Striped Bass and Other Morone Culture, Volume 30 (Developments in Aquaculture and Fisheries Science)

Developments in Aquaculture and Fisheries Science, 30 STRIPED BASS AND OTHER MORONE CULTURE D E V E L O P M E N T S ...

Author: R.M. Harrell

69 downloads 935 Views 20MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!

Report copyright / DMCA form

DOWNLOAD PDF

Developments in Aquaculture and Fisheries Science, 30

STRIPED BASS AND OTHER MORONE CULTURE

D E V E L O P M E N T S IN A Q U A C U L T U R E AND FISHERIES SCIENCE The following volumes are not available anymore: 5, 6, 10-12 and 14 1.

F A R M I N G M A R I N E ORGANISMS L O W IN T H E FOOD CHAIN

A Multidisciplinary Approach to Edible Seaweed, Mussel and Clam Production by P. KORRINGA 1976 xvi + 264 pages 2.

F A R M I N G CUPPED OYSTERS OF T H E GENUS CRASSOSTREA A Multidisciplinary Treatise by P. KORRINGA 1976 x + 224 pages

3.

F A R M I N G T H E FLAT OYSTERS OF T H E GENUS OSTREA

A Multidisciplinary Treatise by P. KORRINGA 1976 xiv + 238 pages 4.

F A R M I N G M A R I N E FISHES AND SHRIMPS

A Multidisciplinary Treatise by P. KORRINGA 1976 xii + 209 pages

7.

MUSSEL C U L T U R E AND HARVEST: A N O R T H A M E R I C A N P E R S P E C T I V E edited by R.A. LUTZ 1980 xiii + 350 pages

8.

C H E M O R E C E P T I O N IN FISHES edited by T.J. HARA 1982 x + 434 pages

9.

W A T E R QUALITY M A N A G E M E N T FOR POND FISH C U L T U R E by C.E. BOYD 1982 xii + 318 pages

13.

B I O E C O N O M I C S OF A Q U A C U L T U R E by P.G. ALLEN, L.W. BOTSFORD, A.M. SCHUUR and W.E. JOHNSTON 1984 xvi + 351 pages

15.

CHANNEL CATFISH C U L T U R E

edited by C.S. TUCKER 1985 xvi + 657 pages 16.

17.

S E A W E E D C U L T I V A T I O N FOR R E N E W A B L E RESOURCES editedby K.T. BIRD and P.H. BENSON 1987 xiv + 382 pages DISEASE DIAGNOSIS AND C O N T R O L IN N O R T H A M E R I C A N MARINE A Q U A C U L T U R E

editedbv C.J. SINDERMANN and D.V. LIGHTNER 1988 xv + 412 pages 18.

BASIC FISHERY SCIENCE P R O G R A M S : A C O M P E N D I U M OF M I C R O C O M P U T E R P R O G R A M S AND MANUAL OF O P E R A T I O N S by S.B. SAILA, C.W. RECKSIEK and M.H. PRAGER 1988 iv + 230 pages

19.

C L A M M A R I C U L T U R E IN N O R T H A M E R I C A edited by J.J. MANZI AND M. CASTAGNA 1989 x + 462 pages

20.

DESIGN AND O P E R A T I N G GUIDE FOR A Q U A C U L T U R E S E A W A T E R SYSTEMS by J.E. HUGUENIN and J. COLT 1989 iv + 264 pages

21.

SCALLOPS: BIOLOGY, E C O L O G Y AND A Q U A C U L T U R E editedby S.E. SHUMWAY 1991 xx + 1095 pages

22.

F R O N T I E R S OF S H R I M P R E S E A R C H edited by P.F. DeLOACH, W.J. DOUGHERTY, and M.A. DAVIDSON 1991 viii + 294 pages

23.

M A R I N E S H R I M P C U L T U R E : P R I N C I P L E S AND P R A C T I C E S by A.W. FAST and L.J. LESTER 1992 xvi + 862 pages

24.

T H E MUSSEL MYTILUS: E C O L O G Y , PHYSIOLOGY, G E N E T I C S AND C U L T U R E by E. GOSLING 1992 xiv + 589 pages

25.

M O D E R N M E T H O D S OF A Q U A C U L T U R E IN JAPAN (2ND REV. ED.) edited by H. IKENOUE and T. KAFUKU 1992 xiv + 274 pages

26.

P R O T O Z O A N PARASITES OF FISHES by J. LOM and I. DYKOV,~ 1992 xii + 316 pages

27.

A Q U A C U L T U R E W A T E R REUSE SYSTEMS: E N G I N E E R I N G DESIGN AND M A N A G E M E N T edited by M.B.TIMMONS and T. M. LOSORDO

28.

F R E S H W A T E R FISH C U L T U R E IN CHINA: P R I N C I P L E S AND P R A C T I C E edited by J. MATHIAS and S. LI 1994 xvi + 446 pages

29.

P R I N C I P L E S OF SALMONID C U L T U R E edited by W. Pennell and B.A. Barton 1996 xxx + 1040 pages

30.

STRIPED BASS AND O T H E R M O R O N E C U L T U R E edited by R.M. Harrell 1997 xx + 366 pages

Developments in Aquaculture and Fisheries Science, 30

S T R I P E D B A S S A N D OTHER M O R O N E CULTURE

Edited by

R E G I N A L M. H A R R E L L University of Maryland System, Horn Point Environmental Laboratory, Centerfor Environmental and Estuarine Studies, Cambridge, MD, U.S.A.

1997 ELSEVIER Amsterdam-

Lausanne - New York - Oxford - Shannon - Tokyo

ELSEVIER SCIENCE B.V. Sara Burgerhartstraat 25 P.O. Box 211, 1000 AE Amsterdam, The Netherlands

Library

Striped

oF C o n g r e s s

Cataloging-in-Publication

bass and o t h e r morone c u l t u r e / e d i t e d by R e g i n a l M. H a r r e ] ] . p. cm. - - ( D e v e ] o p m e n t s in a q u a c u ] t u r e and f i s h e r i e s science

; 30) Inc]udes bib]iographica] references ISBN 0 - 4 4 4 - 8 2 5 4 7 - 9 1. S t r i p e d b a s s . 2. F i s h - c u ] t u r e . II, Series. SH351.B3S76 1997 639.3'7732--dc21

ISBN

Data

and

index.

I.

Harre]],

Regina]

M.

96-29983 CIP

0-444-82547-9

9 1997 Elsevier Science B.V. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without the prior written permission of the publisher, Elsevier Science B.V., Copyright & Permissions Department, P.O. Box 521, 1000 AM Amsterdam, The Netherlands. Special regulations for readers in the USA - This publication has been registered with the Copyright Clearance Center Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923. Information can be obtained from the CCC about conditions under which photocopies of parts of this publication may be made in the USA. All other copyright questions, including photocopying outside of the USA, should be referred to the copyright owner, Elsevier Science B.V., unless otherwise specified. No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. This book is printed on acid-free paper. pp. 185-216, 329-3: i6: Copyright not transferred. Printed in The Neth erlands

PREFACE Striped bass and other Morone culture has been successful in the United States since the mid 1960s, primarily for the purpose of population enhancement and establishing recreational fishing opportunities. However, with the decline in coastal populations of striped bass and the increasing success of commercial aquaculture with other species, the past decade has seen a tremendous growth in the commercial foodfish production of Morone, particularly hybrids of the genus. This growth has developed into what is today one of the fastest growing and expanding segments of the U.S. aquaculture industry. This book is a comprehensive compilation of the biology and rearing of Morone and its hybrids. It is intended to be an in-depth analysis of the scientific aspects of Morone culture, not a "techniques manual" as such. The latter already exists and is frequently referred to throughout the text. In recruiting the authors for this book I specifically requested they steer away from the "how to" aspects and focus on the science and biology. I think they have done an excellent job. I feel I was fortunate to be able to assemble some of the best Morone researchers in the country -- scientists who not only conduct good investigative research but also have a feel for the practical aspects of husbandry. Thus the reader will be able to see that even though much of the book is directed toward practicing scientists and graduate students, it also contains a wealth of pragmatic information. Many people have contributed to the completion of this work. I wish to personally thank each of the contributors for their efforts. A special thanks is extended to Deborah Weber for the tireless reviewing and constructive suggestions on all the chapters. I am indebted to the many external reviewers who took the time to make suggestions to improve the scope and content of the various chapters, especially Jim Anderson, Paul Bowser, Gary Carmichael, John Colt, Jim Easley, John Grizzle, Eric Hallerman, Frank Hetrick, John Hochheimer, Steve Hughes, Patricia Mazik, Tom Rippen, Bill Simco, Ted Smith, Joseph Soares, Jennifer Specker, Michael Timmons, William Van Heukelem, Bamaby Watten, and Yonathan Zohar. Appreciation is extended to my students John Jacobs, Joe Schutz, and Jackie Tackas; and to my colleague, William Van Heukelem, for their patience and consideration during the process of developing and editing this work. This work could have never been completed without the love and support of Christ, my savior, my wife Ann, and my daughter Kaitlyn, who sacrificed considerable quality time that will never be able to be regained-- I Love You and Thank You! Cambridge, Maryland September, 1996

RegimlM. Han~ll

vi LIST OF CONTRIBUTORS G.R. Ammerman, Department of Food Technology, Mississippi State University, Mississippi State, Mississippi, 39762 D.L. Berlinsky, Department of Zoology,North Carolina State University, Raleigh, North Carolina 27695-7615 D.E. Brune, Department of Agricultural and Biological Engineering, Clemson University, Clemson, South Carolina 29634-0357 C.F. Femandes, Department of Food Science, Virginia Polytechnic I: :stitute and State University, Blacksburg, Virginia 24061 G.A. Flick, Department of Food Science, Virginia Polytechnic Institute and State University, Blacksburg, Virginia 24061 D.M. Gatlin, II, Department of Wildlife and Fisheries Science, Texas A & M University, College Station, Texas 77843-2258 C.M. Gempesaw, Department of Food and Resource Economics, University of Delaware, Newark, Delaware

19717 R.L. Hodson, Department of Zoology and North Carolina Sea Grant College, North Carolina State University, Raleigh, North Carolina 27695-7615 R.M. Harrell, University of Maryland Center for Environmental and Estuarine Studies, Horn Point Environmental Laboratory, and Maryland Cooperative Extension Service, Sea Grant Extension Program, Cambridge, Maryland, 21613 J.N. Hochheimer, Piketon Research and Education Center, The Ohio State University, Piketon, Ohio 45661 C.C. Kohler, Fisheries Research Laboratory, Southern Illinois University, Carbondale, Illinois, 62901 D.A. Lipton, Department of Agriculture and Resource Economics and Maryland Cooperative Extension Service, Sea Grant Extension Program, University of Maryland, College Park, Maryland 20742 J.A. Plumb, Southeastern Cooperative Fish Disease Laboratory, Department of Fisheries and Allied Aquacultures, Alabama Agricultural Experiment Station, Auburn University, Auburn, Alabama, 36849 D.D. Rawles, Department of Food Science, Virginia Polytechnic Institute and State University, Blacksburg, Virginia 24061 C.V. Sullivan, Department of Zoology, North Carolina State University, Raleigh, North Carolina 27695-7615 J.R. Tomasso, Department of Aquaculture, Fisheries, and Wildlife, Clemson University, Clemson, South Carolina, 29634-0362

o~ Vll

F.W. Wheaton, Biological Resource Engineering Department, University of Maryland, College Park, Maryland, 20742-5711 D.W. Webster, University of Maryland Cooperative Extension Service, Sea Grant Extension Program, Wye Research and Education Center, Wye Mills, Maryland, 21658 C.R. Weirich, Aquaculture Research Facility, Louisiana State University, Baton Rouge, Louisiana, 70820

This Page Intentionally Left Blank

TABLE OF CONTENTS CONTRIBUTORS

. .

.

.

.

.

.

.

.

.

.

.

PREFACE

.

.

C h a p t e r 1.

An O v e r v i e w of Morone Culture . . Reginal M. Harrell and Donald W. Webster

1.1

H i s t o r y o f striped bass a q u a c u l t u r e

1.2

Commercial production .

.

.

1.2.1

.

.

References C h a p t e r 2. 2.1 2.2

Industry survey .

.

.

.

.

.

. .

. .

.

. . .

.

.

1.2.1.1 T y p e s o f o p e r a t i o n s 1.2.1.2 P r o d u c t i o n t y p e s

. .

. .

. .

. .

1.2.1.3 1.2.1.4 1.2.1.5 1.2.1.6

. . . .

. . . .

. . . .

. . . .

9

Hatchery production Fingerling production F o o d - f i s h g r o w out R e s e a r c h priorities 9

9

.

.

9

~

.

Reproduction . . . . . . Craig V. Sullivan, David L. Berlinsky, and Ronald G. Hodson

11

Introduction . . . The reproductive system 2.2.1 Gonads . . 2.2.1.1 O v a r y . . . . . . 2.2.1.2 Testis . . . . . . 2.2.2 Neuroendocrine system . . . . . 2.2.2.1 B r a i n a n d h y p o t h a l a m u s . . 2.2.2.1.1 Gonadotropin-releasing hormone 2.2.2.1.2 Dopamine . . . . 2.2.2.2 P i t u i t a r y g l a n d . . . . . 2.2.2.2.1 Gonadotropins . . 2.2.2.2.2 G r o w t h h o r m o n e and p r o l a c t i n 2.2.2.2.3 O t h e r pituitary h o r m o n e s . 2.2.2.3 T h e e n d o c r i n e g o n a d . . . . 2.2.2.3.1 Estrogens . . . . 2.2.2.3.2 Androgens . . . .

11 11 11 12 17 19 19 21 21 22 23 24

9

9

o

,,

. .

. . .

.

.

27 28 28 30 30 31 31

.

.

31

2.2.2.3.3 P r o g e stins . . . . 2.2.2.4 O t h e r e n d o c r i n e o r g a n s . . . . 2.2.2.4.1 Stress h o r m o n e s . . . 2.2.2.4.2 Thyroid hormones . Insulin and insulin-like g r o w t h f a c t o r s 2.2.2.4.3 2.3

R e p r o d u c t i v e function . . . 2.3.1 The reproductive cycle . 2.3.1.10ogenesis

.

25 25 27 27

.

2.3.1.1.1 2.3.1.1.2

Oocyte growth . Final o o c y t e m a t u r a t i o n

2.3.1.1.3

Ovulation

.

31 33 34

2.3.1.2 S p e r m i o g e n e s i s . . . 2.3.1.3 Spawning behavior . . 2.3.1.4 Fertilization and egg activation . 2.3.2 Environmental influences . . 2.3.3 Puberty and maturity schedules . 2.3.3.1 Puberty 2.3.3.2 Assessing maturity . 2.3.3.3 Maturity schedules . 2.4 Captive breeding . . . 2.4.1 Broodstock acquisition and conditioning 9 2.4.1.1 Broodstock sources . . 2.4.1.1.1 Captive broodstock . 2.4.1.1.2 Domestic broodstock 2.4.1.2 Conditioning broodstock 2.4.2 Induction o f spawning . . . 2.5 Acknowledgments References . . Chapter 3.

Morone

P o n d P r o d u c t i o n . . . . M. H a r r e l l 3.1 Introduction . . . . . . 3.1.1 General c o m m e n t s . . . . 3.1.2 Striped bass versus hybrid culture . 3.1.3 Hatchery production . . . . 3.2 Phase I production . . . . . 3.2.1 Receipt o f fry . . . . . 3.2.2 Pond preparation . . . . 3.2.3 Stocking fry for phase I . . . . 3.2.4 Feed and feeding phase I fish . . 3.2.5 Harvesting phase I fish . . . . 3.3 Phase II production . . . . . 3.3.1 Fingerling availability . . . . 3.3.2 Pond preparation . . . . 3.3.3 Stocking phase I fish for phase II g r o w out 3.3.4 Feeds and feeding phase II fish . . 3.3.5 Harvesting phase II fish . . . . 3.4 Phase III production . . . . . 3.4.1 Stocking densities for phase III fish . 3.4.2 Feeds and feeding phase III fish . . 3.4.3 Harvesting phase III fish . . 3.5 Summary . . . . . . References

9

.

.

.

.

.

9

.

.

.

.

.

9

.

.

9

.

.

9

.

.

9

.

.

9

.

.

9

.

.

9

.

.

9

.

.

9

.

.

9

.

.

9

.

.

.

.

.

.

.

. .

36 38 39 41 45 45 45 46 49 49 49 49 51 51 53 59 60 75

.

R e g i n a l

9

Chapter 4.

W a t e r D a v i d

4.1 4.2 4.3

9

Q u a l i t y E.

.

D y n a m i c s

9

as

the

9

B a s i s

9

for

.

. .

.

.

.

.

.

.

. .

. . . .

.

. . .

.

.

.

.

. . .

. . . .

. .

.

.

.

.

.

.

. . .

. .

.

.

. . .

. . .

9

A q u a c u l t u r e

D e s i g n

9

75 75 75 76 76 77 77 78 80 80 82 82 82 82 83 87 91 91 91 93 95 96 99

B r u n e

Introduction . . . . . . . Water reuse systems . . . . . . Pond aquaculture systems . . . . . 4.3.1 Primary pond production . . . .

. . . .

99 99 100 101

xi

4.4

4.3.2 P o n d o x y g e n production . . 4.3.3 Algal standing crop and secchi disk D i s s o l v e d o x y g e n concentration . . . 4.4.1 Equilibrium gas concentration . . 4.4.2 G a s transfer rate . . .

.

.

.

. .

. .

. .

. .

.

.

.

. .

4.4.3 M a n u f a c t u r e r s o x y g e n a t i o n capacity . . . . 4.4.4 Surface reaeration . . . . . . 4.5 C a r b o n a t e equilibrium and p H . . . . . . 4.6 Prediction o f aquaculture p o n d p e r f o r m a n c e and water quality . 4.7 Prediction o f aquaculture reuse system p e r f o r m a n c e and w a t e r quality 4.8 Summary . . . . . . . . References 9

9

9

9

,

9

9

9

. 9

C h a p t e r 5.

Intensive Culture of Striped Bass . . . J o h n N. H o c h h e i m e r a n d F r e d r i c k W. Wheaton 5.1 Introduction . . . . . . 5.1.1 W h y intensive culture? . . . . 5.2 W a t e r supplies . . . . . . 5.2.1 Aeration . . . . . 5.3 Solids r e m o v a l . . . . . . 5.4 Intensive culture s y s t e m s . . . . 5.4.1 Pond systems . . . . . 5.4.1.1 O v e r v i e w o f pond s y s t e m s . . 5.4.1.2 T y p e s o f p o n d s and typical c o n s t r u c t i o n . 5.4.1.3 C o m p o n e n t s o f intensive p o n d systems . 5.4.1.3.1 W a t e r supply . 5.4.1.3.2 Drains . . 5.4.1.3.3 Aeration . . 5.4.1.3.4 Harvesting . 5.4.1.3.5 Feeding . . 5.4.2 Flow-through systems . . . 5.4.2.1 T y p e s o f f l o w - t h r o u g h s ys tems . 5.4.2.2 C o m p o n e n t s o f f l o w - t h r o u g h systems 5.4.2.2.1 W a t e r supply . 5.4.2.2.2 Drains . . 5.4.2.2.3 Tanks . . 5.4.2.2.4 Aeration . . 5.4.2.2.5 Harvesting . . 5.4.2.2.6 Feeding . . 5.4.3 Recirculating s y s t e m s . . . 5.4.3.1 C o m p o n e n t s o f recirculating systems 5.4.3.1.1 Biofiltration 5.4.3.2 M a n a g e m e n t o f b i o f i l t e r s 5.4.3.3 O t h e r c o m p o n e n t s . . References

C h a p t e r 6. 6.1

White

Bass

Christopher Introduction

Production

C. Kohler . .

and

.

Broodstock

.

Development

.

.

.

102 103 104 104 104 107 108 109 111 121 123 124 127

9

.

127

.

.

127

9

.

130

9

.

130

9

.

135

9

.

140

9

.

140

9

.

140

9

.

140

9

.

143

9

.

143

9

.

143

9

.

144

9

.

144

9

.

146

9

.

148

9

.

149

9

.

151

9

.

151

9

.

151

9

.

152

9

.

152

9

.

154

9

.

154

9

.

155

9

.

156

9

.

157

9

.

162

9

.

164

9

.

165

169

.

.

169

xii

6.2

B i o l o g y and e c o l o g y

.

.

.

9

.

.

169

.

.

9

.

.

169

Morphological characteristics

.

9

.

.

169

6.2.3

Age and growth

.

.

9

.

.

170

6.2.4

F o o d habits

.

.

.

9

.

.

170

6.2.5

Natural spawning

.

.

9

.

.

170

9

.

.

172

6.2.1

Geographical range

6.2.2

6.3

Collection and transportation of broodstock

6.4

B r o o d s t o c k a c c l i m a t i z a t i o n to c a p t i v i t y . 6.4.1 R e a r i n g facilities .

9

.

.

172

9

.

.

172

6.4.2

9

.

.

172

9

.

.

172

9

.

.

172

9

.

.

173

9

.

.

173

9

.

.

173

9

.

.

175

9

.

.

178

9

.

.

180

9

.

.

180

9

.

.

180

9

.

.

181

9

.

.

182

6.5

Water quality requirements

6.4.3 Disease control. 6.4.4 T r a i n i n g to f o r m u l a t e d f e e d 6.4.5 Separating sexes . Controlled spawning . . . 6.5.1 Use of hormones . 6.5.2 Manual spawning .

. . .

6.5.3 Incubation techniques . 6.5.4 Out-of-season spawning 6.6 Growout . . . . 6.7 Broodstock development . 6.8 Summary . . . . References 9

C h a p t e r 7. 7.1 7.2

7.3

T r a n s p o r t a t i o n

7.5 7.6

7.8

a n d

. .

. .

. .

.

.

9

Stress

. .

9

9

.

M i t i g a t i o n

.

Fish t r a n s f e r e q u i p m e n t . T r a n s p o r t units . .

.

.

.

.

.

.

Water quality considerations during 7.6.1 Dissolved oxygen . 7.6.2 Carbon dioxide . . 7.6.3 Temperature . . 7.6.4 Ammonia . . pH

.

.

.

.

.

. .

. .

Use of water additives during transport . . . 7.7.1 A d d i t i o n o f salts . . . . 7.7.2 Anesthetics . . . . . 7.7.3 Practical recommendations . . . P o s t - c o n d i t i o n i n g o f fish after t r a n s p o r t . . .

.

. .

. . . .

. . .

. .

.

.

.

.

. . .

. . .

. .

. .

.

. . .

.

201 201 202 203 203 203

. .

185 185 185 186 187 187 189 192 195 196 196 198 200

.

. .

.

. .

.

.

fish t r a n s p o r t . . . . . . . . . . . .

185

.

.

.

7.3.3 O x y g e n a t i o n and a e r a t i o n s y s t e m s . 7.3.4 Water quality monitoring equipment . P r e c o n d i t i o n i n g o f fish before t r a n s p o r t . . 7.4.1 P r e c o n d i t i o n i n g fish in p o n d s . . 7.4.2 P r e c o n d i t i o n i n g fish in h o l d i n g t a n k s . L o a d i n g rates . . . . . .

7.6.5 7.7

9

. .

C.R. W e i r i c h Introduction . . . . . . . . Stress and stress m i t i g a t i o n . . . . . . 7.2.1 E f f e c t s o f h a n d l i n g and t r a n s p o r t - i n d u c e d stress on fish 7.2.2 M i t i g a t i o n o f h a n d l i n g and t r a n s p o r t - i n d u c e d stress Transport equipment . . . . . . . 7.3.1 7.3.2

7.4

9

.

. .

204 204 207 208 208

xiii

7.9 Transport of larvae References . .

. .

.

.

.

.

.

.

Introduction . . . . Hybridization . . . . 8.2.1 Natural hybridization . . 8.2.2 Artificial hybridization . . 8.2.3 Backcrosses and F2s . . 8.3 Broodstock domestication . . 8.4 Strains . . . . . 8.4.1 Ancestry . . . 8.4.2 Strain evaluation . . 8.5 Qualitative and quantitative traits 8.6 Cytogenetics and genetic manipulations 8.6.1 Karyology . . . 8.6.2 Polyploidization . . 8.6.3 Gynogenesis . . . 8.6.4 Sex reversal . . . 8.6.5 Cryopreservation 8.6.6 Genetic engineering 8.7 Conservation genetics . References . . .

. .

8.1 8.2

9.1

9.2

9.3

9.4

9.5

9.6

.

.

. .

. .

. . . .

. . . .

.

. . . .

. .

. .

.

.

. . .

.

. . .

. .

.

.

. . .

217

.

.

. .

. .

9

9

9

9

9

9

9

9

9

9

9

9

9

Chapter 9.

.

.

. .

209 211

.

.

Hybridization and Genetics Reginal M. Harrell

Chapter 8.

.

.

9

.

217 217 219 219 220 220 222 222 223 223 223 224 224 226 227 227 228 229 230

Nutrition and feeding of striped bass and hybrid striped bass Delbert M. Gatlin, I I I

235

Introduction . . . . . . 9.1.1 Feeding habits . . . . . 9.1.2 Digestive system structure and f u n c t i o n . 9.1.3 Role o f nutrition in aquaculture . . Energy . . . . . . . 9.2.1 Metabolism . . . . . 9.2.2 Partitioning o f energy . . . . 9.2.3 Factors affecting energy requirements . Protein and amino acids . . . . . 9.3.1 Structure and classification . . 9.3.2 Functions . . . . . 9.3.3 Requirements . . . . . 9.3.4 Biological availability . . . . Carbohydrates . . . . . . 9.4.1 Structure and classification . . 9.4.2 Functions . . . . . 9.4.3 Requirements . . . . . Lipids . . . . . . . 9.5.1 Structure and classification . . 9.5.2 Functions . . . . . 9.5.3 Requirements . . . . . Minerals . . . . . .

235 235 235 236 236 236 236 237 237 237 237 238 239 239 239 239 240 240 240 240 240 241

. . .

.

.

.

.

. . . . .

.

. . . . .

.

. . . .

.

. . . .

xiv

9.6.1

9.7

9.8

Structure a n d c l a s s i f i c a t i o n

9.6.2 Macrominerals . 9.6.3 Microminerals . Vitamins . .

. . .

Structure a n d c l a s s i f i c a t i o n

9.7.2

Water-soluble vitamins .

9.7.3 Fat-soluble vitamins F e e d s and f e e d i n g p r a c t i c e s

9.8.3 9.8.4 References

.

.

242

.

242

.

.

.

.

242 242

. .

. .

. .

. .

243 243

.

.

.

9

.

. .

Diet f o r m u l a t i o n a n d m a n u f a c t u r e Feeding practices . . . .

. .

.

Satisfying nutritional requirements Feedstuffs . . . .

o

241 241

.

. .

.

. .

.

. .

.

9.7.1

9.8.1 9.8.2

. . .

. .

.

.

.

243

.

.

245 245

. .

9

9

9

246 248

Environmental Requirements and Noninfectious Diseases Joesph R. Tomasso

253

10.1

Introduction

10.2 10.3

Stress and d i s e a s e r e s i s t a n c e . Environmental requirements . 10.3.1 G e n e r a l c o m m e n t s . 10.3.2 T e m p e r a t u r e . . . 10.3.3 D i s s o l v e d o x y g e n . 10.3.4 Salinity . . . 10.3.5 H a r d n e s s a n d c a l c i u m . 10.3.6 p H and a l k a l i n i t y . 10.3.7 L i g h t . . . . Environmental related noninfectious 10.4.1 G e n e r a l c o m m e n t s . 10.4.2 A m m o n i a . . . 10.4.3 Nitrite . . . .

253 253 254 254 255 255 257 258 259 259 260 260 260 261 261 262 262 262 262 263 264

C h a p t e r 10.

10,4

.

.

.

.

.

.

. . .

.

. . .

. . .

. .

. .

. .

. . .

. . . .

. . .

. . .

. .

. . diseases . . . . . . . .

. . . .

10.4.4 S u s p e n d e d s e d i m e n t s . . . . . 10.4.5 G a s and o x y g e n s u p e r s a t u r a t i o n . . . . 10.5 Toxicants . . . . . . . 10.5.1 G e n e r a l c o m m e n t s . . . . . 10.5.2 A n e s t h e t i c s . . . . . . 10.5.3 A l g i c i d e s . . . . . . 10.5.4 F o r m a l i n . . . . . . 10,5.5 C h l o r i n e . . . . . . 10.6 Concluding remarks . . . . . . References 9

Chapter 11.

.

.

9

.

9

Infectious Diseases of Striped Bass . John A. Plumb

11.1

Introduction

11.2 11.3

Predisposing factors Virus diseases . 11.3.1 11.3.2 11.3.3

.

.

. .

.

. .

.

. .

.

.

. .

. .

9

.

.

271 .

.

. . .

. . .

264 265 266 271

. .

Lymphocystis virus . . . I n f e c t i o u s p a n c r e a t i c n e c r o s i s virus Striped b a s s a q u a r e o v i r u s . .

.

.

.

.

271 272 272 274 274

XV

11.4

11.5

11.6

Bacterial Diseases

.

septicemia

275

. .

.

.

275

Columnaris

.

.

.

.

.

278

11.4.3

Pasteurellosis

.

.

.

.

.

279

11.4.4

Edwardsiellosis

.

.

.

.

.

280

11.4.5

Vibriosis

.

.

.

.

.

282

11.4.6

Streptococcosis

.

.

.

.

.

284

11.4.7

Enterococcosis

.

.

.

.

.

284

11.4.8

Mycobacteriosis

11.4.9

Other bacteria

.

.

Fungal diseases .

.

.

11.5.1

Saprolegniosis

11.5.2

Branchiomycosis

.

. .

.

288

. . .

.

.

.

11.6.1

.

.

.

.

.

Icthyobodiasis

11.6.1.2

Amyloodiniasis

Ciliates .

.

290 290

.

.

.

.

.

Chilodonella Epistylis

.

.

.

.

11.6.2.3

Trichodina

.

.

.

.

11.6.2.4

Ichthyophthiriasis

Helminths

.

.

.

.

.

.

Digenetic trematodes

.

290 291 293 293 293

.

293 296

.

.

11.7.1.2

.

. .

Monogenetic

. .

.

11.7.1.1

.

292

11.6.2.2

parasitic diseases

. .

11.6.2.1

Metazoan

290

.

.

11.6.1.1

288

.

.

.

Flagellates

288

.

.

.

285

.

.

.

.

.

.

Protozoan diseases

11.7.2

11.9

.

11.4.2

11.7.1

11.8

.

Motile

11.6.2

11.7

.

Aeromonas

11.4.1

.

296

trematodes

.

.

296

.

.

297

11.7.1.3

Cestodes

.

.

.

.

300

11.7.1.4

Nematodes

.

.

.

.

300

.

.

300

.

301

11.7.1.5

Acanthocephalans

11.7.1.6

Leeches

Crustacean

parasites

.

. .

.

.

. .

Control therapy .

.

.

.

.

.

11.8.1

Viruses .

.

.

.

.

.

11.8.2

Bacteria

11.8.3

Fungi

11.8.4

Protozoa

11.8.5

.

.

.

.

.

Helminths

.

.

.

11.8.6

Leeches

.

.

.

11.8.7

Crustacea

.

.

.

Summary

. .

.

.

.

.

.

303 305 305 306

.

.

300

306 9

.

307 307 307

.

307

References

308

Economics and Marketing . . . . Douglas W. Lipton and Conrado M. Gempesaw, II

315

12.1

Introduction

315

12.2

Production

C h a p t e r 12.

12.2.1

. costs

.

.

.

.

Costs of production

.

. .

. .

systems

315

. .

.

.

316

12.2.1.1

Hatcheries

.

.

.

.

316

12.2.1.2

Static ponds .

.

.

.

316

xvi

12.2.1.3 . . . Tanks . . 12.2.1.4 . . . Net-pen and cage culture 12.2.2 Operating costs . . . . . . 12.2.2.1 Stocking costs . . . 12.2.2.2 Feed costs . . . . . 12.2.2.3 Other costs . . . . . 12.3 Demand . . . . . . . . 12.3.1 Seafood demand . . . . . . 12.3.2 The striped bass market . . . . . . 12.3.2.1 The wild fishery . . . . . 12.3.2.2 Hybrid striped bass . . . . . 12.4 Financial performance o f striped bass aquaculture . 12.5 The hybrid striped bass market . . . . . . 12.5.1 Hybrid striped bass m a r k e t i n g studies . . . 12.5.2 Marketing implications . . . . . . References . . . . . . . . . Chapter 13. 13.1 13.2

13.3

13.4

13.5

13.6

13.7

. . .

.

.

.

317 317 317 318 318 319 319 319 320 320 321 321 322 322 325 327

Processing and Food Safety . . . . . . 329 Dafne D. Rawles, Custy F. Fernandes, George J. Flick, and Gale IL Ammerman Production statistics . . . . . . Laws and regulations . . . . . . 13.2.1 Processing facilities . . . . . 13.2.2 H A C C P . . . . . . Processing . . . . . . . 13.3.1 Harvesting methods and transport . . 13.3.2 Receiving fish at the plant . . . 13.3.3 Initial handling . . . . . . 13.3.4 Filleting . . . . . . 13.3.5 Meat-bone separation . . . . . 13.4.6 Dressing yields . . . . . . Preservation methods . . . . . . 13.4.1 Ice-packing and refrigerating . . . 13.4.2 Chill-packing . . . . . . 13.4.3 Freezing . . . . . . 13.4.4 Smoking . . . . . . Packaging . . . . . . . 13.5.1 Bulk packaging . . . . . . 13.5.2 Labeling . . . . . . 13.5.3 Coding . . . . . . . Factors affecting quality and s h e l f life . . . 13.6.1 Microbiological considerations . . . 13.6.2 Microbial growth during storage . . 13.6.3 Chemical considerations . . . Composition . . . . . . . 13.7.1 Amino acid composition . . . 13.7.2 Elemental composition . . . . . 13.7.3 Fatty acid analysis . . . . . 13.7.4 Effect o f diet on composition . . . 13.7.5 Proximate analysis o f striped and hybrid bass

. . . . . .

.

.

. . . . . .

.

. . . . . . . .

. .

. . .

.

.

. .

.

. . .

. .

. .

.

329 329 330 330 330 330 331 332 332 332 332 334 334 335 335 335 336 337 338 338 338 339 340 340 341 341 341 341 341 342

xvii

13.8 13.9

Marketing . . . Cleaning and sanitizing . . 13.9.1 Cleaning and sanitizing 13.9.2 Cleaning and sanitizing 13.9.3 Cleaning and sanitizing 13.9.4 Pest management 13.10 Waste handling . . . 13.10.1 Liquid disposal . . 13.10.2 Solid disposal . . 13.10.3 Solids utilization References . . . . . Appendix

.

.

Subject index

.

.

.

. .

. .

.

. . . principles . plant equipment. plant employees. . . . . . . . . . . . . .

. .

343 344 346 348 349 349 350 351 351 352 353

. .

357 .

.

.

.

361

This Page Intentionally Left Blank

To Ann and Kaitlyn

This Page Intentionally Left Blank

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

Chapter 1

An Overview of Morone Culture Reginal M. Harrell and Donald W. Webster 1.1 HISTORY OF STRIPED BASS AQUACULTURE The genus Morone is comprised of four species found in North America and is a member of the family Percichthyidae, the temperate basses (Setzler et al., 1980). Johnson (1984) placed the Morone in their own family, Moronidae, but they still are recognized by Robins et al. (1991) as belonging to Percichthyidae. Of the four species, the striped bass, M. saxatilis, and white perch, M. americana, are principally Atlantic coast drainage species. Striped bass also are found in the coastal tributaries of the Gulf of Mexico from western Florida to Louisiana (Meriman, 1941; Raney, 1952; Brown, 1965). The other two species, white bass, M. chrysops, and yellow bass, M. mississippiensis, are principally Mississippi River drainage species (Lee et al., 1980). There are also two similar European species, Dicentrarchus labrax and D. punctatus, which have had their nomenclature fluctuate between Morone and Dicentrarchus (Setzler et al., 1980). Historical and current vernacular retains the genus Dicentrarchus. Habitat, spawning, and culture requirements of Dicentrarchus and Morone are very similar, and there are a few excellent publications covering this information in detail. Most notable among the references are Pickett and Pawson (1994) covering the biology and conservation, and Bamab6 (1980, 1990) for biology, life history, and culture requirement information. Due to man's movement of these species, today Morone can be found in almost all 48 contiguous United States. Striped bass have also been exported to the former USSR (Doroshev, 1970), France and Portugal (Setzler et al., 1980), and most recently Israel, Taiwan, and Germany (Striped Bass Growers Association, Raleigh, NC, personal communication). Several synopses of the biological data of striped bass are available (Westin and Rogers, 1978; Setzler et al., 1980; Hill et al., 1989). Artificial culture of Morone was initiated in the early 1880s when S.G. Worth constructed a hatchery on the banks of the Roanoke River in Weldon, NC (Worth, 1884). Eggs were collected from gravid females as they were spawning in the river below the hatchery, fertilized with captured males, and incubated in standard MacDonald incubating jars. During the first year of operation over two million eggs were collected and almost 300,000 fry were hatched and stocked into the river (Worth, 1884). It was not until the mid 1960s in South Carolina that a means to hormone-induce spawning was developed, which allowed culturists to artificially ovulate gravid females taken from areas removed from their natural spawning grounds (Stevens et al., 1965; Stevens 1966, 1967). A good review of the history and overview of striped bass culture and management can be found in Stevens (1984) and Whitehurst and Stevens (1990). See Chapter 8 of this volume for an overview of hybridization. The seminal reproduction efforts in South Carolina were subsequently refined and the information developed into a variety of publications including Bayless (1972), Bonn et al. (1976), Kerby (1986), Harrell et al. (1990a), and Chapter 2 of this volume. Essentially all the knowledge we have today regarding culture requirements of Morone, including that necessary for food-fish production, came from the expansion of a concerted effort throughout the southeastern United States to provide striped bass and interspecific hybrid Morone for stocking public reservoirs for recreational fishing and fisheries management (Whitehurst and Stevens, 1990; Harrell et al., 1990b). Readers interested in the more practical aspects of culture should refer to these references. This book, although it will cover considerable aspects of the culture of the fish, is more

focused on the science of Morone biology and culture rather that the "how to" approaches that can be found in those referenced publications. 1.2 COMMERCIAL PRODUCTION Due to the success associated with Morone population enhancement efforts in inland systems and an associated decline in wild commercial harvests from coastal populations, a food-fish industry began to develop in the mid 1980s. As a result of its success and market acceptance, today striped bass and hybrid commercial aquaculture is considered the fastest growing segment of the U.S. aquaculture industry (USDA, 1992). The industry also is heavily supported by its own producers association, the Striped Bass Growers Association. Millions of Morone fingerlings are produced annually in state and federal hatcheries for population enhancement and in private hatcheries as seed stock for food-fish production and fee fishing operations (Table 1.1, Figure 1.1 ). These fingerlings are stocked in earthen ponds, flow-through and closed recirculating tanks, and net-pens throughout the U.S., and currently yield millions of pounds of food-fish annually (Table 1.2, Figure 1.2). Upon examining the information provided in Table 1.2, one quickly realizes that there has been over a 1,400% increase in production in 10 years, and the industry is still growing annually in the number of producers and production. Today, the food-fish industry is based primarily on the production and rearing of hybrid Morone. Of the different types of Morone that have been cultured, only the palmetto bass (striped bass ~ x white bass cr) and sunshine bass (white bass r x striped bass o') are reared for production (see below). Other types of Morone hybrids that have been created for the potential of food-fish production or have been used in stocking programs for recreational fishing are Maryland bass (white perch ~ x striped bass cr), Virginia bass (striped bass ~ x white perch cr), and paradise bass (striped bass ~ x yellow bass ~) (Harrell et al., 1990; see Chapter 8).

Table 1.I. Swiped bass and hybrid striped bass phase I fingerling production records for state and private hatcheries. State records represent cooperating states reporting to the Striped Bass Technical Committee of the Southern Division of the American Fisheries Society and includes state, federal, and university rearing facilities. Private hatchery data combines palmetto bass and sunshine bass into a general category of hybrid striped bass. Private hatchery data was obtained from Rhodes and Sheehan (1991) and Kahl (1995). NA: data not available.

YEAR Striped Bass

Public Hatcheries Palmetto Bass

Sunshine Bass

Private Hatcheries

TOTAL

1991

10,733,923

4,454,004

4,265,718

18,920,000

38,373,645

1992

6,915,512

3,505,805

3,135,437

19,560,000

33,117,101

1993

15,468,901

4,931,133

4,347,626

NA

24,474660

1994

23,314,272

7,198,387

2,347,235

17,900,000

50,759894

1995

9,132,688

7,091,580

6,967,598

22,100,000

39,021,866

Fig. 1.1. Production estimates for phase I fingerling production. Data obtained for striped bass, palmetto bass, and sunshine bass is from the Striped Bass Technical Committee, Southern Division of the American Fisheries Society, Bethesda, MD, and represents hatchery production from state, federal, and university facilities. Information from private hatcheries combines both palmetto and sunshine bass into a "hybrid bass" category and was obtained by a mail survey to producers of the Striped Bass Growers Association and state Aquaculture Coordinators (Rhodes and Sheehan, 1991; Kahl, 1995). The values for the private hatcheries represent midpoint values between the high and low reported values reported on the surveys (see Table 1.1 for specifics). Private production data was not available for 1993.

1.2.1 Industry Survey In 1995 a Northeast Regional Aquaculture Center funded survey o f producers was conducted by the Striped Bass Growers Association and the University of Maryland in an effort to collect information on the status of the industry, identify concems, and establish industry defined research needs (Harrell and Webster, unpublished data). The information that follows in this section summarizes that information. A total of 75 surveys were sent out to all the striped bass and/or hybrid producers listed in AquacultureMagazine's 1994 Annual Buyer's guide. From those mailings, 27 or 36% were returned.

Table 1.2. Estimated production of food-fish size striped bass hybrids. Kent Sea Farms information represents information gathered from telephone surveys of producers from the Striped Bass Growers Association (SBGA) (J. Carlberg, Kent Sea Farms Corporation, San Diego, CA, personal communication). Values from Rhodes and Sheehan (1991) and Kahl (1995) represents information obtained from mail surveys to SBGA producers and state Aquaculture Coordinators. All values are presented as 1000s of pounds. Missing data indicates information was not available.

Year

Kent Sea Farms

Rhodes and Sheeh,_an (1991) Low High

Kahl (1995) Low High

1986

10

1987

405

1988

880

1989

1,020

1990

1,590

1,150

1,560

1991

2,250

3,270

3,810

1992

3,550

6,910

8,390

1993

5,950

1994

7,625

7,800

9,100

1995

8,450

10,800

12,100

13,700

15,400

1996

1.2.1.1 Types of operations Regarding the type of system being used to culture fish, 65.6% of the producers used earthen ponds, 15.6% used tanks, 9.4% used net-pens, and 9.4% used raceways. Of those producers utilizing ponds, 86.4% used freshwater while 4.6% had saltwater. The remaining 9% had access to either salt or freshwater. Those culturing fish in tanks used primarily flow-through operations (66.7%), 22.3% were closed recirculating systems, and 11.1% had the capacity for both. Two thirds of the net-pen operators worked in private and freshwater systems, while one-third was located in saltwater or public systems. 1.2.1.2 Production _types Morone culture can be broken down into three components: hatchery, fingerling, or food-fish production. Producers can be practitioners of these components individually or in any combination. Of the responses obtained in the survey, 55.6% of the producers were involved in grow-out to market-sized fish, 25.9% were in the hatchery business, and 18.5% were fingerling producers. Of those in hatchery production, 70% claimed to also invest into fingerling production, or grow-out, or both. Likewise, 55.6% of the fingerling producers and 25% of the grow-out producers had diversified into another component of culture.

16 14 12 er O

10

Q

8

9

6

~

j

4 2 0

__.._._.-.R--8g 8~/ 8~:

819

90

9'1 Year

9~2

9~3

9~

93

9~5

Fig. 1.2. Production of food fish size hybrid striped bass. Values for 1986-1989 and 1993 were provided by J. Carlberg, Kent Sea Farms Corporation, San Diego, CA (personal communication) and were based on telephone surveys conducted with producers from the Striped Bass Growers Association (SBGA). Values for 1990-1992 were obtained from Rhodes and Sheehan (1991), and 1994-1996 were taken from Kahl (1995) and represent midpoint values from high and low estimates (see Table 1.2).

1.2.1.3 Hatchery oroduction Just under 50% (47.4%) of the individuals involved in hatchery production used wild fish as a source of broodstock. Surprisingly, 36.8% of the producers claimed to use captive broodstock and 15.8% have domesticated fish. It was not clear what generation level of captive fish are being used as fish are not considered as "domesticated" until at least the second generation has been reared in a captive environment (see Chapters 2 and 8 for more discussion on broodstock). Regarding the types of fish spawned, 26.3% of the producers reported to make pure striped bass, 21.1% pure white bass, and 52.6% hybrids of the two. One hundred percent of the white bass producers manually stripped and artificially fertilized the eggs, while 60% of the pure striped bass were produced manually. Forty percent of those spawning pure striped bass used tank spawning as their method of choice. All hybrids were produced by manual spawning. Ten percent of the hatcheries reported having the capability to spawn year round (see Chapter 2), while 20% reported using some form of temperature control (i.e., cooling) as a means to prolong the spawning season.

It was interesting that 60% of those hatcheries producing hybrids produced both sunshine bass and palmetto bass, while only 20% were exclusively palmetto bass or sunshine bass producers. There were no reports of Maryland, Virginia, or paradise bass production. Of the larvae produced, 80% were for domestic purposes while the remaining 20% were exported. As much as 10% of the annual larval production was exported in 1994. 1.2.1.4 Fingerling vroduction - -

v

.

Almost all fingerling production occurs in earthen ponds (see Chapter 3) although the trend is shifting to an increasing effort of rearing larvae to fingerlings in tanks (see Chapter 5). Only 11% of fingerling producers cultured phase I (30-100mm)exclusively. The majority (89%) produced both phase I and phase II (6-12 month old fish) fingerlings. No individual reported exclusive production of phase II fish. Average length of the phase I production season was three months, while the average phase II production was eight months. Target harvest size for phase I fish was 1,760 fish/kg (800/lb), with a range of 220-4,000/kg. Target harvest weight for phase II fish was 18 g or 55/kg (range of 8.8 to 220 fish/kg). Seventy-eight percent of the producers initiate feeding phase I fish in the pond before harvest and start as early as 14 days post-stocking. As many as two thirds of the producers reported they export fingerlings, and as much as 60% of their production is exported. One of the largest problems mentioned by producers was with the rate of swimbladder inflation in larvae and its impact of harvest success. Twenty-two percent of the fingerling producers mentioned having this type of problem with as much as 30% of the harvested fish exhibiting deformities such as scoliosis and lordosis. Another problem mentioned was with aquatic vegetation. Fifty-six percent of the phase I producers and 78% of the phase II producers stated that aquatic weeds were a significant problem. Diseases were reported as being problematic (see box), but few of the producers actually took their fish to a disease diagnostic service. Most producers performed cursory examinations themselves. For more specific information on infectious diseases and parasites refer to Chapter 11. For information on noninfectious diseases found in Morone refer to Chapter 10.

FINGERLING DISEASE INCIDENCE (% Producers Reporting) Parasites

Bacterial or Funeal

Ichthyobodo sp.

33%

Flexibacter columnaris 56%

Trichodina sp.

33%

Saprolegnia sp.

Grubs, (White, Yellow, and Black) 22%

Aeromonas sp.

1.2.1.5 Food-fish grow out Production facilities varied in size for ponds from 0.5 to 6 acres, with the average size grow-out pond being two acres. There was no reported average size for production tanks, and the net-pens averaged 28.3 cubic meters. Average target size fish for market was 680g, with 50% of the producers preferring a fish 800 g or larger. Market season was reported as ranging from 3 to 12 months, with 50% of the producers having less than a 12 month marketing season. There were a variety of marketing outlets with quite a range in the effort placed by the farmer in a specific area (see box). Most sold fish to distributors, shipping them on ice (60% of the producers moved as much as 100% of their inventory).

11%

11%

Ichthyophthiriasis 11% Epistylis sp.

11%

Ambiphrya sp.

11%

MARKET OUTLET (% of Producers) (Range of Production Sales) On Ice

60% (10-100%)

Live Haul

40% (1-I00%)

Wholesale

45% (75-100%)

Retail

35%

(3-30%)

Direct to Restaurant

30%

(1-15%)

Pond Bank

20% (5-100%)

Gel Pack

20% (5-100%)

Export

10% (5-30%)

Filleted by Producer

5%

(5%)

Most farmers (85%) quit feeding their crops before harvest, with as much as 50% discontinuing feeding at least two days before harvest. Only 25% of the farmers reported having off-flavor problems with their fish. All of them felt that algae was the biggest contributor to off-flavor, with other causes listed as feed, bacteria, and an "unknown" category. Sixty percent of the growers stated they had moderate to serious problems with aquatic weeds and the lack of availability of the chemicals needed to control them. Associated with the aquatic weed problem is the nutrients added to the water through uneaten and partially digested feeds. The average percent protein used in the grow-out diets reported was 38% with a range of 32-40%. Fat composition of the diets ranged from 4 to 12%, with an average of 8%. Maximum feeding rate per acre per day was 80 pounds, with a range of 40 to 200 pounds. Similar to the fingerling production phase of Morone culture, there were a variety of disease problems in the grow-out segment. However, unlike the fingerling stage, most farmers submitted sick or diseased fish to a diagnostic lab for analysis as evidenced by the increased number and type of organisms causing problems (see box). This practice was interesting because it indicated producers felt that smaller fish were not as much of an economic risk as larger fish nearing market size. It is clear that in both cases more attention must be given to health management of the production crop irrespective of whether it is a 60 mm phase I fish or a 600 g market fish. The loss is the same -- there will be no economic return from dead fish. 1.2.1.6 Research priorities

GROW-OUT DISEASE INCIDENCE (% of Producers Reporting) Parasites

Bacterial or Funga!

Aeromonas sp. 30%

lchthyobodo sp.

Flexibacter columnaris 15%

Grubs (White, Yellow, and Black) 25%

Pseudomonas sp. 5%

Trichodina sp.

15%

Mycobacteria 5%

Epistylis sp.

15%

Gill Disease 5%

Chilodonella sp.

10%

Saprolegnia sp.

5%

Trichophrya sp. Amb iphrya sp.

25%

10% 10%

Included in the survey was an opportunity for External Parasites 10% the producers to prioritize what they conceive as the greatest needs for research to improve their production Ichthyophthiriasis 5% ...... returns. First on the list was development of complete Gyrodactylus sp. 5% diets, and second was a reduction in the cost of fingerlings to producers. After the first two items there i~ was a consensus that several items were of high priority and needed attention but were not as immediate as the first two issues. Among these other research priorities were genetic selection, drug and therapeutant availability, year-round spawning capabilities, broodstock development, improved transportation capabilities, and marketing. All of these areas of concern are addressed in this volume and the reader is encouraged to review the appropriate chapter(s) associated with their specific interests. Below is a breakdown of the breeding and selection priorities as listed by the producers as they saw the descending rankings of need. In other words, if there is a genetic basis for these traits and that trait can be exploited, these would be the areas in which research should be directed.

9

9 9 9 9 9 9

9

9 9 9 9

Stress Tolerance 1) Low dissolved oxygen 2) Crowding 3) Handling 4) High ammonia 5) High CO2 6) High nitrite 7) Fluctuating pH 8) High temperature 9) Low temperature 10) Low hardness 11) Low alkalinity 12) No salinity Feed Conversion Efficiency Fast Growth Disease Resistance Uniform Growth Protein Utilization Efficiency Reproduction 1) Early reproduction 2) Delayed reproduction 3) Sterility Body Configuration 1) Yield and dress-out percentage 2) Coloration 3) Striping patterns Ease of Harvest Adaptability to Tank Spawning Age at Weaning Larval-egg size

Obviously, some of these waits may not have an exploitable genetic basis because many are influenced by environmental conditions such as diet, temperature, and photoperiod. As can be seen in Chapter 8 little is known about the additive genetic variation of Morone, thus basic information is needed about the genetics of the group before breeding programs can be developed. It does, however, provide an excellent guideline for researchers to focus future industry prioritized needs for research. What is apparent in these selections is the industry is most interested in those traits that will have a positive affect on their margin of return. Chapter 12 discusses the economics and marketing aspects of Morone culture. When reviewing that chapter, the reader should realize the importance of these specific traits toward improving profitability.

References

Barnabr, G., 1980. Expos6 synoptique des donnres biolgiques sur le loup ou bar Dicentrarchus labrax (Linnaeus 1758). FAO. Fisheries Synopses 126, 70 p. Barnabr, G., 1990. Rearing bass and gilthead bream. Pages 646-686 in G. Bamabr, editor. Aquaculture, volume 2. Ellis Horwood Series in Aquaculture and Fisheries Support, New York, London. Bayless, J.D., 1972. Artificial propagation and hybridization of striped bass, Morone saxatilis (Walbaum). South Carolina Wildlife and Marine Resources Department, Columbia. Bishop, R.D., 1968. Evaluation of the striped bass (Roccus saxatilis) and white bass (R. chrysops) hybrids after two years. Proceedings of the Annual Conference Southeastern Association of Game and Fish Commissioners, 21: 245-254. Bonn, E.W., Bailey, W.M., Bayless, J.D., Erickson, K.E. and Stevens, R.E., editors, 1976. Guidelines for striped bass culture. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Brown, B.E., 1965. Meristic counts of striped bass from Alabama. Transactions of the American Fisheries Society, 94: 278-279. Doroshev, S.I., 1970. Biological features of the eggs, larvae and young of the striped bass [Roccus saxatilis (Walbaum)] in connection with the problem of its acclimatization in the USSR. Journal of Ichthyology, 10: 235-248. Harrell, R.M., Kerby, J.H. and Minton, R.V., editors., 1990a. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Harrell, R.M., Kerby, J.H., Smith, T.I.J. and Stevens, R.E., 1990b. Striped bass and striped bass hybrid culture: the next twenty-five years. Pages 253-261 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Hill, J., Evans, J.W. and Van Den Avyle, M.J., 1989. Species profiles: life histories and environmental requirements of coastal fisheries and invertebrates (South Atlantic)-- striped bass. U.S. Fish and Wildlife Service Biological Report 82(11.118). U.S. Army Corps of Engineers TR EL-82, Vicksburg, MS. Johnson, G.D., 1984. Percoidei: Development and relationships. Pages 464-498 in H.G. Moser, W.J. Richards, D.M. Cohen, M.P. Fahay, A.W. Kendall, Jr., and S.L. Richardson, editors. Ontogeny and systematics of fishes. American Society of Ichthyologists and Herpetologists, Special Publication 1:464-498. Kahl, K.H., 1995. Results of the hybrid striped bass producer and state aquaculture coordinator survey. South Carolina Department of Agriculture Report. Columbia. Kerby, J.H., 1986. Striped bass and striped bass hybrids. Pages 127-147 in R.K. Stickney, editor. Culture of nonsalmonid freshwater fishes. CRC Press, Boca Raton, FL. Lee, D.S., Gilbert, C.R., Hocutt, C.H., Jenkins, R.E., McAllister, D.E. and Stauffer, Jr., J.R., 1980. Atlas of North American freshwater fishes. Publication #1980-12, North Carolina Biological Survey, North Carolina State Museum of Natural History, Raleigh. Meriman, D., 1941. Studies on the striped bass (Roccus saxatilis) of the Atlantic coast. U.S. Fish and Wildlife Service, Fisheries Bulletin, 50: 1-77.

10

Pickett, G.D. and Pawson, M.G., 1994. Sea bass. Chapman and Hall, Fish and fisheries series number 12, London. Raney, E.C., 1952. The life history of the striped bass, Roccus saxatilis (Walbaum). Bulletin of the Bingham Oceanographic Collection, Yale University, 14: 5-97. Rhodes, R.J. and Sheehan, B. 1991. Estimated annual production of commercial hybrid striped bass growers in the United States, 1990-1992. Striped Bass Growers Association Report, South Carolina Department of Agriculture, Columbia. Robins, C.R., Bailey, R.M., Bond, C.E., Brooker, J.R., Lachner, E.A., Lea, R.N. and Scott, W.B., 1991. Common and scientific names of fishes from the United States and Canada. Fifth edition. American Fisheries Society Special Publication 20. Bethesda, MD. Setzler, E.M., Boynton, W.R., Wood, K.V., Zion, H.H., Lubbers, L., Mountford, N.K., Frere, P., Tucker, L. and Mihursky, J.A., 1980. Synopsis of biological data on striped bass, Morone saxatilis (Walbaum). U.S. Department of Commerce, NOAA Technical Report, NMFS Circular 433, FAO Synopsis No. 121. Washington, DC. Stevens, R.E., 1966. Hormone-induced spawning of striped bass for reservoir stocking. The Progressive Fish-Culturist, 28:19-28. Stevens, R.E., 1967. A final report on the use of hormones to ovulate striped bass, Roccus saxatilis (Walbaum). Proceedings of the Annual Conference Southeastern Association of Game and Fish Commissioners, 18: 525-538. Stevens, R.E., 1984. Historical overview of striped bass culture and management. Pages 1-15 in J.P. McCraren, editor. The aquaculture of striped bass: a proceedings. Maryland Sea Grant Publication UM-SG-MAP-84-01. University of Maryland, College Park. Stevens, R.E., May, Jr., O.D. and Logan, H.J., 1965. An interim report on the use of hormones to ovulate striped bass (Roccus saxatilis). Proceedings of the Annual Conference Southeastern Association of Game and Fish Commissioners, 17: 226-237. USDA. 1992. Aquaculture situation and outlook report. United States Department of Agriculture Commodity Economic Division Economic Research Service, March 1992, AQUA-8, Washington, DC. Westin, D.T., and Rogers, B.A., 1978. Synopsis of biological data on the striped bass, Morone saxatilis (Walbaum). University of Rhode Island Marine Technical Report 67, University of Rhode Island, Marine Advisory Service, Narragansett. Worth, S.G., 1884. Report upon the propagation of striped bass at Weldon, N.C., in the spring of 1884. Bulletin of the United States Fish Commission, 4(15): 225-230.

Striped Bass and Other Morone Culture R.M. HarreU (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

11

Chapter 2 Reproduction Craig V. Sullivan, David L. Berlinsky, and Ronald G. Hodson 2.1 INTRODUCTION This chapter covers developments over the last 10 years in our understanding of the reproductive biology of striped bass (Morone saxatilis). Striped bass reproduction has been studied and manipulated for over 100 years, beginning in the 1870s at the U.S. Fish Commission hatchery on the Roanoke River in North Carolina (Whitehurst and Stevens, 1990; see also Chapter 1). Methods for propagating striped bass were recently reviewed in detail (Harrell et al., 1990a). Over the past few years, interest in the biology and control of striped bass reproduction has intensified as a consequence of declines in fishery landings and growth of aquaculture for restocking fisheries and producing marketable fish. Fishery biologists need detailed knowledge of striped bass reproduction and maturity schedules to effectively manage exploited stocks (Specker et al., 1987; Berlinsky et al., 1995b). Environmental scientists require the same kind of information to assess anthropogenic impacts on natural populations. Aquaculturists increasingly need to understand and control the reproductive cycle of striped bass and related species. Development of domesticated broodstocks and their controlled maturation and spawning will be necessary to support continued growth of hybrid striped bass farming (Harrell et al., 1990b), one of the fastest growing forms of foodfish culture in the United States. This chapter should serve as a guide to what is currently known about reproduction of striped bass and as a catalyst for future studies in this area. It focuses on results of original research relevant to the needs of natural resource managers, environmental scientists, and aquaculturists. Information from studies of striped bass is emphasized, supplemented by results of recent research on the congeneric white bass (M. chrysops)and white perch (M. americana). We are just beginning to develop a rudimentary understanding of the reproductive biology of these fascinating and important species. 2.2 THE REPRODUCTIVE SYSTEM The reproductive system of striped bass is fairly typical of teleost fish. Redding and Patino (1993) recently published a succinct review of what is known about teleost reproduction. More detailed reviews of the functional morphology of fish gonads are also available (Grier, 1981; Wallace and Selman, 1981, 1990; Nagahama, 1983; Guraya, 1986; Callard, 1991). In spite of some reports of hermaphroditism in this species (Schultz, 1931; Dorfman, 1976; Moser et al., 1983), striped bass are normally sexually dioecious. They are iteroparous fish, capable of spawning annually for many years once maturity is reached. 2.2.1 Gonads Groman (1982) published a general review of the anatomy and histology of striped bass including the gonads and associated structures. The gonads are paired, elongated organs found along the dorsal wall of the body cavity. They are suspended from the ventral surface of the gas bladder by short peritoneal mesenteries, termed mesovaria in females and mesorchia in males. Blood vessels, lymph ducts, and nerves enter the two lobes of the gonad in the rostral area of these attachments, branching posteriorly as they course through the

12

dorsal portion of the peripheral gonad wall (tunica albuginea). The wall of the gonad extends caudally and eventually fuses into a short common genital duct, the oviduct or sperm duct, exiting the body at a genital pore located between the anus and urinary pore. 2.2.1.10var~ The ovarian wall consists of fibrous connective tissue containing bundles of circular and longitudinal muscle, collagen, blood vessels, lymph ducts, and nerves. The ovary contains a lumen (ovocoele) into which mature oocytes are released at ovulation. The luminal border consists of a squamous epithelium covering the ovarian lamellae and ovarian wall. The lamellae (ovigerous folds) are made up of germinal epithelium, follicular epithelium, vascular elements, and connective tissue. They branch extensively as they project from the ovarian wall into the ovocoele. Oogenesis occurs in ovarian follicles located within the lamellae. In histological sections, the lamellae appear full of oocytes at various stages of development (Figure 2.1A). The ovarian follicle (Figure 2.2) is the fundamental structural and functional unit of the ovary. It is morphologically similar in all teleosts (Patino and Takashima, 1995). The oocyte is located in the center of the follicle and is surrounded, in turn, by several acellular and cellular layers. First is the acellular zona radiata (chorion), which will develop into the structure commonly referred to as the "egg shell" after spawning. Next are the steroidogenic granulosa cells, usually a monolayer of cuboidal cells. One or more layers of squamous theca cells surround the granulosa cell layer. In many teleosts, this thecal layer is known to contain specialized cells involved in production of sex steroids, as well as fibroblasts and capillaries. A basement membrane separates the granulosa and theca cell layers. Microvilli emanating from both the oocyte and the granulosa cells penetrate the zona radiata through pore canals (Figure 2.3) and make cytoplasmic connections with one another via gap junctions. These connections likely provide for chemical exchange and direct communication between the oocyte and granulosa cells. Such connections have also been reported to exist between adjacent theca or granulosa cells in some teleosts. Primary germ cells in the epithelium of the ovarian lamellae undergo mitosis and give rise to undifferentiated oogonia. The oogonia occur in nests and contain a lightly acidophilic staining cytoplasm. Their most striking feature is a large central nucleus with a single prominent nucleolus (Figure 2.1D). The oogonia subsequently initiate meiosis to form oocytes. During follicle development and most of oocyte growth, the oocytes are arrested in meiotic prophase I. Just before ovulation, the fully yolked oocytes resume meiosis which includes migration of the nucleus or germinal vesicle to the oocyte periphery, dissolution of the nuclear membrane (germinal vesicle breakdown), chromosome condensation, assembly of the first meiotic spindle, and extrusion of the first polar body (Nagahama et al., 1994). Meiosis is arrested again at metaphase II until the oocytes are ovulated, spawned, and inseminated. The period from resumption of meiosis at prophase I to its arrest at metaphase II is defined as final oocyte maturation. Several investigators have published sets of micrographs depicting the various stages of oocyte development in striped bass (Groman, 1982; Specker et al., 1987; Berlinsky and Specker, 1991; Tao et al., 1993). Stages of oocyte maturation are identical in all temperate basses, but terminology for naming the stages varies somewhat among authors. A simplified nomenclature is used in this chapter. Oocytes are classified in developmental sequence as primary growth, early secondary growth, vitellogenic, final maturation, or atretic (Figures 2.1 and 2.4). Primary growth oocytes have a maximum diameter of---150 #m with a densely and uniformly basophilic staining cytoplasm (Figure 2.1C). A single large and basophilic nucleolus is present initially (chromatin nucleolar phase). Multiple nucleoli later appear along the periphery of the nucleus and the cytoplasm stains less intensely (perinucleolar phase). Early secondary growth oocytes (minimum diameter 120-150/.zm) contain unstained lipid droplets scattered throughout the ooplasm (early yolk vesicle phase).

13

Fig. 2.1. Female reproductive system of striped bass (from Groman, 1982). A. Transverse section of a juvenile ovary (formalin; H&E). B a r - 71 ~zm. B. Wall of the ovary proximal to the oviduct, juvenile fish (formalin; H&E). Bar = 43 gm. C. Primary growth oocytes within the ovarian lamellae, juvenile fish (formol sublimate; H&E). Bar = 43 ~zm. D. Nests of oogonia in a juvenile ovary (Bouin's; H&E). Bar = 43 ~m. E. Maturing oocytes, early vitellogenic oocyte (center) and late vitellogenic oocyte (bottom left), 3-year-old fish (formalin; H&E). Bar = 24/~m. 12. Perinucleoli 1. Cytoplasm of vitellogenic oocyte 8. Chromatin nucleolar phase 13. Red blood cell 2. Nucleoli primary growth oocyte 14. Smooth muscle 3. Fibrous connective tissue stroma 9. Perinucleolar phase primary 15. Tunica albuginea 4. Follicle cells growth oocyte 16. Yolk granules 5. Lumen of ovary 10. Ovarian vein 17. Lipid droplets 6. Nucleus of oocyte 11. Ovarian lamellae 7. Oogonial nests

14

Fig. 2.2. Diagram of a vitellogenic ovarian follicle. The oocyte nucleus or germinal vesicle (GV), ooplasm, zona radiata (ZR), granulosa cell layer (G), basement membrane (BM) and theca cell layer (T) are shown. (Diagram courtesy of S. A. Heppell, Department of Zoology, North Carolina State University).

Fig. 2.3. Electron micrograph of a portion of an ovarian follicle from a white perch sampled in late vitellogenesis (March). The ooplasm (O), zona radiata (ZR), pore canals (arrows), a granulosa cell (G), and the basement membrane (BM) are shown. The nucleus of a theca cell is apparent in the extreme upper right comer of the micrograph. Magnification~ 40,000X. The ovary sample was fixed in 2.5% glutaraldehyde, post-fixed in 1% osmium tetroxide, embedded in Spurr's resin and sectioned at 50-70 nm (Photograph courtesy ofY. Tao, Department of Zoology, North Carolina State University).

15

Fig. 2 4. Oocytes undergoing ~mal oocyte maturation or atresia. A. Oocyte in early f'mal maturation showing evidence of a migrating germinal vesicle (GV). B. Oocyte in late final maturation showing coalescence of lipid droplets (L) and advanced migratiol: of the germinal vesicle (GV). C. Oocyte undergoing atresia showing resorption vacuoles (arrows). (Photograph courtesy of L. F. Jackson, Department of Zoology, North Carolina State University). Later, the lipid droplets become more numerous, and distinct granules (cortical alveoli) that stain positively with periodic acid - Schiffs reagent (PAS) appear in the peripheral cytoplasm (cortical granule phase). The contents of the cortical granules are discharged into the perivitelline space at the time of fertilization in the cortical reaction (Wallace and Selman, 1990). As the cortical granules do not contribute to embryonic growth, they are not really a component of egg yolk. Vitellogenic oocytes, generally >_ 250 ~tm in diameter, exhibit acidophilic yolk globules in their peripheral cytoplasm in addition to the inclusions seen in early secondary growth oocytes (Figure 2.1E). Their ooplasm stains positively with alcian blue and the yolk globules and zona radiata stain selectively with mentanil yellow. By the time vitellogenic oocytes are fully grown, their entire cytoplasm is filled with lipid droplets and egg yolk. In histological sections, the yolk is present both as yolk globules and as amorphous yolk masses that are likely artifacts of globule breakdown during histological processing (Figure 2.4A). The follicle

16

cell layers surrounding vitellogenic oocytes are well developed, as is the zona radiata, which may have a striated appearance. Most oocyte growth in teleosts occurs by receptor-mediated uptake into the oocyte of the yolk precursor protein, vitellogenin (Mommsen and Walsh, 1988; Specker and Sullivan, 1994). Vitellogenin is a large phospho-lipo- glycoprotein secreted by the liver in response to elevated levels of estrogen circulating in maturing females. Striped bass vitellogenin has recently been isolated, characterized, and used to generate specific antisera (Sullivan et al., 1991; Kishida et al., 1992). Immunocytochemical staining of striped bass oocytes verified that the yolk globules and amorphous yolk masses are vitellogenin products and that the vitellogenic oocytes described above are aptly named (Tao et al., 1993). Although there can be considerable variation between individual females or populations of striped bass, oocytes in early stages of final maturation are generally >750 #m in diameter. They appear similar to large vitellogenic oocytes but their germinal vesicle is eccentrically located, having already initiated migration (Figure 2.4A). Oocytes in later stages of final maturation (Figure 2.4B) are difficult to section using routine methods of fixation and embedment in paraffin, as they become misshapen and their ooplasm becomes brittle and porous. Better results can be obtained if the oocytes are embedded in glycol methacrylate for sectioning (Figure 2.4; Jackson and Sullivan, 1995). The progression of final oocyte maturation can most easily be followed in oocytes chemically cleared just after they are removed from the ovary by dissection or biopsy. Fresh oocytes are placed in a fixative solution (ethanol:formalin:acetic acid; 6:3:1 v/v) for a few minutes, after which the ooplasm becomes transparent and the germinal vesicle becomes visible (Pankhurst, 1985). When the sample is viewed under a dissecting microscope, the degree of germinal vesicle migration is readily apparent and germinal vesicle breakdown can be used as a definitive marker for resumption of meiosis (Figure 2.5). It is sometimes

Fig. 2.5. Fresh oocytes chemically cleared to show migration and breakdown of the germinal vesicle as a marker for progression of final oocyte maturation. From upper left to lower right, examples of oocytes exhibiting a central germinal vesicle, migrating germinal vesicle, peripheral germinal vesicle, and germinal vesicle breakdown are shown. (Photograph courtesy of W. King V, Department of Zoology, North Carolina State University).

17

necessary to post-fix the sample in 100% glycerol for complete clearing of the ooplasm to occur. It is important to observe the clearing process continuously, because the germinal vesicle will remain visible for only a short time after oocytes are placed in glycerol. Fresh oocytes can also be characterized as to their degree of maturity using a scale developed to predict the number of hours (h) until ovulation of females injected with human chorionic gonadotropin (hCG). This scale, based largely on the extent of lipid droplet coalescence in the ooplasm and the degree ofooplasm clarity, has been repeatedly published (Bayless, 1972; Bonn et al., 1976; Kerby, 1986; Rees and Harrell, 1990). It was recently calibrated to the status of final oocyte maturation based on the position and condition of the germinal vesicle by King et al. (1994a), who compared fresh and chemically cleared oocytes at various stages of maturity. The least mature (>15-h stage) oocytes have an opaque cytoplasm and a centrally located germinal vesicle and have not yet initiated final maturation. Oocytes in the 14- to 11-h stages have a progressively increasing degree of lipid droplet coalescence, ooplasm clarity, and migration of the germinal vesicle to the oocyte periphery. By the 10-h stage, their ooplasm has cleared, lipid is present in one or a few large droplets, and the germinal vesicle has migrated to a peripheral position. Germinal vesicle breakdown occurs at, or just after, the 9-h stage. Fully mature and ovulated oocytes are considered to be at the 0-h stage. Atretic oocytes (Figure 2.4C) are in the process of degradation and reabsorption. They have a highly vacuolated cytoplasm and usually lack a germinal vesicle. Often they appear to be collapsing inward and are highly irregular in shape, as opposed to the regular polygonal or circular profile of developing or maturing oocytes. The structure of the follicle cell layers may be disorganized, including what appears to be hypertrophied phagocytic cells, possibly granulosa cells (Jackson and Sullivan, 1995). 2.2.1.2 Testis Anatomical and histological features of striped bass testicular maturation were reviewed by Groman (1982). Corresponding information recently became available for the congeneric white perch (Jackson and Sullivan, 1995). The thin testis wall is made up of fibrous connective tissue, melanocytes, smooth muscle cells, collagen fibers, vascular elements, and nerves. Its periphery is covered by the mesorchia. The testis contains an organized system of seminiferous tubules (lobules) that radiate perpendicularly from a central longitudinal collecting duct (ductus deferens) and terminate blindly beneath the peripheral testis wall (Figure 2.6A-C). Seminiferous tubules are encapsulated by a basement membrane, which may be juxtaposed to the basement membranes of adjacent tubules. Interstitial (intertubular) cells are few in the recrudescing testes. The tubule lumen, often filled with spermatozoa in mature males (Figure 2.6C), connects with the collecting duct near the center of the testes lobe. The wall of the collecting duct contains smooth muscle and connective tissue. Its lumen is lined by low cuboidal epithelium containing mucous cells near its connections with seminiferous tubules. As noted earlier, the two collecting ducts merge caudally near the genital pore. The striped bass testis is of the unrestricted spermatogonial type typical of Salmoniformes, Perciformes, and Cypriniformes (Grier, 1981; Groman, 1982). Spermatogonia are distributed randomly within the germinal epithelium of the tubule, and spermatogenesis occurs along its entire length. Intratubular cell types include Sertoli cells and the germ cells. The Sertoli cells encapsulate cysts of germ cells developing within the tubule (Figure 2.6C). Teleost Sertoli cells are known to regulate release of spermatozoa into the tubule lumen and phagocytize cytoplasm (residual bodies) cast off by spermatids as they transform into spermatozoa. They may have an endocrine function since they possess steroidogenic enzymes (Grier, 1981). The Sertoli cells likely also act as nurse cells, providing nutrition and a suitable environment for germ cell

18

Fig. 2.6. Male reproductive system of striped bass (from Groman, 1982). A~ B.

C. Do E.

Transverse section of an immature testis, 3-month-old fish (formalin; H&E). Bar = 259 #m. Oblique section through the radial testis, 3-year-old fish; fixed during February (Bouin's; H&E). Bar = 81 ~m. Transverse section of the seminiferous tubule, 3-year-old fish; fLxed during February (Bouin's; H&E). Bar = 17 ~zm. Germinal cells of the testes, 3-month-old fish (formalin; H&E). Bar = 9.5 ~zm. Mature spermatozoa, 3-year-old fish (Bouin's; H&E). Bar = 10/~m.

1. Collecting duct (ductus deferens) 2. Fibrous connective tissue of the tunica albuginea 3. Gas bladder wall 4. Genital artery 5. Genital vein

6. Head of spermatozoa 7. Interstitial cells 8. Mesorchia 9. Primary spermatocytes 10. Secondary spermatocytes 11. Primary spermatogonia

12. 13. 14. 15. 16.

Secondary spermatogonia Seminiferous tubule (lobule) Spermatids Spermatozoa Tails of spermatozoa

19

differentiation. Peritubular boundary cells form an incomplete layer over the basement membrane of the tubule, which allows blood vessels and interstitial Leydig cells to border directly on the membrane. The major function of the Leydig cells is to secrete androgens. Spermatogenesis involves the transition of mitotically active primordial stem cells into mature spermatozoa (reviewed by Nagahama, 1986; Lofts, 1987). A residual stock of stem cells divides mitotically to produce primary spermatogonia that undergo a series of mitoses to produce a cyst of secondary spermatogonia (Figure 2.6C). The secondary spermatogonia transform mitotically into primary spermatocytes and then undergo the first meiotic division to produce secondary spermatocytes (Figure 2.6C). The second meiotic division produces spermatids. During the final stage of spermatogenesis (spermiogenesis), spermatids differentiate into flagellated spermatozoa (Figure 2.6E). The developing germ cells are enclosed within germinal cysts formed by enveloping processes from the Sertoli cells. Maturation of the germ cells is synchronous within each cyst. As spermatogenesis proceeds, the cysts enlarge and extend toward the tubule lumen. When spermiogenesis is completed, the Sertoli cell processes forming the cyst wall separate and the cyst and tubule become continuous, liberating spermatozoa into the tubule lumen (Grier et al., 1980). Mature sperm are stored in the collecting ducts. Spermiation (production of milt) involves a hormone-dependent thinning (hydration) of the semen (Nagahama, 1986). Striped bass and related species are seasonal spawners that undergo a characteristic cycle of structural changes within the testis associated with maturation. Stages of maturation can be distinguished by the cytoplasmic and nuclear morphology of the germ cells present, the relative intensity ofspermatogenesis versus spermiation, and the degree of collecting duct development. Grier (1981) identified six stages of testicular development in annual spawning teleosts: (I) spermatogonial proliferation, (II) early recrudescence- spermatogonia and spermatocytes present, (III) mid recrudescence - all stages of sperm development present, (IV) late recrudescence - tubules filled with sperm and the number of developing sperm cysts is declining, (V) functional maturity- tubules filled with sperm, little if any spermatogenesis occurring, and (VI) post spawning mitotic stem cells restock the spent tubules. We are unaware of any published report on changes in testicular histology during the annual gametogenic cycle of striped bass. However, Jackson and Sullivan (1995) verified that male white perch have a seasonal pattern of gonadal maturation consistent with the above scenario. -

2.2.2 Neuroendocrine System Most reproductive processes in teleosts are subject to neuroendocrine control. The hypothalamus-pituitary-gonad axis is the central pathway involved (Figure 2.7). This section reviews the structure and function of the teleost reproductive neuroendocrine system emphasizing what is known about striped bass and related species. 2.2.2.1 Brain and hypothalamu.s Under the influence of environmental and endogenous factors, the brain orchestrates reproductive functions (Kah et al., 1993). The hypothalamus forms part of the base of the brain and is located just above the pituitary gland. Neurons originating locally or in other brain centers course through the hypothalamus and release secretory products into the blood supply of the pituitary gland or directly into the gland itself(Gorbman et al., 1983). These include neuropeptides, monoamines, and amino acids which act directly or indirectly to regulate gonadotropin (GTH) release by the pituitary (Peter et al., 1991). Gonadotropin-releasing hormone (GnRH) plays a pivotal role in regulation of GTH release and has by far received the most attention. It is a small peptide hormone composed of 10 amino acids.

20

Environmental stimuli

Internal stimuli

(SENSO ~ RY STRUCTURE~ S )

I

BRAIN

Neurotransmitters

+/-

HYPOTHALAMUS ,

GnRH

Dopamine

+/-

+/- 4.. Other pituitary hormones +/-

Non-pituitary Factors

GTH-I, GTH-I! +

+/-

r

+ l _ L"

GONAD

Other gonadal factors

E~s/Sper~ Fig. 2.7. Diagram of the hypothalamus-pituitary-gonad axis in teleosts. Environmental or internal stimuli are transduced by the brain to activate neuronal pathways resulting in secretion of neurotransmitters that regulate pituitary function. Pituitary gonadotropin (GTH-I or GTH-II) release is stimulated (+) by GnRH and inhibited (-) by dopamine, as well as being regulated by other brain factors. Gonadotropin and other pituitary and non-pituitary factors regulate steroidogenesis and gametogenesis by the testis and ovary. Gonadal steroid hormones and other factors produced by the gonads exert positive or negative (+/-) regulatory effects on the brain, hypothalamus, and pituitary in addition to the gonad itself. Full activation of the axis culminates in production of gametes and spawning.

21

2.2.2.1.1 Gonadotrop&-releasinghormone Several different forms of GnRH have been identified in teleost brains, usually with more than one form detected in a single species (Sherwood et al., 1994). GnRH shows a complex pattern of distribution in the brain and the relative abundance of specific forms of GnRH also changes with brain region (Kah et al., 1993). Of special interest is identification of the specific form of GnRH delivered to the pituitary gland in a given species. Individual GnRHs are generally named for the species in which they were first discovered. Until recently, most teleosts were believed to have two major forms of GnRH in their brain; a species-specific form that in most fish is salmon GnRH (sGnRH) and a form that appears to be present in all non-mammalian vertebrates, chicken GnRH-II (cGnRH-II). Recently, it was discovered that highly evolved perciform fish like sea bream and striped bass possess three forms of GnRH (Table 2.1); sGnRH, cGnRH-II, and a newly discovered form of GnRH (Powell et al., 1994). The novel GnRH has been characterized, its gene cloned from both sea bream and striped bass, and has been named sea bream GnRH (sbGnRH). The sbGnRH appears to be the endogenous stimulator of GTH secretion and spawning in sea bream and striped bass (Powell et al., 1994; Gothilf et al., 1995a, b; Zohar et al., 1995a). Zohar and colleagues demonstrated that, although all three forms of GnRH are present in the brain of sexually mature females, sbGnRH is the dominant GnPJ-I found in the pituitary gland of fish undergoing final oocyte maturation. The sbGnRH was present at levels 500-1000 times greater than sGnRH, whereas cGnRH-II was below detection limits. There is good evidence that the lack of natural final maturation and volitional spawning by striped bass and many other fish in captivity reflects a lack of GTH secretion by the pituitary gland (Zohar, 1989; Zohar et al., 1995b). The block to reproduction likely results from some disruption of GnRH synthesis or release. Maturation and spawning of captive striped bass can be induced by injected or implanted GnRH analogues (Hodson and Sullivan, 1993; Woods and Sullivan, 1993; King et al., 1994a; Mylonas et al., 1995). Discovery of the form of striped bass GnRH most relevant to GTH secretion (sbGnRH) paves the way to understanding neuroendocrine mechanisms underlying the block to spawning and sets the stage for developing new super-active GnRH analogues for spawning induction. 2.2.2.1.2 Dopamine Our general knowledge of how the brain regulates GTH release in striped bass is still rudimentary. Of the several brain factors other than GnRH that are known to regulate GTH release, only dopamine has received any attention in this species. Dopamine is the primary neuroendocrine factor recognized as an inhibitor of GTH release in teleosts. It directly inhibits GnRH-stimulated GTH release and likely desensitizes the pituitary to GnRH by down-regulating GnRH receptors (Peter et al., 1991). Dopamine antagonists such as pimozide and domperidone have been shown to be potent stimulators of GTH release, capable of inducing

Table 2.1. Amino acid sequences of the GnRH peptides identified in striped bass (data from Powell et al., 1994; Gothilf et al., 1995a).

Sea Bream

1 2 3 4 5 6 7 8 9 10 pGlu-- His -- Trp -- Ser-- Tyr-- Gly-- Leu -- Ser-- Pro -- Gly-- NH2

Salmon

pGlu-- His-- Trp-- Ser-- T y r - - G l y - - Trp-- Leu-- P r o - - G l y - - N H 2

Chicken-II

pGlu -- His -- Trp -- Ser-- His -- Gly -- Trp -- Tyr-- Pro -- Gly -- NH2

22 maturation and spawning of a variety of teleosts, especially when injected in combination with GnRH preparations (Peter et al., 1986). Domperidone (DOM) is commonly administered along with a potent synthetic analogue of GnRH (GnRHa). The combined treatment, termed the "Linpe" method (Peter et al., 1988), is now marketed as Ovaprim| by Syndel Laboratories, Canada. Ovaprim| or a solution of GnRHa plus DOM at the same concentrations used in the commercial preparation (GnRHa + DOM) were both shown to be effective inducers of final oocyte maturation and ovulation in striped bass (King et al., 1994a). However, GnRHa alone was equally as effective as GnRHa + DOM, whereas DOM alone failed to induce maturation or its associated changes in circulating levels of sex steroids. The possibility that dopaminergic inhibition of GTH release may not be strongly operative during the periovulatory period in striped bass warrants further investigation. 2.2.2.2 Pituitary gland The striped bass pituitary gland is structurally typical of teleost fish (Gorbman et al., 1983). Its anatomical regions and histochemistry have been illustrated and most ofthe major cell types at least tentatively identified (Groman, 1982; Huang and Specker, 1994). The pituitary is made up of two parts, the neurohypophysis and the adenohypophysis. The adenohypophysis consists of three distinct regions, including the rostral pars distalis (RPD), proximal pars distalis (PPD), and pars intermedia (PI). The neurohypophysis (NH), rich with neurons that regulate pituitary function, interdigitates with the PPD and deeply invades the PI (Figure 2.8). Excellent micrographs of the striped bass pituitary gland were recently published by Huang and Specker (1994). They definitively identified the somatotrops (growth hormone-producing cells or GH cells) and prolactin cells (PRL cells) by immunocytochemistry. The GH cells are mainly present in the PPD as isolated cells, clusters of cells, or multilayered cords of cells proximal to neural tissue extending into the PPD. The RPD is dominated by PRL cells forming a compact mass. Some isolated or clustered immunoreactive (ir) GH cells were found in the RPD and some irPRL cells were noted in the PPD.

Fig. 2.8. Diagram of a mid-parasagittal section of the striped bass pituitary gland. The major areas of the gland are shown. NH, neurohypophysis; RPD, rostral pars distalis; PPD, proximal pars distalis; PI, pars intermedia.

23

Table 2.2. Distribution of pituitary cell types of striped bass (data from Huang, 1992; Huang and Specker, 1994). Stain and Color Location

Cell type H&E

Tetrachrome

Al-PAS-orange G

RPD

PRL cells ~ putative 2 ACTH cells

red -

pale red pale orange

pale red pale orange

PPD

GH cells ~ putative GTH cells putative TSH cells

red -

orange blue blue

orange violet blue

PI

putative MSH cells putative SL cells

red

-

purple

RPD, rostral pars distalis; PPD, proximal pars distalis; PI, pars intermedia. H&E, hematoxylinand eosin; Tetrachrome, Herlant's tetrachrome;A1-PAS-orangeG, alcian blue-periodicacid Schiff's reagent-orangeG. 1identityconfirmed by immunocytochemicalmethods 2tentativelyidentified from morphology,tinctorial properties, and distribution.

Other cell types in the striped bass pituitary have been identified only on the basis of location, morphology, and histochemical staining in comparison with the same cells in other teleosts (Table 2.2). The PRL cells are separated from neural tissue by a border of cells tentatively identified as adrenocorticotropic hormone-secreting cells (ACTH cells). Putative gonadotrops (GTH-producing cells; GTH cells) were noted by Groman (1982) in the ventral region of the PPD and along its junction with the PI. Huang and Specker (I 994) also noted unstained degranulated cells amongst cords of GH cells in the PPD of pituitaries from fish taken on their spawning grounds and speculated that they might be exhausted GTH cells. In salmonids, GTH cells are located in and around the glandular cords Of the PPD in close association with GH cells (Nozaki et al., 1990a). Putative GTH cells, identified by Groman (1982) from their staining properties, are also present in islands toward the periphery of the RPD. They possibly correspond to the unstained cell clusters located amongst PRL cells noted by Huang and Specker (1994). The PPD also contains putative thyrotrops (TSH cells), cells that secrete thyroid-stimulating hormone (TSH). The PI is dominated by chromophobic putative melanotrops (MSH cells), cells that secrete melanocyte-stimulating hormone (MSH), along with PAS-positive cells tentatively identified as somatolactin cells (SL cells). Somatolactin is a novel pituitary hormone in the GH/PRL family recently discovered in the teleost pituitary (Rand-Weaver et al., 199 la, b). 2.2.2.2.1

Gonadotropins

Recently, it was discovered that most teleost fish possess two GTHs: GTH-I and GTH-II (Kawauchi et al., 1989; Xiong et al., 1994). The biochemistry and physiology ofthe two GTHs is best known in salmonid fish, where they show structural homology and functional analogy to the mammalian GTHs, follicle stimulating hormone and luteinizing hormone, respectively. The two salmonid GTHs are chemically distinct, the products

24

of different genes and pituitary cell types. They are elevated in the pituitary gland and blood plasma during different phases of the reproductive cycle in both sexes where they have distinct but overlapping spectra of physiological action (Swanson, 1991). GTH-I appears to be involved primarily in regulating gametogenesis, whereas GTH-II seems to be mainly involved in control of final oocyte maturation, spermiation, and spawning (Xiong et al., 1994). Studies of GTH physiology in non-salmonid teleosts have been largely confirmatory of the above scenario for sequential control of the reproductive cycle by GTH-I followed by GTH-II. Until recently, research into the reproductive physiolog3' of striped bass has been hampered by lack of methods to measure their GTHs. Vertebrate GTHs are members of a family of glycoprotein hormones, including TSH and chorionic gonadotropin (CG). They are heterodimers, each made up of an tx and [3 subunit. In a particular species, the subunit is shared by GTHs, TSH, and CG if it is present. The 13subunit is unique to each hormone and provides biological specificity (Pierce and Parsons, 1981). Using chromatographic techniques adapted from Rand-Weaver and Kawauchi (1992), the [3-subunit of GTH-II (GTH-II[3) was purified from pituitaries of mature striped bass and biochemically characterized (Swanson and Sullivan, 1991; P. Swanson, National Marine Fisheries Service, Seattle, and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). The striped bass GTH-III3 had an apparent mass (--21.4 kDa) similar to previously isolated fish GTH-III3 and its N-terminal amino acid sequence, determined over a stretch of 32 residues, showed a high degree of homology (< 85%) with GTH-II[3 from some other teleosts. Subsequently, Mananos et al. (1995) produced a purified preparation of intact GTH-II from hybrid striped bass. Biochemical and functional (steroidogenic) properties of the hormone were characterized and it was used as the basis for a sensitive enzyme-linked immunosorbent assay (ELISA). Using the ELISA, it was found that levels of GTH-II circulating in female striped bass were low (3-4 ng/mL) during late vitellogenesis, but they increased several fold (to >20 ng/mL) within 20 hours after administration of GnRHa to the fish. This is consistent with the expected maturational profile and response to GnRH of a teleost GTH-II. To date, striped bass GTH-I and its [3 subunit have eluded purification, presumably because they are never present at very high levels in the pituitary gland. Hassin et al. (1995) recently cloned and sequenced the cDNAs encoding striped bass GTH-I and -II, verifying the presence of a dual GTH system in striped bass. 2.2.2.2.2 Growth hormone and prolactin Growth hormone has been strongly implicated in regulation ofteleost reproduction, especially oocyte growth and final maturation (Le Gac et al., 1993). Information on reproductive actions of PRL in fish is more limited (Hirano, 1986; Mazzi and Vellano, 1987). A recent study of pituitary immunocytochemistry explored potential involvement of GH and PRL in striped bass reproduction (Huang and Specker, 1994). In pituitaries from striped bass maturing in seawater, GH cells were strongly labeled with a specific GH antiserum. GH cell density and intensity of staining were much lower in pituitaries from fish collected at spawning areas in fresh water. In pituitaries from fish maturing in seawater, PRL cells were strongly labeled with a specific PRL antiserum, but they had kidney-shaped nuclei suggesting inactivity. In pituitaries from fish taken on their spawning grounds, the PRL cells showed decreased intensity of staining and had polymorphic nuclei. These findings suggest there could be maturational changes in GH or PRL cell activity in striped bass. When GH and PRL were purified from pituitaries of striped bass sampled at spawning, yields of PRL were always several-fold greater than GH (Swanson and Sullivan, 1991; P. Swanson, National Marine Fisheries Service, Seattle, and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). Conversely, yields of GH were several-fold greater than PRL when the hormones were purified from pituitaries of striped bass maturing in seawater (Huang and Specker, 1994). These findings suggest that the ratio of GH to PRL in the pituitary changes during migration in anadromous striped bass. Both

25

hormones have important roles regulating salt and water balance (Hirano, 1986; Bern et al., 1991), so it is not surprising their pituitary levels would change with environmental salinity. Whether or not these changes are relevant to reproduction remains to be verified. Recent availability of purified striped bass GH and PRL sets the stage for measuring these hormones in fish maturing under constant salinity and will make it possible to test direct their direct effects on reproductive processes. 2.2.2.2.3 Other pituitary hormones Putative ACTH cells and MSH cells are present in the pituitaries of all teleosts (Schreibman, 1986), including striped bass (Table 2.2). ACTH is relevant to fish reproduction because it has an important function eliciting organismal responses to stress (Sumpter et al., 1994). It is the main pituitary factor regulating secretion of glucocorticoid hormones by the teleost interrenal gland (head kidney), although GTH may also regulate secretion of interrenal corticosteroids (Schreck et al., 1989). The actions of stress hormones and the impact of stress on reproduction is discussed further below (see section 2.2.2.4.1) The potential reproductive importance of other factors secreted by ACTH or MSH cells is still poorly understood in fishes, and completely unexplored in striped bass. TSH cells have been tentatively identified in the striped bass pituitary (Table 2.2), and TSH may regulate reproduction by controlling synthesis and release of thyroid hormones. Thyroid hormones are known to influence reproductive processes in some teleosts (Leatherland, 1987; see section 2.2.2.4.2). It will be interesting to discover if SL is produced by the striped bass pituitary and influences maturation. SL has been detected in all teleosts examined (Rand-Weaver et al., 1991 a), and there is some evidence for its involvement in control of fish reproduction (Rand-Weaver et al., 1991 a,b; Planas et al., 1992). 2.2.2.3 The endocrine gonad Steroid hormones produced by teleost ovaries and testes regulate function of the brain and hypothalamus-pituitary axis, pubertal development, acquisition of secondary sexual characteristics and behavior, gametogenesis, somatic growth, and general metabolism (Fostier et al., 1983). GTH plays a central role in regulating gonadal steroidogenesis via its interaction with receptors on specific cell types in the gonad (Idler and Ng, 1983; Nagahama, 1987b, 1993; Swanson, 1991; Planas et al., 1991, 1993; Yan et al., 1992). The main steroidogenic targets for GTH are the theca and granulosa cells of the ovarian follicle and the testicular Leydig cells. It is still unclear whether Sertoli cells contribute much to GTH-induced testicular steroidogenesis (Grier, 1981; Fostier et al., 1987; Lofts, 1987; Callard, 1991). Gonadal and extragonadal factors other than GTH can also modulate gonadal steroidogenesis in fishes (e.g., GH or PRL; see section 2.2.2.4). Teleost gonads produce a complex spectrum of steroid hormone precursors, steroid hormones, and hormone metabolites (Fostier et al., 1983). The major and most studied products are estrogens, androgens, and progestins (Figure 2.9). These typically trigger physiological effects by binding to specific receptors in the cytoplasm or nucleus of target cells (Callard and Callard, 1987), although regulation of final oocyte maturation involves steroid receptors on the oocyte plasma membrane (Nagahama et al., 1994). In addition to directly regulating maturation of the ovary and testis, gonadal steroids can indirectly control reproduction via feedback modulation of hypothalamus and pituitary function (Goos, 1987). Responses of the brain to environmental cues, hypothalamic GnRH secretion, sensitivity of the gonadotrops to GnRH, and GTH secretion are all subject to positive or negative feedback effects of steroid hormones (Peter et al., 1991; Kah et al., 1993). Some gonadal steroids and steroid metabolites have been verified to act as reproductive pheromones, coordinating mating behavior and spawning (Liley and Stacey, 1983; Sorensen et al., 1991; Stacey et al., 1991, 1994; Scott and Vermeirssen, 1994).

b~

(•Hs

H 0

HC~OH

HO

Estradiol- 1 713

1 7a,2013, di hyd ro xy-4- preg ne n- 3 -one (DHP)

H

HC~OH

a

0

Testosterone

I

HC~OH

1 7a,2013,21 -trihydroxy-4-pregnen-3-one

(2013-S)

Fig 2.9. Some gonadal steroids implicated in control of reproductive physiology in striped bass. Shown are an estrogen (estradiol-1713), an androgen (testosterone), and two progestins (DHP and 2013-S). See text for discussion of the roles these hormones play in fish reproduction

27

2.2.2.3.1 Estrogens In female teleosts, estrogens play a central role in regulating hepatic vitellogenesis (Specker and Sullivan, 1994), which entails synthesis and secretion of the egg yolk precursor, vitellogenin (VTG). VTG is then taken up into the oocytes, accounting for most oocyte growth (Selman and Wallace, 1989). VTG in the blood of female fish can be taken as a marker for the onset and stage of maturation. Estradiol- 1713(Figure 2.9) is strongly implicated as the estrogen controlling vitellogenesis in most teleosts, although other estrogens (e.g., estrone) may be important, especially early in the reproductive cycle (Mommsen and Walsh, 1988). The annual cycle of circulating estradiol-17[3 (E2) levels in relation to ovarian maturation has been described for free ranging wild striped bass (Berlinsky and Specker, 1991; see sections 2.3.1.1 and 2.3.1.1.1 and Table 2.3) and corroborated in studies of captive broodstocks (Tao et al., 1993; Woods and Sullivan, 1993; Blythe et al., 1994c). It has been established that E 2 can induce vitellogenesis in striped bass, and VTG has been isolated, biochemically characterized, and measured along with E2and testosterone in blood plasma of adult females at all stages of ovarian maturation (Sullivan et al., 1991; Kishida et al., 1992; Tao et al., 1993; Blythe et al., 1994c). Testosterone (Figure 2.9) serves as a precursor for E2 synthesis in the ovary (Nagahama, 1987a), and both steroids have important direct effects on functioning of the hypothalamus-pituitary axis (Goos, 1987) and reproductive behavior (Fostier et al., 1983) as well as other aspects of maturation. 2.2.2.3.2 Androgens Dominant androgens in male teleosts include, but are not limited to, testosterone (Figure 2.9) and 11-ketotestosterone (11-KT). Typically, circulating levels of these androgens are low in regressed fish, increase to peak levels during spermatogenesis, and then decline just before or during spermiation (Fostier et al., 1983, 1987; Liley and Stacey, 1983). Their individual contribution to different aspects of male reproductive physiology is not fully understood (Billard et al., 1982, Lofts, 1987). Testosterone is thought to be a stimulus for the onset of spermatogenesis and can serve as an intermediate in biosynthesis of 11-KT. 11-KT has been linked to induction of spermiation (milt production) in some species. Miura et al. (1991, 1994) recently demonstrated that 11-KT can support all aspects of spermiogenesis in in vitro cultures of eel (Anguillajaponica) testes, including mitosis of spermatogonia, spermatogenesis, and cytological activation of Sertoli cells. Both of these androgens are believed to influence acquisition of secondary sexual characteristics and reproductive behavior (Liley and Stacey, 1983, Lofts, 1987). Maturational changes in circulating levels of testosterone and 11-KT have recently been described in relation to the developmental status of the testis for captive striped bass broodstocks (Woods and Sullivan, 1993; Blythe et al., 1994c) and wild fish (see section 2.3.1.2 and Figure 2.12). 2.2.2.3.3 Progestins Progestins, specifically C2~ steroids structt;rally related to progesterone (Figure 2.9), are well known to regulate final oocyte maturation (FOM) in teleosts. In salmonids, 17t~-2013-dihydroxy-4-pregnen-3-one (DHP) is the final maturation-inducing steroid hormone (MIH) (Nagahama and Adachi, 1985; Nagahama et al., 1987a-c). Evidence is accumulating that in sciaenids, highly evolved perciform fish like striped bass, 17t~-2013-21-trihydroxy-4-pregnen-3-one (2013-S)is the MIH (Patino and Thomas, 1990; Thomas, 1994). Increased blood titers of GTH-II are known to induce a surge in MIH levels during FOM (Nagahama, 1993). In maturing females of several teleost species, circulating MIH levels peak during FOM or in the periovulatory period (Scott and Canario, 1987). During FOM, the ovarian follicle undergoes a shift in steroid biosynthesis favoring MIH production at the expense of biosynthesis of E2 and testosterone, a process best characterized in salmonids (Nagahama et al., 1994). The shift in steroidogenesis is reflected in circulating levels of estrogens, androgens, and MIH. The MIH binds to specific receptors on the oocyte plasma membrane,

28

initiating production of a maturation promoting factor (MPF) that mediates FOM. The cell and molecular biology of MPF regulation and action is complex and beyond the scope of this chapter. The reader is referred to Nagahama et al. (1994) for a recent review of this topic. In striped bass, circulating levels of both DHP and 2013-S increase in females pharmacologically induced to undergo FOM and ovulation (King et al., 1994a). At the same time, E2, and testosterone concentrations decrease to low levels. The same pattern of change was seen in naturally-maturing striped bass captured on their spawning migration (see sections 2.3.1.1 and 2.3.1.1.2, and Figure 2.11). Similar results were obtained in studies of white perch and white bass (King et al., 1995a). In striped bass undergoing FOM, levels of immunoreactive 2013-S are generally two to three times greater than DHP levels, but about half of the 2013-S measured is a putative metabolite of 2013-S lacking ability to induce FOM (King et al. 1994a,b). Collectively, these findings suggested that either DHP or 2013-S might be the hormone responsible for inducing final maturation in Morone species. An in vitro culture system was established to investigate endocrine regulation of FOM in striped bass (King et al., 1994b). It was found that cultured ovarian fragments produce both DHP and 2013-S in response to GTH (hCG) in vitro, and they do so at the expense of E2 and testosterone production. The steroidogenic response paralleled the course of FOM and generally mirrored the profile of circulating steroid hormones seen in maturing females. Of 16 structurally-related steroids, only DHP and 2013-S could induce FOM in vitro within a physiologically relevant range of concentrations. Cultured ovarian fragments from white perch and white bass also produce appreciable quantities of DHP and 2013-S as they undergo hCG-induced FOM, and both hormones were equipotent inducers of in vitro FOM in white perch follicles (King et al., 1995a). The question of which of these two steroids is the MIH in Morone species may now be resolved. King et al. (1995b) recently found that specific binding sites on ovarian membranes with characteristics of the oocyte MIH receptor exist for 20~-S but not DHP, strongly suggestingthat 20~-S is the MIH in striped bass. The MIH may also play an importantrole in the final stagesof maturation in males (Nagahama, 1986). For example, DHP is a major product of testicular steroidogenesisduring spermiationof salmonids and has been strongly implicated in regulationof sperm maturationand hydrationofthe seminalplasma(Sakai et al., 1989a,b; Miura et al., 1994).

2.2.2.4 Other endocrine organs 2.2.2.4.1 Stress hormones The two main neuroendocrine components involved in responses of teleosts to stress are the hypothalamus-pituitary-interrenal (HPI) axis and the sympathetico-chromaffin system (SCS). A diagram of these components is shown in Figure 2.10. The effects of stress are initially mediated by the central nervous system; stresses are perceived and the information is integrated in the brain. In the HPI axis, stress signals are transduced through the hypothalamus to the ACTH cells in the form of corticotropin-releasing hormone (CRH). ACTH cells and other endocrine cells in the pituitary may also be activated to respond to stress by other brain secretions or via direct innervation (Donaldson, 1981). ACTH secreted by the pituitary gland elicits interrenal production of glucocorticoids, mainly cortisol. An important function of cortisol is mobilization of glucose (metabolizable energy) by stimulating glycolysis and gluconeogenesis from glycogen or protein and lipid stores, respectively. In the SCS, direct sympathetic nervous stimulation of the chromaffin cells in the head kidney (interrenal) leads to almost immediate release of catecholamines, principally adrenaline and noradrenaline (Mazeaud and Mazeaud, 1981). These mainly target the respiratory and cardiovascular system, increasing gill ventilation and heart rate. The principal physiological effect of dual short-term activation of the HPI and SCS axes is delivery into the circulation of increased oxygen and glucose to provide energy for dealing with the emergency (stress).

(stress

Stimuii ) .

.

.

.

Brain "~ ~.~ypothalam ~ u s'~

(r

~nterior Pitui'tary~ (ACTH Cells) .... J

(~ Head Kidney ~ L (Chromaffin Ce!ls)J

(~Other P-tuitary ' ) k~ Ce Is

~nterrenai. CellS) Q~lucocorticoicls) .. (Cortisol)..

Catecholamines -"~

drenaline, nora.drenaline~

1

,

z"

.'e0ro0uctiv 'e'

I

Dysfunction

Fig. 2.10. Diagram of the neuroendocrine responses to stress that influence reproduction in teleosts.

~

30

The above scenario illustrates the adaptive nature of neuroendocrine responses to stress. However, it is well known that profound or prolonged stress can severely disrupt fish reproduction. Symptoms can include reduced pituitary and plasma GTH levels, lowered plasma concentrations of gonadal steroids, reduced gonadal growth, failure to reproduce, or a decrease in the number or quality of gametes if spawning is achieved (Sumpter et al., 1994). Non-reproductive effects of continuous exposure of fish to high circulating titers of cortisol and catecholamines include reduced survival, decreased body growth, and impairment of immune function leading to increased susceptibility to disease (Pickering, 1981). 2.2.2.4.2 Thyroid hormones Thyroid hormones have been reported to act synergistically with GTH in teleosts, enhancing gonadal steroidogenesis and oocyte development (Leatherland, 1987; Cyr and Eales, 1988a, b; Sullivan et al., 1989; Soyano et al., 1989, 1993). Specific receptors for thyroid hormones have been detected in the ovaries of some teleosts (Chakraborti et al., 1986; Soyano et al., 1989), lending credence to the view that the ovary is a thyroid hormone target. The striped bass thyroid gland, like that in most other teleosts, consists of individual and clustered follicles scattered around the ventral aorta (Groman, 1982). The teleost thyroid normally produces mainly thyroxine (T4), which is enzymatically deiodinated in peripheral tissues to form triiodo-thyronine (T)~ "I"3is the principal thyroid hormone that binds to nuclear receptors in the target tissues to exert cellular effects (Eales, 1985). Thyroid hormones have been extensively studied in regard to teleost embryogenesis and larval development. They are: 1) elevated in maternal plasma at specific times during oogenesis, 2) deposited at high levels in egg yolk, and 3) taken up and cleared by developing embryos (Brown and Nunez, 1994). Several experiments performed on striped bass indicated that thyroid hormones of maternal origin may be important to embryos and larvae (Brown et al., 1987, 1988, 1989). Injection of female striped bass with T3 at capture elevated "1"3concentrations in ovulated eggs and led to increased larval survival, growth, and swimbladder inflation. Beneficial effects of thyroid hormone treatment on developing fish have been demonstrated for a number of other teleosts as well (Brown and Nunez, 1994). However, detrimental effects can also be observed, especially when very high doses of hormone are administered to embryos or larvae. The route and time of hormone administration may also be of consequence. For example, immersion of larval striped bass in solutions containing various doses ofT3 had a clear detrimental effect on growth and survival (Huang et al., 1996). More needs to be known about the biology of thyroid hormones in developing striped bass before thyroid hormone treatments are broadly applied in aquaculture. 2.2.2.4.3 Insulin and insulin-like growth factors The main known mechanism for GH action is through its stimulation of localized production of insulin-like growth factor-I (IGF-I) which, in turn, exerts effects on target tissues. The mechanism of GH action in fish reproduction is not known, but plasma levels of immunoreactive IGF-I increase in fish after GH administration (Funkenstein et al., 1989; Drakenberg et al., 1989) and salmon GH stimulates production of IGF-I mRNA in cultured salmon hepatocytes in vitro (Duan et al., 1993a, b). It was recently discovered that purified striped bass GH stimulates in vitro IGF-I mRNA production by cultured salmon hepatocytes with a potency similar to salmon GH (Swanson and Sullivan, 1991; P. Swanson and C. Duan, National Marine Fisheries Service, Seattle, WA, personal communication). This finding implies that maturational effects of GH in striped bass could be mediated by increased IGF-I. Insulin, GH, and IGF-I are all known to regulate VTG uptake (growth) by vertebrate oocytes (Specker and Sullivan, 1994). For example, Tyler et al. (1987) reported that insulin stimulates VTG uptake by trout oocytes in vitro in a range of doses comparable to GTH-I. Insulin and IGF-I receptors have been demonstrated in carp ovaries, with substantial levels oflGF-I receptors predominating and increasing as the spawning season is approached (Gutierrez et al., 1993). As noted, GH cells in vitellogenic striped bass were strongly labeled with a specific GH antiserum, and GH cell density and

31

intensity of staining decreased in mature (spawning) fish. It remains to be verified whether GH acts directly or via IGF-I induction to regulate hepatic vitellogenesis, oocyte growth, gonadal steroidogenesis, or final oocyte maturation in striped bass. 2.3 REPRODUCTIVE FUNCTION 2.3.1 The Reproductive Cycle 2.3.1.10ogenesis Oogenesis in striped bass is of the single clutch, group synchronous type (Wallace and Selman, 1981). One clutch of oocytes is recruited through development, maturation, and ovulation for the single annual spawning. The remaining oocytes are retained in the ovary in early stages of development for recruitment in subsequent reproductive cycles (years). This pattern of ovarian maturation differs from that seen in the congeneric white bass and white perch, which are multiple clutch, group synchronous species that simultaneously recruit several batches ofoocytes for repeated spawning events during a brief annual spawning season (Jackson and Sullivan, 1995; Berlinsky et al., 1995a). 2.3.1.1.10ocyte growth Berlinsky and Specker (1991) were first to describe the reproductive cycle of female striped bass in any detail. Changes in circulating levels of estrogen (E2) and androgen (testosterone) were evaluated in relation to gonadosomatic index (GSI; [gonad weight x body weight~] X 100) and ovarian lipid content (%) at defined stages of ovarian development in wild Atlantic coast striped bass. The pattern of change in circulating steroid and vitellogenin (VTG) levels was subsequently evaluated in relation to gonadal growth or histology of captive striped bass broodstocks (Tao et al., 1993; Woods and Sullivan, 1993; Blythe et al., 1994c), and the results were largely confirmatory. Table 2.3 shows representative levels of E2, testosterone, and VTG in females sampled in various seasons and stages of ovarian maturation. Circulating levels of E2, testosterone, and VTG are low to non-detectable in female striped bass during Summer. Early secondary growth oocytes may be present in the ovary during much of the year as this stage of oocyte development can be initiated more than a year in advance of when the oocytes will be spawned (Specker et al., 1987). Adult female striped bass in the Chesapeake Bay area may initiate vitellogenesis as early as late-September, although there can be an appreciable difference between stocks with latitude. The onset ofvitellogenesis coincides with the main phase of oocyte growth and is characterized by minor increases in plasma E2 and testosterone levels with substantial and sustained elevation of circulating VTG levels (see Figure 2.15). The low plasma steroid levels are maintained until the prespawning period, when E2 levels peak sharply with or without a corresponding increase in testosterone (see also Figure 2.15). Usually, circulating E2 and testosterone levels are well correlated. All of the reproductive parameters are ata minimum in spent females just after the spawning season. The sharp increase in plasma E 2 and testosterone levels before spawning is a regular feature of gametogenesis in female white perch (Jackson and Sullivan, 1995) and white bass (Berlinsky et al., 1995a), as well as striped bass (Blythe et al., 1994c). This prespawning increase in gonadal steroids may be involved in feedback regulation of pituitary GTH secretion (Goos, 1987) because it is not linked to increases in circulating VTG levels (Tao et al., 1993;Woods and Sullivan, 1993)although it is accompanied by an increase in oocyte growth (Blythe et al., 1994a, c). Presumably, oocytes at this time are engaged in a rapid terminal phase of VTG uptake or entering early stages of final maturation -- processes mediated by GTH-I or -II in teleosts (Swanson, 1991; Nagahama et al., 1994).

32

Table 2.3. Levels of estradiol- 1713(ng-mL4), testosterone (ng'mL~), and vitellogenin (#g.mL~) in blood plasma of captive female striped bass broodstock (N=3) repetitively sampled on various dates and at different stages of oocyte development (PG, primary growth; ESG, early secondary growth; E-VTG, early vitellogenesis; L-VTG, late vitellogenesis). The mean + SEM is shown (data from Tao et al., 1993). Date

Estradiol- 1713 Testosterone

Vitellogenin

Oocyte stage

Aug 20

0.1 + 0.1

0.2 + 0.1

ND ~

PG

Oct 29

0.4 + 0.1

0.3 + 0.2

104 + 79

ESG

Jan 04

0.7 + 0.1

0.4 + 0.2

689 + 92

E-VTG

Mar 01

2.4 + 0.7

0.7 + 0.2

840 + 54

L-VTG

~ND indicates non-detectable (< 4.5 ~g-mL "~) Striped bass undergo much of oogenesis without a major elevation in E2 or testosterone levels. This pattern of gonadal recrudescence in the face of low circulating steroid levels seems typical of highly advanced perciform fish (Pankhurst and Carragher, 1991). In studies cited above, low steroid levels do not appear to be related to reproductive dysfunction associated with stress of handling or confinement because: 1) gonadal steroid levels were similar between wild fish bled immediately after capture (Berlinsky and Specker, 1991) and captive fish sampled immediately after being netted from home tanks where they had lived for months to years (Woods and Sullivan, 1993), and 2) selected captive females were induced to spawn and produced viable progeny at rates comparable to wild fish (Woods and Sullivan, 1993; Blythe et al., 1994a). The gametogenic cycle described above appears to represent normal reproductive development of female striped bass. Vitellogenesis in striped bass is induced by E2 (Kishida et al., 1992), which poses the question of how females generate and sustain high circulating VTG levels in the face of low plasma E2 titers (Tao et al., 1993; Blythe et al., 1994c). One possibility is that other estrogens, such as estrone, may prime the liver to respond to E2 or otherwise potentiate vitellogenesis. Although substantial evidence for this scenario has been developed for rainbow trout (van Bohemen and Lambert, 1981; Van Bohemen et al., 1982a,b), analysis of organic extracts of plasma from vitellogenic striped bass by gas chromatography and mass spectroscopy failed to detect appreciable quantities ofestrone (J.G.D. Lambert, Department of Experimental Zoology, University of Utrecht, and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). The answer may lie in the fact that E2 has the capacity to sensitize the liver by up-regulating its own receptor (Mommsen and Walsh, 1988) so that a relatively constant estrogenic stimulus can provide an increasing vitellogenic response. As the main precursor to egg yolk, VTG is centrally important to the process of oocyte growth. Most oocyte growth can be attributed to VTG uptake. Evidence is accumulating that VTG is sequestered by teleost oocytes via receptor-mediated endocytosis (Specker and Sullivan, 1994). An ovarian VTG receptor from white perch with properties similar to VTG receptors in Xenopus, chickens, and some other teleosts, was recently characterized (Berlinsky et al., 1995c; Tao et al., 1996). After being taken up into the oocytes, teleost VTG is cleaved into yolk proteins, which may include lipovitellin (Lv), phosvitin (Pv), and the lY-component (Specker and Sullivan, 1994). In addition to providing proteins for embryogenesis, VTG carries ions and minerals, such as phosphorus and calcium, that are needed for proper metabolism and skeletal development. VTG also transports lipids to growing oocytes, being about 20% lipid by weight in most teleosts examined.

