of quantitative data between studies was greater for Skeletonema than for Chaetoceros and related to culture conditions (Barrett et al. 1995). The weight of sterols per cell also differed for those two species, showing 70 and 54 fg cell⫺1 for Chaetoceros and Skeletonema, respectively. Thalassiosira (Table 6.9) usually has the same sterols as Skeletonema, but 100 weight % of total sterol

90

S. costatum (literature) S. costatum (hatchery)

80 70 60 50 40 30 20 10

D eh yd ro D es ch ol m o C ste ho ro l le s D ih tero y Br dro l ch as 24 ol s -m ica st et er hy ol le C ne am ch pe ol st St er ig o m as l te βr Si to ol st Fu ero co l M s et hy tero l l M et por hy ife lp r av a Et lo hy l p vol av lo vo l

0

Fig. 6.7 Sterol profile of the diatom Skeletonema costatum. Comparison of data reported in the literature (mean and standard deviation, n ⫽ 5) with results from cultures in the IFREMER-Brest hatchery (mean and standard deviation, n ⫽ 8). Table 6.9 Main sterol composition (weight in % of total sterols) of some microalgae: comparison of data in the literature with results from routine hatchery cultures. Species Thalassiosira sp. Cholesterol Desmosterol Campesterol 24-Methylene ␤-Sitosterol Fucosterol Rhodomonas sp. Cholesterol Brassicasterol Tetraselmis sp. Cholesterol Campesterol 24-Methylene

Data from literature

IFREMER-Brest hatchery

28.5 ⫾ 29.4 4.7 ⫾ 0.3 17.8 ⫾ 13.12 45.8 ⫾ 23 6.5 ⫾ 4.8 10.1 ⫾ 7.3

3.5 ⫾ 3 52.9 ⫾ 17.9 43 ⫾ 17.1

Data are mean values ⫾ SD.

Commercial hatcheries

5 2.5 5.6 79.7 3.1 4.1 2.2 ⫾ 0.9 97.8 ⫾ 0.9

7.5 ⫾ 4.3 92.5 ⫾ 4.3

1.7 ⫾ 0.4 82.6 ⫾ 2.8 14.7 ⫾ 0.9

5.9 ⫾ 3 78.3 ⫾ 2.3 15.6 ⫾ 3.9


227

100 90 weight % of total sterol

80 70 60

Isochrysis sp. (literature) T-isochrysis (hatchery)

50 40 30 20 10

D

eh yd r D es och m ol o C ste ho ro l le D ste ih ro y l Br dr o 24 ass ch o -m ica l s et hy ter ol le C ne am ch pe ol st St e ig m rol as βSi tero to s l Fu tero l c M et ost er hy o M l et po l hy rif l p er Et av a hy lov lp o av l lo vo l

0

Fig. 6.8 Sterol profile of the prymnesiophyte Isochrysis sp. Comparison of data reported in the literature (mean and standard deviation, n ⫽ 7) with results from cultures in the IFREMER-Brest hatchery (mean and standard deviation, n ⫽ 16) for the strain Isochrysis galbana (clone T-iso).

with higher and lower percentages of 24-methylenecholesterol and cholesterol, respectively (Gladu et al. 1991). Diatoms are the only algal group containing a high level of cholesterol, the sterol essential for mollusc growth (Trider & Castell 1980). Chaetoceros shows the highest level (50%). 6.3.3.2 Prymnesiophyceae The sterol composition of the Isochrysideae Isochrysis galbana and Isochrysis affinis galbana (clone T-iso) is simpler (Fig. 6.8) and characterised by brassicasterol (⬎90%). Results in the literature are generally similar, with only one study (Véron et al. 1998) showing a different pattern (equilibrated levels of brassicasterol and cholesterol). A peculiarity of Isochrysis is the presence of alkenones, i.e. C37 and C39 ketones (Patterson et al. 1994) that elute after sterols on the chromatogram. Alkenones may be more concentrated than sterols, with 31.9 and 2.8 mg g⫺1 dry weight, respectively (Knauer et al. 1999). The prymnesiophyte Pavlova lutheri (Fig. 6.9) shows a very complex composition with campesterol, stigmasterol, methylporiferasterol, ␤-sitosterol, methyl and ethyl pavlovol (Patterson et al. 1993a). The last two sterols are diols, i.e. with an additional 4␤-hydroxyl group in the structure, found exclusively in Pavlova sp. This species contains much higher levels of total sterols than Isochrysis sp. (290 and 30 fg cell⫺1, respectively). Data obtained routinely in hatcheries are very similar to those described in the literature. 6.3.3.3 Prasinophyceae Two main sterols are evident in Tetraselmis suecica, namely campesterol (52.9%) and 24methylenecholesterol (43%), together with a low amount of cholesterol (Ballantine et al. 1979; Lin et al. 1982; Patterson et al. 1993b). Data obtained in routine hatchery cultures

228


100

weight % of total sterol

90 80

P. lutheri (literature) P. lutheri (hatchery)

70 60 50 40 30 20 10

D eh yd ro D es ch ol m o C ste ho ro le l D ste ih r o y Br dro l ch as ol 24 si c -m as et te hy ro l l C ene am c h p o St est l er ig ol m a β- ste Si r to ol s Fu ter ol c M et oste hy ro l l M et por ife hy r l Et pav a hy lo l p vo av l lo vo l

0

Fig. 6.9 Sterol profile of the prymnesiophyte Pavlova lutheri. Comparison of data reported in the literature (mean and standard deviation, n ⫽ 4) with results from cultures in the IFREMER-Brest hatchery (mean and standard deviation, n ⫽ 2).

are different (Table 6.9), showing higher levels of campesterol in the authors’ hatchery (82%) and commercial hatcheries (78%). The mean content of total sterols in hatchery production is 250 fg cell⫺1. 6.3.3.4 Cryptophyceae No data were found in the literature concerning Cryptophyceae. Those obtained for Rhodomonas sp. in routine hatchery production (Table 6.9) show a very similar profile to that of Isochrysis.

6.3.4 Fatty acids Fatty acids are named here following the C:Xn-Y formula, in which C is the number of carbon atoms, X the number of double bonds, and Y the position of the first double bond counted from the CH3-terminal. The fatty acid composition of microalgae generally shows a very consistent pattern in each group (Napolitano et al. 1990). Fatty acids are distributed among three lipid classes, neutral, glycolipids and phospholipid. Glycolipids, the main constituent (Bergé et al. 1995), are represented by monogalactosyl and digalactosyl glycerides, which are distinctive of plants and located in the thylakoids. Polar lipids (glycolipid and phospholipid), which constitute the membranes, show the highest level of unsaturation. Taxonomic groups exhibit a specific fatty acid composition. Total lipid content and distribution vary considerably with the status of the culture or the strain used, as well as with the environmental conditions of the culture. Decreasing growth rate and temperature increase the lipid content. Temperature mainly affects the galactolipid composition (Henderson & McKinlay 1989), whereas nutrients mainly modify triglyceride and


229

weight % of the total FA

35 30

Chaetoceros sp. literature

25

C. calcitrans hatchery

20 15 10 5 0 14

:0

15

: 0 6: 0 8: 0 -9 -7 -9 -7 -7 -6 -4 -4 -3 -1 -3 -6 -4 -3 -6 -3 -3 -6 -6 -3 -6 -3 -3 1 1 : 1n : 1n : 1n : 1n : 2n : 2 n : 2 n : 3 n : 3n : 4 n : 4n : 2 n : 2 n : 3n : 3n : 4 n : 5n : 2 n : 4n : 5n : 5 n : 5 n : 6n 16 16 18 18 16 16 16 16 16 16 16 18 18 18 18 18 18 20 20 20 22 22 22

Fig. 6.10 Fatty acid (FA) profile of the diatom Chaetoceros sp. Comparison of data in the literature (mean and standard deviation, n ⫽ 7) with results obtained in the IFREMER-Brest hatchery (mean and standard deviation, n ⫽ 6) for the species C. calcitrans.

phospholipid composition (Fernandez-Reiriz et al. 1989; Saoudi-Helis et al. 1994). Decreasing temperature increases the PUFA levels in marine phytoplankton, although the response is moderate (Thompson et al. 1992) and is species specific. Light also exerts some changes, but to a lesser extent. According to Brown et al. (1993a), irradiance caused very little change in the proportions of lipid classes and fatty acid composition of T-iso, with increased levels of docosahexaenoic acid (DHA, 22:6n-3 only). Thompson et al. (1996) obtained the same result for Pavlova lutheri and Thalassiosira pseudonana. However, in Nannochloropsis sp., Sukenik et al. (1989) found that the longest fatty acid, eicosapentaenoic acid (EPA, 20:5n-3), which occurs mainly in galactolipid, increases in low light conditions when galactolipid concentrations increase. 6.3.4.1 Bacillariophyceae The mean fatty acid content for diatoms is 1.7 pg cell⫺1, with a higher content for Chaetoceros (2 pg cell⫺1) than for S. costatum (1.4 pg cell⫺1). The fatty acid composition of diatoms is generally homogeneous within each genus. Saturated fatty acids are represented predominantly by 14:0 and 16:0, monounsaturated fatty acids mainly by 16:1n-7, and PUFA mainly by 16:3n-4, 16:4n-1 (in S. costatum only) and EPA (Ackman et al. 1968; Volkman et al. 1989). The spectra are very similar for the genera Chaetoceros sp. and Skeletonema sp., except for 16:4n-1 (Ackman et al. 1968; Bergé et al. 1995). Although the latter fatty acid was not found in the present cultures, the data are similar to those described in the literature (Figs 6.10, 6.11), showing that 20:4n-6 and the EPA:DHA ratio are higher in Chaetoceros sp. than in Skeletonema sp. 6.3.4.2 Prymnesiophycaea According to data obtained at the IFREMER laboratory, fatty acid content is about 1.8 pg cell⫺1 and 1.1 pg cell⫺1 for T-iso and P. lutheri, respectively, which is in good agreement with results in the literature (Volkman et al. 1991; Wikfors & Patterson 1994). The

230



40 35

S.costatum (literature)

30

S.costatum (hatchery)

25 20 15 10 5 0 14

:0

15


Fig. 6.11 Fatty acid (FA) profile of the diatom Skeletonema costatum. Comparison of data in the literature (mean and standard deviation, n ⫽ 3) with results obtained in the IFREMER-Brest hatchery (mean and standard deviation, n ⫽ 3).


25

T-iso (literature) T-iso (hatchery)

20

15

10

5

0 14

:0

15


Fig. 6.12 Fatty acid (FA) profile of the prymnesiophyte Isochrysis galbana (clone T-iso). Comparison of data in the literature (mean and standard deviation, n ⫽ 9) with results obtained in the IFREMER-Brest hatchery (mean and standard deviation, n ⫽ 7) for the strain Isochrysis affinis galbana (clone T-iso).

major fatty acids of Isochrysis sp. are 14:0, 16:0, 18:1n-9, 18:4n-3 and DHA. Pavlova lutheri exhibits some differences, in that the monounsaturated fatty acid is 16:1n-7 and EPA replaces 18:4n-3. Therefore, P. lutheri provides high levels of both essential n-3 HUFA, with an EPA:DHA ratio ⬎1, and Isochrysis sp. has lower levels of EPA and an EPA:DHA ratio ⬍1. Low levels of ARA (20:4n-6) are also found in both species. The IFREMER hatchery data are quite similar to those in the literature (Figs 6.12, 6.13), but these culture conditions seem to favour the PUFA content of P. lutheri. 6.3.4.3 Prasinophyceae The genus Tetraselmis has a mean fatty acid content of 5.8 pg cell⫺1. The main saturated fatty acid is 16:0, the monounsaturated fatty acids are 16:1n-9 and 18:1n-9, and the main


231

30

P.lutheri (literature) P.lutheri (hatchery)


25 20 15 10 5 0 14

:0

15

: 0 6: 0 8: 0 -9 -7 -9 -7 -7 -6 -4 -4 -3 -1 -3 -6 -4 -3 -6 -3 -3 -6 -6 -3 -6 -3 -3 1 : 1n : 1n : 1n : 1n : 2n : 2 n : 2 n : 3 n : 3n : 4 n : 4n : 2 n : 2 n : 3n : 3n : 4 n : 5n : 2 n : 4n : 5n : 5 n : 5 n : 6n 1 16 16 18 18 16 16 16 16 16 16 16 18 18 18 18 18 18 20 20 20 22 22 22

Fig. 6.13 Fatty acid (FA) profile of the prymnesiophyte Pavlova lutheri. Comparison of data in the literature (mean and standard deviation, n ⫽ 7) with results obtained in the IFREMER-Brest hatchery (mean and standard deviation, n ⫽ 4). 45

T. suecica (literature)


40

T. suecica (hatchery)

35 30 25 20 15 10 5 0 14

:0

15


Fig. 6.14 Fatty acid (FA) profile of the prasinophyte Tetraselmis suecica. Comparison of data in the literature (mean and standard deviation, n ⫽ 4) with results obtained in the IFREMER-Brest hatchery (mean and standard deviation, n ⫽ 2).

polyunsaturated ones are 16:4n-3 and 18:3n-3, together with lower amounts of 18:4n-3 and EPA. Low levels of ARA have also been found (Wikfors et al. 1996). Fatty acid content is similar for Pyramimonas virginica (Chu & Dupuy 1980). On average, Prasinophyceae provide fewer unsaturated fatty acids than Prymnesiophyceae and exhibit total DHA deficiency. The data obtained under IFREMER experimental conditions are in accordance with the literature. However, cells appeared to be enriched in saturated and monounsaturated fatty acids in a reproducible manner, as indicated by the low reported standard deviation (Fig. 6.14). 6.3.4.4 Chlorophyceae This class is not suitable for feeding to molluscs, probably because of its PUFA deficiency. The most unsaturated fatty acids are 16:4n-3 and 18:3n-3 (Ackman et al. 1968; Chu &

232


Table 6.10 Mean fatty acid composition (weight in % of total fatty acids) for two chlorophyceae, a eustigmatophycea and an unknown cryptophycea.

Fatty acid 14:0 15:0 16:0 18:0 16:1n-9 16:1n-7 18:1n-9 18:1n-7 16:2n-7 16:2n-6 16:2n-4 16:3n-4 16:3n-3 16:4n-1 16:4n-3 18:2n-6 18:2n-4 18:3n-3 18:3n-6 18:4n-3 18:5n-3 20:2n-6 20:4n-6 20:5n-3 22:5n-6 22:5n-3 22:6n-3

Dunaliella tertiolecta Compilationa 1.3 0.7 15.7 ⫾ 1.0 1.0 1.7 1.2 ⫾ 0.4 4.4 ⫾ 2.9 1.2 ⫾ 0.1

Chlorella sp. Chu & Dupuy (1980)

Nannochloropsis sp. Sukenik et al. (1989)

Crytophycea Ackman et al. (1968)

2.5

9.0

1.2

27.2 1.3

28.8 0.9

3.6 7.6

26.5 5.5

14.5 1.1 0.5 1.1 3.0 7.2

1.2 6.5 3.4 18.2 ⫾ 7.1 6.6 ⫾ 2.1

11.0

3.5

16.6

23.0 0.8 16.1

32.4 ⫾ 6.4 3.8 ⫾ 0.4 0.6 ⫾ 0.0 0.4

10.5

6.2 19.5

13.8

0.7

Data for Dunalliela tertiolecta are a compilation of mean values ⫾ SD from Ackman et al. (1968), Thompson et al. (1992) and McCausland et al. (1999).

a

Dupuy 1980). The fatty acid composition of Dunaliella tertiolecta and a species of Chlorella is given in Table 6.10.

6.3.4.5 Cryptophyceae Ackman et al. (1968) reported data referring to an unknown cryptophycea (Table 6.10). This species contains 16:0 as saturated fatty acid, 18:1n-9 and 18:1n-7 as predominant monounsaturates, and 18:2n-6, 18:3n-3, 18:4n-3 and EPA as PUFA. This genus provides more EPA (13.8%) than Prasinophyceae.

6.3.4.6 Eustigmatophyceae This class provides EPA at a high level (19.5%) and some ARA (6.2%), but lacks intermediary PUFA (Sukenik et al. 1989). However, it has a high level of monounsaturated fatty acid 16:1n-7 (Table 6.10).


233

6.4 Production Methods for Aquacultural Microalgae The biotechnology of microalgae and cyanobacteria has developed constantly since the 1960s, but more slowly than that of bacteria and fungi. This lag could be due to the special nature of the energy source, since light attenuates while penetrating the reaction medium. This progressive attenuation of light accounts for most of the difficulties encountered during the development of photoautotrophic biotechnology. The first research studies on microalgal production in the 1960s led to the development of most of the culture media and growth models still in use today. These studies were concerned successively with the production of new food resources, the production of energetic hydrocarbons, the preservation of the environment and the production of commercially valuable molecules. Since then, industrial production has developed for human nutrition with Chlorella and Spirulina, for pharmaceuticals with ␤-carotene from Dunaliella salina, and for aquaculture with a dozen species currently cultivated. The value of microalgae for aquaculture has been constant and of a strategic nature since the 1970s, providing food for molluscs, and diets for the live prey species essential for small-mouthed fish larvae and shrimps. With some 2000–4500 t per year depending on sources, Spirulina is the most widely produced genus. World production for aquaculture is difficult to assess, mainly because the microalgae are not harvested, but fed live to animals as raw culture. Initially, aquacultural microalgae were produced by inducing phytoplankton development in ponds with enriched natural water in which the animals to be fed were then directly hatched. These rudimentary rearing methods, which are still used in family farms raising penaeid shrimp in south-east Asia, were the first forms of ‘green water’ and mesocosm techniques. Later, in the 1970s, microalgae were cultured in specialised facilities and then transported to larval or live prey rearing tanks to be used as food and/or for medium stabilisation. The culture techniques developed then have remained relatively unchanged. This section considers the state of the art of microalgal cultivation techniques, with an emphasis on possible improvements.

6.4.1 State of the art of microalgal production techniques in hatcheries Typically (Fig. 6.15), culture units consist of vertical cylindrical tanks (100–500 litres) made of transparent plastic (acrylic, fibreglass polyester, polyethylene), which are flat or conical at the bottom and open at the top or closed by an unsealed cap. Fluorescent tubes placed around the tank in a power:volume ratio of 1 W l⫺1 of culture provide artificial lighting. Natural light is used whenever possible, but the high latitude of most sites requires supplementary use of artificial light. Injection of compressed air at the bottom of each tank produces bubbles that rise to the surface, ensuring mixing as well as gas–liquid exchanges by direct contact. These exchanges consist of oxygen stripping and carbon dioxide enrichment as those gases are produced and consumed, respectively, during photosynthesis. The airflow rate is between 0.1 and 3.0 volume per culture volume and per hour (v/v/h). Although the carbon dioxide content in air is 0.03% in natural conditions, it is often added to the compressed air at a concentration of 1–3% (volume/volume).

234


Fig. 6.15 Typical microalgal production facility for aquaculture at Great Bay Aquafarms, Portsmouth, New Hampshire, USA. This hatchery produces the flounder, Paralichthys dentatus, and the black sea bass, Centropristis striata, two species of small larvae fish. (Photograph: IFREMER/Muller-Feuga).

Rectangular tanks (0.5–1.0 m depth) are also used, with surface lighting provided by artificial sources similar to those used in greenhouses (Fig. 6.16). Once again, air is injected at the bottom of the water column for mixing and gas exchange. These tanks are easier to clean than the transparent ones as they are largely open and do not require tilting for access to the inside. However, production per unit of time and volume is slightly lower. 6.4.1.1 Asepsis and quality controls Certain precautions are needed to maintain culture asepsis and prevent contamination by organisms naturally present in the environment that could act as competitors, predators or toxic agents for microalgae. All fluids and surfaces that come into contact with the culture must be sterilised. If those precautions are not taken, contamination can occur and destroy the cultures, inhibit growth or degrade the nutritional quality of microalgae. Natural water pumped near the facilities is the basis for the culture medium and larval rearing. Thus, the site needs to be carefully chosen in an area with water that is free of pollution and stable in quality, in particular at some distance from towns, harbours and estuaries. Once these precautions are taken, the pumped water is prepared by passage through successive filtering devices. A sand filter reduces the size of suspended particles to 10–20 ␮m, thereby


235

Fig. 6.16 Microalgae production in opaque-walled tanks with an artificial light source above the surface at SATMAR, Barfleur, France. (Photograph: IFREMER/Barbaroux).

eliminating most of the zooplankton. These filters are easily cleaned by regular reverse flushing. Additional filtration by means of cartridges or sieves reduces particle size to 1 ␮m, thereby eliminating most of the protists. Although filters are generally recyclable after washing, they represent an important expenditure item. Water preparation by filtration often remains at this stage, still allowing prokaryotes to pass through. Additional filtration by active carbon is sometimes necessary to eliminate organic substances such as phytoplankton toxins. In some cases, chemical sterilisation is performed within the culture tank once it is filled with filtered water. This consists of the addition of a product whose sterilising toxicity can be eliminated by adding a neutralising agent after a suitable contact delay, e.g. oxidant– reductant or acid–base pairs. Hypochlorite is generally reduced by thiosulfate after overnight contact. The other procedures for keeping cultures aseptic are filtration of the gas injected at the bottom of the tanks and cleaning of intermediate surfaces and containers. The staff can contribute to cleanliness by wearing disposable or regularly cleaned gloves, overalls, headdresses, and overshoes. A footbath filled with sterilising product should be placed at the entry to the culture room to avoid ground contamination. In practice, these precautions are often resisted by a workforce better accustomed to handling a scoop than Pasteur pipettes. Microalgal cultures for aquaculture are not monoseptic because the means available in hatcheries and the capabilities of the staff are generally inadequate to achieve this quality level. For instance, the systematic use of low melting temperature plastics such as polyethylene and polyvinyl chloride (PVC) for culture confinement and piping impedes the use of heating as a sterilising agent. Moreover, even though a culture room may be equipped with dust-control devices to prevent contamination by air, especially at the surface of tanks, this measure is seldom applied. However, as bacteria from microalgal cultures are seldom

236


harmful for algae and larval rearing, aquaculture operators have chosen to consider the current quality of their algae as acceptable. 6.4.1.2 Culture medium and temperature Natural filtered water is enriched by the addition of the mineral salts required for photosynthesis, i.e. metabolisable nitrogen, phosphorus, and trace elements including iron and silicon for diatoms. A chelating agent, generally ethylenediaminetetra-acetate (EDTA), is often added to prevent precipitation of ferric hydroxide. Vitamins such as thiamine or biotin, for which some algae are auxotrophic, should be added with due caution because of their rapid degradation with heat. These salts constitute the enrichment media, the most commonly used being f/2 medium (Guillard & Ryther 1962) and the medium of Conway (Walne 1966). The operators can prepare enrichment media from the constitutive salts. However, some of these media are also marketed in the form of several solutions to be mixed just before use. For example, the salts required for f/2 medium can be purchased at a cost of approximately US $0.5 per 1000 litres of final solution. The same enrichment medium is generally used for all species of microalgae at the same facility. Moreover, as all of these cultures are grown in the same room, this common environment does not make it possible to take the physiological features of each species into consideration. For example, temperature is often controlled between 18 and 25°C for reasons of comfort, whereas this important growth factor should be fine-tuned to each species. Qualitative levelling is likely to occur in these common conditions, thereby reducing the advantages of biodiversity. 6.4.1.3 Running the cultures Batch cultures are generally run according to production cycles of 3–7 days. Once illuminated tanks have been cleaned and filled with filter-sterilised water, enrichment medium is added and compressed air supply plugged, and an inoculum is introduced into the tank in such an amount that the resulting concentration prevents the culture from failure and ensures a quick start. These initial concentrations should be just sufficient for providing photoprotection by self-shading and for outrunning possible competitors. They differ according to the light source and the species. For example, 400,000 cells ml⫺1 are necessary for Tetraselmis suecica and 106 cells ml⫺1 for Pavlova lutheri with 150 ␮Einstein m⫺2 s⫺1 from fluorescent sources. The algal strains are provided from large culture collections or specialised public organisations in the form of a few millilitres of culture in a test-tube (generally not bacteria free). Starting from this sample, successive volumes of increasing size are inoculated in order to prepare the biomass required to reach inoculation concentrations in the production tanks. The intermediate vessels generally used are 250 ml conical flasks, 2 litre bottles and, finally, 10–20 litre bottles or carboys (Fig. 6.17). The densities reached in these intermediate cultures vary from 2–3 ⫻106 cells ml⫺1 for T. suecica to 5–15 ⫻ 106 cells ml⫺1 for P. lutheri. 6.4.1.4 Efficiency The cultures obtained in current hatchery facilities seldom exceed a density of 6 ⫻ 106 cells ml⫺1 at the end of 5 days, i.e. a dry weight concentration of 0.12 g l⫺1 (the dry weight


237

Fig. 6.17 Intermediate cultures for the inoculation of production tanks at Satmar, Barfleur, France. (Photograph: IFREMER/Barbaroux).

of a cell is on average 20 pg). Thus, a facility with a capacity of 5000 litres operating on 200 days per year and producing 24 kg of dry microalgae requires a full-time technician, uses 23,000 kWh and salts to manufacture 200 m3 of culture medium, and needs to amortise about US $10,000 worth of equipment. The total production cost is approximately $1400 kg⫺1 of dry matter of microalgae in the conditions existing in western Europe. These figures are in agreement with those of Bennemann (1992). Meanwhile, other industrial facilities specialised in the production of microalgae and exploiting production systems in highly controlled conditions, such as photobioreactors, can market their product at prices lower than US $200 kg⫺1 of dry weight. This suggests the possibility of a considerable reduction in costs, which should be of interest to aquaculture operators. The cost price of producing hatchery microalgae includes labour (90%), amortisation (6%), energy (3%) and miscellaneous expenses (1%). Thus, a decrease in production costs depends especially on a reduction of staff time, with no very significant increase in other expenses. An increase in production yields can also contribute to a reduction in cost price.