33

A summary of recent analyses of the composition of striped bass VTG is provided in Table 2.4. It has been confirmed that striped bass VTG is a phospho-lipo-glycoprotein (Kishida et al., 1992; Tao et al., 1993), but it is still uncertain as to whether there is more than one form of VTG in this species. Analysis of native VTG by polyacrylamide gel electrophoresis (PAGE) reveals two closely-spaced protein bands, but VTG appears as a single band (apparent subunit) after reduction and sodium dodecylsulfate (SDS)-PAGE (Tao et al., 1993). Comparison of the relative electrophoretic mobilities of the native protein and its apparent subunit suggest that VTG normally circulates as a dimer in striped bass (Kishida et al., 1992). Maturation of female striped bass is accompanied by massive deposition of lipids in the ovary (Berlinsky and Specker, 1991). Most of the lipids carried by VTG are phospholipids, the majority of these in the form ofphosphatidyl choline (Table 2.4). Changes in circulating lipids were recently evaluated during the gametogenic cycle of captive striped bass broodstock (Lund et al., 1995; A. Place and E.D. Lund, Center of Marine Biotechnology, University of Maryland, and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). Phospholipids are the dominant class oflipids in maturing females. Their profile of change during the gametogenic cycle mirrors that previously reported for VTG in the same fish (Tao et al., 1993), but is shifted somewhat out of phase from the VTG cycle. Circulating triglycerides undergo a similar cycle in maturing females but are usually present at levels less than half those seen for phospholipids. Interestingly, phospholipids do not accumulate much in the ovary. The increase in ovarian lipids during maturation is primarily due to wax esters, which are not detected in the plasma of vitellogenic fish. Triglycerides accumulate to a lesser extent in the ovary, and tend to vary inversely with wax esters in abundance. It has been suggested that wax esters are synthesized de novo by the growing oocytes and triglycerides may act as a temporary intermediate for storage of lipids delivered to the oocytes (as phospholipids) by VTG and other very low density plasma lipoproteins (Lund et al., 1995). 2.3.1.1.2 Final oocyte maturation This chapter previously dealt with cytological changes in striped bass oocytes during final maturation (see section 2.2.1.1and Figures 2.4 and 2.5) and our knowledge of hormones regulating these changes (see Table 2.4. Composition of striped bass vitellogenin (data from A. Place and E.D. Lund, Center of Marine Biotechnology, University of Maryland, and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). Parameter Molecular Mass Subunit mass A=280 nm, 1 mg.mL Protein-bound phosphorous (wt %) Total lipids (wt %) *Phospholipids (wt%) Cholesterol (wt %) Fatty acids (wt %) Triglycerides (wt %) Cholesterol esters (wt%) "Greater than 95% phosphatidyl choline

Value 373,000 daltons 170,000 daltons 0.783 2.1 + 0.2 20.1 + 1.8 15.9 + 0.38 0.62 + 0.18 1.2 + 0.39 1.2 + 0.39 0.7 + 0.21

34

section 2.2.2.3.3). Results from a recent study of naturally maturing striped bass illustrate and integrate the two topics (Figure 2.11). The fish were captured on their spawning migration into tributaries of Chesapeake Bay at various stages of FOM (King et al., 1994a). In females whose most mature oocytes were in early FOM, circulating DHP and 2013-S levels were low, whereas and E2 and testosterone levels were high. Levels of the both progestins peaked coincident with oocyte germinal vesicle breakdown (GVBD), marking resumption of meiosis, with concomitant decreases in circulating E2 and testosterone. DHP and 2013-S levels decreased substantially by ovulation, at which time E2 and testosterone levels were also low. This same profile of circulating gonadal steroids was seen in fish pharmacologically induced to undergo FOM (King et al., 1994a). It is indicative of the switch from follicular E2 and testosterone production toward synthesis of the MIH known to occur in numerous other teleosts during FOM (Scott and Canario, 1987). As noted, we strongly suspect that 2013-S is the MIH in striped bass, and DHP may be its biosynthetic precursor. Striped bass ovarian follicles must acquire "maturational competence" before their oocytes are capable of undergoing final maturation in response to MIH (Redding and Patino, 1993). Prior studies of in vitro FOM in Morone species utilized as ovary donors mostly captive broodstock or wild fish collected remote from spawning areas. Their most advanced follicles contained fully grown oocytes with a central germinal vesicle, oocytes that had not yet initiated FOM. Such oocytes do not reliably mature in response to DHP or 2013-S in vitro. To routinely obtain oocytes competent to respond to DHP or 2013-S, it was necessary to "prime" the donor females with an injection ofGTH (hCG) many hours before their ovaries were taken for in vitro culture (King et al., 1994b, 1995a). Thus, development of maturational competence in striped bass appears to be GTH-dependent. Ovaries from wild females captured on their spawning ground contain oocytes already matured to a MIH-sensitive stage (King et al., 1994b). Such fish do not require an hCG injection to induce maturational competence of their oocytes for in vitro maturation, presumably because they have already been exposed to high levels of endogenous GTH. These results infer that there is a surge in maturational gonadotropin (GTH-II) in striped bass before spawning, similar to that reported for salmonids and some other teleosts ('Nagahama et al., 1994). Recent studies indicate that acquisition of maturational competence may involve several components, including the preovulatory surge in circulating GTH-II levels, de novo synthesis ofoocyte MIH receptor, and establishment of cytological connections between the follicle granulosa cells and the oocyte (Redding and Patino, 1993). 2.3.1.1.3 Ovulation Ovulation involves expulsion of the mature oocyte from the follicle into the lumen of the ovary in preparation for spawning (Goetz, 1983). In salmonids, several days may elapse between ovulation and spawning, during which time ovulated eggs may continue to mature in the female (Springate et al., 1984; Mylonas et al., 1992). In contrast, striped bass probably spawn shortly after ovulation. Hatchery production of striped bass often involves manually stripping gametes from both parents for in vitro fertilization, and it is well known that females must be stripped of their eggs within a few minutes of ovulation to achieve adequate fertility. Oocyte maturation, ovulation, and spawning probably occur in rapid succession in naturally maturing striped bass. GTH is likely the proximal stimulus for ovulation, as a single injection of GTH (hCG) is all that is necessary to induce suitably mature females to complete maturation, ovulate, and spawn within two days (Rees and Harrell, 1990). However, ovulation in teleosts can be quite complex, involving: 1) a preovulatory surge of plasma GTH levels, 2) GTH-induced MIH production by the follicles, 3) action of the MIH on the follicle itself and on extra-follicular cells, 4) induction of intrafollicular proteases that weaken or breach the follicle wall, 5) intra- and/or extra-follicular production of prostaglandins required to stimulate ovulation, and 6) contraction of specialized follicle cells to expel the mature oocyte (Goetz et al., 1991). Virtually nothing is known about these mechanisms in striped bass.

7 7

A

9

C

Et 5 2

r./3 I

O

eq

1 2 14

3

1 7

12

7

10

1 2

m. m. 8

" 6

9

9

4

.

9

" 2

I= v 3

2

[.-,

mm

m

t l m m m m m m ,

" 0

1;4 " 1"2" 1 0 "

.

' 6

.

.

.

' 4

.

' :2 " 0

,7

zl

B

8

1 2

D

'-,-4

~.

2 ~.

3

t:l

1 2 2

14

31 12

77

10

8

6

4

2

0

[-r~ 1

lllll

14

12

l

12

2

i

10

$

6

4

2

0

H o u r s to O v u l a t i o n Fig. 2.11. Plasma levels (closed bars) of A) 17tt,2013,2 I-trihydroxy-4-pregnen-3-one (2013-S), B) 17tt,2013-dihydroxy-4-pregnen-3-one (DHP), C) testosterone (T), and D) estradiol-1713 (E2) in female striped bass captured near their spawning ground at various natural stages of FOM or at ovulation. Numbers above the bars = N; nd = non-detectable. Vertical brackets indicate SEM where N>3 or the range where N=2. "Hours to Ovulation" refers to specific stages of striped bass FOM, identified by the degree of lipid droplet coalescence and ooplasm clarity, used to predict the number of hours (h) until ovulation of striped bass (Rees and Harrell, 1990). For comparison to studies of FOM in other fishes, 15-11 h-stage oocytes show increasing degrees of germinal vesicle migration (GVM), I0 h-stage oocytes have a peripheral germinal vesicle (PGV), germinal vesicle breakdown (GVBD) occurs at the 9 h-stage, and 0 h-stage oocytes are fully mature, ovulated and fertilizable (data from King et al., 1994a).

36

2.3.1.2 Soermio~enesis

The reproductive cycle of male striped bass has received much less attention than the female cycle, possibly because fishery management schemes and broodstock management practices have largely targeted females. The gametogenic cycle of wild males from Chesapeake Bay was recently characterized. Figure 2.12 shows the percentage of germinal tissue, interstitial tissue, and sperm filled efferent ducts in their testis compared to circulating levels of androgens, testosterone and 11-KT. Histological examination of the testes in June revealed spermatogonia in lobules leading from just below the testis wall toward the interior. The testes of 80% of the fish contained residual sperm in June, whereas by July the efferent ducts of all fish were devoid of sperm. From June to September, there was a continuous proliferation of spermatogonia such that germinal cords were formed. Spermatogonia first began differentiating into spermatocytes in October, and by November large numbers of germinal cysts containing either primary or secondary spermatocytes were associated with the spermatogonial cords. The efferent ducts began to fill with sperm in December, and by the spawning season in April the testes consisted principally of sperm filled ducts. Although samples were not obtained in the months of January, February, and May, the general profile of circulating testosterone and 11-KT indicates a progressive rise in levels of both androgens between July and December, mainly during the period of spermatogonial proliferation and differentiation into secondary spermatocytes (Figure 2.12). Androgen levels were sustained between December and March, during which time the efferent ducts were developing and filling with sperm. Interestingly, there appeared to be a prespawning peak in circulating androgen levels in males, similar to the late-cycle spike of testosterone and E2 seen in female striped bass. This annual cycle in circulating androgen levels, including the mid-cycle plateau and late-cycle peak, has been confirmed by studies on captive broodstocks (Woods and Sullivan, 1993; Blythe et al., 1994c). It appears nearly identical to the pattern of circulating steroids seen in maturing male white perch (Jackson and Sullivan, 1995) and white bass (Berlinsky et al., 1995a). Testicular recrudescence in striped bass and the other Morone species follows a pattern similar to that observed in tilapia (Oreochromis aureus, Grier and Abraham, 1983) and redfish (Scienops ocellatus, Grier et al., 1987). Early testicular growth is due almost entirely to proliferation of the germinal epithelium, as shown by increases in the percentage of germinal epithelium in the testes between June and November (Figure 2.12). Serum androgen levels also increase during this time. The greatest increases in serum androgens are associated with meiotic differentiation of the germinal epithelium, development of the efferent ducts, and appearance of sperm in the ducts. Testicular regression occurs following spawning and is associated with low or non-detectable levels of serum androgens, even though the testes may remain full of sperm for months after spawning. The lack of hormonal support likely leads to resorption of unused sperm (Jackson and Sullivan, 1995), and then the cycle of germ cell proliferation and differentiation repeats itself. Circulating levels of testosterone and 11-KT covary in male striped bass during the gametogenic cycle. Testosterone can act as a biosynthetic precursor to 11-KT, and the two hormones likely interact to regulate spermatogenesis. Their individual roles controlling the male gametogenic cycle could not be inferred in the studies cited above, because of the simultaneous changes in plasma levels of both androgens. The late-cycle spike in circulating androgens coincides with the time when semen would be undergoing hydration, and either hormone may contribute to control of this process. As noted, it is possible that the striped bass MIH (2013-S) also regulates seminal hydration and spermiation, analogous to the action of DHP in salmonids (Nagahama, 1987b). Alternatively, the late-cycle spike in androgen levels could be involved in feedback regulation of pituitary GTH secretion, as proposed for females.

37

Atl

E

I

11-KT

F-1T

2

v

%

I-1IF]/

0

100

B~

80

....

J F-1

I

[~

Germinal Tissue E f f e r e n t Duct I n t e r s t i t i a l Tissue

6O

r CD r ~r 40 r~

20

Jul

Aug Sep Oct Nov Dec Jan Feb Mar Apr May Jun

Fig. 2.12. A) Serum concentrations of testosterone (T) and 11-ketotestosterone (11-KT) during the annual reproductive cycle of adult male striped bass (total leng~th46-70 cm) from the mid-region of Chesapeake Bay collected by hook and line at monthly intervals between May and December and by gill net between January and April. Blood was sampled from the fish by caudal amputation within 5 rain of capture and allowed to clot on ice. The serum was separated, extracted, and analyzed for T and 11-KT content using specific radioimmunoassays (Cochran, 1987; Cochran et al., 1988). 13) Testicular tissue composition during the annual reproductive cycle of the male striped bass. Testicular tissue samples were fixed, embedded in glycol methacrylate, sectioned, and stained (Jackson and Sullivan, 1995). The sections were examined by phase contrast microscopy at 100X, and a 10 x 10 ocular grid was used to quantify the amount of germinal epithelium, interstitial tissue, and sperm filled efferent ducts. The type of tissue under the intersection of two lines from four random locations in the peripheral portion of each of 3 sections from each testis was recorded. N = 4-12 fish per month (data from a previously unpublished study by H. Grier, Florida Marine Research Institute, Florida Department of Natural Resources, and R. Cochran, California Department of Pesticide Regulation).

38

2.3.1.3 Spawning behavior. Although much is known about artificially inducing spawning of striped bass in tanks (Smith and Whitehurst, 1990), very little is known about natural spawning. The time of peak spawning activity varies with latitude, usually coinciding with water temperatures of 16-17 ~ around the optimum for larval development and survival (Fay et al., 1983). In North Carolina, spawning occurs primarily in April and May. Peak spawning activity appears to be influenced by tidal stage, temperature, and rainfall (Westin and Rogers, 1978; Fay et al., 1983; Hocutt et al., 1990). It remains unclear whether spawning occurs predominantly during the day or at night (Setzler et al., 1980). Males arrive in spawning areas before females (Fay et al., 1983). A recent radio-tagging study in the Chesapeake Bay area (Hocutt et al., 1990) indicated that males remain on the spawning grounds for as along as 30 days while females spend about 7-10 days there. Males are more active than females, moving between freshwater spawning ~ounds and brackish estuarine areas. The estuary may serve as an intercept area where males locate gravid females. In many locales, striped bass are known as "rockfish." Most published descriptions of striped bass spawning events, commonly referred to as "rockfights," suggest that from 10 to 50 fish are involved. They refer to one female and many males darting about and splashing at the water surface. These short anecdotal descriptions indicate there is some brief and intense physical contact between fish, but they otherwise appear to swim around aimlessly for long periods, occasionally rolling over on their sides while throwing water about (Worth, 1903; Merriman, 1941; Woodhull, 1947; Lewis and Bonner, 1966; Bishop, 1975; Smith and Whitehurst, 1990). These accounts are clearly incomplete. Recently, reproductive behavior of captive striped bass broodstock was evaluated through systematic observation and videotaping of their courtship and spawning in seven meter diameter outdoor tanks (S. Salek and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). The fish were induced to mature with implanted GnRHa pellets and injected hCG (Hodson and Sullivan, 1993; Woods and Sullivan, 1993). A brief account of these observations follows. Male and female striped bass exhibit apparent courtship for more than fifteen hours before spawning. This consists of various behaviors, including fish flashing (rolling over on their side), occasionally following one another, attending (close and continuous following), aggressive chasing, and rising together in groups. Males approach and follow females for a few seconds and then break off to swim on their own. Following becomes more intense as it develops into attending. Attending later predominates and males sometimes place their nares close to the female's vent (anus and urogenital pore). Attending develops into periodic chasing when the female attempts to swim away from attending males, sometimes breaching the water surface. Males show intermittent flashing behavior as they follow, attend, and chase the female. As the female draws close to ovulation, she has a clearly distended abdomen, becomes more docile, and sometimes develops a "headstand" posture while males aggregate around her. The group (males and female) swims slowly and all rise together to the surface periodically. There is often some physical contact at this point. Spawning can involve multiple males and more than one female, but it is characterized by one female and many males releasing gametes at the water surface. Spawning begins with the female typically in the center of a group of males. They appear to guide her to the surface, her caudal fin thrashes violently from side to side often emerging from the water, but she remains stationary by "sculling" backward with her pectoral fins. Other males rapidly approach and join the group when this behavior is exhibited by the female. The female then releases one long continuous cloud of eggs for less than ten seconds. Milt release is shorter in duration than egg release but it occurs two or three times and is done by more than one male as they also thrash their caudal fins about. It is this thrashing from which the term "rockfight" probably arises. Males and the female sometimes turn on their sides (flash) before and during gamete release. After spawning, the female rapidly

39

swims away from the males but they continue to vigorously chase her. The female usually stays far enough ahead to avoid further physical contact. Sometimes another (minor) synchronous gamete release can be observed. Once a group of males has spawned, they will continue to chase the spent female or court the next available one. The observations described above, although based on numerous individual spawns, are preliminary and the data is still being subjected to quantification and statistical analyses. The goal is to construct a standard ethogram for spawning behavior of striped bass. It is already clear that mating behavior may last longer and be more complex than previously imagined. Detailed knowledge of spawning behavior by this species could be of significant benefit to aquaculture. For example, fish behavior might be used to predict the time of ovulation. This would reduce or eliminate need for repetitive handling for ovarian biopsy or abdominal palpitation of the fish to detect ovulation as required for production of hybrids by in vitro fertilization (Rees and Harrell, 1990). Normal spawning behavior could also be used as reference (control) in studies aimed at discovering and overcoming the behavioral block to hybridization between striped bass and white bass (Woods et al., 1995). It is highly desirable to achieve volitional tank spawning for hybridization of these species, because stress associated with methods for detecting ovulation and manually stripping eggs from the females can lead to high mortality rates in valuable captive broodstock. 2.3.1.4 Fertilization and egg activation Egg fertilization and activation involves numerous processes including sperm activation, physical and biochemical interactions of the sperm and egg leading to fertilization, and activation of the fertilized egg, which involves several marked changes in its structure and composition. To our knowledge, these processes have not been investigated directly in striped bass or related species. Redding and Patino (1993) provide an excellent concise treatment of these subjects based on what is known in some other teleosts. A synopsis emphasizing what little is known of striped bass follows. Striped bass spermatozoa are immotile until they make contact with the water. Spawning or diluting sperm in water activates the spermatozoa. Striped bass spermatozoa can remain active (motile) for as brief as one or two minutes after contact with the water, so it is important to avoid premature contact when attempting in vitro fertilizations. Activation may involve dilution of factors or conditions present in the seminal plasma, such as cations (e.g. K§ acidic pH, other chemical factors, or the isotonic environment. The chorion (zona radiata or "egg shell") surrounding the ovulated oocyte contains a small opening over the animal pole, the micropyle, which allows sperm to access the surface of the egg. There is evidence that the micropyle is the source in some fish eggs of proteins released into the water that lead sperm toward the egg. In some species, the surface of the egg under the micropyle may be specialized as a sperm entry site. Sperm enter the micropyle to contact the surface of the egg, triggering a number of chemical and structural changes. In some species, a "fertilization cone" forms almost immediately at the base of the micropyle, blocking fertilization by multiple sperm (polyspermy). As part of egg activation, the cortical alveoli (cortical granules) underlying the plasma membrane fuse with the membrane, discharging their contents into the perivitelline space between it and the chorion. This response is termed the "cortical reaction." Cortical alveoli in a mature striped bass oocyte are illustrated in Figure 2.13A. The contents of the cortical alveoli increase osmolality (osmotic pressure) in the perivitelline fluid drawing extracellular water into the perivitelline space and dramatically increasing the volume of the egg. This increase in volume, known as "water hardening" of the egg, is illustrated in Figure 2.13B. In some species, the cortical alveoli may also release into the perivitelline space bactericidal substances or sperm-agglutinating substances that block polyspermy. Polymerization of proteins at the outer egg surface during activation hardens the egg against physical injury.

40

Fig. 2.13. A) Apical ooplasm of mature oocytes of striped bass immunostained with an antiserum to striped bass vitellogenin (Tao et al., 1993). The yolk granules (G) and amorphous yolk (Y) show positive immunoreactivity, whereas the cortical alveoli (A), follicle cells (F), chorion (C), extracellular space (E), and lipid droplets (L) do not. Horizontal Bar -- 15 #m. B) Photograph of fertilized and water-hardened striped bass eggs. Illustrated are their chorion (C), perivitelline space (P), two or four cell-stage embryos (E), ooplasm (O), and lipid droplet (L). Horizontal bar = 500/~m. (Photograph courtesy of L.C. Woods III, Crane Aquaculture Facility, University of Maryland).

41

The net result of activation is to protect the fertilized egg against polyspermy, pathogens, and physical injury. Within minutes after fertilization, the egg completes meiosis including extrusion of the second polar body. During this time various chemical or physical (e.g., hydrostatic pressure) treatments can be used to block extrusion of the polar body, resulting in production of triploid progeny that may have advantages for commercial culture (Kerby and Harrell, 1990; see also Chapter 8). 2.3.2 Environmental Influences

Full development of commercial striped bass farming will require continuous year-round production of fingerlings in intensive aquaculture. Spawning will need to be induced at several times during the year. The ability to reproduce striped bass at any time of year will also accelerate research on problems with broodstock nutrition, induced spawning, hybridization, and developmental biolog2r (see section 2.4. Captive Breeding). This section covers recent research on controlled maturation of striped bass for out-of-season spawning. This type of research principally involves manipulation of annual cycles of daylength and water temperature to which the fish are exposed (see also Chapter 6). Fish reproductive cycles are controlled by endogenous rhythms ("biological clocks") entrained by environmental factors. In the temperate zone, the annual cycle of daylength (photoperiod) is the primary environmental cue influencing initiation of gonadal recrudescence and the time of spawning (Lam, 1983; Zohar, 1989). Water temperature also plays an important role, especially around the spawning season. Where large seasonal fluctuations in temperature occur, radical shifts in spawning time induced by artificial photoperiods can be accompanied by abnormal gonadal development or failure to ovulate unless water temperature also cycles appropriately. Both environmental variables are usually cycled together to mature fish outside the normal season. We refer to coordinated change in both daylength and water temperature as the photothermal cycle. Artificial photothermal cycles are routinely used for spawning farmed fish out-of-season (Carrillo et al., 1995; Thomas et al., 1995; Zohar et al., 1995b). For example, the peak spawning season of different groups of sea bream in Israel is shifted to one of four times during the year, three months apart. One (control) group is exposed to a year-long natural photothermal cycle. The other groups are each exposed to a similar 12-month cycle phase-shifted three, six, or nine months from the control cycle. The fish are implanted with GnRHa to initiate spawning (see section 2.4). Because individual sea bream can spawn daily for up to four months, it is possible to obtain fertilized eggs every day of the year using this approach (Zohar et al., 1995b). Early studies of striped bass demonstrated that they would mature under artificial photothermal cycles, but lack of reliable methods for inducing spawning hampered full development of the technique. HendersonArzapalo and Colura (1987) matured striped bass under a simulated natural cycle. Two females were induced by hCG injection to spawn with good egg fertility but poor embryo viability. Smith and Jenkins (1986) used a slightly modified and shifted (+three months) natural cycle. Females matured in January and early-February, two to three months before normal. Some fish were induced with hCG injections to ovulate, but few viable fry were produced. These studies indicated striped bass will mature under artificial photothermal cycles and it is possible to shift their spawning time, but practical protocols still needed to be developed for commercial aquaculture. Experimental manipulation of the timing of the gametogenic cycle should be undertaken with detailed knowledge of how it normally progresses. Accordingly, changes in circulating levels of sex steroids (E2 and testosterone) and VTG were measured monthly through two reproductive cycles of captive striped bass and evaluated in relation to the natural photothermal cycle (Tao et al., 1993; Woods and Sullivan, 1993). Blood

42

and gonadal biopsy samples were subsequently obtained from the same fish over the course of a third cycle (year) and analyzed for gonadal steroids and VTG, as well as oocyte diameter and ovarian histology (D.L. Berlinsky, L.F. Jackson, and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). Elevated plasma levels of E2 testosterone, and VTG, along with increased oocyte diameter and histological evidence of vitellogenic oocyte growth, marked the start of the annual gametogenic cycle. Subsequent decreases in gonadal steroids and VTG combined with evidence of FOM or atresia in the biopsy samples marked the end of the cycle. Results of these studies can be summarized simply with reference to common astronomical landmarks (Figure 2.14). Female striped bass in the mid-Atlantic region held under natural photoperiod and ambient Chesapeake Bay water temperature initiate ovarian maturation near the autumnal equinox. Maturation (oocyte growth) proceeds at a steady rate through the winter solstice until the vernal equinox when the oocytes are almost fully grown. There is a surge in oocyte growth and circulating levels of sex steroids around this time. Increases in water temperatures from 10 to > 16~ coincide with, and likely trigger, final maturation and spawning in April and early-May. After spawning, the ovary regresses and is quiescent with minimum maturational change around the summer solstice. At this time most oocytes in the ovary are in primary growth. Maturation is reinitiated near the next autumnal equinox. In nature, the interval between spawning and initiation of ovarian recrudescence coincides with recovery from spawning and replenishment of energy reserves. A similar interpretation of the annual reproductive cycle of males was made based on their circulating levels of testosterone and 11-KT and incidence of spermiation (Woods and Sullivan, 1993). In another study, domestic striped bass were induced to mature under compressed six- or nine-month photothermal cycles or under a 12-month control cycle (Blythe et al., 1994a, b,c). Female maturation was monitored by monthly measurement of oocyte diameters in biopsy samples, ovary diameters (measured by ultrasonic imaging), and circulating gonadal steroid and VTG levels. Ovary diameter was highly correlated with VTG levels and oocyte diameter. Some results of this study are shown in Figure 2.15. In fish held under the control cycle, changes in gonadal steroid and VTG levels, and oocyte diameter were nearly identical to those described above. Although changes in these parameters were similar in all groups of fish, the gametogenic cycle was shortened in fish held under compressed photothermal cycles. For each group of fish, the length of the reproductive cycle was clearly proportional to the length of the photothermal cycle to which they were exposed. The main endocrine effect of compressing the photothermal cycle was to shorten the period of sustained low hormone levels, during which time most oocyte growth occurs. Interestingly, rates of oocyte growth were similar in all groups of females regardless of whether they were held under compressed or normal cycles (Figure 2.15). Photothermal cues appeared to initiate and terminate the reproductive cycle, but between these times oocyte growth rate and plasma steroid or VTG levels seemed relatively insensitive to photothermal cues. At times, oocytes were growing at the same rate in different groups of fish exposed to very different prevailing daylength and water temperature. This finding illustrates the fact that it is the change (cycle) in photoperiod and perhaps water temperature, not the absolute prevailing daylength or temperature, that is relevant to controlling the timing of fish reproductive cycles (Sumpter, 1990). As a consequence of their shortened reproductive cycle, the final oocyte diameter of fish held on compressed cycles was reduced, being proportional to the length of the cycle under which they were held. Although fecundity was not effected, ovary diameter and the proportion of females that could be successfully spawned were reduced for the group held under a six month cycle, and no females from this group could be induced to spawn during a subsequent short cycle (Blythe et al., 1994c). It can be concluded from these results that there is a threshold cycle length (< nine months) below which striped bass are incapable of completing normal maturation. It was this kind of experience with compressed cycles that led to use of out-of-phase, 12-month cycles for year-round production of sea bream (Zohar et al., 1995b).

43

More research is needed to develop techniques for routine out-of-season spawning of striped bass. It seems probable that some combination of out-of-phase or compressed cycles will finally be employed. For example, the "biological clock" of juvenile fish may be shifted using compressed cycles so they can later be matured as adults at different times of year under out-of-phase, 12-month cycles. In the end, practical methods for routine photothermal conditioning of striped bass broodstock may be quite different from those used for some other species. Compressed cycles are usually used for red drum (Thomas et al., 1995), out-of-phase, 12-month cycles are routinely employed for sea bream (Sparaus aurata, Zohar et al., 1995b), and alternating light cycles of constant length are preferred for sea bass (Dicentrarchus labrax, Carrillo et al., 1995). Continuous production of hybrid striped bass will probably also require techniques for extending the spawning season so that all fish on a given photothermal cycle do not spawn at the same time. This problem was recently addressed by adapting female white bass with nearly fully-grown oocytes and spermiating male striped bass to low temperature (-~10~ early in the spawning season (R.G. Hodson, M. Hopper, S. Salek, and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). The fish were held in cold water for prolonged periods to prevent atresia so that they could be spawned late in, or after, the usual season. They were then reacclimated to appropriate temperatures for spawning and induced to mature with injected hCG. This procedure is referred to as "cold banking." Using the technique, female white bass

Autumnal Equinox Decreasing

Minimum

Decreasing

Increasing

(12L:12D)

Stage of Maturity

Winter Solstice

Initiate Spermiogenesis Vitellogenesis

Rapid Phase of Gonadal Growth and Maturation

Vernal Equinox Increasing

(12L'12D) o

16 C Spermiation and Final Oocyte Maturation

Fig 2.14. The main phases of gametogenesis in striped bass (stage of maturity) at specific prevailing conditions of water temperature (thermometer symbol) and daylength (sun symbol) shown in relation to common astronomical landmarks (autumnal equinox, winter solstice, vernal equinox). At summer solstice the gonads are quiescent, oocytes are in primary growth, and spermatogonia are the dominant germ cells in the testis.

44

I

'I l:!:}:::i:::::l

i

1.6

E t-

1.2

D

I

ii

to (/}

o

(/)

#. 0.4 ~ , ~ _ ~ ~ _ ~ . T 0.0

.9

-~'["~

- "

~'

-

0.8-

"'

"c}

0.6 W

~_

A_,~

0.0

c

1.0-

B T

s

~

T

o .=...

- -

0.0

I

E

'-- 900

/

i5 600 ~

300

o.-~

0

81

162

244

325

406

Experiment Duration (days)

Fig. 2.15. Monthly oocyte diameter (A) and plasma levels of vitellogenin (B), estradiol-1713 (C), and testosterone (I)) in female striped bass broodfish held under 6-month (closed squares), 9-month (open circles), and 12-month (closed triangles) photothermal regimes. Symbols represent the mean value for four fish, and vertical brackets indicate SEM. Heavy horizontal bars at the top of the figure indicate the interval during which females from the 6-month (open bar), 9-month (shaded bar), and 12-month (closed bar) cycles were spawned (data from Blythe et al., 1994c).

45

were successfully spawned with high fertility to produce hybrids up to three months after the normal spawning season. Similar techniques were recently used for propagation of white bass broodstock (Smith et al. 1996).

2.3.3 Puberty and Maturity Schedules Puberty involves transition of a juvenile to an adult stage and is characterized by functional maturation of the gonads (Goos, 1994). In practical terms, a maturity schedule is a demographic table for a stock of fish, showing the proportion of individuals of each gender in each age class that have undergone puberty and can be considered to be sexually mature. This kind of information is critical to fishery managers for designing harvest strategies that can sustain yields. Maturity schedules are also important to hatchery managers maintaining captive broodstock. Availability of mature adults must be known for programmed spawning (see section 2.4). Mechanisms responsible for pubertal development have not been investigated directly in striped bass, but mechanistic models have been developed for other teleosts. However, striped bass maturity schedules have been extensively studied for over fifty years. 2.3.3.1 Puberty_ By the onset of puberty, striped bass have already undergone sexual differentiation into males and females. The subjects of sexual differentiation and practical control of gender in cultured fishes were recently reviewed (Zohar, 1989), but almost nothing is known of these processes in striped bass. In immature (prepubertal) fish, the germ cells are arrested in early development as primary growth oocytes or spermatogonia, and the morphology or functional capacity of one or more components of the HPG axis is not fully developed. Platyfish (genus Xiphophorus) are best understood as regards neuroendocrine control of fish puberty (Schreibman et al., 1994). Puberty in platyfish and other teleosts involves a sequence of developmental changes in the brain, hypothalamus, pituitary gland, and gonad. These changes include: 1) the appearance of neurons containing one of various forms of GnRH or other neurotransmitters and neuropeptides in specific regions of the brain, 2) differentiation and secretory activity of pituitary gonadotrops, 3) initial activation of gonadal steroidogenesis, 4) positive feedback of steroids to further mature the hypothalamus and pituitary, 5) full functioning of the HPG axis, and 6) activation of gametogenesis culminating in production of mature eggs or sperm. The general subject of puberty in fishes was recently reviewed in detail (Schreibman et al., 1991; Goos, 1994). 2.3.3.2 Assessing Maturity_ Until recently, microscopic examination of the gonads was the only sure means for identifying whether or not an individual striped bass was mature. This procedure usually involved killing the fish and dissecting the gonads to obtain and process tissue for histological examination (Berlinsky et al., 1995b). This approach is still useful and can provide definitive verification of maturity. However, declines of certain stocks of striped bass coupled with development of domestic broodstocks has made non-lethal or non-invasive techniques for assessing maturity highly desirable. Female striped bass can be subjected to non-lethal ovarian biopsy by inserting a glass or plastic catheter (1.0-1.2 mm bore) through their genital pore and withdrawing a small sample of tissue. Ovaries of white bass and white perch can be biopsied with a fire-polished glass hematocrit tube (Berlinsky et al., 1995a; King et al., 1995a). The oocytes are viewed directly under a dissecting microscope, or they can first be chemically cleared to obtain more precise information (Figure 2.6). This procedure is routinely used to identify suitable candidates for hCG injection and spawning and to predict the time of ovulation (Rees and Harrell, 1990). To acquire definitive information on maturity, the biopsy samples can be subjected to routine

46

histological examination (Figure 2.5). Ovaries of adult females can be successfully biopsied throughout the year. Around the spawning season, maturity of males can be verified by simply manually palpitating the fish to express semen (Woods and Sullivan, 1993). Males can be biopsied outside the spawning season using a length of polyethylene tubing (PE 50) inserted through their genital pore. However, the sperm ducts degenerate or become occluded during Summer, and males begin spermiating within only a short time after the ducts reopen in Winter and the testis can be biopsied. Repeated gonadal biopsy suffers from the drawback that it can scar or damage the sensitive gonadal ducts, sometimes leading to blockages at spawning. It can also be difficult to maintain reasonably sterile conditions when biopsying large numbers of broodfish, so there is risk of infection and transfer of disease between individuals. The method is not applicable to male fish except during a few months around the spawning season. These shortcomings led to evaluation of circulating gonadal steroid hormone or VTG levels as markers of maturity (Berlinsky and Specker, 1991; Tao et al., 1993; Woods and Sullivan, 1993; Blythe et al., 1994c). Because of the clearly biphasic profiles of E2 and testosterone circulating in maturing female striped bass (see Figure 2.15), plasma levels of these steroids are not well correlated with oocyte growth and cannot be used to accurately gauge maturation of individual fish except around the spawning season. Conversely, plasma VTG levels are highly correlated with oocyte growth and have strong promise as a serological marker of female maturity (Tao et. al., 1993; B lythe et al., 1994c). Kishida et al. (1992) discovered that VTG is present on the body surface of striped bass in the scale mucus and went on to develop a sensitive ELISA for detecting VTG in mucus and blood plasma. Because mucus and plasma levels ofVTG covary, the technique appears applicable to female striped bass as a non-invasive test of maturity (Specker and Anderson, 1994). Regarding males, plasma levels of testosterone and 11-KT mainly increase around the time of spermiation and do not appear to be useful indicators of maturity at other times (Woods and Sullivan, 1993). Investigations of white bass and white perch generally confirm these conclusions (Berlinsky et al., 1995a; Jackson and Sullivan, 1995). Ultrasonic imaging of the gonads was recently evaluated as a means for detecting gender and maturity of captive striped bass broodstock (Blythe et al., 1994b,c). The gender of 5-year-old adult fish could be distinguished with 95% accuracy throughout the reproductive cycle. Cross-sectional measurements of ovary diameters on ultrasound scans were highly correlated with oocyte diameter and plasma VTG levels. Overall, ultrasonic imaging appears to be a simple, effective, and non-invasive method for verifying gender and assessing maturity of both females and males. Portable ultrasonic imaging equipment is widely available, although expensive, so the technique has potential for use by hatchery managers or fishery biologists. 2.3.3.3 Maturity Schedules Most studies of striped bass maturation have been directed at fishery management concerns and focused on the maturity schedule of wild fish. Numerous maturity schedules have been generated for females, and comparisons of the more comprehensive studies are provided in Table 2.5. Wild male striped bass generally mature within three years and have been the subject of less investigation. While most maturity schedules agree that all females in their seventh year of life (age-class seven) are mature, estimates of the incidence of maturity in age-classes three to five vary considerably. These discrepancies may be due to real differences between the populations sampled, changes in schedules within stocks over time, dissimilar criteria between studies for assessing maturity, or some combination of these variables. Most investigations of striped bass have relied on the presence of oocytes in early secondary growth or later stages of development as a marker of maturity. As noted, ovaries ofjuvenile fish usually contain only primary growth oocytes, characterized in the early literature as "smaller diameter" or "Type I" oocytes.

47

Because striped bass have group synchronous oocyte development, ovaries of adult fish contain predominantly early secondary growth, vitellogenic, final maturation, or atretic oocytes, in addition to the omnipresent primary growth oocytes. Earlier studies refer to growing oocytes by diameter or by the designations "Type II" and "Type III," which correspond to early secondary growth and vitellogenic oocytes, respectively. These are derived from primary growth oocytes and proceed to maturity and ovulation through the developmental sequence previously described. The term "secondary growth" refers to the rapid increase in diameter, mainly resulting from uptake of VTG and/or lipid, seen in early secondary growth and vitellogenic oocytes (see sections 2.2.1.1 and 2.3.1.1.1). Both types of oocytes are in secondary growth. Using secondary growth oocytes as the criterion for maturity suffers from the assumption that initiation of secondary growth is always followed by spawning within the same year. Small striped bass whose most mature oocytes were in early secondary growth have been sampled during Winter when larger fish were in mid-vitellogenesis (Berlinsky, 1989). Large, presumably adult, striped bass whose biggest oocytes (---250/am diameter) were just initiating vitellogenic growth have been sampled in or around the spawning season (Berlinsky et al., 1995b). As vitellogenesis requires several months to complete, it is unlikely that oocytes from these fish could have developed enough for spawning in the year they were sampled. On the basis of these and similar observations, it has been suggested that some striped bass may require more than one year from initiation of secondary oocyte growth until spawning (Merriman, 1941; Chadwick, 1965; Specker et al., 1987). Maturity schedules based on the presence of secondary growth oocytes during the prespawning and spawning season (Olsen and Rulfison, 1992) likely overestimate the number of fish capable of spawning. Estimating the spawning potential of virgin striped bass is further complicated because these fish may reabsorb rather than ovulate secondary growth oocytes and produce eggs of high quality inconsistently. The reproductive output of young fish would be overestimated if the frequency of this pre-ovulatory atresia is high and/or viability of the eggs spawned is low. Widespread pre-ovulatory atresia has been noted in other teleosts and was shown to be associated with fish spawning for their first time (Tam et al., 1986; Bromage and Cumaranatunga, 1987). Because teleosts can reabsorb atretic oocytes quickly, it may be difficult to determine the frequency of pre-ovulatory atresia or its influence on the accuracy of striped bass maturity schedules. Some small striped bass (fork length = 480 mm) sampled from the Roanoke River, North Carolina, were found to have ovaries containing a few atretic or fully-grown vitellogenic oocytes intermixed with predominantly primary growth oocytes. Tank spawning trials were conducted with several of these fish and most did not produce fertile eggs, although large females spawned successfully (see Table 2.6). In one case, only three fertile eggs were recovered from a few dozen in the tank after such a small female spawned (E. Atstupenas, Edenton National Fish Hatchery, Edenton, NC, and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). What possible significance can this kind of reproductive output have for the population under study? Compared to older fish, younger fish may also produce small eggs and larvae which are less likely to survive (Knutsen and Yilseth, 1985; Hislop, 1988). It appears that for striped bass, a correlation between female weight and egg size or larval survival can be detected where the range in size of mature females is large (e.g. Chesapeake Bay) but not where it is small (e.g. Santee-Cooper reservoirs) (Zastrow et al., 1989; Monteleone and Houde, 1990; Secor et al., 1992). Differences between striped bass maturity schedules probably also reflect differences between populations, sampling distance from the spawning grounds, and seasonal effects on sampling (Table 2.5). Striped bass migratory patterns are known to be complex and influenced by population and season, as well as fish age, gender, and size (Kohlenstein, 1981; McLauren et al., 1981; Boreman and Lewis, 1987; Waldman et al., 1990; Dorazio et al., 1994). McLauren et al. (1981) found more than double the frequency of mature females in age-class 6 during the prespawning and spawning season at one location on the Hudson River as

48

Table 2. 5. Comparison of some maturity schedules for striped bass. The percentage of fish reaching maturity in each age-class is shown.

Investigator(s)

3

Age-Class 5 6

4

7

Location

Berlinsky et al. (1995b) a

0

12

34

77

100

Coastal Northeast

Olsen & Rulifson (1992) b

44

93

95

100

100

Roanoke River Albemarle Sound

Harris et al. (1985) r

18

60

85

92

100

Roanoke River Albemarle Sound

Harris et al. (1985) d

17

54

83

100

100

Roanoke River Albemarle Sound

Lewis (1962) e

4

92

100

100

100

Roanoke River Albemarle Sound

Wilson et al. (1975) f

9

78

99

100

100

Potomac River

McLauren et al. (1981 )g

0

0

50

89

90

Hudson River

McLauren et al. (1981)h

2

6

17

41

87

Hudson River

Merriman (1941)~

0

27

74

93

100

Coastal Northeast

aMay-Jun, Oct-Nov 1985-87; bFeb-Jun 1989-90; 'Nov-May 1980-81; aNov-May 1982-83; eOct-Apr 1956-1958; fFeb-May 1975; gMar-Jun, spawning grounds 1976-77; hMar-Jun, below spawning grounds 1976-77; iApr-Nov 1937-1937.

compared to another location further removed from the spawning grounds (Table 2.5). The probability of sampling immature fish increases the further distant fish are sampled from spawning areas because immature fish are less likely than mature fish to migrate upriver during the spawning season. Maturity schedules based on mixed-stock coastal migrants (Merriman, 1941; Berlinsky et al., 1995b) give more conservative estimates of the frequency of maturity in younger age-classes as compared to schedules based on samples from spawning areas. Clearly there are serious complications in developing striped bass maturity schedules arising from methods for assessing maturity, the potential for preovulatory atresia, the dubious reproductive contributions of young or virgin females, and problems with developing sampling programs unbiased with respect to time and location. Fishery management decisions should be based on the most conservative schedule, one that defines the first age of reproduction to be that for which 100% of females can be confidently considered to be mature. Based on a consensus summary of the data in Table 2.6, this would correspond to age-class seven. Information on maturation of captive striped bass is more limited. Our most extensive knowledge of such fish comes from studies done over the last 16 years at the University of Maryland Crane Aquaculture Facility (Woods et al., 1990, 1992; Woods and Sullivan, 1993; L.C. Woods, III, Crane Aquaculture Facility, University of Maryland, personal communication). During the course of producing three generations of domestic broodstock, fish reared from larvae to mature adults in captivity, it has become apparent that captive

49

striped bass can mature at younger ages than their wild counterparts. Numerous females of the F 1 generation have been spawned at the facility and, during ovarian biopsy associated with spawning, it was discovered that ---25% of four-year-olds, --75% of five-year-olds, and all six-year-olds were mature. Comparable data on domestic broodstock produced at the South Carolina Marine Resources Research Institute indicated no females matured by age two, 16% were mature by age three, and 59% were mature by age four (Smith and Jenkins, 1988). These observations are in general agreement with studies performed at the North Carolina State University Pamlico Aquaculture Field Laboratory, involving rearing and spawning F~ generation broodstock (Hodson and Sullivan, 1993). Most females matured at four to five years of age. There is also evidence that the process of domestication can lead to earlier maturation. Recently, several F2 generation females were spawned at the Crane Aquaculture Facility as three-year-olds to produce an Fs generation. These females from the 1989 year class were spawned in the spring of 1992, just before their third birthday, and so must have become vitellogenic as two-year-olds. Subsequent biopsy of scores of females from this year class during springtime spawning activities in 1993 revealed all to be mature. At all ofthe facilities mentioned, some males mature in captivity during their second year and all are mature by age three. 2.4 CAPTIVE BREEDING

2.4.1 Broodstock Acquisition and Conditioning This section reviews current information on establishing striped bass broodstocks, maintaining them in captivity, and conditioning the fish for reproduction. Corresponding information on treatment of wild fish captured annually for spawning was reviewed by Harrell et al. (1990a). 2.4.1.1 Broodstock Sources Broodstock can be developed using fish captured from the wild or by rearing larvae or fingerlings to adulthood in captivity. We refer here to wild broodfish acclimated to captivity as "captive" broodstock and larvae or fingerlings reared to adulthood in captivity as "domestic" broodstock. The terms are somewhat arbitrary, but useful. It should be recognized that development of truly domesticated striped bass selected for desired pertbrmance characteristics will require that several filial generations of domestic broodstock be produced through selective breeding over many years (see Chapter 8). 2.4.1.1.1 Captive broodstock The fastest way to develop a broodstock is to adapt wild striped bass to captivity and prepared diets. The fish can be captured by hook and line or other methods that are not harmful, and transported to the laboratory or hatchery (Harrell, 1984; see Chapter 7). However, it is well known that wild fish of several species exhibit an immediate and complete "shut-down" of the reproductive system when brought into captivity. Recent evidence indicates that this response involves disruption or cessation of production of virtually every pituitary hormone involved in reproduction (Sumpter et al., 1994). Striped bass may be a classic example ofthis phenomenon. Indeed, the routine procedure of injecting wild fish with hCG to maintain spermiation or induce ovulation can be viewed as a kind of"replacement therapy." It likely serves mainly to offset loss of endogenous GTH secretion resulting from the stress of routine hatchery operations associated with spawning. Other responses of wild striped bass to handling and confinement may include elevated plasma cortisol levels, reddening of the ventrum and fins commonly referred to as "red-tail," osmoregulatory dysfunction, and failure to ovulate after hCG injection (Rees and Harrell, 1990; Harrell, 1992; Harrell and Moline, 1992; Harms et al., 1996). At present, the only recourse researchers or hatchery managers have to mitigate stress in striped bass is to minimize handling of broodfish and add anesthetics or salt to the water

50

when fish are handled (Rees and Harrell, 1990; Chapters 7 and 10). Salting the water offsets dilution of the blood plasma and ionic imbalances resulting from stress-induced hyperventilation and increased functional surface area of the gills. Recent studies suggest that, for mitigating stress associated with capture and transport of wild striped bass, use of salt (10 g/L) alone may be preferable to use of anesthetic (tricaine methane sulfonate, 25 moJL) either alone or in combination with salt (Harrell, 1992; Chapter 7). Many experiments on striped bass and white bass reproduction conducted at the North Carolina State University Pamlico Aquaculture Field Laboratory utilized wild fish acclimated to captivity (Hodson and Sullivan, 1993; Tao et al., 1993, 1996; King et al., 1994a, b, 1995b; Berlinsky et al., 1995a,c). A brief summary of the approach used to establish the striped bass broodstock follows (R.G. Hodson and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). Striped bass are captured on their spawning migration in commercial pound nets near the mouth of the Roanoke River, transported to the hatchery, and adapted to captivity and pelleted commercial feeds. Capture is easier during the spawning migration and the gender of the fish can be readily identified then so the needed sex ratio can be obtained. Young fish less than five kilograms in weight are preferred because older and larger fish usually do not adapt well to captivity. Ideally, small males that are spermiating (age class 2-3) and the youngest females with secondary growth oocytes (age class 3-4) are obtained. Spawning females in the season of capture, especially by manually stripping them of eggs for in vitro fertilization, is not advisable. Approximately half of spawned females die during acclimation to captivity, presumably as a result of injury when their eggs are taken, stress associated with handling, and infections. It seems advantageous to allow females to reabsorb their fully grown oocytes as a bioenergetic reserve to carry them through adaptation to captivity until they resume feeding. Using this approach, up to 90% survival has been obtained with several different groups of females. Survival of males is similar whether or not they are spawned after capture, and it is unusual for them not to adapt well to captivity. The males will spermiate during the next spawning season and be available as broodstock. However, females generally require two years before they will produce good quality eggs. Striped bass brought in from the wild are held in large circular tanks supplied with adequate amounts of brackish surface water (5 to 12 ppt) or fresh well water (Hodson and Sullivan, 1993). Brackish water is used to help the fish recover from capture stress and retard development of fungal infections. Strict temperature control is not necessary during acclimation but is required to successfully spawn striped bass. The fish can be trained to accept pelleted feeds within four to six weeks. Within a few days of capture, pieces of fresh fish are offered three times weekly. If the fish do not begin to feed within a couple of weeks, then live food is provided to stimulate feeding activity. Wild gizzard shad or various tilapia species produced at the facility are used as forage. Once fish begin to feed aggressively, they are fed only twice weekly to ensure they are hungry when prepared food is presented. Each time the fish are fed, a pelleted floating feed (--12 mm diameter) is presented first. If the pellets are not eaten within 10-15 minutes, then the broodstock are fed a ration of chopped fish. Floating pellets are used to facilitate observation of feeding behavior. Once a few fish in a tank are accepting pelleted feed, it usually takes only a few days before all the fish are accepting it. When a group of fish is having difficulty converting to prepared feed, actively feeding fish adapted to captivity in prior years are stocked into the same tank to stimulate feeding behavior. Within one week wild fish are usually accepting chopped fish. By the end of three or four weeks they begin to accept some pelleted feed, and after six weeks they are usually completely converted to pelleted feed. The general methods described above can also be used to establish captive white bass broodstocks (see Chapter 6).

51

2.4.1.1.2 Domestic broodstock The other way to develop broodstock is to rear larvae or fingerlings in captivity until they are of reproductive age. Various methods for producing domestic striped bass broodstocks have been reviewed (Smith and Jenkins, 1985, 1987, 1988; Woods et al., 1990, 1992). Similar information is available for white bass (Kohler et al., 1994; Smith et al., 1996; see also Chapter 6). For striped bass, it will generally require two to three years for males and four to five years for females before some of the domestic fish can be spawned. Developing domestic broodstock is a major undertaking involving considerable expense and a long-term commitment of personnel and resources. Furthermore, not all of the environmental or nutritional requirements of adult striped bass are known (see Chapter 9), nor has the methodology for inducing spawning of such fish been perfected. When starting a broodstock program from larvae or fingerlings one also needs to take care not to create a genetic "bottleneck" by using fish that are all the progeny of one or a very few females (Kerby and Harrell, 1990). Similar care must be taken in the production of subsequent generations of broodstock, which means that multiple tanks or rearing systems will be required. Unless the grower has adequate space and resources to make this kind of commitment, it is probably best to begin with a captive broodstock based on wild fish. Acquiring gravid captive or domestic stock from another facility may not be advisable because even domestic broodfish do not adapt well to new environments if they are more than four or five years of age. Various intensive culture methods can be used to establish and propagate domestic white bass broodstock (see Chapter 6). White bass broodstock can also be created within two years by stocking larvae produced from wild parents into ponds and producing fingerlings using routine methods for spawning, pond fertilization, and phase-I fingerling production (Harrell et al., 1990a; Chapter 3). These fingerlings will learn to accept pelleted feed and can be reared in ponds in just the same manner as hybrid striped bass. Within two years, females will produce viable eggs and the males will be producing sperm (R.G. Hodson and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). It should be possible to spawn these fish two or three times (years) before they have to be replaced. Most commercial striped bass culture is based on growout of hybrids (see Chapters 1 and 12) because the hybrid is widely viewed as a superior cultivar relative to either parent. Hybrids themselves are not suitable as broodstock, because the F2-generation is too variable with respect to growth rate, body conformation, body coloration, and other production characteristics (see Chapter 8). Unless the producer is prepared to maintain and propagate domestic broodstocks of both parental lines, the simplest solution may be to use captive striped bass males crossed with captive or domestic white bass females that are replaced every few years. Such a production system can be in place within two years at only modest cost compared to other options. 2.4.1.2 Conditionin~ broodstock Broodstock conditioning is probably the single most important aspect of captive breeding. Conditioning principally involves providing the fish with environmental conditions and a diet that optimize reproductive performance. Although we have learned a great deal about the reproductive biology of striped bass and are developing improved methods for hormonal induction of ovulation and spawning (see section 2.4.2), we still know very little about what constitutes optimal reproductive conditioning for this species. It is certain that the annual photothermal cycle regulates the timing of reproductive processes, but we still have not identified the limits of daylength or water temperature, or the timing in changes thereof, that maximize reproductive efficiency. Grossly out-of-phase temperature and photoperiod cycles can lead to problems with ovulation. For example, striped bass held under a natural annual cycle of photoperiod but at a constant spring water

52

temperature of---18~ routinely completed oocyte maturation but failed to ovulate (R.G. Hodson and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data), even after combined treatment with GnRHa implants and injected hCG (Hodson and Sullivan, 1993). A clear increase in water temperature is a signal for final maturation and ovulation in some fishes (Goetz, 1983), and in this case such a signal would be lacking. Although natural induction of striped bass ovulation and spawning probably involves an abrupt increase in springtime water temperature, very high temperature (> 25 ~ interfere with ovulation. Failure of striped bass to ovulate at temperatures grossly inappropriate for embryo and larval survival is a well known phenomenon. There is good evidence from studies of wild fish that extreme summer water temperature (>30 ~ can stress adult females, alter the composition of their oocytes, and lead to failures of recruitment (Grimes, 1993; Coutant, 1987a, b, 1990a, b). Some recent observations suggest that high water temperature during Winter can also lead to reproductive problems (L.C. Woods III, Crane Aquaculture Facility, University of Maryland, personal communication). In the spring of 1995, several cohorts of larvae exhibited precocious absorption of their yolk sacs to the point where they were completely absent before the larvae had functional mouth parts and could commence exogenous feeding. Egg size and appearance were otherwise normal. The domestic broodstock involved are held under ambient Chesapeake Bay conditions and it is noteworthy that the prior winter was one of the mildest on record for this region. Like high summer water temperature, excessive water temperature in Winter has the potential to disrupt normal vitellogenesis and yolk formation. As noted, annual photothermal cycles less than nine months in periodicity seem inadvisable because they may not allow sufficient time for oocyte growth (see section 2.3.2 and Figure 2.15). Collectively, the observations made above indicate that broodstock should be exposed to an annual photothermal cycle (> 9 months) that includes low water temperature around winter solstice (< 10~ Woods and Sullivan, 1993; Blythe et al., 1993a-c), a temperature appropriate for larval survival around spawning time (--16~ enough scope in springtime (several ~ for an abrupt temperature increase to stimulate final maturation without blocking ovulation, and only moderately high water temperature (<< 30~ in summer. So far, there is no evidence that annual changes in salinity are required for successful reproduction. The phenomenal success of some landlocked striped bass populations (e.g., Santee Cooper Reservoir, South Carolina) suggests salinity changes are not required. The nutritional requirements of striped bass are not complete, and much research remains to be done in this area (see Chapter 9). Studies ofbroodstock nutrition for this species have been few and limited in scope (Gallagher, 1987). The nutritional status of broodstock is not strictly a function of diet. It is also influenced by water temperature and possibly other environmental factors that impact bioenergetics in maturing females. The environmental conditions of captivity and the content ofbroodstock diets likely interact to determine the composition and quality of eggs produced. Harrell and Woods (1995) compared the fatty acid composition of eggs from domestic and wild striped bass. There were distinct differences in lipid composition between the eggs of wild fish captured from Chesapeake Bay and the eggs of domestic fish at the Crane Aquaculture Facility. Wild fish had significantly higher levels of total lipid, n-3 highly unsaturated fatty acids (n-3 HUFA), eicosapentanoic acid 20:5n-3 (EPA), and docosahexanoic acid 22:6n-3 (DHA). Striped bass cannot synthesize these essential long-chain n-3 fatty acids d e n o v o from shorter-chain fatty acids (Tuncer and Harrell, 1992). They must be obtained in the diet and appear especially important to developing larvae (Tuncer et al., 1993). Although levels of these n-3 fatty acids were reduced in domestic females as compared to wild fish, they were still much higher than reported minima needed for larval growth and survival (Watanabe, 1982). The fatty acid composition (n-3/n-6 fatty acid ratio) of eggs from wild fish reflected what would be expected of a marine fish, whereas the composition of eggs from domestic females appeared typical of freshwater fish. This is consistent with the origins of the females involved and with their respective diets.

53

Most striped bass broodstocks are fed commercial diets designed for salmon, containing considerable quantities of corn and soy oils, which do not have a lipid profile typical of marine fish. It remains to be verified whether or not the lower levels of n-3 fatty acids in these diets, or in the eggs of striped bass broodstock fed the diets, impairs egg quality or larval performance. Similar differences in diet composition have been shown to influence reproduction of European sea bass (Carrillo et al., 1995), a close relative of striped bass. Sea bass fed a commercial trout diet grew at a slower rate and exhibited significant decreases in relative fecundity and the incidence and duration of spawning relative to sea bass fed a control diet of chopped baitfish. The decrease in fecundity was associated with an increased incidence of pre-ovulatory atresia. Eggs of fish fed the commercial diet were also smaller and produced lower rates of hatching and larval survival. Spawning was also delayed. For sea bass fed chopped fish at either full ration or half ration, eggs size and fecundity were decreased by reduced ration. Spawning was delayed and the frequency of spawning was also decreased by the half ration. These findings demonstrate that ration and diet composition may have a strong impact on reproductive processes in striped bass and related species. This is an area of striped bass broodstock conditioning that begs for further research. 2.4.2 Induction of Spawning Originally, only female striped bass that were ovulated at the time of capture could be spawned successfully. The "ripe roefish" were purchased from fishermen during their annual spawning run in the Roanoke River, North Carolina, and immediately stripped of their gametes for in vitro fertilization and incubation of the eggs (Worth, 1884). In the early 1960s, techniques were developed for inducing spawning of less mature females (Stevens et al., 1965; Stevens, 1966, 1967). Wild broodfish were collected during the spawning run and induced to ovulate with injected hCG. There was no significant advance in development of methods for reproducing wild striped bass for over 20 years, and hCG injections are still the principal means for inducing ovulation and spawning (Rees and Harrell, 1990; Smith and Whitehurst, 1990). In general, the hCG injection method is only applicable to wild females captured in late stages of ovarian maturation. Females are selected that have oocytes showing some degree of lipid droplet coalescence ("eligible" or_<15 hr stage oocytes; Rees and Harrell, 1990). We now know that such oocytes have a migrating germinal vesicle and have initiated final maturation (Figure 2.6). Females with such mature oocytes probably already have elevated blood levels of maturational GTH-II (see section 2.3.1.1.2). Such fish are seldom available to commercial growers who are legally restricted from using the most efficient gear (e.g., electrofishing) to capture fish on or near their spawning grounds. Females captured further downstream are less mature and typically have not yet initiated final oocyte maturation. Attempts to use hCG injections for spawning the less mature females usually fail. The fish either do not ovulate completely, ovulate underripe and infertile eggs, or yield eggs that give rise to embryos which exhibit asymmetrical cleavage during early development and later die (E. Atstupenas, Edenton National Fish Hatchery, North Carolina, and R.G. Hodson and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). Occasionally such females can be spawned successfully if they are captured near the end of the natural spawning season. These "late" fish are probably maturing very quickly as they rapidly migrate upstream to spawn, having already been exposed to elevated titers of endogenous GTH-II. With these exceptions, hCG injections are usually not useful for ovulating the less mature wild females. Because only wild fish captured at an advanced stage of maturation can be routinely induced to spawn successfully with hCG, many must be released unspawned, which is wasteful and inefficient. Domestic striped bass, like cultured females of several other teleost species, produce fully grown oocytes and then rapidly undergo atresia unless they are pharmacologically induced to complete oocyte maturation and spawn (Zohar, 1989). With some exceptions (Woods et al., 1992; Blythe et al., 1993a-c), they

54

do not spawn successfully unless their oocytes have initiated final maturation by the time they are injected with hCG. Locating females mature enough for hCG injection requires repetitive handling of the fish during the spawning season, which stresses the broodstock and impairs reproductive performance and survival. By the time a sufficient number of candidates for hCG injection are found most other females in the stock will already have initiated atresia and be unspawnable. Therefore, the hCG injection technique is also inadequate for propagating captive broodstocks. Based on their effectiveness in other teleosts, researchers evaluated injections of GNRHa or a dopamine antagonist for inducing maturation and ovulation of striped bass (see section 2.2.2.1). A potent synthetic analogue ofsGnRH (sGnRHa; [D-Argr-Pro9-NEt]-sGnRH]) or the dopamine antagonist, DOM, were used singly or in combination (King et al., 1994a). Synthetic analogues of GnRH, injected alone or with a "priming" dose of carp pituitary extract (CPE), had been used before to induce ovulation or spermiation in scores of fish species that could not be spawned successfully using first generation techniques such as simple hypophysation or hCG injection (Donaldson and Hunter, 1983). Dopamine antagonists had been used in conjunction with GnRHa to successfully ovulate several species of Chinese carp that proved "resistant" to hCG, pituitary extracts, or GnRHa alone (Lin et al., 1986). However, results with striped bass showed that, although injected sGnRHa could induce final maturation and ovulation, DOM did not appear to facilitate the response to sGnRHa or be effective when injected alone. Furthermore, the time between hormone injection and ovulation was up to 10 hours longer for the sGnRHa preparations as opposed to hCG (King et al., 1994a), a highly undesirable response since wild fish must be spawned as soon as possible before they succumb to stress and die or cease maturation. The delayed response to sGnRHa was not entirely unanticipated, because it must first act to elicit release of endogenous GTH from the fish's own pituitary gland, as opposed to direct action on the ovary of injected hCG. On this basis, injecting solutions of GnRHa or dopamine antagonists is not recommended as a technique for inducing ovulation and spawning of wild striped bass. In the late 1980s a technique for chronic administration of GnRHa from implanted pellets became available and showed strong promise as a means for inducing maturation in several species of teleosts (Sherwood et al., 1988; Crim et al., 1988; Almendras et al., 1988). The method involves incorporation of GnRHa into pellets made of cholesterol (CH) and cellulose (CL). The CH/CL ratio can be changed to regulate the rate and duration of hormone release from the pellets, for periods lasting from days to weeks. A synthetic analogue of mammalian GnRH (mGnRHa; [D-Alar-Prog-NEt]-LHRH) was incorporated into such pellets (-20 ~zg mGnRHa/Kg fish body weight) containing either 80% cholesterol (80; fast-release, days) or 95% cholesterol (95; slow-release, weeks) that were modified so as to be injectable through a 10 gauge syringe needle (Hodson and Sullivan, 1993; Woods and Sullivan, 1993). The mGnRHa was chosen over sGnRHa because it is relatively inexpensive and widely available. The implants were injected subdermally into the dorsal lymphatic sinus of striped bass, a few scale rows down from the posterior insertion of the second (caudal) dorsal fin. The mGnRHa implants were highly effective for inducing maturation and spawning of striped bass too immature for hCG injection. In initial trials involving manual spawning and in vitro fertilization to produce hybrid striped bass, paired implants (80/95) administered to female striped bass were found to induce ovulation (Figure 2.16A). In later trials, females were allowed to mature after implantation with mGnRHa pellets until they initiated final oocyte maturation, at which time they were given a "resolving" injection with hCG (330 IU/Kg body weight) to hasten ovulation (Figure 2.16B). This method also synchronizes spawning of implanted broodfish. Synchronization of spawning is highly desirable if extensive production of fingerlings is anticipated, because resulting fry will all be nearly the same age when stocked (Brewer and Rees, 1990; see Chapter 3).

55

Spawn

A

.

5

10-

O

Fish 1 Fish 2

O

Fish 3 >15

' 4'0 ' 6'0 ' 8'0 ' I 0"0

' 2b

Hours

Spawn

B

Inject 330 IU/Kg / hCG

Implant mGnRHa

~ ~

~/

/

//

O

O

12

Fish 2 >15 i

0

i

!

8

!

i

!

16

i

24

i

i

32

i

i

40

!

i

48

Hours Fig 2.16. Maturation and spawning of wild female striped bass using mGnRHa implants alone or implants followed by a resolving injection with hCG. A. Females (2.0-3.4 Kg body weight) whose ovaries contained fully grown oocytes that had not yet initiated final maturation (> 15 hr stage; Rees and Harrell, 1990) were implanted with an 80% cholesterol (fast-release) and a 95% cholesterol (slow-release) pellet, each containing 100/xg of mGnRHa, the remainder of the pellet matrix being cellulose. The fish were subjected to ovarian biopsy at irregular intervals to follow the course of maturation. The oocyte stages and time of spawning are shown. Refer to text (2.2.1.1) for a description of the oocyte stages. Females with ovulated (0 hr stage) eggs were manually spawned and their eggs were fertilized with milt from several white bass males to produce hybrid striped bass. The percentages of eo,~s~,from females 1, 2, and 3 that produced viable 1-day-old fry were 45.5%, 15.3% and 6.1%, respectively. B. Two females (4.1-4.4 Kg body weight) were given dual mGnRHa implants as in A, but were subsequently given a resolving injection with hCG (330 I.U./Kg body weight) once their oocytes had initiated final maturation (oocyte stage > 14 hr). The oocyte stages, time of injection with hCG, and time of spawning (ovulation) is shown. Biopsy, spawning and fertilization of the eggs was done as in A. The percentage of eggs from fish 1 and 2 that produced viable 5-day-old fry were 26.4% and 13.7%, respectively (data from Hodson and Sullivan, 1993).

56

Using the mGnRHa implant technique, the less mature wild striped bass have been repeatedly spawned with egg fertility and embryo hatching rates comparable to mature females undergoing final oocyte maturation at the time ofhCG injection (Hodson and Sullivan, 1993). The technique has been used since 1992 at the Edenton National Fish Hatchery (North Carolina) to induce volitional spawning in tanks by scores of the less mature wild females. A single 95% cholesterol implant (20 gg mGnRHa~g body weight) can substitute for the paired implants described above, and injection of mGnRHa (20 gg m G n R H a ~ g body weight in 0.7% NaCI) or implantation with either a whole or half 80% cholesterol implant (10 #g mGnRHa/Kg body weight) can substitute for the resolving hCG injection. A representative example of results from spawning trials conducted at the Edenton National Fish Hatchery during Spring of 1991-92 is shown in Table 2.6. Domestic broodstocks are also routinely spawned using the mGnRHa implants, including those at the North Carolina State University Pamlico Aquaculture Field Laboratory, the University of Maryland Crane Aquaculture Facility, and the South Carolina Marine Resources Research Institute (Hodson and Sullivan, 1993; Woods and Sullivan, 1993; C.V. Sullivan, Department of Zoology, North Carolina State University, and T.I.J. Smith, South Carolina Marine Resources Research Institute, unpublished data). An advantage of using the mGnRHa Table 2.6. Results of spawning trials using mGnRHa implants at the Edenton National Fish Hatchery (North Carolina) in 1991-92. Wild female striped bass (3.5 - 7.0 Kg body weight) were captured from commercial pound nets near the mouth of the Roanoke River, about 100 Km from the spawning grounds. All had full grown oocytes that had not yet initiated final maturation. They were given 95% cholesterol (95; slow-release) or 80% cholesterol (85; fast-release) implants containing 100~zgmGnRHa, the remainder of the implant matrix being cellulose. After implantation, females were stocked into circular tanks with 2-3 spermiating males for spawning (Smith and Whitehurst, 1990). They were anesthetized (quinaldine sulfate, 50 mg/L) and biopsied at irregular intervals to evaluate the progress of oocyte maturation. Females whose most advance oocytes had initiated final maturation (oocyte stage < 14 hr; Rees and Harrell, 1990) were given a resolving injection ofhCG (330 IU/Kg body weight) or a single 80 implant. The time of spawning, number of eggs spawned and their percentage viability (live embryo after 24 hr of incubation), and the number of resulting 3-day-old fry is shown (data from a previously unpublished experiment by E. Atstupenas, Edenton National Fish Hatchery, North Carolina, and C.V. Sullivan, Department of Zoology, North Carolina State University). mGnRHa Implant Type (Day 0) 95

Secondary Injection (Day, Type) 6, hCG

Spawn (Day)

Eggs (#)

Viable (%)

Fry (#)

7

538,000

55.0

142,500

95

6, hCG

8

408,000

75.0

261,500

95

6, hCG

8

954,800

25.0

57,700

95

3, 80

7

769,000

83.0

345,300

80/95

None

2

546,508

70.0

390,000

80/95

5, hCG

6

616,143

76.5

396,200

Mean ~

6

638,742

64.1

265,530

(0.8)

(79,429)

(8.7)

(56, 945)

(SEM)

57

implants with domestic broodstock is that females can be induced to mature earlier in the spawning season, and before they become atretic. In nature, white bass typically spawn earlier than striped bass in most locales. Earlier spawning of striped bass also provides for synchronization of striped bass females with white bass males for production of hybrids. The mGnRHa implants described above are simple and inexpensive to manufacture, as well as being very reliable for inducing maturation and spawning of striped bass. However, hormone dosages and release rates for this kind of implant have not yet been optimized, nor have other types of sustained hormone-delivery systems been fully explored for spawning striped bass and related species. Especially promising are the new polymer- based hormone or drug delivery systems recently developed for biomedical applications. These have been demonstrated to be highly effective for inducing maturation and ovulation or spawning of sea bream and several other teleosts (Zohar et al., 1995b). Recently, Mylonas et al. (1995) utilized for spawning striped bass a delivery system for GnRHa based on a biodegradable copolymer of a fatty acid dimer and sebacic acid (p[FAD-SA]) formed into microspheres that can be injected using a conventional syringe. In an experiment conducted just before the spawning season when water temperature was still suboptimal (< 16 ~ injection of domestic female striped bass that had fully grown oocytes with GnRHa-loaded p[FAD-SA] microspheres induced 60% of the fish to ovulate over the course of 20 days, whereas no controls did (Figure 2.17A). In a later experiment performed at more suitable spawning temperatures (17-20~ the microspheres induced 100% of the females to ovulate within 11 days as compared to only 65% of females given two bolus injections of GnRHa solution spaced four days apart (Figure 2.17B). The faster and more complete response to the GnRHa-loaded p[FAD-SA] microspheres in the second experiment could be due to the greater quantity of hormone administered (150 #g versus 50/.zg GnRHaJKg body weight) or the higher prevailing water temperature during the second experiment. As noted, a surge in water temperature likely triggers natural final maturation and spawning of striped bass, and this could potentially involve increased responsiveness to endogenous or exogenous GnRH. It is interesting that, in the first experiment, administration of the microspheres could sustain continuous maturation of female striped bass for over 20 days. Development of GnRHa systems and protocols that can prevent onset of atresia and ensure complete maturation of females will contribute greatly to management of domestic striped bass broodstocks. The GnRHa microspheres were also found to be effective for stimulating sperm production of male striped bass (Figure 2.18). The mean total expressible sperm volume was significantly elevated in males injected with the microspheres for up to two weeks relative to control males. Males treated with the microspheres produced four times as much sperm as controls two days after treatment and 10 times as much sperm as controls 14 days after treatment. Not only was sperm volume enhanced, but the spermatozoid count (number of spermatozoa/mL of sperm) was also enhanced by injection with the GnRHa microspheres (Mylonas et al., 1995). These kinds of results underscore the need to fully evaluate the new polymer-based long-term GnRHa delivery systems in striped bass culture. It should be noted that no method for pharmacological induction of maturation will be effective when applied to severely stressed or improperly conditioned broodstock, females that have initiated atresia, or fish too young for spawning. Such fish regularly exhibit abnormal ovulation syndromes after hormone treatment. These include: 1) failure to ovulate (all oocytes retained), 2) incomplete ovulation (some mature oocytes retained), 3) delay of ovulation until the oocytes are overripe and infertile, and 4) failure to spawn shortly after ovulation with subsequent spawning of overripe eggs (R.G. Hodson and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). Failure to ovulate is commonly seen in severely stressed wild females exhibiting the "red tail" syndrome, sometimes accompanied by osmoregulatory problems

58

GnRHa-microspheres 100

~

GnRHainjections

80

=

60

o ~

40

=

20

Vehicle

f

-

25

-

A. = o

---O-

r

s S

B. 20

,,ml

E I= ~

/ ///

"-. 10

.d

5

0

I

A

iliiIi

0

....

5

| ....

| ....

| ....

| ....

| ....

, ....

I

10 15 20 25 30 35 40

|

,

'

0

|

2

I

4

r~ I~ '

,

6

!

8

b~

I

|

10

i

,

12

i

14

Time after t r e a t m e n t (days)

,

,

16

,,,

Fig. 2.17. Cumulative percentage ovulation of domestic striped bass over time in response to injection with GnRHaloaded p[FAD-SA] microspheres (30 gg GnRHa/mg microspheres). A. Females (N=6) were kept under ambient temperature and given 1.6 mg microspheres/Kg body w e i ~ t (--50 ~zg GnRHa~g body weight). B. Fish (N=8) were kept under controlled temperature and given 5 mg of microspheres/Kg body weight (~ 150 #g GnRHa/Kg body weight), or two injections of GnRHa dissolved in saline (20 gg GnRHa/Kg body weight) on clays 5 and 9 (arrows) (data from Mylonas et al., 1995). 20 -

[7] Vehicle

18 GnRHa-microspheres 16 -

E E o

14 12 10 8-

g-

:6

6-

2 0 0

2

4

7

14

Time after treatment (days) Fig. 2.18. Mean total volume of sperm (+ SEM) expressible from domestic striped bass for up to 14 days after treatment with GnRHa-loaded p[FAD-SA] microspheres (30 #g GnRHa/mg microspheres). Fish (N=8 per group) were held at 18 ~ and injected with either 5 mg of microspheres (shaded bar; GnRHa-microspheres) or with vehicle only (hatched bar; Vehicle). The asterisk (*) indicates significant differences between treatments on a sampling day (P < 0.01; ANOVA) (data from Mylonas et al., 1995).

59

and elevated plasma cortisol levels (Harms et al., 1996). Partial or incomplete ovulation is often observed in captive brood stock exhibiting some degree of follicular atresia in biopsy samples taken at the time of hormone injection. Anatomical structures and biosynthetic capacities of their follicles necessary for ovulation are likely compromised. Sometimes very small broodfish exhibit abnormally low weight-specific fecundity, failing to ovulate completely after hormone treatment, a phenomenon we have termed "virgin syndrome" (E. Atstupenas, Edenton National Fish Hatchery, Edenton, North Carolina, and C.V. Sullivan, Department of Zoology, North Carolina State University, unpublished data). Hatchery managers routinely avoid using the smallest females because of this phenomenon. Female striped bass may need to undergo more than one reproductive cycle before becoming competent to reproduce, and do not spawn but instead reabsorb mature oocytes at the end of their first one or two cycles (see section 2.3.3.3). As noted, inappropriate photothermal conditioning cycles or excessive water temperature around the time of spawning can also interfere with ovulation (see section 2.4.1.2). In addition, virgin or atretic females and fish subjected to excessive stress or abnormal conditioning often produce progeny exhibiting abnormal development or high mortality during the early life stages. 2.5 ACKNOWLEDGMENTS The authors are grateful to several individuals who contributed to this chapter. E. Atstupenas, R.C. Cochran, C. Duan, H.J. Grier, L. Huang, L.F. Jackson, D.B. Groman, W. King V, E.D. Lund, C.C. Mylonas, A.R. Place, J.L. Specker, P. Swanson, Y. Tao, Y. Zohar, and L.C. Woods III are acknowledged for allowing us to present their previously unpublished or published results. We are grateful to T.I.J. Smith, C.C. Mylonas, W. King V, and G. M. Weber for proofreading and insightful editorial comments. S.A. Heppell and L.F. Jackson are acknowledged for help with the illustrations and L. Peacock is appreciated for assistance in searching the available literature. One of us (C.V.S.) expresses appreciation of J. Daniels, J. Beam, and J.W. Black for help addressing the editors requests for revision. Our research presented in the chapter was supported by grants from the University of North Carolina Sea Grant College Program (NA86AA-D-SG046, NA86AA-D-SG062 and NA90AA-D-SG062) and the National Coastal Resources Research and Development Institute (grant NA87AAA-D-SG065, contract 2-5606-22-2). This chapter is dedicated to V.A. Sullivan on the occasion of her thirteenth birthday.

60

References

Almendras, J.M., Duenas, C., Nacario, J., Sherwood, N. and Crim, L.W., 1988. Sustained hormone release. III. Use of gonadotropin releasing hormone analogues to induce multiple spawnings in sea bass, Lares calcarifer. Aquaculture, 74:97-111. Bayless, J.D., 1972. Artificial propagation and hybridization of striped bass, Morone saxatilis (Walbaum). South Carolina Wildlife and Marine Resources Department, Columbia. Berlinsky, D.L., 1989. Changes in steroid hormones during oocyte development in the striped bass Morone saxatilis. Ph.D. Dissertation. Department of Zoology, University of Rhode Island, Kingston. Berlinsky, D.L., Jackson, L.F., Smith, T.I.J. and Sullivan, C.V., 1995a. The annual reproductive cycle of the white bass, Morone chrysops. Journal of the World Aquaculture Society, 26: 252-260. Berlinsky, D.L., Fabrizio, M.C., O'Brien, J.F. and Specker, J.L., 1995b. Age-at-maturity estimates for Atlantic coast female striped bass. Transactions of the American Fisheries Society, 124:207-215. Berlinsky, D.L. and Specker, J.L., 1991. Changes in gonadal hormones during oocyte development in the striped bass, Morone saxatilis. Fish Physiology and Biochemistry, 9:51-62. Berlinsky, D.L., Tao, Y. and Sullivan, C.V., 1995c. Isolation and characterization of a vitellogenin receptor in white perch, Morone americana. Page 358 in F.W. Goetz and P. Thomas, editors. Proceedings of the fifth international symposium on the reproductive physiology of fish. University of Texas, Austin. Fish Symposium 95, Publishers, Austin. Bern, H.A., McCormick, S.D., Kelley, K.M., Gray, E.E., Nishioka, R.S., Madsen, S.S. and Tsai, P.I., 1991. Insulin-like growth factors "under water:" role in growth and function of fish and other poikilothermic vertebrates. Pages 85-96 in: E.M. Spencer, editor. Modem concepts of insulin-like growth factors. Elsevier Science Publishing Co., Inc., New York. Billard, R., Fostier, A., Weil, C. and Breton, B., 1982. Endocrine control of spermatogenesis in teleost fish. Canadian Journal of Fisheries and Aquatic Sciences, 39: 65-79. Bishop, R.D., 1975. The use of circular tanks for spawning striped bass (Morone saxatilis). Proceedings Annual Conference Southeastern Association of Game and Fish Commissioners, 28: 35-44. Blythe, W.G., Helfrich, L.A. and Libey, G., 1994a. Induced maturation of striped bass Morone saxatilis exposed to 6, 9, and 12 month photothermal regimes. Journal of the World Aquaculture Society, 25:183-192. Blythe, W.G., Helfrich, L.A., Beal, W.E., Bosworth, B. and Libey, G.S., 1994b. Determination of sex and maturational status of striped bass (Morone saxatilis) using ultrasonic imaging. Aquaculture, 125:175-184. Blythe, W.G., Helfrich, L.A. and Sullivan, C.V., 1994c. Sex steroid hormone and vitellogenin levels in striped bass (Morone saxatilis) maturing under 6-, 9-, and 12-month photothermal cycles. General and Comparative Endocrinology, 94: 122-134. Bonn, E.W., Bailey, W.M., Bayless, J.D., Erickson, K.E. and Stevens, R.E., editors., 1976. Guidelines for striped bass culture. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Boreman, J. and Lewis, R.R., 1987. Atlantic coastal migration of striped bass. American Fisheries Society Symposium 1" 331-339.

61

Brewer, D.L. and Rees, R.A., 1990. Pond culture of phase I striped bass fingerlings. Pages 99-120/n R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Bromage, N. and Cumaranatunga, R., 1987. Oocyte development in the rainbow trout with special reference to vitellogenesis and atresia. Page 194 m D.R. Idler, L.W. Crim, and J.M. Walsh, editors. Proceedings of the third intemational symposium on the reproductive physiology of fish. Memorial University of Newfoundland, St John's. Brown, C.L. and Nunez, J.M., 1993. Hormone actions and applications in embryogenesis. Pages 333-339 in K.G. Davey, R.E. Peter, and S.S. Tobe, editors. Perspectives in comparative endocrinology. National Research Council, Ottawa, Canada. Brown, C.L., Sullivan, C.V. Bern, H.A. and Dickhoff, W.W., 1987. Occurrence of thyroid hormones in early developmental stages of teleost fish. American Fisheries Society Symposium, 2:144-150. Brown, C.L., Doroshov, S., Nunez, J., Hadley, C., Nishioka, R.S. and Bern, H.A., 1988. Maternal triiodothyronine injections cause increases in swimbladder inflation and survival rates in larval striped bass, Morone saxatilis. Journal of Experimental Biology, 248: 168-176. Brown, C.L., Cochran, M., Doroshov, S. and Bern, H.A., 1989. Effect of maternal T3 treatment on survival of striped bass (Morone saxatilis) fingerlings. Fish Physiology and Biochemistry, 7: 295-300. Callard, G.V., 1991. Spermatogenesis. Pages 303-335/n P.K.T. Pang and M.P. Schreibman, editors. Vertebrate endocrinology: fundamentals and biomedical implications, volume 4A. Academic Press, San Diego. Callard, I.P. and Callard, C.V., 1987. Sex steroid receptors and nonreceptor binding proteins. Pages 355-384 in D.O. Norris and R.E. Jones, editors. Hormones and reproduction in fishes, amphibians, and reptiles. Plenum Press, New York. Carrillo, M., Zanuy, S., Prat, F., Caerda, J., Ramos, J., Mananos, E. and Bromage, N., 1995. Sea bass (Dicentrarchus labrax). Pages 138-168 in N.R. Bromage and R.J. Roberts, editors. Broodstock management and egg and larval quality. Blackwell Science, Ltd., Oxford. Chadwick, H.D., 1965. Determination of sexual maturity in female striped bass (Morone saxatilis). Califomia Fish and Game 51:202-206. Chakraborti, P., Mautra, G. and Bhattacharya, S., 1986. Binding of thyroid hormone to isolated ovarian nuclei from a freshwater perch, Anabas testudineus. General and Comparative Endocrinology, 62: 239-246. Coutant, C.C., 1987a. Poor reproductive success of striped bass from a reservoir with reduced summer habitat. Transactions of the American Fisheries Society, 116:154-160. Coutant, C.C., 1987b. Thermal preference: when does an asset become a liability? Environmental Biology of Fishes, 18: 161-172. Coutant, C.C., 1990a. Temperature-oxygen habitat for freshwater and coastal striped bass in a changing climate. Transactions of the American Fisheries Society, 119: 240-253. Coutant, C.C., 1990b. Summer habitat suitability for striped bass in Chesapeake Bay: reflections on a population decline. Transactions of the American Fisheries Society, 119: 757-778.

62

Crim, L.W., Sherwood, N.M. and Wilson, C.E., 1988. Sustained hormone release. II. Effectiveness of LHRH analog (LHRHa) administration by either single time injection or cholesterol pellet implantation on plasma gonadotropin levels in a bioassay fish, the juvenile rainbow trout. Aquaculture, 74: 87-95. Cyr, D.G. and Eales, J.G., 1988a. Influence of thyroidal status on ovarian function in rainbow trout, Salmo gairdneri. Journal of Experimental Zoology, 248:81-87. Cyr, D.G. and Eales, J.G., 1988b. In vitro effects of thyroid hormones on gonadotropin-induced estradiol- 1713secretion by ovarian follicles of rainbow trout, Salmo gairdneri. General and Comparative Endocrinology, 69: 80-87. Donaldson, E.M., 1981. The pituitary-interrenal axis as an indicator of stress in fish. Pages 11-76/n A.D. Pickering, editor. Stress and fish. Academic Press, New York. Donaldson, E.M. and Hunter, G.A., 1983. Induced final maturation, ovulation, and spermiation in cultured fish. Pages 351-404 in Fish physiology, volume 10B. Academic Press, New York. Dorazio, R.M., Hattala, K.M., McCollough, C.B. and Skjeevelaand, J.E., 1994. Tag recovery estimates of migration of striped bass from spawning areas of the Chesapeake Bay. Transactions of the American Fisheries Society, 123" 950-963. Dorfman, D., 1976. Occurrence of a hermaphroditic white perch (Morone americana). The Progressive Fish-Culturist, 38:45. Drakenberg, J.K., Sara, V.R., Lindhal, K.I. and Kewish, B., 1989. The study of insulin-like growth factors in tilapia, Oreochromis mossambicus. General and Comparative Endocrinology, 74:173-180. Duan, C., Duguay, S.J. and Plisetskaya, E.M., 1993a. Insulin-like growth factor I (IGF-I) mRNA expression in coho salmon, Oncorhynchus kisutch: tissue distribution and effects of growth hormone/prolactin family proteins. Fish Physiology and Biochemistry, 11" 371-379. Duan, C., Hanzawa, N., Takeuchi, Y., Hamada, E., Miyachi, S. and Hirano, T., 1993b. Use of primary cultures of salmon hepatocytes for the study of hormonal regulation of insulin-like growth factor I expression in vitro. Zoological Science, 10: 473-480. Eales, J.G., 1985. The peripheral metabolism of thyroid hormones and regulation of thyroidal status in poikilotherms. Canadian Journal of Zoology, 63:1217-1231. Fay, C.W., Neves, R.J. and Pardue, G.B., 1983. Species profiles: life histories of coastal fishes and invertebrates (Mid-Atlantic). Striped bass. U.S. Fish and Wildlife Service, Division of Biological Services, FWS/OBS-82/I 1. U.S. Army Corps of Engineers, TR EL-82/I I. Fostier, A., Jalabert, B., Billard, R., Breton, B. and Zohar, Y., 1983. The gonadal steroids. Pages 277-372 in W.S. Hoar, D.J. Randall and E.M. Donaldson, editors. Fish physiology, volume 9A. Academic Press, New York. Fostier, A., Le Gac, F. and Loir, M., 1987. Steroids in male reproduction. Page 239 in D.R. Idler, L.W. Crim and J.M. Walsh, editors. Proceedings of the third international symposium on reproductive physiology of fish. Memorial University of Newfoundland, St. John's. Funkenstein, B., Silbergeld, A., Cavari, B. and Yaron, Z., 1989. Growth hormone increases plasma levels of insulin-like growth factors (IgF-I) in a teleost, the gilthead seabream (Sparus aurata). Journal of Endocrinology, 120: 1219-1221.

63

Gallagher, M., 1987. Nutritional requirements of striped bass and hybrids. Pages 53-60 in R. Hodson, T. Smith, J. McVey, R. Harrell, and N. Davies, editors. Hybrid striped bass culture: status and perspective. University of North Carolina Sea Grant Publication, UNC-5-87-03, North Carolina State University, Raleigh. Goodyear, C.P., Cohen, J.E. and Christensen, S.W., 1985. Maryland striped bass: recruitment declining below replacement. Transactions of the American Fisheries Society, 114:146-151. Goetz, F.W., 1983. Hormonal control of final oocyte maturation and ovulation in fishes. Pages 117-170 in W.S. Hoar, D.J. Randall, and E.M. Donaldson, editors. Fish physiology, volume 9B. Academic Press, New York. Goetz, F.W., Bemtson, A.K. and Ranjan, M., 1991. Ovulation: mediators at the ovarian level. Pages 127-202 in P.K.T. Pang, and M.P. Schreibman, editors. Vertebrate endocrinology: fundamentals and biomedical implications, volume 4A. Academic Press, San Diego. Goos, H.J.Th., 1994. Pubertal development: big questions, small answers. Pages 11-20 in E.A. Faccinetti, editor. Cellular communication in reproduction. Journal of Endocrinology, Ltd., Bristol. Goos, H.J.Th., 1987. Steroid feedback on pituitary gonadotropin secretion. Pages 16-20 m D.R. Idler, L.W. Crim, and J.M. Walsh, editors. Proceedings of the third international symposium on reproductive physiology of fish. Memorial University of Newfoundland, St. John's.. Gothilf, Y., Chen, T.T. and Zohar, Y., 1995a. Molecular cloning and characterization of a novel gonadotropin-releasing hormone from gilthead seabream (Sparus aurata). Marine Molecular Biology and Biotechnology, 4: 27-35. Gothilf, Y., Elizur, A. and Zohar, Y., 1995b. Three forms of gonadotropin-releasing hormone in gilthead seabream and striped bass: physiological and molecular studies. Pages 52-54 /n F.W. Goetz, and P. Thomas, editors. Proceedings of the fifth international symposium on the reproductive physiology of fish. University of Texas at Austin. Fish Symposium 95, Publishers, Austin. Gorbman, A, Dickhoff, W.W., Vigna, S.R., Clark, N.B. and Ralph, C.L., 1983. Morphology of the pituitary gland system. Pages 45-93 in Comparative Endocrinology. John Wiley and Sons, New York. Grier, H.J., 1981. Cellular organization of the testes and spermatogenesis in fishes. American Zoologist, 21: 345-357. Grier, H.J. and Abraham, M., 1983. A model for testicular recrudescence in Oreochromis aureus. Pages 200-209 in L. Fishelson, and Z. Yaron, editors. International symposium on tilapia in aquaculture. Tel Aviv University Press, Tel Aviv. Grier, H.J., Linton, L.R., Leatherland, J.F. and de Vlaming, V.L., 1980. Structural evidence for two different testicular types in teleost fishes. American Journal of Anatomy, 159:331-345. Grier, H.J., Taylor, R.G. and Reese, R.O., 1987. The mechanism of tubule elongation during testicular recrudescence in the redfish, Sciaenops ocellatus (Perciformes). Pages 285-291 in S.O. Kullander, and B. Femholm, editors. Proceedings of the fifth congress of European ichthyologists. Swedish Museum of Natural History, Stockholm. Grimes, D.V., 1993. Vitellolipid and vitelloprotein profiles of environmentally stressed and nonstressed populations of striped bass. Transactions of the American Fisheries Society, 122:636-641. Groman, D.B., 1982. Histology of the striped bass. American Fisheries Society Monograph 3. Bethesda, MD. Guraya, S.S., 1986. The cell and molecular biology of fish oogenesis. Monographs in Developmental Biology (H.W. Sauer, editor), volume 18. Karger, New York.

64

Gutierrez, J., Parrizas, M., Carneiro, N., Maestro, J.L. Maestro, M.A. and Planas, J., 1993. Insulin and IgF-I receptors and tyrosine kinase activity in carp ovaries: changes with reproductive cycle. Fish Physiology and Biochemistry, 11: 247-254. Harms, C.A., Sullivan, C.V., Hodson R.G. and Stoskopf, M.K., 1996. Clinical pathology and histopathology characteristics of net-stressed striped bass with "red-tail." Journal of Aquatic Animal Health, 8: 82-86. Harrell, R.M., 1984. Review of striped bass brood stock acquisition, spawning methods and fry production. Pages 45-57 in J.P. McCraren, editor. The aquaculture of striped bass: a proceedings. Maryland Sea Grant Publication UM-SG-MAP-84-01. University of Maryland, College Park. Harrell, R.M., 1992. Stress mitigation by use of salt and anesthetic for wild striped bass captured for brood stock. The Progressive Fish-Culturist, 54: 228-233. Harrell, R.M., Kerby, J.H. and Minton, R.V., editors, 1990a. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Harrell, R.M., Kerby, J.H., Smith, T.I.J. and Stevens, R.E., 1990b. Striped bass and striped bass hybrid culture: the next twenty five years. Pages 253-261 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Harrell, R.M. and Moline, M.A., 1992. Comparative stress dynamics of brood stock striped bass Morone saxatilis associated with two capture techniques. Aquaculture, 23: 58-63. Harrell, R.M. and Woods, III, L.C., 1995. Comparative fatty acid composition of eggs from domesticated and wild striped bass (Morone saxatilis). Aquaculture, 133: 225-233. Harris, Jr., R.C., Bums, B.L., Johnson, H.B. and Rulifson, R.A., 1985. An investigation of size, age and sex of North Carolina striped bass. North Carolina Department of Natural Resources and Community Development, Division of Marine Fisheries, Completion Report, Project AFC-18, Morehead City. Hassin, S., Elizur, A. and Zohar, Y., 1995. Molecular cloning and sequence analysis of striped bass (Morone saxatilis) gonadotropin-I and -II subunits. Journal of Molecular Endocrinology, 15:23-35. Henderson-Arzapalo, A. and Colura, R.L., 1987. Laboratory maturation and induced spawning of striped bass. The Progressive Fish-Culturist, 49: 60-62. Hirano, T., 1986. The spectrum of prolactin action in teleosts. Pages 53-74 in C.L. Ralph, editor. Comparative endocrinology: development and directions. Alan Liss, New York. Hislop, J.R.G., 1988. The influence of maternal length and age on the size and weight of eggs and the relative fecundity of haddock, Melanogrammus aegelefinus in British waters. Journal of Fish Biology, 32: 923-930. Hocutt, C.H., Seibold, S.E., Harrell, R.M., Jesien, R.V. and Bason, W.H., 1990. Behavioral observations of the striped bass (Morone saxatilis) on the spawning grounds of the Choptank and Nanticoke Rivers of Maryland, USA. Journal of Applied Ichthyology, 6:21-22. Hodson, R.G. and Sullivan, C.V., 1993. Induced maturation and spawning of domestic and wild striped bass (Morone saxatilis) broodstock with implanted GnRH analogue and injected hCG. Journal of Aquaculture and Fisheries Management, 24: 271-280.

65

Huang, L., 1992. GH cells and PRL cells in the pituitary gland of striped bass. M.Sc. Thesis, Department of Zoology, University of Rhode Island, Kingston. Huang, L. and Specker, J.L., 1994. Growth hormone- and prolactin-producing cells in the pituitary gland of striped bass (Morone saxatilis): immunocytochemical characterization at different life stages. General and Comparative Endocrinology, 94: 225-236. Huang, L., Specker, J.L. and Benison, D.A., 1996. Effect of T3 on the growth and survival of striped bass larvae. Fish Physiology and Biochemistry, 15: 57-64. Idler, D.R. and Ng, T.B., 1983. Teleost gonadotropins: isolation, biochemistry, and function. Pages 187-221 in W.S. Hoar, D.J. Randall, and E.M. Donaldson, editors. Fish physiology, volume 9A. Academic Press, New York. Jackson, L.F. and Sullivan, C.V., 1995. Reproduction of white perch: the annual gametogenic cycle. Transactions of the American Fisheries Society, 124: 563-577. Kah, O., Anglade, I., Leprete, E., Dubourg, P. and de Monbrison, D., 1993. The reproductive brain in fish. Fish Physiology and Biochemistry, 11: 85-98. Kawauchi, H., Suzuki, K., Itoh, H. and Swanson, P., 1989. The duality of teleost gonadotropins. Fish Physiology and Biochemistry, 7: 29-38. Kerby, J.H., 1986. Striped bass and striped bass hybrids. Pages 127-147 in R.R. Stickney, editor. Culture of nonsalmonid freshwater fishes. CRC Press, Inc., Boca Raton, FL. Kerby, J.H. and Harrell, R.M., 1990. Hybridization, genetic manipulation, and gene pool conservation of striped bass. Pages 159-190 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. King, W.V., Thomas, P., Harrell, R.M., Hodson, R.G. and Sullivan, C.V., 1994a. Plasma levels of gonadal steroids during f'mal oocyte maturation of striped bass, Morone saxatilis. General and Comparative Endocrinology, 95: 178-191. King, W.V., Thomas, P. and Sullivan, C.V., 1994b. Hormonal regulation of final maturation of striped bass oocytes in vitro. General and Comparative Endocrinology, 96:223-233. King, W.V., Berlinsky, D.L. and Sullivan, C.V., 1995a. Involvement of gonadal steroids in f'mal oocyte maturation of white perch (Morone americana) and white bass (M. chrysops): in vivo and in vitro studies. Fish Physiology and Biochemistry, 14: 489-500. King, W.V., Ghosh, S., Thomas, P. and Sullivan, C.V., 1995b. Ovarian receptors for 17a,2013,21-tfihydroxy-4-pregnen3-one (2013-S) in striped bass. Page 314 in F.W. Goetz and P. Thomas, editors. Proceedings of the fifth international symposium on the reproductive physiology of fish. University of Texas at Austin. Fish Symposium 95, Publishers, Austin. Kishida, M., Anderson, T.R. and Specker, J.L., 1992. Induction by b-estradiol of vitellogenin in striped bass (Morone saxatilis): characterization and quantification in plasma and mucus. General and Comparative Endocrinology, 88: 29-39. Knutsen, G.M. and Tilseth, S., 1985. Growth development and feeding success of Atlantic cod larvae Gadus morhua related to eoo:,~size. Transactions of the American Fisheries Society, 110:168-179.

66

Kohlenstein, L.C., 1981. On the proportion of the Chesapeake Bay stock of striped bass that migrate into the coastal fishery. Transactions of the American Fisheries Society, 114:507-511. Kohler, C.C., Sheehan, R.J., Habicht, C., Madison, J.A. and Kayes, T.B., 1994. Habituation to captivity and controlled spawning of white bass. Transactions of the American Fisheries Society, 123: 964-974. Lam, T.J., 1983. Environmental influences on gonadal activity in fish. Pages 81-154 in W.S. Hoar, D.J. Randall and E.M. Donaldson, editors. Fish physiology, volume 9B. Academic Press, New York. Leatherland, J.F., 1987. Thyroid hormones and reproduction. Pages 411-431 in D.O. Norris and R.E. Jones, editors. Hormones and reproduction in fishes, amphibians, and reptiles. Plenum Press, New York. Le Gac, F., Blaise, O., Fostier, A., Le Bail, P.-Y., Loir, M., Mourot, B. and Wei, C., 1993. Growth hormone (GH) and reproduction: a review. Fish Physiology and Biochemistry, 11: 219-232. Lewis, R.M., 1962. Sexual maturity as determined from ovum diameters in striped bass from North Carolina. Transactions of the American Fisheries Society, 91: 279-282. Lewis, R.M. and Bonner, Jr., R.R., 1966. Fecundity of the striped bass, Roccus saxatilis (Walbaum) Transactions of the American Fisheries Society, 95:328-331 Liley, N.R. and Stacey, N.E., 1983. Hormones, pheromones and reproductive behavior in fish. Pages 1-49 in W.S. Hoar, D.J. Randall, and E.M. Donaldson, editors. Fish physiology, volume 9B. Academic Press, New York. Lin, H.R., Van Der Kraak, G., Liang, J.-Y., Peng, C., Li, G.-Y., Lu, L.-Z., Zhou, X.-J., Chang, M.-L. and Peter, R.E., 1986. The effects of drugs which block the effects of dopamine on gonadotropin secretion and ovulation in fish cultured in China. Pages 139-150/n R. Billiard and J. Marcel, editors. Aquaculture ofcyprinids. INRA, Paris. Lofts, B., 1987. Testicular function. Pages 283-325 in D.O. Norris and R.E. Jones, editors. Hormones and reproduction in fishes, amphibians and reptiles. Plenum Press, New York. Lund, E.D., Place, A.R., and Sullivan, C.V., 1995. Lipid contents of female striped bass plasma and ovaries exhibit seasonal changes associated with oocyte maturation. Page 371 in F.W. Goetz and P. Thomas, editors. Proceedings of the fifth intemational symposium on the reproductive physiology of fish. University of Texas at Austin. Fish Symposium 95, Publishers, Austin. Mananos, E., Swanson, P., Stubblefield, J. and Zohar, Y., 1995. Purification of striped bass (Morone saxatilis) gonadotropin II and development of a homologous enzyme immunoassay for its measurement. Page 36 m F.W. Goetz and P. Thomas, editors. Proceedings of the fifth international symposium on the reproductive physiology of fish. University of Texas at Austin. Fish Symposium 95, Publishers, Austin. Mazeaud, M.M. and Mazeaud, F., 1981. Adrenergic responses to stress in fish. Pages 49-76 in A.D. Pickering, editor. Stress and fish. Academic Press, New York. Mazzi, V. and Vellano, C., 1987. Prolactin and reproduction. Pages 87-116/n D.O. Norris and R.E. Jones, editors. Hormones and reproduction in fishes, amphibians, and reptiles. Plenum Press, New York. Merriman, D., 1941. Studies on the striped bass (Roccus saxatilis). Annals and Magazine of Natural History Series 11(5): 55-64. McLauren, L.B., Cooper, J.C., Hoff, T.B. and Lander, V., 1981. Movements of Hudson River striped bass. Transactions of the American Fisheries Society, 110:158-167.

67

Miura, T., Kobayashi, T. and Nagahama, Y., 1994. Hormonal regulation of spermatogenesis in the Japanese eel (Anguilla japonica). Pages 631-635 in K.G. Davey, R.E. Peter and S.S. Tobe, editors. Perspectives in comparative endocrinology. National Research Council, Ottawa, Canada. Miura, T., Yamauchi, K., Takahishi, H. and Nagahama, Y., 1991. Hormonal induction of all stages of spermatogenesis in vitro in the male Japanese eel (Anguillajaponica). Proceedings of the National Academy of Sciences, USA 88: 5774-5778. Mommsen, T.P. and Walsh, P.J., 1988. Vitellogenesis and oocyte assembly. Pages 347-406 in W.S. Hoar, D.J. Randall and E.M. Donaldson, editors. Fish physiology, volume 9A. Academic Press, New York. Monteleone, D.M. and Houde, E.D.,. 1990. Influence ofmatemal size on survival and growth of striped bass Morone saxatilis Walbaum eggs and larvae. Journal of Exploratory Marine Biology and Ecology, 140: 1-11. Moser, M., Whipple, J., Sakanari, J. and Reilly, C., 1938. Protandrous hermaphroditism in striped bass from Coos Bay, Oregon. Transactions of the American Fisheries Society, 112: 567-569. Myionas, C.C., Hinshaw, J.M. and Sullivan, C.V., 1992. GnRHa-induced ovulation of brown trout (Salmo trutta) and its effects on egg quality. Aquaculture, 106: 379-392. Mylonas, C.C., Tabata, Y., Langer, R. and Zohar, Y., 1995. Preparation and evaluation of polyanhydride microspheres containing gonadotropin-releasing hormone (GnRH) for inducing ovulation and spermiation in fish. Journal of Controlled Release, 35" 23-34. Nagahama, Y., 1983. The functional morphology ofteleost gonads. Pages 223-275/n W.S. Hoar, D.J. Randall and E.M. Donaldson, editors. Fish physiology, volume 9A. Academic Press, New York. Nagahama, Y., 1986. Testes. Pages 399-437 in P.K.T. Pang and M.P. Schreibman, editors. Vertebrate endocrinology: fundamentals and biomedical implications, volume I. Academic Press, New York. Nagahama, Y., 1987a. Endocrine control of oocyte maturation. Pages 171-202 in D.O. Norris and R.E. Jones, editors. Hormones and reproduction in fishes, amphibians and reptiles. Plenum Press, New York. Nagahama, Y., 1987b. Gonadotropin action on gametogenesis and steroidogenesis in teleost gonads. Zoological Science, 4: 209-222. Nagahama, Y., 1987c. 17tx,20[3-Dihydroxy-4-pregnen-3-one: A teleost maturation-inducing hormone. Development, Growth and Differentiation, 29: 1-12. Nagahama, Y., 1993. Molecular biology of oocyte maturation in fish. Pages 193-198 in K.G. Davey, R.E. Peter and S.S. Tobe, editors. Perspectives in comparative endocrinology. National Research Council, Ottawa, Canada. Nagahama, Y. and Adachi, S.. 1985. Identification of maturation-inducing steroid in a teleost, the amago salmon (Oncorhynchus rhodurus). Developmental Biology, 109:428-435. Nagahama, Y, Yoshikuni, M., Yamashita, M. and Tanaka, M., 1994. Regulation of oocyte maturation in fish. Pages 393-439 in N.M. Sherwood and C.L. Hew, editors. Fish physiology, volume 13. Academic Press, New York. Ng, T.B. and Idler, D.R., 1983. Yolk formation and differentiation in teleosts fishes. Pages 373-404 in W.S. Hoar, D.J. Randall and E.M. Donaldson, editors. Fish physiology, volume 9A. Academic Press, New York.

68

Nozaki, M., Naito, N., Swanson, P., Miyata, K., Nakai, Y., Oota, Y., Suzuki, K. and Kawauchi, H., 1990a. Salmon pituitary gonadotrophs I. Distinct cellular distributions of two gonadotropins, GTH I and GTH II. General and Comparative Endocrinology, 77: 348-357. Olsen, E.J. and Rulifson, R.A.,. 1992. Maturation and fecundity of Roanoke-Albemarle Sound striped bass. Transactions of the American Fisheries Society, 121: 524-537. Pankhurst, N.W., 1985. Final maturation and ovulation of oocytes of the goldeye, Hiodon alosoides (Raf'mesque), in vitro. Canadian Journal of Zoology, 63:1003-1009. Pankhurst, N. and Carragher, 1991. Seasonal endocrine cycles in marine teleosts. Pages 131-135 in A.P. Scott, J.P. Sumpter, D.E. Kime and M.S. Rolfe, editors. Proceedings of the fourth international symposium on reproductive physiology of fish. Fish Symposium 91, Publishers, Sheffield, U.K. Patino, R. and Takashima, F., 1995. Gonads. Pages 128-152 in F. Takashima and T. Hibiya, editors. An atlas of fish histology, second edition. Kodansha-Gustav Fischer Verlag, Tokyo-New York. Patino, R. and Thomas, P., 1990. Gonadotropin stimulates 17~,2013,21-trihydroxy-4-pregnen-3-one production from endogenous substrates in Atlantic croaker ovarian follicles undergoing final maturation m vitro. General and Comparative Endocrinology, 78: 474-478. Peter, R.E., Chang, J.P., Nahomiak, C.S., Omeljaniuk, R.J., Sokolowska, M., Shih, S.H. and Billard, R., 1986. Interactions of catecholamines and GnRH in regulation of gonadotropin secretion in teleost fish. Recent Progress in Hormone Research, 42: 513-547. Peter, R.E., Lin, H.R. and Van Der Kraak, G., 1988. Induced ovulation and spawning of cultured freshwater fish in China: advances in application of GnRH analogues and dopamine antagonists. Aquaculture, 74: 1-10. Peter, R.E., Trudeau, V.L. and Sloley, B.D., 1991. Brain r%malation of reproduction in teleosts. Bulletin of the Institute of Zoology, Academia Sinica. Monograph, 16: 89-118. Pierce, J.G. and Parsons, T.F., 1981. Glycoprotein hormones: structure and function. Annual Review of Biochemistry, 50: 465-495. Pickering, A.D., editor, 1981. Stress and fish. Academic Press, New York. Planas, J.V., Swanson, P. and Dickhoff, W.W., 1991. Regulation of testicular steroidogenesis by coho salmon gonadotropins, GTH I and GTH II, in vitro. Page 103 in A.P. Scott, J.P. Sumpter, D.E. Kime and M.S. Rolfe, editors. Proceedings of the fourth international symposium on reproductive physiology of fish. Fish Symposium 91, publisher, Sheffield, U.K. Planas, J.V., Swanson, P. and Dickhoff, W.W., 1993. Regulation of testicular steroid production in vitro by gonadotropins (GTH I and GTH II) and cyclic AMP in coho salmon (Oncorhynchus kisutch). General and Comparative Endocrinology, 91: 8-24. Planas, J.V., Swanson, P., Rand-Weaver, M., and Dickhoff, W.W., 1992. Somatolactin stimulates in vitro gonadal steroidogenesis in coho salmon, Oncorhynchus kisutch. General and Comparative Endocrinology, 87: 1-5. Powell, J.F.F., Zohar, Y., Park, C., Fischer, A.G., Craig, A.G., Rivier, J.F., Lovejoy, D.A. and Sherwood, N.M., 1994. Three forms of gonadotropin-releasing hormone characterized from brains of one species. Proceedings of the National Academy of Sciences, USA 91: 12081-12085.

69

Rand-Weaver, M., Baker, B.I. and Kawauchi, H., 1991a. Cellular localization of somatolactin in the pars intermedia of some teleost fishes. Cell and Tissue Research, 263:207-215. Rand-Weaver, M., Noso, T., Muramato, K. and Kawauchi,. H., 1991b. Isolation and characterization of somatolactin, a new protein related to growth hormone and prolactin from Atlantic cod (Gadus morhua) pituitary glands. Biochemistry, 30:1509-1515. Rand-Weaver, M., and Kawauchi, H., 1992. A rapid procedure for the isolation ofbioactive growth hormone. General and Comparative Endocrinology, 85:341-345. Rees, R.A., and Harrell, R.M., 1990. Artificial spawning and fry production of striped bass and hybrids. Pages 43-72 in R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southem Division, American Fisheries Society, Bethesda, MD. Redding, J.M. and Patino, R., 1993. Reproductive physiology. Pages 503-534 in D.H. Evans, editor. The physiology of fishes. CRC Press, Inc., Boca Raton, FL. Sakai, N., Ueda, H., Suzuki, N. and Nagahama, Y., 1989a. Steroid production by amago salmon (Oncorhynchus rhodurus) testes at different developmental stages. General and Comparative Endocrinology, 75:231-240. Sakai, N., Ueda, H., Suzuki, N. and Nagahama, Y., 1989b. Involvement of sperm in the production of 17~,20[3-dihydroxy- 4-pregnen-3-one in testis of spermiating rainbow trout, Salmo gairdneri. Biomedical Research, 10: 131-138. Scott, A.P. and Canario, A.V.M., 1987. Status of oocyte maturation-inducing steroids in teleosts. Pages 224-234 in D.R. Idler, L.W. Crim and J.M. Walsh, editors. Proceedings of the third international symposium on reproductive physiology of fish. Memorial University of Newfoundland, St. John's, Canada. Scott, A.P. and Vermeirssen, E.L.M., 1994. Production of conjugated steroids by teleost gonads and their role as pheromones. Pages 645-654 in K.G. Davey, R.E. Peter and S.S. Tobe, editors. Perspectives in comparative endocrinology. National Research Council, Ottawa, Canada. Schreck, C.B., Bradford, C.S., Fitzpatrick, M.S. and Patino, R., 1989. Regulation of the interrenal of fishes: non-classical control mechanisms. Fish Physiology and Biochemistry ,7: 259-265. Schreibman, M.P. 1986. The pituitary gland. Pages 11-47 in P.K.T. Pang and M.P. Schreibman, editors. Vertebrate endocrinology: fundamentals and biomedical implications, volume 1. Academic Press, New York. Schreibman, M.P., Holtzman, S. and Margolis-Nunno, H., 1991. Maturation and aging of the reproductive system. Pages 343-378 in P.K.T. Pang, and M.P. Schreibman, editors. Vertebrate endocrinology: fundamentals and biomedical implications, volume 4A. Academic Press, New York. Schreibman, M.P., Schartl, M., Kallman, K. and Magliulo-Cepriano, L., 1994. Molecular approaches to study the genetic regulation of the fish reproductive system. Pages 343-352/n K.G. Davey, R.E. Peter and S.S. Tobe, editors. Perspectives in comparative endocrinology. National Research Council, Ottawa, Canada. Schultz, L.P., 1931. Hermaphroditism in the striped bass. Copeia, 1931:64. Secor, D.H., Dean, J.M., Curtis, T.A. and Sessions, F.W., 1992. Effect of female size and propagation methods on larval production at a South Carolina striped bass (Morone saxatilis) hatchery. Canadian Journal of Fisheries and Aquatic Sciences, 49:1778-1787.

70

Selman, K. and Wallace, R.A., 1989. Cellular aspects of oocyte growth in teleosts. Zoological Science 6:211-231. Setzler, E.M., Boynton, W.R.. Wood, K.V., Zion, H.H., Lubbers, L., Mountford, N.R., Frere, P., Tucker, L. and Mihursky, J.A., 1980. Synopsis of biological data on striped bass, Morone saxatilis (Walbaum). NOAA (National Oceanic and Atmospheric Administration) Technical Report, NMFS (National Marine Fisheries Service) Circular 433 (Also: FAO (Food and Agriculture Organization of the United Nations) Fisheries Synopsis 121. Sherwood, N.M., Crim, L.W., Carolsfeld, J. and Waiters, S.M., 1988. Sustained hormone release. I. Characteristics of in vitro release of gonadotropin-releasing hormone analogue (GnRH-A) from pellets. Aquaculture, 74: 75-86. Sherwood, N.M., Parker, D.B., McRory, J.E. and Lescheid, D.W., 1994. Molecular evolution of growth hormone-releasing hormone and gonadotropin-releasing hormone. Pages 3-65 in N.M. Sherwood and C.L. Hew, editors. Fish physiology, volume 13. Academic Press, New York. Smith, T.I.J. and Jenkins, W.E., 1985. Status of aquaculture of striped bass (Morone saxatilis) and its white bass (Morone chrysops) hybrids, and current research in South Carolina. Pages 553-582 in Proceedings of the second international conference on warm water aquaculture fm fish. Brigham Young University, Laie, Hawaii. Smith, T.I.J. and Jenkins, W.E., 1987. Aquaculture research with striped bass (Morone saxatilis) and its hybrids in South Carolina. Proceedings of the Annual Conference of the Southeastern Association of Fish and Wildlife Agencies, 39:217-227. Smith, T.I.J. and Jenkins, W.E., 1988. Culture and controlled spawning of striped bass (Morone saxatilis) to produce striped bass and white bass (M. chrysops) hybrids. Proceedings of the Annual Conference of the Southeastern Association of Fish and Wildlife Agencies 40:152-162. Smith, T.I.J., Jenkins, W.E. and Heyward, L.D. 1996. Production and extended spawning of cultured white bass broodstock. The Progressive Fish-Culturist 58:85-91. Smith, J.M. and Whitehurst, D.K., 1990. Tank spawning methodology for the production of striped bass. Pages 73-77 in R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Sorenson, P.W., Goetz, F.W., Scott, A.P. and Stacey, N.E., 1991. Recent studies indicate that goldfish use mixtures of unmodified hormones and hormonal metabolites as sex pheromones. Pages 191-193 in A.P. Scott, J.P. Sumpter, D.E. Kime and M.S. Rolfe, editors. Proceedings of the fourth international symposium on the reproductive physiology of fish. Fish Symposium 91, publisher, Sheffield, U.K. Soyano, K., Saito, T., Nagae, M. and Yamauchi, K., 1993. Effects of thyroid hormone on gonadotropin-induced steroid production in medaka, Oryzias latipes, ovarian follicles. Fish Physiology and Biochemistry, 11: 265-272. Soyano, K., Yamada, H. and Yamauchi, K., 1989. Uptake of thyroid hormone and its generation in the rainbow trout ovarian follicle. Pages 107-108 in D.K.O. Chan, editor. Proceedings of the international symposium on hormones and the environment. University of Hong Kong, Hong Kong. Specker, J.L. and Anderson, T.R., 1994. Developing an ELISA for a model protein-vitellogenin. Pages 567-578 in P. Hochachka and T. Mommsen, editors. Biochemistry and molecular biology of fishes. Elsevier Science Publishing Co., Inc., New York.

71

Specker, J.L., Berlinsky, D.L., Bibb, H.D. and O'Brien, J.F., 1987. Oocyte development in the striped bass: factors influencing estimates of age at maturity. American Fisheries Society Symposium, 1:162-174. Specker, J.L. and Sullivan, C.V., 1994. Vitellogenesis in fishes: status and perspectives. Pages 304-315 in K.G. Davey, R.E. Peter and S.S. Tobe, editors. Perspectives in comparative endocrinology. National Research Council, Ottawa, Canada. Springate, J.R.C., Bromage, N.R., Elliot, J.A.K. and Hudson, D.L., 1984. The timing of ovulation and stripping and their effects on the rates of fertilization and survival to eyeing, hatch, and swim up in the rainbow trout, Salmo gairdneri. Aquaculture, 43:313-322. Stacey, N.E., Cardwell, J.R., Liley, N.R., Scott, A.P., and Sorenson, P.W., 1994. Hormones as sex pheromones in fish. Pages 438-448 in K.G. Davey, R.E. Peter and S.S. Tobe, editors. Perspectives in comparative endocrinology. National Research Council, Ottawa, Canada. Stacey, N.E., Sorenson, P.W., Dulka, J.G., Cardwell, J.R. and Irvine, A.S., 1991. Fish sex pheromones: current status and potential applications. Bulletin of the Institute of Zoology, Academia Sinica Monograph, 16:189-227. Stevens, R.E., 1966. Hormone-induced spawning of striped bass for reservoir stocking. The Progressive Fish-Culturist, 28:19-28. Stevens, R.E., 1967. A final report on the use of hormones to ovulate striped bass, Roccus saxatilis (Walbaum). Proceedings of the Annual Conference of the Southeastern Association of Game and Fish Commissioners, 18: 525-538. Stevens, R.E., May, Jr., O.D. and Logan, H.J., 1965. An interim report on the use of hormones to ovulate striped bass (Roccus saxatilis). Proceedings of the Annual Conference of the Southeastern Association of Game and Fish Commissioners, 17: 226-237. Sullivan, C.V., Bemard, M.G., Hara, A. and Dickhoff, W.W., 1989. Thyroid hormones in trout reproduction: enhancement of gonadotropin-releasing hormone analogue and partially purified salmon gonadotropin-induced ovarian maturation in vivo and in vitro. Journal of Experimental Zoology, 250:188-195. Sullivan, C.V.~ Tao, Y., Hodson, R.G., Hara, A., Bennett R.O. and Woods, III, L.C., 1991. Vitellogenin and vitellogenesis in striped bass (Morone saxatilis) broodstock. Pages 315-317 in A.P. Scott, J.P. Sumpter, D.E. Kime and M.S. Rolfe, editors. Proceedings of the fourth international symposium on reproductive physiology of fish. Fish Symposium 91, publishers., Sheffield, U.K. Sumpter, J.P., 1990. General concepts of seasonal reproduction. Pages 13-32 in A.D. Munro, A.P. Scot and T.L Lam, editors. Reproductive seasonality in teleosts: environmental influences. CRC Press, Boca Raton, FL.. Sumpter, J.P., Pottinger, T.G., Rand-Weaver, M. and Campbell, P.M., 1994. The wide-ranging effects of stress on fish. Pages 535-538 in K.G. Davey, R.E. Peter and S.S. Tobe, editors. Perspectives in comparative endocrinology. National Research Council, Ottawa, Canad Swanson, P., 1991. Salmon gonadotropins: reconciling old and new ideas. Pages 2-7 in A.P. Scott, J.P. Sumpter, D.E. Kime and M.S. Rolfe, editors. Proceedings of the fourth international symposium on reproductive physiology of fish. Fish Symposium 91, publishers, Sheffield, U.K. Swanson, P. and Sullivan, C.V., 1991. Isolation of striped bass, Morone saxatilis, pituitary hormones. American Zoologist, 31(5): 3A.

72

Tam, W.H., Roy, J.J. and Makaran, R., 1986. Ovarian cycle and plasma concentrations of estrogen and vitellogenin in the brook trout, Salvelinusfontinalis. Canadian Journal of Zoology, 64: 744-751. Tao, Y., Hara, A., Hodson, R.G., Woods, III, L.C. and Sullivan, C.V., 1993. Purification, characterization and immunoassay of striped bass (Morone saxatilis) vitellogenin. Fish Physiology and Biochemistry, 12:31-46. Tao, Y., Berlinsky, D.L. and Sullivan, C.V., 1996. Characterization of a vitellogenin receptor in white perch, Morone americana. Biology of Reproduction, 55: 646-656. Thomas, P., 1994. Hormonal control of f'mal oocyte maturation in sciaenid fishes. Pages 619-625 in K.G. Davey, R.E. Peter and S.S. Tobe, editors. Perspectives in comparative endocrinology. National Research Council, Ottawa, Canada. Thomas, P., Arnold, C.R. and Holt, G.J., 1995. Red drum and other sciaenids. Pages 118-137 in N.R. Bromage and R.J. Roberts, editors. Broodstock management and egg and larval quality. Blackwell Science, Ltd., Oxford. Tuncer, H. and Harrell, R.M., 1992. Essential fatty acid nutrition of larval striped bass (Morone saxatilis) and palmetto bass (M. saxatilis x M. chrysops). Aquaculture, 110: 105-121. Tuncer, H., Harrell, R.M. and Chai, T.-J., 1993. Beneficial effects of n-3 HUFA enriched Artemm as food for larval palmetto bass (Morone saxatilis x M. chrysops). Aquaculture, 110: 341-359. Tyler, C.R., Sumpter, J.P. and Bromage, N.R., 1987. The hormonal control of vitellogenin uptake into cultured ovarian follicles of the rainbow trout. Page 220 in D.R. Idler, L.W. Crim and J.M. Walsh, editors. Proceedings of the third international symposium on reproductive physiology of fish. Memorial University of Newfoundland, St. John's, Canada. Van

Bohemen, C.G. and Lambert, J.G.D., 1981. Estrogen synthesis in relation to estrone, estradiol, and vitellogenin plasma levels during the reproductive cycle of the female rainbow trout, Salmo gairdneri. General and Comparative Endocrinology, 45:105-114.

Van

Bohemen, C.G., Lambert, J.G.D., Goos, H.J.T. and van Oordt, P.G.W.J., 1982a. Estrone and estradiol participation during exogenous vitellogenesis in the female rainbow trout, Salmo gairdneri. General and Comparative Endocrinology, 46:81-92.

van Bohemen, C.G., Lambert, J.G.D. and van Oordt, P.G.W.J., 1982b. Vitellogenin induction by estradiol in estrone-primed rainbow trout, Salmo gairdneri. General and Comparative Endocrinology, 46:136-139. Waldman, J.R., Dunning, D.J., Ross, Q.E. and Mattson, M.T., 1990. Range dynamics of Hudson River striped bass along the Atlantic coast. Transactions of the American Fisheries Society, 119:910-919. Wallace, R.A. and Selman, K., 1981. Cellular and dynamic aspects of oocyte growth in teleosts. American Zoologist, 21" 135-140. Wallace, R.A. and Selman, K., 1990. Ultrastructural aspects of oogenesis and oocyte growth in fish and amphibians. Journal of Electron Microscopy, 16:175-201. Watanabe, T., 1982. Lipid nutrition in fish. Comparative Biochemistry and Physiology, 73B: 3-15. Westin D.T. and Rogers, B.A., 1978. Synopsis of biological data on the striped bass Morone saxatilis (Walbaum). Marine Technical Report Number 67. University of Rhode Island, Kingston.

73

Whitehurst, D.K. and Stevens, R.E., 1990. History and overview of striped bass culture and management. Pages 1-5 in R.M. Harrell, Kerby, J.H. and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Wilson. R.A., Morgan, R.P., Jones, P.W., Lunsford, Jr., H.R., Lawson, J. and Murphy, S., 1975. Potomac River fisheries study: striped bass spawning stock assessment. University of Maryland, Chesapeake Biological Laboratory, UMCEES 76-14 CBL, Solomons, Maryland. Woodhull, C., 1947. Spawning habits of the striped bass (Roccus saxatilis) in California waters. California Fish and Game, 33: 97-102. Woods, III, L.C., Bennett, R.O. and Sullivan, C.V., 1992. Reproduction of a domestic striped bass brood stock. The Progressive Fish-Culturist 54:184-188. Woods, III, L.C. Kohler, C.C., Sheehan, R.J. and Sullivan, C.V., 1995. Volitional tank spawning of female striped bass with male white bass produces hybrid offspring. Transactions of the American Fisheries Society, 124: 628-632. Woods, III, L.C. and Sullivan, C.V., 1993. Reproduction of striped bass (Morone saxatilis) broodstock: monitoring maturation and hormonal induction of spawning. Aquaculture and Fisheries Management, 24:213-224. Woods, III, L.C., Woiwode, J.G., McCarthy, M.A., Theisen, D.D. and Bennett, R.O., 1990. Noninduced spawning of captive striped bass in tanks. The Progressive Fish-Culturist, 52:201-202. Worth, S.G., 1884. Report on the propagation of striped bass at Weldon, N.C., in the spring of 1884. Bulletin of the United States Fish Commission, 4 (15): 225-230. Worth, S.G., 1903. Striped bass hatching in North Carolina. Transactions of the American Fisheries Society, 32: 98-102. Xiong, F., Suzuki, K. and Hew, C.L., 1994. Control of teleost gonadotropin gene expression. Pages 135-158 in N.M. Sherwood and C. L. Hew, editors. Fish physiology, volume 13. Academic Press, New York. Yan, L., Swanson, P. and Dickhoff, W.W.. 1992. A two-receptor model for salmon gonadotropins (GTH I and GTH II). Biology of Reproduction, 47:418-427. Young, G., Ueda, H. and Nagahama, Y., 1983. Estradiol-1713and 17~,2013-dihydroxy-4-pregnen'3-one production by isolated ovarian follicles of amago salmon (Oncorhynchus rhodurus) in response to mammalian pituitary and placental hormones and salmon gonadotropin. General and Comparative Endocrinology, 52:329-33 5. Zastrow, C.E., Houde, E.D. and Saunders, E.H., 1989. Quality of striped bass (Morone saxatilis) eggs in relation to river source and female weight. Rapports et Proces-Verbaux des Reunions, Conseil rnternational por 1Exploration de la Mer, 191: 34-42. Zohar, Y., 1989. Fish reproduction: its physiology and artificial manipulation. Pages 65-119 in M. Shilo and S. Sarig, editors. Fish culture in warmwater systems: problems and trends. CRC Press, Boca Raton, FL.. Zohar, Y., Elizur, A., Sherwood, N.M., Rivier, J.F. and Zmora, N., 1995a. Gonadotropin-releasing potencies of the three native forms of gonadotropin-releasing hormones present in the brain of gilthead seabream, Sparus aurata. General and Comparative Endocrinology, 97: 289-299. Zohar, Y., Harel, M., Hassin, S. and Tandler, A., 1995b. Gilt-head sea bream. Pages 94-117 in: N.R. Bromage and R.J. Roberts, editors. Broodstock management and egg and larval quality. Blackwell Science, Ltd., Oxford.

This Page Intentionally Left Blank

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

75

Chapter 3

Morone Pond Production Reginal M. Harrell 3.1 INTRODUCTION

3.1.1 General Comments Production strategies for pond culture of striped bass (Morone saxatilis) and its hybrids are dependent on the intended use of the final product. If the intent is to produce large numbers of fish for population enhancement, stocking and harvest protocols will be different from those of production for food-fish grow out. This chapter will focus on the underlying production strategies for the production of food fish, from the larval stage to market size. It will build on the techniques outlined in Harrell et al. (1990) as those guidelines are still the basis for most production techniques for striped bass and hybrid production today. I will provide an overview of those areas where food-fish production differs from producing fish for population enhancement or restoration. For details on the production techniques of the latter, the reader should reference those chapters in Harrell et al. (1990) most appropriate to the context of pond production, in particular Brewer and Rees (1990), Geiger and Tumer (1990), Minton and Harrell (1990), Parker et al. (1990), and Smith et al. (1990). There are four distinct phases of Morone culture (Brewer and Rees, 1990): the hatchery phase, which is covered in Chapter 2 ofthis volume;phase I, whichis rearing larval fish from the hatchery to a 25 to 75 mm fingerling; phase II, where phase I fish are reared an additional five to nine months (yearling) and a size of 75 to 250 mm; andphase 111,where phase II fish are reared to subadult market or adult sizes. These four phases also correspond to different market niches that commercial producers enter for supplying the food fish industry. Chapter 1, Section 1.2.1.2 discusses these various niches and how the producers are divided between the hatchery, fingerling, and food-fish production aspects. 3.1.2 Striped Bass Versus Hybrid Culture For the most part, the culture requirements for hybrids are the same as those for striped bass. The only exception being in the phase I culture protocols for sunshine or Maryland bass (see Chapter 8) where the female of the hybrid is a white bass (M. chrysops) or white perch (M. americana). Resultant larvae of these hybrid crosses are considerably smaller than larvae of striped bass and therefore need special considerations for manipulation of zooplankton in production ponds (see Section 3.2.2). Outside of these differences, culture and feeding strategies are identical for all Morone. One of the attractive features about hybrid striped bass over that of pure striped bass is that the former exhibits heterosis expressed as faster growth, greater hardiness and adaptability to culture conditions, and higher production yields in ponds and in processed fish (see Chapters 8 and 13). It is interesting to note however that this heterosis is not generally evident in the first 30 days of life (Houde and Lubbers, 1986; Tuncer et al., 1990), but it is expressed between 30 and 100 days of age, or by the time the fish are well into phase II production (Tuncer et al. 1990). It appears that hybrids have a heterotic advantage over striped bass in that they have a lower metabolism, thus more energy can be partitioned into growth instead of being allocated to routine maintenance (Tuncer et al., 1990).

76 3.1.3 Hatchery Production Reproductive aspects of Morone culture are detailed in Chapter 2, Rees and Harrell (1990), and Smith and Whitehurst (1990) and will not be expanded upon in this chapter. In general terms, the hatchery phase includes collecting broodstock, spawning manipulation, fertilizing the eggs with the appropriate male, incubating the eggs, allowing the embryos to hatch, and stocking them in the appropriate rearing environment. This chapter focuses exclusively on the pond as a rearing environment (refer to Chapter 5 for information on intensive tank and raceway culture). Unless captive or domesticated broodstock are available in the rearing facility (see Chapter 2), gravid females (Figure 3.1) are captured on, or near, the spawning grounds, and induced to ovulate. If captive females are used, some form of photothermal conditioning is usually required in concert with maturational hormonal implants (see Chapter 2). When larvae first hatch they are known as prolarvae and receive their nourishment from their yolk-sacs (Figure 3.2). By their fifth day (temperature dependent) their mouth parts are formed and they begin exogenous feeding. At this stage they are known as larvae or fry (Figure 3.2), and will remain as such until they metamorphose into juvenile phase I fish (~ 25-30 ram). 3.2 PHASE I PRODUCTION

The key to successfully culturing of phase I striped bass or hybrids is in providing the right food type, size, and quantity; good water quality conditions; and the proper timing of stocking, supplemental feeding, and harvesting. These specific requirements are detailed in a variety of publications (Geiger, 1983a,b; Geiger et al., 1985; Woods et al., 1985; Fitzmayer et al., 1986; Brewer and Rees, 1990; Geiger and Turner, 1990; Harrell and Bukowski, 1990; Peterson, 1991). Because phase I Morone are essentially planktivorous in their feeding habits, it is essential that the timing of zooplankton successional patterns be manipulated to the optimize the

Fig. 3.1. Gravid Chesapeake Bay female striped bass (Photo by J.H. Kerby, National Biological Service, Leetown, WV).

77

Fig. 3.2. Top: striped bass prolarvae at hatch, Bottom: striped bass feeding larvae (note straight digestive tract) (Photo by: Top: R.M. Harrell, Bottom: J.H. Kerby.) right sized prey organism for the fish (Parmley and Geiger, 1985; Harrell and Bukowski, 1990). Increasing your understanding of plankton dynamics and nutrient-phytoplankton-zooplankton interactions are crucial to the success of this approach to pond management (see Chapter 4). 3.2.1 Receipt of Fry Most phase I fingerling producers do not have a production hatchery from which they could produce their own seed stock, thus they purchase their fry from a hatchery. Morone fry are generally shipped in sealed plastic bags with a minimal amount of water that contains pure oxygen (Figure 3.3). Chapter 7 (Section 7.9) discusses larval shipment in detail. The biggest concern before stocking the fry is in water quality differences. The fingerling producer who is receiving the fry should let the shipper know at what densities he prefers having the fish shipped (see Table 7.3), receiving water conditions, and good flight schedules with at least one-hour layover time if the shipment is scheduled to change aircraft. The last thing a producer or shipper wants is for the fish to miss a flight connection and sit in the hot sun for several hours. Once the fry are received, the producer should take care to stabilize water quality and temperature differences between the shipping water and the receiving water (see Section 7.9). 3.2.2 Pond Preparation As mentioned, one of the keys to successful phase I fingerling production is in providing the proper size and type of food. The key to producing the right type and size is contingent upon your water source temperature, the fertilizers used, whether or not you inoculate your pond with zooplankton, and which Morone

78

Fig. 3.3. Striped bass fry being prepared for shipping. (Photo by R.M. Harrell)

you are rearing. Geiger and Turner (1990) and Brewer and Rees (1990) provide a review of these considerations, and with the exception of Ludwig and Tackett (1991) and Daniels and Boyd (1993) who examined the use of differing organic and inorganic fertilizers, very little information has been added since these publications. The research by Ludwig and Tackett (1991) also includes a cost estimate of production of fingerlings using rice bran and cotton seed meal as the organic fertilizers. Fish reared in ponds fertilized with rice bran were significantly larger and cost considerably less to produce compared to fish reared in ponds with cotton seed meal. A special consideration should be made if you are rearing larvae of white bass, white perch, or yellow bass (M. mississippiensis). Larvae from these females are considerably smaller than striped bass (--3.5 vs 5 mm at hatch), and consequently have much smaller mouth gapes. Thus, while the strategy for zooplankton succession for striped bass is to proceed quickly through rotifers or ciliates into cladocerans and copepods, with the progeny from the other Morone it is crucial to maintain the rotifer and ciliate phase much longer to afford the larvae time to grow large enough to consume first instar copepod and cladoceran larvae. This is very difficult in ponds that have been filled with surface water from rivers or reservoirs. Conversely, ponds filled with ground water afford much better manipulation, but they may also require zooplankton inoculation. 3.2.3 Stocking Fry For Phase I Production Striped bass and hybrid fry are light sensitive and become stressed in bright sunlight (Rees and Cook, 1985). Therefore it is better if fry are stocked just after sunset when the oxygen levels are at their highest level in the pond and temperatures are beginning to cool for the night. A few culturists prefer to stock in the early morning before sunrise to take advantage of the coolest water of the day and lower pH values. In either case, do not stock in direct sunlight. Most producers prefer to stock fry into ponds just as they change from a prolarvae to a larvae and begin exogenous feeding (---day 5 post-hatch (PH)). With white bass this would be around day four as their mouth parts usually form a day earlier than striped bass at the same temperature.

79

If the producer wishes to determine the percentage of swimbladder inflation of his fry, he will have to feed them for at least a couple of days as the swimbladder usually inflates around day 7 PH at 20~ This would mean the producer would have to either hold and feed the larvae in tanks until swimbladder inflation could be determined, or delay having the hatchery producer ship the fish until the inflation rate is known. Delaying shipping until the fish are 7§ days PH would require the hatchery producer to set up a feeding strategy, which would add to the cost of larval seed stock. It is still unknown what factors contribute to swimbladder inflation success or failure. In an effort to minimize handling within the hatchery complex and equate early larval survial in nature to pond production, Harrell (1985) determined that survival, growth, and total number of fish harvested was two to three times better in larvae stocked in ponds at one day PH (24*) compared with sibling larvae stocked at 7 days PH. This finding was significant from the perspective that if swimbladder inflation is a function of surface area and surface tension, then fry stocked in ponds have more surface area and less surface tension than a tank with oily surfaces from the breakdown of dead larvae (Rees and Harrell, 1990; Peterson, 1991). Stocking densiy is the one area where considerations for food fish production differs from population enhancement efforts. In general, the goal of population enhancement programs is to in a 30 to 60 day season produce as great a quantity of the largest fish possible to meet management objectives of stocking a certain number of fish per hectare of a reservoir or number of fish per mile of a river. Therefore phase I fingerling ponds are stocked at very high densities and harvested at a smaller size than in food-fish grow out operations. Some of the numbers of striped bass larvae stocked in enhancement production ponds range from ---125,000 to 1,500,000 per hectare in freshwater and brackish water ponds, with harvest sizes of 450 to 3500 fish per kilogram (Smith, 1988; Brewer and Rees, 1990). Fingerling buyers for grow out want as large a fingerling as possible so that the fish can reach market size quicker. They :.lso want a uniform size so that feed sizes do not have to be mixed. In meeting this demand, producers can expect a premium price, while at the same time providing a much better product to the buyer. There are often price breaks for quantity and established customers. Therefore, the low end stocking rates for population enhancement ponds equates to the high end stocking rates for maximizing growth in foodfish production phase I ponds. In other words, stocking between 75,000 and 150,000 larvae per hectare should yield harvest ranges in the 100 to 300 fish per kilogram range (7 to 10 cm fish) in 30-60 days. Not only will the fish be larger, but survival rates will be higher. The buyer will be getting fish that handle shipping better, convert to artificial diets better, and have the capability of approaching 100-150 g at the end of the first growing season. This in turn relates to market fish in the second growing season approaching 1 kg, which bring a premium price (Tony Mazzacarro, Hyrock Fish Farms, Princess Anne, MD, personal communication). The other choice is to stock the ponds at a higher density, harvest the fish earlier, bring them in a hatchery building into tanks, grade the fish, and get them on artificial feed (Mike Freeze, Keo Fish Farm, Inc., Lonoke, AR, personal communication), then sell or restock ponds with the larger fish. The 30-60 day phase I season is a limiting factor because this is the time and size range when the fish are primarily planktivorous in their feeding behavior. Toward the end of phase I production when they start reaching 40-50 mm in size, they begin to become piscivorous in nature. When this happens, unless supplemental feeding is initiated (Fitzmayer et al., 1986), cannibalism can cause tremendous production losses in the ponds. It is at this time when the fish should be harvested. A good observational signal to the fingerling producer is to observe the edges of the ponds. When the fish begin "running the edges" there's a good chance they have exhausted their primary food source in the pond and are either looking for additional sources of food or staying in the shallows to escape larger siblings in the pond.

80

3.2.4 Feeds and Fee ring Phase I Fish As a routine ~ractice in enhancement production ponds, phase I fish are usually harvested without initiating artificial fi eding. However, since this phase of production is just the first stage of grow out to market-sized fish, m~ ay commercial fingerling producers start providing small amounts of feed to the pond beginning the secor t or third week of production. While there is still debate as to whether this is an economical approach one cannot deny the fact that starter fish meal is an excellent supplemental fertilizer high in nitrogen, and it c aes help maintain phytoplankton and zooplankton blooms (Fitzmayer et al., 1986). Routinely 2 to 7 Kg c f feed per hectare is added on a daily basis at least twice a day (Parker and Geiger, 1984; see Chapter 9 "~r m( re detailed information on feeds and feeding). In some instances, once some of the fish begin to accept the artificial feed, the schooling response of fish in the pond helps increase the percentage of those fish in the pond accepting the diet. One of the problems associated with this practice is that the larger, more aggressive fish tend to "hang-out" below where the other fish are being fed and selectively prey on the smaller fish. This is one argument for harvesting when plankton in the pond "crashes," and then grading the fish in the hatchery before initiating a feeding protocol. 3.2.5 Harvesting Phase I Fish Phase I striped bass ponds are generally specifically designed specialized ponds that are constructed to facilitate fertilizing and harvesting, especially those ponds operated by government agencies (Figures 3.4, 5.4). Enhancement production ponds are generally one quarter of a hectare to rarely one hectare in size. Because phase I ponds for food-fish operations are also used for phase II production, they are often in the half to two hectare size range (see Chapter 5).

Fig. 3.4. Typical pond layout for phase I and phase II production. (Photo by South Carolina Department of Natural Resources).

81

Fig. 3.5. Top: Seining harvest kettle for phase I striped bass, Bottom: preparing to put harvested fish in bucket for transfer to transport unit. (Photo by R.M. Harrell). In ponds specifically designed for fingerling production, the producer usually has the ability to rapidly dewater the pond toward one end. Normally a harvest structure (kettle) or catch basin is present to facilitate harvest (Figures 3.5 and 5.4) and supplemental water is added to provide fresh oxygen and cool water (Warren et al., 1990). Harvesting is usually conducted with fine mesh seines either seining the harvest kettle or with seines with mud lines slowly hauled through the catch basin area (Figure 3.5). Fish are collected in buckets with water and placed on a transport unit for hauling either to the hatchery or directly for stocking into the body of water scheduled to receive the fish (Figure 3.5).

82

3.3 PHASE II PRODUCTION The definition of phase II striped bass and hybrids is an arbitrary term that encompasses fingerlings from the time of phase I harvest until they are one year old. The size range is generally considered to be in the 75-250 mm range. Ponds are usually stocked at densities much lower than the larval to fingerling phase I ponds and much greater than the phase III, or market growout. In general, growth usually occurs from June through October or November, depending on site location and local weather conditions. Phase II fish are generally healthy enough to overwinter in ponds, although they rarely feed until early spring when the pond temperatures begin to rise above 7-8~ Excellent publications are available on the culture techniques for phase II fingerling grow out and include Kerby et al., (1987), Jenkins et al., (1989), Smith et al., (1990), and Kelly and Kohler (1996). When comparing hybrids to striped bass during phase II production, the heterotic advantage of the hybrid begins to become more evident. The hybrids develop a deeper body, express a more rapid growth rate, better food conversion, and handle better. 3.3.1 Fingerling Availability While most grow-out producers purchase phase I fish and stock them in their ponds for phase II and phase III grow out, some producers market phase II fish as well. These larger, advanced fingerlings are sold primarily to governmental agencies for enhancement stocking or net-pen operations. Some commercial pond producers who have had some die-off during the first year of growth will purchase these advanced fingerlings to replenish their stocks. In general, a buyer can expect to pay about $0.04-0.05 per centimeter for these fish if they are purchased in the fall of the year. There is little difference in price between striped bass and hybrids, however, if the producer overwinters the fish and the buyer purchases them the following spring, he can expect to pay a slightly higher price because the producer has both more time and risk involved in the production. Advanced fingerlings can be purchased anytime of the year, however most buyers prefer not to purchase fingerlings during summer months due to the risk in transport and handling losses (see Chapter 7). 3.3.2 Pond Preparation In comparison to phase I production, little pond preparation is needed for phase II production (Smith et al., 1990). Fertilizers are not used in phase II production as feeds and fish wastes contribute significant amounts of nitrogen and phosphorous to the ponds to keep phytoplankton blooms viable. Because the fish are being fed an artificial diet, there is no need to stimulate zooplankton population growth. In fact, excess plankton or algal growth can become a problem if it results in diel oxygen and pH fluctuations outside the optimal range for fish growth. However, the culturist wants a sufficient plankton bloom to shade out submerged aquatic vegetation. 3.3.3 Stocking Phase I For Phase II Grow Out Stocking rates for phase II fish are similar to phase I operations from the sense of matching stocking rates with the intended purpose of the final product. If the purpose for stocking is population enhancement at harvest, then stocking rates are quite dense and the fish size at harvest is smaller. If however, the harvest is intended for food-fish grow out then the stocking rates are considerably less. In general terms, for every hectare of a phase II operation a producer can expect to supply three hectares of a phase III operation.

83

Stocking densities for phase II production range from 10,000-250,000/ha, however densities in the range of 25,000 to 60,000/ha have resulted in more uniform fish and better overall survival (Jenkins et al., 1989). Densities greater than 60,000haa result in a greater biomass but will probably require excess amounts of feed, supplemental aeration, water exchange, and close monitoring of the fish's overall health. Grow-out producers who purchase phase I fish as their seed stock are quite sensitive to market demand of their final product. In other words, the food-fish market demand in size offish dictates the stocking strategy of the phase I buyers. If the market is looking for a wide size range of product (i.e., 0.5 to 1Kg fish) the phase I buyer will not be as picky about the size of the fish and uniformity as if the market demand was closer to 0.8 to 1Kg fish. In the former case, the buyer would probably settle for phase I fish in the 450 to 1000/Kg range and stock at densities as high as 50,000/ha. However, in the latter case, where the market demand is for the larger fish, the buyer would purchase fish no more than 650/Kg and stock his phase II ponds at densities around 20,000 to 30,000/ha. The goal in the latter case would be to produce as uniform fish as possible in the 100 to 150 g range at the end of the first year. 3.3.4 Feeds and Feeding Phase II Fish One of the keys to the success of culturing phase II fish is getting as much growth during that first growing season as possible. To accomplish this task, excellent water quality and an aggressive feeding schedule are required. The fish should be fed at least twice a day with a good high quality feed (see Chapter 9). Most grow-out operations are too large to feed by hand, thus mechanical blower-operated feeders and bulk food storage bins are part of the operation (Figures 3.6, 5.8). In net-pen operations or those operations where the fish are used for population enhancement efforts, simpler feeders such as demand or automatic feeders may be the best choice (Figures 3.7, 5.7). High quality feeds are important for success in striped bass production and, while complete diets are not yet available for Morone, diets high in protein (36-50%) and fat (10-16%) have proven successful (Smith et al., 1990; see Chapter 9). Most Morone diets are formulated similar to trout and salmon diets, and the trend in production operations for grow-out has shifted over the past few years from a sinking to a floating feed. Producers prefer the floating feed because they can observe their fish feeding, and observation of feeding behavior is paramount to determining the general health of your stock. If fish go offfeed, it is a sure indication that something is wrong either with the fish themselves (i.e., disease, see Chapters 10 and 11), the water quality (see Chapters 4 and 5), or the feed (see Chapter 9). In any case, immediate action is prudent. Sinking feed is the feed of choice during winter months when feeding has slowed down because you do not want the fish to rise to the surface and be exposed to dramatic changes in water temperatures where the surface water could be several degrees C lower than deeper pond water. A trade-off exists between biological maximum growth and economic maximum growth. Commercial producers have to be aware of these two maxima and how one influences the other. Obviously when you surpass the economic maximal growth you diminish your return on your investment (see Chapter 12). Biological maximal growth not only addresses the efficiency of food conversion for the fish, which interplays with the economic maximal growth, but it also impacts water quality. Overfeeding fish in a pond can have a drastic affect on the success of the operation because when you exceed that "breaking point" the end result is catastrophic losses of fish due to oxygen depletion, toxic algal blooms, and/or metabolite toxicity (i.e., ammonia or nitrite). To prevent these water quality losses from occurring, the producer must have the ability to supplementally aerate or flush with new water, or both (Figure 3.8). Operation of an aeration system or pumps to flush water requires energy and that is a cost to production. Smart producers work out "off-peak" rate deals with their local utility companies and operate during that time period. Fortunately, these "off-peak" times correspond most often with the times the mitigation efforts are required, such as late at night and early

84

Fig. 3.6. Top: A gas-powered blower feeder with hopper for feeding small-sized feeds for phase II fish. Bottom" Bulk storage bins for fish feed and PTO driven emergency aerators. (Photos by R.M. Harrell).

85

Fig. 3.7. Top: A demand feeder for net-pen culture of phase II and phase III fish. Bottom: An automatic feeder for feeding fish in ponds for grow-out. (Photos by D.W. Webster (top) and J.H. Kerby (bottom)).

86

Fig. 3.8. Aerating and flushing water in striped bass production ponds. (Photo by R.M. Harrell).

87

morning hours. At other times, even flushing and aeration may not be sufficient to prevent oxygen depletions and emergency aeration is required. Emergency aerators are usually portable devices that are driven off the power take-off (PTO) on tractors (see Figure 3.6). For the first month after stocking phase II fish are generally fed 15-25% of their body weight per day allocated in two separate feedings -- one in the early morning just after sunrise and one in late afternoon. After the first month, feed rates can be decreased monthly until the fish are being fed approximately 3% of their body weight per day (Table 3.1; Smith et al., 1990). Table 3.1. Suggested feed sizes and feed rates for differing size phase II Morone ~. Fish weight (g)

Total length (mm)

Feed size

Feed rate (% Biomass)

>_0.4

30

#2

25.0

>--0.8

44

#3

15.0

>__1.5

51

#4

10.0

>--9.0

90

>_25

3/32"

7.5

127

1/8"

5.0

>_75

192

5/32"

3.0

>-_150

238

3/16"

3.0

~From Smith et al., (1990)

3.3.5 Harvesting Phase II Fish Harvesting small (_<1 ha) ponds can be accomplished by draining and seining the fish in a manner similar to that of phase I fish. This practice is fairly common with fish used for enhancement purposes, where the fish are loaded directly onto a transport unit and taken to the water system of choice and released. However, when the phase II fish are going to be restocked into larger ponds for phase III grow out, a different approach is becoming common practice. Larger ponds are too expensive to completely drain and harvest at one time. They contain so many fish that harvesting and restocking becomes a materials handling problem. To solve this, many phase II and grow-out producers have starting using large haul seines similar to those used by the catfish industry and auger or fish loading pumps to move fish between ponds (Figures 3.9, 3.10, 5.6, 7.1). In addition to facilitating harvest, these pumps can be connected to graders that will automatically sort the fish by preset sizes and distribute them to the grow-out ponds while returning the culls to the pond being harvested (Figure 3.11). This automation greatly relieves stress in the fish thereby improving survival rates. It also helps the producer in feed management in that the fish are all of a uniform size and should not require mixing feeds. These automation systems are usually rented from contract harvesters or are available from some feed manufacturers as a service to their customers.

88

Fig. 3.9. Harvesting large production ponds with haul seines. Top: deploying net from reel. Bottom: working the net back into the reel. (Photo by, Top: D.W. Webster, Bottom" D.W. Meritt).

89

Fig. 3.10. Deployment and operation of a fish auger pump harvesting phase II striped bass hybrids for redistribution into phase III grow-out ponds. Notice pipe in pond for returning undersized fish. (Photos by, D.W. Webster)

90

Fig. 3.11. Operation of an auger fish pump (top) connected to automatic sorter and grader (bottom) for distribution of graded fish to grow-out ponds. (Photos by D.W. Webster).

91

3.4 PHASE IIl PRODUCTION

Phase III production is rare in enhancement production efforts thus most of the information that is available in the scientific literature is on experimental grow-out for research purposes or for the development of broodstock programs (Kerby et al., 1983, 1987; Smith, 1988; Smith et al., 1989, 1990). Most of the information that is available on actual production efforts in ponds comes from the industry itself and is not exactly considered proprietary, nor is it is well documented. Phase III grow out is essentially the second year production of striped bass and its hybrids to a market size fish. Almost always the fish are harvested before the beginning of the third growing season and the ponds are prepared to receive a new crop of phase II fish to repeat the cycle. Production has to be conducted at intensive levels to make it economically feasible (Smith et al., 1990). Success, therefore, is going to be dependent on the producers' ability to manage for good water quality, provide good quality feed on a daily basis, minimize losses from predators such as birds, otters, alligators, etc., and good harvest and marketing strategies. All too often the novice producer may be an excellent culturist, but fails to undertake adequate marketing efforts to get the best price for his crop and move that crop in a timely fashion. It is a serious mistake to think that just because you have a quality product that the wholesale and retail industry is going to "beat a path to your door." Marketing is just as important as the biological and economic management of your operation (see Chapters 12 and 13). 3.4.1 Stocking Densities for Phase III Fish Production ponds for final grow out to market size are usually considerably larger than phase I and phase II ponds. While the range in size is between 0.5 and 4 ha, the majority of the production ponds are between 2 and 2.5 ha. As with phase II ponds, there is no additional fertilization required as food and fish wastes are sufficient to provide nutrients for plankton production. Deciding at what level to stock phase III ponds is contingent on market demand, pond design and size, harvesting strategies, feeding rates, the availability of aeration, and the ability to flush with clean water when necessary (Smith et al., 1990). Because the current trend is for fish that yield a fillet of approximately 170 g, fish will have to be in the 700 to 900 g range. To achieve this target harvest size, the producer is going to have to plan on a loading capacity between 5,600 and 6,600 Kg/ha. When back calculating the number of phase II fish needed for stocking assuming a 90% survival from stocking to harvest, the producer should be stocking approximately 8,600 fish/ha. 3.4.2 Feeds and Feeding Phase III Fish Feeding strategies for phase III grow out is an extension of phase II production -- it just requires a greater bulk of feed and larger feeding equipment. As with phase II production, a good quality diet containing at least 38~ protein and 10% fat appears to be what is needed at this stage of our nutritional knowledge (see Chapter 9 for more details on nutritional requirements). Fish should be fed twice daily between 2% and 3% body weight. The feed allocation should be divided into at least two feedings per day with half in the morning and half the ration in the evening. Feeding in phase III ponds is also most often done with blower operated feeders that are powered either by a gasoline engine or off the tractor's PTO (Figure 3.12). Ideally, the afternoon feeding should be around 4:00 to 5:00 PM when the oxygen level is still around its daily peak. The reason for this early afternoon feeding is because late in the production season when the pond biomass is nearing its maximum and the amount of feed is nearing or surpassing the pond's ability to

92

Fig. 3.12. Tractor operated feed blowers for phase III production. The tractors and the hoppers are driven underneath the feed bins and the feed is discharged into the hopper for pond distribution. (Photos by R.M. Harrell)

93

assimilate all this organic material even with supplemental aeration, feeding later in the day will only stress the oxygen demand of the system that much more. When striped bass feed there is an almost immediate rise in metabolic oxygen demand that requires a minimum of 10-12 hours to return to basal rates. If fish are fed late in the evening after photosynthetic activity has shut down and pond respiration is increasing, the pond culturist runs a serious risk of depleting oxygen reserves before sunrise when photosynthesis starts again. Therefore it is prudent to feed when the daily dissolved oxygen is highest so by the time the photosynthetic production stops, the fish's metabolism will already be in a declining stage and not overly tax the total biological oxygen demand on the pond system. Unfortunately, the fish do not always cooperate and feed most actively during the peak oxygen times of the day because the sun is still high in the sky. Normally M o r o n e are most active in their feeding just before daylight and just after sunset. This is a management decision that the producer must make and it requires knowing what your pond's oxygen demand is under normal conditions and what supplemental and emergency aeration capabilities are available to make it through the night. A general rule of thumb is not to exceed a feeding rate of 250 Kg/ha on a daily basis even with supplemental aeration. Exceeding this level may be the crucial deciding factor in increasing return on your investment but, just like any form of gambling, you may end up with nothing. This is just another one ofthose management decisions that have to be made. 3.4.3 Harvesting Phase III Fish Harvesting phase III fish, like that of phase II fish, is both a materials handling problem and a timing problem. Most grow-out operations do not have the facilities to completely dewater a pond and hold the harvest in tanks until the fish can be sold. Just the shear number of fish in a 2 to 4 ha pond limits the feasibility of using the tank holding approach. Therefore, most pond culturists partially harvest their ponds on a weekly or biweekly basis using selective seines and live cars (Figures 3.9 and 3.13). Haul seines are pulled through the pond and fish are crowded into live cars or the fish in the holding net are harvested, usually with a boom operated net (Figures 3.13 (top) and 5.5). Once in the boom net, the fish are either loaded on a hauling truck for transport to a processing plant, or they are quickly killed with an ice brine and/or electrical shock then individually graded and sorted into shipping containers (Figure 3.13 bottom) at about 23 kg/box. Fish are usually graded in quarter pound (---115 g) lots (see Chapter 13). Once the fish are graded and loaded, they are then delivered to the buyer or picked-up by the buyer at the farm, depending on your prior arrangement. Many M o r o n e are sold on the live fish market for specialty restaurants and ethnic markets (see Chapters 1 and 3 for market information). It is critical for the producer to expedite the harvest as quickly as possible and minimize damage to the fish by bruising, cuts, or puncture wounds. Any blemishes could potentially impact the market value of the fish. One advantage to partial harvest of a pond is that you are selectively removing the larger fish and reducing the total pond biomass at a time when the pond' s biomass could be on the "edge" of causing a major loss due to management's inability to maintain water quality. It also provides an opportunity for the smaller fish to express compensatory growth when the larger more aggressive competitors have been removed from the pond. In any case, lowering the pond biomass by partial harvest should lower the biological oxygen demand stress on the pond thereby lowering the potential for catastrophic loss.

94

Fig. 3.13. Top: Harvesting market-size fish with a boom net with live car in background holding surplus fish. Bottom: Hand grading fish into shipping boxes prior to being iced down. (Photos By, Top: D.W. Meritt, Bottom, J.H. Kerby)

95

3.5 S U M M A R Y Pond production for striped bass and Morone hybrids has its roots in the population enhancement programs of the various governmental agencies for natural bodies of water. The specifics of techniques and culture requirements are available in a variety of different references, in particular Harrell et al., (1990), and are found throughout the entirety of this book. Most of our knowledge of phase III production comes from the industry itself and has not been significantly detailed in any specific format. Some ofthe subtle nuances were covered in this chapter but are not intended to be a replacement for site specificity as each operation has its own unique way of producing fish. Thus there are not absolutes in market production of fish and the producers have to rely on intuition and trial and error. Growth of the industry and continued expansion of the research community into the area of production scale efforts may someday soon afford industry standardization as much as the animal and the environment will allow.

96

References

Brewer, D.L. and Rees, R.A., 1990. Pond culture of phase I striped bass fingerlings. Pages 99-120 in R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southem Division, American Fisheries Society, Bethesda, Maryland. Daniels, H.V. and Boyd, C.E., 1993. Nitrogen, phosphorous, and silica fertilization of brackish water ponds. J. Aqua. Trop., 8" 103-110. Fitzmayer, K.M., Broach, J.I. and Estes, R.D., 1986. Effects of supplemental feeding on growth, production, and feeding habits of striped bass in ponds. The Progressive Fish-Culturist, 48:18-24. Geiger, J.G., 1983a. Zooplankton production and manipulation in striped bass rearing ponds. Aquaculture, 35:331-351. Geiger, J.G., 1983b. A review of pond zooplankton production and fertilization for the culture of larval and fingerling striped bass. Aquaculture, 35" 353-369. Geiger, J.G. and Turner, C.J., 1990. Pond fertilization and zooplankton management techniques for production of fingerling striped bass and hybrid striped bass. Pages 79-98 in R..M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society,, Bethesda, Maryland. Geiger, J.G., Turner, C.J., Fitzmayer, K.M. and Nichols, W.C., 1985. Feeding habits of larval and fingerling striped bass and zooplankton dynamics in fertilized rearing ponds. The Progressive Fish-Culturist, 47:213-223. Harrell, R.M., 1985. Survival and production of one- and seven-day-old striped bass larvae in hatchery ponds. Journal of the World Mariculture Society, 16: 82-86. Harrell, R.M. and Bukowski, J., 1990. The culture, zooplankton dynamics and predator-prey interactions of Chesapeake Bay striped bass, Morone saxatilis (Walbaum), in estuarine ponds. Aquaculture and Fisheries Management, 21 195-212. Harrell, R.M., Kerby, J.H. and Minton, R.V., 1990. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, Maryland. Houde, E.D. and Lubbers, III, L., 1986. Survival and growth of striped bass, Morone saxatilis, and Morone hybrid larvae: laboratory and pond enclosure experiments. Fishery Bulletin, 84" 905-914. Jenkins, W.E., Smith, T.I.J., Stokes, A.D. and Smiley, R.A., 1989. Effect of stocking density on production of advanced juvenile hybrid striped bass. Proceedings of the Annual Conference Southeastern Association of Fish and Wildlife Agencies, 42: 56-65. Kelly, A.M. and Kohler, C.C., 1996. Sunshine bass performance in ponds, cages, and indoor tanks. The Progressive Fish-Culturist, 58: 55-58. Kerby, J.H., Hinshaw, J.M. and Huish, M.T., 1987. Increased growth and production of striped bass x white bass hybrids in earthen ponds. Journal of the World Aquaculture Society, 18: 26-34. Kerby, J.H., Woods, III, L.C. and Huish, M.T., 1983. Pond culture of striped bass x white bass hybrids. Journal of the World Mariculture Society, 14:613-623. Ludwig, G.M. and Tackett, D.L., 1991. Effects of using rice bran and cotton seed meal as organic fertilizers on water quality, plankton, and growth and yield of striped bass, Morone saxatilis, fingerlings in ponds. Journal of Applied Aquaculture, 1:79-94.

97

Minton, R.V. and Harrell, R.M., 1990. The culture of striped bass and hybrids in brackish water. Pages243-251 in R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, Maryland. Parker, N.C. and Geiger, J.G., 1984. Production methods for striped bass. Pages 106-118 in H.K. Dupree and J.V. Huner, editors. Third report to the fish farmers: the status of warm-water fish farming and progress in fish farming research. U. S. Fish and Wildlife Service, Washington, D.C. Parker, N.C., Klar, G.T., Smith, T.I.J. and Kerby, J.H., 1990. Special considerations in the culture of striped bass and striped bass hybrids. Pages 191-215 in R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, Maryland. Parmley, D.C. and Geiger, J.G., 1985. Succession patterns of zooplankton in fertilized culture ponds without fish. The Progressive Fish-Culturist, 47:183-186. Peterson, R.H., 1991. Consideration for design of culture facilities for early stages of striped bass (Morone saxatilis). Bulletin of the Aquaculture Association of Canada, 91(3): 86-88. Rees, R.A. and Cook, S.F., 1985. Effect of sunlight intensity on survival of striped bass x white bass fry. Proceedings of the Annual Conference Southeastern Association of Fish and Wildlife Agencies, 36:83-94 Rees, R.A. and Harrell, R.M., 1990. Artificial spawning aadn fry production of striped bass and hybrids. Pages 43-72 /n R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, Maryland. Smith, J.M. and Whitehurst, D.K., 1990. Tank spawning methodology for the production of striped bass. Pages 73-77 /n R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, Maryland. Smith, T.I.J., 1988. Aquaculture of striped bass and its hybrids in North America. Aquaculture Magazine, 14(1): 449. Smith, T.I.J., Jenkins, W.E. and Minton, R.V., 1990. Production of advanced fingerling and subadult striped bass and striped bass hybrids in earthen ponds. Pages 121-139 in R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Smith, T.I.J., Jenkins, W.E., Stokes, A.D. and Smiley, R.A., 1989. Semi-intensive pond production of market-size striped bass (Morone saxatilis) x white bass (M. chrysops) hybrids. World Aquaculture, 20(1): 81-83. Tuncer, J., Harrell, R.M. and Houde, E.D., 1990. Comparative energetics of striped bass (Morone saxatilis) and hybrid (M. saxatilis x M. chrysops) juveniles. Aquaculture, 86: 387-400. Warren, H.J., Harrell, R.M., Geiger, J.G. and Rees, R.A., 1990. Design of rearing facilities for striped bass and hybrid striped bass. Pages 17-27 in R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Woods, III, L.C., Lockwood, J.C., Kerby, J.H. and Huish, M.T., 1985. Feeding ecology of hybrid striped bass (Morone saxatilis x M. chrysops) in culture ponds. Journal of the World Mariculture Society, 16L 71-81.

This Page Intentionally Left Blank

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

99

Chapter 4

Water Quality Dynam&s as the Basis for Aquaculture @stem Design David E. Brune 4.1 INTRODUCTION The primary task facing the fish culturist is to provide an optimal environment to insure maximum growth and reproduction of the aquatic animal of interest while minimizing stress effects. Tomasso, in Chapter 10 of this volume, lists water temperature, dissolved oxygen concentration, water hardness, pH, light levels, ammonia, nitrate, nitrite and suspended solids concentrations as the factors of greatest importance in affecting the culture of striped bass and its hybrids. The design of a successful aquaculture system is heavily dependent on our ability to make accurate predictions ofthe dynamics ofthese important water quality parameters as affected by operator controlled, fish stocking levels and feeding intensity. It is the intent of this paper to demonstrate that relatively simple chemical, physical and biological models can be used to make reasonable long-term predictions of the dynamics of the water quality parameters of importance in recirculating aquaculture systems, as well as, fish culture ponds. The approach used to design a particular aquaculture system generally falls into one of three catagories. First, there is the cookbook approach; this is used for aquaculture system designs in those cases where the behavior of the system is poorly quantified, or so complex, that the system behavior cannot be predicted well enough to insure success of any deviation from past designs. An example of this type of system is the complex technique of pond fertilization, followed sequentially by algal culture, zooplankton culture, and fish rearing which is typical of the current methodology for striped bass fingerling production. The second type of approach is the use of semi-empirical models. In this approach a sufficient data base exists to allow for reasonable predictions of system performance based on correlations between fish production and water quality parameters. Typical of this approach are the models used for prediction of pond dissolved oxygen by Boyd (1979, 1991) and others (Busch et al., 1974) in pond catfish production. The third approach is sometimes referred to as the mechanistic approach. In this latter case, a mass and energy balance is used to represent the nutrient and energy flux in an aquaculture system. This approach is most successfully applied to the design of intensive recirculating aquaculture systems, and is beginning to be used for pond aquaculture systems (Smith 1988; Piedrahita 1991; Brune 1994). This chapter will offer an application of a mixture of these three techniques. A simplified mechanistic representation of an aquaculture system will be used to predict values for the controlling water quality parameters of dissolved oxygen, water pH, nitrogen concentration, and pond algal biomass concentration and rate of photosynthesis. The empirical approach is used to define "typical" field operating conditions and the limits to system performance. Finally, predictions of system behavior based on this field calibrated simplistic model is compared to past "cookbook" based system performance. 4.2 WATER REUSE SYSTEMS The primary design criterions and forcing functions for a water reuse system are fish oxygen consumption rate and the feeding rate. These rates are species and individual fish weight dependent, varying from 0.4 to 10% of body weight fed per day, and 0.5 to 4 g of oxygen per 100 g of body weight per day.

100

Specific values of feed rate and oxygen uptake are available from a number of sources (Piper et al., 1982; Andrews et al., 1975). Furthermore, important water quality parameters, such as CO2, pH, and ammonia levels can be tied to these forcing functions. Generally, 20-25% of feed nitrogen is removed by the fish (Brune and Gunther, 1981; Colt 1986), with typically 60% of feed nitrogen being returned to the culture system as ammonia, with the remaining 15-20% trapped in waste solids. If this waste is not removed rapidly from the system (< 1 day), the solids organic nitrogen will be returned to the aquatic environment as ammonia nitrogen. A useful rule of thumb for calculating the impact of respiration on system pH is to assume that one mole of carbon dioxide will be produced per mole of oxygen consumed (Colt, 1986). Release of ammonia nitrogen into the aquaculture system can further impacts the oxygen concentration and pH as a result of the bacterial process of nitrification. The theoretical balance (Wheaton et al., 1991) for this two step process is summarized as" _

NH 4" + 1.8302 + 1.98 HCO 3 (4.1) 0.021CSH702 N + 0.98NO 3 + 1.883CO 2 This process reduces the oxygen concentration through the process of ammonia oxidation. It affects the water pH by the destruction of alkalinity and production of CO_,. Field observations of the nitrification reactions suggest that 7.1 mg of alkalinity (as CaCO3) is neutralized per mg of NH4-N oxidized. In addition, 4.6 mg of 02 is consumed per mg of NH3 -N oxidized (US EPA, 1975). These forcing functions and relationships, combined with equations describing the rate of system oxygenation and carbonate equilibrium chemistry, can be used to predict system water quality dynamics at a desired fish loading and feeding rate in an aquaculture water reuse system. 4.3 POND AQUACULTURE SYSTEMS The primary driving force for a pond aquaculture system is the level of feed input. For the purpose of predicting system design capacity, the total oxygen demand imposed on the pond is assumed to equal the total ultimate oxygen demand (BODL) of the feed fed to the pond per day. A value of 0.65 mg of oxygen demand per mg of dry feed fed is a reasonable assumption for the purpose of design (Chieng et al., 1989). The addition of nitrogen and phosphorus to a natural water body (as fertilizer, fish food, or fish metabolites) drives the production of organic carbon fixation through the process of phytoplankton photosynthetic biomass production. The elemental composition of this biomass production can vary widely. This composition is often represented as a value referred to as the Redfield ratio (Redfield et al., 1963): 106CO 2 + 16NO 3- + HPO 4 + 122H20 § IgH* $ sunlight

(4.2)

C106 H263 O110 NI6 P1 + 13802 The forward process is referred to as photosynthesis, while the reverse represents the combined processes of respiration and algal cell mineralization. One would expect that since the forward reaction occurs only during the daylight hours and respiration occurs continuously, that a day/night polarization of both oxygen concentration and pH would occur. This is the well known observation of diurnal oxygen concentration

101

fluctuation and diumal pH cycling, resulting from daytime CO2 fixation, and oxygen production followed by nighttime CO2 production and oxygen uptake. The Redfield ratio suggests a algal composition of 35.8% carbon, 6.3% nitrogen and 0.87% phosphorus of algal dry weight. The Redfield ratio was determined from marine phytoplankton. Repeated measurements of freshwater algal biomass suggests slightly higher values of 45% carbon, 8% nitrogen and 1% phosphorus (Goldman, 1980). These latter values will be assumed to best represent the composition of freshwater pond phytoplankton for the purpose of all design calculations. These later values agree with the Redfield Ratio in suggesting an algal C/N ratio of 5.62/1. Furthermore, the Redfield equation suggests an oxygen production of 3.47 mg of O2 per mg of carbon fixed through photosynthesis. 4.3.1 Pond Primary Production Much has been written about the nutrients limiting primary production in freshwater ecosystems (Lehman et al., 1975). Many researchers suggest that phosphorus concentration controls the total algal biomass in most freshwater communities (Schindler, 1971). However, when attempting to predict the primary productivity of an aquaculture pond for design purposes, it is more practical and useful to base predicted rates of algal production on nitrogen loading. In most cases, the nitrogen content of the fish feed is well known, and feed phosphorus content is usually supplied at P/N ratios that are typically equal to, or greater than, the P/N ratio of the expected algal biomass. As a result, the sediment of most aquaculture ponds quickly becomes phosphorus rich, thereby ensuring adequate long-term phosphorus supply to support algal growth in the pond. On the other hand, excessive nitrogen applied to a pond will eventually migrate to the sediment where it will be lost through denitrification. Most importantly, predictions of algal biomass based on nitrogen addition from feeds with carbon, nitrogen, and phosphorus content similar to the C/N/P composition offish biomass correlate well with observations of pond algal biomass, as will be demonstrated in latter worked examples. Calculations of pond productivity, based on daily nitrogen loading rate, should be considered useful

averaged predictions. Day-to-day pond algal biomass can vary widely because of changes in short term nutrient availability, light levels, and zooplankton populations. However, the averaged values yield useful information in terms of predicting overall pond oxygenation requirements and fish carrying capacities. Algal growth rates as high as 2.0 doubling per day have been observed under laboratory and controlled field conditions (Shelef et al., 1973; Goldman and Ryther, 1975). Under such conditions sustained algal productivities in excess of 12 g carbon/m2-day have been observed. However, in the typical aquaculture pond, overall cell turnover is generally limited to maximum rates of 0.5 per day (i.e., a 2 day algal cell age) and a minimum of 0.1 per day (a 10 day algal cell age) with productivities ranging from 1 to 4 g carbon/m2-day (Brune, 1991; Schroeder et al., 1991). Typical values of cell age range from 5-10 days. Therefore in an aquaculture pond, the algal productivity will generally be controlled by availability of nitrogen mass loading up to the level that can sustain a fixation rate of 4 g carbon/m2-day. Typically, water quality parameters are measured and expressed in mg/liter, mmole/liter, or meq/liter, while most US aquaculturists use lbs/acre to describe pond fish carrying capacity and pond feed application rates. It is therefore useful to be able to convert from lb of feed fed to mg/liter of resulting nitrogen in the pond water, and for this reason these english/metric conversions will be incorporated into the rate expressions. Assuming that 75% of the nitrogen contained in the feed added to a pond will be incorporated into algal biomass, (i.e., no waste solids removal from the pond and 25% of fed nitrogen is incorporated into fish flesh) the daily algal fixation can be calculated by multiplying the feed application rate in lb/acre-day by the

102

percent protein contained in the feed and, further by multiplying by the 16% of the feed protein as nitrogen and finally, converting to units of g-N/m2-day: F

lb feed acre -day

p

r [.000134] = IN nitrogen loading g-N protein content m 2-d L percent feed

(4.4)

If the pond is assumed to average 1 meter in depth, then the rate of addition in g-N/m2-d is equivalent to mg-N/L, since 1 m 3 = 1000 liters. The carbon fixation rate can be calculated from the Redfield C/N ratio of 5.62/1: C

g -C m 2-day

= [N] [5.62]

(4.3)

4.3.2 Pond Oxygen Production Once the carbon fixation rate is established, the effect of oxygen production by the algal biomass can be calculated using the Redfield oxygen/carbon ratio of 3.47/1:

O 2

g-oxygen 2 m -day

= [C] [3.47]

(4.5)

In a one meter deep pond, g-oxygen/m2-day is equal to mg/L of 0 2. Total oxygen demand in a pond can also be tied to the feed rate. The ultimate biological oxygen demand of pelleted feeds of 0.65 mg O 2 per mg of feed can be taken to represent the combined fish oxygen demand as well as the pond oxygen demand due to uneaten food or excreted organic waste. The amount of gross oxygen production that results in a net oxygen gain in the pond depends on the average algal cell age. Any algal organic matter that is physically removed from the pond either directly or indirectly by incorporation into filter feeding zooplankton and/or fish, results in algal organic oxygen demand not exerted in the pond, and therefore, a net oxygen addition to the overall pond oxygen balance. As an example, if the pond is fixing a net 3 grn of carbon/m2-day, which is equivalent to 10.4 gm of oxygen/m 2-day with an average algal cell age of 5 days, then 20% of this production is being removed each day either by sedimentation to the pond bottom or incorporation into the pond food chain. Past observations of pond oxygen dynamics suggest that this relatively low rate of algal organic matter removal can be considered to be "lost" from the pond with regard to impact on oxygen dynamics. Therefore, this leads to the expectation of a net oxygen gain in the pond of(0.2) X (10.4 gm) or 2.1 gm oxygen/m2-day. This effect, if controllable, offers one of the greatest potentials to the pond aquaculturist to increase the carrying capacity of his pond. In ponds with unmanaged algal production, the long-term net oxygen yield varies between 10 and 20% of daily production (Brune et al., 1992; Schroeder et al., 1991). For simplicity, and as an added factor of safety in later calculations of pond oxygenation, these low levels of net pond oxygenation from unmanaged algal production will not be routinely included in pond oxygen balances.

103

4.3.3 Algal Standing Crop and Secchi Disk For the purpose of pond design and prediction of system performance, algal standing crop, best represented as total volatile suspended solids (TVSS) and pond total suspended solids (TSS) are the most difficult variables to predict in a aquaculture pond. Algal standing crop is a direct function of the average algal cell age and algal productivity. As indicated above, past observations suggest that this age typically ranges from 5 to 10 days depending on settling rates and zooplankton grazing rates (Brune et al., 1992). Average algal standing crop can be predicted by multiplying the daily carbon fixation rate by the average cell age: SC g-carbon

- [C] [01

(4.6)

where 0 varies from 5 to 10 days in a typical aquaculture pond. For design purposes, it is best to calculate the extremes of expected algal density as an estimation of the range of algal standing crop in the pond. The algal cell density as total volatile suspended (TVSS) solids is obtained by converting from g-carbon/m 2to mg/L of total volatile suspended solids using the ratio of 2.22 g of TVSS per g of carbon: (4.7) rVSS

mg

L

[SC] [2.22]

From the calculation of pond total volatile suspended solids, it is possible to predict average pond secchi disk (SDV in meters) from a correlation between total suspended solids and secchi disk (Almazan and Boyd, 1978) assuming that 100% of TSS is present as TVSS: TSS = 6.03 (SDV) -0"932

(4.8)

Drapcho (1993) later showed that at secchi disk readings of less than 0.5 meter, a more accurate representation of this relationship is: TSS = 11.65 + 297e(-$'22)(SDV)

(4.9)

In both equations 4.8 and 4.9, we are assuming the total volatile suspended solids is equal to total suspended solids as measured by the membrane filtration technique used by both Almazan and Boyd and later by Drapcho. In unmixed ponds this is usually a reasonable assumption, however, in extremely muddy ponds the ratio of TSS/TVSS may often times range as high as 2/1. This effect should be taken into consideration when attempting to predict algal biomass (TVSS) in muddy ponds from secchi disk readings which are correlated to total suspended solids (TSS).

104

4.4 DISSOLVED OXYGEN C O N C E N T R A T I O N 4.4.1 Equilibrium Gas Concentration At a given temperature and salinity, the concentration of oxygen that is soluble in water is a direct function of the partial pressure of gaseous oxygen in contact with the water: Cs

Po~ KH

=

(4.10)

This relationship, referred to as Henry's Law, expresses the equilibrium dissolved oxygen concentration (Cs) as a function of Henry's constant for oxygen (KH) and the contacting partial pressure of oxygen (Po2) . 4.4.2 Gas Transfer Rate If the water is not at equilibrium with regard to gas saturation, then there exists a driving force for oxygen to move into, or out of solution. This non-equilibrium driving force is quantitatively expressed as the oxygen deficit (D) or Cs-C where Cs is the equilibrium value given by Henry's Law and C is the actual solution concentration. The rate at which oxygen moves into solution is controlled by the oxygen transfer at the air water interface, schematically represented in Figure 4.1:

x GAS

PHASE ADVECTION PC,AS

I I GAS FILM

LIQUID FILM

I

I

I

I

WATER FILM

Cs

PHASE ADVECT,ON

CONCENTRATION

Fig. 4.1. Representation of gas/liquid interfacial transfer. The rate of change of oxygen in the bulk water can be described as a function of the interracial surface area (A), the diffusitivity of the gas (D) in the liquid of interest (oxygen in water in this case), the stagnant water film thickness (X) (the water film is the slower, therefore, the rate controlling layer), and the volume of water under the influence of the aeration device (V). The relationship is then expressed as: dc

t i m e r a , e o , change

of oxygen concentration

=

105

Usually, precise values of A, V, X and D are not well known for a particular situation. For this reason, the effects of these parameters are lumped together into a single empirically determined rate constant referred to as the overall gas transfer coefficient (KLa) where: de - KLa (Cs - C) dt where;

(4.12)

KLa = rate of oxygenation, time-X C = system oxygen concentration, m_._gg L C

s

= oxygen saturation concentration, mg L

dc dt

-

rate of change of system oxygen concentration,

mg L -time

This relationship is a first order exponential, illustrated in Figure 4.2:

z o_

CO

t-<~ r~ z

Cs

z 0 z w o X 0

TIME

Fig. 4.2. Exponential time rate of change of oxygen concentration predicted by equation 4.12. A particular value of KLa, which is specific for a given set of conditions, can be used to describe the rate of transfer of oxygen into or out of solution, as shown above. In a system in which fish or other oxygen demand is present, the KLa can be determined by measuring the steady state oxygen deficit (D) and the fish respiration rate (R). This is illustrated in Figure 4.3. With the aeration device on, and the system at steady state, the oxygen concentration in the water remains constant, with the respiration rate (R) equal to the oxygen replacement rate (KL~D). If the aerator is turned off at time equal to tl, the oxygen concentration will begin to fall at a rate equal to R. When the aerator is switched back on, the oxygen concentration will return to C in an exponential fashion described by KLa(CsC) -R. The overall transfer rate (KLa) can be determined from this simple experiment by measuring the rate of respiration and dividing by the driving force or deficit at steady state;

106

KLa

zl

(4.13)

=

Cs - C AERATION OFF

_o ~_. z ,,, (.3 z o o

AEFb~TION ON

(Cs-C)=D R=K

z bJ o>x o

LA

Cs

D LA ( C s - C ) - R

to

i

i

t 1

t 2

,

TIME

Fig. 4.3. Dynamic response of oxygen concentration with simultaneous respiration and aeration.

A detailed presentation that describes how to obtain the KL~value from operating systems is given in Mueller and Boyle (1988). Example 4.1 Kt~ Determination A 1000 L tank containing 100 kg of fish with a known respiration of 1.5 kg of 02 per 100 kg of fish per day must be maintained at an oxygen concentration of 5.0 mg/L. What overall transfer coefficient is required? Solution:

Assuming that dry air is used to aerate this tank at a water temperature of 70~

and a

pressure of 1 ATM pressure, the tank equilibrium saturation value for oxygen can be calculated from Henry's Law directly or obtained from previously calculated tabulated values. The total oxygen demand is: 1.5 kg O~

100 k S fish

100 kg fish day

1 day 24 hr

= 0.01736

C s @ 70~

1 hr

1 min

60 min

60 sec

mg

L -sec

_ dc

dt

and 1 atm pressure = 8.5 m___gg L

106 mg 1000 L

kg

107

With C = 5.0 m~jL, and Cs = 8.5 m g ~ , and dc/dt = 0.01736 mg/L-sec, the K La can be determined: dc dt - KLa (Cs - C)

K~a =

= KLa

(

) mg 8.5 - 5.0 m__gg = 0.0173 L L -sec

0.01736 m8

L

L-sec

3.5 mg

= 0.005 sec -1

4.4.3 Manufacturers Oxygenation Capacity In the case of a small tank, the volume of water under the influence of the aeration device is well known. In a pond however, this volume, referred to as the volume of influence of the aerator, is often not well known. Different aerator designs can produce considerably different oxygen mass transfer rates because of the mixing pattern produced by the device. Manufacturers will frequently rate a particular aeration system with an aeration capacity number which is usually determined by calculating the overall oxygen transfer in clean water at an oxygen concentration of zero and water temperature of 20~ and an assumed, or measured volume of influence (V): No

/

manufacturers

/

oxygenation capacity]

(4.14)

= KLa (Cs2~ - 0 ) V

Example 4.2 OxygenationCapacity The manufacturer lists an aerator as being capable of supplying 4.0 kg of 02 per Kw- h (No) at standard test conditions of water temperature = 20~ What is the field oxygenation capacity (Nf) for fish culture conditions with Cmm= 5.0 mg/L, and Cs = 9.02 mg/L?

Solution: (solving by using ratios),

N O = 4.0

kg - (9.14 - 0)V ~ kw -h

therefore: V o = 0.437

Nf -" (8.0 - 5.0)Vf assuming Vo = Vf Nf = (3.0)(0.437) = 1.31

kg kw -h

This is the rate of oxygenation that can be expected to be supplied to the pond from the aeration device per Kw-h of energy supplied to the aerator, assuming an average pond oxygen concentration of 5.0 mg/L, and a solution oxygen concentration of 9.02 mg/L.

108

4.4.4 Surface Reaeration The rate at which oxygen transfer occurs across the surface of an open body of water has been quantified by Banks and Herrera (1977) as a function of water depth and wind speed:

(4.15) where; KLa = transfer coefficient, day l base e V = wind speed, km/h measured 10 meters above water surface H = water depth, meters This equation was developed from data taken from shallow lakes and assumes that the zone of influence, in this case the entire water depth, is well mixed.

Example 4.3 Surface Reaeration If the early morning (6 AM) dissolved oxygen in a pond is 3.0 mg/L and saturation is 8.0 mg/L, what will the oxygen concentration in the pond be at noon assuming there is no respiration or photosynthesis in the pond?

Solution: Assuming the pond is 1 meter deep and the average wind speed = 9.65 km/h (6 mph) KLa = (1 meter) [(0.384(9.65) '/: - 0.088(9.65) + 0.0029(9.65)2)] KLa = (1)[( 1.193 - 0.849 + .27)] KL, = 0.614 day ~ Converting this value to a per hour rate: 0.614

day

day

24h

= 0.0256h-1

A table can be prepared to estimate the time dependent oxygen concentration by repeatedly solving equation 4.12 for small (1 h in this case) time steps (see Table 4.1). By noon, the oxygen concentration will have increased to 3.72 mg/L as a result of wind reaeration at a constant wind velocity of 9.65 km/h. This value (1.44 mg/L over 12 hrs) compares favorably with the observed values from Schroeder (1987) of 1.69 mg/L oxygen gain over a 12 hour period following a 50% an observed saturation oxygen concentration at dusk.

109

4.5 C A R B O N A T E E Q U I L I B R I U M AND P H One of the most important properties of water is its weak tendency to disassociate into its ionic components: H 2 0 -~- H +

OH-

(4.16)

The ratio of the ionic products to the concentration of water is expressed as the equilibrium constant for water (Kw): Kw = [H+][OH-] [H20]

_= 1 X 10 -~4 @ 25~

(4.17)

Table 4.1. Stepwise solution to reaeration equation. At

Time 6 AM

C

dc/dt = Kea(Cs - C) At

3.0 mg/L

1 hr

(0.0256 hr~)(8.0- 3.000)(1 h r ) = 0.128 mg/L 7 AM

3.000 + 0.128 - 3.128

1 hr

(0.0256)(8.0- 3.128)= 0.125 8 AM

3.128 + 0.125 = 3.253

1 hr

(0.0256)(8.0 - 3.253) = 0.122 9 AM

3.253 + 0.122 = 3.375

1 hr

(0.0256)(8.0- 3.375) = 0.118 10AM

3.375+0.118=3.493

1 hr

(0.0256)(8.0- 3.493)= 0.115 11 AM

3.493 + 0.115 = 3.608

1 hr

(0.0256)(8.0- 3.608) = 0.112 12 Noon

3.608 + 0.112 = 3.720

By convention, the concentration of water [H20] is set equal to 1, and the ion product ([I-F][OH]) in water is seen to vary over 14 orders of magnitude. For convenience, this concentration spectrum is defined as the water pH, where pH is expressed as: pH =

1 = -log [H +] log [H ~]

(4.18)

110

In most natural waters, the factor that exerts the greatest influence on pH is the concentration of inorganic carbon in the water. The presence of approximately 360 ppm CO2 in the atmosphere, and calcium and magnesium carbonate in most rocks and soils, insures that inorganic carbon is present to interact with water pH. Atmosphere CO2 dissolves in water to produce carbonic acid according to: [H2CO 3] H20 + CO 2 ~- H2 CO 3

[CO2]

= KCO'

(4.19)

Carbonic acid undergoes two additional equilibrium reactions:

H 2 CO 3 * H" + H CO a-

[HI[HCO3-]

= K~

(4.20)

[H2CO 3]

HCO; ~ H § + CO;

In "][co~:] [nco~-]

= K2

(4.21)

The end result is that the total inorganic carbon present in water at any time is the sum of the three forms:

C T = [H2CO31 + [HCO3- ] + [CO/]

(4.22)

The total carbon in solution is also related, through the principle of charge balance, to the titratable negative charges (or anions) in solution. A commonly used indication of the total titratable negative charge in solution is called the alkalinity (alk)of the water expressed in either equivalents or milliequivalent per liter (meq/L); [Alk] = C r(tzl + 2tz2) + [OH-] - [H+]

(4.23)

In equation 4.23 0~1and cz2represent the fraction of the total carbon present as (HCO-~ and (CO,respectively: [H2CO3 ] = Crtz ~

(4.24)

[HCO3- ] = C r cz~

(4.25)

[CO3-] = C r tz2

(4.26)

where: tz0 + 0~1 "1-

0~ 2 " -

1

(4.27)

These ionization fractions are a function of the solution pH, temperature, solution ionic strength, and equilibrium constants Kco, K I, and K 2 and can be calculated as der~crib~d in Appendix A, Figure A-2. Other anions such as boron 2(B(OH)T ), ammonium, (NH~), phosphate (PO4-3), or silicate (S iO4-) can affect

III

this relationship; However, in most fresh waters, these concentrations are low relative to the inorganic carbon concentration and can be ignored. For the striped bass culturist, the practical importance of these relationships is summarized in the graphical representation of Equations 4.18 - 4.22 in Figure 1, Appendix A. This figure from Stumm and Morgan 198 l, illustrates that the pH of most natural water can be quantitatively predicted from knowledge of the solution alkalinity and total inorganic carbon content. For example, if a water is at pH - 8.0 with an alkalinity = 2 meq/L (2 meq/L alkalinity = 100 mg/L as CaCO3), then as Figure 1 shows, the water would be expected to have an inorganic carbon content of approximately 2.1 mmole/L. If, as a result of algal photosynthesis, 1 mmole/L of inorganic carbon is taken up and fixed as organic carbon, then the resulting short-term pH can be predicted by tracing a horizontal line from the previous point of intersection in Figure 1 to 1.1 mmole/L of total carbon, yielding a solution pH of 10.35. It should be noted that eventually this pH will again fall as atmospheric CO2 moves back into the solution to return the water CO2(aq) concentration back to equilibrium as predicted by Henry's Law (at atmosphere CO levels the equilibrium pH at 2.0 meq/L alkalinity is approximately 8.3). If both the rate of algal carbon fixation and the K La(C O 2) in a pond or recirculating aquaculture system is known, then the time rate of change of the water pH can be quantitatively predicted. 4.6 PREDICTION OF AQUACULTURE POND PERFORMANCE AND WATER QUALITY The best way to test the validity of the equations and assumptions proposed in this chapter is to evaluate them by comparing the predicted performance of a typical pond to field experience. Because more published pond experience is available for catfish, the examples will be adjusted to account for typical catfish feeding rates, feed nitrogen content, and water temperatures. Three examples are considered; the case of maximum carrying capacity without active aeration, the case of nightly routine aeration with a 1 horsepower per acre aerator (3500 lb/acre carrying capacity), and finally the case of maximum observed feeding capacity of 100 lb/acre fed in routinely aerated catfish ponds.

Example 4.4 Surface Reaeration Limited Fish Carrying Capacity Calculate the fish carrying capacity in an unaerated pond, in upstate South Carolina with an average wind speed of 6 mph (9.65 km/h) assuming an average dissolved oxygen concentration in the pond of 5 mg/L, and a water temperature of 28~ with 1.5% of fish body weight being fed per day with a 35% protein feed.

Solution:

Surface reaeration can be predicted using from the

KLavalue of 0.614 day-~ previously

obtained in example 4.3 and assuming an average acceptable oxygen concentration in the pond of 5.0 mg/L. Pond dissolved oxygen saturation at 28~ is 7.75 mg/L: dc _ (0.614/day_l) (7.75 - 5 . 0 ) = 1.68 rng dt L -day

The pond feed rate is based on the assumption of 0.65 mg 02 demand per mg of feed fed to the pond, which is 1 meter deep:

112

1.68

mgO 2

gO 2

- 1.68

L -day

in a 1 meter deep pond

m 2-day

therefore; 1 $; feed

1.68 g 02 m 2 day

1.0 wet feed wt 0.90 dry feed

0.65 g 02

=

25.6

4048 m 2

1 lb

I acre

454 g

lb feed acre -day

This feeding rate can then be used to predict an algal standing crop and secchi disk using equations 4.3-4.9: First the pond nitrogen loading rate is calculated from equation 4.3" 25.6 lb feed acre

[35% protein] [.000134] = 0.120 mg N L -day

The algal carbon fixation rate is then obtained from equation 4.4: mg N ] g-carbon 0.120 L ' d a y ] [5.62] = 0.674 m2-day

Photosynthetic oxygen production is obtained from equation 4.5:

0.674 g-carbon m 2 -day

[3.47] = 2.34

g 02

m 2-day

The algal standing crop is then predicted from equation 4.6: 0.675 g-carbon m 2-day

[5] = 3.37 g-carbon m

2

0.675 g_-carbo_.....,n [10] = 6.75 g-carbon 2 m 2-day m

typical minimum

typical maximum

113

These values can be converted to algal volatile suspended solids using equation 4.7: 3.37 g_-c_arbo_.___n [2.2] = 7.4 mg TSS L m 2-day

typical minimum

6.75 g_-carbon [2.2] = 14.85 mg TSS L m 2-day

typical maximum

The pond secchi disk value can be predicted using equations 4.8 and 4.9:

0.932 = 0.80 m maximum

14.85 - 11.65 297

= 0.55 m minimum

-8.22

As a "best guess," it is reasonable to assume an average cell age for the algal biomass of 7.5 days (13.3% algal biomass loss each day). This gives an algal standing crop of 11.1 mg/L and a secchi disk of 0.52 m. Actual observed secchi disk readings in ponds receiving from 10 to 70 lb/acre-day of feed (Data from Tucker et al., 1979) correlate well with predicted secchi disk values obtained using this technique (Figure 4.4). The fish carrying capacity and projected pond oxygen concentration can be calculated assuming an average feeding rate of 1.5% of body weight per day, at the end of the season. 25.6 lb feed acre day

100 lb fish day 1.5 lb feed

= 1707 lb fish acre

If 13.3% of the algal biomass is lost each day by being "stored" in the pond food chain (and therefore does not respire away the oxygen produced by fixation), then this suggests that of the 2.34 mg O2/L-day of photosynthetic oxygen production 13.3% or a net addition of 0.31 mg/L or 0.31 g O2/m2-day will be added to the pond. This small addition of 0.31 g O2/m2-day to the pond from algal harvest would suggest an increase in pond carrying capacity of(1.68 mg + 0.31 mg O2)/1.68 or a 1.18 increase, resulting in a prediction of 30.3 Ib of feed and 2014 lb of fish carrying capacity. In a typical pond, in which the algal cultures are unmanaged, it is likely that the actual oxygen yield from algal harvest could vary from lows of 5% per day to highs of 30% per day. To be conservative, this gain in carrying capacity is disregarded in the calculations presented in Figure 4.4.

114

10.0 _

0.9 -

0.8

~

5

2

i 9

0.7~= o . 6 -

.5

x_

p

.2

9

0.5 ~ -T-

o 0

0.4

P

_

39

29

p

0.3

9

0.2 o.iL

,,

0

L

0

10

20

30 FEED

40 RATE

50 (Ib/ocre

i

'

60

70 per

80

90

1 O0

doy)

Fig. 4.4. Observed vs. predicted secchi disk at various feeding rates. (1 = 4,942 fish/HA, 2 = 10,003 fish/HA, 3 = 20,385 fish/HA, p = predicted value from equations 4.3 - 4.9. Data from Tucker et al. 1979. )

Example 4.5

Aeration Requirements for 3500 Ib/acre Fish Carrying Capacity.

Calculate the supplemental aeration needed to support 3,500 lb/acre of catfish. Use a feed rate of 1.5% of fish body weight per day with 35% protein feed with a pond water temperature of 28~

Solution." The rate of nitrogen loading to the pond is based on a feeding rate 1.5% of fish body weight per day or 52.5 lb/acre-day. Using equation 4.3, this feeding rate yields;

52.5

lb [35% protein] [.000134] = 0.246 acre -day

g-N m

2-day

115 Algal production, standing crop and secchi disk are calculated from equations 4.4, 4.6 and 4.9:

0.246 m2_----~~ay ] g - N [5.62] = 1.38 g-carbOnm2_day

1.38 g-carbon m 2-day

average standing crop @ 0=7.5 days

[7.5] = I0.35 g-carb~ m2

r

10.35 g-carb~

[2.2] = /22.77 mg TSS

L

m 2

1n[22.77 - 11.65 297 -8.22

L

= [0.399 m]

average TSS @ 0=7.5 days

average predicted secchi disk

The photosynthetic oxygen production of the pond is calculated from equation 4.5:

I

1.38 g-carbon m 2-day

[3.47] = 4.79

g-O 2 m 2-day

or 4.79

mg 0 2 L

If the pond is aerated in such a manner to insure an average dissolved oxygen of 5.0 mg/L, then the predicted ADO of 4.79 mg/L suggests a maximum pond oxygen concentration of 7.39 mg/L and a minimum value of 2.60 mg/L. The aeration requirement in excess of the 1.68 g Ojm2-day supplied by surface reaeration (example 6.4) is calculated from the BODL loading to the pond from the feed: 52.5 lb feed

454 gm

acre

0.65 g 02

4048 m 2

g feed

= 3.83 acre-day

g

0 2

m 2-day

116

This oxygen requirement from external aeration (reduced by the 1.68 g 02/m2-day from wind driven re-aeration) corresponds to:

3.83 - 1.68 = 2.15

g 02 m 2-day

or

19.2

lb

0 2

acre-day

A one horsepower per acre aerator running 12 hours per night is typically capable of supplying 1-2 lb ofO 2/hp-h-acre or 12-24 lb O ~cre-day. This corresponds to field observations that 3500 Ib/acre represents a carrying capacity that will require routine nightly aeration at maximum oxygen demand. Conversely, ifa 2.0 mg/L dissolved oxygen concentration is defined as the lowest acceptable average concentration in the pond then the wind driven interfacial oxygen transfer can be recalculated from equation 4.12 as" de - (0.614 day-l)(7.75 - 2.0) = 3.53 g-oxygen m 2-day

This oxygenation rate can be converted to a pond feeding rate in the same manner as calculated in Example 4.4: 3.53 g O a

1 ~; feed

1.0 wet feed

4048 m 2

I lb

m 2 day

0.65 g 02

0.9 dry feed

1 acre

454

= 53.8

lb feed acre -day

This value corresponds to almost exactly the feeding rate observed by Cole and Boyd (1986) as the maximum feeding rate tolerated in unaerated ponds (Figure 4.5).

Example 4.6

M a x i m u m Observed Pond Fish Carrying Capacity.

The maximum sustained carrying capacity of ponds is observed to be in the range of 100 lb/acre-day of feed application. Calculate the fish carrying capacity, oxygenation requirements and expected secchi disk reading for a pond at this limit.

Solution:

Fish carrying capacity is calculated from 1.5% of body weight fed per day: 100 lb feed

1O0 lb fish

acre-day

1.5 lb feed

= 6666

lb fish acre

117

Algal productivity and standing crop is predicted from equations 4.3, 4.4, 4.6, 4.7 and 4.9: ] 100 lb feed [35% protein] [.000134] = 0.469 .....g-N . acre -day m 2-day

J

F [5.62] = [2.64 g.-c_arbo.__.._n m 2-day

0.469 g - N m 2-day

2.64 g-carbon m 2-day

[

F [7.5 days] = ~19.80 g-e_arbo_..._n m 2-day L

19.77 g -carbon m2

average standing crop

[2.2] = [43.56 mg TSS]L averageTss

F

In [43.49 - 11.65 L"" "297 -8.22

= [0.27 m] average predicted secchi disk

Pond oxygen production is predicted from equation 4.5"

2.64 g-carbon m _2 -day

or

r [3.471 = [9.16 g._o_xyge_...__.n m 2_day [

9.16 m_._gg in a 1 meter deep pond L

Pond oxygen demand is calculated from feed application:

100 lb feed

0.9 g dry feed

454

acre

0.65 g

0 2 g 0 2 =

6.56

m 2-day acre-day

1.0 g wet feed

lb

4048 m 2

g-feed

118

Pond oxygenation requirement, assuming an average DO of 5.0 mgjL, is calculated by subtracting file wind driven aeration (1.68 g O2/m2-day) from the pond oxygen demand: g 02 :

4 , ,

m 2-day

m 2-da

or 43.5

lb 0 2

oxygenation requirement

acre -day

This corresponds to an active supplemental aeration of 1.81 lb O:/hp-h for a 2 hp/acre aerator running 12 hours every night at maximum carrying capacity. It is interesting to note that the carbon fixation rate of 2.64 g C/m2-day under these conditions is approaching the limit observed in "unmanaged" algal culture ponds. This suggests that the observed 100 lb/acre feed limit may be due to the inability of the pond algal crop to process added nitrogen at rates exceeding this value. Furthermore, if at this carbon fixation rate 20% of the algal photosynthesis is "lost" to the pond each day as is frequently observed in these systems, and if this 20% per day (corresponding to an average algal cell age of 5 days) is stored in biomass, which is ultimately removed from the pond, then 20% of the 9.16 g OJm 2day production will be available to aerate the pond. This amount of photosynthetic aeration would reduce pond supplemental aeration to 27.2 lb OJacre-day. This explains why on certain days the pond may appear to need SO0

.--

0

i

Z 0 ,/

//

! -

/ / / i

I

i

i

2OO f

o@"-

coy)

Fig. 4.5. Field determined aeration requirements vs. feeding rate to maintain a minimum pond dissolved oxygen of 2.0 mg/L (from Cole and Boyd 1986).

119

less aeration than on others. If the algal cell age could be reduced to 1.83 days (53.3% of algal photosynthetic oxygen returned to the pond) by active removal of algal biomass production through the use of filter feeders or physical chemical removal, the supplemental aeration could be reduced to zero as the net algal oxygen production of (9.16) (.533) = 4.88 g O2/m2-day would offset net pond oxygen demand.

Example 4.7 Prediction of Pond pH and Effect of Liming. Predict the pond pH in example 4.5 with 3,500 lb/acre of fish carrying capacity. Assume that the pond has a water alkalinity of 50 mg/L as CaCO> What effect will increasing pond alkalinity to 100 mg/L (as CaCO3) with lime addition have on pond pH?

Solution:

The carbon fixation rate was calculated previously as 1.38 g-carbon/m2-day or 1.38 mg/L-day. The carbon addition rate to the pond is based on the feed addition and is calculated assuming that 85% of added feed carbon is respired into the water: The total carbon released to the pond is calculated from the feed addition rate:

52.5 lb ,,

acre-day

0.9 lb dry feed

0.85 lb carbon released

0.5 lb carbon

1.0 lb added feed

1.0 lb carbon added

Ib dry feed

454 g 1.0 lb

or 2.25

1 acre

4048 m 2

mgC L -day

,,,

= 2.25 g-carbon m Z_day

in a 1 meter deep pond

It is reasonable to assume that pond respiration is constant while photosynthetic carbon fixation occurs only during the daylight hours, therefore, a diumal carbon balance can be constructed:

120

Carbon Balance

Day

Night

Fixation

- 1.38 rag C L

0

Feed/fish respiration

+ 1.125 mg C L

+ 1.125 mg C L

Algal Decomposition

+0.69 mg C L

+0.69 rag C L

TOTAL

+0.435 mg C L

+ 1.815 mg C L

The average net carbon flux rate is out of the pond and occurs at the carbon loss rate of 2.25 mg/L-day. However, because daytime photosynthesis is refixing some of the excess CO2, the intensity of the carbon loss is 4.17 times greater at night than day. This carbon loss rate can be related to the total inorganic carbon concentration by using the gas transfer equations 4.12 and 4.14. The KL~for CO2 can be estimated from the KLa(O2) used in example 4.3 by adjusting for the molecular weight of carbon dioxide:

= [ mwt KL~(C~

32] [o.614 day-l]- 0.446 day-1

02 [rawt CO 2 KLa(O,) = - ~

The average daily pH is obtained by first calculating the total carbon gas transfer driving force from equation 4.12:

dCdt - ( 0 . 4 4 6 d a y - l ) ( A c O 2 ) = (0.435 rag_12 -carbon hrs

During the day

and dedt - (0.446 day -l ) (ACO2) =

1.815

rag-carbon) L"12~rs

During the night

or ACO 2 = 1.95 mg carbon/L A CO 2 = 8.14 mg carbon/L Therefore, the C O 2 transfer rate of 0.435 mg-carbon/L-12 hrs is calculated to occur at an average ACO2 of 1.95 mg-carbon/L or 1.95/12 = 0.1625 mmole/L of carbon. In the worst case, this CQ loss rate increases to 1.815 mg-carbon/L-12 h at night yielding a A C O 2 of 8.14 mg-carbon/L or 0.678 mmole/L of carbon.

121

The calculations required to predict pond pH can be very complex, and a trial and error solution is often required to determine exact values. However, approximate solutions can be obtained from Figure A- 1 in Appendix A. In water at equilibrium with atmospheric CO,, the C for total carbon (in mmole/L) in solution is approximately equal to the alkalinity (in meq/L). Furthermore, at the pH values typically encountered in aquaculture ponds, it is reasonable to assume that the driving force for CO2 gas transfer is C r or the total inorganic carbon concentration (Stumm and Morgan, 1981). Therefore, Figure A- 1 predicts an approximate equilibrium water pH of 8.0 at a water alkalinity of 50 mg/L a s CaCO 3 or 1.0 meq/L. The range of carbon flux rates calculated above suggest an equilibrium Cv of 0.678 mmole/L higher than equilibrium at night, and 0.1625 mmole/L higher than equilibrium at day. Figure one predicts pH's of 6.6, and 7.0, respectively at these CT values of 1.678 and 1.1625 mmole/L and 1.0 meq/L alkalinity. By increasing the alkalinity to 100 mg/L as CaCO3 (or 2 meq/L alkalinity) the effect is to suppress the pH variation from an approximate equilibrium value of 8.5 to a daytime pH of 7.5 and a nighttime pH of 6.8. This compares very well with Tucker's et al. (1979) field observations of average low pH of 6.8 at these feeding rates. However, Tucker also observed occasional daytime high pH's of 9.0. How can this be accounted for? The algal growth in a typical fish pond is often observed to behave in batchwise crashes and blooms. If this were to occur, the algal decomposition may not occur on an average daily basis as shown the carbon balance. In other words, during unbalanced periods of algal bloom, little or no algal decomposition might occur. This could lead to a net gain of carbon into the pond due to photosynthesis of as much as 0.255 mg carbon/L- 12 hrs or a rate of 0.51 mg/L-day (algal decomposition = 0, see carbon balance). This yields (from Equation 4.12) an inorganic carbon concentration 0.091 mmole/L less than equilibrium. Figure 1 Appendix A predicts a pH of 9.5 at this condition ofCr = (1.0 - 0.091) or 0.909 mmole/L and an alkalinity = 1.0 meq/L.

4.7

PREDICTION OF AQUACULTURE REUSE SYSTEM PERFORMANCE AND WATER QUALITY

The design capacity and water quality dynamics in a water reuse aquaculture system is based on the fish loading, feed rate, and fish oxygen demand. A mass balance is used to tie fish metabolism to system design and resultant water quality.

Example 4.8 Reuse System with Nitrification A 264 gal (1000 L) tank holds 0.24 kg/L of fish biomass (2.0 lb/gal). Fish are fed a 42% protein feed at 1.2% of body weight per day. The oxygen uptake rate of this fish biomass is 1 kg O 2/100 kg fish. Water temperature is 28~ If the exit dissolved oxygen must not fall below 5.0 mg/L and the total ammonia must not exceed 0.5 mg/L, what flow rate is required? What is the oxygen demand due to nitrification of the effluent in a nitrifying reactor? What amount of alkalinity must be supplied to the system to offset the alkalinity destruction of the nitrifying filter?

Solution."

The calculation of water flow rate is based on oxygen uptake rate"

1000 L

0.24 kg fish L

1 kg O~ 1O0 kg fish-day

= 2.40

kg

0 2

day

or

2,400,000

mg

0 2

day

122

Assuming that the water is reoxygenated to saturation (7.75 mg/L) on each pass through the system, the water flow is calculated from an oxygen mass balance:

2,400,000 m~ O~

L

day

= 872,727

7.75 - 5.0 mg 02 available

L day

This result is typical for most high density recirculating fish culture systems. Water recirculation for oxygen replacement leads to excessively high flows and pumping requirements. In this case, a flow amounting to 160 gpm (606 L/m) would be required, with a tank hydraulic detention time of only 1.65 minutes. To avoid this situation, oxygen is usually supplied directly to the tank and the water flow is controlled by the maximum allowable ammonia concentration in the tank. Given the maximum total ammonia concentration of 0.5 mg NH3-N, the ammonia limited flow is based on a nitrogen mass balance, assuming that 60% of nitrogen fed to the fish is released to the tank (Gunther et al., 1981):

1000 L

0.24 k~ fish

I

0.012 k~; feed

L

[ 0.9 kg dry. feed

kg fish

[

0.42 kg protein kg dry feed

kg wet feed

0.16 k~ TKN

60% NH~-N released

106 mg

kg protein

Kg TKN

kg

NH 3-N

= 104,509 mg

day

This ammonia nitrogen production rate is transformed into a water flow rate by dividing by the allowable maximum ammonia concentration:

104,509 mg NH~-N day

-- 209,018 0.5 mg NH3-N

L day

This reduces the system flow requirements by a factor of 4.2. The oxygen demand from a nitrification reactor converting the ammonia production into NO3- can be calculated using Equation 4.1"

104,509 m~, NH~-N

4.6 mg O,

day

mg NH3-N

= 480,741

mg 0 2

day

123

Therefore, the nitrification reactor will increase system oxygen demand by 20% over the fish oxygen requirement. In addition, equation 4.1 can be used to calculate the alkalinity destruction due to this nitrification demand:

104,509 mg NH3-N

7.1 mg alkalinity

day

mg NH3-N

= 742,013 ~mg as CaCO 3 L

This corresponds to an alkalinity destruction of [742,013/(1000)(50)] = 14.8 eq/day of alkalinity. If this alkalinity were supplied with NaHCO3 (sodium bicarbonate) this would amount to 2.74 lbs of NaHCO3 needed per day.

4.8 SUMMARY The underlying assumptions of using feed oxygen demand and feed nitrogen loading to a pond as a basis for prediction of pond water quality may, at first, seem foolishly simplistic. However, comparisons of predicted pond pH, pond oxygen levels, pond secchi disk readings, and pond oxygenation requirements are seen to compare favorably to actual field experiences. While these assumptions may be used to make reasonable predictions of the average behavior of these water quality and design parameters, they may not however, be accurate for day-to-day predictions. Pond sediment nutrient dynamics, algal carbon fixation, and surface reaeration can change radically on a day-to-day basis as a result of increases in wind velocity, cloudiness, and changes in zooplankton density. The equations can be adjusted to account for direct effects of wind on surface reaeration. Unfortunately, the effects of such meteorological events on sediment recycling of nutrients is not well quantified. Furthermore, cycling of zooplankton populations has yet to be predicted with any degree of confidence. Therefore, precise day-to-day behavior of these complex pond aquaculture systems will likely remain empirical for some time. However, the technique outlined here is very useful for predicting the averaged behavior of water quality in a pond under different feed loadings or fertilization intensities. These types of predictions are important in that they allow the fish farmer to determine the limits to which he can "push" his pond with respect to nitrogen assimilative capacity, and the resultant impact on water quality, particularly pond dissolved oxygen concentrations and pond aeration requirements. These effects will be the same regardless of whether the pond is growing catfish or stripped bass. Unfortunately, little field data is available from stripped bass pond production systems. For this reason we must turn to observations of catfish pond performance to confirm our predictions. Furthermore, these calculations allow fish farmers to explore the possibilities of increasing production by pond design modification. For example, the calculations and field observations show that the algal growth and oxygen production capacity in conventional fish ponds is currently under-utilized by a factor of 2 to 3. Designing a fish culture pond to take full advantage of the 6-12 g C/m-' day of photosynthetic capacity that is possible from managed algal cultures could theoretically increase fish production to 10,000-20,000 lb/acre year. Research is currently underway at Clemson University (Brune et al., 1995; Schwedler et al., 1994) in an effort to prove this level of predicted carrying capacity in the field, through the utilization of managed algal photosynthesis in aquaculture ponds.

124

References

Almazan, G. and Boyd, C.E. 1978. An evaluation of secchi disk visibility for estimating plankton density in fish ponds. Hydrobiologia, 61 (3): 205-208. Andrews, J.W. and Matsuda, Y. 1975. The influence of various culture conditions on the oxygen consumption of channel catfish. Transactions of the American Fisheries Society, 2: 322-327. Banks, R.B. and Herrera, F.F. 1977. Effect of wind and rain on surface reaeration. Journal of Environmental Engineering Division, ASCE, 103(EE3): 489-503. Boyd, C.E., 1991. Empirical modeling of phytoplankton growth and oxygen production in aquaculture ponds. Pages 363-395 in D.E. Brune and J.R. Tomasso, editors. Aquaculture and water quality. The World Aquaculture Society, Baton Rouge, LA. Boyd, C.E., 1985. Chemical budgets for channel catfish ponds. Transactions of the American Fisheries Society, 114: 291-298. Boyd, C.E., 1979. Water quality in warmwater fish ponds. Auburn University Agricultural Experiment Station, Auburn, AL. Brune, D.E., Drapcho, C.M. and Piedrahita, R.H., 1992. Pond oxygen dynamics; design and management strategies. Aquaculture 92 International Conference, American Society of Agricultural Engineers, Paper No. Aqua 92-101, Orlando, FL. Brune, D.E., Collier, J.A. and Schwedler, T.E., 1995. Nutrient recovery and reuse for water quality control in the partitioned aquaculture system. Proceedings of the Sustainable Aquaculture 95 Conference. Pages 57-68, Honolulu, HI. Brune, D.E. and Gunther, D.C., 1981. The design of a new high rate nitrification filter for aquaculture water reuse. Journal of the World Mariculture Society, 12: 20-32. Brune, D.E., 1991. Fed pond aquaculture. Aquaculture systems engineering, Proceedings of the World Aquaculture Society and American Society of Agricultural Engineers, Jointly sponsored session, ASAE Publication, 02-91. Pages 15-33. Busch, C.D., Flood, C.A., Jr., Koon, J.L. and Allison, R., 1977. Modeling catfish pond nighttime dissolved oxygen levels. Transactions of the American Society of Agricultural Engineers, 17:433-435. Chieng, C.A., Garcia, A. and Brune, D.E., 1989. Oxidation requirements of a formulated micropulverized feed. Journal of the World Aquaculture Society, 29: 24-29. Cole, B.A. and Boyd, C.E., 1986. Feeding rate, water quality, and channel catfish production in ponds. The Progressive Fish-Culturist, 48: 25-29. Colt, J., 1986. An introduction to water quality management in intensive aquaculture, Use of supplemental oxygen. Section 6, Pages 1-16/n H. Lorz, convener, Northwest Fish Culture Conference, Eugene, OR. Drapcho, C.M., 1993. Modeling of algal productivity and diel oxygen profiles in the partitioned aquaculture system, Ph.D. Dissertation, Department of Agricultural Engineering, Clemson University, Clemson, SC.

125

Goldman, J.C. and Ryther, J.H., 1975. Nutrient transformations in mass cultures of marine algae. Joumal of Environmental Engineering Division, ASCE, 101 (EE3): 351-364. Goldman, J.C. 1980. Physiological aspects in algal mass culture. Pages 345-359, in G. Shelef and C. J. Soeder, editor , Algal biomass, Elsevier Press, New York. Gunther, D.C., Brune, D.E. and Gall, G.A.E. 1981. Ammonia production and removal in a trout rearing facility, Transactions o f the American Society o f Agricultural Engineers, 24( 5): 1376-1380. Lehman, J.T., Botkin, D.B., Likens, G.E., 1975. The assumptions and rationales of a computer model ofphytoplankton. Population Dynamics, Limnology and Oceanography, 20: 343-364. Meyer, D.I. and Brune, D.E., 1982. Computer modeling of the diumal oxygen levels in a stillwater aquaculture pond. Aquacultural Engineering, 1" 245-263. Mueller, J.A., and Boyle, W.C., 1988. Oxygen transfer under process conditions, Journal Water Pollution Control Federation, 60 (3): 332-341. Piedrahita, R.H., 1991. Modeling water quality in aquaculture ecosystems. Pages 322-362 in D.E. Brune and J.R. Tomasso, editors. Aquaculture and water quality. World Aquaculture Society, Baton Rouge, LA. Piper, R.G., McElwain, I.B., Orme, L.E., McCraren, J.P., Fowler, L.G. and Leonard, J.R., 1982. Fish hatchery management. U.S. Department of Interior, Washington, DC. Redfield, A.C., Ketchum, B.H. and Richards, F.A., 1963. The influence of organisms on the composition of sea-water. Pages 26-77, in M.N. Hill, editor. The sea. volume 2. New York. Schindler, D.W., 1971. Carbon, nitrogen and phosphorus and eutrophication of freshwater lakes. Journal of Phycology, 7:321-329. Schroeder, G.L. 1987. Carbon and nitrogen budgets in manured fish ponds on Israel's coastal plain. Aquaculture, 62: 259-279. Schroeder, G.L., Alkon, A. and Laher, M., 1991. Nutrient flow in pond aquaculture systems, Pages 489-505 in D.E. Bnme, and J.R. Tomasso, Jr., editors, Aquaculture and water quality, World Aquaculture Society, Baton Rouge, LA. Schwedler, T.E., Brune, D.E. and Collier, J.A., 1994. Development and modeling of the partitioned aquaculture system. USDA funded project, Aquaculture Special Grants Program, Washington, DC. Shelef, G., Schwartz, M. and Schechter, H., 1973. Prediction of photosynthetic biomass production in accelerated algalbacterial wastewater treatment systems. Pages 181-189, in S.H. Jenkins, editor. Advances in water pollution research. Pergamon Press, Oxford, Great Britain. Smith, D.W. and Piedrahita, R.H., 1988. The relation between phytoplankton and dissolved oxygen in fish ponds. Aquaculture, 68: 249-265. Stumm, W. and Morgan, J.J., 1981. Aquatic chemistry, 2nd edition. John Wiley & Sons, New York. Tucker, L., Boyd, C.E. and McCoy, E.W., 1979. Effects of feeding rate on water quality production of channel catfish and economic returns. Transactions of the American Fisheries Society, 108: 389-396.

126

U.S. Environmental Protection Agency, 1975. Process Design Manual for Nitrogen Control. Washington, D.C. Wheaton, F. Hochheimer, J. and Kaiser, G.E., 1991. Fixed film nitrification filters for aquaculture. Pages 272-303/n D.E. Brune and J.R. Tomasso, Jr., editors, Aquaculture and water quality, World Aquaculture Society, Baton Rouge, LA.

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V, All rights reserved.

127

Chapter 5

Intensive Culture of Striped Bass John N. Hochheimer and Fredrick W. Wheaton 5.1 INTRODUCTION Striped bass can be grown using either extensive or intensive culture techniques. Extensive culture includes practices, such as stocking at low densities in ponds, lakes, or other larger bodies of water, that rely on natural food supplies (e.g., plankton or forage fish) and natural water-flow dynamics to maintain water quality and sufficient oxygen. Culturists who use extensive systems have little control over the environment in which their fish are growing. Intensive culture produces fish at densities greater than can be supported in the natural environment. High water flow or exchange rates are used in intensive culture, and the fish feed primarily on formulated feeds (Piper et al., 1982). Intensive culture systems can take the form of ponds stocked at high densities (intensive pond systems); net pens or cages placed in lakes, reservoirs, estuaries, or coastal waters; rectangular raceways or circular tanks with high volumes of water flowing through them (flow-through systems); or closed (recirculating) systems. As the intensity increases, culturists gain more environmental control for their fish production systems and also increase the number and complexity of support systems required to maintain optimal conditions for culture. Most striped bass culturists use some form of intensive culture for production of most or all life stages. Some production aspects are still, however, more suited to extensive culture techniques. For example, most striped bass fry producers rely on naturally reproducing females for broodstock and may also use male fish caught from wild stocks. This trend is slowly changing as improvements are made in the reproductive aspects of striped bass production (see Chapter 2). After fry are hatched in a hatchery (intensive operation), most producers of phase I fingerlings use extensive pond culture techniques. These producers promote the growth of natural plankton in ponds to provide food for the growing fry (see Chapter 3). As the fingerlings mature and the natural food supplies become too small for the growing fish, artificial feeds are introduced to the fish and the culture operation becomes more intensive. It is important to note that some differences between extensive and intensive pond systems are very subtle. In Chapter 3, pond culture was explained in great detail. In the present chapter, some pond components will be reviewed to demonstrate how they relate to more intensive operations. However, the primary emphasis of this chapter will be on flow-through and recirculating systems. 5.1.1 Why Intensive Culture? Striped bass culture, whether for population enhancement or as a farmed crop, requires a high degree of efficiency to be successful. Culturists who use conventional extensive culture techniques have only minimal control over critical culture variables such as water temperature, dissolved oxygen, salinity, and other water quality characteristics; feeding rates; selective harvesting; or disease recognition and treatment (Carlberg et al., 1984). Intensive culture allows the culturist to control one or more of the variables affecting aquacultural production. Many intensive systems allow aquaculturists to see their crop and this can help in the early diagnosis of stress or disease. As the culture operation becomes larger, controlling these variables becomes more important because of the increased investment. Striped bass appear to have a wide tolerance for several water quality variables (e.g., salinity or temperature) and, in general, become more tolerant to a wider range of water quality with increasing age

128

(Kerby, 1993). Carlberg et al. (1984) summarized several advantages of using striped bass and its hybrids as a species for intensive culture: 9 adaptable to controlled environments, 9 exhibits rapid growth rates, 9 has broad physiological tolerances, and 9 has a high market value. Extensive culture of fingerling striped bass in ponds has also been shown by Kerby (1993) to result in wider size variations than those grown in intensive culture systems. Size distributions of fingerling striped bass stocked in ponds at low densitiesm12,350 fish (0.2g to 2.0 g each) per hectare--resulted in a bimodal size distribution where 80% of the harvest were small fish (20g) and 20% were larger fish (100g) (Kerby, 1993). Survival in extensive systems (and in intensive systems) can also be affected by cannibalism. Commercial development of intensively cultured striped bass and its hybrids occurred steadily during the 1980s. Researchers such as Kerby et al. (1987) and Smith et al. (1989) demonstrated that pond production of hybrid striped bass can range from 5600 kg/ha to greater than 7500 ko~qaa if adequate water quality is maintained. Smith et al. (1985) showed that hybrid striped bass could be cultured intensively in tanks under controlled conditions to weights over 0.23 kg in less than one year, with a standing crop at harvest of 43.2 kg/m 3. Comparatively, typical intensive tank cultures of finfish strive to achieve a standing crop of about 120 k~m 3 to justify the costs associated with these often expensive systems. Table 5.1 illustrates some of the production levels achieved for several species of finfish in different intensive systems. Striped bass and hybrid striped bass production is rapidly becoming a significant aquaculture industry in the United States. Commercial production of striped bass and hybrid striped bass in the United States was about 3.2 x 10 6 kg in 1994, with farm prices ranging from $5.33 to $6.67 per kg (Seafood Leader, 1995). Commercial hybrid striped bass producers range from pond culture operations (such as Carolina Fisheries and Hyrock~), to flow-through operations (such as Kent Seafahns ), to recirculating operations (such as AquaFuture3). The demand for striped bass in the market place has often exceeded the supplies available from the traditional commercial fishing sources. As native populations fluctuate, traditional commercial fishing grounds become unavailable, and the demand for fisheries products such as striped bass continue to increase, intensive culture of striped bass becomes more economical and competitive. Additionally, intensive culture practices can potentially offer the marketing advantages of providing a continuous supply of uniformly-sized fish. The largest intensive culture facility for hybrid striped bass, Kent Seafanns (formerly Aquatic Systems, Inc.) who have been producing hybrid striped bass for over 10 years, can be used to illustrate successful hybrid striped bass culture in an intensive system. Kent Seafarms has 48-65m 3 and 48-190m 3circular tanks in a system that uses high quality geothermal water to produce 1.4 million kg of hybrid striped bass annually. This flowthrough system uses state-of-the-art oxygenation, continuous water quality monitoring, and a team of highly trained culturists to produce a premium product (J. Carlberg, Kent Seafarms, Inc., personal communication). Recirculating systems were successfully used by Lewis et al. (1981) to raise striped bass from one day-old larvae to 5 to 10 cm fish at harvest. Over a seven year period, Lewis et al. (1981) used 34-1.7m 3circular tanks

1 Carolina Fisheries, Aurora, NC. Hyrock Aquaculture Farm, Inc., Princess Anne, MD 2 Kent Seafarms, Inc., San Diego, CA 3 AquaFuture, Inc., Turners Falls, MA

129

Table 5.1. Production comparisons for several species of finfish.

Species

Production Method

Density

Carp ~

extensive pond intensive pond

310 kg/ha 8700 kg/ha

(Oncorhynchus mykiss)

flow through cage culture

60 kg/m 3 10-20 kg/m 3

Channel Catfish 2

intensive pond

6725-7846 kg/ha

intensive pond cage tank

1350 kg/ha 3.5-7 kg/m 3 25-50 kg/m 3

intensive pond 3 flow-through 4

7500 kg/ha 43.2 kg/m 3

( Cyprinus carpio) Rainbow Trout ~

(lctalurus punctatus ) Tilapia ~

( Tilapia sp.) Striped Bass

(Morone saxatilis)

Source: ~Pillay (1990); 2 Tucker and Robinson (1990); 3Smith et al. (1989);4 Smith et al. (1985)

and upflow biofilters to produce fingerling striped bass for population enhancement. Survival rates ranged from less than 1% at the beginning of the study to over 46% at its completion (Lewis et al., 1981). This was the first major effort in recirculating system culture of striped bass and many subsequent intensive culture operations of striped bass have benefitted from information collected during the study. In spite of the recent successes in intensive striped bass culture, much remains to be learned about striped bass and its hybrids. Many components and processes in intensive culture systems also need further refinements. Water quality requirements for striped bass culture have not been clearly defined (Kerby, 1993). Because of this, it is difficult for engineers to design components or complete intensive culture systems, with high confidence that the systems are reliable under all conditions. Intensive culture systems have inherent disadvantages (Carlberg et al., 1984), including: 9 may produce wide fluctuations in water quality; 9 can lead to outbreaks of stress related disease, 9 depend on mechanical devices, 9 need to supply all nutritional requirements of fish, 9 need alarms and redundant back-up systems, and 9 require experienced and trained personnel. There are many unanswered design questions facing engineers and biologists responsible for developing intensive culture systems. Costs for designing and constructing intensive systems are often much higher than for extensive systems. Similarly, operational costs in intensive systems are greater than those for extensive

130

systems. However, as the price of land and water, the demand for more seafood, and environmental regulations on discharges from aquacultural facilities increase, intensive systems, especially those that rely on recirculating a large percentage of water, will become more desirable. 5.2 W A T E R SUPPLIES

The water source for the intensive culture system can be groundwater, surface water, or municipal water supplies. Groundwater sources typically have a constant temperature and, as a general rule of thumb, below about 15m, water temperature increases approximately 1 ~ for every 32 m in depth (American Water Works Association, 1951). Regardless of the source, water quality should be checked to determine that it is free from any pollutants that will be harmful to fish or will leave harmful residues in the flesh (see Table 5.2). Most groundwater sources contain little or no oxygen, so they must be oxygenated before use. Some groundwater sources may also contain high levels of hydrogen sulfide, nitrogen gas, carbon dioxide, and/or iron, all of which can be toxic to fish. The three types of groundwater sources are springs, depressions below the water table, and wells. Wells can be either free flowing (artesian) or pumped (Wheaton, 1977). Springs and artesian wells can be a reliable source of water for aquaculture. Many springs and wells are free from pollutants and provide constant temperature water. Pumped wells can also provide a reliable source of constant temperature water, but have the added expense of pumping costs. Depressions below the water table rely on groundwater located near the surface and probably cannot be depended on in times of drought to supply water. Depressional water sources are also influenced more by surrounding land uses and can unpredictably carry pollutants into the aquaculture system. Surface water supplies include water from streams, rivers, lakes, estuaries, and the ocean. Most surface waters do not require pumping to as great a height as wells and can be relatively high in dissolved oxygen. Surface waters, however, are subject to pollution from variety of sources, some of which are intermittent. Systems with surface waters often must be treated to remove unwanted suspended solids and a variety of aquatic organisms, including pathogens. In general, the use of surface waters should be avoided, especially in recirculating systems. Municipal water supplies offer another alternative for some aquaculture systems, although most municipal water users have recirculating systems that require comparatively low volumes of water to operate. Municipal supplies do not have to be pumped, so operational costs for energy are reduced, but this cost reduction may be offset by increased costs for the water. Municipal sources, however, must be treated to remove chlorine, ammonia, fluorine, and other disinfectants or additives. Municipal water supplies tend to be costly when compared to other sources of water. 5.2.1 Aeration Control of dissolved gases is an important consideration for intensive striped bass culture. Oxygen, nitrogen, and carbon dioxide are the gases of principal concern, but other gases such as ammonia and hydrogen sulfide may also be present (Colt and Orwicz, 1991). Aeration equipment is designed to control the concentration of one or more of these gases, either by adding the gas (in the case of oxygen) or by stripping out the gas (in the case of the other gases) (Watten et al., 1989). Most aeration systems are designed to add oxygen and will strip nitrogen efficiently. Special design considerations must be made when attempting to strip other gases, such as carbon dioxide, ammonia, or hydrogen sulfide (Colt and Orwicz, 1991). This section will provide an overview of some of the common aerator types and systems. See Colt and Orwicz (1991), Wheaton (1977), Watten (1994), or Visscher and Dwyer (1990) for more detailed aeration system and gas transfer design information. Parker et al. (1990) provides a discussion of the benefits and disadvantages of several types of aeration devices for use in striped bass culture.