6.4.2 Methods of improvement Productivity is often used to compare various production systems or to define the optimal conditions for operating a given system. This criterion consists of the dry biomass produced per unit of culture space and of time.

238


6.4.2.1 Continuous cultures The main labour-consuming operations are the distribution of cultures when mature, the cleaning of vessels and their filling with culture medium, and the inoculation process. Thus, a reduction in the time devoted to these operations presumes an increase in efficiency or removal of parts or all of these processes. Continuous cultures are the chief means of obtaining such results. Intermediate modes of operation between batch and continuous (called semi-batch or semi-continuous) consist of collecting only part of the culture volume and replenishing with fresh medium. The residual volume of a culture then constitutes the inoculum of a new production cycle. For example, such renewals may comprise two-thirds of the volume every 5 days. Typically, a continuous culture involves supplying a culture tank of volume V with fresh medium according to a renewal flow rate Q. A balance is established at the end of a few days between the renewal of the medium and the growth capacity of the algal population. This balance is characterised by the concentration of the culture C and the dilution rate D (D ⫽ Q/V expressed in % per day). The dilution rate D is a major parameter since it determines the average age of the cells, which has a considerable influence on their biochemical characteristics (Brown et al. 1993b). Another factor is the residence time T, which is the time required for the complete renewal of the volume of culture (T ⫽ 1/D in days). According to the type of facility and species, the dilution rate D is between 20 and 50% per day, corresponding to a residence time T between 5 and 2 days. Volume productivity, which measures the output efficiency of a system, is equal to the dilution rate multiplied by the concentration (P ⫽ CD ⫽ C/T, in g l⫺1 day⫺1). Once a culture has been started in the continuous mode of operation, human intervention is limited to preparation of the nutrient medium upstream, use of the harvest downstream, and a general, rapid monitoring of parameters for the culture and the facility. A substantial reduction in labour requirements may follow (seven-fold according to some estimates). However, this would be accompanied by an increase in investments, particularly for the fresh medium pumps and automatic systems such as carbon dioxide injection under the control of pH. 6.4.2.2 The increase in production yields An increase in concentration of the cultures would result in a more efficient utilisation of the production factors, with a reduction in the share of nutrients and electricity, and amortisation in the cost price of the algae. In most hatcheries, culture concentrations seldom exceed 0.1 g l⫺1 (or approximately 6 million cells ml⫺1 for T-iso), whereas concentrations of photoautotrophic cultures produced in photobioreactors are reported to be some g l⫺1. This high discrepancy can be explained by the difference in light path length of the cultures in those production systems, as it has been stated that this factor has a strong inverse influence on the final concentration (e.g. Grobbelaar 1981; Hartig et al. 1988; Tredici et al. 1991; Pulz et al. 1995). To illustrate this observation, Fig. 6.18 shows a comparison between 500 mm diameter hatchery tanks, with a mean light path length of 392 mm, and photobioreactors, whose light path length ranges between 8 and 30 mm, for the culture of the red microalga Porphyridium cruentum. It is possible to increase concentrations10-fold by reducing the light path length. This example shows that the final concentration C of a culture varies with


239

Concentration (gl⫺1)

2 ALP 2 ALP 1 NLP

1

Tanks 0 0

100 200 300 Mean culture thickness (mm)

400

Fig. 6.18 Culture concentration versus mean optical thickness obtained for different systems used for the production of Porphyridium cruentum (precision bars show SD). The curve corresponds to 42 g m⫺2 areal density. (Reprinted from Muller-Feuga, A., Le Guédes, R., Hervé, A. & Durand, P. (1998) Comparison of artificial light photobioreactors and other production systems using Porphyridium cruentum. J. Appl. Phycol., 10, 83–90: with kind permission of Kluwer Academic Publishers.)

its light path length E according to a simple relation stating that the product of these two factors, also known as areal density S (Soeder 1980), remains constant (S ⫽ CE, in g l⫺1 mm, or g m⫺2), at least for a given species, and provided that their productivity remains at a maximum. Reported areal densities are in the vicinity of 40 g m⫺2 for Dunaliella salina (Grobbelaar 1994) and for Scenedesmus sp. (Livansky et al. 1995; Hartig et al. 1988), and 66 g m⫺2 for Arthrospira platensis (calculated from Torzillo et al. 1986). However, some results do not fit with this simple assertion, and Gudin and Chaumont (1990) proposed detrimental effects of mixing on cells as a possible explanation. It is interesting to examine the practical implications of the stability of the areal density S. According to the Beer–Lambert law, the transmission It/I0 remains constant and equal to e⫺␣S, where I0 is the initial energy hitting the culture, It is the energy transmitted throughout the culture and ␣ is the coefficient of extinction of the culture. In the example of Fig. 6.18, where S was 42 g m⫺2, the mean value of ␣ over the spectrum was equal to 0.115 m2 g⫺1. The calculation of the transmission through the continuous culture of P. cruentum gave 0.8% regardless of the light path length of the production system used, which was confirmed by direct measurements. This suggests that the microalgae cultures in production are unable to utilise all of the light provided and their concentration stabilises before complete extinction. Volume productivity seldom exceeds 20 mg l⫺1 day⫺1 in most hatcheries, whether cultures are operated in batch or continuously. Meanwhile, production values for photobioreactors usually exceed 250 mg l⫺ 1 day⫺1, representing a 10-fold increase. This situation allows a considerable margin of progress for hatcheries, as the technologies developed for the photobioreactors could usefully improve the quality and reduce the cost price of algae. The concept of closed tubular photobioreactors was initially explored in the early 1980s, when their efficiency was assessed for different designs and for species such as A. platensis (Torzillo et al. 1986) and P. cruentum (Chaumont et al. 1988). Although hesitant and

240


burdened by some failures, industrial and commercial developments implementing photobioreactor technology have considered microalgae of interest for the production of natural substances. More recently, attention has been drawn to the production of algae of interest, such as Nannochloropsis sp., for the feeding of aquaculture larvae (Zou & Richmond 1999). This study investigated the productivity of flat plate reactors of light path length ranging between 1.3 and 17.0 cm exposed to natural light. Maximum volume productivity reached 1.7 g l⫺1 day⫺1. The reduction in the light path length of a culture is generally associated with an increase in the concentration and productivity. As observed by Borowitzka (1996), the diameter of the photobioreactor tubes has tended to decrease from 100–300 mm in the first vessels (Torzillo et al. 1986) to 40 mm and less in current types. Appropriate technical solutions for cooling outdoors facilitites are required. There is a risk of damage to the photosynthetic system when exposed to high light intensities. Sunlight can reach more than 2000 ␮Einstein m⫺2 s⫺1 for PAR at midday. Although several mechanisms, such as carotenoid synthesis, have been developed by microalgae to survive what appears to be a recurrent crisis, the most efficient protection against high irradiance remains self-shading. In addition, decay processes and dark respiration may result in a considerable loss of biomass at night. Tredici et al. (1991) mentioned differences between day and night concentrations as high as 42% for the cyanobacterium Anabaena azollae, whereas Molina-Grima et al. (1995) measured night losses smaller than 8% for Phaeodactylum tricornutum. The production of microalgae in photoautotrophy is influenced by a series of biological antagonistic effects such as access to light and self-shading, mixing and cell fragility, high irradiance and thermal stability, requiring the least unfavourable compromises. The closed photobioreactors are surely the most efficient, but designs and operation modes under investigation are still quite diverse, using plates or tubes, sun or artificial light, glass or plastic. As an example among others, the technology developed by Muller-Feuga et al. (1998a,b) uses artificial light, is of stainless-steel and glass modular conception, and allows axenic and mixotrophic cultures (Fig. 6.19). This diversity suggests that microalgal photoautotrophic biotechnology has not yet reached its maturity, even though some commercial products and services have already been marketed.

6.4.3 Heterotrophic production Some microalgae are able to use organic substrates efficiently in darkness and are thus potentially suitable for production by fermentation. This approach offers many technical advantages compared with the photoautotrophic method, especially since concentrations of the organic substrate and oxygen can be as high as their solubility allows and also homogeneous in the whole reaction volume. Fermentation technologies, which are widespread in the food and pharmaceutical industries, provide high-quality products. Such cultures are necessarily monoseptic. Fermentation volumes are huge (up to 500 m3), which reduces costs to levels of a few dollars per kilogram of dry weight. The final concentrations of heterotrophic cultures of Chlorella sp. can reach 100 g l⫺1, but are more often close to 40 g l⫺1 (Running et al. 1994). Doubling times are then drastically reduced compared with those of photoautotrophy (3.5–4.8 h instead of 16–24 h). As a result, volume productivity greater than 200 g l⫺1 day⫺1 can be reached, representing a 100-fold increase compared


241

Fig. 6.19 Example of an artificial light photobioreactor. The culture is impelled by a propeller and an air-lift along a loop passing through eight series-connected light chambers per production set, three of them in parallel being represented here. pH, temperature, dissolved oxygen, pressure and absorption are measured in line and regulated. Liquid and gaseous substrates are filter-sterilised at the inlet and at the outlet. Steam can be admitted into the reactor at several points for initial sterilisation. Continuous cultures can be run according to both turbidostat and chemostat modes. Culture volume: 300 litres; illuminated surface: 4.5 m2.

with photoautotrophy. However, although heterotrophic cultures are of considerable interest because of their high productivity, their application is restricted to those species that are able to grow and to produce the key metabolites in these conditions. The synthesis of PUFA does not seem to be affected, although some biocompounds such as pigments are not synthesised in heterotrophy (Grobbelaar 2000). These biochemical modifications may downgrade the nutritional value of the algae. Mixotrophy, an intermediate solution consisting of heterotrophy in light, is still under investigation.

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The first heterotrophic microalgae that appeared on the market as food for rotifers were chosen from those already in use, such as Tetraselmis suecica (Day et al. 1991) and Chlorella sp. (Lee 1997). The former species had no commercial success, probably because of biomass processing that decreased its nutritional value. However, the latter species is in high demand in Asia in the field of dietetics. The Japanese market represents 500 t per year, and Korea has an ambitious production programme of 1000 t per year (Apt & Behrens 1999). However, as the PUFA composition of this alga is not satisfactory for aquaculture, other species have been proposed because of their high DHA content, such as the diatom Nitzschia, the thraustrochytrid Schizochytrium sp. (Barclay et al. 1994) and the dinoflagellate Crypthecodinium cohnii (De Swaaf et al. 1999).

6.4.4 Discussion The productivity of the microalgal systems used in aquaculture hatcheries is10-fold lower than that of photobioreactors, which is in turn 10-fold lower than that of fermentation techniques (Table 6.11). The cost prices of the biomass produced with these systems are directly related to this evident disparity. Thus, the techniques of microalgal production used in aquaculture hatcheries are far less efficient than those of other sectors of application. Why do aquaculture operators fail to use more efficient technologies allowing both cost reduction and improved quality? First, it should be recalled that microalgae are intermediate products requiring further processing and not a marketable end-product on which the profitability and image of the company are based. Consequently, entrepreneurs are reluctant to take a technological leap that would require significant investment and the recruitment of more qualified staff. Moreover, the technological advances in the production of artificial diets could make this leap unnecessary. The example of shrimp aquaculture shows that microalgae can be replaced by microencapsulated food, a practice that could one day be extended to larval rearing of small-larva fish and most molluscs (Muller-Feuga 2000). Table 6.11 Comparison of the concentration, productivity and cost price of some aquaculture microalgae for various types of production system.

Type of production system Tanks

Species

Volume productivity of dry Concentration biomass (g l⫺1) (g l⫺1 day⫺1)

T-iso 0.1 Skeletonema sp. Pavlova lutheri Nannochloropsis sp.

Order of magnitude of cost price Calculation (US $ kg⫺1) data from

0.02

1000

Bennemann (1992) Brown et al. (1993b)

Photobioreactors Nannochloropsis sp. 1–5

0.5–1.7

100

Chini Zittelli et al. (1999) Zou & Richmond (1999)

Fermentors

100–200

10

Day et al. (1991) Gladue & Maxey (1994) Barclay et al. (1994) De Swaaf et al. 1999

Tetraselmis suecica Cyclotella cryptica Chlorella sp. Crypthecodinium cohnii Schizochytrium sp.

40–60


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The aquaculture operator is persuaded that replacement products for microalgae will eventually appear on the market, offering constant, if not necessarily high, efficiency at acceptable prices. In that case, microalgae would no longer be needed. This accounts for the success of live, concentrated Chlorella produced by fermentation in Japan and proposed as food for rotifers. In fact, algal culture rooms have disappeared from most fish hatcheries in that country. The recent marketing of DHA-rich heterotrophic species is also a response to the demand for suitable food for live prey of fish larvae. Thus, high culture concentrations are essential since they reduce the volume of water handled and transported. Preservation methods at low temperature involving the addition of a preservative are under investigation and will certainly become increasingly efficient. Moreover, it is likely that microalgae for fish aquaculture will be produced in the near future by specialised companies implementing high technology, which will propose highquality products whose shelf-life could reach several months, perhaps in competition with microencapsulated foods. The prospects are different with respect to mollusc production, for which several species of live microalgae are required. Since the preserved microalgae available on the market so far are not suitable for the early stages of these species, on-site production seems unavoidable, at least in the short term. Under these conditions, investment in the improvement of microalgal production systems is beneficial. Moreover, photobioreactors and/or fermentors could be an attractive long-term solution for aquaculture operators who wish to diminish the cost price of microalgae, depending on the applications and considering their capacity to produce the most suitable species examined in Chapter 7.

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Chapter 7

Uses of Microalgae in Aquaculture A. Muller-Feuga, R. Robert, C. Cahu, J. Robin and P. Divanach

7.1 Introduction Unlike their terrestrial equivalents, aquatic animals used as food by humans are rarely herbivorous at the adult stage. Only filtering molluscs and a few other animals are true plankton feeders throughout their lifetime. Most farmed aquatic animals are carnivorous from their postlarval stage and sometimes omnivorous. However, microalgae are required for larval nutrition during a brief early period, either for direct consumption (molluscs and penaeid shrimp) or indirectly as food for live prey fed to small marine fish larvae. Even when necessary for a short period only, microalgae are crucial as they determine (to various extents) the supply of juveniles available for production. Freshwater species such as salmonids do not depend on microalgal production for their culture. Their eggs have sufficient reserves to hatch large larvae capable of feeding directly on dry particles. Certain marine species such as the European sea bass have larvae that are large enough to feed directly on Artemia nauplii. The main microalga-consuming aquaculture groups include filtering molluscs, penaeid shrimps and small larva fish. Overall world production of these microalga-consuming species reached 12 ⫻ 106 t in 1999, i.e. 28% of world aquacultural production (FAO: Shatz 2000). Filtering molluscs constitute in weight the most significant contribution to aquaculture production, with a total of 10 ⫻ 106 t in 1999, and a 23% increase over 5 years. Nutritionally, these harvests are dependent on wild phytoplankton in natural water masses circulating around the livestock, and juvenile supply comes mainly from natural spat collection. However, hatcheries in which larval and juvenile production depends on cultured microalgae are assuming an increasingly important role (Muller-Feuga 2000). The algal requirements of mollusc larvae are considered in Section 7.2. Farmed shrimp reached 1.2 ⫻ 106 t in 1999, and exhibited a 19% increase over 5 years. Production is carried out mainly in subtropical regions of America and south-east Asia. Microalgae are necessary from the second stage of larval development (zoea), and in combination with zooplankton from the third stage (mysis). Although of short duration, these larval stages require microalgal culture facilities, which vary with the size of the hatchery and the extent to which medium parameters are controlled. Although the trend is to substitute cultured microalgae with dry formulated feeds, microalgae are still necessary (Rosenberry 1998). Section 7.3 considers algal requirements for shrimp larvae. Marine finfish aquaculture reached 0.8 ⫻ 106 t in 1999, and is sharply increasing worldwide at a rate of 55% over 5 years. Small larva species, such as sea bream (146 ⫻ 103 t in

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1999) and flatfish (31 ⫻ 103 t in 1999), require small live prey that feed on phytoplankton. They can be fed yeast-based artificial feeds, but this is a much less efficient alternative to phytoplankton. Section 7.4 considers the conditions for using microalgae as food for live prey. The survival and growth of various marine fish larvae are improved by the addition of microalgae. The best results are obtained with live cultures added to, or grown directly in tanks with live prey and/or larvae, and a variety of intensive and semi-extensive technologies has developed along with this practice. Section 7.5 considers other roles of phytoplankton in aquaculture and the possible mechanisms involved.

7.2 Microalgae as Food for Molluscs The culture of microalgae is of fundamental importance to commercial hatcheries rearing marine molluscs, since they are currently the only suitable food source. Molluscs, unlike fish and crustaceans, are fed microalgae directly. Consequently, mollusc development is closely related to the quantity and quality of phytoplankton produced. Microalgal cultures are necessary because the concentration of natural phytoplankton in the seawater used in the hatchery is generally insufficient for optimum growth of the high densities of larvae and juveniles reared. Moreover, to avoid bacterial diseases, the seawater used for rearing is purified, usually by fine filtration (0.2–1.0 ␮m) and/or ultraviolet (UV) treatment (Robert & Gérard 1999). This eliminates almost all of the natural phytoplankton, which must then be replaced by dense artificial cultures. Many attempts have been made to determine which of the microalgal species provide the best food value in terms of mollusc optimal growth and survival (for a review, see Chrétiennot-Dinet et al. 1986). Fifty species have been tested on larvae and juveniles of commercially raised bivalves, including 12 commonly used in mollusc hatcheries. Table 7.1 shows that the relative importance of some species has changed significantly in recent Table 7.1 Utilisation frequency of microalgal species in a mollusc hatchery. Utilisation frequency (%)

Microalgal species

Class

Walne (1970)

Lucas (1980)

Coutteau & Sorgeloos (1992)

Chaetoceros calcitrans Chaetoceros gracilis Skeletonema costatum Phaeodactylum tricornutum Thalassiosira pseudonana, clone 3H Isochrysis galbana Isochrysis affinis galbana (clone T-iso) Pavlova lutheri Pyramimonas virginica Tetraselmis suecica Dunaliella sp. Nannochloropsis occulata

Bacillariophyceae Bacillariophyceae Bacillariophyceae Bacillariophyceae Bacillariophyceae Prymnesiophyceae Prymnesiophyceae Prymnesiophyceae Prasinophyceae Prasinophyceae Chlorophyceae Eustigmatophyceae

40 —a 20 50 40 80 20 70 0 60 0 0

37.5 —a 12.5 12.5 62.5 75 0 62.5 37.5 25 25 25

37 53 14 5 33 19 72 26 —a 35 9 —a

From Robert and Trintignac (1997a). a No data.

Uses of Microalgae in Aquaculture

255

years, reflecting a better knowledge of their food quality and/or feasibility for bulk production. Isochrysis galbana has been replaced by Isochrysis affinis galbana ‘Tahiti’ (clone T-iso), which is more tolerant to high temperatures (Ewart & Pruder 1981). The recognition of Chaetoceros gracilis as a valuable species (Enright et al. 1986) has increased its use, whereas the low larval growth rates obtained with Phaeodactylum tricornutum, Pavlova lutheri and Tetraselmis suecica explain their decreasing use. Coutteau and Sorgeloos (1992) found that 50% of hatcheries produce less than 5 m3 of microalgal culture per day, 25% 10–50 m3 and 25% up to 110 m3, i.e. a daily mean production per hatchery of 13.5 m3. They also found that eight algal species (Chaetoceros calcitrans, C. gracilis, Skeletonema costatum, Thalassiosira pseudonana clone 3H, I. galbana, I. aff. galbana, P. lutheri and T. suecica) were most widely used, representing 90% of the volume of algal culture produced, and that most hatcheries cultured between two and five different algal species. However, this study underestimated the production of S. costatum, which is widespread in France (Sauriau & Baud 1994). Among the hatchery molluscs that do not require microalgae are the giant clams (Tridacna gigas and Hippopus hippopus. They need only yeast extract and vitamins for their larval development and adopt a particular feeding strategy after metamorphosis that consists of symbiosis with the dinoflagellate Symbiodinium microadriaticum (Fitt et al. 1984).

7.2.1 Microalgae as a potential food source in mollusc hatcheries Four criteria are required for a microalga to qualify as a potential food source for bivalves in a hatchery: size, digestibility, good nutritional value and ease of bulk production. 7.2.1.1 Size Under controlled conditions, clam larvae (Mercenaria mercenaria) can actively ingest food particles as small as 0.5 ␮m in diameter, such as the cyanobacterium Synechococcus spp. (Gallager 1988). When offered a 50:50 mixture of T-iso and Synechococcus spp., 2-day-old clam larvae (mean length 100 ␮m) ingested on average 48 Synechococcus spp. cells (mean diameter 0.7 ␮m) for every T-iso cell (mean diameter 4.5 ␮m). This mean ratio dropped to 3:1 in 10-day-old larvae (mean length 234 ␮m), indicating a clear relation between prey and larval size (Gallager 1988). When fed on natural assemblages of phytoplankton, Crassostrea virginica larvae over 300 ␮m were able to graze prey as large as 30 ␮m, such as dinoflagellates (Baldwin & Newell 1991, 1995). However, they also selected small food particles (0.2–0.8 ␮m). Thus, in natural surroundings, C. virginica larvae are able to graze over a wide phytoplankton size range, depending on population blooms (Baldwin 1995; Baldwin & Newell 1995). However, more than half (55%) of all veligers of scallops (Placopecten magellanicus), mussels (Mytilus edulis) and clams (Mya arenaria) collected in the field contained cells of 5–15 ␮m in their stomach, but only 3% had cells of 15–25 ␮m (Raby et al. 1997). Moreover, when fed natural suspensions ranging in size from 1 to 10 ␮m, C. virginica larvae of all sizes (small, 100–125 ␮m; to large, 200–290 ␮m) showed a strong preference for 2–4 ␮m particles (Fig. 7.1a). Conversely, small larvae fed mixtures of four cultured microalgae ranging in size from 1 to 11 ␮m preferred those of 1 ␮m, while large larvae preferred those of 11 ␮m, confirming the relation between prey and larval size (Fig. 7.1b).


(a) 0.7

125 µm-larvae

0.6 0.5

260 µm-larvae

Selectivity (Wi)

256

0.4 0.3 0.2 0.1 0 1–2

2– 4

4– 6

6–8

8–10

Phytoplankton size fraction (µm) 106 µm-larvae

0.6

290 µm-larvae

Selectivity (Wi)

(b) 0.7 0.5 0.4 0.3 0.2 0.1 0 1.0 (Syn)

4.5 (T-iso)

6.2 (Dun)

11.0 (Pro)

Phytoplankton size (µm) and composition Fig. 7.1 Food selection (Wi) patterns of Crassostrea virginica larvae (a) on different sizes of natural phytoplankton assemblages and (b) on mixtures of cultured microalgae. Syn: Synechococcus bacillaris; T-iso: Isochrysis affinis galbana; Dun: Dunaliella tertiolecta; Pro: Prorocentrum mariae-lebouriae. The horizontal line across each graph represents neutral selection. (Reproduced from Baldwin, B.S. (1995) Selective particle ingestion by oyster larvae (Crassostrea virginica) feeding on natural seston and cultured algae. Mar. Biol., 123, 95–107. With permission of Springer-Verlag.)

As noted by Fritz et al. (1984), the critical size of selected cells smaller than 10 ␮m probably depends on mouth and oesophagus diameters, which become larger as molluscs grow, and measure 10–20 ␮m for C. virginica larvae above 160 ␮m, according to Ukeles and Sweeney (1969). Moreover, some microalgae have shapes that make their ingestion more difficult, for example diatoms with long spines (Robert et al. 1989). 7.2.1.2 Digestibility Some authors have considered that the low larval growth rates obtained with some microalgae can be explained by the thickness of their ‘cell wall’ that makes their digestion difficult. This is the case with Chlorella autotrophica (Babinchak & Ukeles 1979), Dunaliella primolecta, Tetraselmis suecica (Le Pennec & Rangel-Davalos 1985), Nannochloris atomus and Stichococcus bacillaris (Robert 1998). On the basis of epifluorescence microscopy observations, these studies showed that those species mentioned above are normally ingested, but not well digested, resulting in poor larval growth for Crassostrea gigas or C. virginica. Conversely, Pavlova lutheri is highly ingested and well digested by the larvae of C. virginica, M. edulis and Pecten maximus (Babinchak & Ukeles 1979; Lucas & Rangel 1981; Le Pennec & Rangel-Davalos 1985), and Chaetoceros calcitrans forma pumilum by 3–4-day-old C. gigas larvae (Robert et al. 1989). A lack of appropriate digestive enzymes


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in young larvae is the current hypothesis to explain why some algal species resist breakdown, passing intact through the gut. 7.2.1.3 Nutritional value: biochemical composition of microalgae Gross composition differs among species, but is generally not the major factor determining their food value. In particular, there is no clear relationship between the whole protein content of microalgae and their nutritional value for bivalves, and amino acid composition cannot explain differences in food quality (Webb & Chu 1983; Brown et al. 1989). The latter authors suggested that larval molluscs in general require 30–60% protein for good growth. Because of the variability in carbohydrate composition, there are few consistent compositional differences between algal classes. However, in some instances, carbohydrates may play an important role in balancing the utilisation of protein and lipid for biosynthesis against catabolism for energy production (Whyte et al. 1989). Indeed, when adequate amounts of protein and lipid are supplied, the use of Chaetoceros muelleri, containing high levels of carbohydrates, enhanced the growth of Ostrea edulis juveniles (Enright et al. 1986). Although the importance of highly unsaturated fatty acids (HUFA) 20:5n-3 and 22:6n-3 is well documented in vertebrate nutrition, reports concerning the importance of lipids as an energy source for the early life stages of bivalves are contradictory (for review see Knauer & Southgate 1999). Most algae are rich in one or both of these fatty acids. In general, microalgae containing 1–20 fg ␮m⫺3 of eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) produce satisfactory growth in bivalves, whereas poor development is often reported when the concentration is below 0.5 fg ␮m⫺3 (Brown et al. 1989). For example, chlorophytes are not only difficult for bivalves to digest, but also lack these fatty acids, which may explain why they are inadequate for young mollusc larval stages. Specific sterols are considered essential for bivalves, but their role in mollusc feeding requirements remains speculative. Microalgae are rich sources of two key vitamins, ascorbic acid and riboflavin, but some species lack specific vitamins (De Roeck-Holtzhauer et al. 1991) and their composition could account for differences in their nutritional value. As microalgae may lack one or more key nutrients, a mixed algal diet increases the chances of achieving a balanced diet. 7.2.1.4 Microalgae bulk production Some species, despite having good food value, are not used by commercial hatcheries because of difficulties in developing large-scale production for these species. For example, C. calcitrans forma pumilum is seldom grown in volumes over 20 litres (Helm 1990). Some other species such as T. suecica are ‘sticky’ and foul culture tank walls.