131

Table 5.2. Water quality for striped bass and hybrid striped bass culture systems (adapted from Warren et al., 1990 and Piper et al., 1982).

Parameter

Standard (mg/L) ~

Parameter

Standard (mg/L) ~

Alkalinity (as CaCO3)

50-400

Aluminum

0.01

Ammonia - unionized

0.0125

Arsenic

0.05

Barium

5.0

Cadmium (Alkalinity _<100) (Alkalinity > 100)

0.0005 0.005

Calcium

10-160

Carbon Dioxide

0-15

Chlorine

0.03

Chromium

0.03

Copper (Alkalinity _<100) (Alkalinity > 100)

Dissolved Oxygen

5-saturation

0.006 0.03

Fluoride

0.5

Hardness

50-400

Hydrogen Cyanide

0.005

Hydrogen Sulfide

0.003

Iron (total) ferrous ferric

0.5

Lead

0.02

Magnesium

15.0

Manganese

0.01

Mercury

0.002

Nitrogen (gas)

110% saturation

Nitrate

3.0

Nitrite

0.1

Nickel

0.01

Ozone

0.005

PCB

0.002

pH

6.5-9

Phosphorus

0.01-3.0

Potassium

5.0

Selenium

0.01

Silver

0.003

Sulfur

1.0

Sulfate

50.0

Total dissolved solids

400.0

Total suspended solids

80.0

Uranium

0.1

Vanadium

0.1

Zinc

0.5

0.05 Zirconium O. 1 All units are expressed in mg/L except for salinity, which is expressed in parts per thousand, and pH, which is expressed as pH units. All values are maximum levels for optimal fish health, but can be exceeded where experience has shown that higher values are acceptable (for example values of nitrate exceeding 100 mg/L have been observed by the authors to cause no adverse effect in striped bass being reared in a recirculating system).

132

Aerator efficiency for oxygen transfer is directly related to its ability to increase the surface area of contact between the water and oxygen (either in air or as pure oxygen) (see Chapter 4). Increasing the oxygen transfer rate in a particular application can be accomplished by either increasing the area of contact between the oxygen and water and/or by increasing the driving force (i.e., the concentration or partial pressure difference between the gas and liquid phases). The area of contact can be increased by increased surface agitation, breaking the water into small droplets, or through the use of thin contact films. The driving force can only be increased by increasing the oxygen concentration in the gas mixture (e.g., using pure oxygen or by pressurizing the air and water). Air-contact aeration systems increase turbulence and water-to-air interface area so that oxygen absorption increases from the atmosphere or from rising air bubbles (Watten, 1994). Gases present in the air and water tend to reach an equilibrium between the gas and liquid phases (Watten, 1994). Thus, when a water supply with a relatively low dissolved oxygen level is turbulently exposed to air, oxygen diffuses into the water in an attempt to reach equilibrium. Similarly, the ability of an aerator to degass excessive nitrogen from a water source is also directly related to the increase in surface area between the water and the atmosphere (Wheaton, 1977). Pure oxygen systems use gas mixtures with higher concentrations of oxygen, typically in the range of 8598% (Colt and Orwicz, 1991). Most aerators can be modified to use pure oxygen as a feed gas rather than air (Colt and Orwicz, 1991). Pure oxygen systems consist of three primary components, including a source of oxygen enriched gas, a flow regulating unit, and an oxygen-to-water contactor (Watten, 1994). Because pure oxygen systems have additional costs for oxygen generation and storage equipment, maximizing oxygen transfer efficiency is essential. Yet, these systems can provide a cost-effective means for maintaining adequate dissolved oxygen levels in an intensive striped bass culture system when they are properly designed and maintained. Sources such as Colt and Orwicz (1991) and Watten (1994) provide detailed design information for pure oxygen systems. Many different types of aeration equipment are used, but there are four main groups of aerator designs (Wheaton, 1977): 9 gravity, * surface, 9 diffusion, and 9 turbine. Some aerators are a combination of one or more attributes of these main groups (Wheaton, 1977). A key distinguishing feature of aerators is the oxygen source; it can be from air or pure oxygen. Which aeration system to use will depend on the oxygen demand and configuration of the culture system, availability and cost of oxygen and energy, and certain physical characteristics such as temperature and salinity. Operational considerations for aeration systems were outlined by Colt and Orwicz (1991) and are listed in Table 5.3. Reliability of the aeration system is an important consideration--systems that fail will lead to fish kills. Table 5.4 lists some of the more common reasons for aeration system failures. Gravity aerators use energy released when water loses altitude (head) to increase the air-water contact area (Wheaton, 1977). Because elevation differences provide the necessary head to power the aeration process, gravity aerators do not require supplemental energy inputs. This tends to increase reliability and eliminate the need for back-up power supplies. Gravity aerators are used in many aquaculture systems, especially flowthrough raceway systems because they are easy to build and inexpensive to operate (Wheaton, 1977).

133

Table 5.3. Operational considerations for aeration systems (after Colt and Orwicz 1991).

Diffuser Fouling

Operation and..Maintenance Action

Problem

A c t i v i ~ '

Algal and bacterial growth may foul aeration diffusers

Rust and scale clogging of fine pore diffusers

- Operate diffusers continuously Periodically clean diffusers by acid washing - Filter air supply Use non-metallic air lines

-

Icing and Slippery Conditions

Safety Hazards

Surface aerators and some gravity aerators produce a spray that accumulates on walkways and roads

- Provide slip-resistant surfaces when possible - Post warnings about possible slippery conditions

In freezing weather, some aerators may freeze-up, which can lead to equipment breakage and insufficient oxygen levels.

- For culture systems that must provide aeration under freezing conditions, ,use aerators that are not affected by icing--e.g., diffusers.

Electrical Hazards

- Use properly installed electrical equipment with adequate grounding - Use caution around water, especially salt water

Fuel Hazards

Mechanical Equipment

Harvesting and Feeding

Aeration equipment interferes with harvesting and/or feeding operations

Repair

Equipment will need routine maintenance and emergency repairs

- Use and store flammable fuels safely - Store all fuels in approved containers and locate all storage areas so that accidental leaks and spills will not contaminate water supplies - Put proper shields and guards on all moving parts--e.g., rotating shafts, moving blades, etc. - Use caution when operating machinery such as tractors on pond banks - use properly sized equipment for a job Locate aeration equipment to minimize interference - Ensure that equipment can be moved if necessary

-

-

-

-

Reliability

Aeration equipment must operate under severe conditions and whenever oxygen levels are low

Ease of repair and maintenance should be a design and operational consideration Have proper tools available for maintenance and repairs Ensure that spare parts and trained repair personnel are readily available

- Back-up power is essential for any electrically powered equipment When adequate head is available, gravity aerators will operate without additional power inputs Equipment design should be simple and highly reliable -

-

134

Table 5.4. Common reasons for aeration system failures (after Colt and Orwicz, 1991). Component

Failure

Diffusers

Clogging from rust, scale, or bacterial and algal growth

Utility lines and equipment--air, water, electrical

Gunshot damage to exposed lines and equipment Vehicle damage to overhead lines Crushing or breakage to buried lines Tree or animal damage to exposed and buried lines Wind or storm damage to exposed lines Freezing of water lines Flooding of electrical conduits Failure of stand-by electrical equipment

Pumps and motors

Gunshot damage Loss of electrical power--complete or one or more phases in three-phase systems Poor regulation of voltage or frequency Poor alignment of motor shafts with pump shafts Clogging of intakes or impellers Foreign objects rubbing or impeding shafts Splashing water causing electrical shorts Salt bridging in electrical motors causing shorts Poor maintenance, including lubrication

Blowers

Flooding when shut down Excessive pressure

Controls

Failure of electrical components Failure of regulators from scale or corrosion Inadequate control, which leads to decreased performance or system failure

Surface aerators spray or splash water into the air to increase the air-to-water interface area and thus increase the oxygen transfer rates (Wheaton, 1977; Colt and Orwicz, 1991). A variety of surface aerators have been devised over the years, including (Wheaton, 1977): 9 pumps that spray water into the air; 9 water pumped through nozzles at high velocity and sprayed onto the water surface; 9 water drawn into a vertical tube by a propeller, pumped upward, and deflected radially over the water surface (spray-type agitators); 9 propellers that draw air through a tube into water at the surface (propeller-air injectors); 9 rotors operated at a shallow depth to create a hydraulic jump, which entrains air; and 9 horizontal rotary brushes or paddle wheels that create turbulence around them and spray water into the air.

135

Diffuser aerators are a class of equipment that inject air or oxygen into the water in the form of bubbles (Wheaton, 1977). Oxygen diffuses from the bubbles into the water and nitrogen diffuses from the water into the bubbles as the bubbles move in the water column. Some of the more common diffuser aerators include (Wheaton, 1977): 9 pipes or tubes (static tube) placed in a water tank; 9 pipes or tubes with diffusers; 9 aspirators that inject air (or oxygen) into a moving stream of water, including venturi, nozzle, and orifice aerators; u-tubes that draw water and air down one leg of the tube and then back up the other to increase the contact time and pressure for diffusing oxygen into the water; air-lift pumps that use air bubbles to lift and move water as well as aerate; and down-flow bubble contactors (aerator cones) that use air or oxygen injected into water as it is pumped downward through a conical hood that increases in diameter with depth. 9

9 9

Turbine aerators are propellers submerged into the water that, when rotated, cause water circulation (Wheaton, 1977). Increased circulation exposes water with a low dissolved oxygen concentration to the surface where oxygen rapidly diffuses from the atmosphere into the water. Some turbine aerators include air or oxygen diffusers with the propellers, which adds air or oxygen that is sheared into fine bubbles by the propeller to enhance aeration (Wheaton, 1977). Turbine aerators should be used with caution in fish culture tanks because propellers can harm fish that come into contact with them. Aeration system design requires analysis of many different variables, including the culture system characteristics, aerator availability, power requirements and availability, capital and operating costs, and other factors (Wheaton, 1977). Table 5.5 provides a comparison of attributes for several aeration system types and Figure 5.1 shows some examples. Wheaton (1977), Boyd and Watten (1989), Colt and Orwicz (1991), and Watten (1994) provide more detail on calculations of loading rates, sizing of aerators, and how they can be designed for striped bass culture systems. 5.3 SOLIDS REMOVAL Solids accumulate in fish culture systems primarily from feces, the breakdown of uneaten food, and dead and decaying algae and bacteria. In pond systems these solids eventually settle on the pond bottom and are broken down by the living bacteria normally found in the pond sediments. Removal of solids is important for maintaining good water quality in both flow-through and recirculating systems. When solids are not removed, they break down into a variety of organic and inorganic substances that can become toxic to the fish being cultured or use up all of the available oxygen in the system. Solids accumulation can become a limiting water quality factor for production systems (Chen et al., 1994). Excessive solids can be problematic in pond, flow-through, and recirculating systems for several reasons, including (Chen et al., 1994): 9 damage to fish gills, 9 clogging of biofilters and other system components, 9 mineralization of organic substances to produce ammonia, and 9 breakdown of organic substances by bacteria that consume oxygen. Solids in effluents from culture system waters can lead to the degradation of receiving waters, such as the formation of sediment deposits in stream channels and in slower moving waterbodies (e.g., lakes, reservoirs,

136

Table 5.5. Design attributes of commonly used aeration systems.

Aerator

Design Factors 4

Examples

Type Gravity

( ~ 02/kw hi')

Most economical choice when sufficient head available No energy requirements Typically used in raceway and flow through systems

Surface

Subsurface

Typical Transfer Rates

Tend to have high oxygen transfer efficiencies

weir

1.5- 1.81

cascading or corrugated inclined plane

1.0- 1.91

overall

0.6- 2.43

Capital and operating costs comparatively low

paddle wheel electric powered tractor powered

Used in many pond culture systems

spray-type

1.3- 1.42

Provide good circulation and aeration

vertical pump

1.2 3

Can be used as pumps to circulate as well as aerate water

propelleraspirator pump

1.7 - 1.92

Simple construction, reliable operation

diffuser

0.9 - 3.03

U-tube aerators

0.7 - 40 ~

1.2- 2.91 1.3 - 2.01

Good for tank culture systems Oxygen Absorption

Very efficient aeration Can be used for oxygen supersaturation Used in flow through and closed systems

Sources: ~Colt and Orwicz (1991); 2 Boyd and Martin (1984); 3 Boyd and Watten (1989); 4 Wheaton (1977).

137

Flow -.- -;,~

,, - - -

~3-?~"~"

o

- ....

=

....

~

:

_- ................ -

s

- o_. _~.. o -__~f:o.._...ioO%__=::: -

.~

_ 2o o'~oO__o_o0

:7

I ~ z f i i Z T . . . . . . .

"i.

i

i

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

!

I

~ _

I

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

C

LOW .....................

DO

~ Water

o_---~_?_

" __~__~ ....~-:_%~____~-~.

......................................... ~--.

Pump

--- o-~S

.....

L_

~::::::::::.:.......--

Air

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

.

.

.

.

.

.

.

. .

....

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

__C._-. . . . . . . . . . . . . . . .

.

.

.

- -

.

_._~_bTJ:___ o . o O

.

~

-

-

.....

~

o oo

~c~ 0

o~ t_~__~-~L~--~f__ -~_ ~~_ _ _ _ ~_ ~ ~_0_ ....

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

.

.

Diffuser~

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

~,~/Woter

e

~

~

o:Ol

'

~

e

~

Liqukl/Air

|r L i f t Pump

Fig. 5.1. Examples of gravity aerators (a - cascading), surface aerators (b - upflow propeller and c - surface nozzle), and diffused aerators ( d - tube with diffuser and e - airlift pump).

138

or estuaries with poor flushing), nutrient enrichment of receiving waters, and oxygen depletion in areas downstream from outfalls. Solids removal is, therefore, important in many intensive system operations. Solids found in fish culture systems can be generally classified as settleable (those greater than 100 ~tm) and those that are nonsettleable (particles less than 100 ~tm) (Chen et al,. 1994). Settleable solids are typically easier to remove and can be done with relatively inexpensive processes and equipment. The finer, nonsettleable solids, are increasingly more difficult to remove as the particle size decreases (Chen et al., 1994). Some of the finer particles are a result of breakdown of larger particles, so quick removal of the larger particles will prevent some of the smaller particles from forming. However, some fine particles will still accumulate and will need to be removed. Fine particles often provide a substrate for bacteria and can result in increased ammonia levels and oxygen demand. There are two predominant classes of filter types used to physically remove solids from fish culture systems, gravitational and mechanical. Gravitational filters use gravity to physically separate solids from liquids (Wheaton, 1977), with sedimentation processes being the most common gravitational method used in aquaculture. Particles that have densities greater than that of the liquid are allowed to settle in quiescent tanks for removal (Wheaton, 1977). Sedimentation tanks or basins are simple to construct and operate and provide an economical method to remove larger-sized particles (Chen et al., 1994). Gravitationally-enhanced processes, such as centrifuges and hydrocyclones, use centrifugal forces to increase the rate of removal of particles from water. Centrifuges and hydrocyclones may be efficient in removing particles that require excessive settling times in gravity settling basins (Chen et al., 1994). Mechanical filters physically trap particles to remove them from water. A variety of different screen and other mechanical filter configurations have been used to mechanically filter culture system waters, including (Wheaton, 1977): 9 Stationary screens are placed across a channel of flowing water. Particles larger than the screen mesh are trapped and collected on the screen; 9 Rotary screens rotate with their axis oriented parallel, perpendicular, or radial to the flow of water. Particles are trapped on the screen and automatically backwashed (removed) from the screen; 9 Vibratory screens vibrate to collect and remove solids that are trapped on the screen surface. Water passes through the screen; 9 Gravity sand filters - water passes down through a bed of sand or other porus media to trap particles larger than the pore size. Collected solids are removed by backwashing and collecting the effluent; 9 Pressurized sand filters - sand is contained in a pressure vessel and water, under pressure, is passed through the sand. Solids are collected in the sand bed and greater flows can be filtered for a given sized sand bed then can be done with only gravity flow. Collected solids are removed by backwashing (see Figure 5.2); 9 Vacuum sand filters - similar to gravity sand filters, except water is pulled through the filter by vacuum; 9 Diatomaceous earth filters - vessel containing a membrane and diatomaceous earth that precoats the membrane to provide a smaller pore size. Particle size removal is a function of the size of diatomaceous earth used in the filter, as with any screening device. Diatomaceous earth filters become more efficient at removing smaller particles as the filter becomes more clogged. Another solids removal process is foam fractionation (Figure 5.3), a device where chemical and physical processes are combined to remove small particles from water. The chemical process occurs when air is bubbled up through a column of water. The air-water interface attracts solids and other charged particles to form a foam that rises to the surface (Chen et al., 1994). The rising foam carries the attached particles with it and can be easily removed from a culture system (Timmons, 1994).

139

Fig. 5.2. A pressurized sand filter.

FOAM )ISCHARGE

FRACTIONATOR COLUMN

_ LIQUID

AIR INLET

J

/

....

.....

iCLEAN

i A I R - LIQUtD MIXER

Fig. 5.3. Foam fractionation unit for the removal of time particulates.

WATER

OUT

140

5.4 INTENSIVE CULTURE SYSTEMS

The remainder of this chapter will focus on the many system components for intensive striped bass culture. Ponds, flow-through, and recirculating systems will be discussed. There is not one system that is better than any others in all cases. Rather, depending on variables such as physical location, climate, water supplies, or market conditions, one system may be preferable over the others. Closed or recirculating striped bass culture systems are relatively new compared to pond or flow-through systems. However, they show great promise as engineering design and operational experience mature. 5.4.1 Pond Systems 5.4.1.1 Overview of pond systems When is a pond intensive? Pond systems are considered intensive when artificial feed is added, the water is aerated to increase carrying capacity, water is exchanged to maintain water quality, or the pond effluents are treated to minimize environmental impacts (Piper et al., 1982). Ponds are successful intensive production systems for striped bass and/or its hybrids (Kerby et al., 1987; Smith et al., 1989; Kerby, 1993). Ponds are often chosen as the culture method of choice when the available land is relatively inexpensive, is uniformly contoured and relatively fiat, and contains sufficient clay to provide a low permeability. Economic considerations require intensive pond systems to be located in areas having a sufficiently long growing season to produce a crop in one to two years, a constant supply of high quality water, and reasonable energy costs. Additionally, intensive pond systems may require the physical and legal ability to discharge large quantities of water, some of which can be high in nutrients and suspended solids. However, current management practices are eliminating this last consideration. 5.4.1.2 Types of ponds and typical construction There are three principal types of pond construction methods, including excavated, embankment, and diversion ponds. Excavated ponds are dug completely below the natural surface of the land. Unless the ground is made-up entirely of heavy clay, excavated ponds usually are linked to groundwater sources and water levels will depend on groundwater levels. These ponds are difficult to drain or flush, may be seasonal because of the groundwater, and can be subjected to flooding. Embankment ponds are typically built to control runoff within a watershed. An embankment pond is constructed by building a berm (or dam) to trap surface waters as they flow from high points to low-lying areas in a watershed. Embankment ponds constructed to capture runoff from a watershed are not suitable for intensive aquaculture because they rely on runoff as a water supply, can be influenced by pollution from other sources within the watershed, and are prone to flooding. Most ponds used for intensive aquaculture are diversion ponds, or ponds that are a combination of excavation and embankment ponds (Figure 5.4). Outside sources of water are diverted away from the pond, including surface and ground waters, thus the operator has more control of the water in the pond. Diversion ponds are normally constructed in relatively flat areas (having a slope of (1-3 %) in one direction) where there is sufficient clay at a sufficient depth to build pond banks that will hold water with a minimum of seepage either down into the ground or laterally through the berms. When there is insufficient clay to avoid seepage, ponds can have a clay core put in the berms or can be lined with a layer of clay or with an impervious liner such as plastic or rubber. Intensive culture ponds need to be located in areas with an abundant supply of high quality water that is inexpensive to pump into the ponds. A general rule

141

Fig. 5.4. Schematic of a typical aquaculture pond. of thumb is 6.3-9.4 liters per second per surface hectare. The water supply can be either ground water or surface waters that are easily treated to remove unwanted materials, including sediment, other aquatic organisms, or harmful chemicals. See pond construction manuals such as Design and Construction of Diversion Pondsfor Aquaculture (Mittlemark and Landkammer, 1990) for more details on pond construction. The size and depth of intensive culture ponds for striped bass production will vary according to the life stage being produced, the local climate, available land, and the preferences of the culturist. In general, smallersized fish will be grown in smaller ponds--both in area and depth. The usual objective is to maximize fish density per unit volume of system water, which also provides for ease of management (e.g., feeding) and

142

harvesting. Pond depths used for aquacultural production usually range from 0.75m to about 2.8m. The depth is chosen depending on winter conditions, species, and other factors. The minimum pond depth is usually chosen to minimize light penetration to the pond bottom to keep aquatic plant growth minimal (Brune, 1991) and is sloped to prevent wading birds from preying on the crop. Shallow ponds are used when fry and fingerlings are being reared to minimize the volume of water needed to be treated for enhancement of plankton blooms, to promote solar warming, and to facilitate easier harvesting of the smaller fish. Shallow ponds are also used if the available water supply quantity is limited and in warmer climates where thick ice covers are not a problem to overwintering fish. As fish get bigger, they can be grown in larger, deeper ponds. The 2.8m maximum depth facilitates easier harvesting and aerating. Deeper ponds are harder to keep uniformly mixed and can become temperature stratified. Ponds that are stratified are harder to aerate and maintain at optimal dissolved oxygen levels. Some ponds can even become anaerobic at the bottom. Stratification is a problem, particularly in summer months when thunderstorms can cause a stratified pond to overturn (flip) and reduce dissolved oxygen levels to dangerously low levels. Pond design considerations for striped bass were summarized by Warren et al. (1990) and Smith et al. (1990): 9 Ponds for fingerling culture should be 0.2 to 0.4 surface hectares in size with a minimum depth of 0.75m gradually sloping to 1.8m at the harvest end. Pond depths should be no greater than 2.8m; 9 Ponds for production of larger fish should range in size from 0.4 to 2.0 hectares with similar depths as for fingerlings as long as proper management control can be maintained; 9 Pond shapes for larger fish grow-out can include square and rectangular shapes, as well as circular and wedge shaped ponds. However, ponds should be relatively uniform in size and shape to facilitate harvesting and maintenance, as well as to reduce construction costs; 9 All production pond berms should contain soils that allow for 2.5:1 to 3:1 side slopes. Steep slopes reduce earth movement and lower construction costs. Also steep slopes on inside berms reduce shallow depths that promote emergent weeds and wading bird predation. Shallower slopes (>_.3:1) on outside berms allow for safe mowing; 9 There should be a minimum freeboard (i.e., height of the berm above the water line) of 0.6m in all production ponds, although more freeboard may be required for larger ponds; 9 Include access up and down the berms to the harvest structure, such as steps or walkways; 9 Berm tops should be graded and preferably covered with gravel to allow all weather access for feeding, observation, and harvest operations; 9 Ponds should be oriented to take advantage of optimal solar radiation and wind velocities to optimize heating (or cooling) and maximize natural aeration; 9 Pond water inlets should be placed to optimize pond flushing capabilities during periods of poor water quality and to facilitate harvesting; 9 Size fill pipes to allow for rapid pond filling (24 hours is ideal); 9 Drainage systems should be located to reduce piping requirements and to reduce the number of settling ponds necessary to improve discharge quality; 9 Size drain lines to allow ponds to be rapidly drained (again, 24 hours is ideal) and include considerations for the labor required during the draining operation; 9 Include dam boards and/or gate or butterfly valves to adjust water flows through the drain structure; 9 For smaller fish, include concrete harvest kettles (catch basins) to facilitate rapid harvesting and reduce fish stress during harvest; 9 Electricity (both 110 and 220 volt service single phase and possibly three phase) should be available along the pond banks to power equipment such as pumps or aerators, especially during harvestwsome operations may want to consider portable generators as their electrical source, especially if located in a remote area where running power lines would be prohibitively expensive.

143

5.4.1.3 .Components of intensive pond systems Intensive pond systems include one or more ponds; a water supply; drains or other water level controls; aeration, feeding, and harvesting equipment; and a means to treat the pond effluent. For those pond systems that require effluent treatments, additional equipment or processes are needed to remove suspended solids and/or nutrients from discharged waters. The next several sections will discuss the various components of intensive pond systems.

5.4.1.3.1 Water supply Successful intensive pond systems must have a continuous supply of good quality water in sufficient quantity to meet system demands. Water quality will depend somewhat on the life stage being reared, but in general will need to be free from harmful chemicals, sediment, and biotic components that can be harmful to the striped bass being cultured and have an adequate oxygen content. The water quality requirements of striped bass are not totally understood (Warren et al., 1990). Table 5.2 provides water quality recommendations for general fish culture that should be applicable to striped bass culture. The 9 9 9 9

quantity of water needed for intensive pond culture of striped bass includes water needed to : fill ponds; make up water losses from evaporation and seepage; flush ponds during periods of poor water quality or low dissolved oxygen; and reduce stress to crowded fish during harvesting.

Warren et al. (1990) recommended a minimum of about 75 liters per minute per surface hectare. Calculations must, however, be made to determine the pumping requirements needed to fill any ponds that may be drained as a normal part of operating the pond system. For example, if several ponds are to be drained after harvesting phase II fingerlings, the water quantity requirements should be calculated to ensure that these ponds can be refilled before restocking.

5.4.1.3.2 Drains All ponds used for intensive aquaculture systems must have adequate drains installed to allow the operator to adequately control the water level in the pond. Drains must be sized so that ponds can be drained rapidly for harvesting, preferably in less than 24 hours. According to Mittlemark and Landkammer (1990) the drainage system must: 9 allow water to be released when necessary and prevent release under normal operating conditions; 9 allow the pond to be drained when necessary; 9 act as an overflow; and 9 allow the operator to draw water from the bottom during pond flushing operations. Drains should be constructed so operators can easily release water from the pond for management purposes, such as drawing down the pond for harvest or for flushing poor quality water from the pond. For example, during periods of low dissolved oxygen or poor water quality, drains must also allow the operator to withdraw water from the pond bottom at a location away from the water supply inlet where the water will almost always be the poorest quality in the pond. Overflows are important to keep rainwater from overfilling a pond.

144

There have been a variety of drain structures developed for pond aquaculture systems. Most include a component for water level control, overflow protection, screens to prevent fish escape, and some kind of valve assembly to keep water in the pond. Except in the southern-most climates, all pond drain structures should be located internally within the pond to prevent freezing and to allow for the operator to use the structure all year long. Controls for drains should be conveniently located for the operator to provide quick and safe access. The internal drain structures include swivel pipes, canfields, monks or other riser-types of structures, and valves. Selection of a particular drain structure is usually based on personal preferences of the operator and costs. The selected drain is acceptable as long as the structure meets the above mentioned requirements. 5.4.3.1.3 Aeration Intensive pond culture systems rely on aeration equipment to mix and oxygenate pond waters. As stocking densities increase, natural aeration cannot continue to meet pond oxygen demands. These demands include respiration requirements of the fish, oxygen for the breakdown of nitrogenous and carbonaceous wastes, and respiration requirements of phytoplankton in the pond. The carrying capacity of a pond without aeration is typically 35-40 kg of feed per hectare without water quality problems (Boyd, 1982). At these densities, natural aeration can usually meet the pond oxygen demands. In addition to meeting the pond oxygen requirements, aeration equipment should be capable of keeping the pond completely mixed to eliminate any thermal stratification and the problems that may result from it. A variety of aeration methods have been proposed for ponds, with surface agitators being the predominantly used method. Surface agitators include paddlewheels, spray-type agitators, propeller-air injectors, and pumps. (see Chapter 3). All of these are commercially available and have published performance data (Boyd, 1982). Other methods for aerating ponds include airlift pumps, diffusers, and oxygen injection systems (Parker et al., 1990). Of these methods, paddlewheel aerators are probably the most prominent method used, especially in larger ponds. Airlifts and diffusers are showing promise as alternatives to surface agitators, especially considering the energy savings that can be achieved. See Parker et al. (1990) for a more complete discussion of designs, loading requirements, and the benefits and disadvantages of several types of pond aeration devices.

5.4.1.3.4 Harvesting There are several methods for harvesting striped bass from ponds, and the methods vary according to the life-stage being harvested. For production of phase I fingerlings, a harvest structure is usually built in conjunction with the drain structure of the pond. This harvest structure, or kettle, allows for the pond to be drained and for the small fish to be captured in the kettle. The fish are then seined from the kettle and transported to their next location. While in the kettle, additions of fresh water and aeration can be used to reduce stress to the fish. For larger fish, ponds are usually seined to crowd the fish. Seining larger ponds requires large seines and heavy equipment, such as tractors, to pull the seine along the banks of the pond (see Chapter 3). Large seines are usually stored on large, mechanically-operating drums or reels. Once the pond is seined--either the entire pond or a portion of the pond for a partial harvest--the fish are crowded as the seine is drawn tighter. Some culturists use a seine and attach a live car (holding net) to the bag end of the seine once the fish are crowded. The crowded fish are then dipped out of the pond with large hand-held dip nets, boom mounted dip nets, or they can be removed from the pond with a piscilator or other commercially available harvesting aids. Hand netting is very labor intensive and can be strenuous with larger fish. Boom mounted dip nets (Figure 5.5)

145

Fig. 5.5. Boom operated dip net for harvesting fish from ponds.

offer several advantages including rapidly moving a large volume of fish and direct weighing as the fish are harvested. Some culturists use piscilators, fish elevators (Figure 5.6), to move fish from a pond to either other ponds or to harvest trucks. The piscilator is either a large rotating drum with internal vanes or a fixed drum with an internal auger to move water and fish. Piscilators are becoming more popular, especially when coupled with graders. The major advantage of using piscilators is the minimal damage done to the fish during the transference form the pond and the lower amount of physical labor required to handle and grade fish. This is especially advantageous when moving large fingerlings from smaller ponds into final grow-out ponds. Intensive striped bass pond systems must be capable of being completely harvested at the end of a growing cycle. Because striped bass are cannibalistic, leaving one or more larger fish in a pond can be very detrimental to future crops in the same pond. Most striped bass culturists completely drain production ponds after a crop is harvested. This allows for the capture of any fish that were missed during the seining operation. The pond bottom can be cultivated to oxidize some of the accumulated organic matter before refilling. Proper pond design can aid in harvest operations. Ponds that have gradually sloping bottoms and regular sides are much easier to completely seine than those with irregular sides and bottoms. Another design feature that facilitates harvesting is water supply inlets at the harvest ends of the ponds. This feature allows the operator to add fresh, oxygenated water to the crowded fish thereby reducing stress.

146

5.4.1.3.5

Feeding

Intensive pond systems require large and mechanized equipment to handle the volumes of feed needed on a daily basis. Most larger intensive operations use a tractor-drawn, blower-type feeder for delivering feed to the ponds (Piper et al., 1982, see Chapter 3). Typical feeding routines with tractor-drawn blowers consist of an operator driving down two pond banks while blowing feed out into the pond (Smith et al., 1990). As the feed is broadcast out into the pond, the operator can observe feeding behaviors and reduce feeding rates when fish are not actively feeding. Decreased feeding levels can also be used to indicate potential problems and direct observations of feeding can serve as a valuable warning. Feed blowers also broadcast feed over large areas, which reduces competition among the fish to produce more uniformed-sized fish (Smith et al., 1990). Commercially-made blowers are available, although many culturists configure their own blowers with a variety of options to make the feeding tasks easier. These modifications include tractor-mounted controls to operate the blower motor and feed shut-off, scales for the hopper (with displays on the tractor) for easier recording of amounts of feed delivered to each pond, and electrical or hydraulic actuators to adjust the feed delivery tube (Varadi, 1984). Other feeder types can be used for intensive ponds, including demand and automatic feeders. Demand feeders (Figure 5.7) are activated by the fish and have the potential advantage of providing only the amount of feed required by the fish (i.e., the fish only eat when hungry) (Zeigler et al., 1984). Striped bass and hybrids are sometimes not easily trained to use demand feeders in ponds (Smith et al., 1990). Multiple demand feeders must be used in ponds so that larger, more aggressive fish do not dominate the feeding site (Robinson and Wilson, 1985). Automatic feeders are designed to broadcast feed out into a pond at preset intervals, without the presence of an operator. With automatic feeders, feed can be presented to the fish at multiple times during the day. Demand and automatic feeders have the disadvantage of removing operator observation from the feeding routine (Robinson and Wilson, 1985). As previously stated, decreased feeding can be a sign of fish

Fig. 5.6. Piscilator for harvesting fish from ponds.

147

Screw

Fig. 5.7. A demand feeder. stress, which will not be observed if operators are not present during feeding. Automatic and demand feeders can also clog and do not feed properly unless constantly maintained (Smith et al., 1990). Regardless of the type of feeder used, it is important that feeding rates be closely monitored. Underfeeding results in lost production and can be as economically damaging to the operation as overfeeding where feed is wasted (Piper et al., 1982). Overfeeding can also result in poor water quality as decomposing feed contributes increased ammonia loads and additional biological oxygen demand to the pond. Careful attention must be given to the feeding rates on a daily basis to achieve maximum production during the growing season (Piper et al., 1982). Storage of feed is also an important consideration for intensive pond culture operations. Because formulated feeds contain high percentages of protein and oils, proper storage is essential to prevent spoilage (Piper et al., 1982; Robinson and Wilson, 1985). Feeds rfust be stored in cool, dry places to prevent spoilage and the formation of molds. Some molds produce toxins that are lethal to fish and rancid oils can also lead to fish health problems (Piper et al., 1982). At the very least, improperly stored feeds will lose some nutritive value and decrease production performance and increase costs. Feed handling considerations must also consider the fragile nature of feed pellets. Improperly handled pellets will break and produce large amounts of fines, which are not readily accepted by larger fish. Feed is typically stored in elevated bins (Figure 5.8) so that it can be loaded directly onto the feeding distribution equipment. Bin capacity should not exceed on-farm storage of greater than 90 days under optimal storage conditions, less for suboptimal conditions (Piper et al., 1982). Feed can also be purchased in bags (either multi-walled paper or plastic), but for larger operations the handling of bags can become a labor-

148

intensive proposition. Feeding and feed handling at a large, intensive operation is a materials handling problem that requires proper equipment for the safe and efficient handling of the large quantities of feed required on a daily basis. The equipment includes properly erected bulk storage bins, feeders of sufficient size or number to allow the operator to feed all of the ponds in a reasonable time period, and tractors that are large enough to handle the intended loads. Pond bank tops also need to be of sufficient width that tractors and feeders can travel safely around the ponds, even in inclement weather. See Chapter 3 for further details on feeding.

Boo

Fig. 5.8 Feed storage bin for bulk storage of fish food. 5.4.2 Flow-through Systems Flow-through or single pass systems rely on increased water flows to maintain water quality throughout the system. As a result, they must be built in areas having abundant, inexpensive water supplies that are at desired temperatures. Flow-through systems also require oxygenation, and discharge waters must either be treated to remove suspended solids and nutrients or discharged to a facility that can use these constituents (e.g., agriculture, golf course, etc.). Intensive systems that use flow-through technologies use the flowing water to reduce metabolite concentrations in the system (Wheaton, 1977). Another important advantage of flowthrough systems is that they often force the fish to swim, which produces healthier and leaner fish. Flowthrough systems consist of culture tanks or raceways, aeration systems, water conveyance systems, feeding equipment, and harvest equipment. For those flow-through systems that require effluent treatments, additional equipment is needed to remove suspended solids and/or nutrients from the discharged waters. Water is often reused in flow-through systems until it becomes ammonia limiting to the fish and then it is discharged. The amount of water needed for a particular system configuration can be determined based on the amount of ammonia produced per unit of feed given to the fish. As a rule ofthumb, lkg of feed yields 0.03

149

kg of ammonia (Wheaton et al., 1994b). This relationship includes breakdown of uneaten feed and fish metabolites and can be used to determine the total pounds of fish that can be supported for a given flow of water. Additional calculations are made to determine the oxygen requirements of the system. The oxygen requirements for the oxidation of nitrogenous and carbonaceous wastes and fish respiration can be calculated as approximately 0.21 kg of oxygen per kg of feed fed (Watten, 1991; Wheaton et al., 1994b). Some flowthrough systems, such as series raceways, reoxygenate the system water one or more times before discharge to maximize water use. Care must be taken in design of such systems to assure that fish near the end of the raceway get adequate water quality to achieve maximum growth rates. Flow-throu~ system design requires species-specific information including behavioral, swimming speed, oxygen requirements, rate of fecat settling, responses to temperature and tolerance to pollutants (Youngs and Timmons, 1991). Water velocities in flow-through systems should be sufficient to provide waste removal (especially solids), but below allowable cruising swimming velocities for striped bass (Wheaton, 1977; Youngs and Timmons, 1991). Some other design considerations for flow-through systems according to Youngs and Timmons (1991) include: 9 System size is determined by topography, soil type, construction materials, and total water available; 9 Ease of access and protection from predators is necessary; 9 Systems operated in series often require water quality enhancement (e.g., aeration and solids removal) between raceways to maintain optimal growth conditions; 9 Ammonia concentration is the major water quality factor that determines system capacity. Ammonia accumulation should be calculated for maximum stocking densities to determine system water volume requirements; 9 Systems should be designed such that individual culture tanks can be isolated for treatment of diseases or cleaning purposes. Advantages of flow-through systems include the ease of handling and feeding the fish, maintaining water quality, and exercising the fish (Youngs and Timmons, 1991). Circular tanks can be designed to be selfcleaning and to provide sufficient current to exercise the fish (Youngs and Timmons, 1991). Design information for flow-through systems can be found in Wheaton (1977), Youngs and Timmons (1991), and Westers (1991). 5.4.2.1 Types of flow-through systems Flow-through systems typically use either raceway, circular tanks, or other regular shaped tanks as culture chambers (see Figure 5.9). Raceways are long rectangular tanks--usually they are constructed so that the length is considerably greater than the widthmand are rather shallow in depth. Dimensions of raceways vary, but a length:width:depth ratio of 30:3:1 has been popular throughout the United States (Piper et al., 1982). Circular tanks used for striped bass culture include fry rearing tanks that are typically about one meter in diameter and about one meter in depth, to large, permanently installed units 10 m or more in diameter and 2-3 m in depth. Other configurations have been used for a variety of species, including oval tanks, cross-flow rearing units (rectangular raceways fitted with flow manifolds to produce a side-to-side flow as compared to flows along the length of the racewayusee Watten and Johnson, 1990 for more information), Swedish Ponds (a square tank with rounded comers that operates much like a circular tank--see Wheaton, 1977 or Piper et al., 1982 for more details), and recirculating rectangular ponds (Wheaton, 1977; Piper et al., 1982). The exact

150

Fig. 5.9. Schematic of raceway (a) and circular tank (b) flow-through systems.

151

design of the tanks or raceways often defers to species preferences and personal preference, but any design should allow for efficient water movement through the tank, minimize dead spots (portions of the tank or raceway with little or no flow rate), and be easy to construct and maintain. Tanks should also be smooth on the inside to prevent damage to the fish and reduce bacteria harborage. Design of a flow-through system can use either parallel or series water flows. Parallel designs pass the water through one raceway or tank and then discharge it. Advantages include minimal head loss (reduced pumping requirements), less buildup of metabolites, and efficient aeration (Wheaton, 1977). Disadvantages include increased flow requirements for a given mass of fish, which could mean increased pumping costs and a greater volume of water required for the operation. Series designs pass water from the outlet of one tank or raceway to the inlet of another tank or raceway. This requires some elevation difference between the successive tanks or raceways to maintain water flow, otherwise water must be pumped. Series arrangements can use natural aeration between units to minimize operational costs (Wheaton, 1977), but suffer from increasing ammonia concentrations from one unit to the next. The capacity of flow-through systems is a function of the most limiting water quality variable, typically oxygen or ammonia. Design information such as that presented in Losordo and Westers (1994) present several options for flow-through systems, including single pass systems, serial reuse of water in multiple raceways or tanks, serial reuse with added oxygen, and serial reuse plus oxygen and biofiltration. Designs are based on the metabolic requirements of the fish, with oxygen consumption and ammonia mass balances predominating the design considerations. Examples of possible production capacities for flow-through systems are 41,000 kg annually at a maximum density of 110 kg/m 3 with a flow of 51,000 L of water required per kg of annual production in a system with serial reuse and oxygenation (Losordo and Westers, 1994). Using similar assumptions (i.e., 4,000 L/pm available water, and the production of 630 g fish), Losordo and Westers (1994) estimated that a flow-through system with serial reuse, oxygenation, and biofiltration added to remove 0.2 g ammonia per square meter per day (see following sections on recirculation systems for more information on biofiltration) would be capable of producing 170,000 kg annually at a maximum density of 110 kg/m 3, with a flow of 12,000 L of water required per kg of annual production. 5.4.2.2 Components of flow-through systems 5.4.2.2.1 Water supply Water quality requirements for flow-through system water sources are similar to those for ponds. Either groundwater or surface water sources can be used for the production of striped bass in flow-through systems. Two primary differences in water supplies between pond systems and flow-through systems are temperature and quantity. Flow-through systems require about two orders of magnitude more water volume to produce a given quantity of fish than do pond systems (Stickney, 1994). Water temperature is also important when considering flow-through systems for striped bass. Water supplies that produce a continuous supply at the proper temperature for the life stage(s) being produced is essential for successful operation of flow-through systems. Water temperature adjustment is usually not economically feasible because of the large water volumes required in flow-through systems. Therefore, a source that is free from pollutants, is the right temperature, and provides a consistent volume is required. In most situations, this will require a groundwater source. 5.4.2.2.2 Drains Flow-through system drains should be designed to maintain approximate plug-flow conditions throughout the tank, which is not possible in circular tanks. Plug flow can be defined as water throughout a given crosssection of the tank flowing at a constant, non-turbulent velocity. Plug flows eliminate dead spots where solids

152

accumulate and dissolved oxygen levels are below critical values (Wheaton, 1977). Typical drains for raceways are constructed from dam boards to allow for a uniformly horizontal flow from the system. This promotes plug flows in the tanks and allows for adjustments to the system water depth. Other tank configurations (e.g., circular, oval, etc.) use drain structures that give adequate circulation throughout the entire tank volume to eliminate dead spots. When standpipes are used, both external and internal standpipes add redundancy to help prevent complete draining when larger fish dislodge the internal standpipe. Sleeves over internal standpipes also help to promote self-cleaning (see Figure 5.9b). 5.4.2.2.3 Tanks The tank shape in flow-through systems is typically rectangular for raceways, but can be circular or oval. The most important consideration in tank shape is choosing a tank design to minimize stress on the fish. Circular or oval-shaped tanks are generally recommended in striped bass culture (Nicholson et al., 1990; Yeager et al., 1990), because they offer unrestricted swimming paths for the fish. Additionally, tanks must be self cleaning, have minimal dead space, and be harvestable (Wheaton, 1977). Tanks and raceways should be constructed from materials that are chemically inert with the culture water, provide a smooth finish, and are relatively inexpensive. Concrete (often coated with a smooth finish), fiberglass, and earthen structures are often used for raceway and other tank construction. Other materials used for construction include plastic, wood or metal with plastic liners, and in some cases aluminum. Ideally, culture tanks will have the following characteristics (Wheaton, 1977): 9 finished with smooth interior surfaces to prevent abrasive damage to the fish; 9 designed to be self cleaning to remove solid wastes; 9 uniformly provides high quality water throughout the tank; 9 has sufficient mechanical strength to withstand any stresses from water filling, backfilling, or moving the tank; 9 is durable to last many growing seasons and resistant to weathering and ultraviolet degradation; 9 easily cleaned and sterilized; 9 has nontoxic interior surfaces that do not harbor disease organisms; 9 are inexpensive to purchase and construct; and 9 will not corrode under normal operating conditions. 5.4.2.2.4 Aeration Aeration is an important consideration in flow-through systems. Aeration methodologies can be completely passive aeration systems that rely on gravity to aerate and reaerate water as it flows through the tanks or raceways. Examples of these aerators include weirs with splash boards, transversely corrugated inclined planes (with and without holes), lattice aerators, perforated tray cascade aerators, and risers with perforated aprons (Wheaton, 1977). These types of aerators limit the amount of oxygen that can be added to the system and rely on elevation differences to accomplish the aeration. Other types of aerators used in flow-through systems include surface aerators or agitators, packed columns and spray towers, U-tubes4, and several configurations of liquid oxygen injection. Surface agitators have been successfully used to add oxygen from atmospheric air and pure oxygen when fitted with a cover (Watten, 1994). Spray towers and packed columns both with and without pure oxygen supplies can be very efficient for oxygenation and removal of other gases, especially nitrogen (Watten, 1991, 1994). U-tubes (Figure 5.10a) 4 Consists of a gas diffuser, a vertical u-shaped conduit to provide oxygen-water contact at greater than atmospheric pressures, and off-gas collection. Oxygen and off-gas are diffused into water entering the u-tube. The oxygen-water mixture flows down one side of the U and back up the other. While in the lower portion of the U. the oxygen-water mixture experiences increased hydrostatic pressure, which accelerates oxygen absorption (Boyd and Watten 1989).

153 have been found to be very efficient for oxygenation of flow-through system waters, but must be carefully designed to operate properly and efficiently. Several other pure oxygen configurations have been used in flowthrough systems, including side stream oxygen injection, low-head oxygenators (Figure 5.10b), and down-flow bubble contactors (Watten, 1994). Oxygenation and aeration systems must be designed to be reliable and cost-effective. Because many flowthrough systems rely on supplemental aeration or oxygenation to accommodate increased carrying capacities,

Air

II,

Off-G~s Off-

Low

| Purnp I

D0 Water

~ ~ 1 1

K o o o o - =--J--"" | o o o%~176 % ~ 0o ~

o ::

Diffuser

II

| ~ - . H i g h DO ~,,...-'-">; Water

~Oo~ 0 ~

0

" Io~ I 0 0 o

~ ~ol~

D ~ 1 7 60 Oo --~1o i ~

H:0R

Low DO Water - - - - ~ ~ . ~

Air - - ' ~ - -

Independent Spray =~

'b

~

-

L

~

_ 2;

2

-

;

-ZI

I

_~ 7-

22 2

. -__

.

. j

~J

High DO Water

Fig. 5.10. U-tube aerator (a) and low head oxygenator (b) used in raceways.

Gas Vent

154

it is essential that these vital components are adequately designed and have sufficient redundancy or emergency backups available in the event of system or component failures. The system must also provide sufficient oxygen to the fish at a reasonable cost--both capital and operational (Watten, 1994). 5.4.2.2.5

Harvesting

Harvesting of fish in flow-through systems is usually accomplished by crowding the fish into one part of the tank. Fish are then dipped out of the tank with nets or pumped out. Handling of the fish for grading and harvesting in flow-through systems is often overlooked in the design phase and is an area that needs more attention. 5.4.2.2.6

Feeding

Feeding equipment for flow-through systems is similar to that used in pond systems. Demand feeders have been used by some flow-through system operators, but are not the most popular system. Feeding can be accomplished by hand feeding, or the use of a variety of mechanical and automatic feeders. Hand feeding is labor intensive and only practical for small systems. Like demand feeders, hand feeding is not too common in most commercial systems. Mechanical feeders are typically used in flow-through system culture operations. Some types of mechanical systems used include towed blowers, stationary broadcast or blower feeders, or automated feed delivery systems (Figure 5.11). Most types of feeding systems are commercially available and can be designed with the help of feed manufactures or equipment dealers. Some important considerations for mechanical systems include minimal grinding of the feed (which produces unwanted fines), uniform delivery of the feed to the culture tanks, adequate cleanouts to alleviate clogged lines, manual overrides for automatic systems, and reliable operation under varying weather conditions. Feed in mechanical systems must also be protected from excessive moisture to prevent spoilage and mold growth that could be toxic to the fish.

Fig. 5.11. Example of an automated feed delivery system.

155

5.4.3 Recirculating Systems Recirculating systems (Figure 5.12) are defined as those systems that recycle water (i.e., water that is treated and reused in the same culture tank) without exchanging the water with new water from outside sources (Wheaton, 1977). Recirculating, or closed, systems offer several potential advantages over pond or flowthrough systems, including extended (year-around) growing seasons, minimal water and land requirements, effluent control for discharged waters, and more control over water quality (Wheaton, 1977; Losordo and Timmons, 1994). As land and water become more expensive and markets more demanding, recirculating systems should become more competitive with pond and flow-through systems for the culture of striped bass. Depending on the system configurations, energy requirements in closed systems may be less than those for flow-through systems that rely on pumps to move water through the system (Wheaton, 1977). Perhaps the most important benefit of recirculating systems is the ability to control system water quality, especially temperature. Because most of the water is recycled, it becomes more cost-effective to heat or cool the system water than can be done in ponds or flow-through systems. Thus, unless ponds can be located in geographic areas that provide optimal temperatures for striped bass production or flow-through systems have a continuous source of high quality water that is constantly at optimal growth temperatures, recirculating systems offer the only economical solution for continuous production throughout the year.

- -

..,.

CULTURE

TANK ,,

,,,

,,

l

1

1

GAS EXCHANGER

t

I~fiow Fig. 5.12. Schematic of recirculating system.

Dischorge

.,o

1

156

Complete (100%) recycle systems are nearly impossible to achieve as some water is lost to evaporation and the treatment processes. Evaporation is difficult to control, even when tanks are covered. Evaporation rates will vary according to ambient relative humidities and air exchange rates in buildings housing recirculating systems. As the system water is treated to remove fish metabolites, some water may be lost in processes such as solids removal. Many systems are operated to exchange a small percentage of water (5 to 10%) on a daily basis to compensate for inefficiencies in the treatment processes and maintain ion balances in the water. Recirculating systems allow diseases to be controlled and treated more economically than do ponds or flow-through systems (Wheaton, 1977). Fish to be raised in recirculating systems can be isolated before introduction into a system for quarantine purposes. Recirculating systems can also be designed to allow operators to isolate fish for treatment. Other design aspects ofrecirculating systems that improve performance include modifications to make management tasks easier, such as feeding, grading, and/or harvesting. There are several disadvantages in recirculating systems. Most importantly, recirculating systems have high capital and operating costs compared to ponds and some flow-through systems. For example, Losordo and Timmons (1994) estimated that commercial scale recirculating systems have investment costs on the order of $4.00 to $8.00 per kg of annual production capacity. This compares to investment costs for commercial ponds or raceways of about $2.20 to $3.30 per kg (Losordo and Timmons, 1994). Operating costs for all three types of systems will vary according to site-specific factors such as feed costs, land and building costs, electricity and other energy rates, water treatment costs (for both supplies and effluents), and personnel requirements. Recirculating systems tend to have greater technical skill requirements for operational personnel than do pond and flow-through systems, but this also depends on the degree of system sophistication. As recirculating system components become more reliable and economical, costs for producing fish in these systems will begin to approach or fall below costs for pond and flow-through systems (Losordo and Timmons, 1994). Assuming a desired production of 630g fish at 18~ Losordo and Westers (1994) estimated that a 90% recycle system (i.e., 10% daily water exchange) would be capable of producing 71,000 kg annually at a maximum density of 110 k#m s, with a flow of 120 L of water required per kg of annual production. Although designed for rainbow trout, Wheaton et al. (1994b) present design estimates for recirculating systems to produce 1 kg fish in 12~ water at a density of 50 kg/m 3. Hochheimer (unpublished data) has successfully produced hybrid striped bass at densities of about 60 kg/m 3 in a recirculating system operated at 20~ It is important to note that the production capacity of a recirculating system is a function of the filter design and densities and annual production rates will not necessarily be directly comparable to those in flow through or pond systems. Annual production capacities of commercially-sized recirculating systems are presently unavailable in the literature. 5.4.3.1 Components of recirculating systems Design ofrecirculating systems is similar to the design of flow-through systems for tank design, aeration, feeding, fish handling, and solids removal. Therefore, these aspects of recirculating systems will not be discussed here and the reader is referred back to the section on flow-through systems. Although water supply quantities will not be as great in closed systems as in flow-through systems, water quality requirements will be similar. The quality of water used to fill recirculating systems should be as good as water used for flow-though or pond systems (see Table 5.2). However, in recirculating systems municipal water supplies can be used as long as additives such as chlorine and fluorine are removed before use in fish

157

culture. Some pretreatment of water can be cost effective in recirculating systems, because the volume to be treated is rather small and treatment flows should also be low. Examples of pretreatment options include filtration with activated carbon, sterilization with ultraviolet lights, or removal of particulate matter. The most up-to-date design information for recirculating systems can be found in Aquaculture Water Reuse Systems: Engineering Design and Management (Timmons and Losordo, 1994). The design of a recirculating system should include: 9 Define the production goals and determine the system operating limits; include fish carrying capacities per tank to achieve desired production levels and calculate the necessary water flow rates and aeration rates to maintain acceptable water quality; 9 Develop a management strategy for the production levels desired; 9 Design the solids management system; based on feeding levels, calculate solids removal system to continuously remove solids from the system; 9 Design the biological nitrification system; determine the ammonia production rates, select a biofilter type, and calculate the size of biofilter needed to meet the production goals; 9 Design the aeration system to provide sufficient oxygen for the production levels desired; 9 Determine if carbon dioxide control is needed in the system; if so, design a carbon dioxide stripper; ~ Determine pH and alkalinity requirements for the desired production levels; develop a process for adding alkalinity to the system to maintain the desired alkalinity and pH levels; 9 Design a system to remove fine solids from the culture water; 9 Develop an alarm and monitoring system for the appropriate, key water quality and operational variables; ~ Develop a final management strategy based on the desired production levels and final system design. 5.4.3.1.1 Biofiltration Biological filtration (biofiltration) can be defined as any process that uses living organisms to remove unwanted substances from the culture system water (Wheaton et al., 1994a). Many different processes have been developed that could be included in a discussion of biofiltration, but the primary design purpose of biofilters in recirculating systems is the removal of ammonia. These other processes include denitrification (the conversion of nitrate into nitrogen gas), removal of organic carbon by heterotrophic bacteria, and removal of nitrogenous compounds by plants (Wheaton et al., 1994a). Ammonia accumulation in recirculating systems becomes the most limiting factor after oxygen requirements are met. Nitrification in biofilters is the process where ammonia is converted to nitrite and then into nitrate. Ammonia and nitrite are toxic to striped bass at relatively low levels (ca. 1-10 mg/L, depending on many other water quality variables) while nitrate is relatively nontoxic to striped bass, even at levels exceeding 200 m g ~ (Nicholson et al., 1990). Although many species of bacteria are responsible for the bioconversion of ammonia into nitrate, the two most important genera are Nitrosomonas for the conversion of ammonia into nitrite and Nitrobacter for the conversion of nitrite into nitrate (Hochheimer, 1990). Biofilters used for ammonia removal are simply chambers used to promote the colonization of nitrifying bacteria. Optimal conditions for bacterial growth include the presence of oxygen, the substrate to be removed (ammonia and nitrite), and carbon in the form of alkalinity. Conditions that reduce or hinder the growth of nitrifying bacteria include excessive sunlight, the presence of toxic materials, and pH extremes. Hochheimer (1990) and Wheaton et al. (1994a) present a complete discussion on the kinetics of nitrification, which includes information on optimal and suboptimal conditions for biofilters.

158

There are many different configurations that have been used for biofilters. They can be generally grouped as upflow (water flows into the bottom and out of the top) and downflow (water flows into the top and out of the bottom) (Wheaton, 1977). Some of the more common configurations ofbiofilters include (Wheaton et al., 1994a): 9 Submerged downflow (Figure 5.13) - water flows down into a bed of media, such as sand, gravel, or other porus material; 9 Submerged upflow (Figure 5.14) - water flows up through a bed of media, such as sand, gravel, or other porus material; 9 Trickling filters (Figure 5.15) - similar in design to submerged filters, except that the media is kept wet, but not submerged. Water trickles over the media to supply substrates to the bacteria; 9 B iodrums (Figure 5.16) - perforated cylindrical container filled with media that is mounted on a shaft and rotates through water to be treated; 9 Biodisk (Figure 5.17) - similar to biodrums, except that the media is circular plates that rotate through the water; 9 Fluidized beds (Figure 5.18) - tube partially filled with media, such as sand, that expands as water flows up through it; 9 Bead filters (Figure 5.19) - similar to a downflow fluidized bed only the media floats and the flow is downward. The selection of a biofilter depends on system specific requirements and often on preferences of the designer and/or operator. Detailed design information for several different biological configurations can be found in Wheaton et al. (1994b).

E " INFLOW

OUT FLOW WATER LEVEL CONTROL

I i,

Fig. 5.13. Submerged downflow biological filter.

159

OUT FLOW

......

~

~,'~!~

J

,.FLow

Fig. 5.14. Submerged upflow biological filter.

~

In Flow

i //~\ ~\\ ~/q~x ~ 4bX ~ 4~k, b//~\\ ~///'~\//(~\\

"6~0

0 o'.~'~'0" . . . . .~.,A ,,,.~ ~..:~.~ ~...,..,_~ . .... . ~ ~~ , .... ~.~..~

(~ (~ 0 ~) ~ "~ (~ u

'~ 0'

Out Flow Fig. 5.15.

Trickling biological filter.

Manifold

6 ( ~ ' ~ (~ O" 6 d . k r" ~ , ~ .. ~. . . . . ...~~. , . .,.. ~ ~.,,. . . , ~ ~:... . :~: . : ~

D

(~ 0 6

- S u p p o r t Plate

160

,------- M e d i o

Out Flow

Water

Level

r'n Flow

Fig. 5.16. Biodrum biological filter.

S

Fig. 5.17. Biodisk biological filter.

161

Out Flow

~

-

~ "

-

:

_: , ~

:..

.....

: . .:~.~,

FIuid F~d "','~".S o h d ~ : ~

,

~m |-

Distributor Plate

~,dP---.-- In Flow

Fig. 5.18. Fluidized bed biological filter.

Outflow

Hold- Down Screen

Screen

~ BEAD

Fig. 5.19. Bead filter.

Inflow FILTER

162

5.4.3.2 Management of_biofilters A properly designed biofilter will operate over a long period of time keeping ammonia and nitrite concentrations at levels that are not harmful to the fish. The filter's effectiveness can be measured by the reduction in ammonia and continued low levels of nitrite in the system (Petit, 1990). The continued effectiveness of well-designed biofilters in recirculating systems can only be achieved by proper management of the system and care of the filters. The following points were presented in Petit (1990) for the design and management of biofilters: 9 the water must be rich in nitrogen compounds with a C:N:P ratio of approximately 10:20:1; 9 the minimum pH should be in the range of 6.5-7.0; 9 the filter area-specific nitrification rate decreases as the concentration of ammonia decreases; since the desirable concentrations in fish culture systems is low (less than 1.0 mg/L total ammonia), the efficiency of the filter is therefore low (typically about 25%); 9 about 5 g of oxygen are required to oxidize each gram of ammonia, therefore a sufficient supply of oxygen is required for the biofilters in the system; 9 the minimum concentration of oxygen in the biofilters is about 3.0 mg/L; 9 most biofilters can treat approximately 0.5 to 1 g of ammonia nitrogen per square meter of filter media per day (the removal rate depends on temperature and other variables); 9 cleaning the filter media (of excessive solids from heterotrophic bacteria and accumulation of suspended solids) will result in the loss of filter efficiency that varies according to the degree of cleaning and the type of media; 9 smaller, lower-density media are easier to clean, will clog more, require more frequent cleaning, and suffer greater efficiency losses; 9 smaller media offer greater surface area per unit volume of media for the growth of bacteria, but become clogged more frequently, which reduces the flow of water and air in the filter; 9 the time of contact in the filter can be short (e.g., several minutes) in recirculating systems; and 9 in systems with soft water (i.e., low alkalinity) it is necessary to account for the acidification that results from nitrification by adding buffers such as sodium bicarbonate. The hydraulic loading rate is the amount of water pumped through the filter per unit area of filter top surface per unit time, usually expressed in m3/mE~ For a given filter configuration, changes in the hydraulic loading rate will result in changes to the velocity of the water flowing through the filter. One way of expressing this important design consideration in biofilters is the apparent velocity, which is the ratio of the hydraulic loading at the surface of the filter divided by the depth. The minimum apparent velocity should be between 3 and 5 m/h to avoid irreversible fouling and below a maximum of 17 m/h to avoid scouring of the biofilm (Petit, 1990). Other authors report minimum and maximum hydraulic loadings as important design factors. The minimum hydraulic loadings are set as the rates that keep all of the filter material wet, which is necessary for the bacteria to survive. Minimum hydraulic loadings are usually more limiting in trickling or submerged filters than in rotating filters (Wheaton et al., 1994a). In trickling filters, Grady and Lim (1980) recommended minimum hydraulic loading rates of 29 m3/mZ.d for randomly packed Norton Actifil| media and Roberts (1985) reported minimum hydraulic loadings of 32 to 55 m3/m2.d for random packed plastic pall rings in trickling filters. The maximum hydraulic loading rate is the velocity the scours bacteria from the filter media or that causes excessive head loss (Wheaton et al., 1994a). In trickling filters, Roberts (1985) reports a maximum hydraulic loading rate of 72 to 188 m3/m2.d and Grady and Lim (1980) found a maximum hydraulic loading rate of 234 to 350 m3/m2-d for Dow Surfpac| a fixed trickling filter medium. Both the maximum and minimum hydraulic loading rates will vary considerably depending on the media size, configuration, manufacturer,

163

material, and other variables such as the filter depth, shape, and type (Wheaton et al., 1994a). Refer to the manufacturer of a particular media type for pertinent information regarding hydraulic loading rates. A related design consideration for biofilters is the hydraulic residence time (or detention time). Traditional wisdom in biofilter desi~ called for long residence times for efficient operation (Petit, 1990). This reflects the operation ofbiofilters in wastewater treatment plants where the substrate loadings are considerably higher than those in aquaculture. Kaiser and Wheaton (1983) found that when low ammonia concentrations are present, shorter filter detention times (e.g., higher hydraulic loading rates) produce greater ammonia mass removal rates. Hochheimer (1990) reports that loadings in biofilters for aquaculture are in the range where they become substrate limiting for ammonia (or nitrite) rather than for oxygen as is the case for wastewater treatment. Because of the ammonia substrate limitation, Hochheimer (1990) concluded that biofilters would operate much more efficiently at higher hydraulic loading rates to overcome the mass loading problem associated with the low substrate concentrations. There are many variables that can affect the performance of biofilters in a recirculating system from a management perspective. These factors can be grouped as chemical (e.g., pH, salinity, alkalinity, oxygen, dissolved organics, and toxic compounds) or physical (e.g., particulates or temperature). The pH in recirculating systems can range from 6.0 to 9.0. Slow changes in pH over the range (slow enough to allow the bacteria to acclimate) will produce relatively minimal effects on nitrification rates. Rapid changes in system pH (i.e., 0.5 to 1.0 pH units) will greatly reduce the filtering efficiency of a biofilter. Ideally, the system should be maintained at pH levels at or just slightly below 7.0. This keeps high levels of unionized ammonia that occur at higher pH levels from affecting the fish and also maintains the ammonia in the ionized form; the form used by the bacteria. Somewhat related to pH is the system alkalinity, which serves two purposes--buffering and as an essential nutrient for the bacteria. Nitrifying filters require alkalinities of at least 1.5 meq/L to ensure maximum efficiency (Gujer and Boiler 1986). Recirculating (and most intensive) systems must be continually checked for alkalinity levels to keep both biofiltration process and fish health optimal. Alkalinity, preferably in the form of bicarbonate or carbonate (e.g., sodium bicarbonate or sodium carbonate), must be added on a regular basis to ensure adequate alkalinity for both nitrification and pH buffering. Other forms of alkalinity, such as calcium carbonate (limestone), calcium hydroxide (lime), other calcium products, or sodium hydroxide, may produce undesirable pH shifts or lead to the accumulation of excessive levels of calcium in systems. Systems with alkalinities of 200 to 400 mg/L (as CaCO3) can be considered adequate. As previously stated, biofilters require sufficient oxygen (generally about 5 g oxygen per gram of ammonia oxidized and at minimum concentrations of 3 mg/L) to function properly. Oxygen will become limiting in a biofilter as a result of saturation oxygen concentration, the concentration of organic substances in the water, and the total bacterial, fish and plant biomass present. The saturated dissolved oxygen content of water depends primarily on temperature and salinity, with higher temperatures and salinities decreasing the possible saturation concentration. The breakdown of organic substances and the respiration from the total biomass in the system compete for available oxygen levels and must be factored into the oxygen demand for the system. Trickling filters can compensate for some oxygen deficit problems as they are usually designed to be self aerating. Biofilters can be acclimated to operate in any salinity range tolerable by the fish. However, nitrifiers are intolerant to rapidly changing salinities and will not usually tolerate abrupt changes of more than about 5 ppt (Hochheimer, 1990). Treatments that require additions of salt should be made with caution to prevent shocking the biofilter.

164

Ammonia removal rates in biofilters are directly influenced by organic loading rates (Wheaton et al., 1994a). Grady and Lim (1980) reported that nitrification rates are optimal at BODs/TKN55 ratios of about 0.25. BOD5 levels below 30 mod%allow nitrifiers to become established in a filter, whereas at higher BOD5 levels, the slower growing nitrifying bacteria cannot compete with the heterotrophic bacteria that oxidize organic compounds in the system (Antonie, 1976). There are a variety of substances that are inhibitory or toxic to nitrifying bacteria, including copper, nickel, vitamins, amino acids, chromium, alcohols, silver, and many antibacterial agents used for the treatment of diseases in fish (Wheaton et al., 1994a). Kaiser and Wheaton (1983) reported that therapeutic levels of formalin, copper sulfate, potassium permanganate, and sodium chloride did not adversely affect nitrification in biological filters. Particulates in system waters can have several effects on biofilters (Wheaton et al., 1994a). Particles that are larger than the pore sizes in the filter media can clog the filter, and lead to reduced filtering capacity and efficiency. Some nitrifiers will grow on particles that reside in the system for extended periods of time and may actually perform the majority of nitrification occurring in the system (at the expense of the nitrifier populations on the biofilter media). If the system is flushed or filtered for particulates, the nitrifiers are removed from the system and nitrification may essentially cease for a period of time. Most of the particulates are made up of organic compounds that will break down rapidly in the system from heterotrophic bacterial activity. This break down consumes oxygen needed by the nitrifiers and fish in the system. There is considerable disagreement in the literature regarding the influence of temperature on nitrification (Wheaton et al., 1994a). In general, the nitrifiers will perform better at higher temperatures (e.g. from 30 to 35~ but can perform adequately at low temperature once acclimated. Like other operational variables, biofilters should be protected from rapid changes in temperature to prevent sudden efficiency changes in the nitrification rate. The operator must remember that the biological filter is a living system and must be managed to reflect the dynamics of the bacterial population. Abrupt changes in the quality of water flowing to the biofilter can disrupt its operation and lead to more serious problems for the fish being cultured in the system. Ideally, changes in temperature, salinity, pH or other water quality variables will be made slowly over the period of several days to allow the nitrifier populations to adjust. Perhaps the most significant management consideration should be made in the changes to feeding levels. Feed amounts should be gradually increased over time (usually no more than about 10% per day) as the fish grow, not incremented in steps on weekly or biweekly adjustments. Careful records should be kept for feeding levels and small adjustments made daily, instead of large changes made weekly. 5.4.3.3 Other comoonents Recirculating systems may contain one or more other system-specific components. These include pumps, heaters, sterilization equipment, aeration and oxygenation equipment, packed columns for carbon dioxide control, ultraviolet or ozone disinfection units, metering systems to add chemical supplements to control pH and alkalinity, foam fractionation units, water quality monitoring and control equipment, and alarms. There are several good references available that include up-to-date design information related to all of these components, including Wheaton (1977), Spotte (1979), Huguenin and Colt (1989), NRAES (1991), and Timmons and Losordo (1994).

5

Five-day biochemical oxygen demand (BOD~)/total Kjeldahl nitrogen (TKN)

165

References

American Water Works Association., 1951. Water quality and treatment. 2nd edition, cited in Wheaton, F.W. 1977. Aquacultural Engineering, John Wiley and Sons, Inc., New York. Antonie, R.L., 1976. Fixed biological surfaces-wastewater treatment: the rotating biological contactor. CRC Press, Cleveland, OH. Boyd, C.E., 1982. Water quality management for warmwater fish ponds. Developments in aquaculture and fisheries science, volume 9, Elsevier Science, Amsterdam. Boyd, C.E. and Martinson, D.J., 1984. Evaluation of propeller-aspirator-pump aerators. Aquaculture, 36: 283-292. Boyd, C.E. and Watten, B.J., 1989. Aeration systems in aquaculture. Reviews in Aquatic Sciences, 1: 425-473. Brune, D.E., 1991. Pond aquaculture. Pages 118-130 in Engineering aspects of intensive aquaculture. Northeast Regional Agricultural Engineering Service (NRAES-49), Proceedings from the Aquaculture Symposium, Comell University, Ithaca, NY. Carlberg, J.M., Van Olst, J.C., Massengil, M.J. and Hovanec, T.A., 1984. Intensive culture of striped bass: a review of recent technological developments. Pages 89-127 in J. P. McCraren, editor. The aquaculture of striped bass: a proceedings. Maryland Sea Grant Publication. UM-SG-MAP-84-01. University of Maryland, College Park. Chen, S., Stechey, D. and Malone, R.F., 1994. Suspended solids control in recirculating aquaculture systems. Pages 61-100 in M.B. Timmons and T.M. Losordo, editors. Aquaculture water reuse systems: engineering design and management. Developments in aquaculture and fisheries science, volume 27, Elsevier Science, Amsterdam. Colt, J. And Orwicz, C., 1991. Aeration in intensive culture. Pages 198-271 in D.E. Brune and J.R. Tomasso, editors. Aquaculture and water quality. The World Aquaculture Society, Baton Rouge, LA. Grady, C.P.L. and Lim, H.C., 1980. Biological wastewater treatment, Marcel Dekker, Inc., New York. Gujer, W. and Boiler, M., 1986. Design of a nitrifying tertiary trickling filter based on theoretical concepts. Water Research 20: 1353-1362. Hochheimer, J.N., 1990. Trickling filter model for closed system aquaculture. Ph.D. Dissertation. University of Maryland, College Park. Huguenin, J.E. and Colt, J., 1989. Design and operating guide for aquaculture seawater systems. Developments in aquaculture and fisheries science, volume 20, Elsevier Science, Amsterdam. Kaiser, G.E. and Wheaton, F.W., 1983. Nitrification filters for aquatic culture systems: state of the art. Journal of the World Mariculture Society, 14: 302-324. Kerby, J.H., 1993. The striped bass and its hybrids. Pages 251-306 in R.R. Stickney, editor. Culture of nonsalmonid freshwater fishes, 2nd edition, CRC Press, Boca Raton, FL. Kerby, J.H., Hinshaw, J.M. and Huish, M.T., 1987. Increased growth and production of striped bass x white bass hybrids in earthen ponds. Journal of the World Aquaculture Society, 18: 35-43. Lewis, W.M., Heidinger, R.C. and Tetzlaff, B.L., 1981. Tank culture of striped bass production manual. Fisheries Research Laboratory, Southern Illinois University, Carbondale.

166

Losordo, T.M. and Timmons, M.B., 1994. An introduction to water reuse systems. Pages 1-8 in M.B. Timmons and T.M. Losordo, editors. Aquaculture water reuse systems: engineering design and management. Developments in aquaculture and fisheries science, volume 27, Elsevier Science, Amsterdam. Losordo, T.M. and Westers, H., 1994. System carrying capacity and flow estimation. Pages 9-60 in M.B. Timmons and T.M. Losordo, editors. Aquaculture water reuse systems: engineering design and management. Developments in aquaculture and fisheries science, volume 27, Elsevier Science, Amsterdam. Mittlemark, J. and Landkammer, D., 1990. Design and construction of diversion ponds for aquaculture. Minnesota Sea Grant Extension Publication number AQUA 17.90, University of Minnesota, St. Paul. Northeast Regional Agricultural Engineering Service, 1991. Engineering aspects of intensive aquaculture. NRAES-49, Proceedings from the Aquaculture Symposium, Comell University, Ithaca, NY. Nicholson, L.C., Woods, III, L.C. and Woiwode, J.G., 1990. Intensive culture techniques for the striped bass and its hybrids. Pages 141-158/n R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Parker, N.C., Klar, G.T., Smith, T.I.J. and Kerby, J.H., 1990. Special considerations in the culture of striped bass and striped bass hybrids. Pages 191-216 /n R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids, Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Petit, J., 1990. Water supply, treatment, and recycling in aquaculture. Pages 63-196 in G. Bamabe, editor. Aquaculture, volume 1, Ellis Horwood Series in Aquaculture and fisheries support, Ellis Horwood, New York. Pillay, T.V.R., 1990. Aquaculture: principles and practices. Fishing news books, Blackwell Scientific Publications, Ltd., Oxford, England. Piper, R.G., McElwain, I.B., Orme, L.E., McCraren, J.P., Fowler, L.G. and Leonard, J.R., 1982. Fish hatchery management. U.S. Department of the Interior, Fish and Wildlife Service, Washington, DC. Roberts, J., 1985. Mathematical models for trickling filter process. Pages 243-324 in S.E. Jorgensen and M.J. Gromiec, editors. Mathematical models in biological wastewater treatment. Elsevier, Amsterdam. Robinson, E.H. And Wilson, R.P., 1985. Nutrition and feeding. Pages 323-404 in C.S. Tucker, editor. Channel catfish culture. Developments in aquaculture and fisheries science, volume 15, Elsevier Science, Amsterdam. Seafood Leader., 1995. Striped bass update. Seafood Leader. 15(2):140. Smith, T.I.J., Jenkins, W.E. and Snevel. J.E., 1985. Production characteristics of SB (Morone saxatilis) and F~, F: hybrids (M. saxatilis and M. chrysops) reared in intensive tank systems, Journal of the World Aquaculture Society, 16: 57-70. Smith, T.I.J., Jenkins, W.E., Stokes, A.D. and Smiley, R.A., 1989. Semi-intensive pond production of market-size striped bass (Morone saxatilis) X white bass (M. chrysops) hybrids. World Aquaculture, 20(1): 81-83. Smith, T.I.J., Jenkins, W.E. and Minton, R.V., 1990. Production of advanced fingerling and subadult striped bass and striped bass hybrids in earthen ponds. Pages 121-140 m R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD.

167

Spotte, S., 1979. Seawater aquariums: the captive environment. John Wiley and Sons, New York. Stickney, R.R., 1994. Principles of aquaculture, John Wiley and Sons, New York. Timmons, M.B., 1994. Use of foam fractionators in aquaculture. Pages 247-280 in M.B. Timmons and T.M. Losordo, editors. Aquaculture water reuse systems: engineering design and management. Developments in aquaculture and fisheries science, volume 27, Elsevier Science, Amsterdam. Timmons, M.B. and T.M. Losordo, editors., 1994. Aquaculture water reuse systems: engineering design and management. Developments in aquaculture and fisheries science, volume 27, Elsevier Science, Amsterdam. Tucker, C.S. and Robinson, E.H., 1990. Channel catfish farming handbook. Van Nostrand Reinhold, New York. Varadi, L., 1985. Mechanized feeding in aquaculture. Pages 446-460 in T.V.R. Pillay, editor. United Nations Development Program. Inland Aquaculture Engineering, Food and Agriculture Organization of the United Nations, ADCP/REP/84/21, Rome. Visscher, L. and Dwyer, W.P., 1990. Oxygen supplementation: a new technology in fish culture, volume 2. Information Bulletin Number 2, U.S. Department of the Interior, Fish and Wildlife Service, Region 6, Denver, CO. Warren, H.J., Harrell, R.M., Geiger, J.G., and Rees, R.A., 1990. Design of rearing facilities for striped bass and hybrid striped bass. Pages 17-28 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Watten, B.J., 1991. Application of pure oxygen in raceway culture systems. Pages 311-332 in Northeast Regional Agricultural Engineering Service. Engineering aspects of intensive aquaculture, NRAES-49, Proceedings from the Aquaculture Symposium, Cornell University, Ithaca, NY. Watten, B., 1994. Aeration and oxygenation. Pages 173-208 in M.B. Timmons and T.M. Losordo, editors. Aquaculture water reuse systems: engineering design and management. Developments in aquaculture and fisheries science, volume 27, Elsevier Science, Amsterdam. Watten, B., Colt, J. and Boyd, C., 1989. Impact of dissolved nitrogen and carbon dioxide on the operation of pure oxygen systems. Fisheries Bioengineering Symposium, American Fisheries Society, Bethesda, MD. Watten, B.J. and Johnson, R.P., 1990. Comparative hydraulics and rearing trial performance of a production scale cross-flow rearing unit. Aquacultural Engineering, 9: 245-267. Westers, H., 1991. Modes of operation and design relative to carrying capacities of flow-through systems. Pages 151159 in Northeast Regional Agricultural Engineering Service. Engineering aspects of intensive aquaculture, NRAES-49, Proceedings from the Aquaculture Symposium, Cornell University, Ithaca, NY. Wheaton, F.W., 1977. Aquacultural engineering. John Wiley and Sons, Inc., New York. Wheaton, F.W., Hochheimer, J.N., Kaiser, G.E., Krones, M.J., Libey, G.S. and Easter, C., 1994a. Nitrification filter principles. Pages 101-126/n M.B. Timmons and T.M. Losord0, editors. Aquaculture water reuse systems: engineering design and management. Developments in aquaculture and fisheries science, volume 27, Elsevier Science, Amsterdam.

168

Wheaton, F.W., Hochheimer, J.N., Kaiser, G.E., Malone, R.F., Krones, M.J., Libey, G.S. and Easter, C.C., 1994b. Nitrification filter design methods. Pages 127-172 in M.B. Timmons and T.M. Losordo, editors. Aquaculture water reuse systems: engineering design and management. Developments in aquaculture and fisheries science, volume 27, Elsevier Science, Amsterdam. Yeager, D.M., Van Tassel, J.E. and Wooley, C.M., 1990. Collection, transportation, and handling of striped bass broodstock. Pages 29-42 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Youngs, W.D. and Timmons, M.B., 1991. A historical perspective or raceway design. Pages 160-169 in Northeast Regional Agricultural Engineering Service. Engineering aspects of intensive aquaculture, NRAES-49. Proceedings from the Aquaculture Symposium, Comell University, Ithaca, NY. Zeigler, T.R., Woods, L.C. and Gabaudan, J., 1984. Striped bass feeds and feeding. Pages 151-176 in J.P. McCraren, editor. The aquaculture of striped bass: a proceedings. Maryland Sea Grant Publication UM-SG-MAP-84-01. University of Maryland, College Park.

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

169

Chapter 6 White Bass Production and Broodstock Development Christopher C. Kohler 6.1 INTRODUCTION White bass are the primary congeneric fish used to cross with striped bass to produce hybrid striped bass. However, due to its lesser stature as a sportfish, considerably less research has been focused on aspects of its biology, ecology, and husbandry. Nevertheless, white bass do usually contribute one-half of the gene pool of hybrid striped bass (palmetto and sunshine bass), and research directed at this species has as much application and potential benefit for the hybrid striped bass industry as does research conducted on striped bass. This chapter summarizes what is known about white bass culture. Much of the knowledge regarding white bass emanates from experiences with sunshine bass. At least for the hatchery and larval rearing phases, white bass and sunshine bass appear to be reasonable surrogates for one another.

6.2 BIOLOGY AND ECOLOGY 6.2.1 Geographic Range White bass are native to North America with a natural range extending from the St. Lawrence River west through the Great Lakes (excluding Lake Superior) to South Dakota, and the Mississippi and Ohio River drainages south to the Gulf of Mexico (Becker, 1983; Lee et al., 1980; Scott and Crossman, 1973). The white bass range has been greatly expanded by stocking and now includes all the Gulf and south Atlantic states, as well as New Mexico, Utah, Colorado (Jenkins, 1970), California (yon Geldern, 1966), and Nevada (Trelease, 1970). 6.2.2 Morphological Characteristics White bass (Figure 6.1) resemble other members of the genus Moronewith horizontal black stripes laterally, and an overall silver color with upper sides olive gray shading to white on head and belly (Becker, 1983). The body is robust, deep, and strongly compressed laterally with the following meristic characters (from Bayless, 1968): 9 9 9 9 9 9

Lateral line Scales above lateral lineSoft anal rays Soft dorsal rays Teeth on tongue Parr marks -

52 to 58 7 to 9 (usually 8) 12 to 13 12 to 13 1 patch absent.

Fingerling white bass are easy to distinguish from fingerling striped bass which have parr marks (Bayless, 1968) and typically have two tooth patches (Williams, 1976). Conversely, it is sometimes difficult to distinguish the young of white bass and hybrids of striped bass, including FI, F:, and backcrosses (Kerby and Harrell, 1990).

170

Fig. 6.1. Adult female white bass in spawning condition. (Photo by V. Sanchez) 6.2.3 Age and Growth In the wild, white bass are relatively fast-growing fish that can live up to nine years of age, but usually only five to six years (Table 6.1). Young-of-the-year white bass grew about 0.1 cm/day in Lewis and Clark Lake, South Dakota (Ruelle, 1971), but likely grow considerably faster in their southern range. In general, white bass grow to a maximum of about 42.5 cm total length (TL), with weights rarely exceeding 2 kg. 6.2.4 Food Habits Newly hatched white bass initially feed on rotifers and similar size organisms for two to three weeks when they reach a size that allows them to feed on microcrustaceans. Copepods and cladocerans comprise the bulk of the natural diet, at least through midsummer (Ruelle, 1971; Priegel, 1970). However, insects were found to predominate in young white bass during spring in Beaver Reservoir, Arkansas (Olmsted and Kilambi, 1971). By midsummer, young white bass will switch to a largely piscine diet, provided forage fish are available in suitable size and abundance. Alternatively, the fish may continue to consume invertebrates (Voightlander and Wissing, 1984). Adults are primarily piscivorous. 6.2.5 Natural Spawning Female white bass in the wild usually mature at age 3 while males generally mature a year earlier (Ruelle, 1971). The spawning temperature for white bass is typically between 14.4 ~ to 18.3 ~ C (Ruelle, 1971). White bass migrate up tributaries when available and spawn in shallow waters on firm gravel or sand (Becker, 1983). They will spawn on any suitable shoreline structure in the absence of tributaries. Spawning occurs during both day and night (Chadwick et al., 1966), but spawning fish are most active crespuscularly (Voightlander and Wissing, 1984). Eggs are adhesive, demersal, and increase little in diameter when waterhardened (Rees and Harrell, 1990). No nest construction or care is provided to the eggs which stick to gravel

Table 6.1. Calculated total length (cm) of white bass from various waters (from Davin et al., 1989).

Age Body of Water

1

2

3

4

5

6

7

8

9

Lake Erie

11.9

20.8

27.7

31.5

33.5

34.5

35.6

Lewis & Clark, SD

10.9

24.4

30.2

32.8

35.8

36.8

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

Ruelle (1971)

McConaughy, NE

11.7

25.1

32.0

35.8

39.4

41.4

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

McCarraher et al. (1971)

Oklahoma a

19.1

31.0

36.6

40.9

43.4

45.2

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

Jenkins and Elkin (1957)

Oneida, NY

13.5

26.2

31.4

33.8

35.6

37.3

39.1

........

40.6

References

Van Oosten (1941)

44.2

Forney and Taylor (1963)

a

Unweighted means.

"--.I

172

and vegetation. Mature white bass females can each produce several hundred thousand eggs ranging in diameter between 0.61 and 0.68 mm at ovulation (Bayless, 1972). Hatching occurs in about two days, with fry being approximately 3.0 mm TL at hatch, growing in 15 days (at 17.8~ to 8.0 mm TL (Ruelle, 1971). 6.3 COLLECTION AND TRANSPORTATION OF BROODSTOCK Adult white bass can readily be caught by hook-and-line (Kohler et al., 1994a) or trap nets (Yeager et al., 1990). Electrofishing can be used for capturing white bass, but with varying degrees of success. Ideally, hauling tanks should be filled with water from where the broodfish are collected. Salt (NaCI or synthetic sea salts) should be added to the water to raise the salinity to 5 ppt, and depending on water temperature where fish are collected, hauling tank water temperature should be 18.3 ~ or less. Ice can be added if needed. The hauling tank should be equipped with pure oxygen regulated to maintain dissolved oxygen concentrations at a minimum of 7 ppm. Depending on fish condition and hauling duration, approved therapeutic treatments to control disease should be considered (see Chapter 7 ). The use of the anesthetic MS222 during hauling does not appear to be necessary, but can be used at appropriate doses based on experience of haulers. Care should always be exercised when using MS-222 due to possible shifts in water pH. 6.4 BROODSTOCK ACCLIMATIZATION TO CAPTIVITY 6.4.1 Rearing Facilities At the hatchery, white bass broodstock should be placed in a stress-recovery tank containing 5-7.5 ppt NaCI or synthetic sea salts for several hours. Salinity can be slowly reduced to 3-5 ppt by flushing. White bass seem to perform best in circular tanks, but can be held in rectangular tanks if necessary. Excess broodstock can be maintained in earthen ponds. South Carolina researchers have demonstrated that wild white bass can be captured in the fall, matured in outdoor tank culture systems using ambient conditions, and spawned in the spring (Smith and Jenkins, 1988) or summer (Smith et al., 1996). 6.4.2 Water Quality Requirements White bass do not have any peculiar water quality requirements. In general, water quality acceptable for other coolwater fishes will suffice (Stickney and Kohler, 1990; Mullis and Smith, 1990; Piper et al., 1982). Kohler et al. (1994a) recommend maintaining brood white bass at 2 ppt salinity. 6.4.3 Disease Control Adult white bass captured from the wild or from culture ponds are often infested with ciliated protozoans and/or monogenetic trematodes. Fungal infections or injuries sustained during collection and hauling are also common. Therapeutic treatments of salt and formalin-F at standard concentrations (Stickney and Kohler, 1990) are usually sufficient to treat these diseases (Kohler et al., 1994a). It is often necessary to initially subject the fish to a concentrated bath solution followed by a prolonged treatment for 7 to 10 days. 6.4.4 Training To Formulated Feed Kohler et al. (1994a) demonstrated that adult white bass can be trained to accept and maintained on dry formulated feed for over two years. They stressed newly collected white bass should not be fed until all therapeutic treatments are completed. This delay assures the fish will be hungry when feed is first offered, and

173

prevents habituating fish to ignore feed by presenting it to them when stress or disease agents impede their appetites. The fish are initially trained to formulated feed by hand-feeding moist pellets (Figure 6.2), prepared by mixing equivalent amounts of commercial dry trout feed (broodstock diet; 40:11 protein:fat) with raw gizzard shad Dorosoma cepedianum, and a vitamin premix (coolwater fish; U.S. Biochemical, Cleveland, Ohio, USA). Healthy white bass readily take this feed. By the second day of active feeding, a small proportion of the dry trout feed (floating) is interspersed with the moist pellets. The proportion of dry feed is slowly increased in the diet until fish accept 100% dry feed, a process usually complete in two weeks or less. 6.4.5 Separating Sexes It is sometimes advantageous to separate broodfish by gender. Sexes can readily be distinguished when the fish are in spawning condition by simply exerting abdominal pressure to extrude a small amount of gametes. However, the gender of white bass not in spawning condition is not as easily distinguished. Sex determination of these fish can be achieved with about 90% accuracy by external examination of the genital regions (Kohler et al., 1994a). Female white bass usually have a highly convoluted genital region as opposed to a smooth appearance in most males (Figure 6.3). 6.5 CONTROLLED SPAWNING 6.5.1 Use of Hormones White bass females are normally injected with human chorionic gonadotropin (hCG) to induce ovulation. Males may also be injected with hCG to increase semen volume. The dosages traditionally injected are considerably higher than those used for striped bass (Smith, 1990). Bonn et al. (1976) stated that white bass females can be successfully induced to spawn at dosages between 500-1000 IU/pound (1100-2200/kg). This was not a recommendation, but has been taken to be as such. Recently, Kohler et al. (1994b) showed that

Fig. 6.2. Training adult white bass to formulated feed requires feeding by hand and close observation. (Photo by D. Russell).

174

Fig. 6.3. Urogenital regions and gonads of male (16) and female (13) white bass not in spawning condition, showing greater convolutions between the anus and urogenital opening on the female (arrows). (Photo by D. Russell; reproduced from Kohler et al., 1994a with permission).

175

hCG dosages as low as 50 IU/kg are more efficacious. Accordingly, it appears white bass should be injected at dosages similar to those commonly used for striped bass, i.e., 275-330 IU/kg for females and 110-165 IU/kg for males. Only females in spawning condition (based on plumpness and/or visual examination of oocytes collected by a 1.5 mm outside diameter plastic catheter; Smith and Jenkins, 1988) should be injected with hCG. Reinjection of females with hCG usually results in abortion of eggs (Rees and Harrell, 1990). However, multiple injections using lower hCG dosages have not been well examined. Males have been shown to continuously spermiate for several months when held at spawning temperatures of about 16~ (Kohler et al., 1994a). A monthly dosage of 100 IU/kg is recommended if males are to be reused as broodstock over an extended period of time. Before injection with hCG, the fish should be anesthetized with 50-100 mg/L MS-222 with an equivalent amount of sodium bicarbonate to serve as a buffer. The fish can then be weighed, and the proper dosages administered intramuscularly just ventral to the first dorsal fin above the lateral line (Figure 6.4). 6.5.2 Manual Spawning Female white bass should be checked for ovulation every 2 hours starting 16 hour post-hCG injection by lightly exerting abdominal pressure (Figure 6.5) to extrude a small amount of eggs. Oocytes can be staged using similar procedures described for striped bass (Kerby, 1986; Rees and Harrell, 1990; see Chapter 2). Ovulation generally occurs between 24-36 hours post-injection at 16~176 but, as with striped bass, a window of only 1-2 hours is available to obtain properly ripened eggs. In general, ovulation is indicated by the occurrence of clear, free-flowing, uniform-shaped, yellowish-tinged eggs with fully intact inner chorion surfaces (Figure 6.6). Ovulated females should be anesthetized with 50-1 O0 mg/L MS-222 buffered with

Fig. 6.4. Injection of female white bass with human chorionic gonadotropin. (Photo by C. Habicht).

176

Fig. 6.5. Manually removing white bass eggs into a Teflon| pan. (Photo by C. Habicht).

Fig. 6.6. Properly ovulated white bass eggs shown at 100x. (Photo by V. Sanchez)

177

an equal amount of sodium bicarbonate. Ripe males usually do not need to be anesthetized because of the relative ease with which their gametes can be expressed. Before manually removing the gametes, fish should be dried with a paper towel to avoid water contamination which might prematurely activate sperm. Eggs are removed from females by firmly exerting abdominal pressure starting just above and slightly posterior to the pelvic fins, and progressing posteriorly and vertically toward the genital opening. Semen can be expressed in a similar fashion. To avoid any contamination with urine, semen can be collected by inserting a Pasteur pipette in the urogenital opening and applying suction (Kohler et al., 1994a; Figure 6.7). It is advisable to collect semen from two or more males for fertilizing each egg batch to improve genetic heterogeneity. This also decreases the possibility of fertilizing eggs with low-quality semen (e.g., sperm of low motility). The "wet" method for fertilization in which semen is added to a mixture of eggs and water is often used when males are manually stripped because urine oftentimes prematurely activates spermatozoa. Some hatcheries use a modified "wet" fertilization method in which semen and water are added simultaneously to the eggs (Kerby and Harrell, 1990). The "dry" method in which semen is mixed with eggs followed by water should probably be limited to when semen is collected by pipette. Semen collection by pipette minimizes any possibility of urine or water contamination. Regardless of method employed, at least one minute of gentle stirring in a Teflon | pan (eggs are adhesive) should be allowed for fertilization to take place. White bass oocyte development is often mixed with up to 40% being in various stages of development when the majority of eggs have been artificially induced to ovulate. Kerby and Harrell (1990) suggest that this condition probably occurs because white bass are naturally intermittent spawners and may take several days to complete spawning.

Fig. 6.7. Collection of white bass semen using a Pasteur pipette. (Photo by C. Habicht)

178

6.5.3 Incubation Techniques White bass embryos can be incubated in aquaria (Bonn et al., 1976), Heath trays (Figure 6.8; Kohler et al., 1994a), or in MacDonald-type jars (Figure 6.9). However, they cannot be incubated in jars unless their adhesiveness is neutralized. Fuller's earth, silt, clay, starch, sodium sulfite, and tannic acid have all been used with varying degrees of success (Kerby and Harrell, 1990). Rottmann et al. (1988) provide a detailed protocol that is summarized in Table 6.2. Key elements of the protocol are use of sodium chloride and urea to clear the embryos so that developmental events can be seen, and tannic acid to reduce adhesiveness. Formalin-F can be used at 50 ppm to reduce fungal infections if hatchery water is being recycled. Otherwise, concentrated bath treatments of 286-429 ppm can be used by injecting 2 or 3 mL of full strength formalin into the top of the shad tubes to provide a rapid flush treatment (Kerby and Harrell, 1990). Dead embryos usually turn opaque after 30 minutes and when practical should be removed. Development is temperature dependent. At 16 ~ 18~ the embryos will begin to hatch 36 to 48 hours post-fertilization and will usually be complete within 24 hours (Kohler et al., 1994a). Depending on temperature, an additional 72 to 96 hours is required for the larvae to absorb their yolksacs. Yellayi and Kilambi (1970) and Bayless (1972) provide characteristics of various stages of development for white bass.

Fig. 6.8. Heath tray used for incubating white bass embryos (see Kohler et al., 1994a for details). (Photo by J. Rudacille)

179

Fig. 6.9. Use of MacDonald-type jars for incubating white bass ..~,~oos. (Photo by C. Kohler)

Table 6.2. Clearing and removal of adhesiveness of newly fertilized white bass eggs (modified from Kerby and Harrell, 1990).

Step One:

Make following solutions just prior to fertilizing eoos'=~. Clearin~ Eoos _

20 g 15 g 5L

~--~,

NaCI Urea Hatchery Water

Removing Adhes.iveness 0.75 g Tannic Acid 5 L Hatchery Water Step Two:

Add fertilized eggs to salt-urea mixture and aerate vigorously 7-10 minutes with airstone.

Step Three:

Remove airstone, let the eoos settle, and decant the water.

Step Four:

Add tannic acid solution to fertilized eggs and vigorously aerate for 6-7 minutes.

Step Five:

Incubate fertilized ..~,~oosin MacDonald-type jars. (Note: Steps 1-3 can be omitted once experience has been attained.)

180

6.5.4 Out-of-Season Spawning Kohler et al. (1994a) successfully spawned captive white bass acclimated to indoor tanks for over one year and entrained to simulated temperature and light cycles. They maintained one water recycle system under an ambient photoperiod and temperature regime, one on a 9-month (compressed) annual cycle of changing photoperiod and temperature, and one held at a temperature at or above spawning temperature (20 ~ -1-5 o C) and constant photoperiod (14 h L: 10 h D). Using hCG injections (1100 and 275 IU/kg for female and male fish), they induced fish to spawn at the prescribed times. Constant-cycle fish injected with hCG failed to spawn early, but did spawn at the same time as the ambient-cycle fish following additional hormone stimulation. Annual rhythms of serum levels of estradiol- 1713and testosterone, as well as gonadal growth and histology of wild and the three captive populations of white bass, were documented and correlated with actual spawning events. The authors demonstrated that wild adult white bass can readily be habituated to captivity to serve as broodstock and that the fish are amenable to manipulation of spawning time. 6.6 G R O W O U T Larval white bass can be reared in ponds in the same fashion as sunshine bass (see Chapter 3). Intensive culture of larval white bass is currently being attempted at several research stations, but to date has met with only limited success. White bass in tanks require at least two weeks of feeding on rotifers or some other suitable-size food before they are large enough to accept brine shrimp (Artemia). Surface water spray (Barrows et al., 1993) should be used during intensive larval rearing to increase swim bladder inflation rates. The procedure entails spraying water on the surface of the tank water in the direction of flow by drilling 1-mmdiameter holes in a supply pipe placed above the water surface. Considerably more research is needed in intensive larval rearing, particularly if culturists want to avail themselves of established out-of-season spawning techniques. Procedures for production of fingerling white bass to adult (broodstock) size should closely parallel those for the hybrids (see Chapter 3). However, no studies have been published on the nutritional requirements of white bass and, assuming equivalence with hybrid striped bass, may prove to be a fallacy. 6.7 BROODSTOCK DEVELOPMENT Forty-seven potential hybrid striped bass fry/fingerling producers were identified from the 1994 Aquaculture Magazine Buyer's Guide and from the directory of the Striped Bass Growers Association. A mail survey on broodstock use was sent to each in fall 1994, and 23 (49%) were returned. Of the 23 respondents, 15 indicated that they were involved in hybrid striped bass fry/fingerling production. Their average time in business is 6.7 years with a range of 1 to 18 years. Sunshine bass are produced by 67% of the responding producers, while 53% produce the palmetto bass (some produce both crosses). Broodstock of white and striped bass are maintained by 40% of the responding producers. However, only 27% maintain broodstock in which they have conducted some genetic selection. Broodstock tend to be collected from local populations where available (Table 6.3), but several respondents indicated that white bass broodstock are also obtained from Sandusky Bay, Lake Erie (Ohio). These fish are presumably obtained via the commercial fishery there. Seventy-three percent of the responding producers indicated that having domesticated broodstock would be beneficial to their businesses. The remaining 27% were uncertain. Although some genetic selection is apparently being undertaken, the questionnaire was not designed to allow determination of the actual status of this work. Recently, Ehtisham et al. (1994) reported significant differences in growth rates of young striped bass originating from various geographical regions, but grown at

181

a single station under identical conditions. Differential growth rates among different geographical populations of white bass might also occur. There is clearly a need for systematic broodstock selection programs for all Morone species being commercially cultured.

Table 6.3. Water bodies and states where white bass broodstock are obtained for hybrid striped bass producers (based on responses to fall 1994 survey conducted by author). WATER BODY

STATE(s)

Walter George Reservoir

Alabama-Georgia

Lake Erie (Sandusky Bay)

Ohio.

Arkansas River

Arkansas

Tennessee River

Tennessee-Alabama

Wateree River

South Carolina

Santee-Cooper Drainage

South Carolina

Coosa River

Alabama

Mississippi River

Mississippi

6.8 SUMMARY Adult white bass can be collected from the wild, acclimated to captivity, trained to eat formulated feed, and, by manipulating temperature and photoperiod, induced to spawn (both in and out of season) using hCG injections. Hatchery and pond fry production techniques are basically the same as for sunshine bass. Considerably more research is needed to perfect intensive culture of larval white bass. To date, only a few attempts are being made to develop domesticated broodstocks, and virtually no systematic genetic selection programs exist. It is clear these areas must be adequately addressed if hybrid striped bass culture is to meet its full potential.

182

References

Barrows, F.T., Zitzow, 1LE. and Kindschi, G.A., 1993. Effects of surface water spray, diet, and cost of production of intensively reared larval walleyes. The Progressive Fish-Culturist, 55 224-228. Bayless, J.D., 1968. Striped bass hatching and hybridization experiments. Proceedings of the Annual Conference Southeastern Association of Game and Fish Commissioners, 21:233-244. Bayless, J.D., 1972. Artificial propagation and hybridization of striped bass, (Morone saxatilis) (Walbaum). South Carolina Wildlife and Marine Resources Department, Columbia. Becker, G.C., 1983. Fishes of Wisconsin. University of Wisconsin Press, Madison. Bonn, E.W., Bailey, W.M., Bayless, J.D., Erickson, K.E. and Stevens, R.E., 1976. Guidelines for striped bass culture. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Chadwick, H.K., von Geldern, Jr., C.E. and Johnson, M.L., 1966. White bass. Pages 412-422/n A. Calhoun, editor. Inland fisheries management. California Department of Fish and Game, Sacramento. Davin, W.T., Heidinger, R.C., Sheehan, R.J., Kohler, C.C. and Tetzlaff, B.L., 1989. Supplemental stocking and spillway escapement literature review. Project F-77-R Completion Report. Illinois Department of Conservation. Ehtisham, A., Brown, J.J. and Conover, D., 1994. Evidence for countergradient variation in growth rate of larval and juvenile striped bass, or why striped bass growers migJat want to select broodstock fish fi'om New York and not South Carolina. Aquaculture '94, World Aquaculture Society, New Orleans, LA, Abstract. Fomey, J.L. and Taylor, C.P., 1963. Age and growth of white bass in Oneida Lake, New York. New York Fish and Game Journal, 10:194-200. Jenkins, R.M. and Elkin, R.E., 1957. Growth of white bass in Oklahoma. Oklahoma Fisheries Research Laboratory Report Number 60, Oklahoma City. Jenkins, R.M., 1970. Reservoir fish management. Pages 173-182/n N.C. Benson, editor. A century of fisheries in North America. American Fisheries Society Special Publication Number 7, Washington, DC. Kerby, J.H., 1986. Striped Bass and striped bass hybrids. Pages 127-147 in R.R. Stickney, editor. Culture of nonsalmonid freshwater fishes. CRC Press, Boca Raton, FL. Kerby, J.H. and Harrell, R.M., 1990. Hybridization, genetic manipulation, and gene pool conservation of striped bass. Pages 159-190/n R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Kohler, C.C., Sheehan, R.J., Habicht, C., Madison, J.A. and Kayes, T.B., 1994a. Habituation to captivity and controlled spawning of white bass. Transactions of the American Fisheries Society, 123: 964-974. Kohler, C.C., Sheehan, R.J., Sanchez, V. and Suresh, A., 1994b. Evaluation of various doses ofhCG to induce final oocyte maturation and ovulation in white bass. Aquaculture '94, World Aquaculture Society, New Orleans, LA. Abstract. Lee, D.S., Gilbert, C.S., Hocutt, C.H., Jenkins, R.E., McAllister, D.E. and Stauffer, J.R., 1980. Atlas of North American freshwater fishes. North Carolina Biological Survey Publication Number 1980-12, Raleigh.

183

McCarraher, D.B., Madsen, M.L. and Thomas, R.E., 1971. Ecology and fishery management of McConaughy Reservoir, Nebraska. American Fisheries Society Special Publication 8:299-311. Mullis, A.W. and Smith, J.M., 1990. Design considerations for striped bass and striped bass hybrid hatching facilities. Pages 7-27 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD Olmsted, L.L. and Kilambi, R.V., 1971. Interrelationships between environmental factors and feeding biology of white bass of Beaver Reservoir, Arkansas. American Fisheries Society Special Publication 8: 397-409. Piper, R.G., McElwain, I.B., Orme, L.E., McCraren, J.P., Fowler, L.G. and Leonard, J.R., 1982. Fish hatchery management. United States Department of the Interior, Fish and Wildlife Service, Washington, DC. Priegel, R.R., 1970. Food of the white bass, Roccus chrysops, in Lake Winnebago, Wisconsin. Transactions of the American Fisheries Society, 99: 440-443. Rees, R.A. and Harrell, R.M., 1990. Artificial spawning and fry production of striped bass and hybrids. Pages 43-72 /n R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Rottmann, R.W., Shireman, J.V., Starling, C.C. and Revels, W.H., 1988. Eliminating adhesiveness of white bass eggs for the hatchery production of hybrid striped bass. The Progressive Fish-Culturist, 50: 55-57. Ruelle, R., 1971. Factors influencing growth of white bass in Lewis and Clark Lake. Transactions of the American Fisheries Society Special Publications, 8:411-423. Scott, W.B. and Crossman, E.J., 1973. Freshwater fishes of Canada. Fisheries Research Board of Canada Bulletin 184, Ottawa. Smith, T.I.J., 1990. Aquaculture of striped bass, Morone saxatilis, and its hybrids in North America. Pages 53-61 in R.S. Svrjcek, editor. Genetics in aquaculture: proceedings of the sixteenth U.S.-Japan meeting on aquaculture. NOAA Technical Report NMFS 92, Washington, D.C.. Smith, T.I.J. and Jenkins, W.E., 1988. Culture and controlled spawning of striped bass (Morone saxatilis) to produce striped bass and striped bass x white bass (Morone chrysops) hybrids. Proceedings of the Annual Conference Southeastern Association ofFish and Wildlife Agencies, 40:152-162. Smith, T.I.J., Jenkins, W.E. and Heyward, L.D., 1996. Production and extended spawning of cultured white bass broodstock. The Progressive Fish-Culturist, 58:85-91. Stickney, R.R., and Kohler, C.C., 1990. Maintaining fishes for research and teaching. Pages 633-663 in C. Schreck, and P. Moyle, editors. Methods for fish biology. American Fisheries Society, Bethesda, MD. Trelease, T.R., 1970. Fish planting and transfer statistics. Nevada Outdoors and Wildlife Review, 4: 5-10. Van Oosten, J., 1941. The age and growth of the Lake Erie white bass, Lepibema chrysops (Rafinesque). Michigan Academy of Science, Arts, and Letters, 27: 307-334. Voigtlander, C.W. and Wissing, T.E., 1984. Food habits of young and yearling white bass, Morone chrysops (Rafinesque), in Lake Mendota, Wisconsin. Transactions of the American Fisheries Society, 103:25-31.

184

von Geldem, C., 1966. The introduction of white bass (Roccus chrysops) into California. California Fish and Game, 52: 303. Williams, H.M., 1976. Characteristics for distinguishing white bass, striped bass, and their hybrid (striped bass x white bass). Proceedings of Annual Conference of the Southeastern Fish and Wildlife Agencies, 29:168-172. Yeager, D.M., Van Tassel, J.E. and Wooley, C.M., 1990. Collection, transportation, and handling of striped bass brood stock. Pages 29-42 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Yellayi, R.R. and Kilambi, R.V., 1970. Observations on early development of white bass, Roccus chrysops (Rafinesque). Proceedings Annual Conference of the Southeastern Association of Game and Fish Commissioners, 23:261-265.

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 1997 Elsevier Science B.V.

185

Chapter 7 Transportation and Stress Mitigation C. R. Weirich 7.1 INTRODUCTION A prerequisite for successful production of most aquaculture species is the existence of efficient, economical procedures to facilitate the transfer of culture animals from one point to another with minimal losses. This axiom is especially true in striped bass and hybrid striped bass culture because, as described in Chapter 3, current pond production practices may involve fish being moved at least three times before harvest: 1) newly hatched larvae are transferred to fingerling ponds to produce phase I fish, 2) phase I fish are moved to initial growout ponds to produce phase II fish, and 3) phase II fish are transferred to final ~owout ponds to produce harvestable phase III fish. In addition to on-site transfers, fish of all sizes and life stages, from eggs to broodfish, may require transportation to other facilities or markets depending on the individual culture operation or marketing strategy. Although a necessary component of striped bass and hybrid striped bass culture, handling and transport procedures can potentially expose fish to a number of stressors (physical or chemical agents perceived as harmful, discussed below) which can cause or contribute to the development of significant mortalities if proper measures are not taken to alleviate the effects of stress. This chapter provides the reader with the current knowledge of handling and transport stress on striped bass and its hybrids, and discusses procedures to mitigate stress. 7.2 STRESS AND STRESS MITIGATION 7.2.1 Effects Of Handling and Transport-induced Stress On Fish Stressors associated with handling and transport procedures are generally due to either adverse physical or environmental factors. Examples of physical stressors include confinement of large numbers of fish in a small area and abrading or injuring fish during net transfer. Environmental stressors include less than optimal water quality conditions such as low dissolved oxygen levels, high ammonia levels and temperature extremes. Stressors can range in intensity from mild (chronic) to severe (acute). If fish are exposed to a severe stressor, for example extremely low levels of dissolved oxygen, mortality usually occurs immediately and will be directly attributable to that particular stressor. In most cases, however, mortalities associated with transport procedures are due to several mild stressors, such as handling and marginal water quality conditions which act in concert to elicit a "stress response" by the fish. The stress response, also described in Chapter 10, is a complex set of physiological changes that occur in response to the detection of a stressor or stressors by fish (Picketing, 1981; Wedemeyer et al., 1990; Barton and Iwama, 1991). These physiological changes, such as increased cardiac output, increased gill vascularity, and increased glycogenolysis (Pickering, 1981, 1993; Wedemeyer et al., 1990; Barton and Iwama, 1991) are affected through the release of catecholamines and corticosteroid hormones (Mazeaud et al., 1977), and are directed toward enabling fish to escape from or better cope with the stressor(s).

186

Because of relative ease of measurement, corticosteroid levels are often used as a primary indicator of the stress response in fish. Increased corticosteroid levels have been detected in a number of fish species exposed to stressors (Mazeaud et al., 1977; Strange et al., 1977, 1978; Specker and Schreck, 1980; Barton and Peter, 1982; Carmichael et al., 1984; Davis and Parker, 1986; Robertson et al., 1988; Mazik et al., 1991a) including striped bass (Davis and Parker, 1990; Mazik et al., 1991b; Harrell, 1992; Harrell and Moline, 1992; Young and Cech, 1993) and sunshine bass (Tomasso et al., 1980) exposed to handling and transport-induced stress. Although physiological changes brought about by release of catecholamines and corticosteroid hormones during the stress response are immediately beneficial, in most aquaculture systems, especially transport units, fish cannot escape from, and may be exposed to, the stressor(s) for an extended period of time. Depending on the severity of the stressor(s) and exposure time, mortality usually results from osmoregulatory dysfunction and immunosuppression: detrimental processes that are attributable to hormones released during the stress response. Increased metabolic activity induced by stress hormones eventually depletes energy reserves and, when coupled with the direct hormonal effects on osmoregulatory processes (e.g., epinephrine increases water permeability of the gills), leads to the inability of fish to effectively osmoregulate (Lewis, 1971; Wydoski and Wedemeyer, 1976; Eddy and Bath, 1979; Carmichael et al., 1983, 1984; Davis and Parker, 1983). Osmoregulatory dysfunction has also been observed in striped bass (Mazik et al., 1991b) and sunshine bass (Tomasso et al., 1980; Weirich et al., 1992) subjected to handling and transport. While the exact mechanism of immunosuppression due to activation of the stress response is still unknown (Wedemeyer, 1970; Barton and Iwama, 1991; Pickering, 1993), corticosteroid hormones are thought to be involved (Pickering, 1993). Immunosuppression greatly increases the fish's susceptibility to a wide variety of disease agents; even those normally considered to be relatively non-virulent. Most warmwater fish culturists are familiar with the almost routine outbreak of disease (often due to a facultative bacterial infection), which occurs after fish are exposed to handling and transport-induced stressors in warmwater temperatures. The mortalities resulting from the increased susceptibility of fish to facultative pathogens are then, although occurring some time after the transportation, directly caused by stress incurred during the transport event. 7.2.2 Mitigation Of Handling and Transport-induced Stress Before considering methods to alleviate stress associated with handling and transport procedures, the culturist should realize that the presence of stressors is an inherent feature of the intensive production of fish. In other words, fish usually will exhibit a stress response to some degree during most normal culture practices, especially handling and transport. Therefore, the goat of culturists transporting fish should be to attempt to reduce or mitigate the effect of stressors, wherever possible. Mitigation of handling and transportinduced stress can be achieved by; 1) reducing the severity of stressors, 2) minimizing the duration of stressors, 3) reducing the number of stressors, 4) minimizing plasma ion disturbances, and 5) minimizing increases in metabolic rate. In the remainder of this chapter general descriptions of how culturists can adhere to these guidelines will be discussed. These include use of proper transport equipment, preconditioning of fish before transport, use of proper fish loading rates, maintenance of water quality characteristics, use of water additives, and post-conditioning of fish after transport. Practical guidelines currently in use by striped bass and hybrid culturists will also be presented.

9

187

7.3 TRANSPORT EQUIPMENT Equipment required to successfully transport striped bass and hybrid striped bass (exclusive of harvesting equipment) includes fish transfer devices, such as buckets and dip nets, transport units (hauling tanks), oxygenation and aeration systems, and meters or testing kits to monitor water quality characteristics including dissolved oxygen, temperature, pH, and ammonia. Lack of, or improper use of, any of these items can stress fish either directly (e.g., faulty or inadequate oxygenation system) or indirectly (e.g., lack of oxygen meter kit to detect low oxygen levels). For additional information concerning transport equipment and usage the reader is referred to the following sources: Johnson (1979), Piper et al. (1982), Busch (1985), and Carmichael and Tomasso (1988). 7.3.1 Fish Transfer Equipment Striped bass and hybrid striped bass may be moved from culture units to transport tanks and vice versa using both "wet" and "dry" transfer methods. As the term implies, wet transfer methods utilize water as the medium to move fish with little or no contact with nets. Because of the reduced extraneous contact associated with wet transfer there is less handling stress incurred and less potential for skin injury associated with this method compared to dry transfer, which is normally only used when phase III food fish are harvested and immediately ice-packed or slaughtered. The most common wet transfer method consists of quickly and gently netting fish and placing them into a suitable container (such as a bucket or tub) filled with water. Fish and water are then emptied into the destined transport or culture unit. Wet transfer is widely used by producers to move fish of all sizes with the exception of market-ready food fish and brood fish. Wet transfer may also be accomplished through the use of fish loading pumps or auger-equipped loading devices (Figure 7.1). Auger-equipped loaders are widely used in trout and salmon aquaculture and are becoming increasingly popular as a method to transfer phase II and phase III fish successfully from ponds to transport units, especially in the mid-Atlantic region. Unloading fish directly from transport to culture units through the use of chutes or pipes attached to the transport unit's drain system is also used to transfer fish without net contact. Considerations concerning the use of this technique will be discussed later. As mentioned previously, dry transfer involves exposure of fish to increased physical stress and is therefore usually only used to transfer fish immediately before they are ice-packed or slaughtered. In this method, fish are concentrated into a small area and then netted and placed directly into transport units or placed in a loading bucket or loading net that is then hoisted by a boom-equipped tractor or truck and unloaded into the transport unit. Regardless of the method used, stress and injury can be reduced during fish transfer by using the proper type of net or other loading device and through the avoidance of overcrowding. Dip nets and loading nets should be constructed of soft, uncoated or slightly coated, knotless netting material. While less resistant to everyday wear and tear than coated, knotted netting material, knotless netting is considerably less injurious to scaled fish such as striped bass and hybrid striped bass. Netting materials should be an appropriate mesh size for the size fish to be transferred to avoid "gill netting." If loading buckets or boxes are used, they should be constructed with interior surfaces, including welds, ground and smoothed to avoid abrading fish. Bottom-mounted tray doors should be constructed so they can be quickly and easily opened. Overcrowding of fish can easily occur during fish transfer. Fish are present at high densities and make it tempting to overload dip nets or loading nets, buckets or boxes. To avoid overcrowding, multiple transfers of small numbers of fish should occur instead of a few transfers of large numbers of fish.

188

Fig. 7.1. Fish loading devices for movement of juvenile and sub-adult fish: Top: Auger-equipped loading device Bottom: Fish loading pump (Photo by: Top, C. Weirich, Bottom, R. Harrell).

189

7.3.2 Transport units Fish transportation tank designs have undergone considerable improvement during the past decade. Although current transportation unit designs may vary depending on the manufacturer, most are suitable for transporting striped bass and hybrid striped bass. Transport units are usually rectangular and constructed of aluminum, steel, or fiberglass. Yeager et al. (1990) recommend that broodfish be transported using circular or oval transport tanks (Figure 7.2) to allow unrestricted swimming. Regardless of the material used, the interior of the tank should be smooth to avoid physical injury to fish. Individual tank capacities generally range from approximately 200 to 3,000 liters.' Relatively small (< 2,000 liters) removable transport units carried on medium to heavy duty (450-900 kg) trucks are commonly used to transport fish short distances or for on-farm fish movement. A recent trend has been to mount transport units on trailers (e.g., gooseneck trailers) that allows trucks to be easily used for other purposes when fish are not being transported. Long distance transport of large numbers of fish is usually achieved through the use of multiple tank transport units. These units may be mounted on large double-axle trucks (Figure 7.3) or may be placed on a heavy duty trailer (Figure 7.4). Regardless of the type of unit used, it must be properly sized to the vehicle. It is important to keep in mind that water is heavy (one liter weighs one kilogram), therefore, truck mounted units when loaded should fall within the guidelines of the truck manufacturer. The weight of loaded trailermounted units should be within the guidelines of the trailer manufacturer, and both the total weight and the tongue weight of the trailer and loaded tank should be within the guidelines of the towing vehicle. Transport units may be insulated or non-insulated. Insulated tanks are commonly used in the southern United States for long distance transport. Urethane foam used as a filler between tank walls is the most common insulation material. Non-insulated tanks are used mainly for short distance or on-farm transport. The absence of insulation makes these units lightweight and easy to handle. Fish are loaded through doors in the top of the transport unit, which are at least 0.75 m by 0.75 m to facilitate loading of fish by mechanical equipment. Large door openings also allow fish to be transferred easily via net or by water-filled containers. Drainage ports may be square or circular and located either in the rear or side of the tank. Side outlets allow fish to be unloaded by wet transfer through the use of chutes or extended discharge pipes (Figure 7.5). Drain openings should be large enough to unload the largest fish hauled without physical damage. Openings of 20 cm square or 20 cm in diameter are sufficient to unload phase I and phase II fish. Larger openings (e.g., 25 to 30 cm square or in diameter) may be required for phase III food fish. Square drainage ports are normally sealed with a cam-locking door, while circular drainage ports are sealed with inside rubber stoppers, outside expansion plugs, or threaded caps. An interior sliding door is desirable to prevent fish loss while drainage ports are being opened and unloading chutes or pipes are positioned. Sloped tank bottoms which direct fish toward drainage ports are a common feature on most current transport units. Other components of a well-constructed transport unit include air vents, overflow drains, and electrical outlet boxes. Air vents (scoops) permit air circulation to occur between the water surface and the top of the transport tank. They are especially useful to prevent carbon dioxide accumulation, which can occur in long distance transport units using oxygen delivery systems (see Section 7.6.2). Overflow drains are a desired feature on transport units using electric agitators. Drains maintain water levels allowing

Capacities are given as the total capacity of the tank. Tanks are usually filled one-half to two-thirds full with water to allow displacement by fish. One liter of water is displaced by each kilogram of fish.

190

Fig. 7.2. Circular transport tank used for transporting large broodstock. (Photo by J. Van Tassel, Maryland Department of Natural Resources)

Fig. 7.3. Typical large double-axle truck used for long-distance transport of juveniles and sub-adults. (Photo by C. Weirich)

191

Fig. 7.4. Typical heavy-duty "gooseneck" trailer mounted transport unit. (Photo by C. Weirich)

Fig. 7.5. Side outlet drainage port used to unload fish by wet transfer. A 20 cm diameter PVC pipe has been placed in the opened port to allow fish and water to flow into a holding tank. (Photo by C. Weirich)

192

agitators to function at the correct operating depth. Outlet boxes provide a convenient means to operate electrical equipment including agitators and pumps. Electricity (12-volt direct current) is supplied through outlet boxes via the vehicle's battery and alternator. While not part of the transport unit per se, several items should be available in the transport vehicle to facilitate operations. A 12-volt, 0.04-kw submersible pump or gasoline pump allows the addition or removal of water when "tempering" fish (see Section 7.6.3). Hanging or electronic scales should be available to weigh samples of fish when necessary. Sufficient spare parts for oxygenation and aeration equipment should always be accessible. 7.3.3 Oxygenation and Aeration Systems The presence of an effective means to maintain adequate dissolved concentrations in transport units is imperative for successful fish transport. A variety of devices are available including air blowers, circulating pumps, electric agitators, and oxygen delivery systems using bottled oxygen. Air blowers, although widely used in the channel catfish industry to aerate large transport units delivering fish to processing plants, possess several disadvantages that restrict their use in the transport of striped bass and hybrid striped bass. Blower systems which pump large quantities of warm air can raise water temperatures (Wilson, 1950) above suggested levels for hauling (see section 7.6.3). In addition, air blowers normally produce a high degree of water turbulence, which can injure phase I and phase II fish. Air blower systems are also less efficient in oxygenating water than oxygen delivery systems. This characteristic greatly limits their use, especially during long distance transport. Use of circulating pumps is fairly common in the transport of salmonids. These devices, described in detail by Piper et al. (1982), generally consist of a gasoline-powered pump that circulates water from the bottom of the transport tank through spray nozzles mounted above the water line. In most pump systems oxygen is introduced in the intake line(s) leading to the pump. Although circulation pump systems are fairly efficient at aeration and carbon dioxide removal compared to other systems, relatively high maintenance levels are required, and they are usually more expensive to operate. In addition, water temperatures may be increased during transit depending on the type of pump used and the ambient air temperature. These factors normally preclude their use in transporting striped bass and hybrid striped bass. Electric agitators, once commonly used in all phases ofwarrnwater fish transport, are today normally only used to aerate small (< 2000 L) transport units during short-term (e.g., <1 hour) on-site fish movements, or as an emergency back-up system for long distance transport (Figure 7.6). These units, in addition to providing effective aeration including carbon dioxide removed in small transport tanks, are relatively maintenance free and easily powered through the truck's electrical system. They are, however, considerably less efficient in oxygenating water than other systems, such as those employing bottled oxygen. The current predominate oxygenation system on transport units is bottled oxygen. In addition to allowing increased fish loading rates compared to those permitted by the use of other systems (Johnson, 1979), bottled oxygen delivery systems are relatively simple to install and are not prone to mechanical problems. Bottled oxygen is commercially available in two forms: compressed oxygen gas and liquid oxygen. Both forms deliver pure oxygen in the gaseous state to transport units (Busch, 1985). Generally, compressed

193

Fig. 7.6 Standard electric agitator commonly used for short-term on-site fish movement or as an emergency back-up during long-distance transport. (Photo by R. Harrell)

Fig. 7.7. Standard compressed oxygen cylinders mounted on transport unit. (Photo by C. Weirich)

194

oxygen gas is used when small volumes of oxygen are required for short-term transport of fish in small units. Most on-farm movements of striped bass and hybrid striped bass are accomplished using compressed oxygen gas. A standard compressed oxygen cylinder holds approximately 6,910 L of oxygen and weighs about 90 kg (Figure 7.7). Liquid oxygen is stored in vacuum-insulated containers (cryogenic or dewar tanks) with capacities ranging from 80-180 L of liquid oxygen (Figure 7.8). A standard 160 L container weighs about 340 kg and provides approximately 127,400 L of gaseous oxygen -- equivalent to 18 standard compressed oxygen cylinders. The large volumes of gaseous oxygen available in dewar tanks make liquid oxygen the preferred oxygen source for long distance transport. Liquid oxygen tends to cool the transport water and does not cause excessive turbulence because relatively small volumes of gas are introduced as fine bubbles. These characteristics make liquid oxygen supply systems ideal for transporting small phase I and phase II fish. Liquid oxygen is less expensive per liter than compressed oxygen gas, but dewar containers are more expensive. Liquid oxygen containers cannot be stored over an extended period of time because in order to maintain safe internal pressures, oxygen is slowly vented from dewar containers at the approximate rate of 2% daily and must be checked periodically, especially before transport. This factor usually makes the use of liquid oxygen on smaller operations less practical. Regardless of the source, oxygen is delivered to transport units through pressure regulators that supply aeration lines equipped with adjustable flow meters to various types of diffusers. Flow meters are normally set to deliver approximately 1-2 L of oxygen gas per minute for every 100 L of transport water. Diffusers introduce oxygen in the form of fine bubbles and may be constructed of fused minerals (airstones), finely woven hoses, or porous plastic tubing (Figure 7.9). The action of diffusers does not remove carbon dioxide efficiently because of the lack of agitation at the water surface. Colt and Watten (1988) present detailed information on the uses of pure oxygen in fish culture.

Fig. 7.8. Vacuum-insulated liquid oxygen container typically used in long-distance transport. (Photo by C. Weirich)

195

Fig. 7.9. Diffuser constructed of porous plastic tubing. The design showed here allows the diffuser tobe easily placed on the bottom of the transport tank. (Photo by C. Weirich). 7.3.4 Water Quality Monitoring Equipment To ensure that water quality characteristics remain within acceptable levels during transport, the transport vehicle should be equipped (minimally) with a dissolved oxygen meter and a standard water quality test kit to determine pH, temperature, and ammonia. The pH may also be determined through the use of an inexpensive meter. Because striped bass and hybrid striped bass are often transported in water containing dilute concentrations of salts (e.g., sodium chloride and calcium chloride), transporters may also be equipped with a refractometer or electronic meter to determine salinity and a means to determine calcium hardness (included in most water quality testing kits). Because dissolved oxygen is the most critical water quality factor during fish transport, the ability to monitor oxygen levels and the performance of oxygen and aeration equipment during transit is desirable. The status of oxygenationor aeration systems during transit can be determined by simply positioning flow meters, gauges, and other oxygenation and aeration equipment where they can be viewed easily while the truck is in operation, or by more elaborate means, such as the installation of remote sensing devices or alarm systems. For example, an alarm system that is activated by reduced oxygen pressures is described by Busch (1985). Aeration motor performance can be determined through an inexpensive monitor described by Millett et al. (1983). If the transport unit contains only one compartment, dissolved oxygen can be monitored during transit by simply placing the meter in the truck cab while the probe is positioned in the transport compartment. Unfortunately, most transport units utilized to transport fish long distances contain multiple compartments requiring the truck be stopped periodically to allow oxygen to be monitored manually. This

196

practice, however, does not completely ensure that fish losses due to low oxygen levels are avoided. For example, a malfunction in the oxygenation system could occur between stops resulting in rapid oxygen depletion that may not be detected. The recent development of remote, computer-assisted oxygen monitoring systems may help solve this problem. These systems allow oxygen and other selected water quality characteristics to be monitored from the truck cab at all times during the transport event. 7.4 P R E C O N D I T I O N I N G OF FISH B E F O R E T R A N S P O R T Mitigation of stress during fish transport should begin well before fish are actually loaded on the transport unit. The stress response of fish that are not properly preconditioned before transport can often be severe, resulting in significant mortalities. Preconditioning or preparing striped bass and hybrid striped bass for transport can involve several cultural procedures, such as increasing the salinity and/or calcium concentration of water in which fish are being held, withholding feed, monitoring fish with subsequent disease treatment when necessary, reducing the metabolism of fish through the use of anesthetics, and acclimating or "tempering" fish to transport water quality conditions. The extent the fish may be preconditioned varies according to the site from which fish are to be loaded. Due to the large volumes of water present in culture ponds, there are fewer measures culturists can take to reduce stress when loading fish from ponds as compared to loading fish from holding vats or large tanks. Nevertheless, most striped bass and hybrid striped bass currently are grown utilizing pond culture, and fish must be moved from these systems when necessary (see Chapter 3). Although some striped bass and hybrids are marketed live, most market-ready fish are usually simply packed on ice immediately after pond harvest. However, phase I and phase II fish are normally transported to holding vats or tanks after pond harvest where they are prepared for subsequent long distance transport. Methods to mitigate stress by preconditioning fish to be loaded from ponds and from holding vats or tanks are discussed in the following two sections. Although the use of water additives such as salts and anesthetics will be mentioned, a discussion concerning the physiological role of these substances and suggested levels in transport units will be deferred to Section 7.7. 7.4.1 Preconditioning Fish In Ponds There are limited measures culturists can take to prepare fish for transport directly from ponds due to the large volumes of water present in these systems. There are, however, several practices that have been shown to mitigate the stress response and improve survivability of pond-harvested striped bass and hybrid striped bass. Aside from utilizing proper harvesting equipment and techniques (discussed in Chapter 3) stress may be alleviated by increasing the calcium concentration (hardness) and/or salinity of pond water, withholding feed from fish before harvest, and reducing environmental differences (especially temperature) between pond and transport water. The importance of increasing the calcium concentration (total hardness) 2 of pond water before harvest was first illustrated by Grizzle et al. (1985). They used calcium chloride (CaC12) to increase the total water hardness from 20 to 45-100 mg/L as calcium carbonate (CaCO3) five days before harvest of phase I

2 The term total hardness refers to the concentration of divalent metal cations in water and is usually expressed as mg/L calcium carbonate (CaCO3). As described in section 7.4.1, calcium is the cation of physiologic importance. When only total hardness values are given, the calcium concentration may be estimated by dividing the total hardness by 2.5 if insignificant levels of magnesium are present.

197

striped bass and sunshine bass and observed a significant increase in survivability at harvest. In a similar study, Mauldin et al. (1986) compared post-harvest survivability of phase I striped bass from ponds with low (< 2mg~) levels of calcium to ponds with increased levels of calcium (14-40 mg/L calcium) through CaC12 addition. Fish survival from treated ponds was significantly greater than survival from the control ponds. The optimal level of calcium needed to mitigate stress in pond-loaded striped bass and hybrid striped bass has not been determined as calcium levels exceeding the treatment concentrations of these two studies have not been scientifically evaluated in pond situations. Higher calcium levels however, have been shown to mitigate stress, depending on fish size, in studies utilizing holding vats and in transport units (see appropriate sections below). Culturists with relatively hard water supplies (total hardness > 100 mg/L as CaCO3) usually do not add CaC12 to increase calcium levels. The addition of sodium chloride (NaC1) has been shown to alleviate stress due to handling and/or transport in striped bass and hybrid striped bass (Bonn et al., 1976; Tomasso et al., 1980; Cech et al., 1990; Mazik et al., 1991b; Harrell, 1992; Weirich et al., 1992) and is discussed in detail in Section 7.7.1. Application of salt to increase pond salinity to levels described in these studies in an attempt to mitigate stress associated with loading fish from ponds has not been formally evaluated, and is rarely undertaken on commercial facilities due to cost considerations.' However, a survey of striped bass producers in the southeastern USA indicated that a salinity of 0.5 g/L or greater was associated with greater production (Geiger and Parker, 1985). Withholding feed before fish are handled or transported is commonly practiced by aquaculturists, regardless of the species concerned. Withholding feed is undertaken to reduce ammonia buildup in transport tanks resulting from increased rates of ammonia excretion and regurgitated feed, which can occur if fish are handled with feed in the digestive tract. The time required to adequately purge fish before handling varies according to fish size and water temperature. Piper et al. (1982) suggests that small (<10 cm) fish should be starved at least 48 h prior to handling, while larger fish should be starved at least 72 h. Carmichael et al. (1984) recommended that feed be withheld from largemouth bass (Micropterus salmoides) for 72 h before handling. Longer periods of time may be required to adequately purge fish present in cold water temperatures (e. g., < 15 ~ In general, most striped bass and hybrid striped bass producers withhold feed from 48 to 72 h before harvesting and transporting phase II and larger fish, especially if fish are to be transported to another site. Feed should also be withheld from ponds containing phase I fish, however, fish of this size are usually actively consuming zooplankton, making it difficult to completely purge these fish before they are loaded from ponds. During on-farm transit, there is less danger for ammonia buildup because fish are usually only confined in the transport unit for a short period of time. Withholding feed is still recommended however, because regurgitated feed could foul the water in which fish are being transported. To eliminate stress associated with exposure of fish to abrupt temperature changes, the transport water temperature should approximate that of the pond from which fish are being loaded. To achieve this, culturists either use pond water to fill transport tanks or use groundwater adjusted to the temperature of the pond. The most common method to adjust the temperature of groundwater is to fill the transport unit the evening before fish are loaded, which allows the water temperature to equilibrate with ambient air temperature. If pond water is used to fill transport tanks, care should be taken to ensure that pond water quality is within acceptable limits. When using pond water to transport fish for extended periods of time, it is usually necessary to flush or exchange the water with a fresh source due to the relatively high organic load present.

198

7.4.2 Preconditioning Fish In Holding Tanks After phase I and phase II striped bass and hybrid striped bass are harvested from ponds, they are usually transported to an on-farm holding facility where they are held until further transport. Fish are held from 24-72 h or longer depending on several factors including the readiness of ponds for stocking, weather conditions, and market availability. Use of a holding facility provides the culturist with an opportunity to perform a number of procedures for preconditioning fish for long-distance transport, which can aid in overall marketability of fish. For example, fish may be graded to prevent cannibalism and obtain a uniform size class to stock into production ponds or sell to buyers. Feeding is usually undertaken during the holding period until 48-72 h before transport. The practice of feeding fish while they are being held ensures that fish are in good health before transport and is especially useful in "training" phase I fish to accept artificial diets. Fish are normally held in circular or rectangular tanks (Figure 7.10) that are normally flow-through systems supplied by high-quality groundwater. Some holding facilities however, use closed-recirculating systems that allow the culturist to manipulate the concentration of water additives, such as salts and anesthetics, more readily than in flow-through systems. Nevertheless, water additives can be effectively added to flow-through systems by either shutting off the water supply or by periodically adding these substances to achieve "pulsed" concentrations. The former method is normally used to add anesthetics or disease therapeutants, while the latter is used to add various salts. Maintenance of salt levels in flow-through systems may be achieved by continuously adding a concentrated solution of the salt via peristaltic pump or a simple drip delivery system. The addition of salts to increase calcium concentrations and salinity has been shown to increase postharvest survival of striped bass and hybrid striped bass. Survival of phase I striped bass and sunshine bass held in tanks for 26 h after harvest was significantly increased in tanks which received added CaCI2 to increase total hardness from 10 to 70-200 mg/L as CaCO3 (Grizzle et al., 1985). Mauldin et al. (1986) observed that phase I striped bass harvested from ponds with low hardness had significantly increased survival over controls,when they were held in tanks in a solution of 150 mg/L CaCI 2 (approximately 60 m g ~

Fig. 7.10. Rectangular tanks constructed of concrete used to precondition fish before long-distance transport. (Photo by C. Weirich)

199

calcium) for 6 h. Highest survival rates were obtained when fish were held in tanks at this calcium concentration after harvest from ponds with similar calcium levels. The addition of NaCI to increase salinity in holding tanks was shown to increase post-harvest survival of phase I striped bass only if the calcium concentration was also increased (Grizzle et al., 1990). They examined survival rates for fish transported from ponds and held 24 h in untreated (soft) hatchery water (8-10 m ~ total hardness as CaCO3), the same water with addition of either CaC12(278 mg/L-approximately 100 mg/L calcium) or NaC1 (5 g/L), and water where both salts were combined at these concentrations (CaCI2 + NaCI). They concluded that while the addition of CaC12 alone increased post-harvest survival, the combination of CaCI., + NaC1 provided the best transport and holding medium. Survival was reduced in other treatments and was lowest when fish were transported and held in soft water with added NaCI. In an earlier study, phase I striped bass held for 3 h after pond harvest in tanks containing soft water (20 mg/L total hardness as CaCO3) with added NaC1 (10 g/L) exhibited abnormal behavior coupled with a grossly visible, hemorrhagic area adjacent to the vertebrae at the junction of the trunk and tail (Grizzle et al., 1985). When total hardness was increased to 200 mg/L as CaCO3 these clinical signs were immediately alleviated. These results indicate that while NaC1 may mitigate the stress response of phase I fish, increasing the salinity of waters deficient in calcium is deleterious. Although calcium and salinity may be maintained at levels shown to mitigate stress in closed system holding facilities, producers utilizing flow-through systems must either have access to a high-quality water source of sufficient calcium and salinity or add salts periodically to maintain concentrations at acceptable levels. Generally, most flow-through systems are supplied with fresh groundwater, which is usually adequate with respect to calcium concentrations but is deficient in salinity. Salinity is increased in these systems by the addition of NaCl or commercially available sea salts. One producer utilizing a flow-through facility who has had success in holding phase I and phase II sunshine bass supplied by hard groundwater (> 150 mg/L total hardness as CaCO3) routinely adds NaCI to maintain the salinity between 1 and 3 g/L. When fish are being held they should be monitored closely for any mortalities or possible disease outbreaks. Mortalities due to handling stress incurred when fish were loaded and transported from ponds are often detectable within 24 h, and dead fish should be removed to prevent water fouling. The holding period also allows time for any fish weakened from transport-induced stressors to die or recover before they are further transported. If a disease problem is detected, fish may be readily treated. Prophylactic treatments may also occur. For a discussion on diseases and disease treatments see Chapter 11. Before fish are loaded from holding tanks into transport units, feed is normally withheld, approved anesthetics may be added to the water to reduce fish metabolism, and fish are acclimated or "tempered" to transport unit water temperatures. If fish are to be transported for long distances, feed should be withheld 48-72 h before loading. The use of anesthetics before loading has been shown to reduce stress and increase survivability of largemouth bass (CarmichaeI et al., 1984) and sunshine bass (Tomasso et al., 1980), but this practice is not widely used by commercial striped bass and hybrid striped bass producers (with exception ofbroodfish). When used in holding tanks, anesthetics are generally added at higher concentrations than in transport units (see Section 7.7.2). Davis and Parker (1990) evaluated the stress response of yearling striped bass exposed to net confinement at water temperatures of 5, 10, 16, 21, 25, and 30~ Physiological changes, (plasma cortisol, glucose, chloride, hematocrit) were lowest at 10 and 16 ~ temperatures which border the range suggested for transport. These water temperatures are normally cooler than those of most groundwater sources supplying flow-through systems in the Southern U.S. and, depending on the time of year, cooler than ambient

200

temperatures present in closed system holding tanks. Therefore, the water temperature of holding tanks usually must be reduced to match suggested transport temperatures or fish may first be loaded and transport unit temperatures then lowered. If possible, it is best to lower holding tank temperatures before loading. This procedure, in addition to tempering fish to transport conditions, allows fish to be handled at temperatures which have been shown to minimize handling-induced stress. Water temperatures may be lowered through the use of chilling units, addition of groundwater of reduced temperatures, or ice. The use of chilling units is normally restricted to closed-system holding facilities. Ice may be added to flow-through facilities without access to a cold water source, but in most cases fish are first loaded from tanks, then tempered in the transport unit. Recommended tempering and temperature requirements for transport will be discussed further in section 7.6.3. 7.5 LOADING RATES The weight of striped bass or hybrid striped bass which can be safely transported is dependent on several variables including fish size, water temperature, transport duration, and the efficiency of the transport units aeration and oxygenation system. Striped bass and hybrid striped bass are normally transported in two fashions; 1) short-term (< 4h) or on-farm movements utilizing transport units normally equipped with compressed oxygen and/or electric agitators, or 2) long-term transport using transport units equipped with liquid oxygen. Short-term transport events are undertaken to move fish from ponds by using pond water or groundwater allowed to equilibrate with ambient air temperature which, depending on the time of year, is often greater than temperatures used during long-term transport. Although more fish can be transported per unit volume at lower water temperatures the short duration of on-farm transport events allows loading rates to be increased relative to those used in long-term transport. Aside from environmental and physical factors (e.g., transport unit design, type of oxygenation system), the most important variable affecting loading rates is fish size. As a general rule, fewer grams of smaller fish can be transported per liter of water than of larger fish. This is due to the fact that smaller fish have increased oxygen consumption rates relative to larger fish on a per weight basis.

Table 7.1. Suggested loading rates (~JL) of striped bass or hybrid striped bass during short-(< 4h) or long-term transport.

Duration of Transport

Fish Size

(Growth stage)

Short-term

Long-term

Phase I

60

30

Phase II

120

60

Phase III

180-240

90

Broodfish

240

180

201

Suggested loading rates based on fish size (phase I -broodfish) and the duration of the transport event (short or long-term) as described previously are listed in Table 7.1. These values are based on loading rates recommended by several sources (Bonn et al., 1976; Geiger and Parker, 1985; Parker et al., 1990), including several commercial producers. Loading rate estimates are presented only as guidelines to calculate the maximum safe loading rates. Practical experience should serve as the final determinant of actual loading rates to be used on individual production facilities. To realize correct loading rates in grams of fish per liter, the water displaced by fish must be taken into account. As mentioned previously every kilogram (1,000 grams) of fish displaces one liter of water. 7.6 WATER QUALITY CONSIDERATIONS DURING FISH TRANSPORT Maintaining water quality characteristics within acceptable levels is essential for successful fish culture in general and becomes even more critical during transport. Under transport conditions, fish are present at high densities in a closed environment. This, coupled with the increased metabolism of fish normally realized after handling, can result in rapid deterioration of water quality conditions if proper transport equipment and techniques are not used. Detailed information regarding environmental requirements for striped bass and hybrid striped bass is presented in Chapter 10. This section will be restricted to a discussion concerning water quality characteristics of importance during fish transport. These are: dissolved oxygen, carbon dioxide, temperature, ammonia and pH. 7.6.1 Dissolved Oxygen Dissolved oxygen is the primary limiting factor to consider regarding fish transport. Failure to supply adequate quantities of oxygen to fish during transport may cause hypoxia, resulting in either acute or delayed mortality. The amount of oxygen required by fish is dependent on several variables including fish size, metabolic activity, water temperature, and the ambient concentrations of dissolved oxygen, carbon dioxide, and ammonia. Oxygen consumption per unit of fish weight decreases with increasing fish size and increases with increased water temperature, fish activity, and the presence of feed in the digestive tract (Busch, 1985). Increased levels of carbon dioxide and ammonia also increase oxygen requirements, as they interfere with oxygen utilization (see following sections). To ensure adequate dissolved oxygen during transport, concentrations should be maintained near saturation at all times. The amount of oxygen at saturation is determined primarily by water temperature and, to a lesser extent, salinity (see Table 7.2). Care should be taken to ensure that concentrations do not exceed saturation levels since effects due to hyperoxic conditions have been observed to occur in some species (Wood, 1991). Fish activity is usually greatest immediately after handling and loading, therefore, it is imperative that oxygen concentrations are established at acceptable levels before fish are loaded. This is especially important if transport tanks have been recently filled with anoxic groundwater. Groundwater may also contain hydrogen sulfide, which although highly toxic to fish, is easily oxidized into its non-toxic form in the presence of sufficient oxygen.

202

7.6.2 Carbon Dioxide Carbon dioxide (CO2) is a waste product of fish metabolism, and thus tends to accumulate during fish transport. Dissolved CO2 gas is directly toxic to fish as it reduces the oxygen carrying capacity of blood, however if adequate oxygen levels are maintained, fish can tolerate relatively high concentrations of COz For example, Dupree and Huner (1984) observed that channel catfish tolerate CO2 concentrations of 20-30 mg/L if the rate of CO2 buildup is slow and sufficient dissolved oxygen is present. Piper et al. (1982) noted that trout are unaffected by CO2 concentrations below 15 mg/L, but become stressed when levels approach 25 mg/L. Carbon dioxide concentrations may be reduced through vigorous aeration or agitation coupled with adequate ventilation. Since, as described in Section 7.3.3, transport units are normally supplied with liquid oxygen, which provides minimal aeration and CO2 removal. To avoid CO 2buildup in these situations correct loading rates should be used and transport units should be equipped with sufficient ventilation (e.g., air scoops). Commercially available antifoam agents used by some culturists to reduce foaming can inhibit gas transfer and should therefore be used with caution, especially during long-term transport using liquid oxygen.

Table 7.2. Saturation values of dissolved oxygen (mg/L at sea level) at varying temperature and salinity.

Salinity (g/L) Temperature ( ~C)

0

10

20

30

10

11.3

10.6

9.9

9.3

12

10.7

10.1

9.5

8.9

14

10.3

9.7

9.1

8.6

16

9.9

9.3

8.7

8.2

18

9.5

8.9

8.4

7.9

20

9.1

8.6

8.1

7.6

22

8.7

8.2

7.8

7.3

24

8.4

7.9

7.5

7.1

26

8.1

7.7

7.2

6.8

28

7.8

7.4

7.0

6.6

30

7.6

7.1

6.8

6.4

32

7.3

6.9

6.5

6.2

7.0

6.7

6.2

6.0

34 Source: Weiss (1970).

203

7.6.3 Temperature Temperature influences not only fish activity but also several water quality characteristics, and is thus an important factor to consider during fish transport. The metabolism of fish increases with increasing temperature, which results in increased oxygen consumption rates. Increasing temperature also lowers the concentration of dissolved oxygen at saturation, and intensifies the effects of carbon dioxide and ammonia. Because of these factors, most fish are transported in relatively cool water, especially if the duration of transport is for an extended period of time. Parker et al. (1990) recommend that hauling temperatures should be kept below 20~ during long-term transport and 24~ during short-term transport of phase I and phase II fish. Davis and Parker (1990) showed that the stress response of net-contained striped bass was lowest at 10~ and 16 ~ and recommended that, when possible, fish should be transported within this temperature range. Most striped bass and hybrid striped bass culturists transporting fish for long distances establish transport unit temperatures between 13 ~C and 15 ~ During short-term transport, temperature is less crucial, but extreme caution should be used when fish are transported in water temperatures exceeding 24 ~ Another important consideration involving temperature is the avoidance of abrupt temperature changes that can occur when fish are moved from one water source to another without being tempered. In general, most recommendations suggest that if the temperatures between holding and transport unit water differ by more than 3~ tempering should be undertaken. Dupree and Huner (1984) recommend that for every 6~ difference in water temperature, at least 20 minutes of tempering be allowed. Longer periods of temperature acclimation may be required for striped bass and hybrid striped bass. Davies (1973) noted that habituating striped bass for several hours each time they were transferred to a different temperature improved survivability. The process of tempering also helps acclimate fish to other environmental differences such as pH and salinity, although both striped bass and hybrid striped bass tolerate transfer from waters of varying salinity fairly well (Wattendorf and Shafland, 1982; Grizzle et al., 1992). 7.6.4 Ammonia Ammonia, like CO2 is a waste product of fish metabolism and tends to accumulate in transport water. Ammonia is directly toxic to fish in the unionized form, which is favored at high temperatures and pH, and it also reduces the ability of fish to utilize oxygen. To avoid the accumulation of excessive ammonia, feed should be withheld from fish before transport and proper loading rates should be used. To reduce the effects of ammonia, fish should be transported in reduced water temperatures, when possible. Weirich et al. (1993) noted that calcium addition seems to provide some benefits regarding ammonia toxicity, as the 96 h LC50 of ammonia for sunshine bass increased significantly when calcium concentrations were raised from 5 to 80 mg/L. 7.6.5 pH The pH of transport water tends to decrease with time during transport due to CO2 accumulation. While lower pH levels are beneficial with respect to ammonia toxicity, extremely low levels or rapid changes in pH should be avoided. The use of a high quality water source with elevated levels of total alkalinity and total hardness generally provides sufficient buffering capacity or stabilization of pH during transport. Sodium bicarbonate (baking soda) and calcium chloride may be added if needed to increase total alkalinity and total hardness respectively. The organic buffer trishydroxymethyl-aminomethane (Trizma| Sigma Chemical Co.) is quite effective in stabilizing pH levels during fresh and saltwater transport of aquarium fishes. Unfortunately, this compound is not currently registered by the FDA for use in food fish.

204

7.7 USE OF WATER ADDITIVES DURING TRANSPORT Significant mitigation of stress induced by handling and transport procedures can be achieved through the use of various water additives. These additives aid in the reduction of the stress response and generally fall under two broad categories: salts and anesthetics. Various salts can be added to prevent or reduce osmoregulatory dysfunction and resulting ionic imbalances, while anesthetics may be added to reduce locomotor activity and metabolic rate (McFarland, 1960). Bacteriostatic agents (antibiotics) have also been used concurrently with salts and anesthetics during fish transports. Many of these substances, however, are not currently registered for food fish use (See Chapter 11). 7.7.1 Addition of Salts Stressors associated with handling and transport procedures, as discussed in section 7.2, induce various physiological changes through activation of the hormonally-driven stress response. A detrimental consequence of the stress response is osmoregulatory dysfunction, which is often the primary factor causing transport mortality. Osmoregulatory dysfunction has been well documented as a secondary response to stress (Miles et al., 1974; Mazeaud et al., 1977; Eddy, 1981), and in freshwater fish is characterized by decreased levels of plasma ions (Weirich and Tomasso, 1991; Weirich et al., 1992). Fish lose their ability to effectively regulate ionic and osmotic constituents due to several factors including the repartitioning of energy from homeostatic activities to the induced stress response. These ionic imbalances and ensuing mortality can usually be prevented in freshwater fishes by modifying the ionic composition of the water in which fish are handled or transported. Commonly, calcium, sodium, and chloride are added (Tomasso et al., 1980; Carmichael et al., 1984; Carmichael and Tomasso, 1988: Weirich and Tomasso, 1991; Weirich et al., 1992). These ions have at least three general functions: 1) increased calcium concentrations act to decrease gill permeability reducing ion and water flux (Potts and Fleming, 1970; Oduleye, 1975; Potts, 1984; Hunn, 1985), 2) increased levels of sodium and chloride facilitate active transport of these ions across the gills, thus preventing or reducing ionic depletions (Pickering, 1981), and 3) Increased ionic concentrations reduce the osmotic gradient between fish plasma and the environment (Redding and Schreck, 1983; Mazik et al., 1991b; Weirich et al., 1992). To increase concentrations of these ions, various salts are added to transport water. The two most common additives are CaCI2 which is primarily added to increase calcium concentrations and NaC1 which is added to increase total ionic content (salinity). Commercial sea salt mixtures can also be added. Sea salt formulations not only increase salinity but also provide calcium (approximately 12 mg/L calcium per each g/L salinity). Several studies have evaluated the stress mitigating effects of increasing the calcium concentration and salinity, alone or in combination, of handling and transport water. Results from these studies indicate that the efficacy of various ion-enhanced transport media depends primarily on fish size and the type and concentration of ions present. In general, the presence of increased calcium levels seems to be more important than salinity regarding handling and transport of phase I fish, while the opposite is true for larger fish. Phase I striped bass that are approximately 30-40 d old have just completed metamorphosis and are sensitive to low levels of calcium (Grizzle et al., 1992; Grizzle and Mauldin, 1994). As noted in sections 7.4.1 and 7.4.2, post-harvest survival of phase I fish increased significantly when calcium concentrations

205

were increased in ponds and holding tanks (Grizzle et al., 1985; Mauldin et al., 1988; Grizzle et al., 1990). Survival of phase I striped bass was also increased during transport by increasing the calcium concentration of the transport water from <15 mg/L to approximately 100 mg/L (Grizzle et al., 1990). The authors also observed that, while a transport medium consisting of approximately 100 mg/L calcium and a salinity of 5 g/L (achieved through NaC1 addition) proved to be most effective, survivability was reduced in transport tanks containing untreated, low calcium (<15 m ~ ) water and was lowest in untreated water receiving NaCI addition. Similar results were observed for striped bass ranging in age from 16 to 54 d (post-hatch) which were transferred to water containing high or low concentrations of sodium and calcium (Grizzle and Mauldin, 1994). This paradox (increasing salinity in the presence of sufficient calcium is beneficial while increasing salinity in calcium-deficient water is not) may be explained by considering both the detrimental effects of low levels of environmental calcium and high levels of environmental sodium on gill permeability, and the beneficial effects of increased sodium (salinity) in reducing ionic gradients. Low concentrations of environmental calcium have been shown to increase gill permeability to water and various ions (McWilliams and Ports, 1978; Pic and Maetz, 1981; Wendelaar Bonga and Van der Meij, 1981; Ogasawara and Hirano, 1984), thus exacerbating the effects of stress-induced osmoregulatory dysfunction. Gill permeability has also been shown to be increased by the loss of surface-bound calcium from the gills of brown trout, (Salmo trutta), caused by low environmental calcium levels, low pH, or high environmental sodium levels (McWilliams, 1983). High sodium levels may also increase gill permeability indirectly through the action of corticosteroid hormones (Donaldson, 1981; Mazeaud and Mazeaud, 1981; Rankin and Bolis, 1984). Davis et al. (1982) noted that corticosteroid levels increased in striped bass contained in 10 g/L NaCI and were further elevated when fish were exposed to handling stress. Grizzle et al. (1993) evaluated the effects of various concentrations of environmental calcium and sodium on calcium flux of phase I striped bass after harvest and transport. Flux was measured in fish confined in flux chambers within 24 h after transport. Four environmental treatments were present in flux chambers: high environmental sodium-low environmental calcium, high sodium-high calcium, low sodiumhigh calcium and low sodium-low calcium. Treatments containing water of high (1,954 mg/L) sodium and low (5 mg/L) calcium caused calcium efflux to increase and calcium influx to decrease and was the only treatment that resulted in a net loss of calcium. However, the authors did not detect any decrease in the concentration of whole-body soluble calcium in fish contained in this treatment, which is in contrast to the decreased soluble calcium concentrations observed by Grizzle et al. (1990) of fish contained in either water containing high sodium-low calcium levels or high sodium-high calcium levels. Grizzle et al. (1993) suggested that although changes in soluble calcium levels were probably not the cause of death in high sodium-low calcium treatments in either study, the lethal effects of this ion combination could be related to changes in calcium concentrations in pools which were not measured, or due to alterations in sodium flux. The permeability of the gill to sodium and hydrogen ions is inversely related to the environmental concentration of calcium (McWilliams and Potts, 1978). Therefore, low environmental calcium tends to increase sodium efflux (Frain, 1987) through increased gill permeability. This effect is intensified in combined environments of low calcium and high sodium due to the increased loss of surface-bound calcium caused by elevated environmental sodium levels (McWilliams, 1983). This may explain the reduced survival observed by Grizzle et al. (1990) for phase I striped bass handled and transported in water containing insufficient levels of calcium and the potentiation of this effect when sodium is added to low-calcium environments.

206

The enhanced survival of phase I fish transported in water containing both calcium and sodium (salinity) at high levels as compared to water containing only high levels of calcium may be due to the reduced plasma-environmental osmotic gradient present in media receiving NaC1 addition. As discussed below, increasing the salinity to approximate isosmotic conditions increases survival of larger striped bass (Mazik et al., 1991b; Harrell, 1992) and sunshine bass (Tomasso et al., 1980; Weirich et al., 1992) subjected to handling and transport stressors. The increased levels of calcium in media used to transport phase I fish containing high levels of sodium (salinity) may thus negate detrimental effects of increased sodium by reducing gill permeability while at the same time allowing beneficial reductions in osmotic gradients. An interesting manifestation of handling and transport induced stress in striped bass and hybrid striped bass is the development of what is commonly termed "sudden death syndrome" in which fish die immediately after handling procedures are initiated. This syndrome has been observed to occur at harvest of all three phases of striped bass and hybrid striped bass, although phase I and phase II fish seem most susceptible. Affected fish usually exhibit a rigid tetany with mouth open, gills flared, and backbone bent laterally. Grizzle et al. (1985) observed fractured vertebrae in sectioned specimens of affected fish that may have been caused by poorly mineralized bone or extreme muscle contractions associated with tetanic convulsions. Most reports of mortalities due to this syndrome have been anecdotal observations conducted in the field or during related experiments (Grizzle et al., 1985, 1990; Weirich et al., 1992, 1993). In all cases, affected fish were being handled in water sources containing low levels of environmental calcium. The tetanic convulsions and associated clinical signs of affected fish indicate a neuromuscular dysfunction. Both calcium and sodium are important ions in the proper propagation of nerve impulses controlling muscular contraction (Alberts et al., 1983). It may be possible that fish present in low ionic environments are unable to maintain adequate stores of these ions. While low levels of environmental calcium have been shown to not affect production of sunshine bass reared under normal culture conditions (Seals et al., 1994), low calcium (and perhaps sodium) stores may affect the ability of fish to cope with the immediate effects of severe handling stress. Although Mazik et al. (199 l b) observed that striped bass were able to maintain plasma calcium levels during periods of primary stress and osmoregulatory dysfunction associated with transport, calcium in other stores, which have not been experimentally measured under stressful conditions, (e.g., muscle) may be affected. The role of environmental calcium in reducing handling and transport-induced stress seems less important than salinity regarding larger fish. Mazik et al. (1991 b) evaluated physiological characteristics of phase II striped bass during and for one month after transport in water containing either 1.0% NaC1, 0.1% CaC12, or freshwater without the addition of salts. While the addition of NaC1 to transport and recovery water decreased the stress response (decreased rise in plasma cortisol and glucose concentrations, decreased osmoregulatory dysfunction) and improved survival, the use of fresh water or water with added CaCI2 did not reduce the stress response or improve survival. These results are in contrast with the findings of Weirich et al. (1992), who subjected phase II sunshine bass to severe net confinement for 6 h in fresh water containing calcium levels ranging from 5 to 80 mg/L. Although increased environmental calcium concentrations did not inhibit the development of osmoregulatory dysfunction, survival was significantly affected by environmental calcium concentrations and was greatest at 80 mg/L. It is possible that the degree of stress associated with severe net confinement during this study was greater than handling and transport stressors induced by Mazik et al. (1991 b), which may account for the different results obtained. Nevertheless, the results reported by Weirich et al. (1992) indicate that calcium is important in reducing stress-induced mortality in larger fish although its role is probably less than that of salinity since increasing calcium failed to prevent osmoregulatory dysfunction.

207

Increasing the salinity to achieve isosmotic or near-isosmotic environments has been shown to reduce the stress response and improve survival of phase II and larger striped bass and hybrid striped bass subjected to handling and transport stress (Tomasso et al., 1980; Mazik et al., 1991; Harrell, 1992; Weirich et al., 1992; Young and Cech, 1993). In all these studies, with the exception of Weirich et al. (1992), a transport medium with a salinity of approximately 10 g/L significantly reduced the effects of transport-induced stress and/or improved survivability (Weirich et al., 1992 observed similar results in water containing 8 g/L salinity). Based on baseline plasma osmolality values obtained for striped bass by Tisa et al. (1983) and for sunshine bass by Weirich et al. (1992), an environmental salinity of approximately 10 g/L approximates isosmotic conditions. The beneficial effects realized by handling and transporting fish in isosmotic or near isosmotic environments, in addition to reducing the plasma-environmental osmotic gradient resulting in decreased energy costs associated with osmoregulation (Redding and Schreck, 1983), are probably due to the prevention of prolonged increased levels of corticosteroid hormones, notably cortisol, which increases gill permeability resulting in water influx and ion effiux (Donaldson, 1981; Mazeaud and Mazeaud, 1981; Rankin and Bolis, 1984). Although increased plasma corticosteroid levels have been reported for striped bass (Davis et al., 1982; Young and Cech, 1993) and sunshine bass (Tomasso et al., 1980) during handling or transport in isosmotic or near-isosmotic media, recovery of fish, evidenced by the return of corticosteroid levels to baseline values within 4 h, was facilitated through use of the same medium used for handling (Young and Cech, 1993). In contrast, plasma corticosteroid levels of fish handled or transported in, or allowed to recover in, fresh water have been shown to remain elevated for extended periods of time (Tomasso et al., 1980; Mazik et al., 1991 b; Young and Cech, 1993). 7.7.2 Anesthetics The use of anesthetics, alone or in combination with the addition of salts, has been employed in an attempt to mitigate handling and transport stress. The primary function of anesthetics is to reduce the metabolic activity of fish thereby lowering oxygen consumption, carbon dioxide production, and ammonia excretion. Sedated fish exhibit reduced locomotor activity and are thus less prone to cause physical injury to themselves or other fish, which can be especially important when large fish are handled. Several different anesthetics have been evaluated for use during handling and transport procedures. Davis et al. (1982) monitored plasma corticosteroid and chloride levels of phase II striped bass after exposure and net confinement in water with either tricaine methanesulfonate (tricaine), quinaldine sulfate, or etomidate, each alone or in combination with 10 g/L NaC1. While chloride levels were unaffected by all treatments, corticosteroid levels increased in fish exposed to salt alone, and in fish exposed to 25 mg/L tricaine or 2.5 mg/L quinaldine, alone or in combination with salt. Water containing 0.1 m g ~ etomidate, with or without salt addition, were the only treatments which limited the increase of plasma corticosteroids during exposure and confinement. Unfortunately, etomidate and quinaldine are not registered for use on food fish. Tricaine, therefore, is the only legal anesthetic to be used on food fish. A 21-day withdrawal period is required before fish are processed. Tomasso et al. (1980) evaluated plasma corticosteroid and chloride levels of sunshine bass during and after exposure to netting and hauling stress in water containing 25 mg/L tricaine, alone or in combination with 10 g/L NaCI. While, as observed by Davis et al. (1982), increased corticosteroid levels were detected in response to netting and hauling regardless of the medium tested, corticosteroid levels of fish stressed in the combined tricaine-salt medium returned to normal levels within 24 h after stress. Although plasma chloride levels were unaffected during netting and hauling, hypochloremia developed in all groups of fish within 24 h after netting and hauling, except in those exposed to stress in the combined tricaine-salt medium. The stress response was further reduced when fish were exposed to 50 mg/L tricaine for 15 min before

208

hauling in the combined tricaine-salt medium. In contrast, Harrell (1992) reported that stress associated with capture of striped bass broodfish was mitigated (based on the time required for corticosteroid levels to return to baseline levels) most effectively by the use of water containing 10 ~ L salt without tricaine addition. The combination of salt and anesthetic was second in effectiveness. The author, however, noted that based on the prevention of the development of hypochloremia after handling and transport, the combined salt and anesthetic medium appears to be as effective as salt alone. Deleterious affects associated with handling or transporting fish in anesthetic alone noted by this and other (Tomasso et al., 1980; Davis et al., 1982) studies show that tricaine should only be used in combination with NaC1 or sea salt addition. 7.7.3 Practical recommendations Based on the studies described in the previous two sections, phase I striped bass should be handled and transported in water containing 100 mg/L calcium with the addition of 5-8 gr NaCI or sea-salt. It is imperative that calcium levels be sufficient when handling or hauling fish of this size, and the salinity should not be increased in water that is calcium deficient. Phase II and larger fish should be handled and transported in isosmotic or near-isosmotic media (8-10 g/L NaCI or sea salts). While possible benefits of increasing the calcium level of isosmotic or near isosmotic media have not been evaluated in larger fish, it is recommended that, as with phase I fish, calcium should be increased to 100 m g ~ . If the anesthetic tricaine is used, it should be added only in combination with the addition of NaCl or sea salts. Caution should be used in adding tricaine to water used during long-term transport events since oxygen utilization may be impeded due to asphyxiation (Parker et al., 1990). Transport water should also be properly buffered as use of tricaine in poorly buffered water can cause a reduction in pH (Rees and Harrell, 1990). While a concentration of 25 mg/L tricaine is useful in stress reduction (in combination with increased salinity) when handling and transporting phase II and larger fish, Yeager et al. (1990) recommends that to avoid disrupting spawning of broodfish, tricaine concentrations ranging from 3-8 mg/L be used. The use of tricaine when handling and transporting phase I fish has not been evaluated scientifically and is rarely used, however, it is added at concentrations used to sedate larger fish by some culturists. When possible, based on the results obtained by Tomasso et al. (1980), it is recommended that phase I and larger fish be sedated before transport using 50 mg/L tricaine for 15 min. 7.8 POST-CONDITIONING OF FISH AFTER TRANSPORT Full expression of physiological changes due to the stress response induced by handling and transport procedures often does not occur until some time after the actual transport event. For example, hypochloremia did not develop in striped bass transported, depending on the transport medium used, until 24 to 72 h after hauling (Tomasso et al., 1980). In addition, physiological changes occurring immediately during transport, such as increased corticosteroid levels (again depending on the transport medium used), may be evident for several hours to days afterwards. Wagner (1987) stated that recovery from handling or transport stress may take from 6-48 h depending on the length of time and the degree of disturbance, although results reported by Tomasso et al. (1980) and Mazik et al. (1991 b) show that effects may be even more long lasting (up to 72 h for some variables). While proper transport techniques and recommended media can prevent or reduce the stress response, ideally fish should be allowed to recover, preferably in the same or similar medium used for transport, for at least 72 h. This practice, especially useful after long-term transport, may easily be accomplished if culturists have access to holding facilities such as described in section 7.4.2. In some cases however, fish must be transferred from the transport unit directly into ponds. In these situations, every effort should be made to minimize environmental differences (especially temperature) between the transport medium and pond water before fish are unloaded.

209

7.9 TRANSPORT OF LARVAE

Larval striped bass and hybrid striped bass are commonly transported through the use of closed-system transport containers. These containers allow large numbers of fish to be transported with a minimal amount of water and its associated weight, which makes this the preferred method for air shipment. Closed-system transport containers normally consist of plastic bags placed into styrofoam or otherwise insulated boxes or cartons. Polyethylene bags with four comers on the bottom are desirable as they reduce the possibility of trapping and suffocating larvae. Larvae shipped by aircraft should be double-bagged and slightly under inflated to avoid bag leakage due to pressure differentials occurring during flight. After water and fish are added, air is forced out of the bag which is then refilled with pure oxygen gas. Approximately 75% of the volume of the bag should be filled with oxygen (Busch, 1985). The bag is then heat-sealed or bound shut with rubber bands. The amount of water placed in the bag depends on the container size and generally ranges from approximately >7.5-20 L. Larvae may be transported at 1 day of age, however, most are transported at 5-7 days of age (Rees and Harrell, 1990). Recommended temperatures for bag transport of larvae generally range from 14-18~ . Ice packs placed between the carton and bag are used to stabilize temperatures. Suggested loading rates (number of larvae per liter) for bag transport of larvae are listed in Table 7.3 Note that as with larger fish, loading rates decrease with increasing duration of transport. Loading rates used for closed-system transport are primarily limited by the accumulation of ammonia and CO2. Water additives commonly used for bag transport of larval striped bass and hybrid striped bass include salts (e.g., NaCI, sea salts, CaCI2), buffers, and ammonia-removal compounds such as zeolite. Larval striped bass can survive in fresh water but survival is increased in diluted seawater (Albrecht, 1964; Lal et al., 1977) or in fresh water receiving NaCI addition (Germann and Reeves, 1975). Grizzle et al. (1992) observed that adding 4-5 gfL NaCI increased the survival of larval striped bass after a 6 h confinement period when compared to larvae confined in fresh water with or without CaC12 addition. In a similar study, survival of 9 d old striped bass larvae transferred to water containing high or low levels of sodium and calcium was greatest in high sodium treatments and was unaffected by calcium (Grizzle and Mauldin, 1994). The absence of beneficial effects due to calcium addition for larval striped bass are in contrast with results obtained for phase I fish (Grizzle et al., 1990; Grizzle and Mauldin, 1994) and may be due to differences in gill structure. Larval striped bass possess rudimentary gills which are probably limited with respect to ion regulatory capabilities, which may account for the different observations regarding the effect of calcium between larval and phase I fish (Grizzle et al., 1992). Larvae may also be unaffected by low environmental calcium because of yolk-sac reserves of this ion (Grizzle and Mauldin, 1994). Nonetheless, although not statistically different from treatments receiving NaCI alone, Grizzle et al. (1992) observed a combined salinity and calcium (100 mg/L) treatment was the best medium with respect to survival after confinement. In addition, other studies suggest that calcium (hardness) enhances larval survival (Hazel et al., 1971; Bonn et al., 1976; MurrayBrown, 1987; Kane et al., 1990) and thus, while not as important a factor as salinity, may provide some physiological benefit to larval fish. Therefore, it is recommended that larval striped bass and hybrid striped bass be transported in water containing 4-5 g ~ salinity and 100 mg/L calcium. Accumulation of C O 2 in bags can cause a considerable decrease in pH if the water is insufficiently buffered. The buffering capacity of water with low levels of total alkalinity can be enhanced through the addition of sodium bicarbonate. As mentioned in Section 7.6.5, commercially available buffer formulations such as Trizma| while effective, are not currently registered for use on food fish. Ammonia accumulation can be reduced somewhat through the addition of natural (clinoptilolite) or synthetic zeolite: ion exchange

210

media that remove ammonia from water (Marking and Bills, 1982). While natural zeolite is superior to synthetic zeolite (Johnson and Sieburth, 1974; Chiayvareesajja and Boyd, 1993), the efficiency of both forms decreases with increasing concentrations of dissolved cations, which compete with ammonia for exchange sites on the zeolite. Thus the ability of zeolite to remove ammonia is reduced as the calcium concentration or salinity of the transport water is increased. Amend et al. (1982) observed that adding clinoptilolite at a concentration of 14 ~ controlled ammonia accumulation during bag transport of fish (total hardness 160180 mg/L as CaCO3). Slightly higher concentrations (20-30 g ~ ) of clinoptilolite are currently used by several producers in closed-system transport of striped bass and hybrid striped bass. Zeolite is normally added to bags in the form of small (200-450 g -- depending on the volume of water used) packets. Packets are usually rinsed free of zeolite dust before being added. Once shipments of larvae reach their destination, bags should be removed from cartons and placed, unopened, in the receiving water for at least 30 min to allow temperature acclimation. Bags are then opened with small amounts of the receiving water added at intervals for another 30 min. Bags should not be opened and aerated since rapid removal of accumulated CO., will increase pH and subsequently increase the percentage of ammonia in the unionized (toxic) form. Larvae are fairly sensitive to ultraviolet light (McHugh and Heidinger, 1978) and should not be exposed to bright sunlight during any phase of transport. If possible, larvae should be introduced in ponds at dusk (Brewer and Rees, 1990). Phase I striped bass and hybrid striped bass are sometimes transported through the use of closed system containers. The same procedures used for bag transport of larval fish are applicable, with the exception of loading rates, which are somewhat lower than those for larval fish (see Table 7.3). Amend et al. (1982) provide guidelines and suggestions for the closed-system transport of fish which are applicable for phase I bag transport.

Table 7.3. Suggested loading rates (number of larvae or fish/L) for closed-system, bag transport of striped bass or hybrid striped bass based on duration of transport.

Duration of Transport Growth Stage

8h

8 h and above

Larvae

10,00 O-13,000

5,000-6,000

Phase I

100-160

50-80

211

References

Alberts, B., Bray, D., Lewis, J., Raft, M., Roberts, K. and Watson, J.D., 1983. Molecular biology of the cell. Garland Publishing Company, New York. Albrecht, A.B., 964. Some observations on factors associated with survival of striped bass eggs and larvae. California Fish and Game, 50:100-113. Amend, D.F., Croy, T.R., Goven, B.A., Johnson, K.A and McCarthy, D.H. 1982. Transportation of fish in closed systems: methods to control ammonia, pH, and bacterial growth. Transactions of the American Fisheries Society, 111: 603-611. Barton, B.A. and Iwama, G.K. 1991. Physiological changes in fish from stress in aquaculture with emphasis on the response and effects of corticosteroids. Annual Review of Fish Diseases, 1: 3-26. Barton, B.A. and Peter, R.E., 1982. Plasma cortisol stress response in fingerling rainbow trout, Salmo gairdneri (Richardson), to various transport conditions, anesthesia, and cold shock. Journal ofFish Biology, 20:39-51. Bonn, E.W., Bailey, W.M., Bayless, J.D., Erickson, K.E. and Stevens, R.E., 1976. Guidelines for striped bass culture. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Busch, R.L. 1985. Harvesting, grading and transporting. Pages 543-568 in Tucker, C.S. (editor). Channel catfish culture. Elsevier, Amsterdam. Brewer, D. L. and Rees, R.A., 1990. Pond culture of phase I striped bass. Pages 99-120 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Carmichael, G.J. and Tomasso, J.R., 1988. Survey of fish transportation and techniques. The Progressive FishCulturist, 50: 155-159. Carmichael, G.J., Tomasso, J.R., Simco, B.A. and Davis, K.B., 1984. Characterization and alleviation of stress associated with hauling largemouth bass. Transactions of the American Fisheries Society, 113: 778-785. Carmichael, G.J., Wedemeyer, G.A., McCraren, J.P. and Millard, J.L., 1983. Physiological effects of handling and hauling stress on smallmouth bass. The Progressive Fish-Culturist, 45:1 I0-113. Chiayvareesajja, S. and Boyd, C.E., 1993. Effects of zeolite, formalin, bacterial augmentation, and aeration on total ammonia nitrogen concentrations. Aquaculture, 116: 33-45. Colt, J. and Watten, B., 1988. Applications of pure oxygen in fish culture. Aquacultural Engineering, 7:397-441. Cech, Jr., J.J., Bartholow, S.D., Young, P.S. and Hopkins, T.E., 1990. Physiological recovery of striped bass from exercise stress. Physiologist, 33: 4. Davies, W.D., 1973. Rates of temperature acclimation for hatchery reared striped bass fry and fingerlings. The Progressive Fish-Culturist, 35: 214-217. Davis, K.B., Parker, N,C. and Suttle, M.A., 1982. Plasma corticosteroids and chlorides in striped bass exposed to tricaine methanesulfonate, quinaldine, etomidate, and salt. The Progressive Fish-Culturist, 44: 205-207.

212

Davis, K.B. and Parker, N.C., 1983. Plasma corticosteroid and chloride dynamics in rainbow trout, Atlantic salmon, and lake trout during and after stress. Aquaculture, 32:189-194. Davis, K.B. and Parker, N.C., 1986. Plasma corticosteroid stress response of fourteen species of warmwater fish to transportation. Transactions of the American Fisheries Society, 115: 495-499. Davis, K.B. and Parker, N.C., 1990. Physiological stress in striped bass: effect of acclimation temperature. Aquaculture, 91 349-358. Donaldson, E.M., 1981. The pituitary-interrenal axis as an indicator of stress in fish. Pages 11-47 in A.D. Pickering, editor. Stress in fish. Academic Press, London. Dupree, H.K. and Huner, J.V., 1984. Transportation of live fish. Pages 165-176 in H.K. Dupree and J.V. Huner, editors. Third report to the fish farmers, the status of warmwater fish fanning and progress in fish farming research. U.S. Department of the Interior, Fish and Wildlife Service, Washington, D.C. Eddy, F.B. and Bath, R.N., 1979. Ionic regulation in rainbow Irout (Salmo gairdneri) adapted to fresh water and dilute sea water. Journal of Experimental Biology, 83: 181-192. Eddy, F.B., 1981. Effects of stress on osmotic and ionic regulation in fish. Pages 77-102 in A.D. Pickering, editor. Stress in fish. Academic Press, London. Frain, W.J., 1987. The effect of external sodium and calcium concentrations on sodium fluxes by salt-depleted and non-depleted minnows, Phoxinus phoxinus (L.). Journal Experimental Zoology, 131: 417-425. Geiger, J.G. and Parker, N.C., 1985. Survey of striped bass hatchery management in the southeastern United States. The Progressive Fish-Culturist, 47: 1-13. Germann, J.F. and Reeves, W.C., 1975. The effects of added hardness, salinity, and source of fry on the survival and growth of striped bass fry in hatching jars. Proceedings of the Annual Conference Southeastern Association Game and Fish Commissioners, 28:199-208. Grizzle, J.M., Mauldin, II, A.C., Young, D. and Henderson, E., 1985. Survival of juvenile striped bass (Morone saxatilis) and Morone hybrid bass (Morone chrysops x Morone saxatilis) increased by addition of calcium to soft water. Aquaculture, 46: 167-171. Grizzle, J.M, Mauldin, II, A.C., Young, D. and Henderson, E., 1990. Effects of environmental calcium and sodium on postharvest survival of juvenile striped bass. Journal of Aquatic Animal Health, 2: 104-108. Grizzle, J.M., Mauldin, II, A.C. and Ashfield, C.J., 1992. Effects of sodium chloride and calcium chloride on survival of larval striped bass. Journal of Aquatic Animal Health, 4: 281-285. Grizzle, J.M., Cummins, K.A. and Ashfield, C.J., 1993. Effects of environmental concentrations of calcium and sodium on calcium flux in stressed 34-day-old striped bass. Canadian Journal of Zoology, 71:1379-1384. Grizzle, J. M. and Mauldin, II, A.C., 1994. Age-related changes in survival of larval and juvenile striped bass in different concentrations of calcium and sodium. Transactions of the American Fisheries Society, 123: 10021005. Harrell, R.M., 1992. Stress mitigation by use of salt and anesthetic for wild striped bass captured for brood stock. The Progressive Fish-Culturist, 54: 228-233.

213

Harrell, R.M. and Moline, M.A., 1992. Comparative stress dynamics of brood stock striped bass Morone saxatilis associated with two capture techniques. Journal of the World Aquaculture Society, 23: 58-63. Hazel, C.R., Thomsen, W. and Meith, S.J., 1971. Sensitivity of striped bass and stickleback to ammonia in relation to temperature and salinity. Califomia Fish and Game, 57: 154-161. Hunn, J.G., 1985. Role of calcium in gill function in freshwater fishes. Comparative Biochemistry and Physiology A, 82: 534-547. Johnson, S.K., 1979. Transport of live fish. Texas Agricultural Extension Service FDDL-F-14. Texas A&M University, College Station. Johnson, P. W. and Sieburth, J.M., 1974. Ammonia removal by selective ion exchange, a backup system for microbiological filters and closed system aquaculture. Aquaculture, 4:61-68. Kane, A.S., Bennett, R.O. and May, E.B., 1990. Effect of hardness and salinity on survival of striped bass larvae. North American Journal of Fisheries Management, 10:67-71. Lal, K., Lasker, R. and Kuljis, A., 1977. Acclimation and rearing of striped bass larvae in sea water. California Fish and Game, 63" 210-218. Lewis, S.D., 1971. The effect of salt solutions on osmotic changes associated with surface damage to the golden shiner, Notemigonus crysoleucas. Dissertation Abstracts Part B 31:6346. Marking, L.L. and Bills, T.D., 1982. Factors affecting the efficiency of clinoptilolite for removing ammonia from water. The Progressive Fish-Culturist, 44:187-189. Mauldin, II, A.C., Grizzle, J.M., Young, D.E. and Henderson, E., 1988. Use of additional calcium in soft-water ponds for improved striped bass survival. Proceedings of the Annual Conference Southeastern Association Fish Wildlife Agencies, 40:163-168. Mazeaud, M.M., Mazeaud, F. and Donaldson, E.H., 1977. Stress resulting from handling in fish: primary and secondary effects. Transactions of the American Fisheries Society, 106:201-212. Mazeaud, M.M. and Mazeaud, F., 1981. Adrenergic response to stress in fish. Pages 49-75 in A.D. Pickering, editor. Stress in fish. Academic Press, London. Mazik, P.M., Hinman, M.L., Winklemann, D.A., Klaine, S.J., Simco, B.A. and Parker, N.C., 1991a. Influence of nitrite and chloride concentrations on survival and hematological profiles of striped bass. Transactions of the American Fisheries Society, 120: 247-254. Mazik, P.M., Simco, B.A. and Parker, N.C., 1991b. Influence of water hardness and salts on survival and physiological characteristics of striped bass during and after transport. Transactions of the American Fisheries Society, 120: 121-126. McFarland, W.N., 1960. The use of anesthetics for the handling and the transport of fishes. California Fish and Game, 46(4): 407-431. McHugh, J.J. and Heidinger, R.C., 1978. Effect ofli~ht shock and handling shock on striped bass fry. The Progressive Fish-Culturist, 40: 82.

214

McWilliams, P.G., 1983. An investigation of the loss of bound calcium from the gills of the brown trout, Salmo trutta, in acid media. Comparative Biochemistry and Physiology A, 74: 107-116. McWilliams, P.G. and Ports, W.T.W., 1978. The effects of pH and calcium concentrations on gill potentials in the brown trout, Salmo trutta. Journal of Comparative Physiology B, 126: 277-286. Miles, H.M., Loehner, S.M., Michaud, D.T. and Salivar, S.L, 1974. Physiological responses of hatchery reared muskellunge (Esox masquinongy)to handling. Transactions of the American Fisheries Society, 103: 336-342. MilleR, K.M., Trial, J.G. and Stanley, J.G., 1983. Inexpensive monitor for aeration motors and transportation tanks. The Progressive Fish-Culturist, 45: 59-60. Murray-Brown, M.A., 1987. Effects of salinity, hardness and pH on the survival and growth of striped bass larvae, Morone saxatilis (Walbaum). Master's thesis. University of Rhode Island, Narragansett. Oduleye, S.O., 1975. The effects of calcium on water balance of the brown trout Salmo trutta. Journal of Experimental Biology, 63: 343-356. Ogasawara, T. and Hirano, T., 1984. Effects of prolactin and environmental calcium on osmotic water permeability of the gills in the eel, Anguillajaponica. General Comparative Endocrinology, 53:315-324. Parker, N.C., Klar, G.T., Smith, T.I.J. and Kerby, J.H., 1990. Special considerations in the culture of striped bass and striped bass hybrids. Pages 191-215 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Pic, P. and Maetz, J., 1981. Role of external calcium in sodium chloride transport in the gills of seawater-adapted Mugil capito. Journal of Comparative Physiology B, 141:511-521. Pickering, A.D., 1981. Introduction: the concept of biological stress. Pages 1-8 in A.D. Pickering, editor. Stress and fish. Academic Press, New York. Pickering, A.D., 1993. Endocrine-induced pathology in stressed salmonid fish. Fisheries Research, 17: 35-50. Piper, R.G., McElwain, I.B., Orme, L.E., McCraren, J.P. Fowler, L.G. and Leonard, J.R.. 1982. Fish hatchery management. U.S. Department of the Interior, Fish and Wildlife Service, Washington D.C. Ports, W.T.W. and Fleming, W.R., 1970. The effects of prolactin and divalent ions on the permeability to water to Fundulus kansae. Journal of Experimental Biology, 53:317-327. Ports, W.T.W., 1984. Transepithelial potentials in fish gills. Pages 105-128 in W.S. Hoar and D.J. Randall, editors. Fish physiology, volume 10B, Academic Press, London, Orlando. Rankin, J.C. and Bolis, L., 1984. Hormonal control of water movement across the gills. Pages 177-201 in W.S. Hoar and D.J. Randall, editors. Fish physiology. Volume 10B, Academic Press, London, Orlando. Redding, J.M. and Schreck, C.B., 1983. Influence of ambient salinity on osmoregulation and cortisol concentrations in yearling coho salmon during stress. Transactions of the American Fisheries Society, 112: 800-807.

215

Rees, R.A. and Harrell, R.M., 1990. Artificial spawning and fry production of striped bass and hybrids. Pages 43-72 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Robertson, L., Thomas, P. and Arnold, C.R., 1988. Plasma cortisol and secondary stress response of cultured red drum (Sciaenops ocellatus) to several transportation procedures. Aquaculture, 68:115-130. Seals, C., Kempton, C.J., Tomasso, J.R. and Smith, T.I.J., 1994. Environmental calcium does not affect production or selected blood characteristics of sunshine bass reared under normal culture conditions. The Progressive Fish-Culturist, 56: 269-272. Specker, J.L. and Schreck, C.B., 1980. Stress response to transportation and fitness for marine survival in coho salmon (Oncorhynchus kisutch) and steelhead trout (Salmo gairdneri). Canadian Journal of Fisheries and Aquatic Sciences, 37: 765-769. Strange, R.J., Schreck, C.B. and Ewing, R.D., 1978. Cortisol concentration in confined juvenile chinook salmon (Oncorhynchus tshawytscha). Transactions of the American Fisheries Society, 107:812-819. Strange, R.J., Schreck, C.B. and Golden, J.T., 1977. Corticoid stress responses to handling and temperature in salmonids. Transactions of the American Fisheries Society, 106:213-218. Tisa, M.S., Strange, R.J. and Peterson, D.C., 1983. Hematology of striped bass in fresh water. The Progressive FishCulturist, 45:41-44. Tomasso, J.R., Davis, K.B. and Parker, N.C., 1980. Plasma corticosteroid and electrolyte dynamics of hybrid striped bass (white bass x striped bass) during netting and hauling. Proceedings of the World Mariculture Society, 11: 303-310. Wagner, E.J., 1987. Stress in cultured fish. Aquaculture Magazine, 13: 24-29. Wattendorf. R.J. and Shafland, P.L., 1982. Observations on salinity tolerance of striped bass X white bass hybrids in aquaria. The Progressive Fish-Culturist, 44:148-149. Wedemeyer, G., 1970. The role of stress in the disease resistance of fishes. Pages 30-35 in S.F. Snieszko, editor. Diseases of fishes and shellfishes. American Fisheries Society, Bethesda, MD. Wedemeyer, G.A., Barton, B.A. and McLeay, D.J., 1990. Stress and acclimation. Pages 451-489 in C.B. Schreck and P.B Moyle, editors. Methods for fish biology. American Fisheries Society, Bethesda, MD. Weirich, C.R. and Tomasso, J.R., 1991. Confinement and transport-induced stress in red drum: effect of salinity. The Progressive Fish-Culturist, 53:59-61. Weirich, C.R., Tomasso, J.R. and Smith, T.I.J., 1992. Confinement and transport-induced stress in white bass (Morone chrysops) x striped bass (M. saxatilis) hybrids: effect of calcium and salinity. Joumal of the World Aquaculture Society, 23: 49-57. Weirich, C.R., Tomasso, J.R. and Smith, T.I.J., 1993. Toxicity of ammonia and nitrite to sunshine bass in selected environments. Journal of Aquatic Animal Health, 5" 64-72. Weiss, R.F., 1970. The solubility of nitrogen, oxygen and argon in water and seawater. Deep-Sea Research, 17: 721735.

216

Wendelaar Bonga, S.E. and Van der Meij, J.C.A., 1981. Effect of ambient osmolarity and calcium on prolactin cell activity and osmotic water permeability of the gills in the teleost Sarotherodon mossambicus. General Comparative Endocrinology, 43: 432-442. Wilson, A.J., 1950. Distribution units for warm water fish. The Progressive Fish-Culturist, 12:211-214. Wood, C.M., 1991. Branchial ion and acid-base transfer in freshwater teleost fish: environmental hyperoxia as a probe. Physiological Zoology, 64: 68-102. Wydoski, R.S. and Wedemeyer, G.A., 1976. Problems in the physiological monitoring of wild fish populations. Proceedings of The Annual Conference of the Western Association of Game and Fish Commissioners, 56: 200214. Yeager, D.M., Van Tassel, J.E. and Wooley, C.M., 1990. Collection, transportation, and handling of striped bass broodstock. Pages 29-42 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Young, P.S. and Cech, Jr, J.J., 1993. Physiological stress responses to serial sampling and confinement in young-ofthe-year striped bass, Morone saxatilis (Walbaum). Comparative Biochemistry and Physiology, 105A: 239244.

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

217

Chapter 8 Hybridization a n d Genetics Reginal M. Harrell 8.1 INTRODUCTION With respect to other aquaculture industries in the United States, striped bass and hybrid striped bass aquaculture is one of most rapidly expanding segments today. Yet, this industry is based principally on the propagation and rearing of interspecific F~ hybrids. Because interspecific hybridization exploits dominance genetic variance, it offers little chance of selective improvement short of domesticating and selecting parental lines and then hybridizing anew each generation (Hallerman, 1994). The success of these hybrids in the food fish and recreational fishing industry has been phenomenal, so it is surprising to find that compared to other established aquaculture species in the U.S., the amount of genetic information conceming Morone culture is quite small (Harrell, inpress). Other than hybrid performance information, most genetic information is limited to some of the more recent molecular genetic research regarding population identification (see below). There is little published information of cytogenetics, ploidy manipulation, domestication, cryopreservation, and genetic conservation. There is no information on quantitative and qualitative genetics. Currently there are several researchers and producers working in concert on the issues most germane to commercial production of striped bass and its hybrids. Most of these issues involve furthering domestication efforts concerning both striped bass and white bass (which can then be used for creating hybrids of selected lines), genetic manipulation and performance, strain evaluations, and heritability studies. Unfortunately, all of these efforts are in varying stages of evaluation and development and the results are not yet available. 8.2 HYBRIDIZATION Hybrid striped bass were first produced in 1965 in South Carolina by Robert Stevens at the Moncks Corner striped bass hatchery (Bishop, 1968). The original purpose of creating hybrid Morone was to produce a new recreational fish that had the size, longevity, food habits, and angling qualities of the striped bass and the adaptability of the white bass to varying environments (Bayless, 1972; Bonn et al., 1976). Interspecific hybrids within the Morone complex has been the standard production fish upon which the industry has developed since the early 1980s. Farmers were quick to recognize that heterosis exhibited by hybrid Morone directly affected economically important character traits, because these traits include being more robust, faster growing, and more resistant to disease and environmental extremes than pure striped bass (Logan 1968; Williams, 1971; Kerby, 1986; Harrell et al., 1990a; Figure 8.1). Taking advantage of market opportunities created by a declining natural coastal fishery for the striped bass in the late 1970s and early 1980s, the aquaculture industry began as pilot grow-out studies with hybrids in silos, inland ponds, and net-pens (Williams et al., 1981; Kerby et al., 1983a,b). By 1989, almost 275 MT were being produced in intensive tank systems using geothermal water in California (Van Olst and Carlberg, 1990). This production has led to an industry that today is approaching 6,000 MT annually (see Chapter 1). While this number is small compared to other aquaculture commodities in the U.S., it represents well over a 1,300% increase since the mid-1980s when production was less than 4.5 MT (Rhoades and Sheehan, 1991; Carlberg, 1995; Kahl, 1995; Carlberg, J.M., Kent Seafarms Corporation, San Diego, CA, personal communication; Table 1.2).

218

Fig. 8.1. Top: Comparison of palmetto bass (top) and white bass (bottom) and, Bottom: Morone hybrids (top to bottom, left to right) striped bass, palmetto bass x white bass backcross, sunshine bass, palmetto bass x striped bass backcross, palmetto bass, palmetto bass x palmetto bass F2, and palmetto bass x palmetto bass F2. (Photo by South Carolina Department of Natural Resources (top) and R.M. Harrell (bottom)).

219

8.2.1 Natural Hybridization Hybrids of the Morone complex are not sterile, and often are reproduced under captive conditions (Harrell, 1984a; Smith and Jenkins 1984; Kerby and Harrell, 1990; Woods et al., 1995b). Natural and introgressive hybridization within the wild populations of Morone also has been reported (Avise and Van Den Avyle, 1984; Todd, 1986; Crawford et al., 1987; Forshage et al., 1988; Harrell et al., 1993). In those situations where outcrossing or natural hybridization has occurred, most often it has been found in environments where artificially produced hybrids or congeneric fish have been intentionally stocked on top of naturally reproducing striped bass or white bass populations. Thus, it is important that before a domestication or captive breeding program using fish of wild origin is to be initiated, the genetic history of the founder populations should be thoroughly known and, if necessary, founder populations should undergo a genetic screening to detect purity (Harrell et al., 1993; Woods et al., 1995a). The concern ofintrogressive or natural hybridization with naturally reproducing populations of striped bass has been one of the factors limiting the expansion of open-water aquaculture for Morone in the U.S. (Harrell et al., 1990b; Harrell et al., 1993). One possible solution to this problem involves production of sterile fish either through triploidy induction or with administration of hormones (see below). I

8.2.2 Artificial Hybridization There are two commonly produced hybrids used in commercial aquaculture, palmetto bass and sunshine bass (Harrell et al., 1990a; see box). Although other Morone hybrids are produced, such as those between striped bass and white perch, known as Virginia or Maryland bass depending on which fish is the female, and yellow bass known as the paradise bass, the latter hybrid, along with palmetto bass and sunshine bass, are used most often for stocking natural bodies of water for recreational fishing purposes (Harrell et al., 1990a). The paradise bass is used almost exclusively in Louisiana.

Common Names of Morone Hybrids ( H a r r e i l et al., 1990, Robins et al., 1991) Common Name

Female

Male

Palmetto Bass ,,

Striped Bass

White Bass

Sunshine Bass

White Bass

Striped Bass

Virginia Bass

Striped Bass

White Perch

Maryland Bass

White Perch

Striped Bass

Paradise Bass . . . .

Striped Bass

Yellow Bass

There are a variety of publications espousing the production performance of the sunshine bass and palmetto bass under intensive culture conditions (e.g., Kerby et al., 1983a,b; Kerby et al., 1987; Smith et al., 1985; Smith and Jenkins, 1987a; Kelly and Kohler, 1996; Rudacille and Kohler, 1996), including several studies comparing F~s to pure striped bass or to F ~. With the exception of Rudacille and Kohler (1996), who compared the growth of white bass, sunshine bass, and palmetto bass in glass aquaria for eight weeks, none of the investigations compared the two hybrids created between white bass and striped bass under the same environmental conditions at the same time. In the case of Rudacille and Kohler (1996), the white bass and sunshine bass outperformed the palmetto bass, but their study was not done under what would be considered commercial environmental conditions. Thus, there is still no real consensus on whether the palmetto bass or the sunshine bass is the hybrid of choice for commercial culture. Reports from research and industry are too inconsistent regarding growth performance, survival, and uniformity to make an informed recommendation. Most recently, Wolters and DeMay (1996) reported a parallel culture trial wherein palmetto bass outperformed paradise bass in growth (0.94 g/d versus 0.59 g/d). The paradise bass had a higher survival (65% versus 44%). It is important to note however that larvae of white bass, white perch, and yellow bass have a much smaller mouth gape than striped bass and therefore require a much smaller feed supply during the first critical days of growth to insure survival (see Chapters 3 and 6), thereby making their early culture efforts more problematic.

220

While hybrids from striped bass and white bass are the hybrids of choice from the aquacultural and recreational fishing perspectives, from a practical approach there is a larger supply of sunshine bass fingerlings available. This larger supply is due in part to the fact that white bass are easier to hold, spawn, and produce in captivity compared to striped bass (Kohler et al., 1994; Smith et al., 1996; Chapter 6). Maturation size and age between the two species are major factors contributing to this proclivity toward producing sunshine bass because white bass mature between 250-580 g and two years of age, while striped bass typically mature at age 4 or older and 2-7 kg (Kohler et al., 1994). There are also some negative aspects of using hybrid bass as the industry standard. Hallerman (1994) listed some of the concerns from a genetic improvement perspective that should be considered before entering into a long-term breeding program for Morone. One of his first concerns deals with the temporal difference in spawning of the various Morone. For instance white bass and white perch normally spawn earlier than striped bass, and thus there may be a problem in obtaining ripe gametes to produce the hybrids. This problem can potentially be overcome with sperm cryopreservation or cold storage (see below). He also pointed out that hybrids are generally considered a terminal cross and that the genetics being exploited, dominance-based genetic variation, is at a maximum in the F, hybrid. 8.2.3 Backcrosses and F2s In addition to F~ hybrids, research has been undertaken to improve performance of hybrid striped bass by producing and evaluating second generation hybrids and backcrosses. Preliminary efforts with backcrosses (backcrossing a palmetto or sunshine bass to a striped bass to create a "75%" striped bass; Figure 8.2) has shown that the F, crossed with a pure species parent can create viable offspring, which have yielded promising results from an aquaculture perspective (Harrell and Dean, 1987, 1988; Kerby et al., 1987; Heyward et al., 1995). While there are other factors that influence heterotic expression, such as additive, maternal, epistatic, and egg cytoplasmic genetic effects along with maternal heterosis (Tave 1992), it has been demonstrated that F2s exhibit lower hatch, poorer larval viability, and slower growth than the F lhybrid (Bayless 1972; Harrell, 1984a; Smith and Jenkins, 1984; Smith et al. 1985). Figure 8.2 also demonstrates that as the hybrids age physical differences between the fish and pure striped bass is becomes more evident. 8.3 BROODSTOCK DOMESTICATION

The striped bass aquaculture industry is primarily dependent on capture of wild broodstock. As early as 1984, it became apparent that continued regulatory restrictions on capture and holding of wild broodstock was going to be problematic if wild fish were to be a continual source of gametes (Harrell, 1984b). Thus, dependence on wild broodstock is a limitation on further development of the industry (Smith, 1989). Controlled spawning of domesticated broodstock provides the best alternative for a dependable supply of seedstock (Smith and Jenkins, 1984; Woods et al., 1990). Several attempts have been made to establish captive populations from wild-caught broodstock, spawn them, and initiate development of domesticated stocks (Henderson-Arzapalo and Colura, 1987; Kerby, 1987; Smith, 1987; Smith and Jenkins, 1987b; Woods et al., 1990). To date, with the exception of the University of Maryland Crane Aquaculture Facility (CAF) broodstock (Woods et al., 1992, 1995a; Woods and Sullivan, 1993), there are only a few individuals in the industry or research communities that have active, successful broodstock development programs (Woods et al., 1995). The CAF broodstock is the only reported population where multiple F3 generations of domesticated striped bass are being held and where selection efforts have been initiated (L.C. Woods, III, University of

221

Fig. 8.2. Top: Comparison phase II palmetto bass (top), striped bass (middle), and palmetto bass x striped bass backcross (bottom). All fish are 80-100 mm TL. Bottom" Comparison of market-sized palmetto bass (top), palmetto bass x striped bass backcross (middle), and striped bass (bottom). (Photos by R.M. Harrell)

222

Maryland, personal communication). Other research facilities and various private aquaculture operations have initiated broodstock domestication programs with different striped bass populations but have not progressed as far as the C AF program. Efforts also have been initiated to domesticate white bass (Kohler et al., 1994; Smith et al. 1996; see Chapter 6). While only captive populations have been conditioned to spawn to date, the importance of white bass domestication is based on their use to create hybrids. Theoretically, using selected lines of both pure species parents to make interspecific F, hybrids should increase expression of hybrid vigor above that demonstrated in non-selected, domesticated F~ lines (Hallerman, 1994). Domestication is in itself then a de facto form of selection where those individuals who adapt to artificial feeds, handling, and crowding are the individuals chosen to be the progenitors of the next generation. It takes a few generations of this form of passive or unintentional selection to result in individual broodstock that are "conditioned" to hatchery environments and for that trait to be expressed in their progeny (Hallerman, 1994). It is well within the realm of possibility that selected lines of domesticated pure striped bass will outperform the current industry standard of non-selected F~ hybrids. The importance of domesticated lines has already been demonstrated with the U.S. catfish industry. For instance, Dunham and Smitherman (1983) demonstrated that domestication effects alone increased channel catfish (lctalu~. punctatus) growth rates an estimated 2-6% per generation. By combining domestication effects with selective breeding on specific traits of pure strain parents, hybrid yields from these crosses should express an even greater improvement in trait performance. As yet, however, no structured or intentional selection efforts have begun with those populations of domesticated striped bass. 8.4 STRAINS 8.4.1 Ancestry The striped bass is classified as an anadromous species that has a geographical range along the Atlantic coast from the St. Johns River, FL to the St. Lawrence River, Quebec (Raney, 1952). There also is evidence of a remnant native population located in the Gulf of Mexico and its drainages from northwest Florida to Louisiana (Wooley and Crateau, 1983), with the major concentration in the Apalachicola River system, FL (Wirgin et al., 1989). The recent utilization of DNA marker techniques for population identification obviated the more traditional, essentially ineffective tools such as scale counts, body measurements, tagging studies, and protein electrophoresis as means to separate different populations. These traditional tools indicated lack of genetic variability among striped bass populations, as they lacked the sensitivity to detect differences between fish of differing geographical origin (Waldman et al., 1988). In the mid- 1980s, scientists starting using mitochondrial DNA, DNA fingerprinting, RFLP analysis, hypervariable number of tandem repeats, and microsatellite DNA markers to determine if population structuring does indeed exist in striped bass (Waldman et al., 1988; Wirgin et al., 1989; Chapman, 1990; Wirgin and Maceda, 1991; Wirgin et al., 1991; Stellwag et al., 1994; Laughlin and Turner, 1996; Leclerc et al., 1996a). The application of the molecular techniques has progressed from .mitochondrial DNA (mtDNA) to microsatellites. In fact, recently the utility ofmtDNA, one of the earliest uses of molecular information as a means to identify genetic populations, has been questioned (Stellwag et al., 1994; Stellwag and Rulifson, 1995; Waldman and Wirgin, 1995). The most recent effort to determine population structure focuses on polymerase chain reaction (PCR) assays of genomic DNA (Laughlin and Turner, 1996; Leclerc et al., 1996a). Almost all of the molecular genetic studies have revealed some degree of structuring ranging from broad geographical inter-population variation to inter-individual variation within

223

populations. The question that remains however, is that if population segregation is only detectable at the nucleotide level, is it biologically significant from aquaculture or natural resource management perspectives. The answer is a matter of degree of the goals of your intended use of the species and of the information provided. 8.4.2 Strain Evaluation Putative population structuring that has been documented at the molecular genetic level could have an impact on striped bass aquaculture genetics because the industry is still dependent on wild fish for a source of broodstock. As mentioned, with the exception of the CAF (Woods et al., 1993), there are no reported facilities that have captive or domesticated striped bass broodstock subsequent to the second generation. Neither does there exist any hard evidence of which striped bass strain is best suited for commercial aquaculture under differing culture conditions. In research at the University of Maryland Horn Point Environmental Laboratory Aquaculture Facility, we are working with private industry and other academic researchers to evaluate which strain of striped bass will be best suited for aquaculture. Preliminary research with larval striped bass revealed that strains from the more northern latitudes along the Atlantic coast performed better in overall growth when compared to the more southern latitudes (Brown, 1994; Ehtisham et al., 1994; David Conover, SUNY, Stony Brook, personal communication). We are currently evaluating whether this observation holds true for growth from the fingerling stage to market size. Once we are able to delineate which strain performs best under differing culture conditions (i.e., closed-system versus ponds versus flow-through, etc.), future efforts placed into domestication should have a higher probability of success. It is interesting to note that the CAF domesticated broodstock are all of Chesapeake Bay, Maryland founder populations, and this may or may not be the best group with which to work. However, the research being conducted at that facility has demonstrated obvious benefits toward domestication in that it has documented excellent growth performance and has succeeded in producing females that mature in three years (L.C. Woods, III, personal communication). 8.5 QUALITATIVE AND QUANTITATIVE TRAITS There is no published information on the heritabilities of qualitative or quantitative genetic traits in striped bass. This is unfortunate because for effective selective breeding programs to develop, it is essential to know baseline information on the quantitative relationships of various commercially important traits, such as growth rate, disease resistance, and dress-out percentage (Hallerman, 1994). Even more unfortunate, because striped bass have a relatively long generation time (four to six years), it will be at least several more years before the necessary baseline information is available to determine which traits can be exploited. Although it may seem more mundane than molecular genetic research into areas such as population structuring and gene mapping, it is essential that concomitant to ongoing domestication research and development, parallel and collaborative efforts be directed at determining the genetic basis for the more economically important traits of striped bass. Hallerman (1994) called for coordination of efforts of all interested parties to work together toward achieving long-term support of breeding and genetic analysis programs. 8.6 CYTOGENETICS AND GENETIC MANIPULATIONS Besides hybridization and population genetics research, most Morone genetics research has revolved around genetic manipulation, including ploidy manipulation, gynogenesis, sex-reversal, and sperm

224

cryopreservation (Kerby, 1983, 1988; Curtis et al., 1988; Kerby et al. 1985, 1989; Kerby and Harrell, 1990; Leclerc et al., 1996b; Harrell et al., 1996). 8.6.1 Karyology Some of the earliest genetics research with striped bass was on karyotyping the species (Kerby, 1972; Rachlin et al., 1978). Striped bass have 48 chromosomes. Kerby (1972) reported that all but two are acrocentric with the other two being metacentric. In two separate chromosome spreads, Rachlin et al. (1978) did a more detailed characterization, describing 38 acrocentric, 8 subtelocentric, and 2 submetacentric in one spread, and 40 acrocentric, 6 subtelocentric, and 2 submetacentric in the other spread. Recently, Wolters and DeMay (1996) reported in a study comparing growth and survival of sunshine bass to paradise bass that all the paradise bass were all female (the sunshine bass did not significantly deviate from a 1:1 sex ratio). This finding of 100 percent females caused the authors to speculate regarding the sex determining mechanism of Morone. Before their findings, it had always been assumed that sex determination in Morone, at least in striped bass and white bass, was of the XY mechanism, where the male was the heterogametic sex. They postulated that a three gonosome sex determining mechanism may be at work, as proposed for tilapia by Avtalion (1982). Under this sex determining mechanism, striped bass females would be AAXXand white bass males would be AAXY, while yellow bass males (the male crossed with striped bass to create the paradise bass) would be.4,4 WW. Thus a cross between striped bass females and white bass males would have a 1:1 sex ratio (AAXX females and A A X Y males) and a cross between striped bass females and yellow bass males would be 1:0 (all AAXW). While this sex determining mechanism is a possibility, further analysis needs to be undertaken before it is generally accepted. The authors also suggested other possible explanations for the observed skewed sex ratio. 8.6.2 Polyploidization Ploidy manipulation is well recognized as a potential means for improving growth rate in fish species. Triploidy induction (producing fish with three sets of chromosomes) has shown promise for increasing the culture potential of Morone, and in particular hybrid Morone. This increased potential is directly related to the observation that after five year's growth, triploid hybrids were functionally sterile (Harrell et al., 1996). The sterility is putatively due to there being an odd set of chromosomes in tfiploids, with mispairing causing a breakdown in meiotic divisions within the gonads -- rendering the gonads nonfunctional (Figure 8.3). Since triploid hybrid Morone have been demonstrated to be sterile, the concern of resource agencies about introgressive hybridization between the hybrids and naturally reproducing striped or white bass (Harrell, 1984a; Kerby and HarreU, 1990; Harrell et al., 1993) has been essentially eliminated. Thus, triploids are useful for removing one of the major impediments to open water aquaculture or to land-based hybrid Morone culture in areas where there are also populations of naturally reproducing bass. The benefit of reproductive sterility can be extended to non-indigenous strains of striped bass where the agency is concerned about genetic integrity of native populations. Triploid Morone have been created by both heat and pressure shock of recently-fertilized eggs (Kerby 1987; Kerby and Harrell, 1990) wherein the second polar body of the egg is retained after the second meiotic division. Unfortunately, neither technique is 100% effective and requires fish be individually tested if pure populations of triploids are required. Because induction techniques are not yet perfected, large-scale production for industry or management purposes is currently problematic (Hallerman, 1994).

225

Fig. 8.3. Top: Reproductively viable ovaries of a gravid palmetto bass. Bottom: Vestigial gonadal tissue of a triploid sunshine bass. (Photo by, Top: J.H. Kerby, Bottom: R.M. Harrell).

226

An altemative method of producing large numbers oftriploids is to cross diploids with tetraploids (fish with four sets of chromosomes). To date, there is only one report where tetraploid Morone have been produced (Curtis et al., 1988), and while the crossing of diploids to tetraploids may relieve some of the technical problems of creating pure populations of triploids by inducing second polar body retention, there is still the requirement of creating the tetraploids. Tetraploids are created by suppressing the first mitotic cell division, thereby creating four instead of two sets of chromosomes within the individual egg. Future mitotic cell divisions will, therefore, replicate these additional sets of chromosomes, and each new daughter cell will have four instead of the usual two sets of chromosomes. This cell division suppression can be performed either by temperature, hydrostatic pressure, or chemical treatment. Regarding triploid performance under culture conditions, to date, the only production research undertaken has been with sunshine bass hybrids, which resulted in the diploids out-growing the triploids. The triploids had a higher survival, but were significantly smaller than the diploids at harvest, albeit both reached market size (i.e., 700g) in 18 mo (J.H. Kerby, National Biological Service, Leetown, WV and R.M. Harrell, unpublished data). It was hypothesized that the maternal effect resulting from two-thirds of the genome being of white bass origin may have caused the slower growth exhibited in the triploids. White bass are significantly smaller at a given age than striped bass (see Chapter 6). 8.6.3 Gynogenesis Production of gynogenetic striped bass (i.e., destroying paternal DNA and suppressing the second meiotic division, wherein the second polar body is retained producing meiotic gynogens; or suppressing first mitotic division creating a mitotic gynogen) has tremendous potential in population restoration and potential enhancement efforts by increasing reproductive potential (Harrell, inpress). By using all-female populations as 50% of the fish stocked, the ratio of females to males can be increased from 1:1 to 3:1 (H.A. Kincaid, National Biological Service, Wellsboro, PA, personal communication). From a commercial aquaculture perspective, total productivity should be higher when gynogenetic fish are reared because both hybrid Morone and pure striped bass exhibit sexual dimorphism with the female being the larger sex. However in the first generation at least, there may be a higher percentage of deleterious recessive alleles being expressed resulting in poorer hatch or overall survival, especially in the mitotic gynogens. To date, production ofgynogenetic striped bass or its hybrids has been limited to a research scale only (Leclerc et al., 1996b). Meiotic gynogenetic hybrids have been produced successfully using UV-irradiated white perch sperm as a genetic marker to detect paternal contribution and hydrostatic pressure as a means to retain the second polar body (Leclerc et al., 1996b). The success of this study showed that 100% production of gynogens could be achieved. Through the use of molecular markers and PCR analyses, it was possible to determine if the fish were indeed gynogens in the larval stage, circumventing the need to wait until the fish were mature to determine if the treatment was successful. An advantage of using F~ hybrid striped bass to produce gynogenetic fish is that they contain one chromosome of each pair from each species. Thus, the progeny should have a random assortment of striped bass and white bass chromosomes. These chromosomes should contain many genetic differences (being from different species), making them useful for the development of linkage and genetic marker maps (Leclerc et al., 1996b). Linkage analyses would be simplified because all recombinants are heterozygous for the allele in question, while non-recombinants would be homozygous -- thereby immediately providing a screening technique to eliminate the recombinants from the analyses (Leclerc et al., 1996b). If recombinant alleles can be identified and removed from the analysis, linkage results in 100% co-segregation of the two alleles, while non-linkage will result in equal ratios of the four possible genotypes (Leclerc et al., 1996b).

227

8.6.4 Sex Reversal In addition to producing gynogenetic females through genetic manipulation, all-female (or all-male) populations can be produced through administration of hormones to allow a fish to phenotypically express the sex not normally associated with its genetic makeup (i.e., XY(males orXYfemales, assuming sex determination is indeed by the XYmechanism). By first using sex-reversal techniques to produce XX-males and then mating these males to normal females, an all-female progeny distribution can be produced without consumers having to eat hormone-treated individuals. Efforts thus far to effect hormonal control of sex in striped bass have been unsuccessful due to progeny mortality, delayed maturation, or sterility (Harrell et al., 1996). This work is continuing in the University of Maryland Horn Point Environmental Laboratory Aquaculture Facility. 8.6.5 Cryopreservation Preservation of fish gametes has been of interest to fisheries scientists for well over 100 years (Kerby 1983). While there are a variety of reasons to undertake long-term gamete storage efforts, two of the major purposes are for gene banking for genetic conservation and for having gametes available in sufficient quantities for hatchery production at atime when one of the sexes is in short supply. For instance, during the annual striped bass production cycle with wild populations, the males are usually found on the spawning grounds well in advance of the females and are usually scarce toward the latter part of the spawning run (Bayless, 1972). In this situation, it would be advantageous to have cryopreserved eggs during the first part of the season and cryopreserved sperm during the last portion. Likewise, cryopreservation is an excellent tool to assist in congeneric hybridization where there exists a temporal spawning isolation between species, such as striped bass and white bass. It also could be used to create conspecific hybrids among populations that are separated by latitudinal differences, e.g., where strains from Canada spawn later than strains from Florida. To date, no one has been successful in effecting long-term cryopreservation of Morone eggs, and only a few efforts have been successful in sperm preservation. Howard Kerby of the National Biological Service published a series of papers in the 1980s on striped bass sperm cryopreservation and performance success of progeny produced by cryopreserved sperm (Kerby, 1983, 1984; Kerby et al., 1985; Parker, et al., 1990). Common to the success ofcryopreservation protocols is the use ofextenders and cryoprotectants. An extender is usually a solution of salts, and possibly organic compounds, that help maintain cell viability during cryopreservation, while cryoprotectants are organic compounds that protect the cells during the freezing and thawing process (Kerby, 1983). The most successful extender was OH-189, which was taken from a formula developed by Ott (1975) for salmonid sperm (Table 8.1). The most successful cryoprotectant was dimethyl sulfoxide at a concentration of 5% (Kerby 1983). The freezing medium most successful was a ratio of 1:4 sperm to medium (v:v) and frozen in 1 mL aliquots in a 2 mL A/S NLrNC polypropylene cryotube (Union Carbide, Inc.). Kerby mentioned in his seminal work in 1983 on striped bass sperm cryopreservation that some of the biggest hurdles to be overcome were controlling the freezing rate, variation in gamete quality among animals, and control of the thawing process. If the freezing rate was not correct, the cell could rupture due to formation of ice crystals within the cell structure. Mean freezing rates between 5 ~ and 20 ~ were more effective than slower rates. Recently, the work on Morone sperm cryopreservation by Kerby has been continued by George Brown of Iowa State University, Ames, Robert Sheehan of Southern Illinois University, Carbondale, and Terry Tiersch of Louisiana State University, Baton Rouge (personal communications), which includes efforts on

228

short-term cold (4 ~ storage. While none of this research has been published, using their protocols in field trials at the Horn Point Environmental Laboratory Aquaculture Facility, we obtained excellent results with the short-term cold storage extender recommended by Dr. Brown (Table 8.1). We observed sperm viability and effected fertilization two weeks after the sperm was first stored, which was critical to a diallele heritability analysis of swim bladder inflation (HarreU, unpublished data). Interested hatchery managers should contact Dr. Brown for the details. In a progeny performance experiment with cryopreserved sperm, Kerby et al. (1995) reported that in two pond experiments there were no significant differences in growth and survival between progeny stocked with larvae produced with fresh or cryopreserved sperm to the phase I fingerling stage. Table 8.1. Chemical composition of extenders used in cold-storage (# 13) and cryopreservation of striped bass sperm. Values presented as ~ams of solute per liter total solution. ExtenderOH- 189 taken from Parker et al. (1990) and # 13 from George Bro.wn., Iowa State Universit),, personal communication: Chemical

OH- 189

# 13

NaCI

7.30

8.60

KCI

0.38

CaC12o2H20

0.23

NaHCO3

5.00

NaH2PO4oH_,O

0.41

MgSO4o7H20

0.23

Fructose

5.00

Lecithin

7.50

Mannitol

5.00

Dimethyl Sulfoxide

5%

pen-strep *

10 mL

pH 7.60 ~penicillin-streptomycin: 5000 units of penicillin, 5 mg streptomycin per mL of 0.9% NaCI

8.6.6 Genetic Engineering In a report on the application of tmnsgenic fish technology in aquaculture, Chen et al. (1991) alluded to success in aquaculture being dependent on breeding, feeding, disease protection, and management. It is their opinion that molecular biology and biotechnology can influence the results of the first three areas. This enhancement can be in the form of improving growth rates, controlling reproductive cycles, improving nutritional components, and developing vaccines and disease resistant populations (Chen et al., 1991). Growth rate enhancement and controlled reproduction have received the most attention. Information regarding controlled reproduction can be found in Chapter 2.

229

To date, use of introduced growth promoting genes has been performed with striped bass only under carefully controlled laboratory conditions (T. Chen, University of Connecticut, personal communication). Successful transfer and expression of such growth factor genes is, therefore, technically feasible and could contribute significantly to the future success of the aquaculture industry (Hallerman, 1994). Once developed and the expression is quantified in first generation transgenic individuals, it will be necessary to rear them to maturity, spawn them, and rear their transgenic off spring in performance trials to identify those individuals that express the favorable new genotype (Hallerman, 1994). These selected individuals could then serve as the founder population for future production (Kapuscinski and Hallerman, 1990). Both performance enhancement and environmental safety will have to be demonstrated before transgenics will be attractive for routine aquaculture production (Kapuscinski and Hallerman, 1990). 8.7 CONSERVATION GENETICS What is clear from the information about Morone genetics is that we are still in the early stages of learning about the genetics of striped bass and its hybrids. While considerable effort is being placed into this discipline, it will take several years before most results are of practical use. In the meantime, however, the industry will continue to depend essentially on wild broodstock as a seed supply source while improving husbandry techniques, and adapting management with the best information they have available to them. Associated with our advancement of knowledge about the genetics of this group of fish are the facts that native populations can be stressed to the point of nearing extinction and that introgressive hybridization can be a reality. As responsible individuals, we must insure that efforts to develop and expand a food fish industry as well as our zeal to meet the demands of a recreational fishing public do not negatively impact native species or their habitat. It is incumbent upon hatchery managers and farm owners alike to remain within the local jurisdictional laws and regulations governing the operation of either a private or public aquaculture facility. The issue of conserving Morone genetic resources has been amply covered in recent publications (Kerby and Harrell, 1990; Harrell et al., 1993; Hallerman, 1994) and the issues are not discussed in depth here. While it is true that man has served as a conduit for moving Morone well outside of its natural and historical range, many of the efforts to date have been undertaken with F1 progeny. Yet, as we learn more about population structuring and advance our efforts at domestication and selective breeding, thereby further removing the product from a "wild" fish, we must be responsible in protecting our natural resource. After all, it is from this wild resource that the founder populations of all the future selected populations will be developed. Thus, it would be prudent to maintain that original diversity not only for its own sake, but also as a reserve to meet any need that requires supplementing a given population's diversity, be it in a private hatchery or pubic ecosystem.

230

References

Avtalion, R.R., 1982. Genetic markers in Sartherodon and their use for sex and species identification. Pages 269-277 in R.S.V. Pullin and R.H.L. McConnell, editors. The biology and culture of tilapias. ICLARM, MCC, Phillippines. Avise, J.C. and Van Den Avyle, M.J., 1984. Genetic analysis of reproduction of hybrid white bass X striped bass in the Savannah River. Transactions of the American Fisheries Society, 113: 563-570. Bayless, J.D., 1972. Artificial propagation and hybridization of striped bass, Morone saxatilis (Walbaum). South Carolina Wildlife and Marine Resources Department, Columbia. Bishop, R.D., 1968. Evaluation of the striped bass (Roccus saxatilis) and white bass (R. chrysops) hybrids after two years. Proceedings of the Annual Conference Southeastern Association of Game and Fish Commissioners, 21: 245-254. Bonn, E.W., Bailey, W.M., Bayless, J.D., Erickson, K.E. and Stevens, R.E., editors., 1976. Guidelines for striped bass culture. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Brown, J.J., 1994. Variation in growth rate among larval striped bass Morone saxatilis stocks from different latitudes. M.S. Thesis, Marine Environmental Sciences Department, State University of New York, Stony Brook. Carlberg, J.M., 1995. Foodfish production: commercial flow-though tank grow-out systems. Aquaculture '95. Annual Meeting of the World Aquaculture Society, Abstract. Chapman, R.W., 1990. Mitochondrial DNA analysis of striped bass populations in Chesapeake Bay. Copeia, 1990: 355-366. Chen, T.T., Lin, C.M. and Kio~at, K., 1991. Application of transgenic fish technology in aquaculture. Bulletin of the Institute of Zoology, Academia Sinica, 16: 375-386. Crawford, T., Freeze, M., Fourt, R., Henderson, S., O'Bryan, G. and Phillipp, D., 1987. Suspected natural hybridization of striped bass in two Arkansas reservoirs. Proceedings of the Annual Conference Southeastern Association ofFish and Wildlife Agencies, 38: 455-469. Curtis, T.A., Sessions, F.W., Bury, D., Resk, M. and Dunham, R.A., 1988. Induction ofpolyploidy with hydrostatic pressure in striped bass, white bass, and their hybrids. Proceedings of the Annual Conference Southeastern Association of Fish and Wildlife Agencies, 41: 63-69. Dunham, R.A. and Smitherman, R.O., 1983. Response to selection and realized heritability for body weight in three strains of channel catfish, lctalurus punctatus, grown in earthen ponds. Aquaculture, 33: 89-96. Ehtisham, A., Brown, J.J. and Conover, D., 1994. Evidence for countergradient variation in growth rate of larval and juvenile striped bass, or why striped bass growers m i ~ t want to select broodstock fish from New York and not South Carolina. World Aquaculture '94. Annual Meeting of the World Aquaculture Society. Abstract. Forshage, A.A., Harvey, W.D., Kulzer, K.E. and Fries, L.T., 1988. Natural reproduction of white bass x striped bass hybrids in a Texas reservoir. Proceedings of the Annual Conference Southeastern Association of Fish and Wildlife Agencies, 40: 9-14. Hallerman, E.M., 1994. Toward coordination and funding of long-term genetic improvement programs for striped and hybrid bass Morone sp. Journal of the World Aquaculture Society, 25: 360-365.

231

Harrell, R.M., 1984a. Tank spawning of first generation striped bass x white bass hybrids. The Progressive FishCulturist, 46: 75-78. Harrell, R.M., 1984b. Review of striped bass brood stock acquisition, spawning methods, and fry production. Pages 45-57 in J.P. McCraren, editor. The aquaculture of striped bass: a proceedings. University of Maryland Sea Grant College publication UM-SG-MAP-84-01, College Park. Harrell, R.M., In press. Genetics of striped bass and other Morone. World Aquaculture. Harrell, R.M. and Dean, J.M., 1987. Pterygiophore interdigitation patterns and morphometry of larval hybrids of the Morone species. Transactions of the American Fisheries Society, 116:719-727. Harrell, R.M. and Dean, J.M., 1988. Identification of juvenile hybrids of Morone based on meristics and morphometrics. Transactions of the American Fisheries Society, 117: 529-535. Harrell, R.M., Geiger, J.A., Kerby, J.H. and Kincaid, H.A., 1996. Genetic manipulation in striped bass, Morone saxatilis. Aquaculture America '96. Annual meeting of the U.S. Chapter of the World Aquaculture Society, Abstract. Harrell, R.M., Kerby, J.H. and Minton, R.V., editors., 1990a. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Harrell, R.M., Kerby, J.H., Smith, T.I.J. and Stevens, R.E., 1990b. Striped bass and striped bass hybrid culture: the next twenty-five years. Pages 253-261 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Harrell, R.M., Xu, X.L. and Ely, B., 1993. Evidence of introgressive hybridization in Chesapeake Bay Morone. Molecular Marine Biology and Biotechnology, 2(5): 291-299. Henderson-Arzapalo, A. and Colura, R.L., 1987. Laboratory maturation and induced spawning of striped bass. The Progressive Fish-Culturist, 49: 60-63. Heyward, L.M., Smith, T.I.J. and Jenkins, W.E., 1995. Performance of striped bass (Morone saxatilis), palmetto bass and backcross hybrids (sunshine bass ~ X M. saxatilis c~) reared in a tank culture system. Aquaculture 95, Annual Meeting of the World Aquaculture Society, Abstract. Kahl, K.H., 1995. Results of the hybrid striped bass producer and state aquaculture coordinator surveys. Report to the South Carolina Department of Agriculture, Columbia. Kapuscinski, A.R. and Hallerman, E.M., 1990. Transgenic fish and public policy. I. Anticipating environmental impacts of transgenic fish. Fisheries, 15(1): 2-11. Kelly, A.M. and Kohler, C.C., 1996. Sunshine bass performance in ponds, cages, and indoor tanks. The Progressive Fish-Culturist, 58: 55-58. Kerby, J.H., 1972. Feasibility of artificial propagation and introduction of hybrids of the Morone complex into estuarine environments, with a meristic and morphometric description of the hybrids. Ph.D. Dissertation. University of Virginia, Charlottesville. Kerby, J.H., 1983. Cryogenic preservation of sperm from striped bass. Transactions of the American Fisheries Society, 112: 86-94.

232

Kerby, J.H., 1984. Cryopreservation of striped bass spermatozoa: problems and progress. Pages 205-213/n J.P. McCraren, editor. The aquaculture of striped bass: a proceedings. Maryland Sea Grant Publication UM-SGMAP-84-01. University of Maryland, College Park. Kerby, J.H., 1986. Striped bass and striped bass hybrids. Pages 127-147 in R.R. Stickney, editor. Culture of nonsalmonid freshwater fishes. CRC Press, Boca Raton, FL. Kerby, J.H., 1987. Genetic manipulation in Morone. Pages 61-71/n R. Hodson, T. Smith, J. McVey, R. Harrel (sic), and N. Davis, editors. Hybrid striped bass culture: status and perspective. University of North Carolina Sea Grant Publication, UNC-5-87-03. North Carolina State University, Raleigh. Kerby, J.H., 1988. Induction of triploidy in hybrid sniped bass using hydrostatic pressure. Journal of the World Aquaculture Society, 19:41A. Abstract Kerby, J.H., Bayless, J.D. and Harrell, R.M., 1985. Growth, survival, and harvest of striped bass produced with cryopreserved spermatozoa. Transactions of the American Fisheries Society, 114:761-765. Kerby, J.H. and Harrell, R.M., 1990. Hybridization, genetic manipulation, and gene pool conservation of striped bass. Pages 159-190, in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and is hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Kerby, J.H., Hinshaw, J.M., and Huish, M.T., 1989. Induction of triploidy in hybrid sniped bass using thermal shock and comparative growth between diploid and triploid offspring. Journal of the World Aquaculture Society, 20: 47A. Abstract. Kerby, J.H., Huish, M.T., Klar, G.T. and Parker, N.C., 1987. Comparative growth and survival of two striped bass hybrids, a backcross, and striped bass in earthen ponds. Journal of the World Aquaculture Society, 20: 47A Abstract. Kerby, J.H., Woods, III, L.C. and Huish, M.T., 1983a. Culture of the sniped bass and its hybrids: a review of methods, advances and problems. Pages 23-54 in R.R. Stickney and S.P. Meyers, editors. Proceedings of the warmwater fish culture workshop. World Mariculture Society Special Publication 3. Louisiana State University, Baton Rouge. Kerby, J.H., Woods, III, L.C. and Huish, M.T., 1983b. Pond culture of striped bass x white bass hybrids. Journal of the World Mariculture Society, 14:613-623. Kohler, C.C., Sheehan, R.J., Habicht, C., Malison, J.A. and Kayes, T.B., 1994. Habituation to captivity and controlled spawning of white bass. Transactions of the American Fisheries Society, 123: 964-974. Laughlin, T.F. and Turner, B.J., 1996. Hypervariable DNA markers reveal high genetic variability within sniped bass populations in the lower Chesapeake Bay. Transactions of the American Fisheries Society, 125: 49-55. Leclerc, G., Diaz, M. and Ely, B., 1996a. Use of PCR-RFLP assays to detect genetic variation at single-copy nuclear loci in striped bass (Morone saxatilis). Molecular Marine Biology and Biotechnology, 5: 138-144. Leclerc, G., Ely, B., Xu, X.L. and Harrell, R.M., 1996b. Gynogen production in hybrid striped bass. Journal of the World Aquaculture Society, 27:119-125 Logan, H.J., 1968. Comparison of growth and survival rates of striped bass and striped bass x white bass hybrids under controlled environments. Proceedings of the Annual Conference Southeastern Association of Game and Fish Commissioners, 21: 260-263.

233

Ott, A.G., 1975. Cryopreservation of Pacific salmon and steelhead trout sperm. Doctoral dissertation, Oregon State University, Corvallis. Parker, N.C., Klar, G.T., Smith, T.I.J. and Kerby, J.H., 1990. Special considerations in the culture of striped bass and striped bass hybrids. Pages 191-215 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Rachlin, J.W., Beck, A.P. and O'Connor, J.M., 1978. Karyotypic analysis of the Hudson River striped bass, Morone saxatilis. Copeia,1978: 343-345. Raney, E.C., 1952. The life history of the striped bass, Roccus saxatilis (Walbaum). Bulletin of the Bingham Oceanographic Collection, Yale University, 14: 5-97. Rhodes, R.J. and Sheehan, B., 1991. Estimated annual production of commercial hybrid striped bass growers in the United States. Striped Bass Grower's Association Report, Raleigh, NC. Robins, C.R., Bailey, R.M., Bond, C.E., Brooker, J.R., Lachner, E.A., Lea, R.N. and Scott, W.B., 1991. Common and scientific names of fishes from the United States and Canada, Fifth Edition. American Fisheries Society Special Publication 20. Bethesda, MD. Rudacille, J.B., and Kohler, C.C., 1996. Relative performance of white bass, sunshine bass, and palmetto bass fed a commercial diet. Aquaculture America 1996, Annual Meeting of the U.S. Chapter of the World Aquaculture Society, 1996:130 Abstract. Smith, T.I.J., 1987. Hatchery. Pages 17-22 in R. Hodson, T. Smith, J. McVey, R. Harrel (sic), and N. Davis, editors. Hybrid striped bass culture: status and perspective. University of North Carolina Sea Grant Publication, UNC5-87-03. North Carolina State University, Raleigh. Smith, T.I.J., 1989. The culture potential of striped bass and its hybrids. World Aquaculture, 20: 32-38. Smith, T.I.J. and Jenkins, W.E., 1984. Controlled spawning of F1 hybrid striped bass (Morone saxatilis x Morone chrysops) and rearing ofF2 progeny. Journal of the World Mariculture Society, 15: 147-161. Smith, T.I.J. and Jenkins, W.E., 1987a. Aquaculture research with striped bass and its hybrids in South Carolina. Proceedings of the Annual Conference Southeastern Association of Fish and Wildlife Agencies, 39:217-227. Smith, T.I.J. and Jenkins, W.E., 1987b. Broodstock development and spawning of striped bass, Morone saxatilis, and white bass, M. chrysops. Journal of the World Aquaculture Society, 19: 65A. Abstract. Smith, T.I.J., Jenkins, W.E. and Heyward, L.D., 1996. Production and extended spawning of cultured white bass broodstock. The Progressive Fish-Culturist, 58:85-91. Smith, T.I.J., Jenkins, W.E. and Snevel, J.F., 1985. Production characteristics of striped bass (Morone saxatilis) and F~, F2, hybrids (M. saxatilis and M. chrysops) reared in intensive tank systems. Journal of the World Mariculture Society, 16: 57-70. Stellwag, E.J., Payne, E.S. and Rulifson, R.A., 1994. Mitochondrial DNA diversity of Roanoke River striped bass. Transactions of the American Fisheries Society, 123:321-334. Stellwag, E.J. and Rulifson, R.A., 1995. Mitochondrial DNA stability and striped bass stock identification: Response to comment. Transactions of the American Fisheries Society, 124: 956-959.