7.2.2 Microalgal requirements in mollusc hatcheries Microalgae are used in mollusc hatcheries to feed broodstock, larvae and postlarvae. The phytoplankton requirements of the Japanese oyster C. gigas and the king scallop Pecten maximus at different life stages are shown in Table 7.2. From a size of 1.5–2 mm, oyster spat are usually grown in an outdoor nursery. Daily consumption in these tanks is even greater than in indoor tanks, reaching 40–100 m3 of

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Table 7.2 Phytoplankton requirements of Crassostrea gigas and Pecten maximus in a hatchery. Volume of algae refers to Isochrysis galbana equivalents at 8 ⫻ 106 cells ml⫺1. Alga

One breeding individual

106 larvae

106 postlarvae (0.2–3.0 mm)

Daily consumption Multispecific mixture Microbiological quality Rearing period per batch

0.5–1.0 litres ⫹⫹ ⫹ 1–3 months

2–4 litres ⫹⫹⫹ ⫹⫹⫹ 0.5–1 month

10–20 litres ⫹ ⫹⫹ 2–3 months

From Robert and Gérard (1999). ⫹, Low requirements; ⫹⫹, medium requirements; ⫹⫹⫹, high requirements.

large-scale algal culture (106 cells ml⫺1 mean density) for one million 6–12 mm juveniles (Bacher & Baud 1992). 7.2.2.1 Feeding broodstock The effect of food on broodstock conditioning is species specific for molluscs. It is generally recognised that feeding is necessary during conditioning, but it is still difficult to determine the precise role of adult reserves and of additional food. When seawater was enriched with T. suecica, the European oyster O. edulis produced earlier broods, showed more rapid larval growth and gave greater spat yields than if no additional food was provided (Helm et al. 1973). More recently, Millican and Helm (1994) showed that an algal diet representing 3–6% of the initial meat weight of oysters (dry weight/dry weight) per day increased larval production in O. edulis. The number of larvae released by flat oysters was also closely related to diet quality (the poorest result being obtained with a monospecific Dunaliella tertiolecta diet). Wilson et al. (1996) showed that spat produced by starved females of the Chilean oyster Ostrea chilensis exhibited low rates of growth and survival. As the genus Ostrea is larviparous, it is not surprising that food is particularly important to the flat oyster during the considerable biological effort required by the conditioning process. The parent O. edulis must sustain egg development, embryogenesis and larval growth in the gill cavity for at least 1 week. The same is not true for the cupped oyster Crassostrea gigas, which reproduces externally. Additional food for the broodstock appears to be of less importance than the initial content of glycogen reserves before conditioning. In some cases, food availability favours growth and maintenance rather than reproduction (Donalson 1991), whereas Muranaka and Lannan (1984) found that the fecundity of C. gigas broodstock was 60% greater when fed an algal food supplement rather than starved. However, the rate of gonad development and gamete viability did not differ significantly. Similar observations have been reported in IFREMER-Brest hatchery with flow-through systems of sand-filtered water (about 50 ␮m mesh). Chàvez Villalba et al. (2001) showed that a 6% (dry weight of algae/dry meat weight) addition per day of equal quantitities of Chaetoceros calcitrans and T-iso had a positive effect on C. gigas fertility. During spring conditioning, the mean fertility (number of eggs per female) of fed broodstocks originating from six different French oyster areas was 5.23 million ⫾ 2.91 versus 0.77 million ⫾ 1.21 for unfed groups. Similar observations were reported in summer: 30.74 million ⫾ 12.05


259

versus 8.11 million ⫾ 7.20, respectively, for fed and unfed oysters. Endogenous reserves deposited in eggs during vitellogenesis are an important source of energy during embryogenesis. However, in normal conditions, C. gigas eggs contain sufficient reserves (particularly essential PUFA) to ensure survival throughout embryogenesis, although not for subsequent larval growth (Utting & Millican 1997). Devauchelle and Mingant (1991) reported that the fecundity of the hermaphroditic scallop P. maximus at similar gametogenic stages decreased with feeding levels. Gonad activity suffered first at lower feeding levels, but only in the female part of the gonad. No eggs were obtained after 28 and 45 days of conditioning when scallops were kept unfed, whereas fecundity increased by 8–25% with a food ration of 3 ⫻ 109 cells animal⫺1 day⫺1 and by 30–60% with 14 ⫻ 109 cells animal⫺1 day⫺1. Heasman et al. (1996) reported similar results for Pecten fumatus, showing that gonad condition and egg production improved as feeding rates increased from 12.5 to 100% satiation (equivalent to 0.75 to 6 ⫻ 109 cells scallop⫺1 day⫺1 respectively) at all test temperatures in the 12–21°C range. As in C. gigas, the hatching success rate for P. maximus is positively related to lipid levels in eggs, whereas subsequent larval growth and survival are independent of egg lipid reserves (Le Pennec et al. 1990; Delaunay et al. 1993). The effect on broodstock conditioning of the addition of a mixed algal diet to the circulating water seems to be more marked for P. maximus than for C. gigas (Saout 2000). With C. gigas, specialised storage cells appear to compensate for a lack of food, while muscle and the digestive gland play this role in P. maximus. The effect of food on broodstock conditioning is closely related to the reproduction strategy of each mollusc species. Although a suitable algal ration for bivalve broodstock is 6% (dry weight of algae/dry meat weight) per day for most species reared at 20–22°C, 3% may be sufficient for species reared at lower temperatures (Utting & Millican 1997). Broodstock diet has an impact on the fecundity of the broodstock, but not on subsequent larval growth (except for Ostrea sp.). 7.2.2.2 Feeding larvae Quantitative requirements The number of algal cells eaten by a larva per day is related to both species and size. Ostrea edulis shows a higher feeding requirement than C. gigas, while the Manila clam Ruditapes philippinarum is far less demanding (Fig. 7.2). Moreover, with the exception of R. philippinarum (size ⭓ 190 ␮m), the feeding requirement increases as the larva grows. However, the effects of food on larval development are also related to environmental conditions, especially rearing temperature (Robert et al. 1988; His et al. 1989). The combined effects of temperature and food concentration on the growth of C. gigas larvae fed a mixed diet (P. lutheri and I. galbana) showed that the maximum daily growth rate (8.6 ⫾ 0.9 ␮m) was obtained at the highest temperature (25°C) and food concentration (100 cells ␮l⫺1) (Fig. 7.3). Similar combined effects of food and temperature on larval growth have been reported for other mollusc species receiving monospecific diets (e.g. C. virginica, Rhodes & Landers 1973; Mytilus edulis, Sprung 1984; O. edulis, Beiras & Pérez-Camacho 1994; Patinopecten yessoensis, MacDonald 1988). These results clearly indicate that food demand is closely related to rearing temperature.

260


Thousands of cells of algae eaten per larva per day

140 120 100 80 60 40 20 0 100

150

200

250

300

Mean shell length (␮m) Fig. 7.2 Quantitative feeding requirements of some mollusc larvae expressed as the number of algal cells (equivalent in size to Isochrysis galbana) eaten per day at 24°C. (–.–.–.–.) Ostrea edulis; (-------) Ruditapes philippinarum; (_______) Crassostrea gigas. (From Utting & Spencer 1991, with permission.)

8 6 4

13

tur

Te

50 100 tration (c ells µl ⫺1)

era

20 25 Food co ncen

e(

25 0

°C )

2

mp

Daily growth rate (µm day⫺1)

10

Fig. 7.3 Effect of different concentrations (cells ␮l⫺1) of a mixed diet (Pavlova lutheri and Isochrysis galbana) on young Crassostrea gigas larvae at different temperatures. (From Abdel-Hamid et al. 1992.)

Qualitative requirements It has been clearly established (Fig. 7.4) that a mixture of algal species provides a better food supply than any individual species (for reviews, see Ukeles 1975; Webb & Chu 1983; Brown et al. 1989; Robert & Trintignac 1997a; Knauer & Southgate 1999). Most authors consider that the higher growth and survival rates observed when larvae are fed a combination of microalgae are due to a more balanced diet. However, Gerdes (1983) showed that filtration activity and food uptake for C. gigas are also greater when larvae are fed a mixed algal diet (50 cells ␮l⫺1 I. galbana plus 50 cells ␮l⫺1 C. calcitrans) compared with an equivalent monospecific ration (100 cells ␮l⫺1 I. galbana). This was clearly apparent for larvae with a shell length of more than 120 ␮m. Higher growth rates


Larval length (µm)

250

261

Unfed Skeleto Pavlo T-iso PT PTS

200

150

100 0

5 10 15 20 Time since fertilisation (days)

25

Fig. 7.4 Effects of monospecific and plurispecific diets on the growth of Pecten maximus larvae. Skeleto: Skeletonema costatum; Pavlo: Pavlova lutheri; T-iso: Isochrysis affinis galbana; PT: P. lutheri ⫹ T-iso; PTS: P. lutheri ⫹ T-iso ⫹ S. costatum. (From Robert & Trintignac 1997a with permission.)

were achieved as a result of a nearly two-fold increase in filtering activity with a plurispecific compared with a monospecific diet (18.81 ⫾ 7.06 and 9.85 ⫾ 0.69 ␮l larva⫺1 h⫺1, respectively) and a large increase in filtered algae (0.83 ⫾ 0.45 versus 0.66 ⫾ 0.18 ␮g algal dry weight larva⫺1 day⫺1). Higher food uptake may also account for the faster development generally observed when larvae are fed mixed diets. 7.2.2.3 Feeding spat Nursery cultures of spat from settlement to a size suitable for growth in the sea are crucial to the success of hatchery-based mariculture operations. Flow-through systems (up- or downwelling) are generally used for spat development. At this stage, growth is largely influenced by the amount of food available, even though food quality is still important. As noted above for larvae, spat growth and survival rates are affected by factors such as species-specific requirements (e.g. Walne 1970; Beiras et al. 1994; Lu & Blake 1996), physical parameters such as temperature (e.g. Albentosa et al. 1994; Laing 2000; Robert & Nicolas 2000) and effects of mixed diets (e.g. Enright et al. 1986; Laing & Millican 1986; O’Connor et al. 1992; Brown et al. 1998; Nicolas & Robert 2001). Concerning this last aspect, spat are more tolerant than larvae to monospecific diets. Moreover, because of their greater size, and potentially more efficient digestive tract, they can accept a larger phytoplankton size range. However, the rearing systems used cause flow rates to be critical (Rodhouse & O’Kelly 1981). As spat densities are very high, less than optimal flow rates result in a serious reduction in the rate of spat development, whereas higher flow rates result in an increase in energy expenditure and a waste of algal culture. A compromise between biological efficiency and operating costs (heating, pumping) is often found at this stage (Robert & Nicolas 2000).

7.2.3 Microalgal substitutes for bivalve feeding As indicated in Chapter 6, the large-scale production of selected microalgae by conventional photoautotrophic means is expensive. It represents nearly 50% of the operating costs

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in mollusc hatcheries. An effort has been made to find nutritionally adequate alternatives that are more cost-effective than algae produced on-site. The alternatives tested include bacteria, yeasts, microcapsules, liposomes, lipid emulsions and dried microalgae (for reviews, see Coutteau & Sorgeloos 1992; Robert & Trintignac 1997b; Knauer & Southgate 1999). In general, the lower nutritional value of these products makes them unsuitable as complete feeds, although some have proved useful in a mixed diet with live microalgae. Microalgal concentrates represent the most promising off-the-shelf alternatives. Concentrates produced by centrifugation and stored at 2–4°C for 1–8 weeks have been used successfully as part of mixed or complete diets for larval or juvenile bivalves (Nell & O’Connor 1991; McCausland et al. 1999; Heasman et al. 2000; Robert et al. 2001). Species with good nutritional value and the best shelf-life for oyster larvae and spat include C. calcitrans, S. costatum and Tetraselmis spp., whereas the flagellates P. lutheri and Isochrysis sp. (T-iso) are damaged by centrifugation and deteriorate rapidly. A novel method for preparing microalgal concentrates based on chemical flocculation (Knuckey 1998) has recently been applied to seven marine microalgae grown in 300 litre tanks (Brown & Robert 2001). Six species were successfully harvested with flocculation rates of 58–81%, whereas I. aff. galbana exhibited lower values (31%). Five of these microalgae were stored at 4°C for 10–20 days and tested as a diet, together with 20% live microalgae, for C. gigas larvae and spat. The efficiency of these concentrates on larvae varied with the species tested and the experimental conditions. On three occasions, concentrates gave poorer results than live microalgae (S. costatum, I. aff. galbana). On other occasions, concentrates showed an efficiency better than (C. calcitrans) or equivalent to (C. sp. ‘tenuissimus-like’, C. calcitrans forma pumilum) live microalgae (Fig. 7.5a). A 28 day feeding experiment with oyster spat showed no difference between C. calcitrans forma pumilum and C. sp. ‘tenuissimus-like’, whether live or concentrated (Fig. 7.5b). Recent studies have shown that microalgal concentrates, when appropriately harvested and stored, have the potential to replace the fresh microalgal cultures used to rear larvae

C. pum conc

Diet

C. pum

C. ten conc

C. ten

0

T-iso (control)

50

4.0

3.0

2.0

Diet

C. pum conc

100

C. pum

150

Day 16 Day 28

5.0

C. ten conc

200

6.0

C. ten

(b)

T-iso (control)

250

Spat length (mm)

Larval length (µm)

(a)

Fig. 7.5 Effects of live algae and mixed (20% fresh and 80% concentrate) diet using different algal species on the growth (mean ⫾ SE) of (a) Crassostrea gigas larvae 2 weeks after fertilisation and (b) spat 16 and 28 days after the beginning of the postlarval feeding trial. T-iso: Isochrysis affinis galbana; C. ten: Chaetoceros sp. (tenuissimus like); C. pum: Chaetoceros calcitrans forma pumilum; conc: concentrate. (Reprinted from Brown, M.R. & Robert, R. (2002) Preparation and assessment of microalgal concentrates as feeds for larval and juvenile Pacific oyster (Crassostrea gigas). Aquaculture, 207, 289–309. Copyright 2001, with permission from Elsevier Science.)


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and juvenile bivalves. Diatoms and prasinophytes are good candidates for microalgal concentrates, showing satisfactory shelf-life, whereas further research is required for the more fragile prymnesiophyte species. Regardless of the harvesting methods applied today (centrifugation, Heasman et al. 2000; flocculation, Brown & Robert 2002), naked flagellates are difficult to preserve. Thus, there is considerable scope for the development of improved microalgal harvesting (e.g. bulk filtration) and/or preservation techniques (e.g. the use of antioxidants and other additives) or specific storage protocols with regard to light, temperature and the mixing atmosphere.

7.3 Microalgae as Food for Shrimp Penaeid shrimp are omnivorous during juvenile and adult life and show filter-feeding behaviour during larval stages. Microalgae are the most important dietary source during larval stages in the wild and contribute to the nutrient supply for postlarvae and juveniles in estuaries. Although the total world capture of shrimp remains at a constant level, penaeid shrimp farming has steadily increased since the early 1980s and is now an important part of world aquaculture production. The postlarval supply for aquaculture comes partly from the wild, but increasingly from small- or large-scale hatcheries. Despite the development of different inert microparticulate diets, microalgae are still cultured in hatcheries to feed shrimp larval stages and as an additional dietary source for shrimp in extensive and intensive growth ponds.

7.3.1 Development of penaeid shrimp Two days are required for embryonic development of penaeid shrimp in the egg. Larval life then consists of a succession of moults and metamorphoses. The first phase includes five or six naupliar stages, depending on the species, and involves endogenous feeding. After a first metamorphosis, larvae go through three zoea stages, during which filter-feeding behaviour is predominant and hatchery production is based on a supply of microalgae. The total duration of the zoea phase is around 5 days at 28°C. After a second metamorphosis, the larva is in mysis phase for 3 days. The postlarval stages begin after a final metamorphosis. Mysis and postlarvae exhibit predatory behaviour, feeding mainly on small animal prey, such as Brachionus, Artemia or copepods. However, mysis and postlarvae still ingest microalgae, and the presence of microalgae in the rearing water improves survival and growth during these stages (Fig. 7.6).

7.3.2 Selection of algal species used for rearing shrimp larvae Shrimp larvae, unlike some other crustacean filter-feeders, do not appear to select algae according to size or ‘taste’. Preston et al. (1992), who studied gut content in Penaeus esculentus zoea reared in grow-out ponds, concluded that penaeid zoea do not select particular algal species, but ingest a variety in similar ratios to those in plankton. The highest survival rates in that study were associated with high levels of diatom flora, and the lowest with cyanobacterial flora. Therefore, algal species used to feed shrimp in hatcheries are generally chosen according to size and their ability to grow in culture conditions. The algae used are generally 5 ␮m

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Days

Development stages

Daily feeding ration (for 1000 larvae)

1 No feeding

Nauplius 2

0.5 –1.5 mm

3 4 Zoea

5

50–100 mg microparticles ⴙ 15,000 algae cells mlⴚ1

6 7

1.5–5 mm

8 9 10 11

Mysis

120–200 mg microparticles ⫹10,000 Artemia nauplii ⴙ 15,000 algae cells mlⴚ1

5–8 mm

12 300–500 mg microparticlate diet ⫹10,000–20,000 Artemia nauplii ⴙ 15,000 algae cells mlⴚ1

13 14 8 –15 mm

Post-larvae

Fig. 7.6 Development and feeding sequences for penaeid shrimp larvae. (From Cahu, C. 2001, Nutrition and feeding of penaeid shrimp larae. In: Nutrition and Feeding of Fish and Crustaceans. With permission of SpringerVerlag.)

(e.g. Isochrysis sp.) to 10–20 ␮m (e.g. Tetraselmis chuii) in diameter. Larger cells have been tested, such as Prorocentrum micans (32 ⫻ 25 ␮m). Low survival was recorded in zoea stages, but it is unclear whether these poor results were attributable to the biochemical composition or the size of the algae (Sanchez 1986). An experiment based on formulated microparticulate diets showed that zoea I and II stages of penaeid species can ingest spherical particles up to 35 and 50 ␮m in diameter, respectively (Galgani & Aquacop 1988). In addition to selection based on size and culture ability, algal biochemical composition has been studied to determine which algae satisfy the nutritional requirements of shrimp larvae. Few species have been tested in the laboratory or used in intensive shrimp hatcheries. Diatoms such as Chaetoceros and flagellates such as Tetraselmis have provided good results for the growth and survival of penaeid larvae, regardless of the species used. D’Souza & Loneragan (1999) found that survival was two-fold higher when Penaeus


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Table 7.3 Main algal species tested in penaeid larval rearing. Shrimp species

Microalgal species

Observations

References

Penaeus japonicus

Monochrysis lutheri Pheodactylum tricornutum Pseudo Isochrysis paradoxa Tetraselmis suecica

Best growth and survival with T. suecica

Cahu (1979)

Penaeus monodon

Chaetoceros calcitrans Tetraselmis chuii

Better growth and survival with C. calcitrans

Tobias-Quinitio & Villegas (1982)

Penaeus vannamei

Isochrysis sp. Bacteriastrum hyalinum Prorocentrotrum micans

Best growth and survival with Isochrysis sp.

Sanchez (1986)

Penaeus monodon

Tetraselmis chuii Dunaliella tertiolecta Rhodomonas baltica Skeletonema costatum

Very poor survival with D. tertiolecta

Kurmaly et al. (1989)

Penaeus monondon Penaeus japonicus Penaeus semisulcatus

Chaetoceros muelleri Best growth and survival Tetrasemis suecica with C. muelleri and Dunaliella tertiolecta T. suecica Isochrysis galbana, clone T-iso

D’Souza & Loneragan (1999)

japonicus larvae were fed Chaetoceros muelleri rather than Dunaliella tertiolecta. Kurmaly et al. (1989) obtained good growth and survival for Penaeus monodon larvae when Skeletonema costatum and Rhodomonas baltica were used (Table 7.3). Several experiments have been conducted to determine whether a combination of algae was better than a single species, but the results are not conclusive (D’Souza & Loneragan 1999).

7.3.3 Ingestion and filtration rates for shrimp larvae fed microalgae Unlike zoea, which have swimming ability, the mobility of nauplii is poor, and food availability is directly related to its concentration. As for other filter-feeders, an algal threshold concentration can be determined, below which growth is reduced. Above this concentration, shrimp larvae can ingest a sufficient amount of food per time unit to sustain their development. The threshold concentration is estimated at 20,000 cells ml⫺1 for the alga Tetraselmis. At low concentrations, penaeid larvae appear to be less efficient filter-feeders than copepods (Cahu 1979). Conversely, very high concentrations (more than 75,000 cells ml⫺1) may stress the larvae and induce mortality. The filtration rate F and the ingestion rate I can be calculated according to the formulae of Paffenhöffer (1971): F⫽

V [log(C0 ) ⫺ log(Ct )] tn

(7.1)

V (C0 ⫺ Ct ) tn

(7.2)

I⫽

where t is the experimental time (day), C0 and Ct are the initial and final concentrations respectively, n is the number of larvae, and V is the volume of water. Both rates are dependent on the developmental stages of the larvae and on cell concentration. The filtration rate is

266


inversely related to algal concentration. Penaeus indicus can filter 1–4 ml of water day⫺1 at the zoea I stage and 3–10 ml day⫺1 at the zoea III stage when the concentration is between 75,000 and 20,000 cells of Tetraselmis ml⫺1. The ingestion rate increases with cell concentration, reaching a plateau. Beyond this point, the concentration has no effect on the ingestion rate. The maximum ingestion rate calculated for P. indicus fed T. suecica is 70,000 cells larva⫺1 per day for zoea I, 200,000 for zoea II, 250,000 for zoea III and 400,000 for mysis II (Cahu 1979). Similar values were found for P. indicus fed Thalassiosira weissflogii (Emmerson 1980). After the mysis stages, larvae shift towards raptorial feeding. Animal live prey, such as Brachionus and Artemia, need to be added to the diet. Maxillae and maxillipeds, the main parts of the filterfeeding apparatus, become reduced in size, and filtration efficiency declines from mysis to postlarva stages. Postlarval ingestion rate reaches only 150,000 cells algae individual⫺1 day⫺1. Thus, microalgae are still ingested in postlarvae, juveniles and even adult shrimp. The zoea stages constitute a period of intense growth during shrimp development: larval dry weight is 4 ␮g at the zoea I stage and reaches 27 ␮g after 5 days of development to the mysis I stage. A high ingestion rate is necessary to sustain such a growth rate. It appears that a zoea I ingests between three and six times its own dry weight of algae, and a mysis III between one-and-a-half and three times. The following microalgae densities are generally used for shrimp larvae reared in the laboratory as well as in the hatchery: 30,000 cells ml⫺1 of T. suecica and 100,000 cells ml⫺1 of C. muelleri or I. aff. galbana (T-iso).

7.3.4 Nutrient supply from algae in relation to larval shrimp requirements Despite extensive studies on the nutrition of juvenile shrimp, the requirements specific to larvae are poorly known. The global effect of an algal species can be assessed, but it is difficult to understand which of the algal nutritional constituents are essential for larvae. Only the recent use of purified microparticulate diets has allowed a better relationship to be established between larval growth and some biochemical components of algae. Experiments conducted with live food (algae, Brachionus, Artemia) or compound diets have given different results for protein requirement. Optimal protein content in diet dry matter is between 30% (Khannapa 1979) and 50% (Kanazawa 1990). Larval lipid requirements are also unclear. However, there is some evidence that around 30% protein and 20% lipid in the dry matter of algae selected for aquaculture satisfy larval requirements. Nevertheless, algae (e.g. diatoms) inducing good larval shrimp development have a disconcerting biochemical composition in nutritional terms. Ash content is very high (nearly 40% of dry matter) and does not contribute to the energy supply, as it is indigestible. This accounts in part for the very high ingestion rate during filter-feeding stages. The energy value in algae is around 16 kJ g⫺1 dry weight, with carbohydrate supplying up to one-quarter of this energy. The optimal protein level in a shrimp diet depends on the carbohydrate. Teshima and Kanazawa (1984) showed that the protein requirement in P. japonicus larvae decreases from 55 to 45% when carbohydrate concentration in the diet increases from 5 to 25%. Most of the studies conducted to determine whether algae satisfy larval shrimp requirements relate to lipids, especially phospholipids or HUFA. Shrimp larvae have higher


267

phospholipid dietary requirements than juveniles. Shrimp have some ability to synthesise phospholipids, but not at a sufficient rate to sustain high development during larval stages. The minimum phospholipid requirement in larvae is considered to be 3.5% of diet dry matter (Kanazawa et al. 1985), consisting essentially of phosphatidylcholine and phosphatidylinositol. Mourente et al. (1995) found that a mixture of Tetraselmis plus Isochrysis, providing 8.2% phospholipid, in which phosphatidylcholine represented 1.2% and phosphatidylinostol only 0.1%, was adequate to sustain growth and survival of penaeid larvae. The optimal phospholipid level in a diet also depends on the dietary supply of HUFA (Cahu et al. 1994; Kontara et al. 1997). Algae constitute a substantial source of HUFA. These fatty acids, such as arachidonic acid (ARA, 20:4n-6), EPA (20:5n-3) and DHA (22:6n-3), are essential for larval development. Although shrimp larvae have a greater ability than juveniles to desaturate and elongate 18:3n-3 to 20:5n-3 and 22:6n-3 (Teshima et al. 1992), the conversion rate is too low to meet larval requirements. Several authors have shown that fatty acid content at different larval stages reflects that of the food. These fatty acids can be concentrated in larvae, and some, at low concentrations in algae, can be increased up to 20-fold in larvae (Table 7.4). Different studies have led to the conclusion that the gross composition of an alga (protein, total lipids and carbohydrate) does not account for the differences observed in larval growth and survival. However, the composition of fatty acids could be a predominant factor for differences in larval growth and survival. D’Souza and Loneragan (1999) attributed the good results obtained in Penaeus larvae fed C. muelleri and T. suecica to their high content of ARA and EPA. This was confirmed by intermediate results obtained with Isochrysis sp., which has high DHA content, but low ARA and EPA content. The poor results obtained for larvae fed with D. tertiolecta were attributed to a low content of the three most important HUFA (ARA, EPA and DHA). Thus, a mixture of algae with different fatty acid compositions, such as C. muelleri and T. suecica, provides an adequate fatty acid composition. Larvae fed with such an algal mixture have the average fatty acid content of larvae fed each alga separately.