234

Tave, D., 1992. Genetics for fish hatchery managers, second edition. Van Nostrand Reinhold, New York. Todd, T.N., 1986. Occurrence of white bass-white perch hybrids in Lake Erie. Copeia, 1986: 196-199. Van Olst, J.C. and J.M. Carlberg. 1990. Commercial culture of hybrid striped bass: status and potential. Aquaculture Magazine, 16(1): 49-59. Waldman, J.R., Grossfield, J. and Wirgin, I., 1988. Review of stock discrimination techniques for striped bass. North American Journal of Fisheries Management, 8:410-425. Waldman, J.R. and Wirgin, I.I., 1995. Comment: Mitochondrial DNA stability and striped bass stock identification. Transactions of the American Fisheries Society, 124: 954-956. Williams, H.M., 1971. Preliminary studies of certain aspects of the life history of the hybrid (striped bass x white bass) in two South Carolina reservoirs. Proceedings of the Annual Conference Southeastern Association of Game and Fish Commissioners, 24:424-431. Williams, J.E., Sandifer, P.A. and Lindberg, J.M., 1981. Net-pen culture of striped bass x white bass hybrids in estuarine waters of South Carolina: a pilot study. Journal of the World Mariculture Society, 12 (2): 98-110. Wirgin, I.I., Grunwald, C. Garte, S.J. and Mesing, C., 1991. Use of DNA fingerprinting in the identification and management of a striped bass population in the southeastern United States. Transactions of the American Fisheries Society, 120: 273-282. Wirgin, I.I. and Maceda, L., 1991. Development and use of striped bass-specific RFLP probes. Journal of Fish Biology, 39(Supplement A): 159-167. Wirgin, I.I., Proenca, R., Grossfield, J. 1989. Mitochondrial DNA diversity among populations of striped bass in the southeastern United States. Canadian Journal of Zoology, 67:891-907. Wolters, W.R. and DeMay, R., 1996. Production characteristics of striped bass x white bass and striped bass x yellow bass hybrids. Journal of the World Aquaculture Society, 27: 202-207. Woods, III, L.C., Woiwode, J.G., McCarthy, M.A., Theisen, D. and Bennett, R.O., 1990. Noninduced spawning of captive striped bass in tanks. The Progressive Fish-Culturist, 52:201-202. Woods, III, L.C., Bennett, R.O. and Sullivan, C.V., 1992. Reproduction of a domestic striped bass brood stock. The Progressive Fish-Culturist, 54:184-188. Woods, III, L.C., Ely, B., Leclerc, G. and Harrell, R.M., 1995a. DNA evidence for genetic purity of captive and domestic striped bass broodstocks. Aquaculture, 137: 41-44. Woods, III, L.C., Kohler, C.C., Sheehan, R.J. and Sullivan, C.V., 1995b. Volitional tank spawning of female striped bass with male white bass produces hybrid offspring. Transactions of the American Fisheries Society, 124: 628632. Woods, III, L.C. and Sullivan, C.J., 1993. Reproduction of striped bass, Morone saxatilis (Walbaum), broodstock: Monitoring maturation and hormonal induction of spawning. Journal of Aquaculture and Fisheries Management, 24:211-222. Wooley, C.M. and Crateau, E.J., 1983. Biology, population estimates, and movement of native and introduced striped bass, Apalachicola River, Florida. North American Journal of Fisheries Management, 3: 383-394.

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

235 Chapter 9

Nutrition and Feeding of Striped Bass and Hybrid Striped Bass Delbert M. Gatlin, III 9.1 INTRODUCTION 9.1.1 Feeding Habits Striped bass and hybrid striped bass exhibit carnivorous feeding behavior throughout their life stages in nature. Prolarvae use endogenous nutrients from the yolk sac for approximately four to five days after hatching as they undergo metamorphosis and digestive tract development. At the beginning of exogenous feeding, a variety of zooplankton species typically comprise the major food items of larval striped and hybrid bass. In freshwater, cladocerans and cyclopoid copepods are considered to be the most important food of fish less than 30 mm long (Regan et al., 1968; Harper et al., 1969; Harrell et al., 1977; Woods et al., 1985), while in brackish water larvae feed principally on calanoid and harpacticoid copepods (Harrell and Bukowski, 1990). As fish length increases, cladocerans and copepods, as well as insects and mysid shrimp typically constitute major food items (Heubach et al., 1963; Regan et al., 1968; Harper et al., 1969; Markle and Grant, 1970; Harper and Jarman, 1972; Humphries and Cumming, 1972). At approximately 100 mm in length, fish becomes the major food resource for striped bass and hybrid striped bass and dominates their diet as they grow to maturity (Stevens, 1967; Harper et al., 1969). Aquacultural production of striped bass and their hybrids generally requires the use of live food organisms during the early stages of fish development. In pond rearing this generally involves fertilization to promote the production of zooplankton in adequate quantities (see chapter 7); whereas, in more intensive or artificial systems, rotifers and/or Artemia nauplii are produced for the fish (Tuncer et al., 1993). In either type of system, rapid conversion of the fish to prepared diets is desirable. 9.1.2 Digestive System Structure and Function Striped bass and their hybrids have a rather simple gastrointestinal tract similar to many other carnivorous fish (Dabrowski, 1993). A stomach is present to initiate acidic digestion of protein, and in adult sunshine bass the maximum proteolytic activity is at a pH of 3.49 (Eshel et al., 1993). Numerous pyloric caecae extend from the stomach and aid in nutrient digestion and absorption. Proteolytic activity of the pyloric caeca of sunshine bass is optimum at a pH of 9, and this activity is higher than in the upper and lower intestine (Eshel et al., 1993). The intestine of striped bass and their hybrids is a straight tube typically measuring less than the length of the body. The upper intestine of adult sunshine bass has an average pH of 7.32 while that of the lower intestine is approximately 7.94, and several proteolytic enzymes including trypsin, collagenase, aminopeptidase, elastase, carboxypeptidase A and B, and chymotrypsin are present (Eshel et al., 1993). Larval striped bass have measurable levels of trypsin, chymotrypsin, carboxypeptidase A, and t~-amylase activities at day 4 Oust before initiation of exogenous feeding), and these activities can be 25 to 60% of those at day 32 as larval development nears completion (Baragi and Lovell, 1986). The gastric enzyme pepsin has not been detected in larval striped bass until day 16 after hatching, which corresponds to the development of the stomach (Gabaudan, 1984). A complement of digestive enzymes from the stomach, pancreas, and intestine promote the breakdown of complex nutrients to smaller units that are absorbed from the intestine into circulation. Once in circulation, these nutrients may be used in various anabolic and catabolic processes.

236

9.1.3 Role of Nutrition in Aquaculture Nutrition is a critical factor in intensive aquaculture as it is in other types of animal production because it can significantly influence growth, health, product quality, and cost of production. As production intensity in aquaculture increases, fish generally must rely more heavily on prepared diets to provide all required nutrients for proper growth and health. Therefore, development of appropriate nutrition and feeding programs for aquacultural production of fish is of utmost importance. Feeding costs generally constitute the largest variable cost in intensive fish production; therefore, cost-effective diet formulation can significantly influence the profitability of fish production. Development of nutritious and cost-effective diets is dependent on knowing and meeting a fish's nutritional requirements with balanced diet formulations and appropriate feeding practices. Development of appropriate feeding practices is needed to maximize diet utilization and minimize wastage that can adversely affect water quality and possibly fish health. It has been determined with the fish species examined thus far that they require most of the same nutrients as terrestrial animals, although some differences in qualitative and quantitative requirements as well as in nutrient metabolism have been established (NRC, 1993). For aquacultural species that have been studied extensively such as the channel catfish (lctalurus punctatus), as many as 40 chemical compounds are needed for proper metabolic functioning (Robinson and Wilson, 1985). These compounds are generally classified as either protein, carbohydrate, lipid, vitamin, or mineral based primarily on their chemical nature. Compounds belonging to the protein, carbohydrate, and lipid classes typically can be used to provide energy which is not a nutrient but is needed for various metabolic purposes. 9.2 ENERGY

9.2.1 Metabolism Fish, like other animals, derive energy from ingested nutrients to support various types of work in the body including active transport within cells, muscle activity, and synthesis of various biomolecules for maintenance, repair, and growth of body tissues. However, energy is not expended by fish to maintain body temperature because of their poikilothermic nature. In addition to that energy savings, fish use less energy than terrestrial homeotherms to maintain position and move in the aquatic environment as well as to catabolize protein and excrete nitrogenous wastes. As a result, fish require much less energy for protein synthesis than terrestrial animals (Lovell, 1989). Fish are similar to other vertebrates in the way they obtain energy from dietary carbohydrate, lipid, and protein; however, the efficiency with which these nutrients are digested, absorbed, and processed in intermediary metabolism for energy production varies considerably (NRC, 1993). Specific information pertaining to the ability of striped bass and hybrid striped bass to use protein, carbohydrate, and lipid for energy is discussed below in conjunction with each of these nutrient groups. 9.2.2 Partitioning of Energy Dietary energy is not totally available for productive purposes as some will be lost during digestive and metabolic processes. The various ways in which dietary energy is partitioned have been established for fish (NRC, 1993). Digestible energy, which is the amount of dietary energy available to the fish after digestion, is most frequently used to assess energy utilization. This is because it only requires the determination of fecal energy and gross dietary energy to compute, and it represents the most significant and variable energy loss relative to subsequent metabolism (Cho et al., 1982). Energy digestibility of some

237

practical ingredients has recently been determined with palmetto bass (Sullivan and Reigh, 1995). Additional determinations with different kinds and batches of diet ingredients subjected to different processing conditions are needed to obtain a more complete assessment of dietary energy utilization of striped bass and hybrids. There is limited information concerning the partitioning of dietary energy by striped bass and hybrids, although comparisons between juvenile striped bass and palmetto bass have been made (Tuncer et al., 1990). In that study it was determined that both genetic stocks lost approximately 10.8% of dietary energy intake in the feces. Palmetto bass required approximately 10-12% less energy for metabolism than striped bass and thus had more energy available for growth. Consequently, palmetto bass grew about 10-14% more than striped bass (Tuncer et al., 1990). 9.2.3 Factors Affecting Energy Requirements The metabolic rate of fish is largely influenced by water temperature due to their poikilothermic nature, and metabolic rate affects energy requirements. Striped bass can tolerate a wide range of temperatures, but maximal growth and minimum food intake to maintain body weight occurred around 24-25 ~ for juveniles (Cox and Coutant, 1981). Other environmental factors such as dissolved oxygen, pH, and salinity may affect metabolic rate of fish, although these factors have not been thoroughly evaluated with striped bass and their hybrids. Additionally, biological factors such as body size, age, and genetics, as previously noted for striped bass and palmetto bass, can influence metabolism and energy requirements. Many of these factors are considered in developing appropriate diet formulations and feeding schedules as will be discussed in subsequent sections. 9.3 PROTEIN AND AMINO ACIDS

9.3.1 Structure and Classification Proteins are complex nitrogenous molecules composed of amino acids and are the major constituents of body muscle. Therefore, a continuous supply of protein is required in the diet for maintenance and growth. Ten amino acids are classified as indispensable or essential because they cannot be synthesized in sufficient quantities by the body so a dietary source is required. These include arginine, histidine, isoleucine, leucine, lysine, methionine, phenylalanine, threonine, tryptophan, and valine. Ten other amino acids that commonly make up protein can be synthesized by the body and thus are called dispensable or nonessential. These amino acids have nutritional significance because their presence in the diet conserves energy that would be required for synthesis, and some dispensable amino acids can partially replace or spare indispensable amino acids. In particular, tyrosine can spare approximately 50% of phenylalanine in meeting the total aromatic amino acid requirement of most fish species studied to date, and cystine generally can replace about 50% of methionine as part of the total sulfur amino acid requirement (NRC, 1993). 9.3.2 Functions In addition to supplying amino acids for protein synthesis, dietary protein also may be catabolized for energy. Carnivorous fish species in particular appear to be very proficient at using dietary protein for energy. This is largely due to the efficient way in which ammonia from deaminated protein is excreted via the gills with limited energy expenditure. However, fish growth may be adversely affected as protein intake becomes extremely excessive (NRC, 1993). Weight gain of sunshine bass was reduced at protein levels greater than 45% of dry diet, possibly due to limitation of non-protein energy (Brown et al., 1992). Determining a fish's minimum dietary requirement for protein or a balanced mixture of amino acids is critical. Satisfying this

238

requirement is necessary to ensure adequate fish growth and health, but providing excessive levels is generally uneconomical because protein is often the most expensive dietary component. Therefore, determination of the minimum dietary protein requirement for a species is generally a primary consideration. This protein requirement is typically determined by feeding young, rapidly-growing fish graded levels of a balanced protein. It has been established with several species that the dietary requirement for protein as well as other nutrients will generally decrease as the fish grow. Thus, minimum dietary requirements established with small fish typically will provide a conservative estimate of requirements for larger fish. 9.3.3 Requirements Many carnivorous fishes have grown most rapidly when fed diets with rather high protein levels. Striped bass fed practical diets with 55% crude protein and 16% lipid had significantly greater weight gain than fish fed diets with 34 or 44% crude protein and 16% lipid (Millikin, 1982). In a subsequent feeding trial, practical diets containing either 37, 47 or 57% crude protein promoted similar weight gain of striped bass if the energy provided by lipid was not excessive (Millikin, 1983). Berger and Halver (1987) reported maximum weight gain and feed efficiency of striped bass at 21-22~ when a diet with 52% crude protein was fed at a somewhat restrictive rate of 2.5% body weight per day. Juvenile sunshine bass required protein at approximately 40% of dry diet to produce maximum weight gain in fresh and brackish water (Brown et al., 1992). In that study, growth in artificial seawater (32 9/o0)was severely restricted regardless of dietary protein level. Webster et al. (1995) also reported a protein requirement value of 41% of.diet for sunshine bass reared in cages. A slightly lower protein requirement value of 36% of dry diet also was determined for juvenile sunshine bass in separate studies (P.B. Brown, Purdue University, personal communication). Based on results of these studies, striped bass and sunshine bass appear to have rather similar requirements for dietary protein. Protein requirements of various species have been reported to vary due to a number of factors such as fish size, water temperature, protein quality, non-protein energy, and feed allowance (NRC, 1993). These various factors must be considered in developing feeding programs. In addition, the level of dietary protein that produces the most cost-effective gain may be somewhat lower than that which produces maximum weight gain, depending primarily on the costs of ingredients that provide protein. Determination of individual amino acid requirements of a species will provide even more specific information about its protein needs. The lysine and total sulfur amino acid (methionine plus cystine) requirements are typically the most critical to quantify because the levels of these amino acids in feedstuffs are usually most limiting relative to the amounts required by the fish. Development of suitable test diets to quantify amino acid requirements sometimes may be necessary if diets composed of purified ingredients such as casein and gelatin are not readily accepted by the fish as observed with some marine species (Moon and Gatlin, 1989). A diet formulation consisting of some quantity of a fish-muscle protein together with crystalline amino acids to simulate the amino acid pattern of the fish's muscle protein has been used to quantify amino acid requirements of sunshine bass (Keembiyehetty and Gatlin, 1992, 1993, 1994). The total sulfur amino acid requirement of juvenile sunshine bass was determined to be 2.9% of dietary protein (Keembiyehetty and Gatlin, 1993). A lysine requirement for sunshine bass was established at 4.0% of dietary protein (Keembiyehetty and Gatlin, 1992). The same lysine requirement value was determined by Griffin et al. (1992) using diets containing casein/gelatin and crystalline amino acids. Griffin et al. (1994) reported a lower total sulfur amino acid requirement of approximately 2.0% of dietary protein. The arginine requirement of sunshine bass also has been quantified at approximately 4.3% of dietary protein (Griffin et al.,

239

1994). In addition, the threonine requirement was recently determined to be 2.8% of dietary protein (Keembiyehetty and Gatlin, 1994). Unlike other fish species, weight gain of sunshine bass fed diets containing crystalline amino acids in these various studies generally was not reduced compared to that of fish fed diets with only intact protein. The reason for this difference in response of sunshine bass compared to other fish has not been identified. The remaining indispensable amino acid requirements of sunshine bass should be determined like other established aquaculture species to provide more complete information concerning their amino acid nutrition. It is anticipated that this information also should be applicable to striped bass and palmetto bass. Established relationships between patterns of indispensable amino acids in muscle tissue and required dietary levels also may allow all amino acid requirements of a species to be estimated after quantifying the requirement for only one or two of the most limiting amino acids (Wilson and Poe, 1985; Moon and Gatlin, 1991). 9.3.4 Biological Availability Dietary proteins and amino acids generally are not totally available to the organism due to interactions with other dietary components and/or resistance to digestive enzymes. Therefore, assessing the biological availability of protein and amino acids in dietary ingredients should allow for the organism's requirements to be more accurately met. However, there is no information pertaining to the availability of amino acids from practical diets or feed ingredients to striped bass or hybrid striped bass. Digestibility of protein in several practical feed ingredients was recently determined with palmetto bass (Sullivan and Reigh, 1994) and sunshine bass (Keembiyehetty, 1995). 9.4. CARBOHYDRATES 9.4.1 Structure and Classification Carbohydrates are compounds consisting of carbon, hydrogen, and oxygen. These compounds vary considerably in terms of size and complexity. The carbohydrates with greatest nutritional significance for fish are composed of glucose, a six-carbon monosaccharide. Starch is a complex polysaccharide containing more than six glucose molecules bonded together and comprises a major storage form of energy in plants. Glycogen also is a polysaccharide of glucose that is present in muscle and liver of animals. Cellulose is a polymer of glucose with bonding that allows it to serve a structural role in plants and makes it resistant to digestion by animals with simple gastrointestinal tracts such as fish. 9.4.2 Functions The primary function of digestible carbohydrates in diets of fish is to provide energy. Complex carbohydrates, principally in the form of starch in cereal grains, also may enhance floatability and integrity of extruded pellets due to their expansion and gelatinization during extrusion. Fibrous carbohydrates, including cellulose, hemicellulose, lignin, and pentosans are essentially indigestible by fish and do not positively contribute to their nutrition. This dietary fiber may influence material passage rate through the gastrointestinal tract and thus affect nutrient utilization (NRC, 1993). The fiber level in practical fish diets is typically restricted to less than 8% by weight to limit the amount of undigested material entering the culture system. Some products derived from fibrous carbohydrates, such as hemicellulose and lignosulfonates, are used as binding agents in diet manufacturing.

240

9.4.3 Requirements Fish do not have a specific dietary requirement for carbohydrates, but the presence of digestible carbohydrates in the diet may provide a rather inexpensive source of energy. The ability of fish to utilize dietary carbohydrate for energy varies considerably, with most carnivorous species having more limited ability (Lovell, 1989). Striped bass and hybrid striped bass are carnivorous, but show relatively good ability to use dietary carbohydrate for energy. Striped bass were able to effectively use carbohydrate at up to 33% of diet (Berger and Halver, 1987). Juvenile sunshine bass fed diets with 35% crude protein in which dextrin and lipid levels were varied to assess relative energy utilization of these components showed no differences in weight gain, feed efficiency, and protein efficiency over an eight-week period (Nematipour et al., 1992a). At the extremes, a diet containing 42% dextrin and 2.5% lipid produced similar growth of sunshine bass as a diet containing 25% dextrin and 10% lipid. The various diets did produce significant differences in whole-body composition with the higher dietary lipid levels resulting in more lipid deposition in the visceral cavity (Nematipour et al., 1992a). Juvenile sunshine bass also exhibited efficient utilization of carbohydrate for energy in a subsequent feeding trial (Nematipour et al., 1992b). Thus, striped bass and their hybrids appear to be similar to the omnivorous channel catfish in terms of their ability to utilize soluble carbohydrate for energy (Garling and Wilson, 1977). Digestibility of carbohydrate in practical diet ingredients has not been specifically determined with striped bass or hybrids but deserves consideration. 9.5 LIPIDS

9.5.1 Structure and Classification This nutrient group consists of fats and oils characterized by being insoluble in water and soluble in non-polar organic solvents. The lipids of greatest nutritional importance are fatty acids which consist of a long hydrocarbon chain with a methyl group and carboxyl group at either end. These fatty acids are generally consumed as triglycerides consisting of three fatty acids esterified to a glycerol molecule. 9.5.2 Functions Lipids are important components of fish diets because they provide a concentrated energy source that is typically well utilized. In addition, dietary lipids supply essential fatty acids that cannot be synthesized by the fish but are required for normal metabolic functions. These essential fatty acids are important components of complex lipids in cell membranes and may serve as precursors of prostaglandins. Dietary lipid also provides a vehicle for absorption of fat-soluble vitamins. Lipids from the diet also may be deposited in the fish and affect the flavor, composition, and storage stability of processed products. 9.5.3 Requirements Striped bass and sunshine bass, like most other carnivorous fish, have been shown to efficiently utilize dietary lipid for energy (Nematipour et al., 1992a, 1992b). Levels of menhaden fish oil ranging from 5 to 17% ofthe diet produced maximum weight gain of sunshine bass if the energy to protein ratio (E/P) was appropriate (Nematipour et al., 1992b). Maximum growth was attained by sunshine bass fed diets with an E/P ranging from 6 to 8 kcal available energy/g protein. In a separate study, supplementation of practical diets containing 40% protein and 4% endogenous lipid with menhaden fish oil at 12% reduced the weight gain of sunshine bass relative to fish fed diets with supplemental menhaden oil levels of 4 and 8% (Fair et al., 1993). This growth reduction was probably due to an imbalance of the E/P. Supplementation of menhaden fish oil significantly influenced the fatty acid composition of sunshine bass muscle tissue as observed in other studies (Nematipour and Gatlin, 1993a, 1993b).

241

The essential fatty acid requirements of hybrid striped bass are of particular interest because these fish are crosses between freshwater and marine species which generally have been shown to have different essential fatty acid requirements due to varying ability to elongate and desaturate shorter-chain fatty acids (Watanabe, 1982). In one study (Nematipour and Gatlin, 1993a), sunshine bass were fed diets supplemented with 5% of either coconut oil, linseed oil, olive oil, safflower oil, or menhaden fish oil that provided relatively high levels of lauric (12:0), linolenic (18:3n-3), oleic ( 18: ln-9), linoleic (18:2n-6), and eicosapentaenoic (20:5n-3) plus docosahexaenoic (22:6n-3) acids, respectively. After eight weeks, fish fed the diets containing coconut, linseed, olive, and safflower oils had significantly less weight gain and survival compared to those fed the diet containing menhaden fish oil. Excessive lipid accumulation in the liver and fin erosion were predominant deficiency signs observed in fish fed all diets except the one containing menhaden fish oil. Similar signs of essential fatty acid deficiency have been reported for other fish species (NRC, 1993). Based on the study of Nematipour and Gatlin (1993a), highly unsaturated fatty acids (HUFA) of the n-3 series that are typically found in marine lipids were determined to be essential for sunshine bass. This hybrid striped bass, therefore, appeared to be similar to marine fish with regard to essential fatty acid requirements (Watanabe, 1982). Larvae of striped bass and palmetto bass also have been shown to require HUFA of the n-3 series in their diets (Tuncer and Harrell, 1992). Large quantities of these n-3 HUFA have been found in the eggs of striped bass (Harrell and Woods, 1995). In a subsequent study, graded levels of 20:5n-3 plus 22:6n-3 were fed to sunshine bass and approximately 1.0% of diet (20% of dietary lipid) was required for optimum growth and health (Nematipour and Gatlin, 1993b). Similar requirement values have been determined for a number of marine species (Thongrod et al., 1989; Watanabe et al., 1989; Takeuchi et al., 1992a, 1992b). In contrast to marine fish that typically require preformed HUFA in the diet, freshwater fish such as rainbow trout (Oncorhynchus mykiss) and channel catfish can meet their essential fatty acid requirements by elongating and desaturating dietary 18:3n-3 to 20:5n-3 and 22:6n-3 (NRC, 1993). Other freshwater fish, such as common carp (Cyprinus carpio), require a mixture of 18:3n-3 and 18:2n-6 while others, such as the Nile tilapia (Oreochromisniloticus), require only 18:2n-6 (NRC, 1993). 9.6 MINERALS

9.6.1 Structure and Classification This nutrient group consists of various inorganic elements that are required by the body for various purposes. It has been established that fish require the same minerals as terrestrial animals for tissue formation and other metabolic functions as well as for osmoregulation (Lovell, 1989). In contrast to terrestrial animals, dissolved minerals in the aquatic environment may significantly contribute to satisfying the metabolic requirements offish and interact with dietary requirements. Minerals are typically classified as macrominerals or microminerals based on the amounts found in the body and generally required in the diet. 9.6.2 Macrominerals Macrominerals are generally present in the body in appreciable amounts and required in the diet in rather large quantities. Calcium, chloride, magnesium, phosphorus, potassium, sodium, and sulfur are classified as macrominerals. Fish typically can obtain enough calcium from the water so that a dietary supply is not required (NRC, 1993). In contrast to calcium, phosphorus is typically limiting in water and must be provided in the diet to meet a fish's metabolic requirement. Thus, the dietary phosphorus requirement is usually the most critical to determine for fish. The minimum dietary available phosphorus requirement of juvenile sunshine bass for optimum growth and bone mineralization was determined to be 0.43% of dry diet

242

(Brown et al., 1993). Similar requirement values have been determined for other species (NRC, 1993) including striped bass (Dougall et al., 1994) when expressed as available phosphorus. The availability of phosphorus in feedstuffs may vary considerably with plant feedstuffs typically having less available phosphorus than feedstuffs of animal origin (NRC, 1993). Inorganic sources of phosphorus, such as monobasic and dibasic calcium phosphate, typically provide high levels of available phosphorus and are used to supplement diets. Specific requirements of striped bass and hybrids for other macrominerals have not been determined. However, quantitative requirements for these other macrominerals are anticipated to be similar to those established for other species (NRC, 1993). Adequate levels of these macrominerals are typically provided by practical ingredients so that supplementation is not required. 9.6.3 Microminerals Microminerals are present in the body and required in the diet at much lower levels than macrominerals. These include chromium, copper, iodine, iron, manganese, selenium, and zinc. Dietary deficiencies of most of these microminerals have been experimentally produced with some fish species, but not with striped bass or their hybrids. Dietary deficiencies have been produced by feeding purified diets under controlled conditions for extended periods of time. The essentiality of other microminerals such as fluorine, molybdenum, nickel, silicon, tin, and vanadium has not been established with fish. Of the microminerals, selenium and zinc have been demonstrated in some fish species to be most important to supplement in diets due to low levels in practical feedstuffs and/or interactions with other dietary components that may reduce bioavailability (Gatlin and Wilson, 1984a, 1984b). For example, the minimum zinc requirement of channel catfish was determined with purified diets to be 20 mg Zn/kg diet (Gatlin and Wilson, 1983); however, the presence ofphytate and calcium in practical diets reduced zinc bioavailability such that approximately 200 mg supplemental Zn/kg was needed to provide enough available zinc for the fish (Gatlin and Wilson, 1984b; Gatlin and Phillips, 1989). Similar findings have been reported for other fish species (Ketola, 1979; Richardson et al., 1985). Although supplementation of practical diets with other microminerals has not been shown to be necessary in most instances, an inexpensive trace mineral premix is typically added to most nutritionally complete diets to ensure adequacy. Guidelines for supplementing fish diets with minerals are presented in Table 9.1. 9.7 VITAMINS

9.7.1 Structure and Classification There are 15 vitamins established as being essential for terrestrial animals and some of the fish species that have been examined to date. Specific functions of these vitamins and dietary requirement values have been established for some fish species (NRC, 1993). These organic compounds are classified as fat-soluble and water-soluble based on their chemical properties. 9.7.2 Water-soluble Vitamins Water-soluble vitamins include ascorbic acid, biotin, choline, folic acid, inositol, niacin, pantothenic acid, pyridoxine, riboflavin, thiamin, and vitamin B,2. Dietary deficiencies of almost all of these vitamins in fish have been shown to cause reduced growth and other specific deficiency signs (NRC, 1993). Recommended levels of vitamin supplementation in fish diets have been established based on quantitative

243

Table 9.1. Suggested levels of minerals to provide in extruded or pelleted diets for striped bass and hybrids to meet their established or estimated requirements. Mineral

Amount %

Phosphorus, available

0.5

Copper

5

Iodine Iron

30

Manganese

25

Selenium Zinc

0.1 200

requirements of some species for these vitamins. The levels of supplementation listed in Table 9.2 should provide adequate levels of each vitamin independent of levels in dietary ingredients and also provide a margin of safety for losses associated with processing and storage. These recommended supplementation levels should be used as guidelines until specific requirement values can be quantified for striped bass or their hybrids. To date, only the choline requirement of sunshine bass has been quantified at 500 mg choline/kg diet (Griffin et al., 1994). 9.7.3 Fat-soluble Vitamins Fat-soluble vitamins include vitamin A (retinol), vitamin D (cholecalciferol), vitamin E (tocopherol), and vitamin K. At this time there is no information concerning fat-soluble vitamin nutrition of striped bass or hybrid striped bass. 9.8 FEEDS AND FEEDING PRACTICES 9.8.1 Satisfying Nutritional Requirements Once the nutrient requirements of a species are quantified, diets should be formulated with more accuracy to satisfy those requirements without excessive supplementation, which should reduce diet costs. Satisfying the protein and amino acid requirements of a species is of utmost importance. Providing the proper amount of dietary energy relative to protein (and other nutrients) also is critical to ensure adequate nutrient intake because the fish's intake is generally regulated by energy (Lovell, 1989). A balanced ratio of dietary energy to protein also is important to maximize use of dietary protein for protein synthesis and limit it from

244

Table 9.2. Suggested levels of vitamins to supplement in extruded or pelleted diets for striped bass and hybrids~. Vitamin

Amount/kg

Iu

Ds

K Ascorbic acid B 12 Choline chloride (70%) Folic acid

Amount/ton

I__U_u

4400

4000000

2200

2000000

m_.g

m_.g

55

50000

11

10000

376

342000

0.09 550 2.2

81 500000 2000

Niacin (nicotinic acid)

88

80000

Pantothenic acid

36

32700

Pyridoxine

11

10000

Riboflavin

13

12000

Thiamin

11

10000

1Modified from Robinson (1984)

being catabolized for energy. Based on information to date, diets providing from 6 to 8 kcal available energy/g protein have supported maximum growth of sunshine bass (Nematipour et al., 1992b). The accuracy of diet formulation can be improved to a greater extent if the digestibility or availability of the various nutrients from feed ingredients is known. Unfortunately, information on nutrient availability from various feedstuffs has been difficult to obtain with most fish species. Obstacles associated with monitoring feed intake of fish and collecting excretory products in the aquatic environment have complicated digestibility determinations. Problems associated with feed intake have been reduced by using various markers in diets for determining digestibility by indirect methods (NRC, 1993). A variety of fecal collection methods also have been evaluated in an attempt to maximize the opportunity for digestion while minimizing leaching of nutrients from feces into the water (NRC, 1993). Digestibility coefficients of protein, lipid, and energy in some practical ingredients recently have been determined with palmetto bass (Sullivan and Reigh, 1995). Further advancements in this area will be of primary importance in improving the efficiency of diet formulation for striped bass and hybrids.

245

9.8.2 Feedstuffs In formulating diets to meet the nutrient requirements of most fish species, relatively few ingredients have been used. This is primarily because only a limited number of feedstuffs have the nutrient density and physical characteristics required to meet the nutritional needs of fish. Feedstuffs of marine origin, such as various fish meals, have been most effective in diet formulations for carnivorous fish species because they are generally quite palatable and high in protein, lipid, and energy. However, these feedstuffs are usually quite expensive and substantially increase diet costs. Other feedstuffs of animal origin, such as poultry by-product meals and spray-dried blood meal, have been reported to replace 75 and 25%, respectively, of the protein from fish meal without reducing weight gain of hybrid striped bass (Gallagher, 1992a). Meat and bone meal also has some potential for use in diets for striped bass and hybrids. The quality of these various animal by-product meals, however, can be quite variable and should be closely monitored. Protein feedstuffs of plant origin, such as soybean meal and cottonseed meal, are generally less expensive than feedstuffs of animal origin, but are limited in diet formulations of many carnivorous species because they are usually lower in digestibility and palatability, and contain less of the indispensable amino acids. Soybean meal, however, was reported to replace between 50 and 75% of the protein from fish meal in diets of hybrid striped bass without causing growth reduction (Gallagher, 1992b). Keembiyehetty (1995) also found that soybean meal could replace up to 75% of the protein from fish meal without affecting growth or feed efficiency of sunshine bass ifmethionine also was supplemented to meet their total sulfur amino acid requirement. Additional evaluations of practical feedstuffs with striped bass and hybrids are warranted. A variety of cereal grains and their by-products including com, wheat, wheat flour, wheat middlings, and rice bran have been used in fish diets to supply available carbohydrate for energy and increase pellet stability. Quantities of these grain by-products included in diets of carnivorous fish are generally restricted due to the limited ability of these fish to use soluble carbohydrate and the need for greater concentrations of protein and lipid. Supplemental lipid, vitamins, and minerals also are included in nutritionally complete fish diets to ensure that all nutrient requirements are satisfied. Lipids of marine origin are commonly included in the diets of marine fish to provide essential fatty acids and improve diet palatability. Vitamin and mineral premixes for fish diets also are commercially available. 9.8.3 Diet Formulation and Manufacture A variety of methods can be employed to formulate diets for fish. Least-cost diet formulation, which is used extensively with terrestrial livestock, has had more limited application with most fish species due to a restrictive number of potential ingredients and a lack of information concerning the value of those ingredients to fish (Lovell, 1989). As nutritional information about aquacultured species and feedstuffs becomes more readily available, the use of least-cost diet formulation also may become more widely used. Striped bass and their hybrids have been successfully reared with a variety of diet formulations including commercial salmonid diets. Some experimental and practical diets have been evaluated with striped bass (Hughes et al., 1992) and hybrids (Brown et al., 1993). As information on nutritional requirements of these fishes and their utilization of practical ingredients becomes available, diets can be more precisely formulated. A model formulation for production of striped bass and hybrids is presented in Table 9.3. Research to determine the cost-effectiveness of different diet formulations for striped bass and hybrids has been rather limited to date, but deserves further consideration.

246

Table 9.3. Model diet formulation for striped bass and hybrids from juvenile (--10 g) to 1 kg. Ingredient

% as fed

High-quality fish meal

33.1

Soybean meal (49% protein)

31

Wheat flour

30.2

Menhaden fish oil Mineral premix ~

0.1

Vitamin premix ~

0.1

Dicalcium phosphate

0.5

Coated ascorbic acid

0.038

Composition Prote in (%)

40

Lipid (%) Fiber (%)

1.7

Estimated available energy (kcal/g)

3.3

Energy:protein ratio (kcal/g protein)

8.25

~Should provide the levels recommended in Tables 9.1 and 9.2 Another important aspect of fish nutrition and feeding involves the feed manufacturing process. Because fish live in the aquatic environment, care must be taken to ensure they are able to obtain a full complement of nutrients without excessive losses to the water. Thus, feed ingredients are usually agglomerated into larger particles by either pressure pelleting or extrusion processing. Pressure pelleting generally requires a binding agent and results in the formation of sinking pellets. In contrast, extrusion processing, which involves greater heat and pressure, produces low-density particles with greater water stability that may float or be neutrally buoyant in water. Striped bass and hybrids have been fed both pelleted and extruded diets with acceptable results. Specific comparisons of these diet forms have not been made experimentally. 9.8.4 Feeding Practices Appropriate feeding schedules and practices must be employed in aquaculture to ensure that maximum benefit can be derived from prepared diets. It is generally desirable in production aquaculture for fish to consume as much diet as they desire on a regular basis. However, excessive feeding should be avoided because it not only wastes expensive diet, but also deteriorates the quality of water in which the fish live. Pellet size and feeding rates should be regularly adjusted according to fish size.

247

Feeding schedules for the aquacultural production of a species primarily consider the effects of fish size and water temperature. In general, smaller fish consume more diet, when expressed as a percentage of body weight, than do larger fish. Smaller fish also should be fed more frequently than larger fish due to their higher metabolic rate. In addition, water temperature also may significantly influence feed intake of fish with reduced consumption occurring above and below a species' optimal temperature. The optimum feeding rate for striped bass fingerlings (initial weight of 38 g) raised at 19~ was between 1 and 1.5% of body weight per day (Hung et al., 1993). In contrast, juvenile striped bass and palmetto bass initially weighing 2.3 and 1.8 g, respectively, required a feeding rate above 5% of body weight per day for maximum weight gain at 24~ (Tuncer et al., 1990). Specific feeding schedules are usually empirically derived for various aquaculture species. In general, when striped bass and hybrid juveniles are reared in fertilized ponds, a prepared diet in the form of mash or # 1 crumble will usually be added to the pond approximately two to three weeks after stocking the larval fish to complement natural productivity. At four weeks, fish should be readily consuming the prepared diet and deriving most of their nourishment from it. Feeding rate, frequency and diet particle size should be adjusted regularly as the fish grow. The physical characteristics of the diet and means by which it is administered to fish will be dictated by a number of factors including design and size of culture systems and preferences of the fish and aquaculturist.

248

References Baragi, V. and Lovell, R.T., 1986. Digestive enzyme activities in striped bass from first feeding through larva development. Transactions of the American Fisheries Society, 115: 478-484. Berger, A. and Halver, J.E., 1987. Effect of dietary protein, lipid and carbohydrate content on the growth, feed efficiency and carcass composition of striped bass, Morone saxatilis (Walbaum), fingerlings. Aquaculture and Fisheries Management, 18:345-356. Brown, M.L., Nematipour, G.R. and Gatlin, III, D.M., 1992. Dietary protein requirement ofjuvenile sunshine bass at different salinities. The Progressive Fish-Culturist, 54: 148-156. Brown, M.L., Jaramillo, Jr., F. and Gatlin, III, D.M., 1993. Dietary phosphorus requirement of hybrid striped bass, Morone chrysops x M. saxatilis. Aquaculture, 113:355-363. Brown, P.B., Griffin, M.E. and White, M.R., 1993. Experimental and practical diet evaluations with juvenile hybrid striped bass. Journal of the World Aquaculture Society, 24: 80-89. Cho, C.Y., Slinger, S.J. and Bayley, H.S., 1982. Bioenergetics of salmonid fishes: energy intake, expenditure and productivity. Comparative Biochemistry and Physiology B, 73:25-41. Cox, D.K. and Coutant, C.C., 1981. Growth dynamics of juvenile striped bass as functions of temperature and ration. Transactions of the American Fisheries Society, 110: 226-238. Dabrowski, K., 1993. Ecophysiological adaptations exist in nutrient requirements of fish: true or false? Comparative Biochemistry and Physiology A, 104: 579-584. Dougall, D.S., Woods, III. L.C. and Soares, Jr, J.H., 1994. Dietary phosphorus requirement of the striped bass, Morone saxatilis. World Aquaculture '94, New Orleans, Louisiana, p. 59, Abstract. Eshel, A., Lindner, P. Smimoff, P., Newton, S. and Harpaz, S., 1993. Comparative study of proteolytic enzymes in the digestive tracts of the European sea bass and hybrid striped bass reared in freshwater. Comparative Biochemistry and Physiology A, 106: 627-634. Fair, P.H., Williams, W.P. and Smith, T.I.J., 1993. Effect of dietary menhaden oil on growth and muscle fatty acid composition of hybrid striped bass, Morone chrysops x M. saxatilis. Aquaculture, 116:171-189. Gabaudan, J., 1984. Posthatching morphogenesis of the digestive system of striped bass. Ph.D. Dissertation, Auburn University, Auburn, AL. Gallagher, M.L., 1992a. Poultry meals as replacements for fishmeal in diets for hybrid striped bass (Morone sp.). Aquaculture '92, Orlando, FL, p. 99, Abstract. Gallagher, M.L., 1992b. Soybean replacement of fishmeal in diets for hybrid striped bass (Morone sp.). Aquaculture '92, Orlando, FL, p. 99, Abstract. Garling, Jr., D.L. and Wilson, R.P., 1977. Effects of dietary carbohydrate-to-lipid ratios on growth and body composition of fingerling channel catfish. The Progressive Fish-Culturist, 39: 43-47. Gatlin, III, D.M. and Wilson, R.P., 1983. Dietary zinc requirement of fingerling channel catfish. Journal of Nutrition, 113: 630-635.

249

Gatlin, III, D.M. and Wilson, R.P., 1984a. Dietary selenium requirement of fingerling channel catfish. Joumal of Nutrition, 114: 627-633. Gatlin, III, D.M. and Wilson, R.P., 1984b. Zinc supplementation of practical channel catfish diets. Aquaculture, 41: 31-36. Gatlin, III, D.M. and Phillips, H.F., Aquaculture, 79: 259-266.

1989. Dietary calcium, phytate and zinc interactions in channel catfish.

Griffin, M.E., Brown, P.B. and Grant, A.L., 1992. The dietary lysine requirement of juvenile hybrid striped bass. Journal of Nutrition, 122: 1332-1337. Griff'm, M.E., White, M.R. and Brown, P.B., 1994. Total sulfur amino acid requirement and cysteine replacement value for juvenile hybrid striped bass (Morone saxatilis x M. chrysops). Comparative Biochemistry and Physiology A, 108: 423-429. Griffin, M.E., Wilson, K.A. and Brown, P.B., 1994. Dietary arginine requirement of juvenile hybrid striped bass. Journal of Nutrition, 124: 888-893. Griffin, M.E., Wilson, K.A., White, M.R. and Brown, P.B., 1994. Dietary choline requirement of juvenile hybrid striped bass. Journal of Nutrition, 124: 1685-1689. Harper, J.L. and Jarman, R., 1972. Investigation of striped bass, Morone saxatilis (Walbaum), culture in Oklahoma. Proceedings of the Annual Conference of the Southeastern Association of Game and Fish Commissioners, 25: 501-512. Harper, J.L., Jarman, R. and Yacovino, Y.T., 1969. Food habits of young striped bass, Roccus saxatilis (Walbaum), in culture ponds. Proceedings of the Annual Conference of the Southeastern Association of Game and Fish Commissioners, 22: 373-380. Harrell, R.M. and Bukowski, J., 1990. The culture, zooplankton dynamics and predator-prey interactions of Chesapeake Bay striped bass, Morone saxatilis (Walbaum), in estuarine ponds. Aquaculture and Fisheries Management, 21: 195-212. Harrell, R.M. and Woods, III, L.C., 1995. Comparative fatty acid composition of eggs from domesticated and wild striped bass (Morone saxatilis). Aquaculture, 133: 225-233. Harrell, R.M., Loyacano, Jr., H.A. and Bayless, J.D., 1977. Zooplankton availability and feeding selectivity of fingerling striped bass. Georgia Journal of Science, 35: 129-135. Heubach, W., Toth, R.J. and McCready, A.M., 1963. Food of young-of-the-year striped bass (Roccus saxatilis) in the Sacramento-San Joaquin river system. California Fish Game, 49: 224-239. Hughes, S.G., Lemm, C.A. and Herman, R.L., 1992. Development of a practical diet for juvenile striped bass. Transactions of the American Fisheries Society, 121: 802-809. Humphries, E.T. and Cumming, K.B., 1972. Food habits and feeding selectivity of striped bass fingerlings in culture ponds. Proceedings of the Annual Conference of the Southeastern Association of Game and Fish Commissioners, 25: 522-536. Hung, S.S.O., Conte, F.S. and Hallen, E.F., 1993. Effects of feeding rates on growth, body composition and nutrient metabolism in striped bass (Morone saxatilis) fingerlings. Aquaculture, 112:349-361.

250

Keembiyehetty, C.N., 1995. Amino acid nutrition of sunshine bass (Morone chrysops x M. saxatilis). Ph.D. Dissertation, Texas A&M University, College Station, Texas, 84 pp. Keembiyehetty, C.N. and Gatlin, III, D.M., 1992. Dietary lysine requirement of juvenile hybrid striped bass (Morone chrysops x M. saxatilis). Aquaculture, 104: 271-277. Keembiyehetty, C.N. and Gatlin, III, D.M.,. 1993. Total sulfur amino acid requirement ofjuvenile hybrid striped bass, Morone chrysops x M. saxatilis. Aquaculture, 110:331-339. Keembiyehetty, C.N. and Gatlin, III, D.M.,. 1994. Dietary threonine requirement of sunshine bass, Morone chrysops 2 x M. saxatilis c~. World Aquaculture '94, New Orleans, LA., p. 61 Abstract. Ketola, H.G., 1979. Influence of dietary zinc on cataracts in rainbow trout (Salmo gairdneri). Journal of Nutrition, 109: 965-969. Lovell, T., 1989. Nutrition and feeding of fish. Van Nostrand Reinhold, New York, 260 pp. Markle, D.F. and Grant, G.C., 1970. The summer food habits of young-of-the year striped bass in three Virginia rivers. Chesapeake Science, l l: 50-54. Millikin, M.R., 1982. Effects of dietary protein concentration on growth, feed efficiency, and body composition of age0 striped bass. Transactions of the American Fisheries Society, 111: 373-378. Millikin, M.R., 1983. Interactive effects of dietary protein and lipid on growth and protein utilization of age-0 striped bass. Transactions of the American Fisheries Society, 112: 185-193. Moon, H.Y. and Gatlin, III, D.M., 1989. Amino acid nutrition of the red drum (Sciaenops ocellatus): determination of limiting amino acids and development of a suitable amino acid test diet. Proceedings of the third international symposium on feeding and nutrition in fish, Toba, Japan, pp. 201-208. Moon, H.Y. and Gatlin, III, D.M., 1991. Total sulfur amino acid requirement ofjuvenile red drum, Sciaenops ocellatus. Aquaculture, 95: 97-106. (NRC) National Research Council., 1993. Nutrient requirements of fishes. National Academy Press, Washinpon, DC. 114 pp. Nematipour, G.R., Brown, M.L. and Gatlin, III, D.M., 1992a. Effects of dietary carbohydrate:lipid ratio on growth and body composition of hybrid striped bass. Journal of the World Aquaculture Society, 23: 128-132. Nematipour, G.R., Brown, M.L. and Gatlin, III, D.M., 1992b. Effects of dietary energy:protein ratio on growth characteristics and body composition of hybrid striped bass, Morone chrysops x M. saxatilis. Aquaculture, 107: 359-368. Nematipour, G.R. and Gatlin, III, D.M., 1993a. Effects of different kinds of dietary lipid on growth and fatty acid composition of juvenile hybrid striped bass, Morone chrysops x M. saxatilis. Aquaculture, 114:141-154. Nematipour, G.R. and Gatlin, III, D.M., 1993b. Requirement of hybrid striped bass, Morone chrysops x M. saxatilis, for dietary (n-3) hio~hlyunsaturated fatty acids. Journal of Nutrition, 127: 744-753. Regan, D.M., Wellborn, T.L. and Bowker, R.G., 1968. Striped bass, Roccus saxatilis (Walbaum): development of essential requirements for production. U.S. Fish and Wildlife Service, Atlanta, GA, 133 pp.

251

Richardson, N.L., Higgs, D.A., Beames, R.M. and McBride, J.R., 1985. Influence of dietary calcium, phosphorus, zinc and sodium phytate level on cataract incidence, growth and histopathology in juvenile chinook salmon (Oncorhynchus tshawytscha). Journal of Nutrition, 115: 553-567. Robinson, E.H., 1984. Vitamin requirements. Pages 21-25 in E.H. Robinson and R.T. Lovell, editors, Nutrition and feeding of channel catfish (revised), Southern Cooperative Series Bulletin No. 296, Texas A&M University, College Station, TX. Robinson, E.H. and Wilson, 1LP., 1985. Nutrition and feeding. Pages 323-404 in C.S. Tucker, editor, Channel catfish culture. Elsevier, Amsterdam. Stevens, D.E., 1967. Food habits of striped bass, Roccus saxatilis, in the Sacramento - San Joaquin delta. California Department ofFish and Game, Fisheries Bulletin, 136: 68-96. Sullivan, J.A. and Reigh, R.C., 1995. Apparent digestibility of selected feedstuffs in diets for hybrid striped bass (Morone saxatilis ~x M. chrysops cO. Aquaculture, 138: 313-322. Takeuchi, T., Shiina, Y. and Watanabe, T., 1992a. Suitable levels of n-3 highly unsaturated fatty acids in diet for fingerlings of red sea bream. Nippon Suisan Gakkaishi, 58:509-514. Takeuchi, T., Shiina, Y., Watanabe, T., Sekiya, S. and Imaizumi, K, 1992b. Suitable levels of n-3 highly unsaturated fatty acids in diet for fingerlings of yellowtail. Nippon Suisan Gakkaishi, 58:1341-1346. Thongrod, S., Takeuchi, T., Satoh, S. and Watanabe, T., 1989. Requirement of f'mgerling white fish Coregonus lavaretus maraena for dietary n-3 fatty acids. Nippon Suisan Gakkaishi, 55:1983-1987. Tuncer, H. and Harrell, R.M., 1992. Essential fatty acid nutrition of larval striped bass (Morone saxatilis) and palmetto bass (M. saxatilis x M. chrysops). Aquaculture, 101: 105-121. Tuncer, H., Harrell, R.M. and Houde, E.D., 1990. Comparative energetics of striped bass (Morone saxatilis) and hybrid (M. saxatilis x 34. chrysops) juveniles. Aquaculture, 86: 387-400. Tuncer, H., Harrell, R.M. and Chai, T., 1993. Beneficial effects of n-3 HUFA enriched Artemia as food for larval palmetto bass (Morone saxatilis x 34. chrysops). Aquaculture, 110:341-359. Watanabe, T., 1982. Lipid nutrition in fish. Comparative Biochemistry and Physiology B, 73: 3-15. Watanabe, T., Takeuchi, T., Arakawa, T., Imaizumi, K., Sekiya, S. and Kitajima, C., 1989. Requirement ofjuvenile striped jack Longirostris delicatissimus for n-3 hi~hly unsaturated fatty acids. Nippon Suisan Gakkaishi, 55: 1111-1117. Webster, C.D., Tiu, L.G., Tidwell, J.H., Van Wyk, P. and Howerton, R.D., 1995. Effects of dietary protein and lipid levels on growth and body composition of sunshine bass (Morone chrysops x M. saxatilis)reared in cages. Aquaculture, 131: 291-301. Wilson, R.P. and Poe, W.E., 1985. Relationship of whole body and egg essential amino acid patterns to amino acid requirement patterns in channel catfish, lctaluruspunctatus. Comparative Biochemistry and Physiology B, 80: 385-388. Woods, L.C., Lockwood, J.C., Kerby, J.H. and Huish, M.T., 1985. Feeding ecology of hybrid striped bass (Morone saxatilis X M. chrysops) in culture ponds. Journal of the World Mariculture Society, 16: 71-81.

This Page Intentionally Left Blank

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

253

Chapter 10

Environmental Requirements and Noninfectious Diseases Joseph g. Tomasso 10.1 INTRODUCTION Striped bass and its hybrids, like all fishes, have certain environmental requirements and are sensitive to noninfectious disease agents. This chapter examines the known environmental requirements and some common noninfectious diseases of Morone. The reader is also referred to Hall (1991), Bills et al. (1993), and Kerby (1993) which present original information and review existing information related to the environmental requirements and noninfectious diseases of striped bass. Before discussing the specifics of striped bass and its hybrids, a review of the concepts of optimal and stressful environments would be helpful. From the standpoint of aquacultural productivity, the relationship of an animal to its environment is best viewed energetically. Energy devoted by an animal to dealing with the environment (such as for water and ion balance) is not available for growth or reproduction. Fry (1947, 1971), through the use of oxygen consumption rates, described the relationship between energy requirements for physiological maintenance and energy available for other activities as "metabolic scope?' Metabolic scope was used by Schreck (1981 ) as one of the bases for his concept of performance capacity of fishes. Schreck argued that fish have some maximum, genetically determined ability to perform. The concept of performance was viewed by Schreck as an '2q-dimensional geometric physiological capacity." In aquaculture, performance may be viewed in the context of integrated responses (such as growth and reproduction). The genetically based performance capacity (referred to as potential performance capacity) is reduced by nonoptimal environments to a realized performance capacity, which can be further limited by stress. The concepts of metabolic scope and performance capacity can act as guides and tools to produce optimal aquaculture environments -environments that minimize energy requirements for maintenance activities and maximize energy available for production. Nonoptimal environments can be distinguished from stressful environments by the responses evoked in the culture animals. A nonoptimal environment is simply one that limits the performance capacity of an animal but is not actually deleterious. For example, a species with an optimal growth temperature of 25 ~ will probably grow well at 22~ --just a little slower. Stressful environments, on the other hand, induce a specific physiological response by an animal (discussed below). The stress response evolved to help animals deal with stressors (environmental threats) and has an energetic cost associated with it. Decreased resistance to infectious diseases is an additional consequence of stress. Researchers have tended to consider nonoptimal environments and stressful environments separately; however, aquaculturists might do well to consider them as a continuum. 10.2 STRESS AND DISEASE RESISTANCE Stress is a term used to describe the physiological responses of an animal to the presence of a stressor. A stressor may be a physical, chemical, or social force perceived by the animal as a threat to its well being. The responses of fish to stressors have been described in detail (Wedemeyer et al., 1990; Pickering, 1981; Barton and Iwama, 1991; Picketing, 1993). Briefly, the fish must first sense the presence of a stressor. Signals are then sent from the brain by way of the sympathetic nervous system to the chromaff'm cells near the posterior cardinal vein and by way of the hypothalamic-pituitary-interrenal axis to the interrenal tissue. In response to

254

these signals, catecholamines (epinephrine, norepinephrine, and dopamine) are released from the chromaffin cells into circulation, and corticosteroids (cortisol, cortisone, and corticosterone) are released into circulation by the interrenal tissue. The catecholamines and corticosteroid hormones bring about a number of physiological changes (stress) in the animal directed toward allowing the animal to better deal with the stressor. Stress or the "stress response" is characteristically the same qualitatively, regardless of the nature of the stressor. The same set of physiological changes occurs in response to handling, low dissolved oxygen concentrations, high ammonia concentrations, or other aquaculture related stressors. However, the magnitude of the response does differ. Generally, the magnitude is proportional to the severity and the length of application of the stressor until some limit is reached (Barton and Iwama, 1991). One detrimental effect of stress is immunosuppression, which may lead to disease outbreaks caused by pathogens that had previously infected the animals but were held in check by the immune system. The mechanisms of immunosuppression in fishes, while not well characterized, have been much reviewed and discussed (Wedemeyer, 1970; Snieszko, 1974; Ellis, 1981; Adams, 1990; Barton and Iwama, 1991; Pickering, 1993). Most evidence points to the corticosteroids as the modulator of immunocompetence during and following stress. The paradoxical effects ofcorticosteroids - preparing the animal to deal with a stressor while predisposing the animal to infectious disease -- have not yet been satisfactorily explained. Pickering (1993), citing mammalian studies, suggested that cortisol in unstressed fish may modulate the immune system ina way to prevent autoimmune effects. However, elevated circulating corticosteroid levels during and after chronic stress associated with some aquaculture operations may overregulate the immune system resulting in immunosuppression. Intrinsic in this line of thinking is the assertion that fish are not chronically stressed in natural habitats, but rather acutely (short-term) stressed such as in the case of avoiding a predator. Conversely, fish in aquaculture settings may be exposed to chronic (long-term) stressors such as crowding, low dissolved oxygen concentrations, or high levels of ammonia. Stress, then, can be thought of as both a noninfectious disease and as a "trigger" for activating latent infectious diseases. The triggering function of stress is very important in aquaculture because it effectively amplifies the effect of the stressor by allowing an infectious agent to cause disease. Thus, an environmental stressor which may cause limited problems by itself may initiate an epizootic with disastrous consequences. Plasma cortisol concentrations were determined for striped bass from four warmwater reservoirs (Tisa et al., 1983). The 95% confidence limits for cortisol concentrations in these fish were 0-12.5 #g/100mL. Elevated plasma cortisol or total corticosteroid concentrations have been reported in striped bass in response to elevated environmental nitrite (Mazik et al., 1991 a); capture, handling and transport (Davis and Parker, 1990; Mazik et al., 1991b; Harrell, 1992; Harrell and Moline, 1992); exposure to anesthetics (Davis et al., 1982); confinement and forced swimming (Strange and Cech, 1992); and serial sampling of tanks (Young and Cech, 1993). Sunshine bass (white bass x striped bass) also exhibit increased plasma corticosteroid levels following handling and transport (Tomasso et al., 1980). 10.3 ENVIRONMENTAL REQUIREMENTS 10.3.1 General Comments The components of an aquaculture environment may be divided into those that directly affect the culture animal (primary components) and those that have an indirect effect (secondary environmental components). Water quality interactions in warmwater culture ponds offer good examples (Boyd, 1979). The

255

dissolved oxygen concentration of the pond water has a direct effect on the well-being of culture animals. Phytoplankton populations have no direct effect on culture animals (unless the culture system relies on natural food to support aquaculture production or a toxic phytoplankton species is present), but directly affect dissolved oxygen concentrations by net production of oxygen during the day and net consumption at night. Similarly, extreme pH levels can directly affect culture animals (and the toxicity of some nitrogenous wastes), but alkalinity affects the animals in that it influences the daily pH fluctuations of the pond. Culturists should manage for acceptable or optimal, depending on the economics of the situation levels of primary components. In pond aquaculture, this often means managing the secondary component which controls the primary component of interest. For example, extreme pH fluctuations in ponds can be managed by increasing alkalinity through liming (Boyd, 1979). Raceway and cage culturists can usually manage the primary components directly -- often by simply modifying the water flow or aerating the system. Some of the primary environmental components for culture of striped bass and hybrid striped bass are discussed below. 10.3.2 Temperature Temperature is the predominant physiochemical characteristic in aquaculture operations. It affects the amount of oxygen a given amount of water will hold (Boyd, 1979), the amount of oxygen the aquaculture animals consume per unit of time and body mass, the rate at which animals feed, and the rate of photosynthesis and decomposition. Fry (1947, 1971) characterized temperature as an environmental controller of metabolism. Effects of all other environmental variables on animals are affected by temperature. Due to the high specific heat of water, a great deal of energy is required to raise or lower its temperature. Hence, modifying water temperatures at aquaculture facilities is rarely economical, and the importance of siting aquaculture facilities where ambient water temperatures are compatible with the species being cultured becomes very clear. A great deal of research has been conducted on the relationship between striped bass and their thermal habitat. Much of this work has been directed toward understanding the relationship between survival and temperature in striped bass fisheries, particularly in reservoirs of the southern United States. Coutant (1985) discussed the problem of summer mortality and hypothesized that large striped bass are being "squeezed" out of suitable habitat as temperatures rise. He speculated that warming near the surface in conjunction with developing regions of low dissolved oxygen concentrations near the bottom of the reservoir resulted in an absence of suitable habitat. Because the summer mortality problem was limited to larger fish, Coutant inferred that juvenile striped bass were more tolerant to high temperatures than fish in their second year or later. An optimal temperature for the culture of striped bass is difficult to identify. The age of the fish and salinity interact to produce different optima, and hybrids apparently have different optima than striped bass. Table 10.1 summarizes available information on thermal requirements of striped bass and hybrids. Generally (based on the work cited in Table 10.1), striped bass eggs, larvae, and fry do best at about 18~ Fingerlings grow best and prefer temperatures in the mid to upper 20s~ Juveniles and adults grow best and prefer lower to mid 20s ~ Hybrid fingerlings grow best at 31 ~ but best food conversion efficiency is at 19~ Juvenile hybrids grow best at about 27~ in 12 hours of daylight, but changing the photoperiod affects thermal optima. 10.3.3 Dissolved Oxygen Adequate dissolved oxygen concentrations are critical during all phases of striped bass and hybrid culture. Low dissolved oxygen concentrations can result in slower growth and induce the stress response,

256

Table 10.1. Temperatures as a function of survival and growth reported for striped bass and hybrid striped bass. Life stage

Temperature (~

Comments Striped Bass

Eggs

18.3 a 18.1-18.7 b 16.0-18.0 i 17.8-20.tY

Best hatching percentage for non-water hardened eggs Best hatching percentage Best hatching percentages, 0-10 g/L salinity Recommended hatching temperature

Prolarvae

18.0 i 18.0-32.0 !

Best survival after 24 h, 0-10 ~JL Recommended up to 9 d of age

Larvae

18.0-321 17.6 r

Suitable for rearing in ponds Best survival after 24 h

Fingerlings

17.8-20.0 k 26.9-30.3 a 27.0 d Overwintering f

Optimal temperature Optimum growth Final preferendum 99.5% survival over winter under ice in cages

Juveniles

24.0 e >33.5 ~ <10.0" 14.0-22.0"

Maximum growth at 100% satiation No growth No growth Greatest bioenergetic efficiency in fluctuating temp.

Phase III

20.0-24.0 h

Thermal niche based on selection in reservoir

Adults

21.6 s

Selected in reservoir during summer Morone

Hybrids

Fingerlings

Overwintering f 31.0 ~ 19.0 ~

100% survival over winter under ice in cages Optimal growth Best food conversion efficiency

Juveniles

26.8" 21.2 m 27.9 m 25.7 m 6.7" 26.6" 28.0 n

Best growth in 12 h daylig,ht Peak conversion efficiency in 12 h daylight Maximum growth under springtime conditions Maximum growth under autumnal conditions No growth Optimal temperature for growth Peak food consumption

'Turner and Farley (1971), 1 ~ salinity; bShannon and Smith (1968), Roanoke River water; CDavies (1973), flesh water; aKellogg and Gift (1983), test water conductivity=100-226/.zmhos; eCox and Coutant (1981), well water; fHarrell et al. (1988), freshwater, palmetto bass; rv'an Den Avyle and Evans (1990), coastal fiver system; hCoutant and Carroll (1980), freshwater lakes; iMorgan et al. (1981), variable salinities; JRees and Harrell (1990); kBrewer and Rees (1990); tBonn et al. (1976); mWoiwode and Adelman (1991), palmetto bass; "Woiwode (1989), palmetto bass; ~ and Adelman (1984), palmetto bass.

257

predisposing the animals to infectious disease. Monitoring of dissolved oxygen concentrations is complicated by the rate at which they can change. In heavily stocked raceways, tanks, or flow-through systems, for example, an interruption of oxygenation may result in critically low dissolved oxygen concentrations within minutes due to consumption by the culture animals. Management of dissolved oxygen concentrations in ponds must also consider the daily rhythms of concentrations characteristic of ponds (Boyd, 1979). Striped bass and its hybrids have different dissolved oxygen requirements at different stages in their lives. Table 10.2 summarizes much of the available information on dissolved oxygen requirements. Striped bass also appear to require higher concentrations of dissolved oxygen relative to other temperate species. Generally, dissolved oxygen concentrations should be maintained as close to saturation as possible for best survival and growth. Table 10.2. Dissolved oxygen (DO) requirements of striped bass. Life Stage

DO (moSL)

Embryos

>3.0 a

Necessary for normal development

Eggs

Saturationb

Best hatching percentage and larval survival

Fingerlings

5.1 c

3.1 r > 6.0 d

Reduced growth at 20 ~ C compared to 7.4 mg/L Reduced growth at 25 ~ C compared to 6.7 mg/L Recommended for phase I pond culture

> 5.0 ~ <4.9 f 1.0s 0.5 g

Preferred for phase II culture Avoided Fish began dying Nearly all fish dead

Juveniles

Comments

aHarrell and Bayless (1984), well water; bTurner and Farley (1971), freshwater, 18.0-22.6 ~ C; CCech et al. (1984), freshwater, DO values calculated from Torr values; dBrewer and Rees (1990); *Kerby et al. (1983); fHill et al. (1981); gChittenden (1971), 16-19~ C, 0 and 10 g/L salinity, "negligible or absent" effects of salinity

10.3.4 Salinity Salinity (%0) is a measure of the solids dissolved in seawater (Spotte, 1979), and is distinct from sodium chloride concentrations. Some published reports, particularly in the older literature, use the term salinity to describe g/L concentrations of sodium chloride. I will concentrate on salinity requirements during aquaculture operations. The uses of sodium chloride in fish transportation and treatment of infectious diseases is discussed in Chapters 7 and 11, and the use of chloride to inhibit nitrite toxicosis is discussed in section 10.4.3 of this chapter. Striped bass adults are euryhaline and move up rivers into freshwater to spawn in the spring. Table 10.3 summarizes some of the literature on salinity and striped bass and its hybrids. Generally, egg hatching occurs best at 10 %o or less. Larvae and fry should be held in salinities below 15~ Fingerlings and juveniles can apparently be reared in freshwater to near-full strength sea water. Palmetto bass fingerlings grow equally well in freshwater to seawater, but survival is lower in full strength seawater.

258

Table 10.3. Salinity requirements of striped bass and hybrid striped bass. Life stage

Salinity

Comments

(%0) Striped Bass Eggs

0-10.0 ~ 0-5.0 ~

>80% hatch at 16-18~ C Recommended for hatching

Prolarvae

5.0-10.0 e 0-15.0 h

>80% survival after 24 h at 18~ C Habitat requirements for survival of prolarvae

Larvae

0-25.0 h 10.0-20.0 a 24.5 ~

Habitat requirements for survival of postlarvae Larvae survived for 2 weeks, unreplicated All larvae died within 30 h, unreplicated

Fingerlings

>0.5 a 4.0 or 12.0g

Single most important factor influencing hatchery production >80% survival after abrupt transfer from fresh water (18 or 24 ~ C)

Juveniles

0 vs 10.0f

No effect on oxygen requirements at 16-19 ~ Habitat requirement for survival Survived and grew

0-30.0 h

33.8j

Hybrid Striped Bass Fingerlings

8.0 b

0-35.0 c 35.0 c

Optimal survival during confinement-induced stress No effect on growth Survival lower than in salinities of 28 %0 or lower

aPerry et al. (1977); bWeirich et al. (1992), sunshine bass, 25~ C; cSmith et al. (1988), palmetto bass, 23 ~ aGeiger and Parker (1985), survey of striped bass hatcheries; eMorgan et al. (1981); fChittenden (1971); sOtwell and Merriner (1975); hHall (1991), values based on review of literature; ~Mullis and Smith (1990); JLal et al. (1977)

10.3.5 Hardness and Calcium Water hardness is a measure of divalent cations (primarily calcium and magnesium) present in the water and is expressed as m g ~ hardness as calcium carbonate (Boyd, 1979). Calcium is probably the most important component of hardness with respect to striped bass culture due to its ability to reduce water flux across the gills. A reduced water flux lowers the amount of energy required to maintain proper water and ion balance during stress (see discussions in Weirich et al., 1992; Grizzle et al., 1993; Seals et al., 1994). Environmental calcium is also actively taken up by fishes via a calcium ATP-ase system located on the chloride cells of the gills (discussed and reviewed in Seals et al., 1994). Under normal (non-stressful) culture conditions actively feeding sunshine bass apparently derive most required calcium from their feed, even in low calcium (5 rag/L) water (Seals et al., 1994). Environmental calcium in these fish only becomes a factor during stress (Weirich et al., 1993). The calcium concentration in a water sample can be estimated (if little or no

259

magnesium is present) by dividing the water hardness by 2.5. Several studies have addressed the effect of environmental calcium on survival and growth of striped bass and its hybrids with mixed results. Age of the animals, other ions present, and presence of stressors seem to affect the efficacy of calcium in a given situation. The effects of calcium on fingerling production and post-harvest survival has been studied extensively in freshwater ponds. Increasing the water hardness of hatchery ponds did not increase fry survival or production (Reeves and Germann, 1972; Mauldin et al., 1988). However, increasing the calcium concentration in production ponds did increase post-harvest survival (Mauldin et al., 1988) even if the hardness was increased (from 20 to 45-100 mg/L as calcium carbonate) only five days before harvest (Grizzle et al., 1985). Weirich (Chapter 7) discusses the effects of environmental calcium on alleviation of handling and transport-induced stress. Environmental calcium concentrations (5-80 mg~) had no effect on survival, growth, or food conversion in juvenile sunshine bass (Seals et al., 1994). Based on the literature, Hall (1991) recommended hardness levels > 150 mg/L as calcium carbonate for prolarvae, larvae, and juveniles. This value appears to be in line with other formal and informal recommendations. Hardness can be increased in culture water by addition of several compounds such as calcium chloride and calcium sulfate. Calcium chloride has the further advantage of adding chloride, another physiologically active ion, to the culture water (see Chapter 3 and section 10.4.3). 10.3.6 pH and Alkalinity Maintaining an acceptable environmental pH is important in the culture of striped bass and its hybrids. Low environmental pH levels may contribute to the development of acidosis in culture animals and allow the buildup of carbon dioxide in culture water, which may have toxic effects on the fish. Acidic pond pH levels may also allow toxic substances found in pond muds, such as aluminum, to dissolve in the water and have toxic effects on the culture animals. Extremely high environmental pH levels will increase the toxicity of any ammonia present (see below). Development of low environmental pH is a problem in recirculating culture systems due to the consumption of buffering capacity by nitrifying bacteria in the biofilter. Routine additions of buffers, such as baking soda (sodium bicarbonate), will prevent pH decreases. Low alkalinity aquaculture ponds characteristically have large daily fluctuations in pH due to changing rates of photosynthesis by phytoplankton and respiration by all organisms during light and dark cycles. It is not unusual for nutrient rich, low alkalinity culture ponds in the southeastern United States to exhibit daily pH fluctuations from about 7 at sunrise to 10 or higher in late afternoon. Liming is the most common method of decreasing the daily fluctuation in ponds (Boyd, 1979). Liming increases alkalinity (buffeting capacity) which increases morning pH but attenuates the increase during the day, resulting in lower afternoon pHs that would occur in low alkalinity ponds. The effects of pH on striped bass and hybrid striped bass have been reviewed by Hall (1991) and Kerby (1993). Based on his review of the literature, Hall concluded prolarvae and larvae require a pH of 7.0-8.5 for survival. These values are similar to the recommendations by Bonn et al. (1976) and Mullis and Smith (1990) who suggest 7.5-8.5 is optimal and 6.5-9.0 is acceptable for striped bass and hybrid fry production. Hall (1991) also concluded that juveniles can survive over a pH range of 6.0-10.0. 10.3.7 Light Striped bass reproductive cycles are cued by changing environmental components such as day length. Chapter 2 considers the role of photoperiod in reproduction. Photoperiod and light intensity also have an

260

effect on nonreproducing fish. Palmetto bass larvae had higher survival rates in shaded aquaria and ponds than in aquaria and ponds not shaded (Rees and Cook, 1985). Striped bass larvae 5-23 days old exhibited hyperactivity in aquaria when exposed to 1,238 lux after 3 h in the dark (McHugh and Heidinger, 1978). Palmetto bass have been shown to grow faster in increasing photoperiods (spring-like conditions) in both fiberglass tanks (Woiwode and Adelman, 1991) and ponds (Kerby et al., 1987). 10.4 ENVIRONMENT RELATED NONINFECTIOUS DISEASES 10.4.1 General comments Noninfectious diseases are qualitatively different from infectious diseases in that noninfectious diseases have non-biotic primary causes while infectious diseases have biotic causes (viral, bacterial, or parasitic). Noninfectious diseases may have a genetic, nutritional or environmental basis. Below, I review some of the noninfectious disease agents common to striped bass and hybrid striped bass culture. The agents reviewed are what might be considered "endogenous" agents in that they are produced in the pond as a natural aspect of pond dynamics. "Exogenous" noninfectious disease agents such as therapeutants and pond treatments are reviewed in section 10.5. 10.4.2 Ammonia Ammonia is produced in fish during the catabolism of proteins and is excreted across the gills, primarily by diffusion. It is also produced in culture ponds during the bacterially mediated decomposition of dead plants and animals and uneaten fish feed. Normally, ammonia is oxidized to nitrite and then to nitrate by nitrifying bacteria present in ponds or biofilters of recirculating systems. However, ammonia concentrations often increase in heavily loaded culture systems due to either an overloading of the nitrification capacity of the system, or to inhibition of the nitrifying bacteria. High environmental ammonia concentrations may inhibit the outward diffusion of ammonia from the fish and, under the right circumstances, ammonia may actually diffuse into the fish. A buildup of plasma ammonia concentrations in culture animals may have toxic consequences. Tomasso (1994) reviewed ammonia toxicity in animals of interest to aquaculturists. "Ammonia" exists m two forms, the ionized ammonium (NH4*) and the un-ionized ammonia (NH3). To avoid confusion, when referring to both forms collectively, most authors use the term "total ammonia." Ammonia and ammonium exist in solution in an equilibrium determined primarily by pH and, to a lesser extent, by temperature (Emmerson et al., 1975). For example, at 20~ and pH values of 7, 8, and 9, ammonia present in the un-ionized form is 0.39, 3.81, and 28.36%, respectively. The pH dependency of this equilibrium is important because the un-ionized form is almost completely responsible for the toxicity of ammonia due to its ability to enter the fish by diffusion across the gill membrane. Conversely, ammonium cannot diffuse across gill membranes due to its charge. Consequently, knowing the total ammonia concentration of a culture system is of little value unless pH and temperature are also known. If all three values are known, the un-ionized ammonia concentration can be determined using the table in Emmerson et al. (1975). The 96-h median-lethal concentration (LCs0, the concentration of a poison that will kill 50% of the exposed animals in a set period of time)ofun-ionized ammonia to striped bass juveniles was 1.01+ 0.24 m g ~ ammonia-N (mean + SD) in fresh water (20-22 ~ 220 m~L total hardness as calcium carbonate, Oppenborn and Goudie, 1993). The same authors also reported the 96-h LC50 of un-ionized ammonia to palmetto bass to be 0.64+ 0.05 m~L. Weirich et al. (1993) reported the 96-h LCs0 of un-ionized ammonia to sunshine bass to be 0.32 mg/L in water containing 5 mg/L calcium and increasing to 0.60 mg/L in water containing 80 mg/L calcium. Mullis and Smith (1990) recommended that un-ionized ammonia concentrations not exceed 0.02 mg/L during striped bass and hybrid striped bass fry production.

261

10.4.3 Nitrite Nitrite is an intermediate product of the oxidation of ammonia to nitrate. Normally, very little nitrite is present in aquaculture systems. However, nitrite will occasionally accumulate in culture systems due to an imbalance in the nitrification process. Nitrite may accumulate in the blood of some fish species and, among other things, cause the oxidation of iron in hemoglobin producing methemoglobin, which is not capable of transporting oxygen. "Nitrite," like ammonia, exists in two forms, the ionized nitrite (NO2) and the molecular nitrous acid (HNO2). These two forms also exist in an equilibrium primarily determined by pH with higher pH values favoring the formation of nitrite. However, at pH values suitable for fish culture, almost all the total nitrite is in the ionized form. In many species, nitrite is actively pumped into the fish by the transport system that normally transports chloride into the fish (reviewed in Tomasso, 1994). In these species, the addition of chloride to the environment will competitively inhibit nitrite uptake and toxicity. Striped bass are relatively resistant to nitrite with a 24-h LCs0 of 50 mg/L nitrite-N (Mazik et al., 1991 a). The addition of chloride to the environment reduced nitrite-induced methemoglobinemia and plasma nitrite concentrations. Chloride added as calcium chloride was more effective in inhibiting methemoglobinemia and accumulation of plasma nitrite than chloride added as sodium chloride (discussed in Tomasso, 1994). For example, in striped bass exposed to 76 mg/L nitrite-N for 24 h, fish simultaneously exposed to 2,000 mg/L chloride as sodium chloride had plasma nitrite-N concentrations of about 15 mg/L, while fish protected with 2,000 mg/L chloride as calcium chloride had plasma nitrite-N concentrations of about 3 m ~ (Mazik et al., 1991a). Sunshine bass are not as resistant to nitrite as striped bass with a 96-h LCs0 of 12.8 m g ~ nitrite-N (Weirich et al., 1993). As with striped bass, calcium chloride is a more effective inhibitor of toxicity than sodium chloride. For example, a group of sunshine bass exposed to 35 mg/L nitrite-N for 72 h accumulated about 70 mg/L nitrite-N in their plasma, while a second group protected by calcium chloride accumulated only about 10 m g ~ nitrite-N. The 96-h LCs0was in excess of 100 mg/L nitrite-N in fish exposed in 8 g/L seawater (Weirich et al., 1993). 10.4.4 Suspended Sediments Water used in aquaculture production systems often becomes turbid due to suspended sediments, especially in brackish water systems. Sediments may become suspended in streams used for water sources after a rain, or ponds may become turbid following rain or due to mechanical aeration. Hatching percentage was not affected by suspended sediment up to 2,300 m g ~ (the highest concentration tested), but developmental rate was slowed by 890 mg/L (Morgan et al., 1983). Suspended solids of 200-500 mg/L reduced consumption of natural zooplankton prey assemblages (mostly copepods) by larvae by about 40% compared to larvae in 0-75 mg/L suspended solids (Breitburg, 1988). Suspended solids did not affect Daphniaconsumption (Breitburg, 1988). During forced swimming activity, juveniles exhibited reduced oxygen consumption rates in water containing river sediments compared to controls (Neumann et al., 1982). Based on a survey of the literature, Hall (1991) recommended prolarvae and larvae be held in water at or below 1,000 mg/L turbidity, and juveniles be held in water <10 g/L clay or silt and < 2 g/L fine grain sediments.

262

10.4.5 Gas and Oxygen Supersaturation Gas supersaturation occurs when the total dissolved gases in a body of water exceed the concentration of total gases that can be dissolved under normal circumstances given the temperature, dissolved solids, and gas pressure above the water (usually determined by altitude). Gas supersaturation is an inherently unstable state and usually leads to loss of excess gas in the form of bubbles. Gas supersaturation can occur when a gas saturated water sample is heated or when aeration occurs under pressure. Both instances may occur in fish hatcheries if water is heated immediately before being added to fish tanks or if air is trapped in high pressure water lines or pump heads. Gas supersaturation is a problem in fish culture because the gas will diffuse into the fish across the gills just as oxygen does and then may come out of solution in the plasma. The emboli formed can interfere with circulation and crush anatomical structures. High gas tensions can also cause overinflation of the swim bladder (Chamberlain et al., 1980). Striped bass larvae are sensitive to supersaturated water, causing intestinal bubbles, overinflation ofthe swim bladder, and death (Comacchia and Colt, 1984). Total gas pressures as low as 102.9% of saturation caused intestinal bubbles and an increase in swim bladder diameter in 10-d old larvae. Sensitivity decreased as larvae aged (Comacchia and Colt, 1984). Oxygen supersaturation occurs when more oxygen is dissolved in a sample of water than would be expected if the water were oxygenated using air. During oxygen supersaturation, oxygen displaces other dissolved gases (such as nitrogen). This allows the total gas pressure to remain at or below saturation so gas supersaturation is usually not a problem. Oxygen supersaturation may occur in a culture pond with high photosynthetic activity, but more commonly occurs when pure oxygen is used to oxygenate culture or transport tanks. Recently, a transient alkalosis following a hyperoxic experience has been observed in some species (reviewed in Wood, 1991). This has led some investigators to recommend that fish not be exposed to hyperoxic water. However, the practical consequences of hyperoxia have not been definitely established. Nevertheless, culturists using pure oxygen should strive to maintain oxygen concentrations no higher than saturation, just to be safe. 10.5 TOXICANTS 10.5.1 General comments During the course of aquaculture operations, fish are exposed to many chemicals, some advertently and some inadvertently. In this section, I review some of the more common ones. A few words of caution are in order here. The U.S. Food and Drug Administration (FDA) has recently become aggressive in labeling substances for use on food fish and enforcing regulations related to the use of chemicals on food fish. Before using a chemical in aquaculture operations, the culturist should determine the status of the chemical with the FDA. Several of the chemicals reviewed below have been commonly used on fish farms but presently cannot be used legally. Culturists should also be aware that water quality often affects the efficacy or toxicity of chemicals, and they should be very familiar with the specifics for a chemical or test it on a small number of fish before general application. Bills et al. (1993) recently published a comprehensive study of the effects of chemicals used in aquaculture on striped bass. 10.5.2 Anesthetics Anesthetics are commonly used to calm fish during handling and transport (see Chapter 7). However, they can kill the fish if too high a concentration is used. All presently used fish anesthetics are dissolved in

263

the water, and the anesthetic crosses the gills to cause the intended effect. The rate at which the drug cross the gills may be affected by specific water quality characteristics such as temperature, pH, and perhaps hardness. Consequently, it is very important to test an anesthetic on a small group of fish at each facility and for each water source before applying to large numbers of animals. Finquel (commonly referred to as MS-222) is the most common anesthetic in use today. Striped bass juveniles in soft water at 12~ had 96-h LCs0s of 28.2 and 49.0 mg/L determined in two separate locations (Bills et al., 1993). Juvenile striped bass were stressed after 15 min exposure to 25 mg/L MS-222 (a commonly used sedating concentration) at 21 ~ as indicated by elevated plasma corticosteroid concentrations (Davis et al., 1982). Other anesthetics used or tested for use on striped bass are listed in Table 10.4. The 96-h LCso for etomidate to striped bass fingerlings (23 ~ 48 mg/L hardness as calcium carbonate) was 0.68 mg/L (Plumb et al., 1983). Concentrations as low as 0.5 m g ~ caused mortality during a 24 h exposure. All fish that survived anesthesia at any concentration for 96 h recovered when placed in anesthetic-free water and were alive 24 h later (Plumb et al., 1983). Benzocaine has been shown to be an effective anesthetic in juvenile and mature striped bass at concentrations of 55 mg/L at 22~ and 80 mg/L at 11 ~ (Gilderhus et al., 1991). Fish survived exposure to up to twice the effective concentrations for 60 min, indicating a good margin of safety. 10.5.3 Algicides Aquaculture ponds usually become eutrophic due to the high level of nutrient input from uneaten food, fish excretions, and intentionally introduced fertilizers. High nutrient levels favor the production of high algal biomass. This biomass must be controlled or it may interfere with pond oxygenation, feeding and harvesting (Boyd, 1979). Copper sulfate and Aquazine are two commonly used algicides. Copper sulfate toxicity to larval and fingerling striped bass was tested at 21 ~ in fresh water containing 0.055 g/L sodium bicarbonate (Hughes, 1971). All fingerlings survived a 96-h exposure to 0.1 mg/L while all died in 0.25 mg/L. The 96-h LCso was determined to be 0.15 mg/L. All larvae survived a 96-h exposure to 0.05 mg/L while all died in 0.5 mg/L. The 96-h LCso was 0.1 mg/L. Increasing pH and alkalinity decreases copper toxicity (Tucker and Boyd, 1985). Unfortunately, pH was not reported in the Hughes (1971) study, and alkalinity must be estimated from the amount of sodium bicarbonate added to the test water. Another study on fingerlings at 21 ~ (pH = 8.2, total alkalinity - 64 as calcium carbonate) estimated the 96-h LCs0 to be 0.62 m g ~ (Wellborn, 1969). Bills et al. (1993) estimated the 96-h LCs0 of copper sulfate to juveniles (soft water, 12~ to be 0.23 mg/L. The 96-h LCsos of copper sulfate to fingerlings in dilute sea water (26~ were 2.68, 8.08, and 7.88 mg/L in 5,10, and 15 g ~ seawater, respectively (Reardon and Harrell, 1990). The authors suggested that tolerance increases as salinity increases up to the salinity to which the animals are acclimated, and then declines. Corneal damage was reported in larvae exposed to as little as 60 gg/L copper in 5 g/L sea water at 20~ (Bodammer, 1985). The resistance of striped bass to Simazine (also, Aquazine) appears to be high, but some conflicts exist in the literature. Wellborn (1969) reported that the 96-h LCs0 of Simazine to fingerlings (21 ~ C, pH = 8.2, total hardness = 35 mg/L as calcium carbonate, total alkalinity = 64 mg/L as calcium carbonate) to be 0.25 mg/L. However, McCann and Hitch (1980), following a design similar to Wellborn's, determined the 96-h LCs0 to be > 180 mg/L. McCann and Hitch suggested that differences in additives during formulation may explain the

264

Table 10.4. The 96-h median-lethal concentrations (LCs0) of selected anesthetics to striped bass juveniles a reported by Bills et al. (1993). If two values are presented for an anesthetic, tests were conducted in two locations. All tests were conducted in soft water at 12 ~ Anesthetic

LCs0 (m~JL)

Finquel (MS-222)

28.2 49.0

Benzocaine

28.1 20.0

Etomidate Metomidate

0.28 2.00

Quinaldine sulfate

22.4

differences. Bills et al. (1993) estimated the 96-h LCso to juveniles (soft water, 12~ to be 822 mg/L. A similar increase in tolerance with age was reported by Fitzmayer et al. (1985) who reported 48-h LCsos of 16-18 mg/L and >100 mg/L for 3 and 7-day old larvae, respectively (19-21 ~ hardness=120 and 220 mg/L as calcium carbonate). 10.5.4 Formalin Formalin is a commonly used therapeutant. Its uses include treatment of fungal and ectoparasitic infections. Bills et al. (1993) reported a 96-h LCso to juveniles of 75 ~zL/L in soft water at 12~ and 30 ~zL/L in soft water at 22 ~ At 21 ~ (total hardness=35 mg/L as calcium carbonate, pH=8.2) fingerlings had a 96-h LCs0 of 18 mg/L (Wellborn, 1969). In seawater (26~ fingerlings had 96-h LC5o s of 5.0, 13.5, 15.5, and 10.8 mg/L in 0, 5, 10, and 15 g/L salinity, respectively (Reardon and Harrell, 1990). 10.5.5 Chlorine Chlorine is a common disinfectant used on aquaculture operations, and it is also a normal additive to tap water. Consequently, aquaculture animals may come in contact with it if tanks and nets are not thoroughly rinsed after disinfection, or if tap water is used to fill holding tanks without properly neutralizing the chlorine. All larvae survived a 96-h exposure (21 ~ to 0.2 mg/L HTH (a commercial preparation of calcium hypochlorite). All died after exposure to 0.7 mg/L and the 96-h LCs0 was estimated to be 0.5 mg/L (Hughes, 1971). Fingerlings survived a 96-h exposure to 0.2 mg/L, but all fish exposed to 0.3 mg/L died. The 96-h LCso for fingerlings was estimated to be 0.25 mg/L (Hughes, 1971). Bills et al. (1993) reported the 96-h LC 5o of HTH to juveniles (soft water, 12 C) to be 0.35 mg/L.

265

10.6 CONCLUDING REMARKS Successful production of striped bass and its hybrids requires that the culturist understand the environmental requirements and recognizes the noninfectious disease agents of the species and its hybrids. The relationships between environmental needs and the energetics of growth, and between stress and disease resistance require careful environmental management. Poultry, beef and pork producers have long recognized these relationships and combined environmental with nutritional management to maximize production and minimize infectious disease outbreaks. Aquaculturists would do well to follow their example. This is especially true with respect to prevention of infectious diseases. Presently, few vaccines and therapeutants are available for use on fishes and the future does not look bright. The high costs of developing and labeling medicines coupled with the difficulty of delivering the medicines effectively to large numbers of relatively low-value anirrlals in an aquatic environment places limitations on what can be expected. Presently, the most effective way to deal with infectious disease is through prevention of which careful environmental management is an integral part.

266

References

Adams, S.M., editor, 1990. Biological indicators of stress in fish. American Fisheries Society, Bethesda, Maryland. Barton, B.A. and Iwama, G.K., 1991. Physiological changes in fish from stress in aquaculture with emphasis on the response and effects of corticosteroids. Annual Review of Fish Diseases, 1: 3-26. Bills, T.D., Marking, L.L. and Howe, G.E., 1993. Sensitivity of juvenile striped bass to chemicals used in aquaculture. U.S. Fish and Wildlife Service Resource Publication 192, Washington, D.C. Bodammer, J.E., 1985. Corneal damage in larvae of striped bass Morone saxatilis exposed to copper. Transactions of the American Fisheries Society, 114: 577-583. Bonn, E.W., Bailey, W.M., Bayless, J.D., Erickson, K.E., Stevens, R.E., editors, 1976. Guidelines for striped bass culture, Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, Maryland. Boyd, C.E., 1979. Water quality in warmwater fish ponds. Alabama Agricultural Experiment Station, Auburn. Breitburg, D.L., 1988. Effects of turbidity on prey consumption by striped bass larvae. Transactions of the American Fisheries Society, 117: 72-77. Brewer, D.L. and Rees, R.A., 1990. Pond culture of phase I striped bass f'mgerlings. Pages 99-120 in R.M. Harrell, J.H. Kerby, and R.V. Minton., editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, Maryland. Cech, Jr., J.J., Mitchell, S.J. and Wragg, T.E., 1984. Comparative growth ofjuvenile white sturgeon and striped bass: effects of temperature and hypoxia. Estuaries, 7:12-18. Chamberlain, G.W., Neill, W.H., Romanowsky, P.A. and Strawn, K., 1980. Vertical responses of Atlantic croaker to gas supersaturation and temperature change. Transactions of the American Fisheries Society, 109: 737-750. Chittenden, M.E., 1971. Effects of handling and salinity on oxygen requirements of the striped bass, Morone saxatilis. Journal of the Fisheries Research Board of Canada, 28:1823-1830. Comacchia, J.W. and Colt, J.E., 1984. The effects of dissolved gas supersaturation on larval striped bass, Morone saxatilis (Walbaum). Journal ofFish Diseases, 7: 15-27. Coutant, C.C. and Carroll, D.S., 1980. Temperatures occupied by ten ultrasonic-tagged striped bass in freshwater lakes. Transactions of the American Fisheries Society, 109: 195-202. Coutant, C.C., 1985. Striped bass, temperature, and dissolved oxygen: a speculative hypothesis for environmental risk. Transactions of the American Fisheries Society, 114:31-61. Cox, D.K. and Coutant, C.C., 1981. Growth dynamics ofjuvenile striped bass as functions of temperature and ration. Transactions of the American Fisheries Society, 110: 226-238. Davies, W.D., 1973. Rates of temperature acclimation for hatchery reared striped bass fry and fingerlings. The Progressive Fish-Culturist, 35: 214-217. Davis, K.B., Parker, N.C. and Suttle, M.A., 1982. Plasma corticosteroids and chlorides in striped bass exposed to tricaine methanesulfonate, quinaldine, etomidate, and salt. The Progressive Fish-Culturist, 44: 205-207.

267

Davis, K.B. and Parker, N.C., 1990. Physiological stress in striped bass: Aquaculture, 91: 349-358.

effect of acclimation temperature.

Ellis, A.E., 1981. Stress and modulation of defense mechanisms in fish. Pages 147-169 in A.D. Pickering, editor. Stress and fish. Academic Press, London. Emmerson, K., Russo, R.C., Lund, R.E. and Thurston, R.V., 1975. Aqueous ammonia equilibrium calculations: effect of pH and temperature. Journal of the Fisheries Research Board of Canada, 32: 2379-2383. Fitzmayer, K.M., Geiger, J.G. and Van Den Avyle, M.J., 1985. Acute toxicity effects of simazine on Daphnia pulex and larval striped bass. Proceedings of the Annual Conference Southeastern Association of Fish and Wildlife Agencies, 36: 146-156. Fry, F.E.J., 1947. Effects of environment on animal activity. University of Toronto Studies, Biological Series, 55: 1-62, Toronto. Fry, F.E.J., 1971. The effect of environmental factors on the physiology of fish. Pages 1-98 in W.S. Hoar and D.J. Randall, editors. Fish physiology, volume VI. Academic Press, New York. Geiger, J.G. and Parker, N.C., 1985. Survey of striped bass hatchery management in the southeastern United States. The Progressive Fish-Culturist, 47: 1-13. Gilderhus, P.A., Lemm, C.A. and Woods, III, L.C., 1991. Benzocaine as an anesthetic for striped bass. The Progressive Fish-Culturist, 53: 105-107. Grizzle, J.M., Mauldin, A.C. II, Young, D. and Henderson, E., 1985. Survival of juvenile striped bass (Morone saxanlis) and Morone hybrid bass (Morone chrysops XMorone saxatilis) increased by addition of calcium to soft water. Aquaculture, 46: 167-171. Grizzle, J.M., Cummins, K.A. and Ashfield, C.J., 1993. Effects of environmental concentrations of calcium and sodium on the calcium flux in stressed 34-day-old striped bass. Canadian Journal of Zoology, 71: 1379-1384. Hall, Jr., L.W., 1991. A synthesis of water quality and contaminants data on early life stages of striped bass, Morone saxatilis. Reviews in Aquatic Sciences, 4:261-268. Harrell, R.M. and Bayless, J.D. 1984. Effects of suboptimal dissolved oxygen concentrations on developing swiped bass embryos. Proceedings of the Annual Conference Southeastern Association ofFish and Wildlife Agencies, 35: 508-514. Harrell, R.M., Meritt, D.W., Hochheimer, J.N., Webster, D.W. and Miller, W.D., 1988. Overwintering success of swiped bass and hybrid swiped bass held in cages in Maryland. The Progressive Fish-Culturist, 50:120-121. Harrell, R.M., 1992. Stress mitigation by use of salt and anesthetic for wild striped bass captured for brood stock. The Progressive Fish-Culturist, 54: 228-233. Harrell, R.M. and Moline, M.A., 1992. Comparative stress dynamics of brood stock striped bass Morone saxatilis associated with two capture techniques. Journal of the World Aquaculture Society, 23: 58-63. Hill, L.G., Schnell, G.D. and Matthews, W.J., 198 I. Locomotor responses of the striped bass, Morone saxatilis, to environmental variables. American Midland Naturalist, 105: 139-148.

268

Hughes, J.S., 1971. Tolerance of striped bass Morone saxatilis (Walbaum) larvae and fingerlings to nine chemicals used in pond culture. Proceedings of the Southeastern Association of Game and Fish Commissioners, 24: 431-435. Kellogg, R.L. and Gift, J.J., 1983. Relationship between optimum temperatures for growth and preferred temperatures for the young of four fish species. Transactions of the American Fisheries Society, 112: 424-430. Kerby, J.H., 1993. The striped bass and its hybrids. Pages 251-306 in R.R. Stickney, editor. Culture of nonsalmonid freshwater fishes (second edition). CRC Press, Boca Raton, FL. Kerby, J.H., Woods, L.C., and Huish, M.T., 1983. Pond culture of striped bass x white bass hybrids. Joumal of the World Aquaculture Society, 14:613-623. Kerby, J.H., Hinshaw, J.M., and Huish, M.T., 1987. Increased growth and production of striped bass x white bass hybrids in earthen ponds. Journal of the World Aquaculture Society, 18: 35-43. Lal, K., Lasker, R. and Klujis, A. 1977. Acclimation and rearing of striped bass in sea water. California Fish and Game, 63: 210. McCann, J.A., and Hitch, R.K., 1980. Simazine toxicity to fingerling striped bass. The Progressive Fish-Culturist, 42: 180-181. McHugh, J.J. and Heidinger, R.C., 1978. Effect of light shock and handling shock on striped bass fry. The Progressive Fish-Culturist, 40: 82. Mauldin, A.C. II, Grizzle, J.M., Young, D.E. and Henderson, H.E., 1988. Use of additional calcium in soft-water ponds for improved striped bass survival. Proceedings of the Annual Conference Southeastern Association of Fish and Wildlife Agencies, 40:163-168. Mazik, P.M., Hinman, M.L., Winklemarm, D.A., Klaine, S.J., Simco, B.A. and Parker, N.C., 1991a. Influence of nitrite and chloride concentrations on survival and hematological profiles of striped bass. Transactions of the American Fisheries Society, 120: 247-254. Mazik, P.M., Simco, B.A. and N.C. Parker., 1991b. Influence of water hardness and salts on survival and physiological characteristics of striped bass during and after transport. Transactions of the American Fisheries Society, 120: 121-126. Morgan, R.P. II, Rasin, V.J. Jr. and Copp, R.L., 1981. Temperature and salinity effects on development of striped bass eggs and larvae. Transactions of the American Fisheries Society, 110: 95-99. Morgan, R.P., II, Rasin, V.J., Jr. and Noe, L.A., 1983. Sediment effects on eggs and larvae of striped bass and white perch. Transactions of the American Fisheries Society, 112: 220-224. Mullis, A.W. and Smith, J.M., 1990. Design considerations for striped bass and striped bass hybrid hatching facilities, Pages 7-16 in R.M. Harrell, J.H. Kerby, and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Neumann, D.A., O'Connor, J.M., Sherk, Jr., J.A. and Wood, K.V., 1982. Respiratory response of striped bass (Morone saxatilis) to suspended solids. Estuaries, 5: 28-39. Oppenbom, J.B. and Goudie, C.A., 1993. Acute and sublethal effects of ammonia on striped bass and hybrid striped bass. Journal of the World Aquaculture Society, 24: 90-101.

269

Otwell, W.S. and Merriner, J.V., 1975. Survival and growth ofjuvenile striped bass, Morone saxatilis, in a factorial experiment with temperature, salinity and age. Transactions of the American Fisheries Society, 104: 560-566. Perry, W.G., Carver, D.C. and Williams, A.M., 1977. Brackish water culture of striped bass in Louisiana. Proceedings of the World Aquaculture Society, 8:107-114. Pickering, A.D., editor., 1981. Stress and fish. Academic Press, New York. Pickering, A.D., 1993. Endocrine-induced pathology in stressed salmonid fish. Fisheries Research, 17: 35-50. Plumb, J.A., Schwedler, T.E. and Limsuwan, C., 1983. Experimental anesthesia of three species of freshwater fish with etomidate. The Progressive Fish-Culturist, 45: 30-33. Reardon, I.S. and Harrell, R.M., 1990. Acute toxicity of formalin and copper sulfate to striped bass fingerlings held in varying salinities. Aquaculture, 87: 255-270. Rees, R.A. and Cook, S.F., 1985. Effects of sunlight intensity on survival of striped bass x white bass fry. Proceedings of the Annual Conference Southeastern Association ofFish and Wildlife Agencies, 36: 83-94. Rees, R.A. and Harrell, R.M., 1990. Artificial spawning and fry production of striped bass and hybrids. Pages 43-72 /n R.M. Harrell, J.H. Kerby, and J.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division, American Fisheries Society, Bethesda, MD. Reeves, W.C. and Germann, J.F., 1972. Effects of increased water hardness, source of fry and age at stocking on survival of striped bass fly (sic) in earthen ponds. Proceedings of the Annual Conference Southeastern Association of Game and Fish Commissioners, 25: 542-548. Schreck, C.B., 1981. Stress and compensation in teleostean fishes: responses to social and physical factors. Pages 295-321 in A.D. Pickering, editor, Stress and fish. Academic Press, New York. Seals, C., Kempton, C., Tomasso, J. and Smith, T., 1994. Environmental calcium does not affect production or selected blood characteristics of sunshine bass reared under normal culture conditions. The Progressive Fish-Culturist, 56: 269-272. Shannon, E.H. and Smith, W.B., 1968. Preliminary observations of the effect of temperature on striped bass eggs and sac fry. Proceedings of the Annual Conference Southeastern Association Game and Fish Commissioners, 21: 257-260. Smith, T.I.J., Jenkins, W.E. and Haggerty, R.W., 1988. Growth and survival ofjuvenile striped bass (Morone saxatilis) x white bass (M. chrysops)hybrids reared at different temperatures. Proceedings of the Annual Conference Southeastern Association of Fish and Wildlife Agencies, 40:143-151. Snieszko, S.F., 1974. The effects of environmental stress on outbreaks of infectious diseases of fishes. Joumal ofFish Biology, 6: 197-208. Spotte, S., 1979. Fish and invertebrate culture. Wiley-Interscience, New York. Strange, R.J. and Cech, Jr., J.J., 1992. Reduced swimming performance of striped bass after confinement stress. Transactions of the American Fisheries Society, 121:206-210. Tisa, M.S., Strange, R.J. and Peterson, D.C., 1983. Hematology of striped bass in fresh water. The Progressive Fish-Culturist, 45:41-44.

270

Tomasso, J.R., Davis, K.B. and Parker, N.C., 1980. Plasma corticosteroid and electrolyte dynamics of hybrid striped bass (white bass x striped bass) during netting and hauling. Proceedings of the World Mariculture Society, 11: 303-310. Tomasso, J.R., 1994. Toxicity of nitrogenous wastes to aquaculture animals. Reviews in Fisheries Science, 2:291-314. Tucker, C.S. and Boyd, C.E., 1985. Water quality. Pages 135-227 in C.S. Tucker, editor. Channel catfish culture, Elsevier, New York. Turner, J.L. and Farley, T.C., 1971. Effects of temperature, salinity, and dissolved oxygen on the survival of striped bass eggs and larvae. California Fish and Game, 57: 268-273. Van Den Avyle, M.J. and Evans, J.W., 1990. Temperature selection by striped bass in a Gulf of Mexico coastal river system. North American Journal of Fisheries Management, 10: 58-66. Wedemeyer, G., 1970. The role of stress in the disease resistance of fishes. Pages 30-35 in S.F. Snieszko, editor. Diseases of fishes and shellfishes. American Fisheries Society, Bethesda, MD. Wedemeyer, G.A., Barton, B.A. and McLeay, D.J., 1990. Stress and acclimation. Pages 451-489 in C.B. Schreck, and P.B. Moyle, editors. Methods for fish biology. American Fisheries Society, Bethesda, MD. Weirich, C.R., Tomasso, J.R. and Smith, T.I.J., 1992. Confinement and transport-induced stress in white bass Morone chrysops x striped bass M. saxatilis hybrids: effect of calcium and salinity. Journal of the World Aquaculture Society, 23: 49-57. Weirich, C.R., Tomasso, J.R. and Smith, T.I.J., 1993. Toxicity of ammonia and nitrite to sunshine bass in selected environments. Journal of Aquatic Animal Health, 5: 64-72. Wellbom, Jr., T.L., 1969. The toxicity of nine therapeutic and herbicidal compounds to striped bass. The Progressive Fish-Culturist, 31: 27-32. Woiwode, J.G., 1989. The effects of temperature, photoperiod and ration size on the ~owth and thermal resistance of the hybrid striped x white bass. Ph.D. Dissertation, University of Minnesota, Saint Paul. Woiwode, J.G. and Adelman, I.R., 1984. Growth, food conversion efficiency, and survival of the hybrid white x striped bass as a function of temperature. Pages 143-150 in the Aquaculture of striped bass: a proceedings, J.P. McCraren, editor. University of Maryland Sea Grant Publication, UM-SG-MAP-84- 01, College Park. Woiwode, J.G. and Adelman, I.R., 1991. Effects of temperature, photoperiod, and ration size on growth of hybrid striped bass x white bass. Transactions of the American Fisheries Society, 120:217-229. Wood, C.M., 1991. Branchial ion and acid-base transfer in freshwater teleost fish: environmental hypoxia as a probe. Physiological Zoology, 64: 68-102. Young, P.S. and Cech, Jr., J.J., 1993. Physiological stress responses to serial sampling and confinement in young-of-the-year striped bass, Morone saxatilis (Walbaum). Comparative Biochemistry and Physiology, 105A: 239-244.

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

271

Chapter11 Infectious Diseases of Striped Bass John A. Plumb

11.1 INTRODUCTION The common disease causing organisms associated with most aquaculture fish species also occur in cultured striped bass and their hybrids. There seems to be no difference in disease susceptibility between the pure strains of striped bass and their hybrids, therefore, generally no distinction is made between them in this text. With the intensification of rearing methods, and recent expansion of striped bass culture, the list of known infectious diseases of the species has not increased greatly but consequences of disease have become more acute. In some instances, infectious diseases have limited striped bass culture, and in others they have significantly contributed to the demise of their production. Infectious agents that cause disease of striped bass include viruses, bacteria, fungi, protozoa, and metazoan parasites. Except for viruses and helminthic parasites, only a few of these agents are truly obligate pathogens while most are saprophytic, facultative, or opportunistic, and precipitate a disease only after the striped bass are stressed, injured, or otherwise debilitated. There are several reviews on infectious diseases of striped bass but none of these have been comprehensive (Bonn et al., 1976; Sinderman and Lightner, 1988; Hughes et al., 1990; Plumb, 1991). 11.2 PREDISPOSING FACTORS Because of a dearth of therapeutic compounds for diseases of striped bass, it is essential that the aquaculturist be aware of the many predisposing factors to infectious diseases of striped bass that are inherent in the cultural system. Those stressful conditions that exist in most aquaculture environments also exist in striped bass culture. Striped bass and their hybrids are very susceptible to environmental stress, which is often followed by infectious disease; therefore, the best defense against infectious diseases is prevention by following good aquaculture practices and health management. Health management includes all phases of the cultural process, emphasizing maintenance of good water quality, using optimum stocking densities, adequate availability of feed and good nutrition, maintaining optimum temperature and practicing proper fish handling procedures (Plumb, 1994). Prophylactic treatments when appropriate, and judicious, timely use of chemotherapy are also a part of the health management strategy. For a detailed discussion on optimum environmental requirements for striped bass and its hybrids the reader is referred to Chapter 10. Although striped bass and their hybrids grow over a wide range of water quality characteristics, it is essential to maintain the best water quality possible during all phases of the culture cycle, thus reducing the potential of environmental stress, which increases disease susceptibility. In general, oxygen concentrations should remain above 3 mg/L. Striped bass can survive oxygen levels as low as 1 m~q~, but extended exposure to this concentration is highly stressful (Hill et al., 1989). Stressed fish become inactive and lose equilibrium, which may precipitate increased disease susceptibility or death. Low alkalinity (less than 20 to 30 mg/L) may cause striped bass to be less hardy and more prone to handling stress and increased infectious disease susceptibility. Grizzle et al. (1992) showed that survival of 20- to 30-day-old striped bass fry held in extremely soft water (<10 mg/L as CaCO3) could be significantly improved by increasing the calcium concentration in the water (Figure 11.1).

272

A

lOO 80

,~

8

60

0 >

= 40

(/1

C 2O

O,

i

Hatchery Water

!

Calcium Chloride

|

Sodium Chloride

|

Both

Fig. 11.1. Survival of 20- to 30-day-old striped bass fry in water with less than 10 mgft~ C a C O 3 and in the same water with calcium chloride and sodium chloride added (Reprinted with permission from Grizzle et al. 1992). During striped bass handling, avoiding injury to the skin, fins, mucus layer, or scales will help the fish withstand secondary infections by opportunistic organisms. For example Flexibacter columnaris often develops on the fins of striped bass after being seined and/or handled. A discussion of the transportation, stress, and water quality values affecting striped bass is presented in Chapter 7. 11.3 VIRUS DISEASES No known virus diseases are highly pathogenic, or are of importance to striped bass, however, the striped bass aquareovirus may be specific for the species or their hybrids. Two other viruses (lymphocystis virus, and infectious pancreatic necrosis virus) are known to infect striped bass in cultured and possibly natural environments, or have been experimentally transmitted to striped bass (Table 11.1). 11.3.1. Lymphocystis Virus Lymphocystis virus disease is widely spread, and it infects a diverse range of fish species, genera, and family including all members of the Percichthyidae. It was first reported in striped bass in 1950 at the New York Aquarium (Nigrelli, 1950), and since has occasionally been reported in wild and cultured striped bass. The incidence of the disease appears to be low and infrequent. Lymphocystis virus is yet to be reported in hybrid striped bass, but it is unlikely that they are resistant because neither the striped bass nor the white bass (principal species in the hybridization) are resistant. Lymphocystis virus does not cause severe pathology, debilitating disease, or death in striped bass. While not producing a typical viremia, lymphocystis virus manifests itself by producing large, benign, hypertrophied cells forming transient lesions on the skin and fins (Figure 11.2). Infected cells appear in grayish or cream colored clusters on the skin and fins that slough from the surface after a period of time. Virus

273

Table 11.1. Viruses isolated from striped bass. Virus disease

Effect on Striped bass

Lymphocystis virus

Typical surface growths

Infectious pancreatic necrosis virus

Inapparent infections

Striped bass aquareovirus

Inapparent infections

infected cells may be 100 to 1,000 times larger than the 5 to 10 ~m diameter of normal cells. A thick hyaline sheath formes a matrix around infected cells. These cells contain an enlarged, irregularly shaped nucleus that is generally centrally located. Ribbon shaped, basophilic, Feulgen positive, cytoplasmic inclusions give a positive DNA reaction, and are thought to be sites of viral replication. Fish heal after sloughing the lesion, leaving no evidence of the infection, however, these cells disintegrate releasing virus into the water, thus providing a source for infection of other individuals. Lymphocystis virus belongs to the genus Cystivirus, within the family Iridoviridae (Flugel, 1985). Size of the large icosahedral, unenveloped, DNA virus varies from 130 nm to 330 nm. The virus may be isolated from the lesions in cyprinid cells lines.

Fig. 11.2. Lymphocystis virus. (A) Lymphocystis lesions (arrows) on striped bass (photo courtesy of A. Baya). (B) Lymphocystis virus infected cells showing the enlarged nucleus (arrow).

274

Increased susceptibility of other fish species to lymphocystis has been associated with injury to the epithelium. Although a controversial issue, increased environmental pollution has occasionally been associated with increased susceptibility of fish to lymphocystis, and this could be critical in striped bass because of their sensitivity to pollutants. Lymphocystis usually occurs during cool or cold weather at which time the disease seems to persist longer than during warm weather. The lesions are unsightly and may cause rejection of infected fish by anglers and the consumer. 11.3.2. Infectious Pancreatic Necrosis Virus Infectious pancreatic necrosis virus (IPNV) is normally associated with salmonids, however, this virus infects a wide variety of fish species, including striped bass, in the culture and natural environment (Wolf, 1988). Infectious pancreatic necrosis virus often causes high mortality in cultured juvenile salmonids, but it does not cause a disease problem in striped bass. The first reports of IPNV in striped bass occurred in Maryland where the virus was isolated from 4-week old fry originating from eggs taken from brood fish in the Chesapeake Bay (Schutz et al., 1984). The epizootic occurred at temperatures of 17.5 to 21 ~ and stopped when temperature reached 25 ~ Affected striped bass exhibited a darting, random swimming behavior but there were no external lesions. Histopathology of moribund fish showed only focal degeneration of the basal layer of the epidermis. These juvenile fish suffered high mortality in which about 2 million fish died, however, it was not conclusive that the virus actually caused their death. The IPNV isolated from the striped bass was serologically similar to IPN VR 299, the common strain that causes mortality in North American salmonids. Infectious pancreatic necrosis virus is a member of the Birnaviridae; it is icosahedral, and contains a double stranded RNA genome (Dobos et al., 1977). The virus is isolated in several different cell lines including CHSE-214, FHM and others. Wechsler et al. (1987) showed that 26 to 180-day old striped bass were not killed by IPNV after experimental water borne exposure or infection by the oral or intraperitoneal injection route. However, naturally and experimentally infected striped bass did actively shed IPNV (without significant mortality) and became carriers for at least 14 months after exposure. Wechsler et al. (1987) further showed that IPNV was not transmitted to the next generation via gametes from carrier adult striped bass, or by exposing eggs to virus, therefore, the means of transmission from generation to generation of striped bass is not clear. McAllister and McAllister (1988) did show that IPNV carrier striped bass could transmit the virus to brook trout (Salvelinus fontinalis) via contaminated water flowing from the infected striped bass to the trout and resulted in mortality of the brook trout. Cultured fish are of most concem when considering the presence oflPNV, but Wechsler et al. (1987) detected IPNV neutralizing antibody in the sera of wild striped bass caught in the Chesapeake Bay, therefore, it is possible that wild fish do become carriers following exposure to the virus. Toranzo et al. (1983) showed that IPNV can survive for nearly six weeks in estuarine water, so if carrier fish are present, there is ample time for non-infected fish to contract IPNV. Although IPNV has no pathogenic effects on striped bass, care should be taken to prevent possible IPNV carrier striped bass from coming in contact with more susceptible species of fish, brook trout for example. 11.3.3. Striped Bass Aquareovirus Since the late 1970s there have been numerous viruses isolated from fish that have been classified as members of the Reoviridae (Winton et al., 1987). These viruses are unique enough to be considered in a specified genus, Aquareovirus (Holmes, 1991). Baya et al. (1990a) described a reo-like virus from striped bass, however, it could not be clearly demonstrated that it was a major factor in mortality. The fish showed

275

hemorrhagic lesions along the dorso-lateral portions of the body where scales were missing. Hemorrhages were also found in the swim bladder and the liver was pale, enlarged, and mottled. A bacterium, Carnobacterium piscicola, was also isolated from these fish, thus confusing the etiology. Additional characterization of the reo-like virus indicated differences from other reoviruses especially the possession of an RNA genome with 11 segments of double stranded RNA instead of the usual 10 segments. Baya et al. (1990a) named it the striped bass rotavirus (SBR) since the rotaviruses have 11 segments. This virus was thought to represent a new genus within the family Reoviridae, and Samal et al. (1990) suggested the name Aquarotavirus. As more information on the molecular nature of the virus became known, it was clear that SBR should be grouped with the other "aquareovirus" and not separated into a new genus. The RNA electrophoretic patterns are similar to those ofaquareoviruses isolated from four other species of marine fishes from North America and Europe, between which, Samal et al. (1991) found no genetic differences. Striped bass aquareovirus can be isolated from actively infected fish in CHSE-214 cells where it produces a well defined syncytium in two days at 15 ~ and in three days at 20~ total cell sheet destruction occurs in 6 and 13 days respectively (Dopazo et al., 1991). Subramanian et al. (1993) developed a nucleic acid hybridization probe that rapidly detected small quantities of SBR in fish tissue and cell cultures. The effects of SBR on cultured striped bass is not clear at the present time, however, the aquareovirus group is considered to have low virulence and produce little pathology and this virus seems to be no different. 11.4 BACTERIAL DISEASES Bacterial diseases cause some of the most serious problems for cultured striped bass (Table 11.2). Most bacteria that infect striped bass, with a couple of exceptions, are saprophytic, facultative, and opportunistic organisms that often cause debilitating infections following the previously discussed predisposing factors. None of the diseases are unique to striped bass, but species in several genera, namely Streptococcus, Mycobacterium, and Pasteurella, are generally more serious in striped bass than they are in other cultured fishes (Plumb, 1991; J.P. Hawke, Louisiana State University, personal communication). The opportunistic pathogens include species of A eromonas, Pseudomonas, Vibrio, Edwardsiella, and Flexibacter. Other bacteria may also occasionally cause infections in striped bass under certain circumstances. With the exception of lesions caused by F. columnaris, there is very little difference in the gross, external appearance of striped bass infected with most of these bacteria. 11.4.1. Motile Aeromonas Septicemia Motile Aeromonas septicemia (MAS), a disease of many freshwater fish species, is associated with infections caused by the motile members of the genus Aeromonas (Austin and Austin, 1987). Also, several species ofPseudomonas, P. fluorescens primarily, cause the same type of disease syndrome in striped bass as MAS. The disease is also called hemorrhagic septicemia. Striped bass afflicted with MAS will stop or slow their feeding activity, and swim lethargically at the surface. Clinical signs are not specific. Fish may develop mild to severe hemorrhage in the skin and fins which also have pale frayed margins (Figure 11.3). Scales may protrude (lepidorthosis) as result of fluid (edema) in the scale pockets and fluid in or behind the eye causes exophthalmia. Scales may slough in severe cases, and the skin becomes necrotic and exposes underlying musculature to water and potential pathogens therein. Gills are often pale. In systemic infections a cloudy, bloody fluid is often present in the body cavity, and internal organs may be hyperemic or pale depending on the stage of infection. The causative agent of MAS can be one of several different organisms including Aeromonas hydrophila (punctata, liquefaciens) and A. sobria, however, in striped bass A. hydrophila is the most common

276

Fig 11.3. Motile Aeromonas septicemia (Aeromonas hydrophila) and related species. (A) Necrotic and hemorrhaged skin lesion (arrow) on wild adult striped bass (printed with permission of CRC Press). (B) Hemorrhagic septicemia lesions (arrow) on striped bass (photo courtesy of A. Baya).