Table 7.4 Fatty acid contents in algae and in zoea fed these algae, expressed as a percentage of total fatty acids. Penaeus Penaeus Pavlova indicus fed Chaetoceros Penaeus spp. spp. fed lutheri P. lutheria muelleri fed C. muellerib Isochrysis sp. Isochrysis sp.b Total saturated fatty acids Total monounsaturated fatty acids 20:4n-6 20:5n-3 22:6n-3 Total n-6 Total n-3 a

27.7

40.2

17.2

33.2

23.5

32.9

16.9

18.1

21.1

20.5

19.2

17.9

0.3 20.0 7.5 4.2 34.2

2.1 16.2 12.3 6.6 32.5

1.5 23.3 1.8 2.0 25.6

8.0 22.1 5.8 9.7 28.0

nd 0.3 13.7 3.6 45.6

2.0 7.0 21.3 7.7 37.6

From Cahu et al. (1988). From D’Souza and Loneragan (1999). nd, not determined. b

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Algal biochemical composition can be altered by that of the culture medium. D’Souza and Kelly (2000) found that Tetraselmis reared in a low-nitrogen medium had a three-fold higher carbohydrate content and lower protein, lipid and n-3 HUFA content than Tetraselmis grown in a high-nitrogen medium. The gross composition of larvae fed the two diets was not different, but the essential fatty acid 18:3n-3 was 1.6 times as high in larvae fed the second diet, coinciding with faster development of zoea II. Thus, the lower growth observed in larvae fed Tetralsemis grown with a low-nitrogen medium can be attributed in part to the lower protein: energy ratio and mainly to the lower n-3 HUFA content. Moreover, the ratio of n-3 fatty acids to n-6 fatty acids was 3.0 in the algae reared in high-nitrogen medium, whereas it reached only 1.5 in the other algae. This ratio may be important for prawn development. Xu et al. (1994) recorded high survival and development of Penaeus chinensis juveniles fed artificial diet containing linolenic and linoleic acids in a 3:1 ratio. In addition to HUFA, algae supply larvae with vitamins. High concentrations of some vitamins (500 ␮g g⫺1 of diet for ascorbic acid and 300 ␮g g⫺ 1 for ␣-tocopherol) have been found in developing embryos (Cahu et al. 1995), which supports the notion that these micronutrients are of considerable importance during early larval development. Large differences in ascorbic acid levels have been detected in algae, but are more likely to be due to culture conditions than to the species used. Although levels as high as 3800 ␮g g⫺1 dry weight were detected in laboratory-cultured Isochrysis or Chlorella, only one-third of this concentration was found in algae grown in a commercial hatchery (Merchie et al. 1997). Despite the wide range of ascorbic acid concentrations reported in algae cultured in various conditions, it is obvious that microalgae are a rich source of this vitamin for shrimp larvae. Although a concentration of 20 ␮g g⫺1 is sufficient to sustain penaeid juvenile growth, requirements are much higher during larval stages. Merchie et al. (1997) estimated that 130 ␮g of ascorbic acid g⫺1 of diet is needed to sustain requirements related to fast collagen formation during frequent moulting and metamorphosis. The high concentrations found in algae (up to 2 ␮g g⫺1) enhance the resistance of postlarva shrimp to stress conditions and bacterial infections.

7.3.5 Substitution of spray-dried algae or microparticulate compound diets for live algae Attempts have been made to replace live microalgae with spray-dried algae in order to lower hatchery production costs. Biedenbach et al. (1990) showed that total replacement of live algae with spray-dried algae leads to a decrease in growth, metamorphosis and survival. This disappointing result was attributed to an alteration in the physical integrity of algal cells and the rapid loss of soluble nutrients upon placement in water. Nevertheless, it appears that 60% of live algae can be replaced by spray-dried T. suecica. Dried algae are often used in the hatchery as a supplement to live algae or microparticulate diets. Studies have been conducted since the early 1970s to substitute a compound diet for algae. Hirata et al. (1975) reported the first results obtained by feeding P. japonicus larvae with soya-cake particles containing 28% protein and 10% fat. The best survival during zoea stages was obtained with 0.16 mg zoea⫺1 day⫺1. Subsequently, different ingredients were incorporated into particles, such as chicken eggs (Jones et al. 1975), squid meal, mollusc meal (Jones et al. 1979), crab protein (Koshio et al. 1989), yeast (Jones et al. 1987) and fish meal (Teshima et al. 1993). Concurrently, studies conducted with semi-purified


269

casein-based diets have provided a better understanding of the dietary requirements of larvae (Teshima & Kanazawa 1984). The main drawback of compound diets, compared with live prey (including algae), is their low stability in water. Studies have investigated the physical characteristics of microparticles in an attempt to prevent nutrient leaching. Microbound particles were manufactured using binders such as carrageenan, gelatine or zein, but micro-encapsulation using a cross-linked protein has produced more stable particles (Kurmaly et al. 1989). Microparticulate feed may replace algae in the hatchery (Autrand & Vidal 1995), but algae will still be used quite often in co-feeding.

7.3.6 Other roles of algae in shrimp larval growth The beneficial effect of the presence of algae on larval growth and survival has long been known. Kumlu and Jones (1995) showed that a small quantity of algae, in addition to compound diet feeding, improves penaeid larval growth. The concentration of algae was very low in this experiment (15,000 cells ml⫺1), so that a nutritional effect of algae could not be implicated. The authors suggested that this positive effect was caused by alga-induced stimulation of larval digestive enzymes (mainly trypsin). More efficient digestion of a microparticulate diet could account for growth enhancement in larvae reared in co-feeding. Similar results have been obtained in fish: the addition of a very low concentration of I. galbana (20,000 cells ml⫺1) induced 40% improvement in the growth of sea bass larvae fed compound diets as well as 26% survival enhancement. It has been suggested that enhancement of different enzymes (trypsin, alkaline phosphatase and maltase) and improvement of digestive tract maturation are responsible for the improved sea bass larval rearing (Cahu et al. 1998). Other hypotheses have also been postulated to explain the beneficial effect of algal addition in fish and shrimp rearing: algae may stimulate the appetite of larvae by releasing components that act as attractants (Støttrup et al. 1995), or may affect the bacterial population in the rearing water and thus contribute to establishing an early gut microbial flora in larvae (Skjermo & Vadstein 1993).

7.3.7 Feeding microalgae to shrimp juveniles and adults Juvenile and adult shrimp are omnivorous, and microalgae constitute a part of the diet of growing shrimp under natural conditions. Moss and Pruder (1995) showed that the presence of pennate and centric diatoms induced improved growth in Penaeus vannamei reared in intensive ponds. These algae contributed mainly to the particulate organic carbon in intensive ponds, and a highly significant linear relationship was observed between shrimp growth and the particulate organic matter concentration. In extensive ponds, microalgae can supply up to 50% of the feeding ration of shrimp and also provide a means of recycling nitrogen and phosphorus leached from faeces and uneaten feed (Duerr et al. 1998). Nevertheless, the contribution of algae to shrimp nutrition is variable, as the temporal variability in algal cell density follows a bloom and crash cycle. Further ecological research could improve algal productivity and shrimp growth. Benthic microalgae also play an important role in shrimp feeding. Stable carbon, sulfur and nitrogen isotope ratio techniques were used to evaluate the relative importance of algae in the

270


nutrition of two penaeid prawns (Penaeus merguiensis and Parapenaeopsis sculptilis) in Malaysia. Stable isotope analyses suggested that benthic microalgae are the major dietary component for prawns living in tidal creeks (Newell et al. 1995). In a great part of the world, shrimp production is carried out in extensive ponds, where ‘lab-lab’ (a biological plant–animal complex) grows concurrently (Bai & Bensam 1993). A large variety of microalgae is found in ‘lab-lab’, and the dominant species are blue–green algae of the genera Oscillatoria, Phormidium and Spirulina, as well as diatoms of the genera Navicula, Pleurosigma and Nitzschia. Microalgae support shrimp growth better than macroalgae. Moss (1994) showed that shrimp fed a diatom culture composed primarily of Chaetoceros or a culture of Nannochloropsis oculata grew significantly better than those fed the macroalga Ulva or Enteromorpha. In addition, RNA:DNA ratios were significantly greater in muscle tissue of diatom-fed shrimp, which suggests that diatoms make a substantial contribution to shortterm shrimp growth. Some algal species can also be detrimental to shrimp production. For example, toxic effects were suspected in the blue shrimp Penaeus stilirostris exposed to the blue–green alga Spirulina subsala (Lighner 1978). Animals showed necrosis of the epithelial lining of the midgut, dorsal caecum and hindgut gland, resulting in haemocytic enteritis.

7.4 Microalgae as Food for Live Prey The first attempts at culturing live prey for aquaculture mimicked the marine food chain, using phytoplankton to feed zooplankton, which in turn was fed to fish or prawn larvae. A basic method in early research was to enclose and enrich natural water masses, thus encouraging the growth of prey organisms to support cultured fish and shellfish species. Microalgae and animals potentially useful as live prey were identified by such experiments. Among them, the rotifer Brachionus plicatilis is the organism most commonly cultivated and studied today. Some data for B. plicatilis may in fact concern B. rotondiformis, as the two were formerly regarded as a single species. Artemia is used as live prey because of its convenient ‘off-the-shelf’ availability in cyst form. Its use today in larval rearing does not require algal production, as nauplii are not actually grown, but only enriched, i.e. fed to improve their nutritional quality just before being used as live prey. As the hatchery artificial food chain has been simplified to reduce production costs, live prey are now often fed with a single microalgal species, each of which has its own biochemical characteristics. Differences are apparent in the fatty acid profile, sterols and vitamins, whereas protein:lipid:carbohydrate ratios depend on culture conditions as well as the algal species. Thus, the nutritional composition of a given alga offered to a live prey has an influence on growth efficiency as well as on the quality of the prey produced.

7.4.1 Feeding live prey with live microalgae Various authors have studied the optimal feeding rate of live prey in test-tubes containing low animal densities. For the rotifer, Hirayama et al. (1973) found an optimal feeding


271

concentration of around 10 ␮g ml⫺1 for 150 ⫻ 104 cells ml⫺1 of ‘marine chlorella’ (Nannochloropsis), and an ingestion rate of about 1.4 ⫻ 10⫺3 ␮g individual⫺1min⫺1. In optimal conditions, a rotifer population may double or triple daily, rapidly exhausting the algal population. With a filtration rate of up to 10⫺4 ml individual⫺1 min⫺1, the total filtration capacity of 200 rotifers ml⫺1 is equal to the entire rearing volume within less than 1 h. Thus, optimal growth can only be transient, and underfeeding will decrease the growth rate and nutritional status of the animals. Scott and Baynes (1978) found that the mean dry weight per rotifer reached around 600 ng during the first days of culture and then decreased to less than 300 ng after the algal population was consumed. A common means of production is a semi-continuous culture in which a part of the population is withdrawn daily and replaced by algal culture. However, raw algal cultures (around 100 ␮g ml⫺1 dry weight) are generally not concentrated enough to meet the feeding demand. In that case, live prey production rate and density depend on the quantity of food delivered and on algal nutritive quality. However, since the addition of concentrated food could result in detrimental effects on medium quality (oxygen depletion and metabolite toxicity), waste products have to be maintained at low levels. A food supply compromise has then to be found for rotifer rearing, the terms of which are examined later in this chapter. The rotifer B. plicatilis ingests small cells less than 20 ␮m in diameter, with a preference for those between 2 and 12 ␮m. However, another important food selection criterion could be the particle width rather than the mean volume, as rotifers have been observed feeding on Arthrospira trichomes. Conversely, small diatoms with long setae (Chaetoceros) cannot be ingested. Copepod species can feed on diatoms, and the addition of microflagellates may provide better growth (Kraul 1989). Artemia of a given length are able to ingest a wide range of particle sizes; at maximum ingestion rate the same total volume of cells is ingested, regardless of species (Sick 1976). The efficiency of an algal species may also be related to the capacity of live prey to digest cell walls, as mentioned previously for molluscs. Microscopic observation of rotifers feeding on Tetraselmis shows faecal pellets containing undigested and even viable algal cells (Fig. 7.7), which suggests that feeding activity may exceed digestion capacity.

7.4.2 Nutritional value of algae for live prey Various algal species have been compared for their efficiency as food for live prey. The results obtained for these comparisons are only of relative value as they depend on culture conditions. This section will consider the nutritional value of algal species for live prey and, consequently, for fish larvae. 7.4.2.1 Proteins and proximate composition By varying the composition of the alga Brachiomonas submarina grown in a chemostat, Scott (1980) obtained optimal growth/ingestion efficiency of B. plicatilis with algae containing approximately equal amounts of protein, lipid and carbohydrates. According to Frolov et al. (1991), there is a good correlation between protein and lipid content in rotifer and that in the algal diet. The amino acid composition of rotifers is constant and not correlated with the algal amino acid profile. Although the essential amino acid content of algae

272


Fig. 7.7 Rotifer Brachionus rotondiformis fed on Tetraselmis suecica. (Photograph: IFREMER/Robin.)

has never been considered deficient for live prey, some essential amino acids could exist in lower amounts in algae than in live prey, and may reduce nutritive efficiency. Kleppel et al. (1998) found that egg production was poor in copepods fed a strain of Isochrysis unusually deficient in histidine. 7.4.2.2 Fatty acids Most of the literature on the nutrition of live prey concerns fatty acids, which are considered of major importance in larval nutrition. Most marine animals have little or no capability to transform polyunsaturated fatty acids (PUFA) such as linoleic acid or linolenic acid into longer and more unsaturated fatty acids, i.e. HUFA, even though their tissues contain high proportions of n-3 HUFA. The first evidence for the importance of dietary HUFA for marine larval growth and survival came from comparisons of live prey grown on algae or baker’s yeast (Watanabe et al. 1979) or on algal species showing different HUFA levels (Scott & Middleton 1979), or from comparisons of copepods with rotifers or Artemia fed on the same algae (Witt et al. 1984). The ability of rotifers or Artemia to grow on yeast (low lipid content), with an addition of various oil emulsions (during feeding or final enrichment), provided an experimental means for estimating the HUFA requirements of several fish species. These studies indicated that DHA is more efficient than EPA (these have been introduced earlier) as an essential fatty acid for marine fish. A DHA:EPA ratio above 1 and the addition of ARA were considered as optimal for feeding fish larvae (Sargent et al. 1997). The ability to use and incorporate essential fatty acids from algal sources differs widely between prey types, and the DHA:EPA ratio is a key criterion for the efficiency of live prey as food for marine larvae (Table 7.5).


273

Table 7.5 Effect of eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) content (% total fatty acid) of algae on the same contents in prey fed these algae during culture or enrichment.

Alga

EPA DHA Prey

EPA

DHA

DHA: EPA References

Nannochloris Nannochloris Nannochloris Nannochloris Chaetoceros calcitrans Dunaliella tertiolecta Isochrysis galbana Dunaliella tertiolecta Rhodomonas baltica Tetraselmis sp. Isochrysis galbana Pavlova lutheri Cryptomonas Isochrysis (T-iso) Control (starved)

24.9 — 24.9 — 24.9 — 24.9 — 33.7 1.8 0.1 0.3 2.2 25.7 0.4 0.3 11.5 7.9 10.8 0.5 0.9 19.4 28.3 15.5 7.4 3.8 0.5 6.4 — —

16.4 15.5 21.4 6.8 8.3 6.2 6.7 5.6 17.7 9.9 4.4 24.2 7.1 2.9 2.8

10.8 8.6 1.3 — 21.4 12.4 22.9 17.9 32.1 1.7 13.5 11.8 1.5 0.9 —

0.7 0.6 0.1 0.0 2.6 2.0 3.4 3.2 1.8 0.17 3.1 0.5 0.2 0.3 0.0

Copepod nauplii Copepodites Brachionus Artemia (fed 12 h) Tisbe sp. Tisbe sp. Tisbe sp. Tisbe holothuriae Tisbe holothuriae Brachionus plicatilis Brachionus plicatilis Brachionus plicatilis Artemia (fed 24 h) Artemia (fed 24 h) Artemia (unfed 24 h)

Witt et al. (1984) Witt et al. (1984) Witt et al. (1984) Witt et al. (1984) Nanton & Castell (1998) Nanton & Costell (1998) Nanton & Costell (1998) Norsker & Støttrup (1994) Norsker & Støttrup (1994) Reitan et al. (1997) Reitan et al. (1997) Reitan et al. (1997) Thinh et al. (1999) Thinh et al. (1999) Thinh et al. (1999)

Copepods Marine planktonic copepods contain high amounts of n-3 HUFA, which are obtained from their phytoplankton diet in the natural environment. It is questionable whether all species require n-3 HUFA or only C18 PUFA as essential fatty acids. Moreno et al. (1979) showed that Paracalanus parvus has the enzymatic capability to desaturate and elongate 18:3n-3 to n-3 HUFA. Støttrup and Jensen (1990) found that the copepod Acartia tonsa fed with Dunaliella tertiolecta (containing 18:3n-3 but no HUFAs) ceased egg production, while HUFA-containing algae such as Rhodomonas baltica were suitable for egg production. According to Payne and Rippingale (2000), copepod production is positively related to the DHA:EPA ratio in the diet. Harpacticoid copepods are easier to grow than calanoid copepods. Various food sources can be used, which suggests that they do not require dietary HUFA for growth and reproduction. Norsker and Støttrup (1994) found no difference in brood size for Tisbe holothuriae females fed either D. tertiolecta or R. baltica. Harpacticoid copepods contain relatively suitable amounts of 22:6n-3 and ARA, even when raised on a diet deficient in essential fatty acids (Norsker & Støttrup 1994; Nanton & Castell 1998). The fatty acid content of both copepod groups is influenced by dietary fatty acid, but also modified by elongation–desaturation capabilities and/or selective incorporation. The animals will be richer in HUFA if fed on HUFA-rich algae, but will have a higher DHA:EPA ratio than that of their food and are thus eminently suitable diets for marine fish larvae. Rotifers: Brachionus plicatilis Rotifers can also be fed with various food sources, even those nearly devoid of essential fatty acids, such as baker’s yeast. In this case, the fatty acid content will show very low HUFA levels. In an axenic culture of rotifers fed with baker’s yeast, growth improved

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slightly when marine oil was added to the medium (Hirayama & Funamoto 1983). In various experiments comparing the effect of different algae on rotifer culture, population growth appeared to be independent of algal content of n-3 HUFA, while rotifer fatty acid content reflected that of the algae used as food (Frolov et al. 1991). Some elongation of dietary fatty acids was observed in rotifers, but no substantial amount of desaturation products appeared. Moreover, rotifers seem to catabolise more DHA than EPA, so that the DHA:EPA ratio is lower than in their food (Rainuzzo et al. 1994). Unfortunately, algal species providing the best rotifer productivity are not those with the highest HUFA content. For instance, freshwater Chlorella does not contain longer chain fatty acids than C18. Even though Nannochloropsis and Tetraselmis are good food sources for rotifer production, they have a high EPA content, but almost no DHA. Isochrysis affinis galbana has a good DHA:EPA ratio, but provides rather low or unstable rotifer production. In these circumstances, a common practice consists of using a microalgal species suitable for mass production of rotifers, which are then fed a DHA-rich microalga such as I. aff. galbana, before being distributed as food to larvae. Artemia In hatcheries, Artemia are mainly used in their early life-stages, i.e. newly hatched or shortterm enriched nauplii. As nauplii already show a high fat content at hatching, any noticeable modification in their fatty acid profile within a short period requires a high-lipid diet. Artemia show a clear tendency to incorporate less DHA than EPA from dietary fatty acids and demonstrate rapid retroconversion of DHA to EPA (Navarro et al. 1999). Furthermore, nauplii initially contain EPA (high amounts in ‘marine’ strains), but DHA is absent except in a few strains, which contain very low amounts. Since these features all lower the postenrichment DHA:EPA ratio in Artemia, optimal enrichment diets must have very high DHA and low EPA levels. Compared with oil emulsions, 24 h feeding on algae results in a relatively slight HUFA increase, and the DHA:EPA ratio remains low, even though it corresponds to microalgal fatty acid composition (Thinh et al. 1999). Thus, algal enrichment is sometimes performed (mainly with Isochrysis), but oil emulsion enrichment is generally preferred. 7.4.2.3 Other lipid components Crustaceans are incapable of synthesising cholesterol from lower units and require a dietary source of cholesterol or some sterol precursors. Like other crustaceans, Artemia lacks sterol synthesising activity, but can convert some other sterols to cholesterol (Teshima & Kanazawa 1971). Teshima et al. (1981) found that cholesterol is mainly of dietary origin in the rotifer. Nannochloropsis is a good cholesterol source, but freshwater Chlorella or yeast is not, even though rotifers grown on yeast have been successfully used for prawn production. Phospholipids are considered to be essential for the nutrition of prawn and fish larvae (Kanazawa 1993). Unfortunately, live prey can store additional neutral lipids, but phospholipid levels remain constant for a given size of prey, regardless of the diet and/or enrichment products used. The effect of phospholipids on fish seems to be related to their role in intestinal absorption of neutral lipid fatty acids (Geurden et al. 1998). This aspect should also be considered in evaluating HUFA requirements, as oil emulsion enrichment causes a disproportion between neutral and polar lipid contents in live prey.


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7.4.3 Vitamins The nutritional deficiencies that occur when a single algal species is used as a sole food source are often attributed to vitamins. As bacteria are known to produce vitamins, mainly of the B group, studies for the determination of vitamin requirements have used bacteriafree cultures. Tigriopus and Artemia display diverse ability to use different species of algae as food, but none of these species was able to achieve complete development of Artemia or indefinite cultivation of Tigriopus when used alone. However, a mixture of algal species or the addition of a mixture of vitamins can compensate for the nutritional inadequacy of algal food, which suggests that no algal species is able to cover fully the various vitamin requirements (Shirashi 1966). The rotifer Brachionus plicatilis requires vitamin B12, whose complex status in culture tanks depends on the bacterial population (which produces this vitamin at low oxygen levels) and algal content (Hirayama 1987). Maruyama and Hirayama (1993) obtained optimal cultures of rotifer by enriching freshwater Chlorella vulgaris with B12 at 2 ␮g g⫺1 of dry matter. Satuito and Hirayama (1986) have shown that fat-soluble vitamins (A, D, E) have a supplementary effect on rotifers. Algae synthesise ascorbic acid (vitamin C), which stimulates rotifer growth (Satuito & Hirayama 1991) and has a beneficial effect on larval culture. Lie et al. (1997) showed that I. aff. galbana increases ascorbic acid, thiamine and B12 content in rotifers previously grown on yeast (the last two vitamins were incorporated with biotin in ‘f/2’ algal culture medium, used in this study). As well as their nutrient function, some vitamins (e.g. C and E) have been shown to have an antioxidant function. As suggested by Sargent et al. (1997), natural microalgal antioxidants are likely to minimise PUFA peroxidation during the enrichment procedure. Thus, in practical mass cultivation of living prey, vitamin status depends on specific synthesis by the algal species, incorporation of the vitamins into algal culture medium and vitamin production by bacterial populations.

7.4.4 Minerals Seawater is generally considered to be a sufficient source of minerals for most marine organisms. However, in intensive production systems, high biomass levels may lead to the depletion of essential minerals. Watanabe et al. (1983), in an analysis of various components of larval feed, considered that minerals were not a determining factor for dietary value. However, Robin (1989) obtained a significant increase in turbot growth using rotifers enriched with a mineral premix. There is little information in the literature about mineral and trace element requirements, and even less about the effect of algal mineral content on live prey (Lie et al. 1997). The mineral medium used to grow algae should be reappraised to optimise nutritive value through the food chain.

7.4.5 Influence of algae on live feed and larval microbiology Many substances released by marine algae influence the relationship between algae and zooplankton. Van Alstyne (1986) found that exudates of some algal species enhance and

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others inhibit the feeding activity of copepods in cultures. These substances are also involved in the settlement of bacterial microflora within the algal population. As noted above, bacterial communities developing in live prey cultures may provide micronutrients. Douillet (2000) showed that the introduction of bacterial strains into an axenic culture of algae and rotifers could affect rotifer production positively or negatively. In hatcheries, unavoidable contamination by various micro-organisms leads to complex and unstable bacterial associations. In general, modern husbandry methods allow good, stable cultures to be established, but abnormal larval deaths and/or failures in live prey production can still occur. Pathogenic or undesirable bacteria should then be suspected, especially in cases where antibiotic treatment (Gatesoupe 1982) or a sanitary cleaning strategy restored production. A balanced microflora is required in the gut of fish larvae to prevent intestinal opportunistic bacteria from causing disease (Ringo & Birkbeck 1999). In a clearwater system, Nicolas et al. (1989) compared two fish rearing systems using rotifers fed with baker’s yeast and Tetraselmis suecica or Monochrysis lutheri. When M. lutheri was used, turbot larvae ceased to ingest the rotifers and died, while large numbers of bacteria (particularly Vibrionaceae) were found in the rotifer population. Analysis of bacterial communities showed no evidence of a direct relationship between the bacteria in the algal culture and those in the fish larvae. None of the bacteria from the algal culture could be isolated from the corresponding rotifer culture. No common bacteria were observed in the two rearing systems, except in the two batches of larvae. Most of the bacteria from the rotifers were not found again in the larvae, except for some Vibrio. In a later investigation, Austin et al. (1992) showed that T. suecica inhibits bacterial fish pathogens. Bacteria associated with live feed, including pathogenic bacteria, can be transmitted to larval fish during feeding (Benavente & Gatesoupe 1988). As live prey actively ingest bacteria, it is possible to introduce favourable bacteria as a probiotic. Makridis et al. (2000) showed that short-term enrichment with microalgae reduces the total bacteria population harboured by live food, this antibacterial effect amounting to nutritional improvement. The bacteria harboured in the digestive tract of healthy and especially unhealthy larvae belong mainly to Vibrionaceae. Such bacteria are classically found in rotifer and Artemia samples, but not in algal cultures (Verdonck et al. 1994). Microalgae have inhibiting properties on bacteria, particularly on species of the Vibrio group (Salvesen et al. 2000). When algae are added directly to larval tanks (‘green-water’ technique), they not only contribute to maintaining the nutritional quality of live food (Reitan et al. 1993), but also have a positive influence on the settlement of a healthy intestinal microflora in fish larvae by preventing the development of opportunistic bacteria (Skjermo & Vadstein 1993). The direct influence of microalgae on marine fish larvae is discussed further in Section 7.5.

7.4.6 Substitutes for live microalgae As the major drawback in using living microalgae is production costs, their replacement by other food sources is of considerable economic importance for hatcheries. As the algae culture volume requirement is two to three times that of the rotifer rearing volume, most of the area in larval production facilities is used for live feed production, when algae are the only food source.


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Many food sources, including off-the-shelf products and detritus, can be utilised by benthic harpacticoid copepods. These animals may consume the product directly and/or microorganisms growing on it. Mesoplanktonic harpacticoids such as Euterpina or Tigriopus show a more planktonic behaviour, requiring buoyant feed. Microalgae are commonly used by these copepods but can be replaced by yeasts (Fukusho et al. 1980). In contrast, planktonic copepods, such as calanoids, have a greater requirement for live microalgae. Micronised bran has also been used for Artemia rearing, and soluble fish protein concentrate for rotifers (although for a short period only). Most dry products tested as food for rotifer and Artemia are commercially available as single-cell proteins. Spray-dried freshwater algae were tried first, especially Scenedesmus and Spirulina (Person-Le Ruyet 1976) and Chlorella (Hirayama & Nakamura 1976). Dry yeast was also used alone or in formulated diets for rotifers (Gatesoupe & Robin 1981) and Artemia (Robin et al. 1987). Caked live yeast, such as baker’s yeast, has become the most common substitute food for rotifers, either alone or supplemented with living algae or oil emulsions. In general, food efficiency is better with live microalgae than with substitutes, which may have a lower dietary value and/or show potentially detrimental physical properties (buoyancy, leaching from broken cells and packed cells). These properties are conducive to increased waste and thus higher bacterial levels, resulting in an unstable culture and/or contamination by protozoa, with accumulation of packed detritus. This may increase the labour time required for live prey culture and harvesting, and could be disastrous for larval production because of inefficient or collapsed live prey cultures. These problems may be reduced through improvements in feeding methodology. Substitute diets should be carefully prepared, products should be suspended in the water and homogenised, and daily rations fractionated or, even better, delivered continuously. Several artificial diets for rotifer production are available as commercial products, for which a recommended batch culture and feeding methodology (Lavens et al. 1994) help to avoid degradation of the medium. Another type of food consists of marine microalgae produced on a large scale and then concentrated and preserved frozen, spray-dried or chilled. The general purpose of their use is to save labour and space in hatcheries by using products processed in specialised facilities. The nutritional values of freeze-dried and live algae appear to be similar for a number of species. Yùfera and Navarro (1995) obtained good rotifer production with 25–100 mg l⫺1 day⫺1 of freeze-dried T. suecica, whereas 20 years earlier Person-Le Ruyet (1976) used 200–250 mg l⫺1 day⫺1 of the same food. The inhibition of harmful bacteria by microalgae seems effective, even with dried algae (Austin et al. 1992). However, any dead material can be a source of bacterial proliferation. More recently, the use of chilled algae as paste or concentrate slurry has been developed. As observed by Montaini et al. (1995), freezing allows long-term preservation of Tetraselmis cell quality, but results in total loss of viability, whereas concentrated cultures kept in darkness at 4°C show a high capacity for survival. Papandroulakis et al. (1996a) obtained higher rotifer production with these concentrated algae than with frozen algae. According to Yamasaki et al. (1989), similar population growth can be obtained with rotifer fed on either fresh or preserved Nannochloropsis, although frozen algae exhibit three times as much exudation of organic nitrogen (which becomes a pollutant). Heterotrophically grown microalgae and microalgae-like organisms have recently become available on the market. In Japan, freshwater Chlorella, cultured in fermenters on

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organic medium and fortified with vitamin B12, is available commercially in refrigerated form for rotifer culture. This highly concentrated algal biomass (135 g l⫺1) can be used for ‘ultra-high-density’ mass culture of rotifers (Yoshimura et al. 1996). This technique consists of a 2 day batch culture with high food density (3–5 g dry weight l⫺1 day⫺1), resulting in high rotifer concentrations (20,000–25,000 individuals ml⫺1). It requires continuous control of oxygen level, pH (to lower un-ionised ammonia) and accumulation of particulate organic matter in the culture. For enrichment of rotifers and Artemia, such short-duration cultivation does not require cell viability. A high HUFA value and a suitable DHA:EPA ratio are prevalent criteria for live microalgae substitutes. High DHA content can be found in the heterotrophically grown and spray-dried thraustochytrid Schizochytrium (Barclay & Zeller 1996) or the dinoflagellate Crypthecodinium cohnii. Such products can be an alternative to oil emulsions for live feed enrichment protocols (Fig. 7.8). However, regardless of the food used for mass production and enrichment of live prey, live algae are still the best option for strain conservation and production start-up. Microalgae were the first food commonly used for live prey production. However, their importance has decreased since the 1980s, as more cost-effective products can now be substituted as food for rotifer production. Microalgae are still required as food for calanoid copepods, but these cultures have economic disadvantages that limit their use. Increasing the HUFA content of live prey has drastically enhanced the survival and growth of larvae, so it is hardly surprising that current developments in live prey production procedures are mainly focused on providing the n-3 HUFA requirements of larvae. This has led to the extensive use of oil enrichment, which is more efficient than microalgae for attaining high HUFA levels. Microalgae provide other factors than HUFA in live prey production. In particular, their effect on the bacterial environment is now recognised. The ideal algal species, which would be inexpensive to grow and efficient for both live prey production and nutritional quality for larvae, still remains to be discovered. It is likely that new techniques based on ‘off-the-shelf’ products will prevail.

Rotifer grown on Ts

DHA EPA

Rotifer grown on yeast + fish oil Rotifer fed 8 h on Sc Rotifer fed 6 h on oil DHA/EPA = 1 Artemia fed 24 h on oil DHA/EPA = 1 Artemia fed 24 h on oil DHA/EPA = 4 Artemia fed 24 h on 0.4 g l⫺1 Sc Artemia fed 24 h on 0.2 g l⫺1 Sc 0

1

2 3 4 5 Live prey HUFA content (% dry matter)

6

Fig. 7.8 Comparison of docosahexaenoic acid (DHA) and eicosapentaenoic (EPA) contents of Artemia and rotifer using various substitutes to living algae: Ts: chilled Tetraselmis; Sc: spray-dried Schizochytrium; HUFA: highly unsaturated fatty acids. (Data for Schizochytrium from Barclay & Zeller 1996; data for Artemia oil enrichment from Curé et al. 1995; other data from IFREMER-Brest.)


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7.5 Importance of Microalgae in Marine Finfish Larviculture Unlike bivalve and crustacean larvae, which are regular or transient microalgal feeders, most marine fish larvae do not feed directly on microalgae and cannot survive in pure microalgal cultures or on exclusive phytoplankton diets. However, when phytoplankton was included in larval rearing tanks, the survival, growth and food conversion index of more than 40 species were better than in clear-water conditions (Anon. 1970; Howell 1979; Scott & Baynes 1979; Scott & Middleton 1979; Jones et al. 1981; Quinonez Velasquez 1989; Eda et al. 1990; Reitan et al. 1993, 1997; Dhert et al. 1998; Tamaru et al. 1994; Papandroulakis et al. 2000, 2002a, b). The green-water technique (larviculture in an endogenous bloom of phytoplankton and rotifers) and the ‘pseudo-green-water technique’ (larviculture in a tank supplemented daily with exogenous phytoplankton and rotifers), as well as all mesocosm technologies, constitute industrial application of this phenomenon (Divanach & Kentouri 2000). The reasons for the apparently positive effects of microalgae on fish larvae are not yet fully understood. In particular, analysis is difficult because phytoplankton cultures are complex mixtures of suspended (live or inert) and soluble organic and mineral substances. Moreover, the results obtained are often more positive than expected (Tamaru et al. 1994). The following hypotheses have been proposed to explain this phenomenon: stabilisation of or improvement in water quality and light contrast (Naas et al. 1992), the role of direct (via drinking and gill retention) or indirect (via absorption of endogenously enriched prey) nutrition (Moffatt 1981; van der Meeren 1991; Reitan et al. 1993), the micronutrient stimulus for feeding behaviour or physiological processes (Hjelmeland et al. 1988; Cahu et al. 1998); the regulation of bacterial opportunistic populations by antibacterial or probiotic action (in Skjermo & Vadstein 1993), and improvements in rotifer pelagic quality, behaviour and availability (Reitan et al. 1993, 1997; Øie et al. 1997).

7.5.1 Range of microalgal action Positive results have been obtained with a large specific variety of live mature (but not old) microalgal cultures used at low or medium concentration (Divanach & Kentouri 2000), as well as with frozen or lyophilised extracts of microalgae (Scott & Baynes 1978; Papandroulakis et al. 1996a,b; Navarro & Sarasquete 1998). The most spectacular effects are observed during transition from the endotrophic to the exotrophic phase (Table 7.6), particularly for the two ‘mixed’ (i.e. endo–exogenous) and first (i.e. first large zooplanktonic prey) feeding processes. However, endotrophic stages (eggs and prelarvae) and early exotrophic stages are also affected.

7.5.2 Effects on endotrophic larval stages Accounts of positive effects of microalgae on endotrophic larval stages have sometimes been reported, but these are generally unproven. A light microalgal background improves the buoyancy of eggs after handling and/or transfer, as well as the deployment and subsequent survival of newly hatched sea bream prelarvae. Microalgae also reduce the third day

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Table 7.6 Possible early feeding phases in marine fish (shading indicates the predominant feeding mechanism). Life stage

Oocyte

Nutritional mode

Maternal

Important steps

Egg

Prelarva

Larva

Endotrophic 1

Endo–exotrophic 2

3

Exotrophic 4

A Dominant feeding mechanisms

B C D E

1, fertilisation; 2, hatching; 3, mouth opening; 4, resorption of last reserves. A, transchorionic; B, integumental; C, drinking; D, filter-feeding; E, hunting.

post-hatching (DPH) sinking syndrome for sea bream and the lethal consequences of mirror effects of light reflecting walls at the end of the prelarval stage for sea bass and sea bream. The mode of action of microalgae during this phase is still debated, but may be related to a reduction in water loss experienced under stress conditions. Although embryos and prelarvae are dependent on internal nutrient reserves, they interact with the environment through transchorionic (eggs) and integumental (prelarvae) exchanges (water, dissolved minerals and organic substances), as well as through primitive sensorial links provided by auditive capsules, neuromasts, photoreceptors of the pineal gland and future eyes. The major environmental effects on eggs and larvae have been documented in Hoar and Randall (1988). There is thus a range of possible modes through which microalgae could interact with endotrophically feeding stages.

7.5.3 Effects on the yolk-sac drinking stage At mouth opening, fish prelarvae begin a new nutritional relationship with the environment that is more efficient than the previous integumental interaction. Drinking activity (i.e. oral absorption of water) not only contributes to osmoregulation, but also allows intestinal absorption of dissolved organics and ingestion of particulate matter. After mouth opening, hepatic glycogen reserves vary considerably according to nutritional status (Guyot et al. 1993) and environmental changes. 7.5.3.1 Drinking and ingestion of dissolved organics Water uptake, a counterbalance for losses due to hyperosmotic seawater pressure and/or stress conditions, is developed early in marine fish larvae: from 1 DPH in cod (Mangor-Jensen & Adoff 1987), 3–7 DPH in halibut (Tytler & Blaxter 1988; Reitan et al. 1994), 3 DPH in sea bream and 4 DPH in sea bass (Diaz et al. 1994). This activity has also been reported in cod eggs (Mangor-Jensen 1987), for 4 DPH rainbow trout (Tytler et al. 1990) in clear water, and 7 DPH herring in an iso-osmotic environment (Tytler & Bell, in Tytler et al. 1990). Drinking rates range from 0.10–0.27% larval wet weight h⫺1 (7–160 nl larva⫺1 h⫺1) in 20 DPH halibut, with a slight increase in the last part of the yolk-sac stage (Tytler & Blaxter 1988; Reitan et al. 1994), to 0.15–0.59% larval wet weight h⫺1 in cod (Mangor-Jensen & Adoff 1987). Differences in drinking rates for stress compensation have not yet been documented in clear


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and green water. However, some larvae of certain fish, such as the sea bream (personal observations) and halibut (Holmefjord et al. 1993), are very sensitive to any change in water quality during this period and show much better behaviour in green than in clear water. Green water is added empirically to tanks to minimise larval stress and correlated sedimentation during and after transfer and handling. A positive effect of dissolved organic substances on fish larvae has been documented in clear and green water. Absorption of soluble organics through the gut by drinking larvae increases markedly after mouth opening: glucose in sea bream and sea bass (Diaz et al. 1994), glycerol and various neoglucogenic substrates in sea bream (Maurizi et al. 2000), and free amino acids in cod (Fyhn & Serigstad 1987; Fyhn 1989). At correct doses, all of these organics can induce accumulation of large amounts of glycogen in hepatocytes via neoglucogenesis and restore the deficit that occurs in clear-water controls at the end of the prelarval stage. However, none can actually sustain long life. Starved green-water larvae quickly present symptoms of nutritional deficiency (Diaz et al. 1998) and die a few days after the clear-water controls. However, if these few extra days’ survival represent additional time for initiation of successful exogenous feeding, these larvae would seem to have a considerable advantage over their clear-water counterparts. Studies of the influence of microalgae on larval absorption of soluble organics via drinking are still preliminary. As microalgae can absorb nutrients heterotrophically or myxotrophically, produce them photosynthetically, and transform and excrete various soluble organic substances (Brockmann et al. 1983; Admiral et al. 1986), the possibilities for interaction with drinking larvae are multiple, even in the gut. Their role in larval nutrition during this period is probably important. 7.5.3.2 Ingestion of microalgae First ingestion of microalgae begins passively via water intake at mouth opening and becomes progressively passive (filter-feeding) or active (hunting) intake of larger microalgae and microzooplankton. It always precedes the classical first (zooplanktonic) feeding. In the wild or in mesocosms with a natural food chain, microalgae and the greenish remains of semi-digested phytoplankton have been found in the gut of several families of marine fish larvae, such as clupeids (Lebour 1919), engraulids (Scura & Jerde 1977), pleuronectids (Last 1978a), gadoids (Last 1978b) and scophthalmids (Last 1979). In the laboratory or in aquaculture with an oligospecific food chain, ingestion of microalgae by marine fish larvae has been reported in several species, such as the anchovy, Engraulis mordax (Moffatt 1981), menhaden, Brevoortia patronus (Stoecker & Govoni 1984), cod, Gadus morhua (van der Meeren 1991), halibut Hippoglossus hippoglossus (Reitan et al. 1991; Lein & Holmefjord 1992), Diplodus sargus, Sparus aurata, Lithognathus mormyrus and Puntazzo puntazzo (Kentouri 1985). Algal concentrates have systematically been found to be the most dominant prey in the gut 1–10 days post-first feeding (DPF) in P. puntazzo and L. mormyrus (i.e. before intake of copepod nauplii and the rotifer Synchaeta) and as the second most dominant prey (after nude ciliates, tintinids and the rotifer Synchaeta) in the gut of 1 and 2 DPF D. sargus and S. aurata (Kentouri 1985; Kentouri & Divanach 1986; Kentouri et al. 1983). In intensive sea bream larviculture using the pseudo-green-water method, all larvae had Chlorella cells present in the gut when examined under the microscope (Kentouri 1985). Although impossible to count precisely because of their small size (1–2 ␮m in diameter) and large number, these cells never

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represented more than a few per cent of the total volume of ingested material. In yolk-sac halibut, the uptake of Tetraselmis is low before 30 DPH (⬍0.1% larval biomass day⫺1), reaches a peak of 1.2–4.5% larval biomass day⫺1 between 43 and 48 DPH, and then drops again to low values (Lein & Holmefjord 1992). In cod, up to 7% and 40% of larvae ingested Isochrysis galbana and Dunaliella salina respectively, at 1 DPH, and 93–100% ingested Nannochloris atomus at 2 and 3 DPH. In a mesocosm with a variety of phytoplankton and zooplankton organisms, the fraction of algal material larger than 8 ␮m in diameter was 39.2% at 7 DPH and decreased to 12.6% at 12 DPH. In complete darkness, cod larvae were still able to ingest algae, showing a high feeding incidence similar to (or even higher than) larvae on a diurnal photoperiod (van der Meeren 1991). Mechanisms of microalgal ingestion are specific, including passive and active responses conditioned by biological adaptations. In 2–6 DPH cod, small algae (N. atomus, 1–4 ␮m) were found to enter the larval gut as a function of the drinking rate, while concentrations of larger alga (D. salina, 6–10 ␮m) represented 492–7251 times the drinking rate (van der Meeren 1991). In yolk-sac halibut, the clearance rate of Tetraselmis sp. ranged from 100 times (before 30 DPH) to more than 1000 times (between 30 and 43 DPH) and 60–300 times (after 43 DPH) the drinking rate (Lein & Holmefjord 1992; Reitan et al. 1994). Positive correlations were noted between algal size and intergill-arch distance. Ellertsen et al. (1981) considered that microalgal cells (D. salina) enter the mouth cavity by accident and clog the visceral arches before being swallowed. However, van der Meeren (1991) and Reitan et al. (1993) concluded that both cod and halibut are active filter-feeders at this stage owing to specific adaptation of gill-arch spacing and prospective behaviour. As for larger prey, modes of ingestion of microalgae include specific selectivity and quantitative regulation modified by age (a proof of adaptive behaviour). Thalassiosira sp. was found in the mouth cavity of cod, but not in the larval gut (van der Meeren 1991). When offered various prey, 3.9–4.2 mm larvae of the menhaden, B. patronus, began feeding on phytoplankton, dinoflagellates (Prorocentrum micans) and microzooplankton (tintinids), but not on copepod nauplii, which were selected later (Stoecker & Govoni 1984). An increase in D. salina cell density in the medium from 1 to 10 million cells l⫺1 resulted in an increased feeding incidence of only about 20% in 2 DPH cod larvae, but about 40% in 6 DPH larvae (van der Meeren 1991). The most characteristic gut contents in the youngest cod larvae were green spheres (10 ␮m), naked dinoflagellates (20 ␮m) and short chains of Skeletonema costatum, which were actively concentrated from the wild. 7.5.3.3 Digestion and assimilation of microalgae Except for some families (clupeids, engraulids) and to a lesser extent a few other species (Boops salpa, Chanos chanos, Siganus sp.), fish are not biologically equipped to digest microalgae (Juario & Storch 1984). Most microalgae in laboratory tests are not (or only partly) digested. In halibut, the assimilation of ingested Tetraselmis was low through the yolk-sac stage, with no detectable (14C-labelled) respiratory products from the algae and assimilation efficiency below 1% until 55 DPH, and then ranged from 1 to 5% (Reitan et al. 1994). Similar findings were obtained with D. salina after microscopic studies of cod larval gut content (van der Meeren 1991). Conversely, northern anchovy benefited directly and indirectly from the algal supply (Moffatt 1981). In mesocosms with a wider variety of


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microplankton species (algae or other organisms), the filter-feeder adaptations of fish larvae seem to be more efficient. Microalgal cells, naked dinoflagellates and various (greenish, yellowish, brownish) remains were observed in the gut. However, Kentouri and Divanach (1986) noted the importance of microzooplankton [mainly small pelagic ciliates (Amphorella, Stenosemella, Strobilidium) and tintinids (Favella)] as concentrators and predigestors of microalgae in the early larval nutrition of sparids. Similar observations applied to B. plicatilis in green water. Although microalgae probably contribute little energy to larval metabolism at this stage, their ingestion is of major importance for subsequent larval adaptation to the heterotrophic stage.

7.5.4 Resistance to delay in first zooplanktonic feeding Microalgae compensated for delays in successful initiation of first feeding in the sea bream, Sparus aurata (Maurizi 2000). Larvae in clear water showed high cholestase, hepatocytic degeneration, pancreas pathology and depressed glycogenic reserves after a delay in feeding of 8 or 16 h after mouth-opening and needed 16 or 40 h, respectively, more to begin successful feeding and induction of normal intestinal lipid absorption. When first feeding was delayed for 32 h, defects became so great that larvae could not begin to feed at all (the ‘point of no return’) and died at 7–8 DPF, the same time as starved controls. Conversely, even after a 32 h delay in first feeding in green water, larvae did not develop such symptoms of metabolic stress. They ingested rotifers almost immediately after distribution (8 h maximum delay), and could initiate successful intestinal lipid absorption within 40 h after mouth opening and then remain alive, without further feeding for up to 9–10 days (Maurizi 2000). A similar slight increase in survival without feeding was observed with cod in green water (Lein & Holmefjord 1992).

7.5.5 Process and efficiency of first feeding Microalgal background has an important, but specifically different, effect on the timing and intensity of first zooplanktonic feeding. In rearing trials, first (zooplankton) feeding time in sparids was earlier in green than in clear water, and better matched the moment corresponding to mouth opening (Kentouri 1985). With sea bream, 50% of larvae in green water versus 0% in clear water consumed the rotifer B. plicatilis at 5 DPH, and 100% versus 50% at 7 DPH. In clear water, up to 10 days was necessary after mouth opening for all larvae to be engaged in consumption. As a result, there was a high frequency of fasting symptoms (hepatocyte degeneration, cholestase) and a poor (often irreversible) state of health among the population (Maurizi 2000). Halibut larvae showed similar behaviour. At 3 DPF, 50% consumed rotifers in green water compared with 0% in clear water, and 14 DPF was necessary for all batches to show a high feeding incidence on this prey (Naas et al. 1992). Feeding intensity is specific. First-feeding larvae of turbot (Reitan et al. 1993; Øie et al. 1997) and halibut (Reitan et al. 1997; Naas et al. 1992) showed an increased rate of consumption of the rotifer B. plicatilis in tanks containing microalgae rather than clear water. This trend was maintained in 2 and 6 DPF turbot, regardless of the rotifer strain and the type of enrichment (303–400 versus 130–318 rotifers larva⫺1 day⫺1) (Øie et al. 1997).

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Table 7.7 Increased survival (%) with and without microalgae in larval tanks. Species

With algae

Without algae

% Increase

Age

References

Scophthalmus maximus

28–55 5–25 30–36 29–54

4–18 0–2 5–6 6–24

277 1400 500 177

23 DPH 13 DPH 23 DPF 23 DPH

Reitan et al. (1993, 1997) Scott & Baynes (1978) Reitan et al. (1993) Øie et al. (1997)

Mugil cephalus

76–82 34–43

41–54 18–18

66 113

15 DPH 15 DPH

Tamaru et al. (1994) Tamaru et al. (1994)

Sparus aurata

56 ⫾ 16 44 ⫾ 17 21.7

8⫾4 16 ⫾ 6 1.9

600 175 1042

30 DPH 20 DPH 15 DPH

Papandroulakis et al. (2002a) Papandroulakis et al. (2002b) Maurizi (2000)

Hippoglossus hippoglossus Dicentrarchus labrax

30 71 ⫾ 3

1.2 60 ⫾ 4

2400 18.2

21 DPF 32 DPH

Naas et al. (1992) Cahu et al. (1998)

DPH, days post-hatching; DPF, days post-(first) feeding.

However, in 5 DPF sea bream, Papandroulakis et al. (2002b) found only a slight difference in consumption (37–42 ng C larva⫺1 in green water versus 33–43 in clear water, depending on the photoperiod). This trend was also observed in 15 DPF sea bream larvae (343–350 versus 266–414 ng C larva⫺1 in green water versus clear water, respectively). Nonetheless, in terms of the percentage of body weight, larvae in clear water (which were smaller) consumed slightly more than those in green water. Microalgae also play a role in intestinal transit and gut repletion. Øie et al. (1997) found a greater number of rotifers in guts of turbot larvae in clear rather than green water and suggested that a longer digestion time was responsible for the better assimilation rate in clear water. Kentouri (1985) observed similar results for 2–5 DPF sea bream, noting that the ‘gut was distended, giving the false impression of good feeding’. Part of the distension was due to accumulation of empty rotifer lorica at the end or the rectum and a very low rate of excretion.