(Plumb, 1991). The number of different species of bacteria in the motile aeromonads is rapidly expanding due to taxonomic reclassification and molecular identification techniques (Carnahan, 1993a), but how many of these new species cause disease in striped bass is unknown. The motile aeromonads can be isolated from skin and muscle lesions or internal organs of infected fish on brain heart infusion (BHI) agar or tryptic soy agar (TSA) incubated at 25 to 35~ In addition to uniform motility, all are gram-negative, short rods that measure about 0.8 to 0.9 X 1.5 , m , polar flagellated, cytochrome oxidase positive, oxidative and fermentative in glucose media, and resistant to the vibriostat 0/129 (Shotts and Teska, 1989) (Table 11.3). Aeromonas hydrophila forms a yellow-orange colony in 24 to 48 h on Rimler-Shotts media when incubated at 35~ (Shotts, 1991). Pseudomonasfluorescens is weakly motile, oxidative only in glucose media and produces soluble fluorescent pigment that is visible on BHI or TSA media. The bacterial species comprising the MAS are generally detected by isolation on media and identification by conventional bacteriological procedures (Shorts and Teska, 1989). Implementation of rapid identification systems (API-20E and Minitek) for positive identification is increasing (Carnahan, 1993b). This group of organisms is antigenically diverse, therefore, molecular and serological identification methods are generally not used in the diagnosis of MAS in fish. However, DNA polymerase chain reaction (PCR) has been used to detect virulent A. hydrophila strains in other species offish (Baloda et al., 1995), and an enzyme-linked immunosorbent assay was developed to detect A. hydrophila in different foods (Merino et al., 1993).

277

The motile aeromonads constitute a ubiquitous group of bacteria commonly found in most waters that contain any organic material from which they can derive nutrients (Hazen et al., 1978). These bacteria seldom precipitate disease unless the host is debilitated by some stressful condition that allows the normally non-pathogenic bacteria in the water to become established on, or in the host and results in either a skin or systemic infection. In view of this, as long as the striped bass are not stressed and are in good condition they can usually resist exposure to the motile aeromonads. High stocking densities, improper handling resulting in injury to the mucous layer or skin, scale loss, temperature shock, and poor water quality are the principal predisposing factors to infection. Extended periods of exposure to low oxygen (less that 3 mg/L) will reduce the resistance of striped bass to bacterial infections including MAS. Motile Aeromonas septicemia may also be secondary to other less debilitating disease organisms, for example protozoan parasites. The disease has been documented in all types of culture facilities, including intensive and semi-intensive systems, and in freshwater and occasionally brackish water environments with salinities up to 15 ppt.

Table 11.2. Bacterial diseases of striped bass.

Disease

Causative agent

Severity

Motile Aeromonas septicemia (MAS)

Aeromonas hydrophila A. sobria

Moderate

Pseudomonas septicemia

Pseudomonas fluorescens Pseudomonas sp.

Low

Columnaris

Flexibacter columnaris F. maritimus

Moderate to high Unknown

Pasteurellosis

Pasteurella piscicida

High

Edwardsiellosis

Edwardsiella tarda

Low

Vibriosis

Vibrio anguillarum

Moderate

Enterococcosis

Enterococcusfaecium

Moderate

Streptococcosis

Streptococcus sp.

Moderate

Mycobacteriosis

Mycobacterium marinum

High

Camobacteriosis

Carnobacterium piscicola

Low

Corynebacteriosis

Corynebacterium aquaticum

Low

278

Motile Aeromonas septicemia has no geographical restriction other than its propensity for the freshwater environment. In most fishes, the disease does occur seasonally with increased incidence in spring and early summer and again in the fall with low incidence in the winter and late summer (Plumb et al., 1990), however, specific seasonal mortality data are not available for striped bass. Mortality associated with MAS is usually chronic, rarely becoming acute, but with its chronic nature, significant numbers of fish may die over a long period of time. The disease may affect juveniles, market size fish, or adults. 11.4.2. Columnaris Columnaris is an acute to chronic infection of a variety of species of fish including striped bass. The causative agent of columnaris in freshwater is Flexibacter columnaris. This organism has, at one time or another, also been called Chondrococcus columnaris and recently Cytophaga columnaris. However, using DNA relatedness, Bernardet and Grimont (1989) demonstrated that the aetiology of columnaris should be in the Flexibacter genus rather than Cytophaga, therefore Flexibacter columnaris is used in this text. Columnaris occurs in both cultured, and probably wild striped bass, but it is much more severe in cultured populations. Flexibacter maritimus is the "columnaris" equivalent in brackish and salt water (Wakabayashi et al., 1986). Although there is no published record of F. maritimus infecting striped bass in salt water, there is no reason to believe it cannot. Columnaris is usually confined to the skin, fins, and/or gills, and occasionally it occurs systemically. Whitish areas appear on the skin, while lost scales often expose the underlying musculature (Figure 11.4). Fins are usually white and are in various stages of fraying. Necrotic lesions on the gills appear pale compared to normal gills (Figure I 1.4). The margins of lesions may be yellowish because ofthe presence of a large number of the bacteria. Pathology of internal organs in systemic infections is not dramatic. Other organisms such as Aeromonas sp., fungi, or protozoan parasites may also be present in columnaris lesions. Columnaris is detected by recognition of typical lesions on diseased fish and identifying the long slender (0.5 X 4 to 10 ~zm) rod shaped cells in wet mounts made from these lesions (Table 11.3). Upon Gram staining, F. columnaris is gram negative. The cells are motile by flexing or gliding and will form "hay stacks" or columns on wet mounts. Flexibacter columnaris does not grow on conventional laboratory media, but requires a low nutrient, moist medium such as Ordal's (Anacker and Ordal, 1959) or Hsu-Shotts (Shotts, 1991). Cultures should be incubated at 25 to 30~ where discrete, spreading, rhizoid colonies with irregular margins and yellow centers that adhere tightly to the media surface form in 24 to 48 h. Isolation is enhanced by the addition ofpolymyxin B (10 IU/mL) and neomycin (5~zg/mL) which inhibit growth ofnon-columnaris bacteria. Griffin (1992) described a procedure for presumptively identifying the yellow pigmented bacteria from fish, of which F. columnaris is a member, based on its ability to grow in the presence of neomycin sulfate and polymyxin B; forming yellow, fiat, rhizoid colonies; positive for gelatin degradation; binding of Congo red, and production of chondroitin lyase. Flexibacter columnaris can be specifically identified by serological agglutination using specific antisera. Although F. columnaris can survive in water and mud, fish are considered to be the reservoir of the pathogen. Columnaris is common among cultured fishes, usually affecting striped bass following handling or exposure to some other stressful condition such as seining, temperature shock, low oxygen, or crowding. On occasion it may be a primary pathogen. Columnaris can be a chronic infection in which a small number of fish are affected and mortality is comparatively low; contrastingly, the disease can be acute with explosive mortality reaching as high as 90%. Young fish are much more susceptible to columnaris than are older fish, but no age group is totally immune. Columnaris disease occurs at any time of the year, however, it is generally a seasonal problem being most prevalent during spring and fall.

279

Fig. 11.4. Columnaris (Flexibactercolumnaris). Early (upper arrow) and advanced (lower arrow) necrotic lesions on the gills of a largemouth bass. Lesion on striped bass would appear the same.

11.4.3 Pasteurellosis Pasteurellosis is an infrequent chronic to acute septicemia in several species of fish including striped bass and other members of the temperate bass. With increased intensification of striped bass culture, Pasteurellapiscicida infections have become more common in fish raised in brackish waters because of the bacteriums halophilic nature (Robohm, 1983). Following its first implication as the etiological agent of a massive die off of white perch (Morone americana) and a lesser number of striped bass in Chesapeake Bay (Snieszko et al., 1964), P. piscicida has been reported in wild striped bass in New York (Robohm, 1983) and cultured striped bass along the Gulf of Mexico coast of southeastern United States (Hawke et al., 1987). More recently acute infections have occurred among striped bass cultured in salt water ponds, cages, and net pens (J. P. Hawke, Louisiana State University, personal communication). The disease has not been documented in striped bass reared in fresh water. Pasteurellosis occurs most often in high density culture systems where water temperatures range from 22 to 30~ and oxygen concentrations are marginally low. External clinical signs of pasteurellosis in striped bass are rather indistinct. Infected fish lose locomotion and sink in the water column (Robohm, 1983). The skin becomes dark and some petechial

280

hemorrhages appear at the base of the fins and on the opercules. Internally the spleen and kidney are swollen and have white granulomas or miliary lesions giving rise to the name "pseudotuberculosis" (Figure 11.5). Cut sections of the spleen may show white patches (granulomas). Phagocytes become swollen and laden with bacteria; this is thought to contribute to death of infected fish. Mortalities due to pasteurellosis in striped bass may vary but Hawke et al. (1987) reported that 80% of infected cultured fingerling striped bass died during an epizootic in Alabama.

Pasteurellapiscicida survives less than three days in sterile brackish water, but the organism appears to be a normal inhabitant of the estuarine environment where other fish are most likely the natural host (Janssen and Surgalla, 1968). Pasteurellosis is detected by isolation of the organism on BHI agar with 2% salt and incubation at 20 to 30~ This facultative anaerobe does not grow at 37~ but there is disagreement on its ability to grow at 10~ (Robohm, 1983; Hawke et al., 1987). The organism is a gram-negative (stains bipolar), nonmotile, pleomorphic, non-encapsulated, bacillus ranging in size from 0.6 to 0.7 by 1.2 to 2.6 ~zm. Isolates from the U. S. and Japan are morphologically, physiologically and biochemically similar (Table 11.3). 11.4.4. Edwardsiellosis

Edwclrdsiella tarda, while a common pathogen in some cultured fish species, only occasionally infects cultured striped bass. Herman and Bullock (1986) reported that E. tarda infected 4 to 5 cm fingerling striped bass cultured in freshwater. The fish became moribund, swam lethargically at the surface, displayed pale gills and had a slightly discolored area in the cranium. A pure culture of a gram-negative, cytochrome oxidase negative, motile rod, presumptively identified as Edwardsiella tarda was isolated from lateral line and kidney tissues (Table 11.3). Additional biochemical characteristics include production of indole from tryptone and an alkaline slant over an acid butt in triple sugar iron agar with production of H2S and gas. Naive striped bass were infected with this organism by water borne exposure. Histologically, the epithelium of experimentally infected fish was necrotic and the fins were frayed. Numerous abscesses were present in the kidney along with necrosis of the hematopoietic tissue of the trunk kidney. As the disease progressed, the kidney became grossly enlarged with massive inflammation, necrosis and a large number of bacteria being present. The lesion expanded to the adjacent trunk musculature. Recently, a significant E. tarda infection was found in wild adult striped bass in Chesapeake Bay (A. Baya, University of Maryland, personal communication). The most notable clinical signs in these fish were numerous irregular, coalescing, hemorrhagic areas on the skin and fins, with some ulcerations emitting an unpleasant odor. The peritoneal cavity contained a yellowish mucoid fluid and visceral organs showed multiple tiny white foci. The intestines contained a thick, white, opaque mucus. Most internal organs showed extensive ulcerative dermatitis, cardiac endothelial hyperplasia, and necrotic abscesses. In channel catfish (Ictalurus punctatus), E. tarda produces a putrefactive condition resulting in gas filled pockets in the muscle that emit a putrid odor when cut (Meyer and Bullock, 1973). In eels (Anguilla japonica) the disease is a kidney nephritis accompanied by suppurative hepatitis (Miyazaki and Kaige, 1985). Edwardsiella tarda infection in striped bass resembles the nephritic form of eels.

Edwardsiella tarda infections in fishes other than striped bass are usually associated with warm temperatures, poor water quality, and the presence of high concentrations of organic matter. The impact of E. tarda infections on striped bass is not clear but as the culture of this species intensifies, and beating in mind the broad host range of the pathogen and its propensity to infect environmentally stressed fish, it is possible that this organism could become significant. Also, E. tarda has an affinity to brackish water environments, therefore, it could be a potential problem in cages or net pens suspended in such areas.

T a b l e I 1.3. Presumptive biophysical and biochemical characteristics o f gram negative bacteria that cause disease in striped bass and their hybrids ~.

Bacterial Species Character

Aeromonas hydrophila

A. sobria

Pseudomonas fluorescens

Pasteurella piscicida

Vibrio anguillarum

Edwardsiella tarda

Flexibacter columnaris

Cell m o r p h o l o g y

Short rod

Short rod

Short rod

Short rod

Short rod

Short rod

L o n g rod

Isolation media

BHI; T S A

BHI; T S A

BHI; TSA

BHI; Blood

BHI

BHI; T S A

O r d a r s , Hsu-Shotts, etc.

NaC! tolerance (%)

0-4

?

?

0-3

0-7

0-4

0-0.5

Motility

+

+

+

-

+

+

+ (Flexing)

C y t o c h r o m e Oxidase +

+

+

+

+

-

+

Oxid./Ferm.-Glucose

+/+

+/+

+/-

+/+ (weak)

+/+

+

NA

H2S prod. (TSI)

+

?

-

-

+

+

NA

lndole

+

+

?

-

+

+

Esculin

+

-

NA

?

-

-

Sensit. to 0/129

-

-

NA

+

+

NA

+

Pigment production

-

-

Soluble 9 fluorescing

-

-

-

Non-soluble yellow

'"+" = positive "-" = negative "?" = U n k n o w n "NA" = N o t applicable

to oo

282

Fig. 11.5. Pasteurellapiscicida in striped bass. (A) The spleen (arrow) is enlarged and dark red. (13) Liver, and spleen with granulomas (small arrows), and congested blood vessels in the ovaries (large arrows). (Photos courtesy of J. P. Hawke).

11.4.5. Vibriosis Vibriosis is a serious bacterial disease of many marine fish species, and it can be a significant problem in striped bass reared in salt water pens or cages as well as of wild fish. Several Vibrio species are implicated and include V. anguillarum, V. vulnificus, V. alginolyticus, V. cholerae, V. mimicus, V. parahaemolyticus and possibly others (Colwell and Grimes, 1984). Vibrio anguillarum is most often associated with bacterial infections of striped bass. Vibrio spp. infections are usually stress related; high stocking densities, handling, temperature shock and poor water quality are probable predisposing factors. Infections usually occur in salt water because most of the Vibrio spp. are halophilic and require salinities of 5 ppt or greater to cause disease, however, V. cholenae and V. mimicus, and rarely V. anguillarum may cause infections of fish in freshwater. In addition to lethargy, clinical signs of vibriosis in striped bass are hyperemia of the fins and skin that results in scale loss and development of ulcerated epidermal lesions (Figure 11.6). These lesions may appear at any location on the body including the head and gill covers and are quite similar to those associated

283

with motile Aeromonas septicemia. The gills are often pale and eyes may be hemorrhaged and exophthalmic. Internally, the body cavity may contain a bloody fluid, the liver may be pale and/or mottled, the spleen is usually swollen and dark red and the kidney is often swollen and soft. The gastrointestinal tract is usually void of food, flaccid and inflamed. Vibriosis is diagnosed by isolation of the causative organism on BHI or TSA media that is supplemented with 2 to 3% NaCI and incubated at 25 to 30~ Vibrioanguillarum is a gram-negative, slightly curved rod that measures about 0.5 X 1.5 #m; it is motile, cytochrome oxidase positive, ferments carbohydrates without gas and is sensitive to novobiocin and the vibriostat 0/129 (Table 11.3). Vibrio anguillarum is a heterogenous species with at least five different biotypes and serotypes (Pacha and Kiehn, 1969; Hastein and Smith, 1977; Austin and Austin, 1987). Diagnosis of vibriosis is confounded by the fact that several species of Vibrio and other pathogens can cause similar clinical disease. Virulence of different isolates may vary from lowto high but environmental conditions will influence the mortality rate. Vibrio anguillarum may have a significant effect on cultured striped bass and the more intensive the culture system the more severe infections may be. The impact of vibrio infection is increased

Fig. 11.6. Vibriosis (Vibrio) on striped bass. (A) Vibrioanguillarum causing necrotic, hemorrhaged lesions (arrows) on striped bass (photo courtesy of F. Hetrick, and A. Baya). (B) Vibrioparahaemolyticus on fingerling striped bass (arrow) (photo courtesy of A. Baya).

284

during stressful periods such as elevated water temperature, chronically low oxygen, high ammonia concentrations, and crowded conditions. 11.4.6. Streptococcosis

Streptococcus septicemia affects a variety of fish species, but is particularly serious in some striped bass culture operations (Kitao, 1993; J. P. Hawke, Louisiana State University, personal communication). The disease is usually chronic but may on occasion be acute. Clinical signs of Streptococcus infection in striped bass are not particularly specific, but the fish are generally darker than normal, exhibit erratic, spiral swimming, and often display curvature of the body (Figure 11.7). They often have either bilateral or unilateral exophthalmia with hemorrhage in the iris. Hemorrhages develop at the base of fins, in the scale pockets, and on the operculum and mouth; ulcerative lesions seldom occur. A bloody fluid is present in the intestine, which is also hyperemic, and the liver is pale but the spleen is greatly enlarged and dark. Several species of Streptococcus affect fish, but the organism that infects striped bass has not been named; it is a non-hemolytic (alpha), Group B, type Ib Streptococcus sp. (Table 11.4). Kitao (1993) suggested that according to unpublished DNA/DNA hybridization studies that alpha hemolytic Streptococcus sp. are genetically different from S. faecalis and S. faecium, therefore the Streptococcus from fish remains unnamed. Infection is diagnosed by isolation on BHI, nutrient agar, or TSA media with growth being enhanced by the addition of blood or up to 3% NaC1. The organism is a gram-positive, non-motile coccus that appears in chains of up to about nine cells in culture. In infected fish the bacterial cells usually appear singly, paired or in short chains. Although Streptococcus has been reported in wild striped bass in brackish water (Baya et al., 1990b), the infection is more significant in fish cultured in ponds, cages, net-pens, and raceways in brackish water or estuarine waters where the organism apparently occurs naturally. Mortality in intensively cultured striped bass can be high, especially when water temperatures are 25 to 30~ deaths seldom occur at temperatures below 20~ Handling and moving fish seems to trigger an overt epizootic in fish where Streptococcus is endemic. Abrasions, loss of scales and other injuries to the skin are also important precursors to streptococcosis in some species of fish (Chang and Plumb inpress). Other stressful environmental conditions, such as poor water flow chronically low oxygen concentrations, high population density, etc. also contribute to the onset of clinical infections. 11.4.7. Enterococcosis The genus Enterococcus is a relatively new taxonomic group that was previously included in Streptococcus (Mundt, 1986). Enterococcus infections have been noted in a limited number of intensive striped bass rearing facilities using fresh water at temperatures of 28 to 32 ~ (J. P. Hawke, Louisiana State University, personal communication). The causative organism is Enterococcus faecium (formerly Streptococcusfaecium). Adult and juvenile striped bass develop a septicemia with hemorrhages in the scale pockets, and swollen, hemorrhaged or cloudy eyes.

Enterococcus is a gram-positive, non-motile cocci, but more ovoid than round. The organisms are Beta hemolytic (Lancefield's Group D); grow at 45~ in 40% bile and in 6.5% NaCl; these characteristics separate them from the Streptococcus sp. The full impact of E. faecium on striped bass is unknown, but judging from recent epizootics the potential is considerable.

285

Figure 11.7. Streptococcus sp. (A) Striped bass with hemorrhages in the skin, missing scales, and a whitish ring around the eye (Photo courtesy of J. Hawke). (B) Streptococcus associated necrotic lesions in striped bass (photo courtesy of J. Evans).

11.4.8. Mycobacteriosis Mycobacteriosis offish has been known for years, but it has become one ofthe most serious infections in intensive, recirculating culture systems of striped bass. The disease is generally chronic but in certain situations becomes acute (Frerichs, 1993). The causative organism is usually Mycobacterium marinum, however other acid-fast staining bacteria may be involved. When infected fish are in closed, recirculating systems, the bacterium apparently has an opportunity to accumulate and produce a serious infection (J. P. Hawke, Louisiana State University, personal communication).

Mycobacterium marinum infected fish become dark and emaciated with occasional ulcerations and hemorrhaging in the skin. The most striking gross pathology is a pale, granulomatous liver with a rough granular surface (Figure 11.8). Granulomas also develop in the spleen, heart, kidney, and mesenteries; the surface of these organs have a rather rough, sandpaper texture.

to oo

Table 11.4. Presumptive characteristics of gram (G) positive and acid-fast bacterial pathogens of striped bass and their hybrids.

Character

Bacterial Species I

Streptococcus sp.

Carnobacterium p isc icola

Corynebacterium aquat icum

Mycobacterium marinum

Cell morphology

Coccus

Short rod

Short rod

Short rod

Stain reaction

G+

G+

G+

Acid-fast

Isolation media

BHI, Blood Agar

BHI, TSA

BHI, TSA

BHI, Petraynani's agar

NaC! tolerance (%)

0-3

0-2

0-3

?

Motility

.

Hemolysis of blood

None (Group B)

+ Alpha or Beta

None

NA

Catalase

-

NA

NA

NA

CAMP Test (Facklam, 1980)

+

NA

NA

NA

~"+" = "-" = "NA" = "?" =

Positive character Negative character Not applicable Unknown

.

.

.

287

Mycobacteriosis is diagnosed by detection of strongly acid-fast (Ziehl-Neelsen) staining cells in smears, or histological sections of the ~anulomas. The ceils are non-motile, pleomorphic rods that measure about 0.5 X 1.5 to 2.0/.zm (Table 11.4). They can be cultured on Lowenstein-Jensen or Petrignanis's media, but isolation is often difficult because the organism may require up to three weeks at 25~ to form visible colonies. Hedrick et al. (1987) reported that M. marinum grows at temperatures from 20 to 37~ It is likely that a low level ofmycobacterial infection occurs in wild fish, not necessarily striped bass, and are reservoirs of the organism for cultured fish, but mycobacteriosis seldom creates overt disease in wild fish. It has also been theorized that carrier adult striped bass could be the source of infection and infect juveniles through vertical transmission. Sakanari et al. (1983) found the prevalence ofmycobacteriosis in wild striped bass to be 25 to 68% in California and 46% in Oregon. However, months may elapse between exposure of the fish to the organism and actual appearance of clinical infection. Morbidity in striped bass populations may be low at any one time but cumulative mortality may become high. Hedrick et al. (1987) reported that 50% of a yearling striped bass population died due to M. marinum infection within months of stocking into an intensive culture system, and 80% of those remaining were infected. Hybrid striped bass in closed recirculating systems in Louisiana experienced chronic mortalities due to mycobacteriosis, but up to 50% of the fish had characteristic mycobacterial ~anulomas in internal organs (J. P. Hawke, Louisiana State University, personal communication). Mycobacterium marhmm has the potential to cause infections on the extremities of humans coming in contact with infected fish; therefore, caution should be exercised by fish culturists when handling striped bass in facilities that may enhance the expression ofmycobacteriosis, namely, intensive recirculating systems. Skin lesions on humans are usually confined to the hands, wrist, and forearms where hard, raised, calcified, granulomas develop. Apparently, the organism cannot adapt to the higher inner body temperature, therefore, it seldom precipitates systemic problems unless the individual is otherwise debilitated (Frerichs, 1993).

Fig. 11.8. Mycobacteriosis (Mycobacterium marinum). The gills are pale toward the proximal ends of the filaments because of granulomas, the liver (large arrow) is pale and highly granular (granulomas) and the spleen (small arrow) is enlarged and has white granuloma lesions (Photo courtesy of J. Newton).

288

11.4.9. Other Bacteria Several other bacteria have been isolated from diseased striped bass that may or may not be serious pathogens. Baya et al. (1990a) isolated a bacteria that belonged to the genus Moraxella. The organism was a short, gram-negative rod that often appeared in pairs; it exhibited bipolar bodies, was cytochrome oxidase positive, non-fermentative in glucose, and did not produce acid from most carbohydrates. Affected fish had large hemorrhagic lesions on the dorso-lateral body surface and scales had been lost. Internally the liver was pale to mottled and enlarged. Hemorrhages were also found in the swim bladder, and membranous material (adhesions) connected the liver to the body wall. The actual influence of the Moraxella infection in the disease process was somewhat speculative because gills of afflicted fish were heavily parasitized with Trichodina and Ergasilus. Also, a viral agent, tentatively named striped bass reovirus, was isolated from these fish (see Section 11.3.3 of this chapter). Baya et al. (1991) reported the isolation of a Carnobacterium-like organism from moribund and dead striped bass, and other fish in the Chesapeake Bay area of Maryland. No clinical signs of disease were described, but the fish had been stressed before the infection. The organism was similar to Carnobacterium piscicola which is a gram-positive bacillus that tolerates salinities from 0 to 6% and has a growth temperature range from 10 to 37~ (Table 11.4). It was easily isolated on BHI agar or TSA. Toranzo et al. (1993) were unable to kill striped bass with C. piscicola but did kill about 35% of rainbow trout (Oncorhynchus mykiss) that were injected with 4.5 X 10 6 cells. Histologically the organism caused mild infiltration of the liver, hemorrhage in the kidney, and inflammation of the meninges in striped bass. These observations led to the conclusion that C. piscicola possesses low virulence to striped bass and produce only moderate injury to internal organs. However, infected striped bass could be carriers of the organism for at least two months, and it was suggested that perhaps the fish infected with C. piscicola would be more susceptible to invasion by secondary pathogens, or to environmental stress. The full impact of C. piscicola on cultured striped bass is yet to be determined. Baya et al. (1992) isolated a bacterium, identified as Corynebacterium aquaticum (Table 11.4), from the brain of striped bass exhibiting exophthalmia. This is the first report of C. aquaticum, normally a water borne organism, being pathogenic to fish. The LDs0 of the organism in striped bass was 1.0 X 105 colony forming units. Experimentally infected fish developed hemorrhaging in most internal organs with the most severe being in the brain and eyes. It should be noted that C. aquaticum is also a possible infectious agent for homeothermic animals. 11.5 F-YNGAL DISEASES Fungal diseases of striped bass have been recognized for a long time, however, less is known about them than some other types of diseases. Several reasons have contributed to this dearth of knowledge of fungi infecting striped bass, not the least of which is that fungi are difficult to identify and are often secondary pathogens to other diseases, injury, or environmental stress. However, several fungi do cause infections in striped bass including Saprolegniaparasitica and related species (water mold), and Branchiomyces sp. (gill rot). 11.5. I. Saprolegniosis Saprolegniosis is a general term referring to a number of species of fungi that occur in water and infect the epithelium and eggs of a variety of fish species including striped bass (Figure 11.9). The principal species are Saprolegniaparasitica, Jphanomyces sp., and Achlya sp. Other species of fungi may occasionally occur,

289

but nearly all are in the family Saprolegniaceae (Alderman, 1982; Hatai, 1989). The taxonomy of these fungi is such that no effort to taxonomically separate them is made in this text. These fungi are saprophytic organisms that are widely distributed in the aquatic environment and can derive nutrients from any organic source in water. These fungi most often become pathogenic to fish only when fish are stressed, injured, in a poor nutritional state, temperature shocked, or otherwise debilitated. They occur on incubating dead or injured fish eggs, or when incubation temperature is below the optimum.

Saprolegn&reproduces either sexually or asexually (Alderman, 1982). Sexual reproduction is by fusion of two gametes to form a thick-walled oospore, whereas asexual reproduction is by means ofzoospores produced in a zoosporangium, which develops at the end ofnon-septate cells. Zoospores are motile by means of flagella after escaping from the zoosporangia. After going through primary and secondary stages, these reproductive cells attach to dead or injured tissue to produce filaments (hyphae) which, in turn, form the visible mycelium (fungal colony). As the hyphae grow from the dead tissue, they invade surrounding healthy tissue and produce necrosis. Once the fungus becomes established it can kill infected fish. The prognosis of fungal-infected fish is not encouraging because most antifungal medicants have similar toxicities to fish and fungi. Saprolegniosis is identified by the presence of large non-septate, tubular cells in wet mounts from infected tissues. Saprolegniainfections are generally external and appear anywhere on the body surface, fins, or gills. They are often accompanied by pathogenic bacteria or protozoan parasites, which can also be the predisposing factor to fungal infection. Fungal colonies, which appear as tufts of cotton on the fish's body, may be white, grey, or turn brown as the mycelium traps mud or silt. Saprolegniausually does not produce deep lesions in the muscle, but infected fish swim lethargically and may linger moribund for several days before death. Aphanomycessp. are more likely to produce deep, ulcerated lesions in the muscle of infected fish (Noga, 1992).

Fig. 11.9. Fungal infections. Saprolegniainfections (arrows) on the skin and fins of striped bass (Photo courtesy of R. Durborow).

290

Saprolegnia on incubating eggs can develop rapidly and usually begins with the fungus attacking injured, infertile, or dead eggs and then invading the healthy eggs nearby, and eventually totally engulfing all eggs in a hatching unit. The disease may occur at a wide range of temperatures with infections more often developing in water with below optimum temperature for hatching, but any temperature shock, either up or down, can be a predisposing factor to fungal infection on eggs or fish. 11.5.2. Branchiomycosis Branchiomycosis, also known as "gill rot," was first reported to occur in pond raised striped bass fingerlings by Meyer and Robinson (1973). The disease is caused by Branchiomycessanquinis, an obligate pathogen, and is more common in other species of fish than striped bass. Branchiomycessanquinis grows within the branchial blood vessels, but hyphae may protrude from necrotic tissues. Affected gills are necrotic and hyperplastic because of an infarction due to intravascular growth of B. sanquinis. Telangiectasis and vascular necrosis also occurs. Branchiomycessanguinis forms multi branching, nonseptate hyphae that measures 8 to 30 ~zm in length while the spherical spores, within the hyphae, measure 5 to 9/~m in diameter (Hatai, 1989). Branchiomycosis usually occurs in fish in waters with high organic content and during periods when water temperatures are 20~ or higher. Up to 50% morbidity may occur and affected fish may succumb rapidly due to this terminal disease. 11.6 P R O T O Z O A N DISEASES Protozoa (Protista) are single cell organisms that can inflict damage to striped bass when present in sufficient numbers. These ectoparasites, primarily flagellates and ciliates, are often present in low numbers on skin, fins or gills of fish. Fish infected with large numbers of protozoa do not feed actively, they swim lethargically and gasp at the surface, and affected fish do not tolerate handling very well. It is common to see two or more different species of protozoans on a single fish. Individual fish that are emaciated may harbor huge numbers of multiple protozoan species, but whether the fish are emaciated because of the heavy parasite load, or they are present in large numbers because of the poor condition of the fish, is difficult to determine. Protozoan diseases are diagnosed by examining wet mount slides from lesions at 100 or 400X magnification. Although some dead parasites are identifiable when preserved, they are more easily detected and identified in live material. 11.6.1. Flagellates Flagellated protozoa occur on the skin, fins, and gills of striped bass, and occasionally attain sufficient numbers to cause injury, especially during periods of poor water quality and environmentally induced stress. 11.6.1.1. lchth,cobodiasis Ichthyobodiasis is a disease caused by the flagellate lchthyobodo sp., which is often present on the gills or skin (Post, 1987). The tear shaped lchthyobodo is about the size of a red blood cell, measuring 10 to 20 ;zm in length and 5 to 10/.zm in width (Figure 11.10). They have a pair of flagella, one of which modifies for attachment to the gills or skin where they derive sustenance from live cells, lchthyobodo is an obligate pathogen transmitted from host to host through the water, and young fish seem to be more susceptible than older fish. Usually the defenses of the host keeps the protozoan population at a low level, but if resistance is lowered, the parasite can proliferate and cause serious injury and death. A change in the health status of the host, or degradation of environmental conditions, such as low oxygen concentration, high ammonia, etc., or crowding are often associated with infections, but these stressors are not essential. Ichthyobodiasis may occur

291

between 10 and 25~ but optimum temperature for the disease is 18 to 20~ (van Duijn, 1973). Ichthyobodiasis may progress from a latent stage with no mortality of the fish, to severe infection with acute mortality in which losses may approach 90% in a few days.

Ichthyobodo infected fish refuse to eat and may become lethargic. Fish develop a thickening of the mucus on the most heavily infected areas of the skin giving the lesion a gray appearance. Fish with skin infections may scrape (flash) against the sides of tanks or submerged structures in an attempt to remove the irritants. It is also possible to have lchthyobodo infection only on the gills which swell and produce mucus. When material from lesions or gills are microscopically examined, unattached lchthyobodo cells move rapidly and erratically in the water. Attached cells are often side by side on the epithelium and their movement resembles flickering of a candle flame. Unless the parasites are alive it is most difficult to distinguish them from host tissue cells. 11.6.1.2 Amyloodinias.is The dinoflagelate, Amyloodinium ocellatum, causes amyloodiniasis and is of particular concern in striped bass mariculture (Lawler, 1980). Amyloodinium is an obligate parasite of marine fish, butit can tolerate low salinity (Lauckner, 1984; Johnson, 1987). It infects larvae, fry, and fingerling striped bass when the fish are held at high density. Visible, small white spots appear on the fins and skin. Amyloodinium ocellatum infected fish gasp rapidly at the surface, or congregate on the bottom of the container where they swim erratically and lose equilibrium. Infected fish also scratch against submerged objects or the tank sides. The parasite attaches to the gills by way of organelles that cause tissue hyperplasia, fusion of gill lamellae, inflammation, hemorrhages, and eventually gill necrosis. The parasite may also attach to the skin of fish during heavy infestations giving rise to the name "velvet disease."

Amyloodinium ocellatum has a distinct life cycle beginning with the attachment of a dinospore (swarmer) to the host where it develops into a parasitic stage, the trophont (Lawler, 1977). Upon maturity, the reproductive (tomont) stage detaches, divides into as many as 256 free-swimming dinospores, and reattaches to the host where it becomes a mature trophont, and the cycle continues. The speed of the life cycle depends on water temperature and varies from 3 to 6 days, but dinospores retain infectivity for up to 15 days in water. Amyloodiniasis is detected by observing the parasite inwet mounts from infected tissue which appear as brown, circular, or oblong cells of various sizes (mature trophonts measure about 150/~m, Figure 11.10). Secondary bacterial invaders are prominent in the epithelial lesions.

Amyloodinium ocellatum is introduced into the cultural system by carrier fish, or the free living stage gains access via the water supply from natural sources. The parasite may reach concentrations on the skin that severely affect the health and feeding response of striped bass. Heavy infestations ofA. ocellatum are usually terminal, with death being the result of acute anoxia and suffocation (Lawler, 1977). Infections may develop rapidly resulting in death of fish in as little as 12 h after introduction of healthy fish into tanks containing numerous dinospores (Overstreet, 1978). In closed recirculating systems as high as 80% of stocked striped bass may die as result ofA. ocellatum(Johnson, 1987). This disease is not as much of a problem in open water cages, net-pens, or flow through systems because the parasite cannot complete its life cycle as easily under these conditions.

292

Fig. 11.10. Flagellated Protozoa. (A) Ichthyobodo sp. (arrow) attached to the gill epithelium (Photo courtesy of W. A. Rogers). (B) Amyloodinium ocellatum (arrows) on the gills of striped bass. The different size cells are in various stages of development (Photo courtesy of S. K. Johnson).

11.6.2. Ciliates Several species ofciliated protozoa can be present on striped bass, however, very few are devastating. The most common genera of ciliates that infect striped bass are Ambiphyra, Apiosoma, Chilodonella, Trichodina, Epistylis, lchthyophthirius, and Cryptocaryon. All but Ichthyophthirius and Cryptocaryon are symbiotic protozoa that reside on the skin, fins, or gills (Hoffman, 1978). These symbionts are basically ubiquitous, have no host specificity and have little effect on the host unless present in great numbers or when the fish are stressed due to some environmental or water quality disorder. All possess cilia, or tentacles, which are used for mobility or feeding. Reproduction is by binary fission with a simple life cycle that involves the formation of reproductive stages, either on the host or in the water. Most ciliates are more numerous when water temperatures are cool. Discussion of the ciliates that are most common or cause the most serious disease problems to striped bass follows and merely presents examples of the broad group.

293

11.6.2.1. Chilodonella These protozoa are flat, spoon shaped cells that measure about 30 to 70/_zm in length by 20 to 40 gm in width. They occur on the gills or skin and are motile as a result of parallel rows of cilia on the body surface which also has an oral groove. While mortalities due to Chilodonella are usually not severe, infected fish have increased mucus on infected gills or skin. 11.6.2.2. Epis.tylis There are several species of Epistylis that infect the skin, fins, and gills of fish (Esch et al., 1976). The urn shaped organism, which is adorned by a ring of cilia on the distal end, is at the terminus of a dichotomous stalk that, in turn, attaches by a disk to hard surfaces of the host such as spines, scales, or gill covers. Groups of Epistylis form a colony (Figure 11.11). These parasites cause irritation and inflammation of the epithelium of the host at the point of attachment which may provide a site for secondary infections of Aeromonas hydrophila. The parasites feed primarily on bacteria and organic material in the water, but they erode scales and hard spines of fins where they attach. Other than the injury incurred by attachment, they are seldom harmful to the host unless there are large masses of these parasites. 11.6.2.3. Trichodina Trichodinids are complex protozoa that are found in low numbers on the gills, skin, or fins of most fish, but occasionally may be extremely numerous and injurious (Wootten, 1989). Trichodina sp. are round and saucer or bell shaped with a ring of cilia on the margins (Figure 11.I 1). The convex side, or adoral surface, possesses an attachment organ, the adhesive disk. This disk has a complicated tooth-like structure (denticular ring) arranged concentrically, the pattern of which varies between species. Trichodinids move over the surface like a saucer being slid across a table, or they swim upright on their edge. Trichodina sp. are the most often reported parasite on many species of fish, however, they seldom attain the numbers that are detrimental to the health of the host. When present on the gills in large numbers, the epithelium becomes swollen and excess mucus is produced. 11.6.2.4. Ichthyophthiriasis Ichthyophthiriasis, commonly known as "Ich," is the most devastating protozoan disease of fish, and it may cause high losses in cultured striped bass. The disease in fresh water is caused by lchthyophthirius multifiliis and the marine counterpart is Cryptocaryon irritans (Hoffman, 1978), both of which have similar life cycles. These obligate parasites mature while embedded in the epithelium where they form visible white spots (trophozoites) (Figure 11.12). On maturation, the trophont (mature trophozoite) leaves the fish and drops to the bottom or attaches to vegetation. The trophont forms a thick gelatinous coating, forming a cyst in which 250 to 1,000 infecting units, called tomites (swarmers), are produced. Tomites released from the cysts seek and penetrate the skin of the host where they develop into tomonts and mature trophozoites. The infective tomite stage must find a host within about 48 hours or it will die. This reproductive cycle is temperature dependent with an optimum of 24 to 26~ at which the process requires about 4 days, and at 7~ 35 to 40 days are required. The process proceeds more slowly at lower temperatures, but temperatures approaching 28 to 30~ completely shut down the process. The presence of small white spots in the skin or gills is an indication of Ich. Infected fish become lethargic, listless, gasp at the surface, and produce copious amounts of mucus. Diagnosis is achieved by detecting mature cells or trophozoites in wet mount preparations of skin or gill scrapings (Figure 11.12). Adult

294

Fig. 11.11. Ciliated Protozoa. (A) Trichodina sp. from gills with a prominent denticular ring (arrow). (B) Mucoid colony (arrow) of Epistylis on the skin of a centrarchid. (C) Colony of Epistylis (arrow) on dichotomous stalks (Photos courtesy of W. A. Rogers).

295

Fig. 11.12. Ichthyophthiriasis (lchthyophthirius multifiliis). (A) L multifiliis on the skin of spotted bass. The small visible white spots (arrow) are mature trophonts. (B) Developing trophozoites on a fin. (C) Mature trophont with the "C"-shaped nucleus. (Photo courtesy of W.A. Rogers).

296

Ich cells are large (0.5 to 1.0 mm) and possess a "C"-shaped nucleus. Various size trophozoites and tomites are characterized by extensive ciliation and movement similar to a rolling water-filled balloon. A low to moderate lchthyophthirius infection will generally progress into heavy infections that are devastating and can cause explosive mortalities of up to 100% if not treated, particularly in intensive culture systems (Post, 1987). The disease is somewhat seasonal due to its temperature requirements, therefore, it is most often seen in the spring and fall with some epizootics occurring in the winter, but almost never in the summer, unless the temperature of intensive systems is maintained in the optimum range. Fish do not need to be stressed in order to develop clinical disease. Also, injury to the skin by lchthyophthirius often allows multiple secondary invasion of bacteria or fungi. Severe infections may occur on the gills with little evidence of infections on the skin. 11.7 METAZOAN PARASITIC DISEASES Metazoan parasites of striped bass include the monogenetic and digenetic trematodes (flukes), cestodes (tape worms), nematodes (round worms), acanthocephalans (spiny-headed worms); leeches; and parasitic crustaceans (Post, 1987). Many of these organisms infect or infest striped bass and other fishes, but few are specific for striped bass, or cause severe debilitation of infected fish except under unusual circumstances. They are of interest primarily due to their manifestation on the host. Striped bass raised in ponds may be particularly susceptible to metazoan parasites, and if fingerlings so produced become infested before stocking into intensive tanks or recirculating systems, they may create problems. Many metazoan infections can be detected by the unaided eye because of the macroscopic size of the adult worms or the size and nature of the encysted larval forms, however, identification of species requires microscopy and knowledge of the taxa (Hoffman, 1967, 1973). Most trematodes, cestodes, nematodes, or acanthocephalans are not harmful to humans if ingested from freshwater fish. However, if these parasites occur in marine fish the natural final host is often a marine mammal and could cause some problems in humans if consumed raw. Proper cooking eliminates any zoonotic problem in ingesting metazoan parasites from fish. 11.7.1. Helminths Helminthic parasites include the monogenetic and digenetic trematodes, cestodes, nematodes, spiny headed worms, and leaches. 11.7.1.1. Monogenetic trematodes Monogenetic trematodes (gill flukes or body flukes) occur on the gills, skin, and fins where they browse on dermal or gill debris (Hoffman, 1967; Noble and Noble, 1971). According to Hughes et al. (1990) members of four genera of monogenetic trematodes (Diplectanum sp., Gyrodactylus, Microcotyl sp., and Urocleidus sp.) occur on striped bass. Gyrodactylus sp. occurs on the skin and the others on the gills. None of these parasites of striped bass have been identified to species. Monogenes have simple life cycles that involve either laying eggs or bearing live young and requires but one host. Monogenetic trematodes attach to the host by a structure on the posterior end called a haptor, that is equipped with hooks or sucking valves which may cause injury to the fish's epithelium (Figure 11.13). Many fish will have a few monogenes either on the gills or the skin, which are not overly deleterious, but occasionally these parasites occur in sufficient numbers to cause injury. However, the pathogenic effects of monogenetic trematodes on striped bass appears to be minimal (W. A. Rogers, Auburn University, personal communication). When monogenes do cause problems, the fish becomes lethargic, swims near the surface, refuses food, and occasionally results in death (van Duijn,

297

Fig. 11.13. Monogenetic trematode on a gill. The worms are attached to the fish tissue by a haptor (Photo courtesy of W. A. Rogers).

1973). The gills may have areas of thickened mucus, hyperplasia, and necrosis. High populations of trematodes on the skin causes increased mucus production, giving the fish a whitish, patchy appearance and cause the host to rub against the sides of tanks or underwater structures. 11.7.1.2. Di~enetic trematodes Digenetic trematodes have a complex life cycle involving multiple larval generations in different intermediate hosts before developing into adults in a primary host (Noble and Noble, 1971; van Duijn, 1973). The larvae of several digenetic trematodes infect visceral organs, eyes, and muscle of striped bass. Adult digenetic trematodes live in the gastrointestinal tract of fish or fish eating animals, usually birds. Trematode eggs released into the gut of the final host are voided with defecation, and the hatched larvae (miracidium) seek and invade snails where they form a sporocyst and undergo further division to produce hundreds of cercaria. Cercaria escape from the snail and actively seek a second intermediate host, usually a fish, including striped bass. The cercaria penetrates the second intermediate host and develops into a metacercaria (larvae) where the parasite resides until that host is consumed by the final host. These,parasites generally cause little damage to the secondary host, but when small fish contract large numbers of the parasites during a short period oftime, severe injury can occur. These are generally of major concern because of their manifestation in the visceral organs, eyes, or flesh of the intermediate host. It is not unusual to have more than one species of digenetic trematode in an individual fish. All of these trematodes occur primarily in pond cultured striped bass where snails are indigenous and fish eating birds have easy access. Because of their life cycle, there are no geographical restrictions. The most common digenetic trematode is Posthodiplostomum minimum, commonly called the "white grub," that infests the heart, liver, spleen, kidney, and muscle of many fish including striped bass (Spall and Summerfelt, 1970; van Duijn, 1973; Hoffman, 1978). These encysted larvae appear as small white spots and may be numerous (Figure 11.14). In a recent case, heavily infected 5 to 6 cm striped bass fingerlings exhibited

298

Fig. 11.14. Digenetic trematodes (white grub). (A) Striped bass fingerling heavily infected with white grub (P.

minimum). Note the exophthalmia and edematous musculature. (B) Visceral organs of a bluegill heavily infested with white grub cysts in the liver (large arrow) and the heart (small arrow) (Photo courtesy of W. A. Rogers).

extreme exophthalmia as the result of large numbers of larvae behind the eye and internal organs. They exhibited hemorrhage in the eye, severe edema of the musculature, and very pale internal organs (Figure 11.14). About 95% of the fish in the pond population were infected and were obviously in poor health. The yellow grub, Clinostomummarginatum, does cause some problem in cultured striped bass because the larvae form a visible 1 to 2 mm yellow cyst in the muscle (Figure 11.15). Infected fish are usually not harmed, but when large numbers of parasites are present, either just under the skin or in the muscle, they are unsightly. Consequently the fish are unsalable which occasionally has led to destruction of entire populations of striped bass fingerlings infected with yellow grub.

Diplostomum spathaceum, and D.flexicaudum (eye flukes) are digenetic trematodes that occasionally affect the eyes of striped bass. The larvae concentrates in the eye lens causing opaqueness (cataract) and blindness (Figure 11.15). The grub, Neascus sp. or Uvulifer sp., produces a black cyst in the skin and flesh of striped bass, thus the name "black spot disease." It is thought to cause little harm to the fish and does not affect its culinary quality because many of these reside in the epithelium and are removed by skinning.

299

Fig. 11.15. Digenetic trematodes (yellow grub). (A) Yellow grub (C. marginatum) cysts (arrows) in the muscle of striped bass. (B) Black grub (Neascus sp.) (arrow) in the skin and muscle of a centrarchid. (C) Eye fluke (D. spathaceum) producing cataract (arrow) in a largemouth bass (Photos courtesy of W. A. Rogers).

300

11.7.1.3. Cestodes Adult cestodes (tape worms) live in the intestine of piscivorous fish, or other vertebrates, but intermediate larval stages live in the viscera and muscle of a variety of fish species (van Duijn, 1973; Post, 1987). There is no record of adult tape worms in the gut of striped bass. However, there are several types of larval tapeworms that parasitize visceral organs of striped bass. The body of larval cestodes is ribbon-shaped but not divided into the typical short segments (proglottides) characteristic of the adult life stage. Larval cestodes possess a holdfast organ (scolex) at the anterior end, which is useful in identification. The life cycle of tapeworms is varied, with fish taking the role of primary, intermediate, or transport host. The striped bass serves as an intermediate host in most cases where the parasite manifests itself as yellowish cysts in the visceral organs or muscle tissue. Beginning with the adult in the intestine of the host, eggs are deposited from proglottides into the feces and then into water on defecation. A hexacanth embryo develops in the egg, which is then ingested by a copepod where a procercoid forms. The copepod is eaten by a variety of forage fish species or striped bass fry ingest the copepod with the procercoid, and the plerocercoid encysts in the visceral organs where it remains until eaten by the final host. These larval parasites are generally not harmful to the fish but the unsightly cysts may be visible with the unaided eye and cause some concern by the consumer. None of the tapeworms that use striped bass in freshwater as intermediate host will infect humans. 11.7.1.4. Nematodes Nematodes, or round worms, infect the visceral organs of striped bass or the larvae encysts in the eye. Philometra sp. may be seen in the mesenteries of striped bass where these blood red worms resemble large blood vessels (Figure 11.16). Most nematode infections, either larval or adult, cause little harm to the host unless Philometra sp. occurs in the eye, in which case the eye is destroyed when the adult worm and its larvae escapes the host. The life cycle of nematodes in fish may be direct from host to host, but usually involves a micro-crustacean intermediate host. If intermediate hosts are required, the larval stages encyst in the visceral organs along with other larval metazoan parasites. Goezia sp. is a nematode that penetrates the stomach wall where it burrows through the musculature (Figure 11.16) (Gaines and Rogers, 1972). Goezia sp. normally occurs in marine fish, but it can create problems if uncooked marine "rough fish" are chopped and fed to striped bass cultured in freshwater. 11.7.1.5. Acanthocephalans Adult acanthocephalans (spiny head worms) possess an anterior proboscis covered with many hooks (Hoffman, 1967) with which they attach to the epithelium of the intestine of vertebrates. The trunk of these worms is more or less cylindrical in shape. Where the proboscis is imbedded in the epithelium of the host's gastrointestinal tract there may be necrosis, ulceration, and peritonitis. If infections are heavy, death of the host may occur; however, usually their consequence is minor. The final host for each species of acanthocephalan is specific. Pomphorynchus rocci is found in striped bass, but the intermediate host of the larvae are less specific occurring in a variety of forage fish species. Encysted larvae of several species have been detected in the viscera of striped bass. The life cycle of acanthocephalans involves passage through crustaceans, then to fish as an intermediate or final host. 11.7.1.6. Leeches Leeches have occasionally occurred on the skin of striped bass raised in ponds and tanks (L. C. Woods, III, University of Maryland, personal communication). These blood sucking annelid worms are

301

Fig. 11.16. Nematodes. (A) Philometra sp. (arrow) in the mesenteries of the visceral cavity of striped bass. (B) Goezia sp. burrowing through the stomach wall of striped bass (Photo courtesy of W. A. Rogers). transient and usually cause no health problems, however, small fish with large numbers of leeches could suffer (Chitwood, 1969). Leeches have simple life cycles and do not involve intermediate hosts. 11.7.2. Crustacean Parasites Parasitic crustaceans occasionally cause problems in cultured fishes including striped bass. The parasites usually involved are Lernaea sp., Argulus sp., and Ergasilus sp. (Figure 11.17) (Hoffman, 1973). There is virtually no host specificity associated with parasitic crustacea, therefore, any one of those listed may occur on striped bass. Female Lernaea sp. attach to the skin of fish where their modified head is imbedded into the flesh of the fish (Figure 11.17). The attached female Lernaea develops two egg sacs at the distal end of the tubular body. Following hatching, the naupli larvae migrate to the gills where they go through a series of molts in which they become copepodids that resemble a typical crustacean, although they are microscopic. On maturation to the last copepodid stage, the female mates with the male. The male then dies while the female mi~ates to some spot on the fish, attaches to the epithelium, and the reproductive cycle starts again. Adult female Lernaea, with a body of about 0.5 cm in length, are visible to the unaided eye and appear as small sticks attached to the skin. Unless there are a great number of adults on a fish they do little damage, however, the copepodid stages on the gills may cause extensive injury.

302

Fig. 11.17. Parasitic crustaceans. (A) Adult Lernaea sp. attached to the skin. (Aa) Copepodid stage that appears on the gill. (B) Ergasilus sp. attached to the gill epithelium of striped bass. (C) Argulus sp. on the body surface. (Photos courtesy of W. A. Rogers).

303

Ergasilus sp. attaches to the gills of fish and follows a similar life cycle as Lernaea~ but these crustaceans are smaller and are not as visible without microscopy (Figure 11.17). Ergasilus attaches to the gills by imbedding modified appendages in the epithelium. If present in large numbers, they do cause epithelial hyperplasia and necrosis to the gill at the point of attachment. Argulussp. is a large parasite that moves freely on the skin, fins, and gills of the fish. This parasite, called "fish louse," is fiat with adaptations for attachment to the host on which it feeds. The body is broad, fiat, and oval (Figure 11.17). Unlike Lernaea and Ergasilus, Argulus leave the host and lay their eggs on vegetation. Naupli larvae develop in the egg and, on hatching, the copepodid stage seeks a new host. Unless large numbers are present on the fish they cause little damage. 11.8 C O N T R O L THERAPY Because no chemicals can be used in the control of infectious diseases of striped bass intended for food, management, health maintenance and disease prevention are the keys to successful striped bass culture (Plumb, 1994). As cultural systems become more intensive, the need for environmental control, water quality stability through management, reduction of stress on the cultured fish, and a sound health maintenance policy is essential (see Chapters 7 and 10). To reduce the potentials of catastrophic disease outbreaks, the striped bass culturists should maintain the highest water quality possible. This is best carried out by using prudent stocking densities, an adequate flow of freshwater through intensive culture units, removal of metabolites from recirculating water, supplemental aeration in flow through and pond culture systems and the use of high quality feed. Striped bass are sensitive to improper handling, therefore, gentle netting and handling is essential. The use of prophylactic chemotherapy during or following handling or moving fish will reduce the possibility of secondary bacterial infections and reduce the external parasitic population. The use of chemicals and drugs as prophylactics, and the application of therapeutics, if used at all, are mere tools to be employed in health maintenance. Successful aquaculture can not depend on chemotherapeutics to solve all of its infectious disease problems, however, regardless of the manage-ment level, sooner or later chemotherapeutics will be required to control an infectious disease in cultured striped bass. The use of drugs and chemotherapeutics on food fish is becoming more restrictive due to U. S. Food and Drug Administration (FDA) regulations (Meyer and Schnick, 1989; Anonymous, 1994). Concerns for unwanted residues in food, and the potential deleterious effects of these compounds on the fish, the fish culturist, and the consumer further curtail the use of many compounds. In view of these concerns, the FDA has limited the number of therapeutics that can be used on cultured food fishes, and under the current guidelines there are N_..Ochemotherapeutics approved for the "legal" treatment of any disease of striped bass (Table 11.5). It is hoped that the label for Terramycin for bacterial infections and formalin for parasitic infections, will soon be extended for cultured striped bass. With the cost and time required to obtain licenses and product labels that will allow the use of chemotherapeutics on striped bass it is unlikely that new compounds will be forthcoming soon. In some instances the aquaculturists can take advantage of the "extra label use" policy of the FDA which allows a qualified veterinarian to prescribe a drug or chemical to treat diseases of fish under certain conditions. This extra label use allows a veterinarian to prescribe a drug that is labeled for use in one food animal to be used in an identical manner in another food animal group. The following discussion outlines what has recently been used for the various infectious groups of organisms and the diseases they cause. With deference to the FDA regulations, the applications noted herein are for discussion purposes only, and do not constitute endorsement of their use on striped bass or their efficacy. Before any treatment is applied to striped bass the aquaculturist should determine the latest FDA policy and regulations on chemotherapeutics (Anonymous, 1994).

Table 11.5. Chemicals and antibiotics that are used for the control and treatment of infectious diseases of cultured striped bass. None of these compounds are approved by the U.S. Food and Drug Administration for use on striped bass, therefore, their listing here does not constitute a recommendation or indorsement.

Chemical

Purpose

Unit

Concentration

Duration

Comments

Formalin F

Parasiticide Fungicide

Pond Eggs

25mg/L 600mg/L

Indefinite ! 5 min

Low 02, striped bass more sensitive

Copper sulfate

Parasiticide

Pond

0.5-3mg/L

Indefinite

Cone. depends on water alkalinity

Potassium permanganate

Parasiticide & Antimicrobial

Ponds

2-4mg/L

Indefinite

Cone. depends on organic load

Tanks

2-10mg/L

1 hour

Caution at higher concentrations

Salt

Parasiticide & Antibiotic

Tanks

0.5-3%

See. to indefinite At higher cone. move to freshwater at sign of stress

Terramycin

Antibiotic

In feed

50-75mg/kg/day

14 days

21 day withdrawal

Romet

Antibiotic

In feed

50mg/kg/day

5 days

Long withdrawal

Masoten

Parasiticide

Ponds

0.25- I mg/L

Indefinite

305

11.8.1. Viruses Because virus diseases do not currently pose a major threat to striped bass, there is little effort to attempt their control. Lymphocystis is best handled by removing infected fish from a population and disinfection of the facilities after the fish are removed. Thorough drying or disinfection with chlorine (200 mg/L) should eliminate residual virus. Treatment of incoming water with ultraviolet (UV) radiation or ozone should eliminate the virus in water sources. Infectious pancreatic necrosis virus can be handled in a similar manner except it is not practical to eliminate infected fish from a population. Populations that carry IPN virus can be identified by virological sampling (Thoeson, 1994) where the fish may serve as a source oflPNV for cultured salmonids. Insufficient information exists to consider control of the SBR agent. 11.8.2. Bacteria Bacterial diseases of striped bass are best controlled by maintaining a high quality environment, and avoiding stressful conditions and injury to the fish; in other words "fish health management" (Plumb, 1994). Reduction of population density often reduces the effect of infectious diseases. Disinfection of water with UV radiation and ozone will help reduce the bacterial populations in recirculating water or open water supplies. Elimination of uncooked wild fish from diets will help reduce the possibility of mycobacterial infections. Sanitation by routinely sterilizing nets, buckets, and other utensils between use will reduce accidental cross infecting culture units. Either chlorine (200 mg/L) or a solution of a quaternary ammonium compound (i.e., Roccall) (20 to 100 mg/L) can be used for disinfection. When these substances are used, thorough rinsing with clean water is essential so that the disinfectant is not carried into the culture units. In some instances, Pasteurellapiscicida for example, it may be helpful to stock fish that have been shown to be free of certain pathogens (J. P. Hawke, Louisiana State University, personal communication). Although there are no therapeutic agents approved by the FDA for striped bass, some drugs and chemicals have been successfully used prophylactically or in chemotherapy for clinical bacterial infections. Prophylaxis includes baths in NaC1 (0.5 to 2% for various periods of time) and/or potassium permanganate (2 to 5 mg/L for 1 hour to indefinitely). Clinical, systemic, bacterial infections are usually treated with medicated feed containing oxytetracycline (Terramycin) at a rate of 2.5 to 3.5 g/45 kg of fish per day for 10 days. Xu and Rogers (1993a) determined that oxytetracycline injected intraperitoneally into striped bass was cleared from the muscle in 32 days, however, shorter times are required for clearance of Terramycin when applied in the feed. A withdrawal time of 21 days is required for catfish and trout. Resistance of aeromonads, as well as other bacteria, to Terramycin have increased over the past 25 years. Romet-30 (sulfadimethoxine ormetoprim) fed at a rate of 2 to 3 g/45 kg of fish per day is also effective against most systemic bacterial infections, but resistance of some isolates to this drug is emerging. Romet may not be effective against

Streptococcus. It is important to accurately diagnose an infectious disease as soon as possible so that medicated feed can be applied before the majority of the infected population stops feeding. Bacterial infections confined to the skin can be treated with an indefinite treatment of potassium permanganate in the water at 2 to 4 mg/L or 2 to 10 mg/L for 1 hour in tanks. An indefinite treatment with copper sulfate (0.5 to 3 mg/L, depending on water alkalinity) has been used with some success for columnaris (McFarland et al., 1986). Copper sulfate is less expensive than potassium permanganate, but it is also generally less effective for bacterial diseases. Vaccination of fish is becoming another tool to combat bacterial infections, however, few experiments have involved striped bass or diseases that affect this species. Moore et al. (1990) showed that vaccination of channel catfish against F. columnaris reduced losses and the need for chemotherapy; this may be practical for striped bass. Rogers and Xu (1992) reported successful vaccination of striped bass against vibriosis, therefore,

306

vaccination shows some promise as a preventive treatment for V. anguillarum. Some autologous vaccines have been applied to certain populations of striped bass for Streptococcus and Pasteurella, but the results are not conclusive. Nevertheless, vaccination for some bacterial diseases is successful in other cultured fish, salmonids, and to a lesser extent, channel catfish, and should be considered for striped bass. 11.8.3. Fungi Control of fungal infections on striped bass is difficult, therefore, the best approach is to eliminate the predisposing factors that precipitate them. Treatment with formalin, copper sulfate, and potassium permanganate are used, but often with unsatisfactory results. The prognosis offungal infections on fish is such that prevention is the only practical approach. Fungal infections on eggs are preventable by daily treatments with formalin at a rate of about 600 mg/L for a 15 minute flush. 11.8.4. Protozoa Because protozoan parasites are ubiquitous and they cause serious health problems during stressful conditions or following handling, the utilization of the best management practices is important. Insuring good water quality, moderate stocking densities, good nutrition, and proper handling will help prevent most protozoan caused epizootics, or reduce their impact when they do occur. Most ectoprotozoan parasites can be controlled by some chemical bath treatment (Hoffman and Meyer, 1974). Formalin is one of the most widely used broad spectrum parasiticides at 15 to 25 mg/L in static ponds or at a concentration of 1:6,000 (167 mg/L) for 1 hour in tanks or raceways when water temperature is above 20~ Some eviderrce indicates a higher sensitivity of striped bass to formalin than other species of fish, therefore, caution is recommended. Although formalin is not currently an FDA approved chemical for treating striped bass, Xu and Rogers (1993b) showed that striped bass exposed to 25 mg formalin/L for 24 hours, or 250 mg/L for 1 hour, cleared the chemical to normal levels in 96 hours. Copper sulfate at 0.2 to 2 mg/L is also used but the concentration is dependent on the alkalinity of the water. It is generally safe to use 1 mg/L of copper sulfate for every 100 mg/L of alkalinity as CaC03 up to about 400 mg/L of alkalinity. Potassium permanganate at 2 to 4 mg/L indefinitely in static water or 4 to 10 mg/L for 1 hour in tanks may also be used.

Ichthyophthirius multifiliis, and C. irritans require special consideration because their life cycle includes attachment or invasion of the host and free swimming stages. Few chemicals will kill adult/. multifiliis or C. irritans imbedded in the host's epithelium, therefore, elimination of the free swimming form is the logical approach. Formalin and copper sulfate kills the free swimming stage but will not affect imbedded stages, thus, multiple treatments at intervals dictated by water temperature are essential. The intervals may be as short as two days at optimum water temperatures and up to 10 days at cool temperatures. Density of free swimming stages of parasites,/, multifiliis and A. occelatum for example, can be reduced in water supplies or recirculating systems by the use of diatomaceous earth filters, UV radiation, or ozone treatment of the water (Johnson, 1987). Raising the water temperature above the optimum, or increasing water flow in tanks will also reduce the chances of free swimming stages from easily finding a new host. Reduced fish density will lower the impact ofA. ocellatum (Johnson, 1987). Treatment with formalin at 25 mg/L indefinitely or 1:6000 (167 mg/L) for 1 hour in tanks will efficiently treat the clinical disease, however, these treatments require repeated applications to kill the parasites as they leave the host or the free swimming stage. Copper sulfate at 0.2 mg/L as a continuous treatment for two weeks has also been shown to be effective for A. ocellatum (J. P. Hawke, Louisiana State University, personal communication).

307

Experimental vaccines developed for/. multifiliis have shown effective protection in channel catfish against this parasite (Goven et al., 1981; Dickerson et al., 1984). However, none of these preparations are practical at the present time and there has been limited application of these vaccines to non-ictalurid fishes (Wolf and Markiw, 1982). 11.8.5. Helminths There are no approved chemicals for treatment of helminthic parasites of striped bass. However, external infections of monogenetic trematodes (gill flukes and skin flukes) can be removed by application of formalin at 15 to 25 mg/L indefinitely in ponds or at 167 mg/L for 1 hour in flow through or recirculating tanks (Hoffman and Meyer, 1974). Tissue inhabiting helminths have been experimentally treated by bathing in 2 to 4 m ~ L of Praziquantel for up to four hours. This treatment kills many of the yellow grubs in situ and up to 90% of the eye flukes in channel catfish (Lorio, 1989; Plumb and Rogers, 1990). The best control method for the digenetic trematodes is breaking the life cycle by elimination of snails, and blocking access of fish eating birds to the culture ponds. 11.8.6. Leeches Leeches can be removed from infected fish with 0.5 to 1 mg/L of masoten (Dylox, Trichlorfon) indefinitely or by a one hour bath in 3.5% NaC1 where fresh water fish are involved (Post, 1987). Destruction of leech habitat will also reduce their presence. 11.8.7. Crustacea From a management stand point, increased water flow through intensive culture facilities may help flush away the free swimming life stages of crustaceans before they can attach to the host. Crustaceans are difficult to control with chemicals because the host is about as sensitive to most potential chemotherapeutics as the adult parasite. Formalin at the previously suggested concentrations will eliminate juvenile copepodid stages on the gills but will not affect the adults. Masoten at a concentration of 0.25 mg/L indefinitely is a good treatment for parasitic crustaceans. 11.9 SUMMARY Infectious diseases of striped bass and their hybrids do not differ greatly from those of other fishes, however, in intensive culture systems diseases become more acute and may result in high losses. The most severe diseases are caused by bacterial organisms and include members of the genera Flexibacter, Aeromonas, Edwardsiella, Pasteurella, Streptococcus, and Mycobacterium. Fungal infections frequently occur on injured or debilitated striped bass as secondary pathogens. The most severe parasitic diseases include the obligate parasite lchthyophthirius multifiliis and other external protozoa. Parasitic trematodes, nematodes, cestodes and crustacea are fairly common but seldom create serious disease conditions. Striped bass respond negatively to improper handling, stress as result of water quality degradation, and other environmental anomalies. Many serious disease problems can be avoided by good health management practices but drugs and chemicals are sometimes required to arrest infectious diseases. However, it must be emphasized that there are no drugs or chemicals that can be legally used on cultured striped bass at the present time.

308

References

Alderman, D.J., 1982. Fungal diseases of aquatic animals. Pages 189-242, in R. J. Roberts, editor. Microbial diseases of fish. The Society of General Microbiology, Academic Press, London. Anacker, R. L. and Ordal, E.J., 1959. Studies on the mycobacterium Chondrococcus columnaris. I. Serological typing. Journal of Bacteriology, 78: 25-32. Anonymous, 1994. Guide to drug, vaccine and pesticide use in aquaculture. Federal Joint Subcommittee on Aquaculture, Texas Agricultural Extension Service and U. S. Department of A~iculture, Washin~on, DC. B-5085. Austin, B. and Austin, D., 1987. Bacterial fish pathogens: diseases in farmed and wild fish. Ellis Horwood Ltd. Chichester. 364pp. Baloda, S.B., Krovacek, K., Eriksson, L., Linne, T. and Mansson, I., 1995. Detection of aerolysin gene in Aeromonas strains isolated from drinking water, fish and foods by polymerase chain reaction. Comparative Immunology, Microbiology and Infectious Disease, 18:17-26. Baya, A., Toranzo, A.E., Nunez, S., Barja, J.L. and Hetrick, F.M., 1990a. Association of a Moraxella sp. and a reo-like virus with mortalities of striped bass, Morone saxatilis. Pages 91-99, in F.O. Perkins and T.C. Cheng, editors. Pathology in marine science, Academic Press, New York. Baya, A.M., Lupiani, B., Hetrick, F.M., Roberson, B.S., Lukocovic, R., May, E. and Poukish, C., 1990b. Association of a Streptococcus sp. with fish mortalities in the Chesapeake Bay and its tributaries. Journal of Fish Diseases 13: 251-253. Baya, A.M., Toranzo, A.E., Lupiani, B., Li, T., Roberson, B.S. and Hetrick, F.M., 1991. Biochemical and serological characterization of Carnobacterium spp. isolated from farmed and natural populations of striped bass and catfish. Applied and Environmental Microbiology, 57:3114-3120. Baya, A.M., Lupiani, B., Bandin, I., Hetrick, F.M., Figueras, A., Carnahan, A., May, E.M. and Toranzo, A.E., 1992. Phenotypic and pathobiological properties of Corynebacterium aquaticum isolated from diseased striped bass. Diseases of Aquatic Organisms, 14:115-126. Bernardet, J.-F. and Grimont, P.A.D., 1989. Deoxyribonucleic acid relatedness and phenotypic characterization of Flexibacter columnaris sp. nov., nom. rev., Flexibacter psychrophilus sp. nov., nom. rev., and Flexibacter maritimus Wakabayashi, Hikida, and Masumura 1986. International Journal of Systematic Bacteriology, 39: 346-354. Bonn, E.W., Bailey, W.M., Bayless, J.D., Erickson, K.E .and Stevens, R.E., editors., 1976. Guidelines for striped bass culture. Striped Bass Committee of the Southern Division, American Fisheries Society, Bethesda, MD. Carnahan, A.M., 1993a. Aeromonas taxonomy: A sea of change. Medical Microbiology Letters, 2:206-211. Carnahan, A.M., 1993b. Update on current identification methods for clinical Aeromonas isolates. Medical Microbiology Letters, 2:212-216. Chang, P.H. and Plumb, J.A., in press. Effects of salinity on Streptococcus infection on Nile tilapia, Oreochromis niloticus. Journal of Applied Aquaculture.

309

Chitwood, B.M., 1969. The systematics and biology of some nematodes. Pages 223-244/n M. Florkin and B.T. Scheer, editors. Chemical zoology, Academic Press, New York. Colwell, R.R. and Grimes, D.J., 1984. Vibrio diseases of marine fish populations. Helgolander Meeresuntersuchungen, 37: 265-287. Dickerson, H.W., Brown, J., Dawe, D.L. and Gratzek, J.B., 1984. Tetrahymena pyriformes as a protective antigen against Ichthyophthirius multifiliis infections: comparisons between isolates and ciliary preparations. Journal ofFish Biology, 24: 523-528. Dobos, P., Hill, B.J., Hallett, R., Kells, D.T.C., Becht, H. and Teninges, D., 1977. Biophysical and biochemical characterization of five animal viruses with bisegrnented double-stranded RNA genomes. Journal of Virology, 32: 593-605. Dopazo, C.P., Lupiani, B., Rivas, C., Novoa, B., Barja, J.L. and Toranzo, A.E., 1991. Influence of the temperature and cell line on the adsorption and replication of five fish rotaviruses. Fish Pathology, 26: 169-175. Esch, G.W.., Hazen, T.C., Dimock, Jr., R.V. and Gibbons, J.W., 1976. Thermal effluent and the epizootiology of the ciliate Epistylis and the bacterium Aeromonas in association with centrarchid fish. Transactions of the American Microscopy Society, 95: 687-693. Facklam, R., 1980. Streptococci and aerococci. Pages 88-110 in A. Balows, W.J. Hausler, Jr. and J.P. Truant, editors. Manual of clinical microbiology. American Society of Microbiology, Washinpon, DC. FlugeI, R.M., I985. Lymphocystis disease virus. Current Topics in Microbiology and Immunology, 116: 133-150. Frerichs, G.N., 1993. Mycobacteriosis: Nocardiosis. Pages 219-233 in V. Inglis, R.J. Roberts and N.R. Bromage, editors. Bacterial diseases of fish. Blackwell Scientific Press, Oxford. Gaines, J.L. and Rogers, W.A., 1972. Fish mortalities associated with Goezia sp. (Nematoda: Ascaroidea) in central Florida. Proceedings 25th Annual Southeastern Association of Game and Fish Commissioners, 25: 496-497. Goven, B.A., Dawe, D.I. and Gratzek, J.B., 1981. Protection of channel catfish (lctalurus punctatus) against Ichthyophthirius multifiliis (Fouquet) by immunization with varying doses of Tetrahymenapyriformis (Lwoff) cilia. Aquaculture, 23: 269-273. Griffin, B.R., 1992. A simple procedure for identification of Cytophaga columnaris. Journal of Aquatic Animal Health, 4: 63-66. Grizzle, J.M., Mauldin, II, A.C. and Ashfield, C.J., 1992. Effects of sodium chloride and calcium chloride on survival of larval striped bass. Journal of Aquatic Animal Health, 4:281-285. Hastein, T. and Smith, J.E., 1977. A study of Vibrio anguillarum from farms and wild fish using principal components analysis. Journal ofFish Biology, 11: 69-75. Hatai, K., 1989. Fungal pathogens/parasites of aquatic animals. Pages 240-272 in B. Austin and D.A. Austin, editors. Methods for the microbiological examination of fish and shellfish. Ellis Horwood Limited. Chichester. John Wiley and Sons, New York. Hawke, J.P., Plakas, S.M., Minton, R.V., McPhearson, R.M., Snider, T.G. and Guarino, A.M., 1987. Fish pasteurellosis of cultured striped bass (Morone saxatilis) in coastal Alabama. Aquaculture, 65: 193-204.

310

Hazen, T.C., Fliermans, C.B., Hirsch, R.P. and Esch, G.P., 1978. Prevalence and distribution ofAeromonas hydrophila in the United States. Applied and Environmental Microbiology, 36:731-738. Hedrick, R.P., McDowell, T. and Groff, J., 1987. Mycobacteriosis in cultured striped bass from California. Journal of Wildlife Diseases, 23: 391-395. Herman, R.L. and Bullock, G.L., 1986. Patholog), caused by the bacterium Edwardsiella tarda in striped bass Morone saxatilis. Transactions of the American Fisheries Society, 115: 232-235. Hill, J., Evans, W. and Van Den Avyle, M.J., 1989. Species profile: Life histories and environmental requirements of coastal fishes and invertebrates (South Atlantic)-striped bass. U.S. Fish and Wildlife Service and U. S. Army Corps of Engineers. Biological Report 82 (11.118). 35p. Hoffman, G.L., 1967. Parasites of North American freshwater fishes. University of California Press. Berkeley. 486p. Hoffrnan, G.L., 1973. Parasites of laboratory fishes. Pages 645-768 in R.J. Flynn, editor. Parasites of laboratory animals. Iowa State University Press, Ames. Hoffman, G.L., 1978. Ciliates of freshwater fishes. Pages 583-632 in J.P. Kreier, editor. Parasitic protozoa: Intestinal flagellates, histomonads, trichomonads, amoeba, opalinids and ciliates. Academic Press, New York. Hoffman, G.L. and Meyer, F.P., 1974. Parasites of freshwater fishes, a review of their control and treatment. TFH Publications, Neptune, New Jersey. 224pp. Holmes, I.H., 1991. Family Reoviridae. Pages 186-190/n R.I.B. Fraucki, C.M. Fauguet, D.L. Knudson and F. Brown, editors. Classification and nomenclature of viruses. Archives in Virology, Supplement 2. Hughes, J.S., Wellborn, T.L. and Mitchell, A.J., 1990. Parasites and diseases of striped bass and hybrids. Pages 217-238/n R.M. Harrell, J.H. Kerby and R.V. Minton, editors. Culture and propagation of striped bass and its hybrids. Striped Bass Committee, Southern Division American Fisheries Society, Bethesda, MD. Janssen, W.A. and Surgalla, M.J., 1968. Morphology, physiology, and serology of a Pasteurella species pathogenic for white perch (Roccus americanus). Journal of Bacteriology, 96:1606-1610. Johnson, S.K., 1987. Recognition and control of diseases common to grow-out aquaculture of red drum. Pages 8-36 in R.J. Chamberlain, ILL Migot and M.G. Haby, editors. Manual on red drum. Texas Agricultural Extension Service and Sea Grant College Progam, Texas A & M University, Corpus Christi. Kitao, T., 1993. Streptococcal infections. Pages 196-210 in V. Inglis, R.J. Roberts and N.R. Bromage, editors. Bacterial diseases of fish. Blackwell Scientific Press, Oxford. Lauckner, G., 1984. Diseases caused by protophytans (algae). Pages. 169-179 in O. Kinne, editor. Diseases of marine animals, vol IV, part 1, Biologische Anstalt Helgolander, Hamburg. Lawler, A.R., 1977. Dinoflagellate (Amyloodinium) infestation of pompano. Pages 257-264 in C.J. Sinderman, editor. Disease diagnosis and control in North American aquaculture. Elsevier, Amsterdam. Lawler, A.R., 1980. Studies on Amyloodinium ocellatum (Dinoflagellata) in Mississippi Sound: natural and experimental hosts. Gulf Research Report, 6: 403-413. Lorio, W.J., 1989. Experimental control ofmetacercariae of yellow grub (Clinostomum marginatum) in channel catfish. Journal of Aquatic Animal Health, 1:269-271.

311

McAllister, K.W. and McAllister, P.E., 1988. Transmission of infectious pancreatic necrosis virus from carrier striped bass to brook trout. Diseases of Aquatic Organisms, 4:101-108. McFarland, R.D., Bullock, G.L. and McLaughlin, J.J.A., 1986. Effects of five metals on susceptibility of striped bass to Flexibacter columnaris. Transactions of the American Fisheries Society, 115:227-231. Merino, S., Camprubi, S. and Tomas, J.M., 1993. Detection ofAeromonas hydrophila in food with an enzyme-linked immunosorbent assay. Journal of Applied Bacteriology, 74: 149-154. Meyer, F.P. and Bullock, G.L., 1973. Edwardsiella tarda, A new pathogen of channel catfish (lctalurus punctatus). Applied Microbiology, 25:155-156. Meyer, F.P. and Robinson, J.R., 1973. Branchiomycosis: a new fungal disease of North American fishes. The Progressive Fish-Culturist, 33: 74-77. Meyer, F.P. and Schnick, R.A., 1989. A review of chemicals used for the control of fish diseases. Aquatic Science, 1: 693-710. Miyazaki, T. and Kaige, N., 1985. Comparative histopathology of edwardsiellosis in fishes. Fish Pathology, 20: 219-227. Moore, A.A., Eimers, M.E. and Cardella, M.A., 1990. Attempts to control Flexibacter columnaris epizootics in pond-reared channel catfish by vaccination. Journal of Aquatic Animal Health, 2:109-111. Mundt, J.O., 1986. Enterococci. Pages 1063-1065 in H.A. Smeatin, editor. Bergeys manual of systematic bacteriology, vol 2. Williams and Wilkins, Baltimore, MD. Nigrelli, R.F., 1950. Lymphocystis disease and ergasilid parasites in fishes. Journal of Parasitology, 36: 36. Noble, E.R., and Noble, G.A., 1971. Parasitology: The biology of animal parasites. Lea and Febiger, Philadelphia, PA. 617 pp. Noga, E.J., 1992. Fungal and algal diseases of temperate freshwater and estuarine fishes. Pages 278-283 in M.K. Stoskopf, editor. Fish medicine. W.B. Saunders Company, Philadelphia, PA. Overstreet, R.M., 1978. Marine maladies? Worms, germs and other symbionts from the northern Gulf of Mexico. Mississippi-Alabama Sea Grant Consortium. Ocean Springs, MS. Pacha, R.E. and Kiehn, E.D., 1969. Characterization and relatedness of marine vibriosis pathogenic to fish: physiology, serology and epidemiology. Journal of Bacteriology, 100: 1242-1247. Plumb, J.A., 1991. Major diseases of striped bass and redfish. Veterinary and Human Toxicology, 33, Supplement 1: 34-39. Plumb, J.A., 1994. Health maintenance of cultured fishes: Principal microbial diseases. CRC Press, Boca Raton, FL. 254 pp. Plumb, J.A. and Rogers, W.A., 1990. Effect ofDroncit (praziquantel) on yellow grubs Clinostomum marginatum and eye flukes Diplostomum spathaceum in channel catfish. Journal of Aquatic Animal Health, 2: 204-206.

312

Plumb, J.A., Schwedler, T.E., Mitchell, A.J., Rawson, M. and Wellborn, Jr., T.L., 1990. Diseases of warmwater fish in the southeastern United States. Pages 177-183, in R.O. Smitherman and D. Tave, editors. Proceedings: Auburn symposium on fisheries and aquacultures. Auburn University, Auburn, AL. Post, G., 1987. Textbook of Fish Diseases. TFH Publications, Neptune City, NJ, 288p. Robohm, R.A., 1983. Pasteurella piscicida. Pages 161-175 in D.P. Anderson, M. Dorson and P. Dubourget, editors. Antigens of fish Pathogens. Collection Fondation Marcel Merieux, Lyon. Rogers, W.A. and Xu, D., 1992. Protective immunity induced by a commercial vibrio vaccine in hybrid striped bass. Journal of Aquatic Animal Health, 4: 303-305. Sakanari, J.A., Reilly, C.A. and Moser, M., 1983. Tubercular lesions in Pacific coast populations of striped bass. Transactions of the American Fisheries Society, 112: 565-566. Samal, S.K., Dopazo, C.P., McPhillips, T.H., Baya, A., Mohanty, S.B. and Hetrick, F.M., 1990. Molecular characterization of rotavirus-like virus isolated from striped bass (Morone saxatilis). Journal of Virology, 64: 5235-5240. Samal, S.K., Dopazo, C.P., Subramanian, K., Lupiani, B., Mohanty, S.B. and Hetrick, F.M., 1991. Heterogeneity in the genome RNAs and polypeptides of five members of a novel group ofrotavirus-like viruses isolated from aquatic animals. Journal of General Virology, 72:181-184. Schutz, M., May, E.B., Kraeuter, J.N. and Hetrick, F.M., 1984. Isolation of infectious pancreatic necrosis virus from an epizootic occurring in cultured striped bass, Morone saxatilis (Walbaum). Journal of Fish Diseases, 7: 505-507. Shotts, E.B., 1991. Selective methods for fish pathogens. Journal of Applied Bacteriology, Symposium Supplement, 70: 75-80. Shotts, E.B. and Teska, J.D., 1989. Bacterial pathogens of aquatic vertebrates. Pages 164-186 in: B. Austin and D.A. Austin, editors. Methods for the microbiological examination of fish and shellfish. Ellis Horwood, Ltd. Chichester. Sinderman, C.J. and Lightner, D.V., 1988. Disease diagnosis and control in North American marine aquaculture. Second Edition. Elsevier, Amsterdam Snieszko, S.F., Bullock, G.L., Hollis, E. and Boone, J.G., 1964. Pasteurella sp. from an epizootic of white perch (Roccus americanus) in Chesapeake Bay tidewater areas. Journal of Bacteriology, 88:1814-1815. Spall, R.D. and Summerfelt, R.C., 1970. Life cycle of the white grub, Posthodiplostomum minimum (McCallum, 1921: Trematoda, Diplostomatidae), and observations on host-parasite relationships of the metacercaria in fish. Pages 218-230 in S. F. Snieszko, editor. A symposium on diseases of fishes and shellfishes. American Fisheries Society, Special Publication No. 5, Bethesda, MD. Subramanian, K., Lupiani, B., Hetrick, F.M. and Samal, S.K., 1993. Detection of aquareovirus RNA in fish tissues by nucleic acid hybridization with a cloned cDNA probe. Journal of Clinical Microbiology, 31:1612-1614. Thoeson, J., editor, 1994. Blue book. version 1: suggested procedures for the detection and identification of certain fmfish and shellfish Pathogens. 4th Edition. Fish Health Section, American Fisheries Society. Bethesda, MD.

313

Toranzo, A.E., Barja, J.L., Lemos, M.L. and Hetrick, F.M., 1983. Stability of infectious pancreatic necrosis virus (IPNV) in untreated, filtered and autoclaved estuarine water. Bulletin of the European Association of Fish Pathologists, 3: 51-53. Toranzo, A.E., Novoa, B., Baya, A.M., Hetrick, F.M., Barja, J.L. and Figueras, A., 1993. Histopathology of rainbow trout, Oncorhynchus mykiss (Walbaum), and striped bass, Morone saxatilis (Walbaum), experimentally infected with Carnobacterium piscicola. Journal of Fish Diseases, 16:261-267. van Duijn, Jr., C., 1973. Diseases of fishes. Charles C. Thomas Publishers, Springfield. 372pp. Wakabayashi, H., Hikida, M. and Masumura, K., 1986. Flexibacter maritimus sp. nov., a pathogen of marine fishes. International Journal of Systematic Bacteriology, 36: 396-398. Wechsler, S.J., Woods, Kraeuter, J.N., Hetrick, F.M. and McAllister, P.E., 1987. Transmission of infectious pancreatic necrosis virus in striped bass, Morone saxatilis (Walbaum). Journal of Fish Diseases, 10: 29-34. Winton, J.R., Lannan, C.N., Fryer, J.L., Hedrick, R.P., Meyers, T.R., Plumb, J.A. and Yamamoto, T., 1987. Morphological and biochemical properties of four members of a novel group ofreovimses isolated from aquatic animals. Journal of General Virology, 68: 353-364. Wolf, K., 1988. Fish viruses and fish viral diseases. Cornell University Press. Ithaca, NY. 476pp. Wolf, K. and Markiw, M.E., 1982. Ichthyophthiriasis: immersion immunization of rainbow trout (Salmo gairdneri) using Tetrahymena thermopila as a protective immunogen. Canadian Journal of Fisheries and Aquatic Sciences, 39: 1722-1725. Wootten, R., 1989. The parasitology of teleosts. Pages 242-288 in V. Inglis, R.J. Roberts, and N.R. Bromage, editors. Bacterial diseases of fish. Blackwell Scientific Press, Oxford. Xu, D. and Rogers, W.A., 1993a. Oxytetracycline residue in hybrid striped bass muscle. JoumaI of the World Aquaculture Society, 24: 466-472. Xu, D. and Rogers, W.A., 1993b. Formaldehyde residue in striped bass muscle. Joumal of Aquatic Animal Health, 5:306-312.

This Page Intentionally Left Blank

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 9 1997 Elsevier Science B.V. All rights reserved.

315

Chapter 12

Economics and Marketing Douglas W. Lipton and Conrado M. Gempesaw, II

12.1 INTRODUCTION Before the advent of hybrid striped bass aquaculture the market relied on a wild striped bass fishery in the Middle Atlantic and Chesapeake Bay states. As early as 1888, the harvest in these two regions amounted to about three million pounds (National Marine Fisheries Service, 1990). Striped bass harvests peaked in 1973 at 14.7 million pounds when the fishery was valued at the harvest level (ex-vessel) at $4.7 million (National Marine Fisheries Service, 1992). Unfortunately, this harvest level, coupled with extended periods of recruitment failure represented severe over fishing, and landings declined dramatically that by 1983, striped bass harvest was reduced to 1.7 million pounds. In 1985, fishing moratoriums were put into effect in Maryland and Delaware to aid in population rebuilding, and in 1986 and 1987 in New York and New Jersey because of PCB contamination. By 1991, harvests hit a recorded low of 254 thousand pounds. However, the moratoriums have returned dividends with the Atlantic States Marine Fisheries Commission now declaring the population as being recovered. Limited fishing is now allowed, with the 1994 striped bass harvest at 1.7 million pounds with an ex-vessel value of $2.8 million (National Marine Fisheries Service, 1995). Events in the wild fishery for striped bass set the stage for the development of aquaculture of striped bass and its hybrids. The early history of commercial striped bass aquaculture was reviewed by Van Olst and Carlberg (1990), with the earliest commercial production of about 10 thousand pounds dating back to 1973. It was not until 1987 when approximately 395 thousand pounds were produced that there was any substantial production. Current production figures are difficult to come by. Bush and Anderson (1993) conducted a survey of Northeast (from Maine to Maryland) aquaculture producers that estimated 1992 production at 947 thousand pounds, with a farm gate value of $2.3 million. Rhodes (1993) estimated South Carolina production in 1992 to be around 500 thousand pounds, valued at $1.1 million (see Chapter). An earlier study (Rhodes and Sheehan, 1991) conducted for the Striped Bass Growers' Association, estimated that nationwide production of hybrid striped bass in 1992 would be about 7.7 million pounds (Table 1.2, Chapter 1). The most recent production survey revealed that the 1994 production exceeded 8 million pounds (Carlberg and Massingill, 1995; Table 1.2 Chapter 1). These production figures suggest that hybrid striped bass production has grown significantly since 1987, and production is a significant percentage of historic striped bass fishery production levels. Striped bass production still has a long way to go to equal the domestic catfish aquaculture production of 459 million pounds in 1993 (USDA, 1994).

12.2 PRODUCTION COSTS Information on striped bass production costs is extremely limited, reflective of the relatively small size of the industry, its dispersion, and its developing nature. AquacultureMagazine's 1994 Buyers Guide and Industry Directory lists 41 firms involved in culture of market sized striped bass and hybrids. Familiarity with the scant literature makes one wary of its accuracy because of the lack of independence of the various studies. One of the earliest studies (Brown et al., 1988) was based on a couple of operations that had not yet produced market fish in North Carolina, and used published data on catfish production costs. A subsequent evaluation by Strand and Lipton (1989) modified the Brown et al. (1988) data for production in Maryland, and further

316

production cost analyses by Gempesaw et al. (1993) relied heavily on the Strand and Lipton (1989) study. These studies share a common feature which is a reliance on "engineered" cost estimates as opposed to a statistical analysis of real production costs. Because of this lack of actual production performance information, Gempesaw et al. (1993) relied on a stochastic model of financial performance which uses Monte Carlo analysis to provide meaningful risk information to prospective aquaculturists. This approach is discussed further in Section 12.4. 12.2.1 Costs of Production Systems In terms of numbers of operations, evidence suggests that most striped bass aquaculture operations are conducted in open or static pond systems. Rhodes and Sheehan (1991) study estimated that 41% of producers used ponds, 29% used recirculating tank systems, 25% grew fish in raceways and 5% used cages or net-pens. Bush and Anderson's (1993) northeast aquaculture survey found, in terms of production volume, 42% were produced in open ponds and 40% in recirculating systems. They also found that producers expected to significantly increase the share coming from recirculating systems. The reader should keep in mind that these percentages are of a relatively small level of production so that one or two firms producing in one type system or another can greatly affect the ratios. 12.2.1.1 Hatcheries In addition to the grow-out systems discussed above, there are hatchery production systems to produce fry and fingerlings for either an in-house vertically integrated operation or to supply other producers. Of all the production systems for striped bass and hybrid culture, the least amount of economic analysis has been performed on hatchery operations. Gempesaw et al. (I 993), in their attempt to look at degrees of vertical integration in hybrid striped bass production, "engineered" costs for a hatchery facility capable of producing about 36 million eggs or, with 25% survival, about 9 million fry. The capital investment for this operation, on about 5 acres, was less than $100,000. The more common scenario is an integrated hatchery-fingerling producer producing phase I and some phase II fingerlings for sale. The Gempesaw et al. (1993) version of this farm produces an average of 2.3 million phase I fingerlings on an additional thirty acres of ponds. The investment cost for this operation was almost $570 thousand, the highest of any integration scenario described. The major expense in this scenario is the numerous small ponds that must be constructed for the fry and fingerlings as opposed to fewer larger ponds, which elevates the relative costs. 12.2.1.2 Static ponds The cost of static pond grow-out systems is fairly well understood, given that it emulates the more well-established catfish culture operations. Variability in land acquisition costs from farm to farm, variations in topography and water supply make it difficult to generalize about what pond construction costs will be. Generally, costs of pond construction will rely directly on the amount of earth movement needed to construct levees and obtain the proper pond depths. The number and size of wells will also vary greatly depending on the size, quality and depth of the local aquifer. If surface waters are available, their use can decrease the expense of maintaining the pond water supply. Pond construction costs will also vary depending on the individual pond size and layout. For example, a configuration where ponds share levees saves on earthmoving costs. To estimate pond construction costs you need to determine the total linear feet of levees, determine the width of the levees and the slope to estimate the total cubic yardage of earth that will have to be moved,

317

then multiply this value times the cost of moving a cubic yard of soil. The layout of water pipes, wells, pumps, and drainage structures would also be determined at the same time. For a 30 acre operation, Brown et al. (1988) calculated that pond construction and equipment costs, amortized over a ten year period, contributed about $0.15 per pound to the cost of growing fish. Strand and Lipton (1989) computed the cost to be about $0.17-$0.30 for a 25 acre operation in Maryland, and $0.21-$0.35 if the cost of land acquisition was included. 12.2.1.3 Tanks The published literature on the economics of striped bass production in tank systems is almost nonexistent. A number of tank systems are in use nationwide to grow hybrid striped bass, but there is little documentation of their economic production and performance. Gempesaw et al. (1992) "engineered" costs for a small (50,000 pounds per year) recirculating tank system requiring a $271 thousand investment. In the relatively successful Aquatic Systems Inc. (now Kent Sea Farms) operation in California, a geothermal aquifer supplies water for the tank systems (Van Olst and Carlberg, 1990). 12.2.1.4 Net-pen and cage culture Many existing farm ponds cannot be seined to harvest fish, and would be unsuitable for open pond culture. These ponds lend themselves to the placement of cages or net-pens. Because it is assumed that the pond is already in place, the only major cost is construction of the cages. This can be done with inexpensive materials. Strand and Lipton (1989) estimated that a 64 cubic foot cage could be constructed for less than $200. In order to stock the cages at commercial densities, supplemental aeration would be required. The cost of the aerator plus the installation of electric service to the pond would add an additional $0.10 per pound to the cost of production, but the increased production would lower the contribution of the cage costs per pound of fish produced from $0.09 to $0.06. These costs are for production in a 5-acre pond at a production level of 2,500 pounds per acre. Another application of cage or net-pen culture is the culture of striped bass in open estuarine systems. System requirements would be similar to the net-pens or cages placed in ponds, but might require some modification for tethering and access to the pens. Although Asian countries have been culturing marine species (i.e., yellowtail) in cages for decades, Norwegian's first developed the net-pen technology to produce Atlantic salmon, and that technology has been adopted in the United Kingdom, the United States, Chile, Canada and other countries with suitable growing conditions for salmon and other species. Bettencourt and Anderson (1990) documented the costs of these salmon net-pen operations, and costs ranged from $25,000 for four, self-assembled pens to $2 million for a 40 pen system. Currently, there are two small scale net-pen production sites for striped bass in Maryland, but no data on economic costs and retums are available yet. 12.2.2 Operating Costs Operating costs are those that vary from period to period based on the level of production of the aquaculture facility. The lack of a history of commercial scale striped bass production precludes econometric analysis of operating costs. Instead, information has relied on engineered budgets. As a result there can be no testing of hypotheses regarding economic items of interest such as returns to scale and input substitutability. The major operating cost items for aquaculture products are typically feed costs and the cost of the seed stock (i.e., the starting point in the life cycle of the organism being cultured). The contribution of feed

318

and seed costs to production are a direct result of the production relationship, which may vary among production systems, species being cultured, sizes stocked and harvested, and feeding rates. Whenever figures are quoted regarding production costs in aquaculture, there are underlying production assumptions that should be evaluated for their accuracy, such as survival and feed conversion rates. 12.2.2.1 .Stocking costs Stocking costs will depend on what point in the striped bass life cycle production is being initiated. For a hatchery, stocking entails obtaining and maintaining brood stock. Typically, for grow-out operations, stocking of phase I or phase II fish is performed, although some might start with fry. Fry prices are usually about $0.01 a piece, and fingerling prices about $0.10 an inch. Thus, phase I fingerlings can be purchased at $0.20-$0.30 and phase II fingerlings at $0.60-$0.80 apiece, with price breaks for large quantity purchases. The contribution of stocking to the final production cost will depend on the of the fry or fingerling price (or the production cost of fry if the operation includes a hatchery), the average weight of the harvested fish, and the survival rate from stocking to harvest: P r i c e seed C ostseed-

(Weighth .... ~

Survival) .

Using this formula, stocking phase I fingerlings costing $0.25 each, with 70% survival to grow-out at an average of 1.25 pounds, ends up contributing $0.29 per pound to the cost of the final product due to the mortality effect. If survival can be increased to 90%, the cost drops 24% to $0.22 per pound. If in addition to increased survival, fingerling production costs, and hence the fingerling price, falls to $0.15, stocking costs contribute only $0.13 per pound to final production costs. It is easily seen how technological efforts to increase survival and decrease the cost of seed stock reduces the overall production costs. 12.2.2.2 Feed costs Feed costs are typically the greatest operating expense for aquaculture enterprises. The competitiveness of striped bass and hybrid culture will depend greatly on obtaining a low cost feed that is efficiently converted into final product weight by the fish. The feed price will depend on its ingredients, higher protein feeds being more expensive. In addition, the contribution of feed costs will depend on the survival of striped bass in the production system, and the timing of that mortality. Mortality that occurs early in the production cycle has little impact on feed costs because the quantity fed to fish at that point is small. If mortality occurs just before harvesting, then most of the investment in feed has already been made, and the cost of feed per pound of fish ultimately harvested is much greater. If we assume that mortality is uniform over the growing cycle, the following formula can be used to estimate feed cost contributions to fish production:

Pricefeed X WeightGain X ConversionRatio Costfee d -

1 - [0.5 X (1-Survival)]

Using the above formula, a typical calculation for striped bass might entail a survival rate of 70% for phase I fingerlings to grow-out, a feed conversion ration of 2.5:1, a weight gain of 1.25 pounds and feed costs of $0.20 per pound. The result is a contribution to production costs due to feed of $0.74. Changing any of those variables can substantially raise or lower costs. A production technology and feed that results in 80% survival, a 2:1 feed conversion rate and a feed that costs only $0.15 per pound lowers the contribution of feed costs by 43% to just $0.42 per pound.

319

12.2.2.3 Other costs Other important operating costs include labor, utilities, and water monitoring. Labor costs are typically treated as a variable cost, with more labor needed, the greater the production. Some labor may be concentrated during specific periods of the operation such as harvesting, and it may be more effective to contract for this activity. An important fixed cost is that paid for management. In an owner-operator situation, this might not actually entail a cash payment for management, but the opportunity cost of the manager's time -- what the manager could earn in alternative employment -- should be considered in evaluating the economic performance of the aquaculture operation. Often the financial performance of such an operation will be reported as returns to management, so care must be taken in comparing this with other enterprises where management costs are explicitly considered. 12.3 DEMAND

One of the most naive assumptions that can be made about developing aquaculture industries is that the price of the product will be the same several years in the future as it is today. The assumed price of the product in future years of production will determine the rate of return on the investment. Thus, wrong investment decisions may be made by not using all the information available on potential future prices. The major aquaculture products that have experienced large expansions in production over the past decade (i.e., shrimp, salmon, catfish) have all experienced real price declines. An understanding of seafood demand and how the seafood market works in general is essential for the successful aquaculturist. 12.3.1 Seafood Demand There is a widely held expectation about the extent of expanding seafood demand that is not supported by the available data. In part this perception was fueled by a significant expansion in U.S. per capita seafood consumption during the mid- 1980's (Figure 12.1). From 1980 to 1987, per capita consumption increased from 12.5 to 16.2 pounds. Since its peak, however, per capita consumption has fallen and stood at 15.2 pounds in 1993. This is less than the 1985 figure and only 13% greater than per capita consumption in 1978. Strikingly, today's per capita seafood consumption is only 32% greater than it was over 80 years ago in 1913. Although it is clear that there was a significant gain in seafood per capita consumption in the 1970s and 1980s, there is little evidence that this rate of expansion is being sustained. Still, some experts predict that consumption of seafood products will almost double during the next three decades resulting in shortages in seafood products (Dowdell, 1990; Rogness and Weddig, 1991). With these caveats, we can still state that the outlook for seafood demand is positive. It is showing a long-term increase, and U.S. and world population growth will continue the need for increases in seafood production in order to maintain current price levels. In addition, per capita seafood consumption in other industrialized countries is substantially higher than U.S. seafood per capita consumption (Bjorndal, 1990). This correlation between industrialization and seafood demand suggests that as the economy improves in developing countries, their demand for seafood may increase. Demand, however, entails looking at both the quantity consumed and the relative price of seafood. Prices at the consumer level have tended to increase greater than the overall rate of inflation, leading to a real price increase. If demand is not increasing, then a real price increase will lead to a fall in per capita consumption, such as was observed from 1987-1993. Another issue is that retail price increases for seafood have not always translated into price increases for the producer. While retail seafood prices increased by 14% from 1988-1993, producer prices increased only 5%. More relevant to striped bass and hybrids producers, the producer price index for finfish actually fell 3.7% over that time period.

320

Figure 1. U.S. Seafood Per Capita Consumption, 1909-1993

20

Pounds per capita

1909

14

19

24

29

34

39

44

49

54

59

64

69

74

79

84

89

94

Year

12.3.2 The Striped Bass Market Most of what is known about the striped bass market is based on studies before the collapse of the market in the mid to late 1980's. During that period the market became very confused with the plethora of regulations regarding the sale and possession of striped bass and striped bass hybrids in the various states. Some of the restrictions on marketing where due to concerns about PCB contamination of wild striped bass from the Hudson River. Some states were concerned about enforcing a fishing moratorium and forbade the possession of any striped bass or hybrid. Thus, aquaculturists were prevented from initiating the development of their industry by regulations intended to protect wild populations. 12.3.2.1 The wild fishery Yamashita (1981) and Adriance (1982) studied both the marketing practices of striped bass agents and price determination in the wholesale and ex-vessel markets. Yamashita, studying the 1972-1978 period, found New York City's Fulton Market to be an important element in the marketing of striped bass. She found that much of the reported Chesapeake landings were marketed there, especially in the months of April, May and June. Currently, Chesapeake commercial fishing is not allowed in those months. Fifty percent of the Chesapeake wholesalers reported using Fulton prices to establish the prices they paid fishermen. All of them also differentiated their prices based on size, with sometimes small fish and sometimes large fish commanding a premium. It is not known to what extent these pre-moratorium preferences hold today in a highly regulated wild fishery, and an aquaculture industry producing four times the amount of the wild fishery. Adriance (1982) and Norton et al. (1983) estimated a statistical model of price determination for the 1976-1979 period. In 1988 dollars, Fulton and Baltimore wholesale market prices dropped approximately 4.8 cents for each 10,000 pounds marketed.

321

12.3.2.2 Hybrid striped bass Strand et al. (1988) used existing information on the wild striped bass market to determine the expected effect of increased cultured striped bass production on prices. Implicit in this analysis is the assumption that hybrid striped bass is a perfect substitute for striped bass and would trade in the same market channels. An eight block system of equations was developed for striped bass price:

Price i:ao+a~(Production/Population) +a2(~,y~roductionJPopulation ) +a3(U.S. Per Capita Income) +a4PCB

Where, I is the subscript for the state, production is the ith state's striped bass production, population is East Coast and South Atlantic human population and PCB is a dummy variable representing closure of the New York market. Based on the results of this analysis, it was concluded that the price distribution of hybrid striped bass would fall in the $2.00-$3.00 per pound range. This is consistent with the actual prices as have been reported by the Maryland Department of Agriculture's wholesale market report for the past several years. 12.4 FINANCIAL PERFORMANCE OF STRIPED BASS AQUACULTURE Given the uncertainty in the production economics and market for striped bass and hybrids aquaculture, point estimates of financial performance based on deterministic analysis can be very misleading. The computer program Aquasim (Gempesaw et al., 1992; Gempesaw et al., 1993), although generic for different aquaculture species, was initially developed to analyze striped bass hybrid aquaculture. Aquasim is a farm-level dynamic and stochastic capital budgeting computer simulation model, and has been used for analysis of striped bass financial performance under a variety of scenarios and assumptions about prices and production performance. In the two published studies, one comparing tank and pond production (Gempesaw et al., 1992) and the other comparing vertical integration in pond production (Gempesaw et al., 1993), results were characterized by high sensitivity to the price and production scenarios, as well as great variability of results within a scenario. Pond production of hybrid striped bass had a higher internal rate of return than production in recirculating tanks in all scenarios modeled. However, in both production systems the coefficient of variation of the discounted net cash income was close to or exceeded the expected value, indicating that there was a significant probability that these operations would fail, economically. Similar results were found when comparing different levels of vertical integration for the farm. In fact, for all but the extremes of only a hatchery operation and only a fully-integrated operation, the coefficient of variation for discounted net cash income exceeded the expected value by several orders of magnitude. For the most typical operation of raising phase I fingerlings to market size on a 30-acre farm, the internal rate of retum was 29% with a coefficient of variation of 51.6%. The discounted net cash income was $48,500 with a coefficient of variation of $222,550. This huge uncertainty in financial performance has probably contributed to an unwillingness in the investment community to become involved heavily in striped bass aquaculture. Although the figures demonstrate the potential for substantial profit and retum on investments, this is accompanied by significant risk. There has been no published verification based on actual performance of real operations.

322

12.5 THE HYBRID STRIPED BASS MARKET Given the risk involved in investing in the developing hybrid striped bass industry, it is essential that attention be paid to marketing as well as production. The information presented here reviews the existing information on hybrid striped bass marketing. 12.5.1 Hybrid Striped Bass Marketing Studies Numerous studies have been undertaken to investigate the market potential of hybrid striped bass. Before these studies can be reviewed, it is important to understand the various marketing channels for aquaculture products. These various marketing channels include the wholesaler, retailer, restaurants, and finally the consumers themselves. Studies on hybrid striped bass marketing have generally focused on various parts of these marketing channels. Harvey et al. (1990) conducted a study dealing with two distinct market segments, i.e., restaurants and wholesalers across the Mid-Atlantic region with major emphasis on the restaurant market. Two market surveys were conducted. The restaurant survey was designed to obtain specific information about the restaurant market concerning familiarity with hybrid striped bass, present and future use, product form, price, and size. The authors reported that a total of 13,460 restaurants located in New York, Pennsylvania, New Jersey, Virginia, Massachusetts, Maryland, Connecticut, Washington, D.C., Rhode Island, and Delaware were surveyed with a response rate of less than eight percent. The wholesaler survey consisted of 894 wholesalers in Virginia, Maryland, Washington, D.C., New York, and Pennsylvania with a response rate of also less than eight percent. One of the primary objectives of the Harvey et al. (1990) study was to investigate whether hybrid striped bass could occupy a "market niche" in restaurants located in the Northeast region. Of the 948 restaurants responding to the familiarity question, only 22 percent answered with a positive response. Among the factors that would encourage restaurants to offer hybrid striped bass were the possibility of high margins and consistency in supply. Most of the respondents indicated that they preferred product size to be less than two pounds with fillet as the most popular product form. Almost a third of the restaurants responding stated that they were willing to pay between $2.50 to $3.00 per pound and preferred individual negotiations with hybrid striped bass producers in determining the final price. Similar questions were also asked of Northeast seafood wholesalers. Close to two-thirds of the wholesalers responded that they were familiar with hybrid striped bass with the primary sales area identified as New York followed by Maryland. Again the top two factors cited by the wholesalers for buying hybrid striped bass were the possibility of consistent supply and high margins. At least forty percent of the wholesalers preferred a two pound fish. At least a third of the wholesalers noted that they were willing to pay between $2.08 to $3.36 per pound. An important conclusion from the study was that although restaurants indicated a strong willingness to offer the product, less than a fourth of the respondents were familiar with the product. Furthermore, the wholesalers, though more familiar with the product, seemed to find that market demand was not strong enough. A marketing study was conducted by Wirth et al. (1990) to investigate the purchase decision of MidAtlantic seafood buyers when purchasing farm-raised hybrid striped bass. A conjoint design survey questionnaire was developed and sent to wholesalers, retailers, and restaurants located from New York down to North Carolina. Over 2,400 surveys were sent out with a response rate of about 12 percent. The authors found that wholesalers and retailers were more concerned with price while restaurants gave product form more importance in their purchase decisions. This implies that wholesalers and retailers are more price sensitive

323

than the restaurant market. The fillet form was also found to be the preferred product form with the retail and wholesale markets having a preference for a two pound fish. Fuller et al. (1990) conducted a study on the consumer acceptance of hybrid striped bass in a Mississippi Gulf Coast restaurant. The fish were served over an eight-week period starting in November 1988. A total of 192 useable responses were obtained from consumers who ordered the fish during that time period. Approximately 40 percent of the consumers rated hybrid striped bass as better than their favorite fish. The most popular fish for the consumers who participated in the survey were trout, redfish, red snapper, flounder, catfish, and grouper in that order. Consumers were also asked whether they would order hybrid striped bass again at prices comparable to their favorite fish. Nearly 86 percent of the respondents answered that they would order the fish again. As a measure of quality, consumers rated appearance as very important followed by freshness, texture, and flavor. D'Souza et al. (1993) reported a study on the marketing potential of aquaculture products in West Virginia by conducting a survey of consumer, restauranteur, and wholesaler attitudes. The study dealt with several aquaculture products including hybrid striped bass. A random sample of 2,000 West Virginia households were asked to participate in the survey with a response rate of almost 16 percent. A total of 250 restaurants in West Virginia were identified as serving seafood products. The restaurants were sent a survey questionnaire and yielded a response rate of almost 13 percent. In addition, two out of the nine seafood wholesalers in the state accepted the invitation to participate in the marketing study. At least half of the consumer respondents indicated that they have eaten farm-raised fish and identified freshness and/or appearance as the most important factor they consider when buying fish products. Over half of the respondents answered that they are familiar with farm-raised trout, but only 23% responded that they have heard of hybrid striped bass. The respondents stated that they have eaten catfish (78%) and trout (58%) but only 13% have actually eaten hybrid striped bass. The restaurant managers identified consistent quality and dependable supply as important factors in influencing their purchase decisions of farm-raised fish products. Less than one-third of the respondents were offering farm-raised fish in their restaurants citing "availability" and "never heard of it" as the two most common reasons for not offering the product. In the specific case of hybrid striped bass, 81% of the respondents indicated no interest in offering the product. What is more surprising is that none of the restaurants offered hybrid striped bass. Trout, shrimp, scallops, salmon, and catfish were the most popular fish products offered by the restaurants. The wholesalers supported these findings by identifying shrimps, scallops, flounder, cod, orange roughy, and catfish as their best selling species. In 1993, the Northeastem Regional Aquaculture Center funded a regional study entitled "Altemative Marketing Options to Improve the Profitability of Northeast Aquaculture Industry." Among its objectives was to gather information regarding consumer attitudes and preferences for finfish products, one of which was hybrid striped bass. A mail survey questionnaire was developed and sent to 5,000 randomly selected households in the Northeast region. The project was supervised by Cathy Wessels, University of Rhode Island, Alberto Manalo, University of New Hampshire, and Conrado Gempesaw, University of Delaware. The states included in the survey were Maine, New Hampshire, Vermont, Massachusetts, Rhode Island, Connecticut, New York, Pennsylvania, Maryland, New Jersey, Delaware, Virginia and West Virginia, and Washington, D.C. A total of 1,529 responses were received for the finfish survey yielding a response rate of over 30%. Several studies have been undertaken using this data set. For example, Gempesaw et al. (1985) reported that the most common reasons for not consuming hybrid striped bass were: 1) the respondents' lack of familiarity of the species, 2) the respondents had never seen this fish in stores and, 3) a dislike for its taste.