7.5.6 Effect on survival and growth efficiency at first feeding Improvement in survival at first feeding is the main result of larviculture with microalgae (Table 7.7). For species considered difficult to rear in clear water, such as halibut, turbot or sea bream, the gain is generally greater than 100–500% (Naas et al. 1992; Reitan et al. 1993; Papandroulakis et al. 2002a) and may exceed one order of magnitude (Scott & Baynes 1979). For species considered easy to rear in clear water, such as sea bass or mullet, microalgae enhances survival by 18–113% (Tamaru et al. 1994; Cahu et al. 1998), which is sufficient to improve the cost-effectiveness of production. Differences in death rates were evident at less than 10 DPF and often at 2–5 DPF for difficult species. Improvement in growth efficiency during the rotifer period was another result of microalgal background in larval tanks (Table 7.8). In 10 DPF sea bass, this improved growth was apparent as a 40% increase in weight (as measured in formol-preserved larvae) compared with clear-water controls (Cahu et al. 1998), a 82% increase in wet weight of 20 DPF seabream (Papandroulakis et al. 2002a), between 22 and 75% relative difference in


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Table 7.8 Differences in initial growth with and without microalgae in the larval tanks.

Species

Criteria

With algae

Without algae

Rate

Age

References

Turbot Turbot Halibut Sea bass Sea bream Mullet

SGR (% day⫺1) SGR (% day⫺1) SGR (% day⫺1) FW (mg) WW (mg) SL (mm)

28 28 9.5 na 2.0 4.0–4.4

16 23 1.5 na 1.1 3.5–3.7

na na na 40% 82% na

8 DPF 7 DPF 11 DPF 10 DPF 20 DPF 15 DPH

Reitan et al. (1993) Øie et al. (1997) Naas et al. (1992) Cahu et al. (1998) Papandroulakis et al. (2002a) Tamaru et al. (1994)

SGR, specific growth rate; FW, weight after preservation in buffered saline formaldehyde; WW, wet weight; SL, standard length; na, not available; DPH, days post-hatching; DPF, days post-(first) feeding.

the specific growth rate in turbot at 7 and 8 DPF (Øie et al. 1997; Reitan et al. 1993), and 530% relative difference in the specific growth rate in 11 DPF halibut (Naas et al. 1992). These differences in growth were due to two causes: (a) negative or only slight growth during the first 2–5 days (depending on species) for larvae in clear water compared with fast initial growth for larvae in green water; and (b) lesser growth of larvae in clear water compared with those in green water during the rotifer stage. The efficiency of prey assimilation was also affected by microalgae, although not homogeneously, possibly because of differences in feeding rates. Papandroulakis et al. (2001b) found that daily feeding rates were similar for sea bream in green and clear water for a long photoperiod, and that the overall food conversion index (weight of ingested food/gain of larval biomass) for rotifer carbon was better in larvae reared with microalgae than in clear water (6.3–8.2 versus 12.8–20.1, respectively). However, Øie et al. (1997) found that the ingestion rate for turbot was lower in clear than in green water and that more protein (P) and carbon (C) from rotifers were used by larvae in clear (18–28% P, 12–19% C) than in green water (6–9% P, 4–7% C). Similar inverse correlations between the feeding rate and the conversion index have often been found in aquaculture.

7.5.7 Stimulation of digestive functions and gut flora Early enhancement of digestive and assimilative functions improves the survival and growth of fish larvae and favours the transition to exotrophy. The use of microalgae in tanks increases the production of pancreatic and intestinal digestive enzymes, and improves the quality of gut flora. In sea bass larvae, Cahu et al. (1998) found that I. galbana clone T-iso triggers digestive enzyme production at both the pancreatic and intestinal levels, facilitating the onset of hydrolytic functions of cell membranes and early development of brush-border membranes lining the gut. From 8 to 16 DPH, microalgae present in the sea bass larval culture water result in a marked increase in trypsin activity, whereas amylase and chymotrypsin are not affected. At 26 DPH, alkaline phosphatase and maltase assayed in purified brush-border membranes of the intestine are significantly higher for larvae reared in green water than in clear water. Both stimulations are correlated with better survival and growth efficiency. Similar triggering of enzymatic synthesis was reported by Hjelmeland et al. (1988) for

286


herring larvae. Stimulation of trypsin synthesis was also triggered when larval diets were supplemented with free amino acids (Zambonino Infante & Cahu 1994), suggesting that the large amounts of free amino acids in microalgae may be the cause of enhanced trypsin activity in green-water larvae (Admiral et al. 1986). The addition of microalgae to rearing water modified the bacteriology of larval skin and gut (Skjermo & Vadstein 1993). Unlike skin microflora, which is clearly related to the flora of the water and less affected by algal addition than gut flora, the intestinal microflora of larvae kept in green water differed considerably from that of larvae kept in clear water. It consisted mainly of slow-growing bacteria, together with a smaller fraction of opportunistic bacteria (potential pathogens). Selection of bacteria in the gut was more active in green than in clear water, indicating that microalgae produced substances (e.g. lectins, taxins) that enhanced the ability of certain bacteria to grow in the gut.

7.5.8 Effects on early exotrophic larvae Even after the endo–exotrophic phase, microalgae have a positive effect on larviculture and may increase the resistance of larvae to further stressing or adaptive conditions:

•

• •

23 DPF turbot larvae reared in pseudo-green water (Tetraselmis sp. and I. galbana) showed a lower death rate during a stress test (30 s exposure to air) than those reared in clear water (5–6% versus 29%, respectively) (Reitan et al. 1993). However, no statistically significant difference was found beween treatments following a salinity stress test (Øie et al. 1997). Surprisingly, 15 DPH mullet larvae reared with Nannochloris atomus background showed better growth efficiency than those reared in clear water, despite the higher concentration of un-ionised ammonia (0.13 versus 0.03 mg l⫺ 1) in phytoplankton culture (Tamaru et al. 1994). Sea bass larvae reared with I. galbana clone T-iso showed a 26% improvement in survival over those in clear water when they were weaned early (15 DPF) with compound diets (Cahu et al. 1998).

7.5.9 Indirect effects of microalgae on larvae The indirect effects of microalgae on larvae are mainly related to three causes: water quality and luminosity, the bacteriology of water and rotifers, and the quality and accessibility of rotifers. Improved larval rearing efficiency in tanks with microalgae was initially considered an effect of improved water quality due to a counterbalance of larval respiration (oxygen uptake and production of carbondioxide, ammonia and phosphate) with photosynthesis (uptake of carbondioxide, nitrogen and phosphorus, together with oxygen production) and the stabilisation of fluctuations in pH and carbonate–bicarbonate equilibrium. However, the situation is more complex than first believed. Tamaru et al. (1994) found that the input of algal culture fertilisers increased ammonia levels in tanks with green water. With the green-water technique (based on strong lighting), fluctuations in oxygen and pH are generally higher than in clear water because of difficulties in stabilising the microalgal bloom.


287

This can result in water near the tank bottom being deficient in oxygen even though surface water is oxygen saturated. For this reason, various authors (Scott & Baynes 1979; Tamaru et al. 1994) have considered the possible effects of a modified lighting environment in green water. Reduced light intensity and changes in spectrum, contrast and turbidity, which appear to be the main environmental factors involved, are likely to have an effect on behaviour and thus on efficiency. Addition of microalgae to larval rearing water modifies the bacteriology of the water and rotifers both quantitatively and qualitatively (Nicolas et al. 1989; Skjermo & Vadstein 1993; Salvesen et al. 2000). In green water containing Pavlova lutheri, bacterial density was increased by 45%, and the structure of bacterial organisation was more diversified, with a larger fraction of slow growers and fewer opportunistic (potentially pathogens) fast growers than in clear water. The matured green water also had a more stable microflora, which was less affected by modifications of the organic load or antibiotics than the microflora in clear water (Skjermo & Vadstein 1993). With other microalgae (Skeletonema costatum, Chaetoceros meulleri, Nannochloropsis oculata, Isochrysis galbana, Pavlova lutheri and Tetraselmis sp.), higher levels of bacteria were also associated with slowgrowing bacteria. Except for C. meulleri, which was associated with relatively high densities and a high proportion of opportunistic and haemolytic species, low levels of Vibrio sp. were observed with all algae and mainly with Tetraselmis (Salvesen et al. 2000). The rotifer B. plicatilis, when introduced into tanks together with microalgae, showed a bacterial gut flora close to that of the rearing medium, i.e. with fewer opportunistic bacteria than in clear water (Nicolas et al. 1989). The correlations between the bacteriology of water, rotifers, larvae and larval efficiency indicate the importance of this factor. The importance of microalgae for the production, enrichment and nutritional value of rotifers before distribution has been extensively documented (see Section 7.4). Their role in larval tanks is even more apparent. In cultivation trials, the growth rate of rotifers was positive in all tanks with algae and negative in all tanks without algae (Øie et al. 1997). Their lipid and protein content was maintained or enhanced in tanks with microalgae, but lost quite rapidly in tanks without microalgae (Olsen et al. 1993; Reitan et al. 1994). Therefore, the decline in nutritive value of rotifers in clear water before consumption by larvae could be more marked than improvements in composition made by live prey enrichment before distribution (Øie et al. 1997). Since the biochemical composition of live prey is important in pigmentation and metamorphosis of flatfish (McEvoy et al. 1998; Estevez et al. 1999), it is generally believed that green-water culture helps to improve the success of these two parameters (Støttrup et al. 1995). In addition to a direct nutritional action, microalgae in larval tanks can cause behavioural effects in live prey and fish larvae. In green-water systems, rotifers remain pelagic (i.e. accessible), whereas they sink to the bottom or agglutinate on walls within 3–6 h in clear water. With sea bream, this reduced accessibility becomes critical (i.e. detrimental) during the night and the health condition of the larvae is markedly affected in the morning. Therefore, any delay in feeding at this time can have severe consequences. This problem is currently being solved through the use of both automatic computerised feeding and a long photoperiod (Papandroulakis et al. 2000). Through these behaviour modifications and alterations in prey visibility, green-water culture may also have the indirect effect of reducing larval stress levels, a factor thought to be influential in flatfish pigmentation quality (McEvoy et al. 1998).

288


Improvement in, or preservation of the nutritional value and accessibility of rotifers in larval tanks is probably one of the main reasons for successful larviculture with the greenwater technique.

7.5.10 Future developments Strategic use of microalgae in hatcheries during the very early life of marine fish improves the success of first feeding, a prerequisite for efficient survival, growth and quality in fish larviculture. This phenomenon has led to a reconsideration of the three mechanisms involved in early larval feeding:

• • •

the role of drinking and filter-feeding during the transition from endotrophy to exotrophy; the potential direct effect of dissolved organics from microalgae as contributors (probably neoglucogenic) to feeding autonomy and anti-stress responses; the pretrophic (probably oligotrophic or non-trophic) importance of microalgae as a trigger for both physiological and behavioural processes and bacterial probiotic conditioning of water, rotifers and larval gut.

New fields of research have been opened that create closer links between science and production, and aquaculture and experimental marine biology. However, fundamental knowledge is still needed concerning the mechanisms of alga–fish interaction at the various transition points between endotrophy and exotrophy. This should cover areas such as nutrition, metabolism, physiology, feeding, behaviour, environment and bacteriology.

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Appendix I

Procedures for Assessment of Rotifer Cultures

Routine Procedures Daily routine procedures for examining the state of the rotifer cultures are essential for assuring an appropriate supply for raising marine fish larvae in hatcheries. While there are many options for culture methods, daily routine monitoring of general and specific parameters will help to predict the production scale, identify problems before the cultures collapse and improve culture practices under the specific culture conditions prevailing in a hatchery. Routine practice is based on looking at each culture in the morning before initiating any treatment such as harvesting, feeding, change of water or initiation of new cultures. The observations should be recorded in a log-book to help in tracking problems and in formulating more reliable routine culture practices. The daily examination of the culture should be done at several levels: 1.

2.

Take a general look at the culture tanks. Notice the colour, smell, foam and coloured rings at the rim of the culture tank walls. Record the temperature, salinity, pH and oxygen level. In contrast to published data, rotifer cultures require high concentrations of dissolved oxygen, especially in high-density cultures. Take a general look against the light of 30–40 ml samples in glass beakers, from each culture tank. The observations should reveal whether the samples are clear, milky, clean or dirty. A milky appearance of the culture, several hours after the last feeding (this is relevant in cultures using non-continuous feeding methods), usually indicates the presence of excess food in the culture. It may either indicate excess feeding of the culture or some other problems in the culture that prevented the anticipated filterfeeding process that may have consequently reduced reproductive rates of the rotifers. These include: culture temperatures that are not in the optimal range, low oxygen levels, high relative levels of un-ionised ammonia and non-suitable salinity. A clear appearance of the culture indicates an active culture, but additional microscopic observations (see below) are needed to verify the presence of food in the gut and the presence of eggs. In some cases the milky appearance of the culture medium may be caused by other reasons, such as the proliferation of ciliates (usually Euplotes sp.), flagellates (e.g. Oxyrrhis marina) or, in outdoor cultures, blooms of certain algae that cannot be ingested by rotifers (Chaetoceros sp. or Thalasiossira sp.). This is more typical of semi-continuous extensive cultures that are run for several weeks or months.

Appendix I

3.

4.

5.

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Closer observations using a binocular microscope (at ⫻20–50 magnification) should be made on the 30–40 ml sample, including: (a) How dirty is the culture? Are there many particles? Do they float or sink? Are there any protozoa in the culture? Ciliates such as Euplotes sp., Vorticella sp., Paramecium sp., Uronema sp., Fabrea sp. or flagellates (Oxyrrhis sp.)? Are there any filamentous bacteria or other undesired organisms such as amoebas, nematodes or copepods? (b) Are the rotifers swimming or sinking to the bottom of the examination dish? Are the rotifers clean? Do they stick to each other, forming clumps? Are there dead rotifers and/or empty loricas? Are the guts empty or full? (c) Swimming behaviour. How do the rotifers swim: Quickly? Slowly? Nervously? While fast swimming rates are a sign of good fitness, they do not necessarily indicate optimal reproductive rates. For example, the swimming behaviour of Brachionus plicatilis rotifers at 10°C does not differ greatly from their swimming behaviour at 20–22°C, but their reproductive rates at 10°C are significantly lower than those at 20–22°C. Slow swimming is a good and quick indication of problems in the culture such as starvation, low oxygen levels, high un-ionised ammonia levels and too high temperatures for B. plicatilis rotifers. Nervous swimming may indicate the presence of contaminants or external parasites on the rotifers. Sessile ciliate species such as Vorticella sp. or Zoothanmium sp., flagellates or filamentous bacteria may stick to rotifers’ lorica or may be present in the culture medium and hamper their swimming speed. (d) How many eggs are the females carrying? None, one, two or more? Are there any males? What is the relative abundance of young, immature rotifers? Count the number of rotifers and the number of eggs in 1 ml samples (three to five replicate counts per tank, until the difference between replicate counts does not exceed 10%) and calculate the average number of eggs per female (egg ratio). The egg ratio is a powerful tool for predicting problems in rotifer cultures. A ratio of less than 25% may indicate insufficient feeding of the culture or other problems that will lead to its eventual collapse. The number of rotifers and the number of eggs that they carry are used for calculating the food ration for the next 24 h. The amount will range from 1 to 4 g of wet baker’s yeast for every million rotifers (adult females plus amictic eggs). The exact amount has to be determined empirically for each culture facility. After some experience and feeding trials, a certain rate of daily growth in the rotifer population is achieved by using a specific routine method. This value will serve as an indicator for the stability of the cultures kept in the facility and for calculating the food ratio to be supplied to the culture tanks. However, there is an exception to this general concept in the case of young cultures that exhibit log-phase population growth rates. These cultures show a fast growth in density, an abundance of young rotifers and relatively few females bearing eggs. Remove the debris that has accumulated in the culture tank by stopping the aeration for a few minutes and allowing the suspended material to sink to the bottom. The debris is removed from the bottom outlet. Although live rotifers may also be removed this way, it usually has little effect on the total production.

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6.

Appendix I

Undertake routine inspection of food quality provided to rotifers: (a) Live cultures of algae should be inspected for occurrence of contaminating organisms such as bacteria. The colour and shape of the algal cells is also a good indicator for cultures of Nannochloropsis sp., while motility can serve as an additional good indicator for flagellates such as Isochrysis sp. or Tetraselmis sp. (b) Preserved algae (concentrated chilled pastes or frozen concentrated paste) should be inspected for their colour (spoiled algae can appear brownish) and smell. The correct appearance is gained by experience. (c) Fresh yeast: the expiry date is an important indicator. While the yeast may still look fresh, it may not be appropriate as food for rotifers. Using old yeast may lead to a reduction in the pH of the culture. (d) Freeze-dried yeast should be inspected after suspension in fresh water, but its more difficult to identify quickly problematic batches.

Trouble Shooting Contaminant particles and cleaning of rotifer cultures The main problems in rotifer cultures are the occurrence of contaminant organisms and dirt particles. Contaminant organisms may compete for the food with rotifers, increase ammonia levels through their excretion, reduce the dissolved oxygen levels and be harmful to the fish larvae after they are passed with the cultured rotifers to the fish larvae tanks. Dirt particles may either harbour undesired organisms that will be passed on to the fish larvae tanks, or adhere to the walls in the fish larvae tank, creating an optimal habitat for harmful bacteria. The best way of dealing with these problems is to avoid them, by starting rotifer cultures with a clean inoculum and culturing rotifers in clean seawater. The seawater used for mass cultures should be filtered and sterilised by ultraviolet light or ozone. In most places, sand filtration is used to remove most live organisms in seawater, but this method of filtration does not eliminate eggs or cysts of various sizes, and the best way is to filter the water through 0.2 ␮m water filters. However, this step requires an energy source, either in the form of gravitation or by a pumping system. In general, short-term batch cultures (of about 7–10 days at the most) are easier to maintain free of contaminants than long-term semicontinuous cultures. One of the simplest practices is to stop aeration in the culture tank before harvesting of rotifers and allow the settlement of the dirt particles to the bottom of the tank. If there are also floating particles, the rotifer collection hose should be placed below the upper layer. The collected rotifers should be immersed in chlorine-free freshwater, for 5–10 min, followed by suspension in seawater at the culture salinity. This treatment removes several of the parasites, such as ciliates, flagellates and bacteria, and has little effect on survival of the rotifers. These cleaned rotifers can then be used either as an inoculum for new cultures or for nutrient enrichment before being transferred to fish larvae as food. A simple way to avoid the growth of algae is to cover the rotifer culture tanks. The removal of suspended dirt particles is more problematic. Some particles sink after cessation of aeration in tanks and can be removed easily in tanks with a bottom outlet. Settlement of dirt particles can also be encouraged by placing the aeration tubes a short

Appendix I

303

distance off the tank bottom. The settled dirt can be removed either by siphoning or by a bottom outlet. Floating particles can be removed by siphoning the top layer of the culture. However, most particles are suspended in the water column and cannot be removed easily by sieving, being of the same size as the rotifers or causing the rotifers to cling to them. One solution is to pass the water of the culture tank continuously through a filter. This filter is constructed within a container that encloses synthetic nylon fibres or material similar to that used in scouring pads the culture water is forced through a pipe into the container by an air-lift and exits the filter directly to the culture tank. The tank water content passes this filter-bed, which reaches about 1% of the culture tank volume, several times a day. While some of the rotifers are caught in the filter and lost from the culture, the rest of the rotifers are relatively clean. The filter can either be submerged in the culture tank or stand as a separate unit outside the culture tank. The filter material should be washed once a day and it is advisable to construct two filter beds per tank, with one in use and another one in the process of being washed and dried. Another way of eliminating non-motile dirt particles is by using a settling device, which can be attached on the side of each culture tank and the culture water passed through it once or twice a day.

Oxygen supply Aeration of rotifer cultures is usually done through air-stones that provide fine air bubbles to the water. However, in many cases the total amount of air passing these diffusers is very small. Using open-ended pipes will produce large air bubbles with a small surface to volume ratio, but much more oxygen will be dissolved in the water. Adequate oxygen supply is specifically of importance during nutrient enrichment of rotifers, where rotifers are suspended in very high densities in nutrient-rich media. From experience, the rotifers are not affected by the strong aeration, which may reach tens of litres per minute per tube outlet. While the strong aeration will disconnect the eggs from the adult rotifers, this is not important as the rotifers after this treatment serve only as food for the fish larvae.

Appendix II

Decapsulation Procedure for Artemia Cysts

Hydration 1.

Hydrate cysts by placing them for 1 h in water (less than 100 g l⫺1), with aeration, at 25°C. This will enable the process to start with spherical cysts, which improves the physical contact with the decapsulation solution.

Decapsulation 2.

Prepare the decapsulation solution with: (a) hypochlorite, (b) an alkaline product, and (c) seawater. (a) The hypochlorite solution can be made up with either liquid bleach, NaOCl (fresh product; activity normally ⫽ 11–13% w/w), or bleaching powder, Ca(OCl)2 (activity normally ⫾70%). Use an amount equivalent to 0.5 g active hypochlorite product per gram of cysts (the activity is normally labelled on the package, otherwise it must be determined by titration). (b) An alkaline product is necessary to keep the pH ⬎ 10. Use, per gram of cysts: 0.15 g technical grade NaOH when using liquid bleach. Either 0.67 NaCO3 or 0.4 g CaO for bleaching powder; dissolve the bleaching powder before adding the alkaline product; use only the supernatants of this solution. (c) Determine the amount of seawater required to end up with a final solution of 14 ml decapsulation solution per gram of cysts. Collect the now hydrated cysts on a 125 ␮m mesh sieve, rinse and transfer to the hypochlorite solution. Cool the solution to 15–20°C (e.g. by placing the decapsulation container in a bath filled with ice water). Add the hydrated cysts and keep them in suspension (e.g. with an aeration tube) for 5–15 min. Check the temperature regularly, since the reaction is exothermic; never exceed 40°C (if necessary, add ice to decapsulation solution). The solution will turn brown–red and release foam. When it turns whitish-yellow it is time to stop the reaction. Check the evolution of the decapsulation process regularly under binoculars.

• •

3. 4.

Appendix II

305

Deactivation 5.

When cysts turn grey (with powder bleach) or orange (with liquid bleach), or when microscopic examination shows almost complete dissolution of the cyst shell (after 3–15 min), deactivate the hypochlorite with an equivalent amount of 0.1% Na2S2O3 solution.

Washing 6.

Cysts should be removed from the decapsulation suspension and rinsed with water on a 125 ␮m screen until no chlorine smell is detected anymore. It is crucial not to leave the embryos in the decapsulation solution for longer than strictly necessary, since this will affect their viability. Hypochlorite residues can be detected by putting some decapsulated cysts in a small amount of starch-iodine indicator (starch, KI, H2SO4 and water). When the reagent turns blue, washing and deactivation have to be continued.

Use of the Decapsulated Cysts 7. The decapsulated cysts can be incubated immediately for hatching or used directly as food. Alternatively, they can stored in the refrigerator (0–4°C) for a few days before hatching incubation. For long-term storage cysts need to be dehydrated in saturated brine solution (1 g of dry cysts/10 ml⫺1 of brine of 300 g NaCl l⫺1). The brine must be renewed once or twice after 24 h until the salinity of the brine does not drop significantly.

Appendix III

Enrichment Procedure

Seawater Disinfection (Optional) 1. 2. 3. 4.

Add 1 mg l⫺1 NaOCl (100 ␮l bleach solution per 10 litres of filtered seawater). Incubate for 1h without aeration. Aerate strongly overnight. Add 0.5 g l⫺1 NaHCO3 (dissolved in deionised water and filtered).

Cyst Disinfection 5. 6. 7. 8.

Fill a cylindroconical tube. Incubate cysts at a maximum density of 100 g l⫺1. Leave for 20 min with 200 mg l⫺1 NaOCl (⫾2.0 ml bleach solution l⫺1) under strong aeration. Harvest over a 100–150 ␮m sieve and rinse well with tap water or seawater.

Hatching 9.

Hatch and harvest nauplii as described in Sections 3.5.2 and 3.5.3.

Enrichment 10. 11. 12. 13.

Choose a cylindroconical tank with a volume of clean seawater as to obtain a maximum density of 300,000 nauplii l⫺1. Transfer nauplii to the enrichment tank. Add the enrichment product according to specific guidelines, e.g. 0.6 g emulsion l⫺1 at the start and another 0.6 g l⫺1 after 12 h. Keep the temperature around 28°C and oxygen above 4 mg l⫺1.

Harvesting 14. 15. 16.

Remove all aeration. Collect and rinse nauplii over a submerged 120 ␮m sieve as to keep the nauplii submerged at all times. Use the nauplii as such or proceed to cold storage.

Appendix IV

Web Sites for Culture Collections

Provasoli–Guillard National Center for Culture of Marine Phytoplankton in the United States: ccmp.bigelow.org Centro de Investigaciones Biológicas del Noroeste in Mexico: www.cibnor.org Culture Collection of Algae and Protozoa in the United Kingdom: www.ife.ac.uk/ccap

Taxonomic Index

Acanthopagrus cuvieri, 151, 187 Acanthopagrus latus, 150 Acanthopagrus schlegeli, 8, 9, 187 Acartia, 147, 148, 155, 157, 159–161, 165, 167, 172, 175, 177, 179, 181, 194 Acartia clausi, 150, 156, 157, 164, 167, 178–180, 191 Acartia longiremis, 147, 190 Acartia pacifica, 147 Acartia plumose, 147 Acartia sinjiensis, 147, 159 Acartia teclae, 148, 168 Acartia tonsa, 147, 157, 159–161, 164, 165, 176–180, 192, 193, 273 Acartia tsuensis, 147, 171 Aetideus divergens, 159 Amphiascoides atopus, 150, 182, 183, 186 Amphorella, 283 Amonardia, 150, 163 Anabaena azollae, 240 Anarhichas, 4 Anarhichas lupus, 147 Aphanius, 127, 134 Apocyclops, 187 Apocyclops borneoensis, 151, 187, 188 Apocyclops dengizicus, 188 Apocyclops panamensis, 154, 188 Apocyclops royi, 151, 157, 158, 163, 187, 188 Ardea, 134 Argyrosomus hololepidotus, 148 Artemia, 2, 9–13, 17, 49, 65–111, 122–143, 148, 150, 151, 159, 163, 172, 175, 187, 191–195, 263, 264, 266, 270–275, 277, 278 Artemia franciscana, 76, 77, 98, 107, 108, 111, 112, 131 Artemia (franciscana) monica 76 Artemia parthenogenetica, 75, 96 Artemia persimilis, 76, 77, 96 Artemia salina, 75–77, 166 Artemia tunisiana, 75 Artemia sinica, 76, 111, 112 Artemia tibetiana, 76, 96 Artemia urmiana, 76 Arthrospira, 208–210, 218–220, 271 Arthrospira platensis, 208, 212, 220, 239

Arthrospira maxima, 208, 212 Asplanchna, 27 Bacillus, 34 Bacteriastrum hyalinum, 265 Boops salpa, 282 Brachionus, 19, 21, 23, 27, 263, 266, 270, 273 Brachiomonas submarina, 271 Brachionus calyciflorus, 19, 24 Brahcionus plicatilis, 3, 17–19, 21–35, 37, 39, 40, 44–46, 49–52, 270, 271, 273, 275, 283, 287, 301 Brachionus rotundiformis, 17, 18, 21–23, 25, 28–33, 35, 37, 39, 40, 45, 46, 49–51, 270, 272 Brevoortia patronus, 281, 282 Calanus, 155, 157, 190 Calanus finmarchicus, 146, 157, 159, 172 Calanus helgolandicus, 160, 161 Calanus pacificus, 157 Cancer salinus, 75 Candida, 188 Candida utilis, 109, 110 Centropages, 159, 172, 175, 179 Centropages furcatus, 155 Centropages hamatus, 148, 150, 157, 165, 167, 168, 172 Centropages typicus, 148, 165, 172 Centropristis striata, 234 Chaetoceros, 93, 129, 218–220, 225–227, 229, 264, 270, 271, 300 Chaetoceros calcitrans, 208, 210, 212, 222, 225, 229, 254, 255, 257, 258, 260, 262, 265–267, 273, 287 Chaetoceros calcitrans forma pumilum, 210, 222, 256, 257, 262 Chaetoceros gracilis, 183, 208, 212, 216, 217, 225, 254, 255 Chaetoceros muelleri, 182, 183, 225, 255, 257, 258, 262, 265–267, 287 Chaetoceros pumilum, 208 Chlamydomonas, 25, 32, 129 Chanos chanos, 282 Chlorella, 25, 37, 40, 41, 43, 48,

87, 183, 184, 209, 210, 217–221, 232, 233, 240, 242, 243, 268, 274, 277, 281 Chlorella autotrophica, 256 Chlorella fusca var vacuolata, 219 Chlorella grossii, 208 Chlorella minutissima, 184, 208 Chlorella ovalis, 183, 184 Chlorella pyrenoidosa, 212 Chlorella saccharophila, 219 Chlorella sorokiniana, 219 Chlorella stigmatophora, 30 Chlorella virginica, 208 Chlorella vulgaris, 210, 212, 220, 275 Chroomonas fragarioides, 183, 184 Chroomonas salina, 164 Clarias gariepinus, 104 Clupea harengus, 177, 192 Cocochloris, 87 Conchilus, 27 Coryphaena hippurus, 147, 150 Coscinodiscus wailesii, 166 Coullana canadensis (ie, Scottolana Canadensis), 156, 157, 162 Crassostrea gigas, 256–260, 262 Crassostrea virginica, 255, 259 Crypthecodinium, 209, 210 Crypthecodinium cohnii, 208, 212, 242, 278 Cryptomonas, 273 Cyclops oithonoides, 163 Cyclops strenuous, 146, 156 Cyclops vernalis, 189 Cyclotella, 218 Cyclotella cryptica, 32, 242 Cyclotella nana, 165, 166 Cynoscion nebulosus, 10 Daphnia, 189, 194 Dicentrarchus labrax, 4, 9, 17, 284 Diplodus sargus, 281 Ditylum brightwellii, 166, 176 Dunaliella, 76, 85, 129, 185, 254 Dunaliella primolecta, 256 Dunaliella salina, 183, 184, 208, 219, 233, 239, 282 Dunaliella tertiolecta, 161–163, 185, 208, 232, 256, 258, 265, 267, 273

310

Taxonomic Index

Eleutheronema tetradactylum, 8 Elops saurus, 147, 177 Engraulis mordax, 150, 281 Enteromorpha, 270 Epinephelus, 4, 8, 17, 151, 175 Epinephelus coioides, 147, 172 Epinephelus fuscoguttatus, 147 Epinephelus striatus, 150 Euchaeta elongata, 158 Euchaeta norvegica, 159, 190 Eugerres brasilianus, 151, 187 Euplotes, 33, 176, 180, 300, 301 Eurytemora, 175, 179, 181 Eurytemora affinis, 147, 148, 156, 159, 165, 166, 172, 178 Eurytemora hirundoides, 148, 172, 191 Eurytemora longicornis, 178 Euterpina, 277 Euterpina acutifrons, 150, 157, 182, 183 Eutreptiella, 32 Fabrea, 134, 301 Favella, 283 Fugo rubripes, 17 Fundulus, 147, 177 Gadus morhua, 1, 4, 10, 147, 148, 173, 175, 177, 281 Gladioferens imparipes, 147, 156, 157, 159, 161, 166, 176, 178, 180, 187 Glaucosoma, 175 Glaucosoma hebraicum, 147, 192 Glenodinium, 183, 184 Gobionellus boleosoma, 150 Gonyaulax grindleyi, 159 Halectinosoma curticorne, 163 Heterocapsa triquetra, 192 Hippocampus angustus, 147 Hippocampus subelongatus, 157, 192 Hippoglossus hippoglossus, 4, 10, 147, 148, 281, 284 Hippopus hippopus, 255 Homarus, 1 Huntemannia jadensis, 159 Isochrysis, 48, 166, 227, 228, 230, 262, 264, 265, 267, 268, 272, 274, 302 Isochrysis affinis galbana ‘Tahiti’ (T-iso), 166, 185, 208, 210, 212, 216, 218–220, 222, 227, 229, 242, 254–256, 258, 261, 262, 265, 266, 273–275, 285 Isochrysis galbana, 159, 160, 162, 164–166, 176, 182, 184, 188, 192, 208, 210, 212, 215, 217–220, 227, 230, 254, 255, 258–260, 269, 273, 282, 286, 287

Labidocera aestiva, 178 Lates calcarifer, 4, 8, 17, 147, 148 Leucothrix, 92 Limanda yokohamae, 150 Lithognathus mormyrus, 281 Litopenaeus vannamei, 10, 111 Loligo pealie, 147 Lutjanus argentimaculatus, 147, 175 Lutjanus johnii, 147, 157 Lyngbya, 129 Macrobrachium, 12 Macquaria novemaculeata, 148 Melanogrammus aeglefinus, 1 Mercenaria mercenaria, 255 Metapenaeus, 8 Meteridia longa, 147 Moina, 111, 194 Morone chrysops, 10 Morone saxatilis, 10, 147 Mugil cephalus, 17, 150, 284 Mya arenaria, 255 Mylio macrocephalus, 150 Mytilus, 183, 185 Mytilus edulis, 185, 255, 256, 259 Nannochloris, 166, 189, 273 Nannochloris atomus, 208, 256, 282, 286 Nannochloropsis, 25, 35, 39, 45, 48, 51, 129, 209, 212, 218, 219, 221, 229, 232, 240, 242, 271, 274, 277, 302 Nannochloropsis oculata, 25, 32, 39, 159, 208, 210, 218–220, 254, 270, 287 Navicula, 129, 270 Nereis, 184 Nitocra spinipes, 183 Nitzschia, 218, 219, 185, 242, 270 Nitzschia closterium, 32, 209 Ochromonas, 218 Oithona, 147, 151, 152, 167, 172, 181, 187, 189 Oithona ovalis, 157 Oithona plumifera, 163 Oithona similis, 172 Oscillatoria, 129, 270 Ostrea, 258, 259 Ostrea chilensis, 258 Ostrea edulis, 257–260 Oxyrrhis marina, 164–166, 300, 301 Pagrus auratus, 147, 192 Pagrus major, 4, 8, 9, 17 Paracalanus, 150, 167 Paracalanus parvus, 172, 273 Paralichthys dentatus, 10, 234 Paralichthys flesus, 172 Paralichthys olivaceus, 4, 9 Paramecium, 301 Parapenaeopsis sculptilis, 270

Patinopecten yessoensis, 259 Pavlova, 166, 227 Pavlova (Monochrysis) lutheri, 25, 32, 162, 164, 176, 208, 210, 212, 217–220, 222, 227–231, 236, 242, 254–257, 259–262, 265, 267, 273, 276, 287 Pavlova salina, 208 Pecten fumatus, 259 Pecten maximus, 256–259, 261 Penaeus, 10, 267 Penaeus chinensis, 9, 67, 268 Penaeus esculentus, 263 Penaeus indicus, 8, 266, 267 Penaeus japonicus, 2, 8, 9, 265, 266, 268 Penaeus merguiensis, 8, 270 Penaeus monodon, 8, 111, 112, 265 Penaeus semisulcatus, 19, 265 Penaeus stilirostris, 270 Penaeus vannamei, 150, 265, 269 Phaeodactylum, 185, 223 Phaeodactylum tricornutum, 160, 166, 183, 184, 208, 209, 211, 212, 218–220, 223, 240, 254, 255, 265 Phormidium, 270 Placopecten magellanicus , 255 Platicththys flesus, 148, 172 Platymonas, 165 Platymonas suecica, 183, 184 Pleuronectes americanus, 1 Pleuronectes platessa, 177 Pleurosigma, 270 Porphyridium cruentum, 238, 239 Prorocentrum mariae-lebouriae, 256 Prorocentrum micans, 215, 264, 265, 282 Pseudocalanus, 157, 166, 178 Pseudocalanus acuspes, 190 Pseudocalanus elongatus, 147, 161, 172 Pseudodiaptomus, 147, 172, 181 Pseudo Isochrysis paradoxa, 265 Puntazzo puntazzo, 281 Pyramimonas grossii, 208 Pyramimonas virginica, 208, 231, 254 Recurvirostra, 134 Rhabdosargus sarba, 8 Rhincalanus nasutus, 166 Rhinomonas reticulata, 162 Rhodomonas, 164–166, 211, 212, 218–220, 226, 228 Rhodomonas baltica, 161–166, 176, 182, 185, 192, 208, 222, 265, 273 Rhodomonas reticulata, 208, 211 Rhodomonas salina, 208, 211, 212 Robertgurnenya, 159 Ruditapes philippinarum, 259, 260

Taxonomic Index

Saccharomyces cerevisae, 25, 109, 110 Scenedesmus, 239, 277 Sciaenops ocellata, 10 Schizochytrium, 208, 209, 211, 212, 242 Schizopera elatensis, 150, 181, 183 Scophthalmus maximus (ie Psetta maxima), 1, 4, 11, 17, 147, 150, 192, 284 Scottolana canadensis, 156, 157, 162, 184 Scylla serrata, 18 Seriola quinqueradiata, 9, 17 Siganus, 282 Sillago ciliata, 148 Sillago sihama, 150 Simantherina, 27 Skeletonema, 226, 229, 242 Skeletonema costatum, 160, 162, 165, 166, 182, 208, 209, 211, 212, 218, 219, 220, 222, 226, 229, 230, 254, 255, 261, 262, 265, 282, 287 Skeletonema pseudocostatum, 208, 209 Solea solea, 150 Sparus aurata, 4, 9, 17, 148, 150, 281, 283, 284 Spirulina, 162, 208, 209, 233, 270, 277

Spirulina subsala, 270 Stenosemella, 283 Stichococcus, 87 Stichococcus bacillaris, 256 Strobilidium, 283 Symbiodinium microadriaticum, 255 Synchaeta, 281 Synechococcus, 255 Synechococcus bacillaris, 256 Synechococcus elongates, 30, 32 Temora, 110, 159, 172, 175, 179 Temora longicornis, 147, 148, 150, 157, 166, 167, 172, 191 Temora stylifera, 160 Tetraselmis, 129, 165, 211, 212, 217–220, 226, 230, 262, 264–268, 271, 273, 274, 277, 278, 282, 286, 287, 302 Tetraselmis chuii, 188, 208, 211, 264, 265 Tetraselmis striata, 208, 211 Tetraselmis suecica, 24, 164–166, 182, 185, 208, 211, 212, 217, 221, 227, 231, 236, 242, 254–258, 265–268, 272, 276, 277 Tetraselmis tetrathele, 30 Tigriopus, 181, 275, 277 Tigriopus japonicus, 150, 159, 163, 181, 184, 186 Tilapia, 134

311

Tisbe, 150, 155, 157, 158, 163, 167, 172, 181, 182, 185–187, 192, 273 Tisbe battagliai, 155, 162 Tisbe cucumariae, 158, 163, 182 Tisbe furcata, 150, 162, 164, 179, 184 Tisbe gracialis, 146, 158, 182 Tisbe holothuriae, 153–155, 157, 159, 162, 163, 182, 184–187, 189, 190, 273 Tisbe reticulata, 185 Thalassiosira, 226, 282 Thalassiosira fluviatilis, 166 Thalassiosira nordenskjoldii, 172 Thalassiosira pseudonana, 162, 164, 166, 184, 208, 229, 254, 255 Thalassiosira weissflogii, 160, 166, 176, 215, 266 Trachurus, 9 Tridacna gigas, 255 Ulva, 162, 185, 270 Ulva petrusa, 184 Uronema, 301 Vibrio, 5, 34, 174, 180, 276, 287 Vibrio anguillarum, 109, 110 Vorticella, 33, 301 Zoothanmium, 33, 301

Common Names Index

Anchovy, 150, 281 Asian sea bass, 4, 8, 9, 17, 147, 148 Atlantic halibut, 3–6, 10, 67, 107, 111, 112, 147, 148, 192, 193, 280–285 Australian bass, 148 Black sea bream, 8, 9, 46, 187 Black sea bass, 8, 234 Catfish, 67, 94, 104 Chinese white shrimp, 9, 67, 268 Clams, 255 Cod, 1, 3–5, 10, 67, 147, 148, 159, 171–175, 177, 180, 280–283 Darter goby, 150 Dhufish, 175 Dolphin fish, 147, 150 Dover sole, 150 European sea bass, 4, 9, 17, 284, 285

Halibut, 3–6, 10, 67, 107, 111, 112, 147, 158, 192, 193, 280–285 Herring, 177, 192, 280, 286 Jack mackerels, 9 Japanese flounder, 192 King scallop, 256–259, 261 Kuruma prawn, 2, 8, 9, 265, 266, 268 Manila clam, 259, 260 Menhaden, 281, 282 Mud crab, 18 Mud dab, 150 Mullet, 7, 9, 17, 150, 284, 286 Mulloway, 148 Mussels, 2, 255 Nassau grouper, 150 Olive flounder, 4, 9 Oyster, 2 Chilean, 258 cupped, 256,-260, 262 European, 257–260

Red drum, 10 Red snapper, 147, 175 Sand borer, 150 Sand whiting, 148 Scallop, 255 Sea bass, 3, 4, 9, 111, 253, 269, 280, 281, 284–286 Sea bream, 9, 10, 253, 279–281, 283–285, 287 Seahorse, 4, 10, 147, 148, 281, 284 Silver bream, 8 Spotted sea trout, 10 Squid, 147, 268 Striped bass, 10, 147 Striped patao, 151, 187 Summer flounder, 10 Threadfin, 8 Turbot, 1, 3–5, 11, 17, 34, 47, 67, 109–112, 147, 150, 168, 171, 172, 192, 194, 195, 275, 276, 283–286 Winter flounder, 1 Wolfish, 4, 147

Flounder, 3, 4, 67, 148, 172 Giant clams, 255 Gilthead sea bream, 4, 9, 17, 148, 150, 281, 283, 284, 285 Golden snapper, 147, 157 Grey mullet, 17, 150, 284 Grouper, 4, 8, 9, 17, 151, 171, 172, 175, 187

Pacific white shrimp, 150, 265, 269 Pink snapper, 192 Plaice, 177 Pufferfish, 17 Rainbow trout, 280 Red sea bream, 3, 4, 8–10, 17, 45, 46

Yellowfin sea bream, 150 Yellowtail, 9, 17

Subject Index

Adenosine triphosphate, 80, 214 Aeration, 33, 37, 39, 51, 83, 84, 88, 89, 93, 101–103, 106, 111, 178, 186, 189, 222, 301–304, 306 Airflow, 141, 233 Air-water lift, 88, 89 Algae, 19, 127, 129, 133–134, 171, 206–232, 279–281, 307 Algae, culture of, 2, 6, 35, 233–242, 302 Algae, as food, 4, 8, 11, 24, 25, 27, 32, 37, 39, 41, 43, 47–49, 75, 79, 87, 94, 108, 129, 130, 176, 177, 182, 253–276, 281–287, 300 Algae, freeze-dried/preserved, 25, 31, 32, 35, 87, 88, 276–278, 302 Algal blooms, 133, 171 Alkaline phosphatase, 269, 285 Alkenones, 227 Allelopathic, 208 Allelopathy, 208 Amino acid, 47, 49, 79, 97, 98, 104, 109, 110, 161, 191, 194, 217, 223, 272, 281, 286 Amictic egg, 21, 27, 30, 45, 50, 301 Amictic female, 23, 27–30 Ammonia, 32, 33, 38, 43, 44, 85, 92, 153, 180, 186, 278, 286, 300–302 Amphoteric females, 27 Amylase, 26, 191, 285 Anal somite, 149,153 Antenna, 20, 23, 26, 68–70, 92, 147, 149, 151, 152 Antennule, 68 70, 147, 149, 151–153 Antibiotic, 34, 45, 92, 111, 276, 287 Antifoam, 101 Antioxidant, 194, 263, 275 Arabinose, 223 Arachidonic acid; AA, 47, 48, 107–109, 163, 223, 230, 232, 267, 272, 273 Areal density, 239 Artemia Reference Center, 65 Artificial light, 233, 240 Ascorbic acid, 48, 98, 99, 109, 110, 224257, 268, 275

Asepsis, 234 Aseptic, 235 Asexual reproduction, 27, 28, 31, 35, 156 Ash, 266 Aspartate, 223 Attractants, 269 Auditive capsules, 280 Axenic, 86, 162, 221, 240, 273, 276 β-carotene, 219, 224, 233 Bacillariophyceae, 207, 208, 225–227, 229–230, 254 Bacteria, 5, 20, 24, 31, 33, 34, 37–41, 45, 48, 51, 68, 75, 85–87, 92, 102, 129, 130, 140, 161–163, 180, 182, 206, 207, 209, 235, 236, 240, 254, 262, 268, 269, 275–279, 286–288, 301,302 Baker’s yeast, 32, 33, 38, 46, 48, 163, 272, 273, 276, 277, 301, 302 Batch culture, 36–38, 42, 86, 164–167, 177, 179, 183–185, 212, 213, 236, 277, 278, 302 Batches, 35, 77–79, 87, 95, 103, 139–141, 143, 181, 276, 283, 302 Beer-Lambert, 217, 239 Benthic, 21 algae, 127, 129, 134, 269, 270 copepods, 146, 147, 149, 151, 157, 158, 161, 163, 182, 186, 209, 277 grazers, 161 Bicarbonate, 286 Binders, 269 Bioconversion, 221 Bio-encapsulation, 105, 106, 111 Biosynthesis, 108, 214, 257 Biotechnology, 7, 233, 240 Biotin, 99–109, 224, 236, 275 Bivalve, 221, 254, 255, 257, 259, 261–263, 279 Black disease, 92 Bleach, 37, 42, 102, 128, 140, 304, 305 Bleaching, 128, 304 Bloom, 2, 5, 10, 128, 130, 133, 171, 172, 179, 255, 269, 279, 286, 300

Blue, 129, 214, 270, 305 Bond, 228 Brassicasterol, 225, 227 Brine, 82, 84, 91, 105, 106, 123, 125, 131, 137–140, 305 Brood size, 125, 132, 155, 273 Broodstock, 6, 257–259 Bubbles, 84, 89, 178, 233, 303 Buoyancy, 49, 87, 190, 277, 279 Calanoid, 146–153, 155–167, 171, 172, 175–180, 182, 187, 189–193, 273, 277, 278 Campesterol, 225, 227, 228 Cannibalism, 171, 176 Carbohydrate, 45–47, 49, 207, 208, 217, 222, 223, 257, 266–268, 270, 271 Carbon, 38, 160–162, 189, 206, 215, 217, 220, 228, 233, 235, 269, 285 Carbon dioxide, 38, 206, 217, 220, 233 Carbonate,123, 128, 286 Carboxylic acids, 217 Carnivorous, 158, 159, 163, 253 Carotene, 219, 233 Carotenoid, 111, 153, 191, 193, 194, 207, 240 Carrageenan, 269 Casein, 269 Catabolism, 109, 161, 257 Cell wall, 87, 207, 221, 256, 271 Centric diatoms, 269 Centrifugation, 36, 140, 262, 263 Cephalosome, 147, 149, 151 Chelating agent, 236 Chemostat, 39, 40, 241, 271 Chicken eggs, 268 Chitin, 67, 79, 97, 152, 153, 157, 191 Chlorophyceae, 207–209, 231, 232, 254 Chlorophyll, 134, 208, 214, 215 Chlorophytes, 257 Chloroplast, 214 Cholestase, 283 Cholesterol, 221, 223, 225–227, 274 Chorion, 73, 77, 94, 97, 105, 280 Chromatogram, 227 Chromatography, 223 Chymotrypsin, 285

314

Subject Index

Ciliates, 34, 37, 38, 134, 176, 179, 180, 186, 187, 281, 283, 300–302 Cingulum, 23 Clam, 2 255, 259 Clear water, 276, 279–281, 283–287 Clupeids, 282, 282 CO2, 79, 102, 129, 133, 212, 220, 222, 238, 286 Coefficient of extinction, 217, 239 Cold storage, 78, 111–112, 140, 156, 306 Collagen, 268 Compensation energy, 215 Competitor, 75, 102, 128, 131, 134, 234, 236 Concentrator/rinser, 103, 283 Concentrated Chlorella, 41 Contamination, 79, 137, 179, 189, 213, 234, 235, 276, 277 Continuous culture, 36, 38–42, 44, 51, 186, 187, 213, 238, 239, 271, 302 Conversion index, 279, 285 Conway, 217, 218, 222, 236 Copepod, 3–5, 13, 17, 34, 38, 97, 109, 111, 128, 129, 134, 146–195, 208, 263, 265, 271–273, 276–278, 281, 282, 301 Corn bran, 87, 93 Corona, 20, 21, 23, 24, 26, 27, 29 Cost, 13, 17, 32, 37, 40, 42, 43, 52, 65, 67, 74, 83, 86, 87, 91, 100, 104, 112, 125, 126, 128, 129, 179, 263, 237, 240, 242, 243, 261, 262, 268, 270, 276, 278, 284 Cost price, 237, 238, 239, 242, 243 Crustaceans, 2, 3, 6–13, 18, 37, 67, 86, 103, 106, 107, 191, 221, 254, 263, 274, 279 Cryptophyceae, 207, 208, 228, 232 Cyanobacteria, 207–209, 233, 263 Cyclopoid, 151, 154, 158, 188 Cyst, 5, 7, 12, 17, 65–68, 72–82, 94–98, 100–106, 112, 122, 123, 125, 127, 131, 132, 134, 137, 139, 140, 141, 143, 156, 194, 195, 270, 302, 304–306 Cyst, rotifer 27 Cysteine, 223 D1 protein, 215 Dark-light cycles, 178, 216 Day post-hatching, 279, 280, 282–286 Decapsulation, 67, 79, 82, 94, 102, 105, 106, 304, 305 Decapsulated cyst,94–98, 103, 105, 106, 305 Degassing, 220

Desaturases, 221 Desaturate, 267, 273 Desaturation, 221, 273, 274 Diacylglycerol, 47, 49 Diapause Artemia, 72, 73, 77–79, 81, 82, 131, 137, 139, 140 copepods, 156, 178, 195 Diatoms, 2, 129, 160, 161, 171, 206, 209, 217, 222, 225, 227, 229, 236, 256, 263, 264, 266, 269, 270, 271 Digalactosyl glycerides, 228 Digestion, 11, 24–26, 98, 152, 193, 194, 256, 269, 271, 282–283, 284 Digestive tract, 21, 68, 69, 75, 153, 157, 261, 269, 276 Dilution rate, 39, 213, 238 Dinoflagellates, 2, 24, 87, 159, 208, 242, 255, 278, 282, 283 Diols, 227 Dipalmitoyl phosphatidylcholine (DPPC), 109 Dipterex, 129 Disinfection, 101, 102, 103, 105, 138, 156, 306 Docosahexaenoic acid (DHA), 47, 48, 96, 97, 107–112, 161,163, 176, 181, 190, 192–194, 229–231, 242, 243, 257, 267, 272–274, 278 DHA/EPA, 47, 107, 161, 181, 272–274, 278 DNA, 75, 270 Doubling time, 209, 218, 240 Drinking, 279–282, 288 Dry formulated feed, 253 Dry matter, 43, 237, 266, 267, 275 Dry particles, 253 EDTA, 236 Egg production, 30, 31, 43, 50–52, 155, 156, 158, 160–163, 176, 178, 180, 182, 187, 259, 272, 273 Egg ratio, 31, 43–45, 301 Egg sac, 152, 154, 155, 157, 163, 187, 189, 190 Eicosapentaenoic acid (EPA), 47, 48, 66, 96, 97, 107, 108, 112, 161, 163, 176, 181, 190, 192–194,229–232, 257, 267, 272–274, 278 Electrodes, 215 Elongate, 30, 32, 69, 89, 168, 172, 267, 273 Elongation, 221, 273, 274 Embryogenesis, 160, 258, 259 Embryo, 1, 27, 72, 73, 80–82, 140, 141, 268, 280, 305 Endogenous reserves, 259 Endopodite, 69, 70 Endotrophic, 279, 280

Energy, 25, 49, 74, 77, 81, 101, 103–106, 109, 111, 123, 140, 141, 162, 189, 190, 207, 213–217, 233, 237, 239, 257, 259, 261, 266, 268, 283, 302 Energy conversion, 214 Engraulids, 281, 282 Enrich, enrichment, 3, 6, 19, 32, 35, 37, 42, 45–49, 52, 76, 79, 97, 103, 106–112, 193, 194, 220, 223, 236, 272–278, 283, 287, 302, 303, 306 Enteritis, 270 Environmental factors, 50, 82, 213, 287 Enzymes, 24, 25, 109, 137, 191, 194, 256, 269, 285 Epifluorescence, 256 Essential amino-acid, 97, 110, 223, 272 Essential fatty acid (EFA), 37, 47, 107, 193, 194, 268, 272, 273 Eukaryotes, 208 Eukaryotic, 208 Euryhaline, 19, 21, 22, 219 Eustigmatophyceae, 207, 208, 232, 254 Eustigmatophyte, 222 Exopodite, 69, 70 Exotrophic, 279, 286 Eye, 21, 68, 70, 89, 112, 149, 152–154, 214, 218, 280, 287 f/2, 35, 217, 236, 275 Faecal pellets, 153, 159, 180, 271 Faeces, 90, 269 Fasting, 283 Fatty acid, 3, 37, 43, 47–49, 79, 96, 97, 107, 108, 111, 134, 161–163, 176, 190, 193, 194, 208, 217, 228–232, 257, 266–268, 270, 272–274 Fecundity, 30, 155, 159, 160–162, 180, 182, 186, 190, 258, 259 Feeding, 10, 12, 17, 43, 52, 83, 94, 98, 103, 105, 106, 111, 174, 187, 193, 231, 240, 253, 255, 257–259, 161–264, 267–287 Artemia, 84, 86–88, 91, 93, 106, 110, 112 copepods, 147, 149, 151–153, 157, 159–163, 176–178, 182, 186, 276 filter-feeding, 24, 25, 68, 69, 79, 88, 91, 106, 151, 263, 266, 280, 281, 288, 300 first feeding, 3, 5, 11, 49, 97, 157, 172, 283, 284, 288 microphagus feeding, 24 raptorial feeding, 266 rotifers, 19, 20, 23–25, 33, 35, 40, 47–50, 271, 300, 301 Feeding rate, 25, 92

Subject Index

Female, 21, 23, 27, 29, 30, 32, 43, 51, 69, 70, 72, 81, 132, 152, 153, 155, 156, 158, 159, 162, 163, 181, 182, 187, 189, 258, 273, 301 Fermentation, 219, 240, 243 Fermentor, 242, 243, 277 Fertilization, 1, 2, 29, 31, 72, 125, 129, 130, 131, 134, 155, 261, 262, 280 Filamentous bacteria, 301 Filter, 89–91, 128, 136, 168, 170, 171, 177, 180, 235 Filtering, 68, 86, 106, 132, 168, 171, 172, 180, 234, 253, 261 Filtration, 5, 25, 38, 86, 92, 126, 147, 168, 170, 235, 254, 260, 263, 265, 266, 271, 302 Filtration rate, 25, 86, 92, 126, 265, 271 Finfish, 3, 7–11, 13, 17, 253, 279 Fingerlings, 9, 10, 13 Flagella, 210, 211, 221 Flagellates, 2, 3, 171, 208, 262–264, 300–302 Flatfish, 3, 67, 107, 146, 171, 192, 154, 287 Flocculation, 38, 92, 262, 263 Flora, 34, 263, 269, 276, 285–287 Fluorescence, 214 Fluorescent, 44, 214, 233, 236 Folate, 99, 109, 224 Formulated diets, 4, 6, 7, 11–13, 67, 100, 277 Free amino acid, 49, 79, 97, 98, 104, 109, 191, 281, 286 Freeze-dried algae, 25, 277 Artemia, 112 yeast, 302 zooplankton, 168 Fry, 1, 7–11, 66, 162, 168 Fucosterol, 225, 226 Fungi, 34, 38, 207, 209, 233 Gadoids, 281 Galactolipid, 228, 229 Gamete, 1, 207, 258 Gametogenic, 259 Gastric glands, 21, 24, 25 Generation time, 158, 159, 175, 194 Genetically modified organisms (GMO), 7 Gill-arch, 282 Glucose, 46, 163, 217, 223, 281 Glutamate, 223 Glycerol, 101, 103, 219, 281 Glycogen, 46, 80, 214, 258, 280, 281 Glycogenic reserves, 283 Glycolipid, 228 Gonad, 21, 27, 258, 259 Great Salt Lake, 7, 65–67, 73, 76, 78, 82, 100, 122, 123

Green, 281–283 algae, 129, 270 house, 133, 221, 234 radiation, 214 water, 11, 36, 125, 233, 276, 279, 281, 283–288 Gross composition, 222, 257, 267, 268 Growth, 1, 3, 6, 7, 11, 12, 34, 37, 39, 40, 47, 48, 93, 104, 107, 109, 110, 171, 181, 191, 192, 254–272, 279, 284–286, 288 algae, 36, 127, 129–131, 207, 209, 212, 213, 215–221, 223, 227, 228, 233, 234, 236, 238 Artemia, 78, 84–88, 92, 112, 123, 126, 131, 133, 134, 278 copepods, 156, 158, 160, 176, 208, 273 rotifers, 29, 33, 271, 273–275, 277, 284, 287, 301, 302 Growth rate, 255, 256, 259, 260, 266, 271, 285 algae, 209, 215, 216, 228 Artemia, 78, 84–87 copepods, 176 rotifers, 287, 301 Harpacticoid, 3, 146, 147, 149, 150, 152, 153, 155–159, 161–164, 172, 175, 179, 181, 182–187, 189, 190, 192–194, 273, 277 Harvest, 7, 173 algae, 36 Artemia, 65–67, 73, 76, 77, 81, 83, 84, 91, 93, 94, 100, 101, 103, 106, 112, 122, 123, 127, 131, 134–139, 306 copepods, 164–167, 168, 171, 172, 178, 180, 181, 183–189, 194 rotifers, 37–39, 41, 42, 45, 49, 302 Hatch, 3, 4, 22, 27, 28, 32, 28, 44, 45, 51, 52, 65–67, 72, 77–82, 95, 98, 100–106, 108, 111, 112, 131, 137, 139, 140, 141, 143, 152, 155–158, 160, 163, 171, 172, 181, 187, 190, 193, 233, 253, 274, 279, 305, 306 Hatchery, 1, 2, 4–11, 13, 17, 19, 36–38, 42, 43, 50, 52, 66, 76, 79, 102, 106, 137, 172, 187, 191, 194, 195, 206, 217, 221, 222, 225, 227, 228, 230, 233, 235–239, 242, 243, 253–255, 257, 258, 261–264, 266, 268, 270, 274, 276, 277, 288, 300 Hepatocytes, 281 Herbivorous, 146, 159, 163, 190, 191, 253 Hermaphroditic, 259 Heterokont, 208, 211

315

Hetrotrophic, 129,217, 240–243, 278, 281, 283 Heterotrophs, 209 Histidine, 110, 223, 272 Highly unsaturated fatty acids (HUFA/HUFAs), 43, 47, 107, 108, 163, 182, 192, 221, 230, 257, 266–268, 272–274, 278 Hunting, 47, 104, 281 Husbandry, 6, 193, 276 Hyperosmotic, 280 Hypochlorite, 37, 51, 73, 79, 105, 106, 129, 140, 235, 304, 305, IFREMER, 221, 222, 225, 229–231, 258 Infra-red, 214 Ingestion, 12, 280–283, 285 Artemia, 3, 86, 129, 271 copepods, 159, 160, 162, 163, 176, 256 rotifers, 24, 25, 34, 44, 271 shrimp, 265, 266 Ingestion rate, 24, 25, 44, 86, 159, 160, 163, 176, 265, 266, 271, 285 Inhibitory, 208, 219, 220 Inoculation, 39, 78, 86, 125, 131, 212, 220, 236, 238 Inoculum, 51, 177, 236, 238, 302 Instar, 67, 68, 77, 80, 92, 94–97, 103–106, 111, 131 Integumental, 280 Intergill-arch, 282 Iron, 236 Isochrysideae, 227 Isochronal development, 157 Iso-osmotic, 280 Juveniles, 1, 5, 6, 10, 11, 83, 91, 95, 132, 171, 172, 174, 191, 195, 253, 254, 258, 263, 266–269 Ketones, 227 Kinetic, 212, 217 Lab-lab, 270 Labrum, 69 Larvae, 20 crustacean, 12, 37, 67, 69, 85, 86, 103–108, 112, 253, 263, 264, 269, 270, 279 fish, 1,3,5, 10–13, 17–19, 32, 34, 36, 38, 43, 45, 47–49, 52,65,66,97–99, 105–111, 146, 147, 149, 159, 171, 172, 174, 177–182, 187, 191–194, 206, 233, 234, 240, 243, 253, 254, 271–274, 276, 278, 279, 280–287, 300–303 mollusc, 2, 253–263 Larviculture, 5, 7, 8, 12, 13, 66, 67, 79, 94, 96, 106, 108, 111, 112, 149, 208, 279, 281, 284, 286, 288

316

Subject Index

Larviparous, 258 Lectins, 286 Light, 23, 81, 82, 102, 103, 129, 140, 143, 163, 177, 179, 181, 182, 186, 187, 207, 209, 213–217, 219–221, 223, 229, 232–234, 236, 239–241, 263, 279, 280, 286, 287, 300 intensity, 47, 74, 81, 129, 178, 192, 240 level, 177 Liming, 128 Linoleic, 47, 96, 268, 272 Linolenic, 47, 96, 97, 108, 268, 272 Lipid, 45–49, 79, 92, 96, 97, 106–109, 111, 152, 153, 156, 160–162, 176, 189, 190, 192–194, 207, 217, 222, 223, 228, 229, 257, 259, 262, 266–268, 270–272, 274, 283, 287 Lipisomes, 109, 111 Live prey, 5, 7, 12, 112, 147, 168, 172, 174, 175, 180, 194, 195, 206, 233, 243, 253, 354, 266, 269–272, 274–278, 287 Livestock, 253 Lorica/loricated, 18, 20–22, 26, 45, 284, 301 Lumen, 24, 26, 214 Lux, 39, 81, 100, 102, 178, 187, 214 Lyophilised, 279 Macroalgae, 146, 161, 270 Males, 20, 21, 23, 27–30, 32, 51, 52, 69, 70, 152, 158, 159, 189, 301 Maltase, 269, 285 Mandible, 68–70, 151, 152 Mannose, 46, 223 Mariculture, 8, 41, 261 Mastax, 21, 23, 24, 26 Maxillae, 151, 152, 266 Maxillipeds, 152, 266 Media, 2, 3, 31, 33, 39, 44, 49, 50, 69, 83, 84, 92, 212, 217, 221, 233, 235, 236, 301, 303 Medium, 3, 30, 33–38, 44, 74, 77, 81–83, 85, 87, 103, 156, 176, 187, 189, 190, 212–214, 217, 219, 220, 233–235, 238, 248, 271, 274, 275, 277–279, 282, 287, 300 Membranes, 21, 47, 107, 192, 208, 228, 285 Mesocosm, 233, 279, 281, 282 Mesoplanktonic, 277 Metabolites, 34, 83, 89, 91, 103, 175, 208, 219, 241 Metamorphosis, 3, 255, 263, 268 Metasome, 149, 151 Methionine, 97, 109, 110, 223 Methylenecholesterol, 225–227

Methylporiferasterol, 225, 227 Microalga-consuming, 253 Microalgae, 2, 4, 39, 108, 206–243, 253–288 Microalgal feeders, 279 Microbial flora, 269 Microcapsules, 47, 262 Micro-encapsulated, 65, 108, 242 Microflagellates, 271 Microflora, 34, 86, 130, 276, 286, 287 Micronutrients, 268, 276 Microorganisms, 39, 91, 207–209, 219, 276 Microparticulate, 47, 263, 264, 266, 268, 269 Microsatellite, 22 Microzooplankton, 281, 283 Mictic female, 23, 27–30, 51 Mineral premix, 275 Mineral salts, 236 Minerals, 79, 130, 194, 275, 280 Mixing, 81, 88, 90, 105, 141, 143, 171, 209, 213, 220, 221, 233, 234, 239, 240, 263 Mixis, 28, 29, 31, 52 Mixotrophic, 221, 240 Models, 25, 39, 215, 233 Mollusc, 2, 3, 9, 206, 221, 227, 231, 233, 242, 243, 253–259, 262, 268, 271 Monoacylglycerol, 47, 49 Monochromatic, 217 Monogononta, 19, 21 Monoseptic, 235, 240 Monospecific, 3, 258–261 Monounsaturated, 229–232 Mouth, 2, 3, 5, 11, 12, 17, 19, 23, 24, 68, 152, 175, 195, 233, 256, 280–283 Mouthparts, 67 Mouth opening, 23, 280, 281, 283 NADPH, 214 Nannoplankton, 2, 3, 209 Naupliar, 68, 77, 94, 95, 106, 149, 151–153, 156–159, 162, 172, 176, 181, 263 Nauplius, 2, 265 Artemia, 2–4, 17, 65, 67–69, 71, 72, 77–81, 94–98, 100–106, 108, 110–112, 130–132, 146, 154, 253, 270, 274, 306 copepods, 5, 17, 152, 155–159, 162, 163, 168, 171, 172, 174–178, 181, 182, 186, 187, 190–194, 281, 282 Nematodes, 21, 301 Neoglucogenesis, 281 Neoglucogenic, 281, 288 Neuromasts, 280 Neutral lipids, 190, 193, 223, 274 Niacin, 99, 109, 110, 223, 224

Nitrogen, 38, 40, 50, 81, 83, 92, 130, 143, 160, 161, 163, 189, 207, 217, 236, 268, 269, 277, 286 Nitrogenous, 26, 38, 85, 152 Nucleotides, 214 Nutrients, 17, 32, 33, 49, 86, 87, 105, 129, 130, 134, 137, 171, 213, 217, 222, 223, 228, 238, 257, 268, 281 Nutritional value, 5, 17, 45, 65, 76, 79, 95, 106, 111, 134, 182, 191, 194, 222, 223, 241, 255, 257, 262, 271, 277, 287, 288 Nycthemeral, 216 Off-the-shelf, 262, 270, 277, 278 Oil emulsions, 48, 272, 274, 277, 278 Oil sac, 153, 190 Omnivorous, 163, 191, 253, 263, 269 Operating costs, 261 Optimum temperature, 218, 219 Ornamental fish, 19, 112, 175, 195 Osmoregulation, 26, 74, 280 Osmotic, 101, 191, 219 Ovary, 21, 27, 153 Oviparous, 27, 72, 125, 132, 133 Ovisac, 155 Ovoviviparous, 71, 72, 123 Oxygen, 33, 34, 38, 39, 43, 72, 79, 81–85, 88, 93, 100–103, 111, 124, 126, 129, 131, 133, 137, 143, 171, 178, 180, 189, 206, 208, 215, 220, 233, 240, 241, 271, 275, 278, 286, 287, 300–303, 306 P/I, 215 PAR, 214, 240 Parasite, 18, 146, 174, 180, 301 Parthenogenesis, 27, 29, 156 Partial pressure, 220 Particles, 12, 23–27, 30, 33, 67, 68, 83, 85–93, 106, 128, 130, 143, 149, 152, 159, 214, 234, 235, 255, 264, 268, 269, 271, 301–303 Path length, 217, 238–240 Pathogenic bacteria, 5, 34, 39, 276 Pavlovol, 227, 228 Pelagic, 147, 149, 157, 189, 190, 209, 279, 283, 287 Pelleted, 12 Penaeid, 3, 4, 8–12, 17, 105, 112, 233, 253, 263–265, 267–270 Pennate diatoms Peridinians, 206 Peroxidation, 275 pH, 32, 33, 37–39, 43, 44, 81, 83, 84, 100–102, 126, 128, 129, 133, 179, 189, 214, 220, 241, 278, 286, 300, 302 Pheromone, 27, 29

Subject Index

Phosphatase, 269, 285 Phosphatidylcholine, 109, 267 Phosphatidylinositol, 267 Phospholipid, 47, 107–109, 190, 194, 228, 229, 266, 267, 274 Phosphorus, 129, 130, 189, 217, 236, 269, 286 Photo-autotrophic, 129, 209, 213, 218–221, 233, 238, 240, 241, 261 Photoautotrophs, 209 Photobioreactors, 237–239, 240–243 Photon-flux density, 214 Photochemical, 208, 214 Photoinhibition, 215 Photons, 214 Photooxidative, 220 Photoperiod, 178, 182, 187, 282, 285, 287 Photoprotection, 236 Photoreceptor, 26, 280 Photosynthesis, 133, 213, 215, 217, 220, 233, 236, 281, 286 Photosystem, 214, 215 Phototactic, 103, 186, 103 Phototaxis, 178 Phytoplankton, 2, 5, 10, 123, 126, 127, 130, 147, 159, 160, 162, 168, 171, 174, 176, 178–182, 192, 206, 215, 221, 223, 229, 233, 235, 253–255, 257, 261, 270, 273, 279, 281, 282, 286, 307 Phytosterols, 221 Pigment, 25, 27, 47, 74, 79, 84, 105, 107, 153, 193, 241 Pigmentation, 108, 110–112, 191–193, 287 Plankton, 5, 19, 51, 168, 172, 174, 253, 263 Planktonic, 18, 21, 24, 146, 149, 151, 157, 158, 163, 181, 273, 277 Pleuronectids, 281 Plurispecific, 261 Polar lipids, 49, 107, 109, 193, 228, 274 Polycarbonate, 40 Polysaccharides, 207 Ponds, 1, 3–5, 10, 19, 28, 36, 37, 66, 67, 75, 83, 91, 123–135, 137, 139, 156, 171, 172, 174, 178, 179, 194, 233, 263, 269, 270 Population dynamics, 74, 123, 126 Post-larva, 11, 13, 67, 257, 263 Prasinophyceae, 207, 208, 237, 230–232, 254 Prawn, 2, 4, 9, 67, 106, 263, 268, 270, 274 Predator, 1, 5, 18, 20, 75, 83, 93, 104–106, 123, 124, 127–129, 131, 133, 134, 147, 149, 159, 168, 171, 178, 180, 234, 263

Predigestors, 283 Prelarvae, 279, 280 Prelarval stage, 280, 281 Prey size, 24 Probionts, 34 Probiotic, 18, 34, 45, 276, 279, 288 Production systems, 37, 40, 43, 153, 174, 195, 206, 237–239, 243, 275 Productivity, 5, Prokaryotes, 208, 235 Protein, 11, 37, 39, 45–47, 49, 80, 87, 88, 95, 97, 98, 109–111, 152, 153, 160–163, 182, 190, 194, 207, 214, 215, 217, 222, 223, 257, 266–271, 277, 285, 287 Protein enrichment, 109, 110 Protonephridia, 21, 24, 26 Protozoa, 33, 38, 86, 176, 277, 301, 307 Prymnesiophyceae, 207–209, 227, 229, 231, 254 Prymnesiophyte, 222, 223, 227, 263 Pseudocoelom, 19, 21, 26 Pseudo-green water, 279, 281, 286 Polyunsaturated fatty acids (PUFA), 47, 162, 163, 192–194, 221, 229–232, 241, 242, 259, 272, 273, 275 Pyridoxine, 99, 109, 223, 224 Quanta R0, 30 Red, 27, 68, 84, 133, 153, 214, 238, 304 Reproduction, 258, 259 Artemia, 72, 78, 123, 125, 126, 132–134 copepods, 155, 156, 160, 162, 190, 191, 273 microalgae, 207 rotifers, 27–35, 28, 40, 45, 48 Reserves, 68, 81, 95, 103, 104, 140, 141, 152, 153, 253, 258, 259, 280, 283 Residence time, 192, 238 Resting egg copepods, 156, 168, 174 rotifers, 21, 22, 27–32, 34, 38, 43, 44, 50–52 Retinal, 192, 193 Retinol, 223, 224 Riboflavin, 99, 109, 110, 223, 224, 257 Rice bran, 87, 91, 93, 96, 98, 176, 177, 180 RNA, 80, 270 Rotenone, 5, 129, 168, 180

317

Rotifer, 3–6, 8–11, 13, 17–52, 94, 108, 134, 159, 168, 172, 175, 179, 181, 186, 187, 191–194, 242, 243, 270–279, 281, 283–288, 300–303 Saline lakes, 122 Salinity, 18, 22, 29, 31–33, 35, 37–39, 43, 45, 50, 52, 72, 74–76, 78, 80–84, 92, 93, 100, 101, 110, 112, 122–134, 137, 139, 140, 159, 175, 179, 186, 191, 207, 209, 212, 213, 219, 220, 286, 300, 302, 305 Salmonids, 1, 6, 12, 146, 253 Salt production, 123, 124, 131 Saltwork, 66, 75, 123–125, 127, 131 Sand filter, 234 Saturation, 32, 33, 84, 176, 215, 220 Scophthalmids, 281 Seawater, 2, 3, 34, 35, 37, 38, 51, 67, 80, 81, 84, 90–93, 101, 123, 124, 129, 133, 137, 171, 174, 176, 177, 182, 186, 219, 220, 254, 258, 275, 280, 302, 304, 306 Self-shading, 216, 217, 236, 240 Setae, 68, 149, 151, 152, 271 Sexual reproduction, 27–32, 34 Shear forces, 221 Shelf life, 137, 140, 141, 143, 243, 262, 263 Shrimp, 2–4, 6, 8–13, 17, 19, 65–67, 79, 94, 105–108, 112, 124, 125, 128, 129, 135, 137, 162, 206, 221, 223, 242, 253, 263–270 Silicon, 101, 180, 217, 236 Single-cell-proteins (SCP), 87 Sinking syndrome, 280 Sitosterol, 225–227 Small larva fish, 253 Small-mouthed, 17, 19, 175, 195, 233 Sorbitol, 219 Soybean, 93 Soy pellets, 87, 91, 96 Soy-cake, 268 Spat, 253, 257, 258, 261, 262 Specific growth rate, 209, 215, 285 Spermatophore, 152, 155 Spray-dried, 268, 277 Starch, 207, 305 Senescence phase, 213, 223 Steady state, 213 Sterilisation, 235 Sterilising, 235 Sterol, 47, 49, 221, 223, 225–228, 257, 270, 274 Stomach, 11, 21, 24–26, 255

318

Subject Index

Storage, 220, 259, 263 Artemia, 77, 78, 81, 82, 87, 98, 106, 111, 112, 137, 139, 140, 143, 305, 306 copepods, 156, 174, 180, 181, 190 rotifers, 49–51 Strains, 3, 7, 28, 31, 32, 34, 35, 38, 39, 49, 50, 52, 72, 74, 76–79, 81, 82, 84, 94, 96–98, 101, 107, 108, 125, 131, 139–141, 143, 209, 236, 274, 276 Stress, 45, 47, 76, 84, 93, 107, 110, 112, 129, 133, 193, 221, 265, 268, 280, 281, 283, 286–288 Substrates, 85, 86, 126, 207, 213, 217, 240, 281 Survival, 1, 3, 6, 11, 12, 38, 47, 48, 50, 74, 78, 81, 85, 86, 91, 93, 94, 110–112, 133, 135, 156, 157, 159, 162, 172, 180, 181, 191–193, 254, 258–261, 263–265, 267–269, 272, 277–279, 281, 283–286, 288, 302 Swim, 11, 93, 104, 133, 158, 186, 301 Swimming velocity, 44 Taxins, 286 Taxonomic, 20, 21, 22, 24, 228 Telopodite, 69, 70

Thiamine, 98, 223, 236, 275 Thiosulphate, 235 Thorax, 149, 152, 155 Thraustochytrid Thraustochytriidae, 208 Thylakoid, 214, 228 Triacylglycerol, 47–49, 107, 190 Tintinids, 281–283 Tocopherol, 110, 223, 268 Toxins, 44, 213, 235 Trace elements, 79, 236 Transchorionic, 280 Transmission, 239 Transport, 1, 49, 84, 93, 135, 136, 180, 181, 187 Trichomes, 271 Triglyceride, 228 Trophi, 21, 23, 24 Trypsin, 269, 285, 286 Tryptophan, 223 UV light, 106 Ultraviolet, 214 UNIK filter, 168 Unsaturation, 228 Urosome, 147, 149 Veligers, 255 Viability, 79–81, 123, 137, 139, 140, 156, 180–182, 258, 277, 278, 305 Vibrionaceae, 276

Vitamin, 37, 48, 79, 98, 99, 109–111, 162, 182, 191, 193, 218, 221, 223, 224, 236, 255, 257, 268, 270, 275, 278 Vitamin enrichment, 48, 109 Vitellarium, 21, 26, 27 Vitellogenesis, 259 Wavelength, 214, 217 Wax esters, 49, 190, 193 Weaned, 286 Welded-wedge filter, 90, 91, 128 Yeast, 24, 32–34, 37, 38, 43, 45–48, 87, 88, 108, 162, 163, 254, 255, 262, 268, 272–277, 301 Yellow, 214, 283, 304 Yields, 65, 83, 87, 91, 94, 102, 123, 125, 128, 132, 237, 238, 258 Yolk-sac, 159, 172, 280, 282 Zein, 269 Zoea, 18, 253, 263–268 Zooplankton, 1–5, 10–12, 18, 23, 98, 107, 108, 144, 148, 171, 172, 174, 180, 181, 184, 190, 192–194, 233, 270, 275, 279, 281–283

Live Feeds in Marine Aquaculture

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Design and Operating Guide for Aquaculture Seawater Systems (Developments in Aquaculture and Fisheries Science) (Developments in Aquaculture and Fisheries Science)

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Methods in Reproductive Aquaculture: Marine and Freshwater Species (Marine Biology)

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