324

Nauman et al. (1995), using the same survey data, used the evoked set framework to analyze the demand for finfish products, which included hybrid striped bass. Limited dependent variable estimation techniques were used to model the effect of socio-economic variables and other explanatory factors on consumer experiences, perceptions, preferences, and ultimate choice for hybrid striped bass. The model was developed showing that consumer seafood purchase decision is formed in a manner suggested by a four equation recursive system:

(1) (2) (3) (4)

Experience =f(Socioeconomic/Demographic Factors) Perception =f(Socioeconomic/Demographic Factors, Experience) Preference =f(Socioeconomic/Demographic Factors, Experience, Perception) Choice =f(Socioeconomic/Demographic Factors, Experience, Perception, Preference)

The experience variable was represented by a binary variable assuming a value of one if the respondent was a frequent purchaser of seafood products in general (weekly to at least four times per year) and was set to zero if the respondent was an infrequent purchaser of hybrid striped bass (less than four times a year to never purchased). The perception variable represented four classifications of how hybrid striped bass was perceived by the respondents. The four categories were health and nutritional value, adds variety to diet, good taste, and easy to prepare. The four categories were also specified as a binary variable and given a value of one if the respondent perceived hybrid striped bass as having a specific attribute and zero otherwise. The preference variable was represented by the number of individuals in a household who have consumed hybrid striped bass and was specified as a continuous variable. Finally, the choice variable was specified as a binary variable with a value of one if the individual has purchased hybrid striped bass and zero otherwise. The results show that over 77% of the respondents were a frequent purchaser of seafood products in general. The variables that were found to significantly affect seafood purchases were education (higher education had a positive effect), race (non-caucasians tend to buy more seafood), location of residence (the farther from the ocean, the lower the incidence of seafood purchases), and stories from the media about seafood (positively contributed to seafood purchases). In the perception model, only 9% of the respondents indicated that the health and nutritional value was a major perceived reason for their buying hybrid striped bass. The variables that influenced health and nutritional value as a perceived attribute for hybrid striped bass were race (non-caucasians having a positive impact), location of residence and knowledge that the fish is farm-raised (positive impact), and experience (a consumer who has purchased seafood products in general will more likely perceive hybrid striped bass as having a health and nutritional value). Only 11% of the respondents indicated that variety of diet was one of the important aspects of their decision to eat hybrid striped bass. The knowledge that hybrid striped bass is farm-raised and experience with seafood products significantly affected this particular perception. Similarly, around 10% of the respondents answered affirmatively when asked whether taste was a major reason for consuming hybrid striped bass. This perception was significantly influenced by age (consumers between the ages 20 to 40), knowledge that hybrid striped bass is farm-raised, and having experience with seafood products. In the easy to prepare perception model, only 5% of the respondents indicated that hybrid striped bass was easy to prepare. This perception was affected significantly by age (consumers between 40 and 60 years of age did not agree with this perception), knowledge that hybrid striped bass is farm-raised and having experience with seafood products. At least 23% of the respondents indicated that one or more individuals in their household has consumed hybrid striped bass. The variables that were found to positively influenced the respondents' preference for hybrid striped bass were knowledge that hybrid striped bass is farm-raised, having a perception

325

that hybrid striped bass adds variety to diet and is healthy and nutritious, and good taste. In the choice model, approximately 77% of the respondents noted that they have not purchased hybrid striped bass. Thus, hybrid striped bass is currently not part of the evoked set of a large proportion of the respondents. Caucasians were found to be less likely to purchase hybrid striped bass while the attributes of farm-raised, healthy, good taste, and easy to prepare, and having experienced seafood in general contributed to the likelihood of the respondents in consuming hybrid striped bass. In a much larger study, Nauman (1995) compared three finfish products (salmon, trout, and hybrid striped bass) and three shellfish products (mussels, clams, and oysters) using the same survey data set. By far, hybrid striped bass was found to be the least popular among the respondents. In fact, hybrid striped bass rated only slightly higher than tilapia, which was dropped from the study due to insufficient response. It is interesting to note that these two finfish products are incidentally very popular with aquaculture producers in the Mid-Atlantic region. A survey of potential and current producers in Delaware and Maryland conducted by Bacon et al. (1993) showed that catfish followed by hybrid striped bass, trout, crawfish, and tilapia were the preferred species for culture by current and potential producers. Thus, awareness and acceptance of hybrid striped bass by consumers is going to be essential to the successful development of this industry. 12.5.2 Marketing Implications It has been optimistically projected that per capita consumption of seafood products will continue to increase in the future. Several reasons have been offered to support this assertion. First, the aging of the American population will contribute to increased seafood consumption given health and nutrition concerns. Thus, the trend away from red meat to seafood and poultry as sources of low fat animal protein is expected to continue. Second, the minority population in the U.S., particularly Asians and Hispanics, are the fastest growing groups of the U.S. population. These minority groups generally rely heavily on seafood products for their daily diet. Third, a working population depends more on fast food products. Seafood has been identified as a natural fast food and is served readily in fast food restaurants or for dinner at home. Finally, continued improvements in the distribution of food products in general should have beneficial effects in the provision of consistent supply and good quality seafood products across the nation. These factors all point to a continued growth in the demand for seafood products. In addition, the nation's growing trade deficit has already been identified as another major reason for the expansion of the aquaculture industry due to the current huge U.S. seafood imports. It has been estimated that the U.S. may depend on imports for as much as 80% of its seafood consumption in the year 2000 unless domestic aquaculture expands rapidly (Egan, 1990). What do all these trends mean for hybrid striped bass? One can surmise that it offers a good outlook for the growth of the hybrid striped bass industry. However, there are barriers that need to be surmounted by the hybrid striped bass industry. As shown in past marketing studies dealing with hybrid striped bass, there are at least three major problems that need immediate attention. The first problem deals with familiarity. Consumers are not familiar with the species. Hybrid striped bass producers have to implement effective marketing or advertising strategies in order to promote hybrid striped bass. One might argue that all fish are the same similar to the idea that chicken products are generally homogeneous. But it is also possible to follow what Perdue Farms has done in advertising their chicken products as better with their flesh, yellow appearance. A similar product differentiation strategy can be adopted by hybrid striped bass producers in order to position the species as "different" from other seafood products.

326

The second problem identified in previous marketing studies is consistent supply and quality. This is a very critical element in any successful marketing strategy. Unless producers are able to supply good quality hybrid striped bass on a consistent basis, it would be difficult for restaurants, wholesalers, and retailers to promote the product vigorously. Finally, a third problem in the marketing area for hybrid striped bass is high price. Due to rapid changes in required production technology and continued innovation in the development of an efficient and environmentally-sound system of production, hybrid striped bass production costs are relatively higher compared to other traditional seafood, poultry, and red meat products. The American consumer is generally price sensitive particularly for products that are not traditionally served on a frequent basis. With a high price, consumers are not given the chance to "experience" the product so that positive "perceptions" on hybrid striped bass can be developed. It would seem that these problems are difficult to solve. However, statistics 'show that hybrid striped bass production has risen rapidly in the Northeast region (Bush and Anderson, 1993). This increasing production trend will allow consumers to "experience" hybrid striped bass and perhaps result in the willingness of consumers to try it again. This was shown in the Fuller et al. (1990) study wherein 86% of the consumers served hybrid striped bass were willing to try the fish again. With a high consumption repeat rate for first time consumers, the hybrid striped bass industry may have the potential to succeed in the future.

327

References

Adriance, J.G., 1982. A market model for the North Atlantic commercial striped bass industry. M.S. Thesis. Department of Agricultural and Resource Economics, University of Maryland, College Park. Bacon, J., Lussier, W., Dowse, B. and Gempesaw, C., 1993. The potential for aquaculture in Delaware and Maryland: A survey of current and potential aquaculture producers. Agricultural Experiment Station Bulletin no. 500. Department of Food and Resource Economics, University of Delaware, Newark. Bettencourt, S.U. and Anderson, J.L., 1990. Pen-reared salmonid industry in the northeastern United States. NRAC Publication No. 100. Cooperative Extension Service, University of Rhode Island, Kingston. Bjomdal, T., 1990. The economics of salmon aquaculture. Blackwell Scientific Publishers, Brookline Village, MA. Brown, J.W., Easley, Jr., J.E. and Hodson, R.G., 1988. Investment and production costs for the hybrid striped bass x white bass in North Carolina. University of North Carolina Sea Grant Publication UNC-SG-WP-88-2, Raleigh. Bush, M.J. and Anderson, J.L., 1993. Northeast region aquaculture industry situation and outlook report. Rhode Island Agricultural Experiment Station Publication No. 2917. Department of Resource Economics, University of Rhode Island, Kingston. Carlberg, J.M. and MassingilI, M.J., 1995. Foodfish production II: commercial flow-though tank grow-out systems. Aquaculture 95, Annual Meeting of The World Aquaculture Society. San Diego, CA p 79. Abstract. Dowdell, S., 1990. Fish, seafood shortages projected. Supermarket news, Fairchild Publications, New York, 24: 46. D'Souza, G., Vanderpool, A., McCauley, A., Gempesaw, C. and Bacon, J., 1993. The marketing potential of aquaculture products in West Virginia: A survey of consumer, restaurateur, and wholesaler attitudes. R.M. Publication no. 93-01, Division of Resource Management, College of Agriculture and Forestry, West Virginia University, Martinsburg. Egan, J., 1990. The fish story of the decade. U.S. News and World Report. Washin~on, DC, November 26, pp. 5256. Fuller, M., Keenum, M. and Kelly, R., 1990. Consumer acceptance of hybrid striped bass in a Mississippi Gulf coast restaurant. Agricultural Economics Research Report no. 187, Department of Agricultural Economics, Mississippi State University, Mississippi State. Gempesaw, II, C.M., Bacon, J.R., Wessels, C.R. and Manalo, A., 1995. Consumer perceptive of aquaculture products. American Journal of Agricultural Economics, 77:1306-1312. Gempesaw, II, C.M., Lipton, D., Varma, V., and Bacon, J.R., 1992. A comparative analysis of hybrid striped bass production in ponds and tanks. Proceedings of the National Extension Aquaculture Workshop. Ferndale, AR, p. 31-42. Gempesaw, II, C.M., Wirth, F.F., Bacon, J.R. and Munasinghe, L., 1993. Economics of vertical integration in hybrid striped bass aquaculture. Pages 91-105 in U. Hatch and H. Kinnucan, editors. Aquaculture: models and economics. Westview Press, Boulder, CO.

328

Harvey, P., Kirkley, J., Richardson, L. and Sanko, J., 1990. Hybrid striped bass aquaculture survey and market potential. Virginia Department of Agriculture and Consumer Services, Division of Marketing, Richmond, VA. National Marine Fisheries Service, 1990. Historical catch statistics Atlantic and Gulf coast states 1879-1989. Current Fisheries Statistics No. 9010, Historical Series Nos. 5-9 revised, National Marine Fisheries Service, Washinpon, DC. National Marine Fisheries Service, 1992. Historical catch statistics Atlantic and Gulf coast states 1950-1991. Current Fisheries Statistics No. 9210, Historical Series No. 10 revised, National Marine Fisheries Service, Washinpon, DC. National Marine Fisheries Service, 1995. Fisheries of the United States, 1994. Current Fisheries Statistics No. 9400, National Marine Fisheries Service, Washin~on, DC.. Norton, V., Smith, T. and Strand, I., 1983. Stripers: The economic value of the Atlantic coast commercial and recreational striped bass fisheries. University of Maryland Sea Grant College UM-SG-TS-83-12, College Park. Nauman, F., Gempesaw, C., Bacon, J., Wessels, C., Manalo, A. and Lussier, W., 1995. Consumer choice for fresh fish: factors affecting purchase decisions. Aquaculture 95, Annual Meeting of The World Aquaculture Society. San Diego, CA p 166. Abstract. Nauman, F., 1995. Modeling demand for seafood using the evoked set framework. M.S. thesis, Department of Food and Resource Economics, University of Delaware, Newark. Rhodes, R.J. and Sheehan, B., 1991. Estimated annual production of commercial hybrid striped bass growers in the United States. Striped Bass Growers' Association Report, Raleigh, NC. Rhodes, R., 1993. South Carolina triples aquaculture production value. Water Farming Journal, 8(15): .3. Rogness, R. and Weddig, L., 1991. A word to the wise fisherman: imports to play a bigger role for U.S. consumer. National Fisherman Annual. 71 (4): 74-96. Strand, I. and Lipton, D., 1989. Aquaculture: An alternative for Maryland farmers. Pages 37-45 in Proceedings of the governor's conference on the future of Maryland agriculture., University of Maryland, College Park. Swartz, D., 1984. Marketing striped bass. Pages 233-254, in J.P. McCraren, editor. The aquaculture of striped bass. University of Maryland Sea Grant College UM-SG-MAP-84-01, College Park. USDA. (U.S. Department of Agriculture)., 1994. Aquaculture situation and outlook. Commodities Economics Division, Economic Research Service, AQS-12. Van Olst, J. and Carlberg, J.M., 1990. Commercial culture of hybrid striped bass: Status and potential. Aquaculture Magazine, 16(1): 49-59. Wirth, F., Halbrendt, C. and Vaughn, G., 1990. Conjoint analysis of the mid-Atlantic food fish market for farmraised hybrid striped bass. Agriculture Experiment Station Bulletin no. 488, Department of Food and Resource Economics, University of Delaware, Newark. Yamashita, V.F., 1981. Marketing striped bass in the Chesapeake Bay states of Maryland and Virginia. M.S. Thesis. Department of Agricultural and Resource Economics, University of Maryland, College Park.

Striped Bass and Other Morone Culture R.M. Harrell (Editor) 1997 Elsevier Science B.V.

329

Chapter 13 Processing and F o o d Safety Dafne D. Rawles, Custy F. Femandes, George J. Flick, and Gale R. Ammerman 13.1 PRODUCTION STATISTICS Interest in striped bass and hybrid striped bass aquaculture continues to grow in the United States. Before the mid to late 1980s, commercial attempts to raise striped bass for food were experimental or failed. After 15 years of rigorous research and expenditures exceeding $3.0 million U.S., Aquatic Systems Inc. (now Kent Seafarms, San Diego, CA) was the first company in the world to profitably produce hybrid striped bass for food (Van Olst and Carlberg, 1990). Of the hybrid striped bass crosses developed, the palmetto bass (Morone saxatilis x M. chrysops) and the sunshine bass (3//, chrysops x M. saxatilis) are most frequently cultured (see Chapter 1). The sunshine bass is considered a fancy food fish because of its fine flavor, however, limited production has made the fish relatively expensive. The growing demand for premium quality finfish has strengthened interest in hybrid striped bass production, therefore it is considered a commercial product of increasing importance. Producers culture striped bass and its hybrids in tanks (e.g., flow-through circular or rectangular, recirculating systems), net-pens, and ponds throughout the U.S. producers can be broadly classified into three categories: (a) large scale producers harvesting 1,000,000 pounds or more; (b) medium scale producers harvesting between 100,000 and 1,000,000 pounds; and 9 small scale producers harvesting less than 100,000 pounds. Most large scale culturists produce in tanks as well as ponds. The small scale aquaculturist produces mostly in ponds. The majority of hybrid striped bass are sold to major fish distributors located along the east and west coasts as well as in the midwest. A marketable size hybrid striped bass is produced in 8-13 months in tanks, and 18-24 months in ponds. Unlike the catfish industry, where the product is sold by unit weight, hybrid striped bass are frequently sold by weight range. There are eight grades of fish sold in quarter pound increments from one to three pounds (e.g., 1 to 1.25; 1.26 to 1.50; 1.51 to 1.75; 1.76 to 2.00; 2.01 to 2.25; 2.26 to 2.50; 2.51 to 2.75; and 2.76 to 3.00 lb). Generally, producers sell the lower weight range as live or iced whole fish, while the higher weight range is preferred for processed products such as fillets. About 75-80% ofthe product is sold as iced whole fish, 10-15% is sold live, and 5% is processed into fillets. Producers prefer to sell their fish quickly to maintain a high pond turnover rate. They are uninterested in waiting an additional period (e.g., three months) to reach the highest weight range category since the cost of production increases due to reduced feed conversion (Coale et al., 1993). 13.2 LAWS AND REGULATIONS Several agencies monitor the fish industry in the United States Regulations under the Food, Drug and Cosmetic Act of 1938 are enforced by the Secretary of Health and Human Services and updated through publications in the Code of Federal Regulations (CFR). The U.S. Department of Commerce National Marine Fisheries Service (NMFS) administers programs related to grading and voluntary inspections, and establishes market standards for fishery products. Regulations mandating the production of safe, wholesome fishery products are published by the Food and Drug Administration (FDA) in Title 21 CFR and by the NMFS in Title 50, both of which are revised annually. Many standards used internationally, officially or voluntarily, have

330

been developed by the Food and Agricultural Organization (FAO) and the World Health Organization (WHO) of the United Nations (Regenstein and Regenstein, 1991). 13.2.1 Processing Facilities Regulations concerning the safe and wholesome processing of fish and fishery products are contained in the Federal Food, Drug and Cosmetic Act, as Amended (U.S. Department of Health and Human Services, 1993) and Title 21 CFR, Part 110, (U.S. Department of Health and Human Services, 1995). The regulations address personnel, plant and grounds, sanitary operations, sanitary facilities and controls, equipment and utensils, processes and controls, and warehousing and distribution. 13.2.2 HACCP The seafood industry and government regulatory agencies (FDA and NMFS) are placing greater reliance on the Hazard Analysis Critical Control Point (HACCP) system to provide greater assurance of food safety and to maintain wholesomeness and prevent economic fraud such as species misidentification or short weight packages. The FDA published a mandatory food safety regulation based on the HACCP concept in the December 18, 1995 issue of the Federal Register amending Title 21 CFR, Chapter I to add PART 123 Fish and Fishery Products. The seafood industry was given two years to implement the new regulation. Additionally, the European Union (E.U.) passed a requirement that all fish and shellfish products entering member countries after January 1, 1996 be produced under a HACCP program. Although HACCP involves substantial self-monitoring by the industry, federal (FDA and NMFS) programs will ultimately rely on regular monitoring inspections, with periodic verification inspections. Using inspections, agencies will be able to review the plant's self-monitoring data to determine whether each HACCP-based system is in compliance with the plant's approved HACCP plan; they will check overall sanitation and compliance with good manufacturing practices, labeling, and other legal requirements. The HACCP approach is a preventive system where safety (and occasionally wholesomeness and economic fraud) programs are designed by which fish are processed. This is more effective than conventional quality assurance programs, which tend to rely on finished-product testing to provide evidence of compliance. Basically, HACCP is a manufacturing technique in which product quality is designed into the process. Although HACCP is being broadly applied in fish processing to provide greater assurance of food safety, it must be applied in all segments of the industry including growing, ingredient procurement, distribution, retailing, and the home. The HACCP principles should be used as a guideline for processors. Self-inspection, particularly using the HACCP approach, provides processors with the same comprehensive understanding of their operations that is needed to conclude that they are operating within legal requirements. To overlook the opportunity for an effective self-inspection program reduces the ability of processors to control their operations. Lack of a self-inspection technique places them in a position of assuming safety, wholesomeness, and quality, without actually knowing whether they exist. An HACCP-based self-inspection provides this assurance. 13.3 PROCESSING 13.3.1 Harvesting Methods and Transport Different harvesting procedures may have a significant effect on the final product quality and shelf life (Mitsuda et al., 1980; Botta et al., 1987). To assure optimum quality, freshly caught fish must be handled

331

and processed carefully because its flesh is both delicate and highly perishable. In aquaculture production of hybrid striped bass, fish are seined from ponds and frequently transported to a live car. By using a hydraulically operated harvesting basket, the fish are transported into a container filled with water. Fish are then sold live or weighed and packed on ice until delivery to the buyer (Hodson et al., 1987). Because stress and high metabolic activity associated with harvesting and transportation of live fish may affect the final product quality and shelf life, this aspect must be carefully handled. A live-haul truck usually has 2500 to 4500 kg capacity. If there is an abundant supply offish in ponds not over 75 km from the processing plant, keeping the plant supplied is not a serious problem except in cases of extreme weather. In Mississippi, the catfish industry is highly centralized, and virtually all fish are transported alive to the processing plants. Live-holding at the plant until the fish are processed has advantages in that self-grading in the holding tanks is possible. There is a continuous risk of losses in transit, particularly during warm weather. Hauling fish on ice is an alternative to live-hauling, particularly if the fish must be transported long distances. Larger pay loads can be hauled using ice, and during bad weather the fish can be hauled from the ponds in lighter trucks thus keeping heavier loads off the pond levees or dams. If fish are held for several days before processing, the meat may discolor and a loss in flavor could result, however, acceptable processed fish can be produced. Crushed ice should be used to minimize bruising, and alternate layers of ice and fish should be added to insure adequate contact of fish with the ice. The amount of ice required is a function of the initial temperature of the fish, adequacy of the insulation of the carrier walls, and the amount of time the fish are to be iced. With good contact of fish and ice, 0.33 kg of ice will reduce the temperature of 1 kg offish from 27~ to 2~ upon complete melting of the ice over a four to six hour period of time. If the fish are to be held iced 12 h and are initially near 27~ a ratio of one kg of ice per kg of fish has been found to be satisfactory. If the fish are at 10 ~ or colder, a ratio of 0.5 kg of ice per kg of fish will be adequate for holding the fish 12 h (Ammerman, 1985). Freshly caught fish that are iced immediately and held in ice can maintain a high quality for eight to nine days and be edible for up to two weeks. Ideally the temperature should be kept close to 0~ throughout handling, therefore attention must be paid to obtain an intimate contact between fish and ice to assure proper cooling. 13.3.2 Receiving Fish at the Plant A problem often encountered by producers is off-flavor. Off-flavor, described as musty or earthy, is mainly due to algal growth, however, it could be bacterial in nature, Generally, off-flavor is more pronounced in ponds rather than tanks. Off-flavor problems can usually be rectified by purging live fish in holding tanks or ponds for three to seven days after harvest. Reducing off-flavor by chemically treating hybrid striped bass ponds with copper sulfate to inhibit algal growth has proven successful. Also, high pond water turnover rate can reduce off-flavor. Although some producers claim that sodium bicarbonate (baking soda) can alleviate off-flavor problems, its success has not been scientifically proven. Several days before delivery to the processing plant, samples of one to three fish from the pond to be harvested are obtained for flavor evaluation. The fish are dressed and cooked to a cold-spot temperature of 70~ in a covered dish in a microwave oven. If the fish are judged to have acceptable flavor, the pond can be approved for the purchase. On the day the fish are delivered to the plant, they are cooked and tasted again before they are unloaded and received by the processor. Ifjudged to be "on-flavor" the fish are unloaded into the processing plant holding tanks and held alive until processed.

332

13.3.3 Initial Handling Bleeding fish is accomplished by cutting the arteries behind the gills and in front of the heart. After allowing it to bleed either in air or water for approximately 20 min, the fish receive a final wash and are then eviscerated (gutted). This operation is generally done manually and consists of cutting from either the fish's gill cleft to the vent or from the vent to the gills. Once the fish is opened, the viscera are removed. Fish gills may also be removed, although there is controversy about this practice since their removal may result in difficulties in determining quality at retail level (Regenstein and Regenstein, 1991). For fish that will not be skinned, scaling is required before filleting. This operation is most commonly achieved by using a hand-held or mechanical scaling machine and large amounts of water spray. Mechanical damage of the tissue may result from these methods. Hand scaling results in less damage, however it increases processing cost. Consequently, it is effective when small quantities of fish are processed. 13.3.4 Filleting Filleting is done manually and mechanically. When done manually, the fish are brought to the filleters and the finished fillets are moved along manually or by conveyors. Generally, two fillets are obtained from each fish. During filleting, the head of the fish remains attached to the body. Cutting the fish by machine can be faster and more profitable than filleting by hand, however, it requires more capital investment and maintenance (Regenstein and Regenstein, 1991). Mechanical filleting usually delivers lower yields than the hand operation, however, it is still the operation of choice for filleting smaller fish. Some filleting machines require the fish to be decapitated before operation. 13.3.5 Meat-bone Separation During filleting of hybrid striped bass, only 29-50% ofthe meat is obtained from the whole fish. Meat remaining on the fish frames may be recovered by a meat bone separator as mince. The meat bone separator is used in many seafood processing industries to recover the minced meat adhering to the frames. A moving belt presses the fish frames firmly against the surface of a perforated hollow cylinder allowing valuable fish flesh to pass into the drum. Utilization of spent-frames and salvage of portions of the belly flaps for use in further processed products are areas of great interest to processors since the meat can be used to produce fish portions and sticks. The ability to produce a product through by-product recovery increases profitability from both new product sales and the reduction in cost for disposal of a solid waste. The oxidative stabilities of farm raised striped bass and hybrid striped bass minced muscle tissue were evaluated during a nine month fluctuating frozen (- 18~to -6 ~ C) storage period (Erickson, 1993). Hybrid striped bass (Table 13.1) were less stable in the early phases of storage because of lower concentrations of naturally occurring antioxidants (tocopherol, ascorbic acid, manganese, zinc). Also, the higher concentration of iron and the chemical nature of the lipid increased susceptibility to oxidation. At the end of the storage study, both bass samples exhibited significant oxidative deterioration. 13.4.6 Dressing Yields Aquacultured hybrid striped bass are mainly marketed as iced whole fish, but may also be sold as dressed or further processed. Generally, processing of hybrid striped bass augments product value by changing its market form, and raises revenue for the processor.

333

Table 13.1. Initial lipid, metal composition, and oxidative measurements of frozen, minced bass tissue ~ (from Karahadian et al., 1995). Hybrid Striped Bass

Striped Bass Storage ( m o n t h s )

Oxidative Measurement

0

3

5

9

Peroxide.s (nmol/g dry welglat)

31.61 a

0.0 a

31.9 a

631.3 c

Conjugated dienes (A z32 )

1.78 ab

1.7 ab

TBA.R.S .(tamol/g dry welglat) -

0.4 a

5.2 b

53.1 d

46.0 c

337.9 ~

nd b

a-Tocopherol equivalents ( n m o u g dry weight) Ascorbic acid ( n m o l / g dry welglat)

0

3

6

9

2.9 a

0.0 a

117.6 a

656.1 r

1.65 a

2.26 c

1.68 a

1.85 b

1.83 b

2.45 d

33.0 ~

95.3 g

0.0 a

8.8 r

! 5.0 d

80.4 f

45.3 r

13.1 a

84.3 c

85.3 e

89.6 ~

37.7 b

0.0 a

0.0 ~

413.3 d

48.0 b

56.3 b

0.0 ~

Phospholipids PUF.A. ( m g / g dry welglat)

7.3 a

8.3 b

22:6n.-.3 (mg/g dry weight)

1.3 a

2.0 b

Peroxidizability index 3

35.8 a

32.1 b

C o p p e r ( ~ g / g dry weigtiO

0.8 a

1.0 a

Iron ( u g / g dry welglat)

8.0 a

15.2 b

M a n g a n e s e (~tg/g dry welglat)

0.4 a

0.5 b

15.1 a

17.3 b

Z i n c . ( ~ g / g dry weight)

i Values in a row followed by the same letter are not significantly different (P>0.05). 2Thiobarbituric acid - Reactive substances. 3Peroxidizability index = {(sum mg monoenes X 0.025) + (sum mg dienes X 1) + (sum mg trienes X 2) + (sum mg tetraenes X 4) + (sum mg pentaenes X 6) + (sum mg hexaenes X 8)} / g dry weight (Murata & Yamaguchi, 1990).

334

Table 13.2. Processing yields for hybrid striped bass (from Coale et al., 1993). Product Form

Whole fish

Percent Yield

100.0

Dressed, with gills

90.5

Dressed, without gills

85.4

Fillet, with rib bones

45.4

Fillet, with skin

41.9

Fillet, without rib bone, skinless

32.0

Fillet, without rib bone, skinless, trimmed

29.5

Solid waste: Frame

48.8

Skin

19.8

Viscera

9.5

Sunshine bass were processed into whole-gutted fish and fillets (Coale et al., 1993), with the weight of the fish ranging from 225g to 500g. The processing yields of some products are shown in Table 13.2. 13.4 PRESERVATION METHODS A wide variety of preservation techniques can be used for fisheries products, however, most striped bass and hybrids receive little processing before consumption, being sold whole, gutted, or dressed and on ice to ensure good quality and extended shelf life. Different degrees of processing can sometimes result in substantial weight losses. Scaling can result in approximately 3% weight loss, and gutting and heading represent another 15% weight loss (Smith et al., 1985). Most central wholesale market centers deal in fresh fish and purveyors prefer to receive fish in ice, packed in-the-round, or gutted (Carlberg and Van Olst, 1987). Yield of cultured bass, ranging in size from one to two pounds round-weight, processed as boneless fillets with skin on is approximately 42%. 13.4.1 Ice-packing and Refrigerating Deheaded, and eviscerated fish may be sold fresh; iced without packaging, iced in packages, or packaged and refrigerated. Most striped bass are sold fresh in ice. After size grading, fish are either packed in trays or remain unpackaged. The fish are then boxed, with ice, in cardboard shipping cases and transported. Approximately 12 days after slaughter, the normal shelf life of iced fish will be reached whether it is packed in trays or not.

335

13.4.2 Chill-packing This method involves bringing the temperature down, from ambient to -1 ~ and developing crustfreezing. The product is then traded in the crust-frozen state, but it is offered for sale at retail level as fresh fish. Chill-packed fish may retain good flavor and odor for twice as long as ice-packed fish. Chill-packed (-2~ commercially processed hybrid striped bass were compared to refrigerated samples (2 ~ for changes in microbiological, chemical, and sensory evaluations over a 21-day storage period (Boyd et al., 1992). Aerobic plate counts and hypoxanthine (a degradation product of ATP) formation showed evidence of rapid deterioration of refrigerated samples after eight days storage compared to 21 days for chillpack samples. A trained sensory panel found 21 day stored chill-pack samples similar in aroma and flavor but of firmer texture than refrigerated samples stored eight days. The presence of the belly flap did not affect sensory characteristics during the storage study. Hypoxanthine formation and aerobic plate counts appeared to be good indicators of quality deterioration, whereas thiobarbituric acid measurements were not. 13.4.3 Freezing Food chains prefer frozen products for self-service seafood sales (Swartz, 1984). Although microbial changes are inhibited in frozen muscle, physical and chemical changes can still occur. Regenstein et al. (1982) reported that freezing and frozen storage of fish can cause textural changes with decreased water retention of muscle proteins. Freezing consists of rapidly lowering the internal temperature to-18~ or below. Fish freezing is a gradual process that takes place between -1 ~ and -5~ Gutted or whole fish are unpackaged and once freezing is complete, the fish are removed from the freezer and placed in a primary container. The fish are then returned to the freezer until they are placed in a master carton immediately before shipping. Ideally, the frozen storage temperature should be -30~ or lower. To obtain a uniformly frozen product, it is recommended that fish be frozen until the surface temperature is down to -35 ~or-40 ~ so that they will equilibrate without the need for additional heat removal during storage. Gutted fish that has been quick frozen and then thawed after proper cold storage can be handled in the same way as top quality fresh fish. They can be filleted and distributed for retail sale, or brined and smoked, or filleted and used for bulk or tray packs. When fillets are frozen, a less ragged product is obtained compared with fillets obtained from frozen and then thawed whole fish. However, they are usually duller and may not produce a smoke cure of the same high quality (Graham, 1982). 13.4.4 Smoking Smoking is mainly done to meet consumers' taste demands rather than as a preservation method. Smoked fish are obtained through a series of operations that can last from a few hours to few days. There are basically four steps in the process, beginning with brining and ending with heating. In brining, fish (gutted, headed and gutted, or skin-on fillets) are exposed to a 12-20% salt solution (brine). The brine may contain other ingredients such as sugar, lemon juice, bay leaf, onion powder, and garlic powder. Brining before smoking is done to: (1) give a firmer consistency to the product; (2) stop oil from leaking out; (3) impart flavor; and (4) prepare the flesh to accept smoke. The increased salt level helps to inhibit pathogenic bacteria, however, it may act as a pro-oxidant. The product is then hung for smoking or placed on smoking racks. The liquid phase salt concentration in the loin muscle should be 2.5% if an aerobic package is used, and 3.5% for an anaerobic package.

336

The fish must be dried to achieve the characteristic smoked flavor, and a firm consistency, and to aid in the formation of the glossy surface pellicle by sealing the surface. The fish can be dried through placement in refrigeration or in the smoke house. Drying in the smoke house will depend on humidity, air movement, and temperature. Smoke may be generated by making small fires in a smoke house or by a smoke generator. With the latter treatment, smoke at temperatures between 250 ~ to 350~ is generated, avoiding temperatures above 400~ which may cause formation of undesirable potential carcinogenic compounds. The smoke is composed of two fractions: a non-volatile fraction that contains polycyclic aromatic hydrocarbons identified as potential carcinogens; and the volatile fraction containing the flavor compounds, phenols, and carbonyls. Liquid smoke, increasingly in use, duplicates the latter fraction with the advantage of the absence of the carcinogenic component. In any system, the fish is always stationary and the smoke is passed through it. In the middle of the smoking protocol, the fish may be rotated to achieve an even smoking. The internal temperature at the cold point (the center of the thickest part of the fish) of the bass should be a minimum of 63~ and maintained for 30 rain. 13.5 PACKAGING Packaging technology has advanced considerably but is slow in implementation in the fish industry. Most hybrid striped bass producers and processors are small and cannot afford expensive packaging machinery, besides the small quantity processed is generally economical to package manually. An attractive package draws the consumer to a product, however, an attractive packaging cannot rectify the quality of the hybrid striped bass product if it is less than optimal, thus, the product must be of high quality before packaging. Packaging does play an important role in preserving product quality, processors should develop packages with good sales appeal, and the design must be convenient for end use (e.g., retail, restaurant). Processors should design packages appropriate for selling a pricey product such as the hybrid striped bass. Because the product is an extremely perishable food, deterioration in quality is directly proportional to product holding temperature. Packaging is influenced by preservation technique (e.g., refrigerating (2~ chillpacking (-2 ~ ice packing, or freezing), and lowering temperature extends the shelf life of the product. Boyd et al. (1992) studied the quality changes of aquacultured hybrid striped bass during packaging in oxygen permeable bags (Cryovac E ) and storage as refrigerated and chillpacked samples. Aerobic plate counts and hypoxanthine formation showed evidence of rapid deterioration of refrigerated samples after eight days, compared to 21 days for chillpacked samples. A trained panel found 21-day stored chillpack samples were similar in flavor and aroma but of firmer texture than refrigerated samples stored for eight days. Factors affecting the quality of refrigerated or chill packed or ice packed fish include biological (bacterial deterioration), biochemical (autolytic enzymes), and chemical (lipid rancidity). For refrigerated fish products, factors affecting quality are controlled by lowering temperature. Generally, low temperature retards bacterial deterioration, enzymatic breakdown, and lipid rancidity. Oxygen is required for microbial survival and growth as well as lipid rancidity. In modified (MAP) or controlled atmospheric packaging (CAP), the oxygen is replaced with carbon dioxide. Although the growth of spoilage organisms is retarded, product quality may be deteriorated by anaerobic organisms, some of which may produce toxins. In refrigeration, an excessive moisture loss (weight loss) impairs the sensory properties (e.g., texture, flavor, and color). Some liquid is lost and trapped in the packed product and fish odor permeates through the package, resulting in loss of volatiles.

337

Factors affecting the quality of frozen fish products include physical (moisture loss), chemical (lipid rancidity), sensory (odor, flavor, appearance and texture), biochemical (enzymes), and nutritional (vitamins). For a frozen product, the package must provide an adequate barrier to moisture migration. Glazing fish is an excellent protective package, because the protective coating of ice prevents oxygen and moisture migration and freezer bum. Skintight vacuum packaging films can also prevent moisture loss. It is important to avert voids during packaging as they are responsible for moisture migration and accumulation of unsightly "snow." Generally, most fish are sold directly to distributors who sell to the consumer through retail supermarkets or restaurants, therefore, packaging sells the product to consumers. For retail trade, the secondary (shipping container) package must offer convenience to the retailer in storing and handling, whereas the primary (store package) packaging must be weighed, priced, and use-by-date stamped. Some retail chains require the use of UPC codes on prepackaged foods. Additionally, nutritional information may be provided, as well as preparing and holding instructions. For restaurant or institutional trade, bulk packaging must meet their storing and handling needs. Proper packaging will minimize or eliminate product handling. Some consumers are accustomed to evaluating fish quality by sight and smell, therefore fillets should be placed with skin and bone sides up so the consumer can rapidly identify the species and perform a sensory inspection. Permeable packaging may have a beneficial role in this type of sensory evaluation. For chilled products, visibility is important. Clear films with no fogging and dry, fresh products are always appealing. For frozen products, skintight transparent films that do not accumulate frost are best. Most hybrid striped bass are dressed and not thermally processed, except for smoked fish, therefore, temperature control is critical as a packaged product. Strict temperature control (maintaining the product at 2~C or less) is necessary, however, when the barrier film is impermeable to oxygen. Failure to maintain proper temperature control could result in the growth and toxin production ofClostridium botulinum. For refrigerated retail trade, an absorbent pad lined with plastic is inserted between the product and the foam plastic tray (e.g., polystyrene) and overwrapped with transparent plastic film (e.g., polyvinyl chloride). Product weepage is best controlled by constantly maintaining the product below or near (0~ Moisture condenses as droplets and obscures the view of product. Moisture droplets can be dispersed by addition of antifogging (or wetting) agents (e.g., nonionic ethoxylates or hydrophobic fatty acid esters of glyceryl stearate) (Bakker, 1986). For frozen products, the most popular packaging is wax or plastic coated paperboard. The package must provide a good oxygen and moisture barrier as high oxygen permeability results in rapid rancidity development. An end use problem with frozen products is drip, a liquid released from thawing product, which represents a loss of product components. Drip may be caused by several factors including thawing after slow freezing, protein destruction, and storage temperature fluctuations. 13.5.1 Bulk Packaging Chilled whole products are transported on ice in wax or plastic coated 3-ply boxes in 22.7 kg unit size. Chilled fillets are packaged in wax or plastic coated 3-ply boxes in two unit sizes, 4.54 kg and 7.81 kg. For convenience, fillets should be layer packed with plastic film sheet sandwiched between layers to facilitate fillet removal. The product is placed in a snap lid plastic tray which is immersed in a refrigerant. Generally, ice is the most economical refrigerant for land transportation. Melting ice maintains product temperature near 9~ Reusable refrigerants such as blue ice packs have also been employed, but they must be recyclable to be economical. Reusable refrigerants are almost always used during air transportation. For air packages, an additional polyethylene bag box liner is used for liquid containment.

338

13.5.2 Labeling Hybrid striped bass processors are required to follow the U.S. Fair Packaging and Labeling Act of 1966 and its subsequent amendments (Title 15 CFR, Chapter 39). This act requires very specific information to be placed on the packaging label such as the processor's, packer's, and distributor's business name and address. The primary packaging must include information on weight (e.g., pounds), ingredients (e.g., breaded products, fillets), nutritional information, and safe handling instructions. Nutritional information is mandatory for formulated product and the label must contain information such as serving size, which must be written in tangible terms (e.g., weight), and should include calories per serving (total and lipid), protein, and fat (saturated, unsaturated, cholesterol). Minerals and vitamins (A, B series, C, D, and E), and contents per serving should be reported as percent daily values based on a 2000 calorie diet. Food constituents that are less than 2% of daily values do not have to be listed. The safe handling instructions should indicate that the product is perishable and needs appropriate refrigerating or freezing before preparation; good sanitary practices such as cleaning hands and utensils before and after handling raw meat to avoid cross-contamination; and information on thorough cooking before consuming and refrigerating leftovers. 13.5.3 Coding Coding is not required by law but is important as it facilitates identification. Identification enables the buyer, processor, or consumer to separate or recall a specific batch in the event of spoilage, mislabeling, or food poisoning. Essential elements for batch coding are the species type, product style, processor's or distributor's name, and processing date. For production scheduled during an entire working day, the processing time (e.g., AM or PM) should be indicated. As processors expand their business and operate multiple-lines and multiple-processing plants, batch coding information could include line and plant numbers. Batch codes could also include information on the producer. This information is essential for quality control, as some producers may supply fish containing a chemical contaminant. Below is an illustration of a one line batch identification code developed using alphabetical and numerical codes. B-14 05-03-12156 B - the species code for 26 species (e.g., sunshine bass, palmetto bass, etc.) 14 - the product style code for 100 styles (e.g., fresh whole, fillets) 05 - the production line # (for 5 plants) 03 - the production plant # (for 5 plants) The number 12156, is the production date (e.g., Dec. 15, 1996) which is a Gregorian date code, or alternatively the Julian date code could be used which is 3486. The 348 represents the day of the year and 6 represents the year 1996. The batch coding system should be flexible so changes can be made according to needs. 13.6 FACTORS AFFECTING QUALITY AND SHELF LIFE Spoilage of harvested fish may result from a variety of causes including microbiological, causing deterioration of organoleptic properties; chemical, resulting in rancidity; and physical, including freezer burn or dehydration (Curiale, 1991). Spoilage in fish occurs quicker than that of mammalian muscle due to the higher water and free amino acid content, less connective tissue, higher enzyme activity, and higher ultimate pH of fish flesh (Pedrosa-Menabrito and Regenstein, 1988). Of all the flesh foods, fish are the most susceptible to autolysis (post-harvest decomposition), oxidation and hydrolysis of fats, and microbial spoilage (Frazier and Westhoff, 1978).

339

Stress and pre-harvest condition of fish may have a dramatic impact on final product quality. To optimize product quality, post-mortem changes in fish flesh, including bacterial and enzymatic degradation, need to be controlled. The physiological condition of fish before harvest also has an important effect on the ultimate quality. Love (1988) demonstrated that starving fish can enter rigor mortis almost immediately after death, and that spawning fish are in their worst condition with very soft, watery flesh. Enzymatic activity is accelerated in the gut of a fish subjected to struggle, with an even greater effect when the fish has been feeding before capture. Overcrowding of fish in ponds and tanks can result in increased spoilage rate because scale losses favor penetration of bacteria into the skin and flesh. Post-harvest metabolism may be changed because, as a result of energy expended by swimming and struggling during capture, there is a reduction of muscle glycogen in fish which results in accumulation of lactate and pyruvate (Black et al., 1962). This can lead to a rapid onset of rigor mortis with a significant effect in quality deterioration (Pedrosa-Menabrito and Regenstein, 1988). By using procedures to lengthen rigor mortis, shelf life can be extended and fish preservation improved by delaying post-mortem autolysis and bacterial decomposition (Crawford et al., 1970; Korhonen et al., 1990). Post-harvest metabolism can be reduced by placing the fish into cooled water or water with an elevated level of carbon dioxide and by decreasing the pH (Frazier and Westhoff, 1978; Eifert, 1991; Mayer and Ward, 1991). Variations between texture measurements in fish fillets from a singular species have been reported to be mainly due to geographical, seasonal, and feeding factors of live fish, post-mortem biochemical changes, and the filleting process (Johnson et al., 1980). Post-mortem microbial changes are significant because the muscle of fresh fish provides an excellent substrate for the growth of spoilage microorganisms due to high water activity, near neutral pH, and a high level of soluble nutrients (Liston, 1980). During cold storage of fish, bacterial numbers increase reaching levels above 107 colony forming units/g (cfu/g). Predominant spoilage bacteria in this stage are psychotrophic Gram negative organisms of the genus Pseudomonas and Alteromonas (Kraft and Rey, 1979; Venugopal, 1990). These organisms can grow near 0~ and can produce hydrogen sulfide (HES),methyl mercaptan (CH3SH), and dimethyl sulfide ((CH3)2S) from the reaction with amino acids methionine and cysteine. With bacterial levels above 106 cfu/g, production of these volatile sulfur compounds is such that spoilage becomes organoleptically evident (Shewan, 1977; Liston, 1982). Metabolites ofpost-mortem bacterial action on amino acids can also yield objectionable flavors and odors from volatile bases such as diamines, cadaverine, histamine, and ammonia (Jacober and Rand, 1982). Bacterial action also leads to the development of acetic acid, CO2, and H20 from carbohydrates and lactate, and to conversion of glycine, leucine, and serine to esters of acetic, propionic, butyric, and hexanoic acids, and production of fruity odors (Shewan, 1977). Post-mortem chemical and physical changes are significant because lowering of the pH from near neutral to approximately 6.2 to 6.5, due to the anaerobic degradation of glycogen to pyruvic and lactic acid and enzymatic degradation of ATP to glutamic and formic acids are post-mortem changes that occur in fish flesh (Barrett et al., 1965; Bendall, 1972; Korhonen et al., 1990; Sikorski et al., 1990). Accumulation of lactic acid is the principal factor in determining post-mortem muscle acidity (Tart, 1966). If fish struggle or swim excessively before death, depletion of glycogen reserves will occur and post-mortem pH will be higher than in rested fish where lactic acid may accumulate through glycolysis. Flesh texture in fish is toughest during rigor and at low pH (Dunajski, 1979). 13.6.1 Microbiological Considerations Bacteriological contamination in aquacultured fish can result from contact with the environment, feed, birds, animals, and humans. In any production system (pond or tank) where there is a large population of fish, the intensive nature of cultural conditions may cause rapid spread of pathogens and/or high bacterial loads.

340

Little is known about the microflora of healthy, aquacultured hybrid striped bass, therefore the incidence of food borne illness-causing bacteria is not clear. Recently Nedoluha and Westhoff (1993) reported presence of Aeromonas spp. (27%), coryneforms (14%), Pseudomonas spp. (12%), Flavobacterium/Cytophaga/ Sphingobacterium group (8%), Plesiomonas shigelloides (7%), Bacillus spp. (7%), and Enterobacteriaceae (6%) among the main bacterial contaminants in pond farm-raised hybrid striped bass in the Maryland area. Listeria monocytogenes, Staphylococcus aureus, Shigella dysenteriae, Vibrio spp., and Yersinia pseudotuberculosis were among the human pathogens isolated. Fish in recirculating systems have been found to contain Vibrio spp. and Mycobacterium mar&um as well as other Mycobacterium species. The presence of Mycobacterium species presents a greater danger to employees of the producer rather than the consumer (Flick, unpublished data). However, no evidence of a higher risk of food borne illness from pond-raised fish as compared to the wild counterpart was indicated. Fish are an excellent medium for the growth of various microorganisms. As shown above, microorganisms encountered in hybrid striped bass could cause spoilage or be a food-borne pathogen. Food spoilage organisms would reduce the shelf-life of processed fish while food-borne pathogens could induce disease. Food-borne pathogens are of two types: food infections caused by the presence of the viable organism in food (e.g., Escherichia coli O157:H7, Salmonella), food intoxications caused by the presence of toxic microbial metabolite in food (e.g., botulinum from Clostridium botulinum; enterotoxin from Staphylococcus aureus). The most potentially dangerous microorganisms to striped bass processors are Staphylococcus aureus due to human contamination and Salmonella. Another pathogenic bacteria of emerging significance is Listeria monocytogenes, which can survive and grow on fish. It becomes a problem when the monocytogenes is transferred from fresh fish to a cooked ready-to-eat product (e.g., smoked fish). Optimum protection from these and other pathogenic bacteria will result from planning and executing a cleaning and sanitizing program. Fish should be moved through the processing lines rapidly and kept below 4~ as much as possible. Diseased fish should not be processed for human consumption. 13.6.2 Microbial Growth During Storage Storage temperature has a significant effect on surface microbial populations. The higher the microbial populations, the shorter the shelf-life. Hybrid bass stored at three temperatures (-1.67, 0, and 2.2 ~ had the surface counts found in Table 13.3. Each lot processed in a defined period must be examined for microbiological specifications to ensure process control (Liston and Matches, 1976). Microbiological specifications may include aerobic plate count, E. coli count, coagulase positive Staphyloccus aureus count (FDA, 1992). 13.6.3 Chemical Considerations Various agrochemicals including pesticides (White et al., 1985; Armbruster et al., 1987, 1988), drug residues (Plakas et al., 1991), and heavy metals (Rehwoldt et al., 1978; Armbruster et al., 1988) have been identified from wild striped bass fish and fillets. Some heavy metals are toxins, and limits are placed on their content in food. However, there is a paucity of information on the presence of agrochemicals in aquacultured striped bass and hybrid striped bass. Contaminants in fish can be reduced to acceptable levels or eliminated if producers implement a quality assurance program using guidelines described by the National Aquaculture Association.

341

Table 13.3. Surface microbial counts on hybrid striped bass stored at chilled and super-chilled temperatures (Flick, unpublished data). Post-Processing days

-1.67~

Surface Microbial Counts 0~

2.2~

3

102

103

102

8

102

103

103

10

102

10 4

106

13

103

105

107

15

103

107

*

17

104

*

* = fish considered unacceptable by sensory analysis

13.7 COMPOSITION 13.7.1 Amino Acids Composition The composition of free amino acids in the muscle tissue is found in Table 13.4. 13.7.2 Elemental Composition Composition of macro and micro nutrients in the muscle tissue of hybrid striped bass is found in Table 13.5. 13.7.3 Fatty Acid Analysis Fatty acid composition varies according to environmental conditions and other factors. The fatty acids analysis obtained from hybrid striped bass grown in four states in both pond and recirculating systems is presented in Table 13.6. 13.7.4 Effect of Diet on Composition The uptake of omega three fatty acids was reported to be dependent on the amount of unsaturated fatty acids fish fed diet supplemented by fish oil (Fair et al., 1993). Fatty acid profiles in muscle tissue reflected diet concentrations with increases in eicosapentaenoic (EPA) acid directly related to feed lipid composition. Other fatty acids demonstrating muscle concentration increases of 1 percent or greater were 14:0, 16: ln-7, and 22:6n-3, as well as total saturates, total n-3 fatty acids, and the n-3/n-6 ratio. Fatty acids showing decreases of 1% or greater were 16:0, total saturates and monounsaturates. A dietary source of n-3 fatty acids resulted in elevated levels of EPA and docosahexaenoic acid (DHA) in muscle tissue.

342

Table 13.4. Free amino acid content found in the muscle of hybrid striped bass (Flick, unpublished data). Amino Acid

Concentration Range in Nanomoles/gram

Aspartic acid

274-724

Threonine

794-1306

Serine

1781-1954

Isoleucine

211-690

Phosphoserine

NA

Leucine

414-1527

Glutamic Acid

1474-3140

Proline

604-846

Glycine

14171-24376

Alanine

3631-4251

Cystine

0

Valine

552-1326

Methionine

0

Tyrosine

0-842

Phenylalanine

237-700

Lysine

1944 -2595

Histidine

2696 - 4197

Ammonia

13580 - 15621

Arginine

0- 273

Taurine

10498 - 14133

It was reported by Fowler et al. (1994) that a descriptive sensory evaluation of cooked fish fed diets containing at least 0.50% EPA and 0.24% DHA showed a significant increase in fish flavor intensity compared to cooked fish fed diets containing trace amounts of these n-3 fatty acids. Although EPA and DHA do not exhibit fishy flavors and aromas, their oxidation products are known contributors to fishy aromas and flavors. 13.7.6 Proximate Analysis of Striped and Hybrid Bass Striped bass from the Chesapeake Bay and palmetto bass and sunshine bass were analyzed for their chemical composition. The proximate analysis (Table 13.7) and fatty acid compositions of edible tissue from wild catches were similar within a sampling season. Hybrid bass contained a greater variation than the wild

343

Table 13.5. Macro and micro nutrients found in the muscle of hybrid striped bass (Flick, unpublished data). Element

Concentration in ppm

Phosphorus

6558- 8398

Potassium

12860 - 16670

Calcium

130 - 1124

Magnesium

1041 - 1309

Sodium

1562 - 2824

Sulphur

7372- 12220

Zinc

<0.40 - 9.81

Manganese

0.46 - 0.73

Copper

<0.20

Iron

9.09 - 43.09

Aluminum

<2.50 - 5.19

Boron

<0.60 - 10.17

Cadmium

<0.20- 1.90

Nickel

<10.0

Arsenic

< 10.0

Mercury

< 10.0

Lead

< 10.0

Selenium

<8.0

species. The report indicated that, generally, fish sampled during spawning season were higher in moisture and lower in protein, lipid, ash, EPA and DHA contents compared to their winter counterparts. These differences were reported to be influenced by diet and seasonal factors as water temperature and physiological changes. The percent crude protein (dry weight basis) for fish 500 to 770 g ranges from 78.8 to 91,2%. Crude fat or lipids (dry weight basis) for the same fish ranges from 3.0 to 17.4%. 13.8 M A R K E T I N G During 1994, the largest hybrid striped bass producers harvested about 1,360,777 kg annually in tanks with the medium size producers harvesting about 68,039 kg annually in ponds. The live market is mostly for

344

Table 13.6. Fatty acid compositions in the muscle of hybrid striped bass produced in four states (Flick, unpublished data). Fatty Acid Florida a C 18

220.0

Concentration_in ppm Virginia b Mississippi a

Maryland ~

144.0

511.0

160.5

C 18:1

1457.0

691.0

4538.0

1353.5

C 18:2

775.7

432.5

2616.5

565.5

C18:3

123.0

45.0

368.0

92.0

C20

93.0

2.0

389.5

61.0

C22:1

77.0

81.0

---

64.0

5.0

3.0

10.0

---

C 18:4

341.3

194.5

450.0

195.5

C20:4

334.0

---

492.5

201.0

C20:5

898.3

348.5

1860.0

453.0

C22:6

859.0

342.0

1866.0

415.5

C24

_.. pond production b = recirculating system production a

smaller fish (under 2.0 lbs) while the fillet market came from larger fish (over 2.0 lbs). A large fish produces two fillets of 170-225 g each. It takes about 14 to 16 months to produce a 680g fish and 14 to 24 months in ponds to produce a 900g fish. In tanks with controlled temperatures it takes 8 to 10 months and 11 to 13 months respectively. Most of the fish sold live and on ice is distributed in major U.S. cities to individual (e.g., Asian) and institutional (e.g., food service) buyers. The Asian community prefers the whole product for steaming or frying while the Anglo communities consume the fillet as a baked product (Coale, 1993). 13.9 C L E A N I N G AND SANITIZING The U.S. Food and Drug Administration has responsibility for enforcement of the Food and Drug Act, which has as its main purpose..." to prohibit the movement in interstate commerce of adulterated or misbranded foods." The FDA holds a food to be adulterated if it contains any poisonous or deleterious substance which may render it injurious to health, if it contains any unsafe food additive or color additive, if it consists of any filthy, putrid, or decomposed substance, or if it has been prepared, packed, or held under unsanitary conditions whereby it may have become contaminated with filth or rendered injurious. A sanitation program must be aimed at preventing production of fish which would be subject to action under this act, but more importantly the program, in conjunction with the quality control program, must insure a product with adequate shelf-life and good salability and acceptability.

345

Table 13.7. Proximate analysis of wild-captured striped bass and aquacultured hybrid striped bass (Karahadian et al., 1995). Sample Description Moisture

Proximate Analysis Lipid Protein

Ash

% Composition 1 Wild-captured Spring Potomac Upper Bay Choptank

78.86 b 79.36 b 79.71 b

2.63 ~ 2.55 ~ 1.8T

19.6 ~ 19.0 ~ 18.1 b

1.22 ~ 1.56 b 1.09~

Wild-captured Winter Potomac Upper Bay C hoptank

78.86 b 77.88 b 74.70 a

3.25 d 3.51 d 2.65 c

21.8 d 20.4cd 21.9 a

1.15 a 1.64 b 1.20 a

Aquaculture Palmetto Bass Sunshine Bass

72.69 ~ 78.22 b

2.84 c 2.63 c

24.8 e 18.9 bc

1.59 bc 1.17"

1 Values reflect means of four replications. Mean values in the same column with different superscripts are significantly different (P>0.05).

During dressing (manual and mechanical), cleaned and sanitized processing environments and equipment are soiled and contaminated by inherent microflora and processing waste (as flesh trimmings, blood, offal, skin, and slime). These inherent and adventitious microorganisms multiply and decompose processing waste. As dressing progresses, contaminated machinery, knives, cutting boards, totes, and employee hands alter the attributes of aquacultured products. Therefore, plant cleaning and sanitizing are an integral component in attaining and sustaining a high quality product in all food factories, including striped bass processing plants. Plant cleaning and sanitizing are individualistic and interdependent, yet indispensable functions. Plant cleaning encompasses the removal of objectionable matter from the processing environment and equipment, whereas sanitizing encompasses the reduction and elimination of pathogens and food spoilage microorganisms. The processing environment and equipment are considered clean when all the processing waste is removed. Generally, cleaning decreases the inherent and adventitious bacterial count. Cleaning is facilitated by the incorporation of cleaning (detergent) agents. Detergents hydrate and penetrate the soil, solubilize as well as hydrolyze proteins, emulsify and saponify lipids, and solubilize deposited minerals. The cleaning technique and not the cleaning agent differentiates an efficient from an inefficient cleaning. Cleanliness is a visible attribute, and inspection reveals inefficient cleaning. Sanitation is an invisible attribute and, although a processing environment may be clean, it may be contaminated with a diverse bacterial population.

346

Following cleaning, a sanitizer is applied to kill residual microorganisms exposed by the cleaning process. A sanitizing (disinfecting) agent essentially reduces or eliminates the adventitious spoilage microorganisms to a level judged safe by public health requirements. Additionally, it ensures a safe and wholesome product with an adequate shelf-life. An effective disinfectant must possess excellent sanitizing properties to kill microorganisms entrapped in soiled layers. Sanitizer effectiveness should result in a significant reduction in bacterial numbers. Sanitization success depends on the preceding cleaning efficacy, surface composition, sanitizer potency, and application technique. Sanitizing must follow a thorough cleaning process because soiled surfaces cannot be sanitized. 13.9.1 Cleaning and Sanitizing Principles Cleaning and sanitizing processes are accomplished in two ways: (a) Cleaning and sanitizing in-place is a process utilizing pipes for transportation and steam sterilization for aseptic operations, and is used infrequently in the aquacultural industry; (b) Cleaning and sanitizing out-of-place -- is a process utilized most often for batch processes and handling equipment, and requires dismantling of machinery. Cleaning and sanitizing out-of-place could be used most extensively in striped bass processing plants. Independent of practiced protocol, the principal steps are common to both processes. Dry cleaning is the first step in an effective cleaning and sanitizing program. An effective dry clean procedure reduces the amount of time and personnel required for cleaning and can reduce waste disposal cost by restricting the amount of solid and suspended materials discharged into the sewer. Processing wastes are removed from equipment, utensils, and floor by hand, brush, or broom and shovel. If large quantities of the materials are retrieved, they may be recycled into fish feed or another animal ration. The second step is the initial water wash. During this step the maximum percentage of the processing waste is dislodged and the microbial load is reduced. This process is accomplished with potable water that is of good microbiological quality and essentially free of pathogens. Chemical characteristics are used t o broadly classify water types, such as soft and hard waters. Soft water is superior to hard water for preliminary cleaning. The hardness of water is reduced by heating (temporary hardness) or chemical treatment (permanent hardness). Thermal (warm), kinetic (pressure), and mechanical (brush or scrubber) energies could be used in several combinations for effective preliminary cleaning. Pressurized (435-930 psi) warm (above melting point of fat) water would dislodge proteins and liquefy fat. Processing waste however, continues to adhere after the initial wash water is solubilized with the cleaning solutions. Cleaning agents with diverse chemical properties are used for cleaning. The cleaning agents are broadly classified as alkaline and acidic. Alkaline cleaning agents are general in nature with sodium salts of hydroxide, orthosilicate, sesquisilicate, metasilicate, and carbonate being the strongest cleaning agents, while sodium bicarbonate is a mild alkaline cleaning agent. They exhibit similar emulsifying properties. Sodium salts of metasilicate and orthosilicate have excellent cleaning capability in comparison to hydroxide, sesquisilicate, carbonate, and bicarbonate. All alkaline cleaning agents are corrosive, with sodium hydroxide being the most corrosive. Additionally, sodium hydroxide does demonstrate some bacteriocidal properties. Sodium carbonate can produce calcium carbonate scale, which is a major drawback. While most alkaline cleaning agents are highly soluble, sodium metasilicate has a low solubility. An acidic cleaning agent may be necessary if the alkaline cleaning agent is inefficacious. When both alkaline and acid cleaning agents are used, it is necessary to use an additional water wash between the two diverse cleaning agents. Acid cleaning agents are used for specific functions, and they are ineffective against lipid and protein soils. Their most common role is to dissolve divalent mineral salts termed as waterstones,

347

which are hard, tenaciously adherent deposits that support microbial multiplication. Strong mineral acids (e.g., hydrochloric, sulfuric) are used as acidic cleaning agents, however, the compounds are corrosive to metal (e.g., black iron), concrete, and fabric. An intermediate wash rinses the alkaline and/or acidic cleaning agents, and is followed by application of sanitizing solution. The role of sanitizing solution is to reduce or kill residual microorganisms. Sanitizers are applied through automatic dosing, foaming, flooding, or fogging. The most effective and economical method for each operation must be employed. Some sanitizers should not be used in particular areas of the operation because of their chemical composition. Sanitizing agents used in the food industry are broadly classified as thermal, radiant, and chemical agents. Steam and hot water are used as thermal sanitizing agents and can kill bacterial spores. Hot water is an effective, nonselective sanitizing method for food contact surfaces. This method is used for sanitization-inplace in the food industry as well as immersion sanitization of processing accessories (e.g., knives, machine parts, holding containers, etc.). Time-temperature combinations determine the efficacy of the sanitizing process. The use of hot water exceeding a temperature of 63 ~ should be avoided to prevent the soil from adhering to food contact surfaces. Cooked product usually requires hand scrubbing for complete removal. Radiant sanitizing agents such as electromagnetic (e.g., ultraviolet, high-energy cathode, gamma ray) waves can injure and/or kill bacteria. Ultraviolet units are used in food factories to disinfect processing water and can be used in the aquacultural industry to improve the bacterial quality of processing water. Ultraviolet light is ineffective in the presence of suspended solids and turbid solutions. Also, UV lamps become decreasingly effective with age and should be replaced on a scheduled basis. Chemical sanitizing agents used in food processing plants fall into the following categories: halogenated compounds (e.g., chlorine, iodophor), and quaternary ammonium compounds. Chlorine compounds are probably the most commonly used sanitizers in the fish industry. Use of chlorine prevents or greatly reduces the accumulation of microbial slimes on equipment that is continuously or frequently washed with chilled chlorinated water and it prevents the development of malodorous fermentative decay. The use of chlorine also aliows longer time for processing operations by reducing the time for cleanup. Chlorinated compounds are useful in reducing spoilage microorganisms and thus increasing the shelf-life of the product. Hypochlorite dissolved in water (50 ppm of chlorine) is a cheap and competent sanitizer. It is used for sanitizing plant environments and equipment following a thorough cleaning, however, it is inactivated by organic (e.g., protein) compounds. Chlorine dioxide has 2.5 times the oxidizing power of chlorine and its effect is greatest at pH 8.5. It is unaffected by alkaline conditions and organic matter. Chlorinated systems should not be relied on to solve sanitation problems. The indiscriminate use of chlorine cannot compensate for unsanitary conditions in a processing plant. The major iodine compounds used for sanitizing are iodophors and iodine in alcohol, as well as aqueous iodine solutions. Iodophors are complex iodine sanitizers. They are the most popular form of iodine compounds and have greater bacteriocidal activity under acidic conditions. Complexing iodophor with surface active agents and acids gives them detergent properties and qualifies them as detergent-sanitizers. These compounds are bacteriocidal and, in comparison to aqueous and alcoholic suspensions of iodine, have greater solubility in water, are nonodorous, and nonirritating to the skin. Iodophors are used as hand-dips in some fish processing facilities. Quaternary ammonium compounds come in many forms and in a wide variety of commercial products. Theyare less versatile than chlorine, but are better wetting agents and, because they do not evaporate, provide

348

a residual germicidal action. They are particularly effective in controlling mold ~owth on walls and ceilings of coolers. The major disadvantage of the compounds is that they leave a residual film. This residue must be removed from food processing surfaces before they are used if concentrations greater than 200 ppm are applied. Acidic sanitizing compounds are organic (e.g., formic, acetic, lactic, propionic) acids. They are generally regarded as safe for use in food, are stable to heat, organic matter (e.g., fish slime, protein, and fat residues), and are safe for use on most food contact surfaces (e.g., stainless steel, polyester, plastic). However, they are costly and corrode metal (e.g., black iron). Among them, propionic acid has the best sanitizing effect. Rinsing-off sanitizer in the form of a potable water rinse is sometimes required because of potential problems with worker discomfort and odor. The application technique is specific for each sanitizer. After rinsing off a sanitizer, thorough draining or drying is essential. Wet surfaces stimulate and support microbial multiplication whereas dry surfaces discourage it. Sanitizing adjuncts are an important constituent of sanitizing compounds. Chelates are a useful class of compounds with limited versatility. They react with the divalent metal ions (e.g., calcium, magnesium) and staining metals (e.g., iron) and result in water soluble complexes. Sodium salts of ethylenediaminetetraacetic acid (EDTA) and gluconate are the most widely used chelators. Surfactants have advanced the technology of detergents (synthetic soaps). Soaps perform most of the desirable functions of a detergent, however they have some serious shortcomings because they are precipitated by hard water and are limited to the alkaline pH range. An excessive concentration to precipitate the divalent metal ions (e.g., calcium, magnesium) could overcome some shortcomings but are expensive. Surfactants have largely supplemented soaps in the detergent industry. Their chemical composition and classification is very complex, and are broadly classified as anionic, nonionic, and cationic surfactants. 13.9.2 Cleaning and Sanitizing Plant Equipment Equipment such as dressing tables, chill kill tanks, conveyor belts, deheading saws, skinning machines, and floors and walls are cleaned with a high pressure sprayer. Preliminary equipment cleaning with warm water (e.g., 40~ ) working from the top downward until all organic substances have been removed is required. Utensils and removable machine parts, valves, and similar pieces of equipment should be hand-washed and sanitized. A heavy-duty alkaline cleaning agent should be applied, preferably with foaming equipment, however, this type compound should not be used on aluminum or galvanized metal. The nozzle from the soiled surface (approx. 1.0 m), and the cleaning agent should be allowed to remain on the surfaces for 15 min to hydrate the soil. Subsequently, the equipment should be washed with warm water (55~ with a spray nozzle positioned about 15 cm from the surface of the equipment. The surface should be rinsed with potable water (25~ using a hose or pressure gun. A sanitizing solution (e.g., chlorine or quaternary ammonium) should be used just before initiating processing operations. An acidic cleaning solution should be used periodically (weekly) to remove mineral deposits. Floors are best cleaned using a specially designed detergent for that purpose. First, dry clean to remove gross debris and spread the detergent over the floor surface at about 454 g per 4.7 square meters. Scrubbing with a good brush is recommended then rising with ambient water or an acid rinse at pH 5. The floor should be sanitized by applying a 200 ppm chlorine solution at ambient temperature. For the sanitizing procedure, a good iodophor at 50 ppm or 400 ppm quaternary ammonium compound may be substituted for the chlorine solution.

349

At the end of each day, processing accessories (filleting knives, plastic boards, bone clippers, knife sharpeners, etc.) should be rinsed in ambient water, than soaked and scrubbed in warm water. The accessories should be transferred into an alkaline cleaning solution. The processing accessories can then be rinsed with warm water, then immersed in a sanitizing solution. Accessories should be allowed to drip and dry overnight. For rinsing of hands before returning to work by those actually handling fish, a 25 ppm iodine solution is satisfactory. 13.9.3 Cleaning and Sanitizing for Plant Employees Employees working in the processing room should change into their work clothing, then walk through a footbath to sanitize their shoes, clean their hands, and dress in disposable or sanitized clothing (e.g., gloves, apron, hairnet, etc.) before they initiate their workday. Employees leaving their work station for a break or restroom visit must remove their clothing before leaving the processing floor. Employees resuming work after a break or restroom visit must follow the same sequence of operations as at the start of work. Disposable clothing should be discarded at the end of each work day. Faucets which are activated optically or by foot or knee valves are helpful in restrooms and could be installed to reduce microbial transfer (fecal contaminants) from restroom to processing room. Rest-rooms are excellent reservoirs of fecal microbes. It may be necessary for plant maintenance personnel to physically contact processing equipment during processing for maintenance. After repairs, the food contact surface must be cleaned and sanitized before resuming normal operations. A proactive managerial support is essential to execute a successful cleaning and sanitizing program. Because hybrid striped bass processing plants are in their infancy, they are developing at a time when HACCP programs are being implemented thus the cleaning and sanitizing protocols can be established simultaneously. It is desirable that each processing plant initiate its own cleaning and sanitation department which should be independent of production functions. Cleaning and sanitizing personnel must be trained in cleaning, dismantling, and assembling equipment, as well as the economical significance of bacterial spoilage and the liability of hazard. A permanent cleaning and sanitizing schedule should be used to ensure that the entire facility is cleaned and sanitized on a timely basis. Management should evaluate cleaning and sanitizing program efficacy, as well as enforce compliance with guidelines and standards of regulating agencies. The program's practices should be recorded, reviewed, and revised when necessary. 13.9.4 Pest Management The key to good insect control is a well constructed building and a good sanitation program. The necessity for housefly control is pointed by the fact that one housefly carries about 1.25 million bacteria. Many of these bacteria are of the type incriminated in public health problems. For use inside processing plants, the recommended insecticides are pyrethrum, allethrin, or rotenone. Concentrations should not exc~d 1% solutions. To deny insects entrance to the plant, adequate screens on all doors and windows are essential. At conveyor entrances and other openings where screens are not practical, water or air sprays may be used. Air at a velocity of 5 meters per minute will exclude 80% of flying insects. Air screens should be directed at a slight outward angle from the top of the opening. Use of a good residual insect spray on the outside of buildings and the grounds will greatly reduce the numbers of insects available to gain entrance to the plant. Rodent control is best attained by a good maintenance program which eliminates hiding and breeding places near the processing plant. Grasses and weeds should be kept at 10 cm or less in height. The original design of the plant is the most decisive factor in excluding rodents. Sheet metal, 3 mm mesh hardware cloth,

350

or concrete may be used to close openings left by plumbers and electricians. Doors and windows must fit snugly and air intakes and exhaust openings must be tightly screened. Bait boxes may be used outside the processing plant to control rodents in the vicinity when neighboring property owners fail to keep them under control. Many food processors contract with pest control operators for rodent control services. Sanitation in a hybrid striped bass processing plant starts with the architect who designs the facility and it continues with the engineers who build it. Another vital link is the machinery design and lay-out. The management of fish processing facilities are both legally and morally responsible to see that fish are handled so that they are clean and wholesome. A good sanitation program is not really very expensive, but the lack of one can be very costly. The program will be only as good as the degree of support of management. 13.10 WASTE HANDLING Substantial liquid and solid wastes may be generated by hybrid striped bass processors. Federal, state, and local regulatory agencies as well as environmentally conscious citizens are demanding prompt, complete, responsible, and better waste handling from all industries. Food factories discharging high biochemical oxygen demanding effluents have to pay surcharges if discharged into municipal systems. Therefore, solid and liquid wastes must be economically exhausted using the criterion and standards set by federal, state, and local regulators. Processors must explore methods that reduce expenses incurred for disposing liquid and solid wastes. Conversely, liquid and solid constituents may be transformed into profitable products such as minced fish, feed ingredients, silage, and compost before they become wastes. Potable processing water used in aquacultural processing facilities is essential to clean incoming hybrid striped bass, the processing environment, and equipment. Liquid waste is a result of potable processing water, and contains organic (e.g., protein, lipid) and inorganic (e.g., minerals) constituents. Improper discharge of liquid waste is hazardous to terrestrial and aquatic creatures as well as the environment. Organic waste is decomposed by proliferating microbes, and it emits malodorous. Decomposing organic matter reduces the availability of biochemical oxygen and changes the malodorous fermentative decomposition from aerobic to anaerobic, which has greater odor problems. To eliminate these problems, organic and inorganic materials must be treated before discharging effluents into the sewage system. Regulations have been developed to encourage industrial wastewater management programs to prevent, control, or reduce pollutants. Large food processors have initiated liquid and solid waste treatments. To initiate a waste treatment system, the processor must have complete information on liquid and solid wastes, which includes the volume, frequency (e.g., operating schedule), and composition of the waste. Plant design (i.e., piping, equipment layouts), should be analyzed to determine incoming and outgoing wastes. The quality and quantity of products processed in a defined period (e.g., daily, weekly, monthly, seasonally, or annually) is also important. Production and water usage logs are beneficial in waste treatment analysis. A sample of the liquid waste should be compared with the federal, state, and local discharge criterion. Volumetric (e.g., water) and mass (e.g., production) balances would reveal unaccounted losses (e.g., leaks). Sampling the effluents' volumetric flow rate for each unit operation provides information on liquid waste composition. Identification of representative sampling locations and frequencies should be made to avoid erroneous results. An analysis of the samples should be made for biological oxygen demand, chemical oxygen demand, dissolved oxygen, total organic carbon, residue in liquid waste, total suspended solids, total dissolved solids, fats, oils and grease, and total phosphorous contents.

351

13.10.1 Liquid Disposal Processing water is the most common liquid waste in hybrid striped bass processing plants, and the best solution to disposal of processing water is reducing its generation. Liquid waste can be salvaged through recycling (recovery and reuse). Liquid waste, unacceptable for discharge into sewers, may be subjected to physical processes for reducing effluents. Insoluble organic waste can be recovered by screening with dynamic (e.g., vibratory, rotatory) or static screens. Sedimenting solids can be removed with clarifying tanks. Biological and chemical treatment processes follow those ofwastewater treatment for soluble solids, and biological degradation of organic waste through oxidation is the most common technique. Both aerobic and anaerobic lagoons can be used to reduce the biological oxygen demand. Microflora in the lagoon oxidizes the soluble solids and produces sludge solids. Various chemicals are used for chemical oxidation and bacterial disinfection. Ozone, chlorine, chlorine dioxide, and permanganate are used as chemical oxidants. The nascent oxygen generated from ozone breaks down rapidly and oxidizes organic matter. Chlorination produces organochlorides and caution should be used because organochlorides are potential carcinogens. Additionally, over chlorination of wastewater effluents can be toxic to terrestrial and particularly aquatic animals. Electromagnetic waves such as ultraviolet, gamma, and microwave can also be used to disinfect liquid waste and recycle. 13.10.2 Solid Disposal The accumulated solids obtained from retaining devices (e.g., filtering, screens, and settling basins) and solid waste from hybrid striped bass (e.g., offal, skin, head, and frame) must be removed and disposed appropriately. Solid waste usually has a high moisture content, thus, dewatering before disposal is recommended to economize on handling and transport. Most landfills will not accept liquid processing wastes. Dewatering systems can apply heat and/or pressure to reduce moisture content, but economics for dewatering must be compared to reduced costs for transporting and disposing. Occasionally, dewatering costs are unavoidable in providing solids which are compatible with the available disposal option. Disposal methods must consider odor, insect, and decomposition problems, as well as a general adverse public opinion regarding unpleasant aesthetic aspects. Trucking to municipal dump sites is the most common disposal method because of the small processing volume. Alternatively, the solid waste (e.g., head, frame, skin, offal) could be used as fertilizer (e.g., raw, wet or dry) or animal feed after pulverizing. Small processors located in rural locations may spread the solid waste without treatment as raw fertilizer, or the solid waste can also be disposed by composting. The finished compost can then be used as fertilizer and soil amendment. Organic material is stabilized through metabolism and multiplication of microbes, and humus resulting from stabilization of solid waste improves fertility and tillage properties. Developing these markets would depend on competitive cost and any unique features in the fertilizer. Utilization of by-products may prove to be the most economically effective solid waste treatment option, and should be examined as a least-expensive treatment option as well as for profitability. If an individual food industry processes a small quantity of volume, it may not be economical for them to solve their waste disposal problems alone. In that case, it may be more productive for a group of food processors located in the same region to cooperate in solving their liquid and solid wastes disposal problems together.

352

13.10.3 Solids Utilization Solid waste can be used as a source of protein or lipid. Edible product may be recovered for animal feeds, the most popular of which is meal or dehydrated protein. The production of animal feed from solid waste has become a less favored option due to the costs for heating fuels and cost competition from other protein meals (e.g., animal meat, soybean). The development of the striped bass industry will provide an impetus for using more fish waste as feed. As the hybrid striped bass processing industry grows, other by-products may become more useful for human consumption, including minced fish and roe where lipids can be recovered through rendering. There are basically three types of rendering systems: dry-rendering, wet-rendering, and enzymatic hydrolysis. Dry rendering is a high temperature process which heats coarsely ground waste until the moisture content is reduced to 3%. The proteinaceous meal is then ground in a hammer mill and conveyed to a storage bin. Wet-rendering is a low temperature process in which the waste is cooked at atmospheric pressure in a steam jacketed vessel. At cooking, the offal is then expressed through a low-pressure screw press, which results in a 50% moisture press cake. The expressed liquid is then used for recovering the oil. For enzymatic hydrolysis, the offal is pulverized to facilitate enzymatic digestion. Many different commercial enzymes are available, and each is active over a variety of optimal pH and temperature ranges. By adjusting the reaction conditions (e.g., pH, temperature, time, enzyme specificity and concentration), enzymatic hydrolysis can be manipulated. Various unit operations (e.g., separating, concentrating, and drying) can be used to achieve the desired products.

353

References

Ammerman, G.R., 1985. Processing. Pages 569-616 in C. S. Tucker, editor. Channel catfish culture. Elsevier Science Publisher, Amsterdam, New York. Armbruster, G., Gerow, K.G., Gutenmann, W.H., Littman, C.B. and Lisk, D., 1987. The effects of several methods of fish preparation on residues of polychlorinated biphenyls and sensory characteristics in striped bass. Journal of Food Safety, 8: 235-243. Armbruster, G., Gutenmann, W.H. and Lisk, D.J., 1988. The effects of six methods of cooking on residues of mercury in striped bass. Nutrition Reports International, 37(1): 123-126. Bakker, M., 1986. The Wiley encyclopedia of packaging technology. John Wiley & Sons, New York. Barrett, I., Brinner, L., Brown, W.D., Bolev, A., Kwon, T.W., Little, A., Olcott, H.S., Schaefer, M.B. and Schrader, P.P., 1965. Changes in tuna quality and associated biochemical changes during handling and storage aboard fishing vessels. Food Technology, 19 (12): 108-117. Bendall, J.R., 1972. Poslrnortem changes in muscle. Pages 243-309 in G.H. Bourne, editor. The structure and function of muscle. Academic Press, New York. Black, E.C., Connor, A.C., Lam, K. and Chiu, W., 1962. Changes in glycogen, pyruvate and lactate in rainbow trout (Salmo gairdneri) during and following muscular activity. Journal of Fisheries Research Board of Canada, 19: 409-436. Botta, J.R., Bonnell, G. and Squires, B.E., 1987. Effect of method of catching and time of season on sensory quality of fresh raw Atlantic cod (Gadus morhua). Journal of Food Science, 52:928-931,938. Boyd, L.C., Gree, D.P. and LePros, L.A., 1992. Quality changes of pond-raised hybrid striped bass during chillpack and refrigerated storage. Journal of Food Science, 57: 59-62. Carlberg, J.M. and Van Olst, J.C., 1987. Processing and marketing. Pages 73-82 in R.G. Hodson et al., editors. Hybrid striped bass culture: status and perspective. University of North Carolina, Sea Grant College Publication, UNCSG-87-03, Raleigh. Coale, C.W., Anthony, J.P., Flick, G.J., Libey, G.S., Hong, G.-P., Spittle, G.D. and Valley. N.A., 1993. Marketing aquaculture products: a retail market case study for sunshine bass. Virginia Aquaculture Experimental Station Bulletin. 93-3.56 p. Crawford, L., Irwin, E.J., Spinelli, J. and Brown, W.D., 1970. Premortem stress and postmortem biochemical changes in skipjack tuna and their relation to quality of the canned product. Journal of Food Science, 35:849-851. Curiale, M.S., 1991. Shelf-life evaluation analysis. Dairy, Food and Environmental Sanitation, 11: 364-369. Dunajski, E., 1979. Texture of fish muscle. Journal of Texture Studies, 10:301-318. Eifert, J.D., 1991. Quality changes of aquacultured hybrid striped bass fillet meat resulting from reduction of postharvest metabolism. M.S. thesis. Virginia Polytechnic Institute and State University, Blacksburg. Erickson, M.C., 1993. Ability of chemical measurements to differentiate oxidative stabilities of frozen minced muscle tissue from farm-raised striped bass and hybrid striped bass. Food Chemistry, 48:381-385.

354

Fair, P.H., Williams, W.P. and Smith, T.I.J., 1993. Effect of dietary menhaden oil on growth and muscle fatty acid composition of hybrid striped bass, Morone chrysops x M. saxatilis. Aquaculture, 116:171-189. Food and Drug Administration., 1992. Bacteriological analytical manual. 7th edition, PUb. AOAC Intemational, Arlinpon, VA. Fowler, K.P., Karahadian, C., Greenberg, N.J. and Harrell, R.M., 1994. Composition and quality of aquacultured hybrid striped bass fillets as affected by dietary fatty acids. Journal of Food Science, 59: 70-75, 90. Frazier, W.C. and Westhoff, D.C., 1978. Contamination, preservation, and spoilage of fish and other seafoods. Pages 243-255 in Food microbiology. McGraw Hill Book Company, New York. Graham, J., 1982. Freezing. Pages 56-71 in A. Aitken, I.M. Mackie, J. H. Merritt, and M.L. Windsor, editors. Fish handling and processing. Her Majesty's Stationary Office, Edinburgh, Scotland. Hodson, R., Smith, T., McVey, J., Harrell, R. and Davis, N., 1987. Hybrid striped bass culture: status and perspective. University of North Carolina Sea Grant College Publication UNC-SG-87-03, Raleigh. Jacober, L.F. and Rand. A.G., 1982. Biochemical evaluation of seafood. Pages 347-365 in R.E. Martin, G.J. Flick, C.E. Hebard, and D.R. Ward, editors. Chemistry and biochemistry of marine food products. AVI Publishing Co., Westport, CT. Johnson, E.A., Segars, R.A., Kapsalis, J.G., Normand, M.D. and Peleg, M., 1980. Evaluation of the compressive deformability modulus of fresh and cooked fish flesh. Journal of Food Science, 45:1318-1320, 1326. Karahadian, C., Fowler, K.P. and Cox, D.H., 1995. Comparison of chemical composition of striped bass (Morone saxatilis) from three Chesapeake Bay tributaries with those of two aquacultured hybrid striped bass types. Food Chemistry, 54:409-418. Korhonen, R.W., Lanier, T.C. and Giesbrecht, F., 1990. An evaluation of simple methods for following rigor development in fish. Journal of Food Science, 55: 346-348, 368. Kraft, A.A. and Rey, C.R., 1979. Psychotrophic bacteria in foods: an update. Food Technology, 33: 66-71. Liston, J., 1980. Fish and shellfish and their products. Pages 567-605/n J. H. Silliker, editor. Microbial ecology of foods, volume II, food commodities. Academic Press, New York. Liston, J., 1982. Recent advances in the chemistry of iced fish spoilage. Pages 27-37 in R.E. Martin, G.J. Flick, C.E. Hebard and D.R. Ward, editors. Chemistry and biochemistry of marine food products. AVI Publishing Co., Westport, CT. Liston, J. and Matches, J.R., 1976. Fish, crustaceans, and precooked seafoods. Pages 507-521 in M.L. Speck, editor. Compendium of methods for the microbiological examination of foods. American Public Health Association, Washin~on. Love, R.M., 1988. Texture. Pages 121-140 in The food fishes: their intrinsic variations and practical implications. Van Nostrand Reinhold Co., New York. Mayer, B.K. and Ward, D.R., 1991. Microbiology of fmfish and fmfish processing. Pages 3-17 in D.R. Ward and C.R. Hackney, editors. Microbiology of marine foods products. Van Nostrand Reinhold, New York.

355

Mitsuda, H., Nakajima, K., Mizuno, H., Kawai, F. and Yamamoto, A., 1980. Effects of carbon dioxide on carp. Journal of Nutritional Science and Vitaminology, 26: 99-102. Murata, H. and Yamaguchi, K., 1990. Relationship between the 2-thiobarbituric acid values and polyunsaturated fatty acid concentrations of some tissues from cultured red sea breams. Nippon Suisan Gakkaishi 56:761-764. Nedoluha, P.C. and Westhoff, D., 1993. Microbiological flora of aquacultured hybrid striped bass. Journal of Food Protection, 56:1054-1600. Pedrosa-Menabrito, A. and Regenstein, J.M., 1988. Shelf-life extension of fresh fish - a review, Part I - spoilage of fish. Journal of Food Quality, 11:117-127. Plakas, S.M., DePaola, A. and Moxey, M.B., 1991. Bacillus stearothermophilus disk assay for determining ampicillin residues in fish muscle. Journal of the Association of Official Analytical Chemists, 74:910-917. Regenstein, J.M., Schlosser, M.A., Samson, A. and Fey, M., 1982. Chemical changes of trimethylamine oxide during flesh and frozen storage offish. Pages 137-147 in R.E. Martin, G.J. Flick, C.E. Hebard and D.R. Ward, editors. Chemistry and biochemistry of marine food products. AVI Publishing Co., Westport, CT. Regenstein, J.M and Regenstein, C.E., 1991. Introduction to fish technology. Van Nostrand Reinhold, New York. Rehwoldt, R.E., Mastrianni, W., Kelley, E. and Stall, J., 1978. Historical and current heavy metal residues in Hudson River fish. Bulletin of Environmental Contamination and Toxicology 19: 335-339. Shewan, J.M., 1977. The bacteriology of fresh and spoiling fish and the biochemical changes induced by bacterial action. Pages 51-66/n Proceedings Conference on the Handling, Processing and Marketing of Tropical Fish. Tropical Products Institute, London. Sikorski, Z.E., Kolakowska, A. and Burt, J.R., 1990. Postharvest biochemical and microbial changes. Pages 55-76 in Z. E. Sikorski, editor. Seafood: resources, nutritional composition, and preservation. CRC Press, Inc., Boca Raton, FL. Smith, T.I.J., Jenkins, W.E. and Snevel, J.F., 1985. Production characteristics of striped bass (Morone saxatilis) and Fi, Fz hybrids (M. saxatilis and M. chrysops) reared in intensive tank systems. Journal of the World Mariculture Society, 16: 57-70. Swartz, D., 1984. Marketing striped bass. Pages 233-254 in J. McCraren, editor. The aquaculture of striped bass. University of Maryland Sea Grant College Publication UM-SG-MAP-84-01. College Park. Tarr, H.L.A., 1966. Post-mortem changes in glycogen, nucleotides, sugar phosphates, and sugars in fish muscles - a review. Journal of Food Science, 31: 846-854. U. S. Department of Health and Human Services, Public Health Service, Food and Drug Administration., 1993. DHHS Publication No. (FDA) 93-1051, Rockville, MD. U. S. Department of Health and Human Services, Public Health Service, Food and Drug Administration., 1995. 21 CFR 110. Rockville, MD. Van Olst, J.C. and Carlberg, J.M., 1990. Commercial culture of hybrid striped bass: status and potential. Aquaculture Magazine 16(1): 49-59.

356

Venugopal, V., 1990. Extracellular proteases of contaminant bacteria in fish spoilage: a review. Journal of Food Protection, 53:341-350. White, R.J., Kim, H.T. and Kim, J.S., 1985. PCBS in striped bass collected from the Hudson River, New York, during fall 1981. Bulletin of Environmental Contamination and Toxicology, 34: 883-889. SUGGESTED READING Aitken, A., Mackie, I.M., Merritt, J.H. and Windsor, M.L., 1982. Fish handling and processing. Edinburg,h, Scotland: Her Majesty's Stationary Office. Hodson, R.G., 1991. Hybrid striped bass culture in ponds. Pages 167-191 in J. P. McVey, editor. Section IV: Marine f'mfish aquaculture. Handbook of mariculture, volume II. CRC Press Inc. Boca Raton, FL. Norton, V., Smith, T. and Strand, I., editors., 1983. Stripers: the economic value of the Atlantic coast commercial and recreational striped bass fisheries. University of Maryland Sea Grant College Publication, UM-SG-TS-83-12, College Park. Partmana, W., 1963. Post-mortem changes in chilled and frozen muscle. Journal of Food Science 28:15-27. Pigott, G.M. and Tucker, B.W., 1990. Food from the sea. Chapter. 1, in Seafood effects of technology on nutrition. Marcel Decker, New York. Ring, C. and Schlager, B., 1988. Meat quality of CO2- stunned pigs. Fleischwirtschaft 68: 1532-1534. Stickney, ILR., 1979. Handling, processing, and economics. Chapter. 8 m Principles of warmwater aquaculture. John Wiley and Sons, New York. Strand, I.E., Norton, V.J. and Adriance, J., 1980. Economic aspects of commercial striped bass harvest, in H. Clepper, editor. Fifth annual marine recreational fisheries symposium, Boston, Sport Fishing Institute, Zeller, W., Mettler, D. and Schatzmann, U., 1988. Studies on stunning of slauoJater poultry with C O 2. Fleischwirtschaft, 68:1308-1312.

Appendix

358 4.10 APPENDIX A Carbonate Equilibrium Chemistry_(from Stumm and Morgan, 1981)

G,, m

2

m

g

o

.> <_-=

-0.5

1 CT(Total

CH

2

carbonate

C02

carbon;

millimolerJliter|

CaCO3 ,, Na2 CO3

NaHCO3

2 !

ca

. . . . .

,I

_i

1 1

Fig. A-1. Alkalinity vs. Cr at varying pH.

Dilution

359

IHCO;] = CT~ 1

H2CO3"] = CT(go

K t~o=

1+

-

KIK2).-1

: _ [H *] [-H .]2

0~1 =

+1

+

K1 ,12 + [H*I

+ 1

)-1

Fig. A-2. Carbonate ionization fractions as a function of hydrogen ion concentration.

Table A-1. First Acidity Constant: H2CO3= = HCO3- + H § Temp (~

"

-log K~

Table A'2. Second Acidity Constant: H § + CO3 =

HCO3"

-log K2

Temp. (~

,,,

6.579

10.625

6.517

10.557 10.490

10

6.464

10

15

6.419

15

10.430

20

6.381

20

10.377

25

6.352

25

10.329

30

6.327

30 84

35

6.309

35

10.250

40

6.298

40

~0.22o

50

6.285

50

9

o.29o

10.172 _

Fig. A-3. Equilibrium constants for carbonic acid vs. temperature.

This Page Intentionally Left Blank

361

Subject Index Achyla 289,

Acidity (see pH) Aeration -, carbon dioxide and 192, 202 -, emergency 87 -, in fish transport 187, 192-202 -, operation considerations 91, 111-119, 130 -136, 143-165 -, principle 105-110, 261 -, supplemental 83, 93, 118, 301,317 -, water circulation and 108, 261 Aerators -, oxygen transfer rates 104-110, 116 -,types 84, 87, 130-144, 152-153 Aeromonas sp. 6, 7, 275-284, 293,341 Alcian blue 15, 23 Algae 7, 135 -, calculation of standing crop 103, 112-116 Alkalinity -, alkalinity destruction 121 -123 -, carbon dioxide and 100, 209 -, effect on copper toxicity 131,262, 307 -,pH and 100, 119-121,157, 163,203, 255, 259, 359 -, phytoplankton communities and 259 Amino acids (see also Protein) 19, 21,237-245, 340, 342343 Ammonia -, concentrations in tanks 121-122, 135, 162-163 -, effect on fish growth 99-101,130-131,149, 185, 284 -, excretion of 237, 259-261 -, fish transport and 197, 201-203,209 -, nitrification of 151, 157 -, pH and forms of 187, 195, 201-203 -, toxicity 131,149, 151,162-163, 254, 259-261 -, volatization 340 Androgens -, testosterone 19, 25-28, 31-36, 42-46, 180 -, 1 l-ketotestosterone 27, 37 Anesthetics 50, 172, 95-99, 202-208, 254, 262-264 Antibiotics 203,305 Antibodies 274 Antioxidants -, in processed fish 333 Aquasim 321 Aquatic weeds 6-7 Ascorbic acid 242, 244, 246, 333-334 Bacteria -, in processed fish 306-307, 332, 337, 340-341,347, 350 Bacterial diseases -, Aeromonas sp. 6, 7, 275-281,284, 293, 341 -, Chondrococcus sp. 278 -, Carnobacterium sp. 275, 277, 287, 288 -, Corynebacterium sp. 277, 286 -, Cytophaga sp. 278, 341 -, Edwardsiella sp. 275, 277, 280, 282, 309 -, Edwardsiellosis 277, 280 -, Enterococcosis 277, 285

-, Enterococcus sp. 277, 285 -, Flexibacter columnaris 6, 7, 272, 275,277-281,307 -, Moraxella sp. 288 -, Motile aeromonas septicemia 275-278

-, Mycobacteriosis 277, 287-289 , Mycobacterium sp. 275, 277, 286-288, 306, 309, 341 , Pasteurellosis 277, 279-280 -, Pasteurella sp 275, 277, 279-282, 306-309 -, P s e u d o m o n a s s p . 7, 275-277, 281,340-341 -, Streptococcosis 277, 284 -, Streptococcus sp. 275, 277, 284-287, 307 -,Vibrio sp. 275, 277, 281,283-284, 381 -, Vibriosis 283-284, 307 Bicarbonate (see alkalinity) 123, 162, 172-177, 203,209, 259, 263,332, 348 Biofilters 163 165, 260 Biofiltration 151,157, 163 Biotin 242 Body flukes (see Metazoan parasites) 297 Branchiomyces sp. 289, 290 Breeding programs 7-8, 41, 49, 51,219-220, 222-223, 228229 -, hybridization 39, 41, 217, 219, 224, 227, 229 Broodfish -, care of 40, 49-51 -, selection of 53 -, sex determination 46, 173 -, stress in 207, 208 -, transport of 172, 185, 189, 199, 200 "Brown-blood" disease (methemoglobinemia) 261 Buffers 162-163, 175, 177, 203,208-209, 258 Cage culture (net-pens) 2, 4, 6, 82- 85, 129, 217, 317-318 Calcium -, in nutrition 241-242, 344 -, in reproduction 32 -, in water quality 110, 131,195-199, 203-209, 258-264, 271-272 Calcium chloride 195-196, 203,259, 261,272 Calcium hydroxide 163 Calcium sulfate -, and pH 259 Captive breeding -, broodstock sources 49 -, captive broodstock 49-51, 219 -, conditioning domestic broodstock 51-53 -, domestic broodstock 45, 49, 51-53, 56 -, hormonal implants 21, 38, 41, 52, 54-59, 76 -, spawning induction 21, 53-59 Carbohydrates 239-240, 284, 288, 340 Carbon dioxide -, aeration and 100, 130, 157, 192-194 -, concentrations in ponds 100 -, effects on striped bass 131,207 -, and fish transporting 201-203 -, and pH 259 -, removal of 200, 120, 157, 189, 194 Casein 238-239

362

Catch basin 81,143 Channel catfish 129, 192, 222, 236, 240-242, 282, 307-308 Chemical oxygen demand 163, 351-352 Chilodonella sp. 7, 293 Chloride -, dietary 241,244 -, and pH 202 -, incubation 178 -, and methemoglobinemia 250 -, and salinity 257 -, ans stress 204, 207, 272 -, and transport 195-199, 258 Chlorine 130-132, 156, 264, 306, 348-352 Chromosome 12, 224, 226 Chromosome numbers (karyology) 224 Clinoptilolite 209 Conditioning fish -, for breeding 43, 49, 51, 53, 59, 76 -, for transport 186, 196-197 -, feed management 197 -, larvae 208-209 -, loading rates 200-201 -, temperature 199-200, 256 -, treatments 198-199, 203-208 Conservation genetics 229 Copper sulfate 164, 262 Cryopreservation (see Genetic manipulation) 217, 220, 227-228 Cytogenetics 217, 224-229 Design capacity 99, 121 Disease control -, chemical usage 303-308 -, chemotherapeutics 303-308 -, chlorine 262, 306 -, copper sulfate 164, 305-308, 332 -, formalin 170, 173, 178, 263, 305-309 -, masoten 305-309 -, potassium permanganate 164, 305-308 -, preventative medicine 172, 304-308 -, ozone 306-308, 352 -, Roccal1305 -, Romet 305-307 -, salt 305-307 -, terramycin 305-307 -, ultraviolet light 156, 305, 348 -, vaccination 307 Dissolved Oxygen -, equilibrium constants 104 -, Henry's Law 104 -, surface reaeration 108, 111 -, transfer rate 104, 105, 108 DNA hybridization 275, 284 Domestication (see Captive breeding) Economics -, demand 83, 91,320-322, 325, 330 -, seafood demand 320-321 -, striped bass demand 321

-, feed costs 156-319-320 -, financial performance 317, 320, 322-323 -, hatchery production 3, 4, 76, 258, 317 -, market 316, 320, 321-326, 330, 345-353 -, production costs 316-320 -, production levels 128, 157, 316 -, pond production 123, 128, 322 -, net-pen production 318 -, operating costs 135-136, 155, 318-320 -, stocking costs 319 -, tank production 318 Eels 282 Environmental influences on reproduction -, controlled reproduction 41-44, 229 -, photoperiod 8, 41-42, 51,180-181,255-260 -, photothermal cycles 41-44, 52 Environmental requirements -, alkalinity 255-260, 263, 271,305-308 -, ammonia 82, 99-100, 149, 197-203, 209-210, 259-261 -, calcium 131,195-199, 203-209, 258-263 -, dissolved oxygen 185, 201,237, 253-257 -, gas supersaturation 262 -, hardness 203,209, 258-260, 263 -, light 77, 210, 259-261 -, nitrite 83, 157, 254, 257, 261 -, pH 78, 82, 256, 259 -, salinity 258, 263 -, suspended solids 99, 131,148, 261,348 -, temperature 253-256, 260 Extensive culture , definition 127 FDA 203, 262, 306, 307, 330-331,346 Filtration systems -, types of 138-140 Fish lice (see Metazoan parasites) 297 Flow-though systems -, aeration in 152-153 -, and oxygen 149 -, design considerations 149-151 -, feeding in 155 -, harvesting 154 -, requirements of 148-149 -, tanks 152 -, types of 149-151 -, water supply 151 Fungal disease -, Achyla sp. 290 -, Aphanomyces sp. 290 -, Branchiomyces sp. 288-291 -, Saprolegnia sp. 288-291 -, Saprolegniosis 288-291 Genetic engineering (see Genetic manipulation) 228-229 Genetic manipulation -, gynogenesis 226-227 -, polyploidy 224-226 -, sex reversal 227 -, genetic engineering 228-229

363

Gill flukes (see Metazoan parasites) 297 Grubs (see Metazoan parasites) 297-300 HACCP (see processing laws & regulations) 330 Heritability 223 Heterosis 217-218 Hybrids -, backcrosses 221 -, common names of 220-221 -, performance of 220-221 -, sterility in 218, 225-227 Hybridization -, artificial 220 -, natural 220 Infectious disease -, alkalinity and 272 -, dissolved oxygen and 272 -, predisposing factors 272-273 -, stress and stressors 272 -, water quality and 272 Intensive culture -, definition 127, 140 -, disadvantages of 129 -, in flow-through systems 148-155 -, in ponds 140-148 -, in recirculation systems 155-165 -, water quality requirements 129-131 -, water supplies 130, 140 Iteroparous 11 Laws and regulations in processing -, HACCP 331 -, FDA 330-331 -, labeling 339 -, NMFS 330-331 -, processing facilities 331 Leeches (see Metazoan parasites) 293,297, 308 Lipids -, in nutrition 242-243 -, in reproduction 34 Marketing 344-345 Metazoan parasites -, Acanthocephalans 297, 301 -, Argulus sp. 302-304 -, body flukes 297 -, Cestodes 297, 299, 301 -, Clinostomum sp. 299-300 -, Crustacean parasites 297, 302-304, 308 -, Digenetic trematodes297-300 -, Diplectanum sp. 297 -, Diplostomum sp. 299-300 -, Ergasilus sp. 302-304 -, gill flukes 297 -, Goezia sp. 301-302 -, grubs 297-300 -, Gyrodactylus sp. 297 -, Helminths 297-302, 308

-, Leeches 293, 297, 308 -, Lernaea sp. 302-304 -, Microcotyl sp. 297 -, Monogenetic trematodes 297-298 -, Neascus sp.299-300 -, Philometra sp. 301-302 -, Pomphorynchus sp. 301 -, Posthodiplostomum sp. 298 -, round worms 301-302 -, Spiny head worms 297, 301 -, Urocleidus sp. 297 -, Uvulifer sp. 299 Neuroendocrine system -, brain 19 -, adrenocorticotropic hormone-secreting cells (ACTH) 23 -, catecholamines 29 -, cortico-releasing hormone 28 -, cortiso128 -, endocrine gland 25 -, hypothalamus 19 -, insulin 30 -, insulin-like growth factors (IGF-I) 30 -, metabolism 25 -, pituitary gland 19 -, steroid hormones 25 -, steroidogenesis 25 -, stress hormones 25 -, thyroid hormones 25 Noninfectious disease -, algicides 264-265 -, anesthetics 263-264 -, chlorine 265 -, corticosteroids 255 -, environmental requirements -, alkalinity 256-260 -, ammonia 261-262 -, calcium 259 -, dissolved oxygen 256-258 -, gas supersaturation 263 -, hardness 259 -, light 260-261 -, nitrite 262 -, pH 256-260 -, salinity 258 -, suspended solids 262 -, temperature 256 -, formalin 265 -, immunosuppression 255 -, stress, the role of 254-255 -, toxicants -, algicides 264-265 -, anesthetics 263-264 -, chlorine 265 -, formalin 265 Nutrition -, carbohydrate requirements 236- 248 -, energy requirements 237-238, 254 -, feeds and feeding 236, 244-248

364

-, in disease control 306 -, lipid requirements 241-242 -. macro mineral requirements 242-243 -, micro mineral requirements 243 -, protein requirements 238-240 -, role of nutrition 237 -, vitamin requirements 243-245 Oxygen production 102 pH -, dynamics in ponds 109-111 -, relationship to alkalinity 110-111 Phase II production -, auger pumps 87, 89-90 -, feeds 80, 82-83, 87 -, feeding strategies 74, 79 -, fish pumps 91 -, harvesting phase II fish 76, 80-81, 87 -, net-pens 82 -, phase I1 fingerlings 78, 82 -, phytoplankton 77, 80, 82 -, pond preparation 77, 82 -, stocking densities 83 -, stocking for phase II grow-out 74-79, 82-83, 87 -, supplemental aeration 83 Phase III production -, boom nets 93-94, 145 -, compensatory growth 93 -, feeding strategies 91 -, grading 80, 94 -, harvesting phase III fish 91, 93-94 -, haul seines 87-88, 93 -, live cars 93 -, markets 93 -, marketing 91 -, predators 91 -, production ponds 74, 79-80, 86, 88, 91 -, stocking 91 -, supplemental aeration 83, 92 Phosphorous -, in nutrition 243 -, in reproduction 33 -, in water quality 82 Polyploidy 225-227 Pond culture -, feeding in 146-148 -, harvests 144-146 -, types of ponds 140-144 Pond production -, catch basin 81 -, cladocerans 78 -, copepods 78 -, cotton seed meal 78 -, exogenous feeding 76, 78 -, fry 76-78 -, harvest kettle 81 -, harvesting phase I fish 80 -, harvesting phase II fish 87 -, harvesting phase III fish 93

-, hatchery 74-81 -, heterosis 74 -, inorganic fertilizers 78 -, larval shipping 78-79, 93-94 -, light sensitivity 78 -, Maryland bass 74 -, organic fertilizers 78 -, planktivorous 76, 79 -, plankton dynamics 77, 80, 82 -, pond management 77 -, population enhancement 74, 79, 82-83 -, prolarvae 76-78 -, rice bran 78 -, rotifers 78 -, schooling response 80 -, seining ponds 81, 87 -, stocking phase II fish 87 -, striped bass versus hybrids 74 -, supplemental feeding 76, 79 -, sunlight 78 -, sunshine bass 74 -, swimbladder inflation 79 -, zooplankton 74-78, 80 Pond production systems -, nitrogen input -, as a controlling factor 100-101 -, for fertilization 100-101 -, phosphorous inputs 101 -, primary pond production 101 Preservation methods -, chill packing 336 -, freezing 336 -, ice-packing and refrigeration 335-336 -, shelf life 339-342 -, smoking 336-337 -, freezing 336 Processing -, cleaning and sanitizing 345-351 -, employee considerations 350 -, equipment 349-350 -, pest management 350-351 -, principles of 347-349 -, filleting 333 -, handling and bleeding 333 -, harvesting and transport 331-332 -, off-flavor 332 -, meat and bone separation 333 -, packaging 337-339 -, bulk packaging 338 -, coding 339 -, labeling 339 -, receipt offish at the processing plant 332 -, waste handling 351-353 -, liquid disposal 352 -, solid disposal 352 -. solid utilization 352-353 -, yields 333,335-336 Processing and food safety -, chemical considerations 341 -, producer classification 330

365

-, -, -, -,

striped bass markets 330 market grades 330 microbial growth during storage 341-342 microbiological considerations 338-341 -, Aerornonas sp. 341 -, Bacillus sp. 341 -, Clostridiurn botulinurn 338, 341 -, Cytophaga sp. 341 -, Enterobacteriaceae 341 -, Escherichia coil 341 -, Flavobacteriurn sp. 341 -, Listeria rnonocytogenes 341 -, Mycobacteriurn rnarinum 341 -, Plesiornonas shigelloides 341 -, Pseudornonas sp. 340-341 -, Salmonella sp. 341 -, Shigella dysenteriae 341 -, Sphingobacteriurn sp. 341 -, Staphylococcus aureus 341 -, Vibrio sp. 341 -, Yersinia pseudotubercuIosis 341

-, product form 330 Protozoan diseases -, Arnbiphrya sp. 293 -, Amyloodiniasis 292-293 -, Arnyloodiniurn sp. 292-293,307 -, Apisorna sp. 293 -, Chilodonella sp. 293-294 -, ciliates 291-297 -, Cryptocaryon sp. 293-294, 307 -, Epistylis sp. 293-295 -, flagellates 241-293 -, Ichthyobodiasis 291-292 -, Ichthyobodo sp. 291-293 -, Ichthyophthirias 293-296 -, Ichthyophthirius sp. 293-297, 307-308 -, Trichodina sp. 289, 293-295 Puberty -, assessing maturity 46 -, gonadal biopsy 47 -, maturity schedules 48 -, ultrasonic imaging 48 Recirculation systems -, advantages of 156 -, biofilter management 162-164 -, biofilter hydraulic loading rate 162 -, biofilter hydraulic loading residence time 163 -, biofilter oxygen requirements 162 -, biofiltration 157-164 -, design considerations 121-123, 156-157, 164 -, effects of particulates on 164 -, oxygen consumption 99-100 -, nitrification in 157, 163-164 -, requirements of 152-156 -, water quality requirements 163 -, water supply and needs 153-155 Redfield ratio 100-101 Reproduction -, atretic or atresia 12, 15

-, chorion (zona radiata) 11, 13 -, egg 12 -, egg and sperm activation 30 -, embryogenesis 30 -, feedback regulation 31 -, fertilization 31 -, final oocyte maturation 12 -, gametogenesis 24 -, germ cells 12, 17, 19 -, germinal vesicle 12, 14 -, gonads 10 -, Leydig cells 19 -, meiosis 12 -, micropyle 30 -, mitosis 12 -, oocyte 11 - 14 -, oocyte maturation 12 -, ooplasm 12-15 -, ovary 11-16 -, polar body 12, 41,224, 226 -, polyspermy 30 -, primary growth oocyte 12 -, recrudescence 19 -, reproductive cycle 31 -, resorption 36 -, secondary growth oocyte 12, 15 Sertoli cells 17, 19 -, spawning 24 -, spawning behavior 38-39 -, sperm 19, 28, 36, 46, 51, 57-58, 177, 220-228 -, spermatogenesis 17, 19 -, spermatogonia 17, 19 -, spermatatids 17, 19 -, spermatozoa 17, 19 -, spermiation 19 -, spermiogenesis 19 -, testicular maturation 17 -, testis (es) 17, 19 -, vitellogenic (in, sis) 12, 14, 15 -, "water hardening" 41 -, yolk 12, 15 Reproductive hormones -, 11-ketotestosterone 27 -, 17t~-2013-dihydroxy-4-pregnen-3-one (DHP) 27 -, 170t-2013-2l-trihydroxy-4-pregnen-3-one (20-13) 27 -, androgens 25 -, chorionic gonadotropin (CG) 25 -, dopamine 29 -, domperidone (DOM) 21 -, estradiol-1713 (E2) 27 -, estrogen 26 -, estrone 32 -, feedback regulation 32 -, follicle stimulating hormone (FSH) 23 -, gonadotropin (GTH) 19 -, gonadotropin-releasing hormone (GnRH) 19 -, growth hormone (GH) 22-23 -, luteinizing hormone (LH) 23 -, maturation inducing hormone (MIH) 27 -, maturation promoting factor (MPF) 28 -

366

-, melanocyte-stimulating hormone (MSH) 33 , pimozide 21 -, progesterone 27 -, progestins 25 -, prolactin (PRL) 24 -, somatolactin (SL) 22 -, testosterone 27 -, thyroid stimulation hormone (TSH) 33 -, thyroxine (T4) 30 -, triiodo-thyronine (T~) 30 Round worms (see Metazoan parasites) 301-302 Solids in production -, problems with 135, 138-140 -, source of 135 -, types of 138 Spiny head worms (see Metazoan parasites) 297, 301 Strain evaluation 223-224 Stress -, handling and transport 186-187 -, mitigating stress 187, 197-201,203-209 -, stress response 186-187, 253-255 -, types of stressors 186-187, 253-255 Sudden death syndrome 206-207 Swimbladder inflation 181 Tapeworms (see Metazoan parasites) 297, 299, 301 Toxicants -, algicides 264-265 -, anesthetics 263-264 -, chlorine 265 -, formalin 265 Transportation equipment -, oxygenation and aeration equipment 193-196 -, transfer equipment 188-193 Viral disease -, Aquareovirus 275-276 -, Aquarotavirus 276 -, Cystivirus 275

-, Lymphocystis 273-275 -, Infectious pancreatic necrosis 275 Water quality -, during fish transport 202-211 -, ammonia 204, 210-211 -, anesthetics 208, 262-263 -, carbon dioxide 202-203, 210 -, dissolved oxygen 202 -, pH 204-210 -, salts 204-208, 210 -, temperature 203-204 White bass -, biology and ecology 170-172 ,, broodstock collection 171 -, broodstock handling 171, 181-182 -, conditioned spawning 178-180 -, egg incubation 178 -, egg size 171

-, feeding broodstock 173-175 -, hatching rate 178-180 , hormonal spawning 174-178 -, grow-out to market 181 -, intensive culture 181 -, morphology 170 -, natural food habits 171 -, natural spawning 171-172 -, removal of egg adhesive matrix 180 -, sexing 174 -, swimbladder inflation 181 White perch 1, 2, 11, 17, 19, 28, 31-31, 36, 41, 46, 74, 78, 219-220, 226, 229

Principles of Salmonid Culture (Developments in Aquaculture and Fisheries Science)

Design and Operating Guide for Aquaculture Seawater Systems (Developments in Aquaculture and Fisheries Science) (Developments in Aquaculture and Fisheries Science)

Striped Bass and Other Morone Culture, Volume 30 (Developments in Aquaculture and Fisheries Science)

Principles of Salmonid Culture (Developments in Aquaculture and Fisheries Science)

Principles of Salmonid Culture (Developments in Aquaculture and Fisheries Science)

Design and Operating Guide for Aquaculture Seawater Systems (Developments in Aquaculture and Fisheries Science) (Developments in Aquaculture and Fisheries Science)

Principles of Salmonid Culture (Developments in Aquaculture and Fisheries Science)

Biology of the Hard Clam (Developments in Aquaculture and Fisheries Science) (Developments in Aquaculture and Fisheries Science)

Biology and Culture of Channel Catfish, Volume 34 (Developments in Aquaculture and Fisheries Science)

The Knowledge Base for Fisheries Management, Volume 36 (Developments in Aquaculture and Fisheries Science)

Biotechnology and Genetics in Fisheries and Aquaculture

Biotechnology and Genetics in Fisheries and Aquaculture

Biology of the Hard Clam, Volume 31 (Developments in Aquaculture and Fisheries Science)

Private standards and certifications in fisheries and aquaculture

State of World Fisheries and Aquaculture, 2010

Lobsters Biology Management Aquaculture and Fisheries

Aquaculture and fisheries biotechnology. Genetic approaches

Aquaculture and Fisheries Biotechnology, CABI Publishing

Aquaculture: Biology And Ecology Of Cultured Species (Ellis Horwood Series in Aquaculture and Fisheries Support)

Aquaculture Science

Aquaculture and Fisheries Biotechnology: Genetic Approaches Animals Pets

Striped

Stability of Motion, (Mathematics in Science and Engineering, Volume 30)

Striped

Striped

Developments in Block Copolymer Science and Technology

Selective Breeding in Aquaculture: an Introduction (Reviews: Methods and Technologies in Fish Biology and Fisheries, 10)

Science Education Issues and Developments

Ecological and Genetic Implications of Aquaculture Activities (Reviews: Methods and Technologies in Fish Biology and Fisheries)

Striped

Striped

Striped

Managing Sea Cucumber Fisheries With an Ecosystem Approach (Fao Fisheries and Aquaculture Technical Paper)

Striped Bass and Other Morone Culture, Volume 30 (Developments in Aquaculture and Fisheries Science)

Principles of Salmonid Culture (Developments in Aquaculture and Fisheries Science)

Principles of Salmonid Culture (Developments in Aquaculture and Fisheries Science)

Design and Operating Guide for Aquaculture Seawater Systems (Developments in Aquaculture and Fisheries Science) (Developments in Aquaculture and Fisheries Science)

Principles of Salmonid Culture (Developments in Aquaculture and Fisheries Science)

Biology of the Hard Clam (Developments in Aquaculture and Fisheries Science) (Developments in Aquaculture and Fisheries Science)

Biology and Culture of Channel Catfish, Volume 34 (Developments in Aquaculture and Fisheries Science)

The Knowledge Base for Fisheries Management, Volume 36 (Developments in Aquaculture and Fisheries Science)

Biotechnology and Genetics in Fisheries and Aquaculture

Biotechnology and Genetics in Fisheries and Aquaculture

Biology of the Hard Clam, Volume 31 (Developments in Aquaculture and Fisheries Science)

Private standards and certifications in fisheries and aquaculture

State of World Fisheries and Aquaculture, 2010

Lobsters Biology Management Aquaculture and Fisheries

Aquaculture and fisheries biotechnology. Genetic approaches

Aquaculture and Fisheries Biotechnology, CABI Publishing

Aquaculture: Biology And Ecology Of Cultured Species (Ellis Horwood Series in Aquaculture and Fisheries Support)

Aquaculture Science

Aquaculture and Fisheries Biotechnology: Genetic Approaches Animals Pets

Striped

Stability of Motion, (Mathematics in Science and Engineering, Volume 30)

Striped

Striped

Developments in Block Copolymer Science and Technology

Selective Breeding in Aquaculture: an Introduction (Reviews: Methods and Technologies in Fish Biology and Fisheries, 10)

Science Education Issues and Developments

Ecological and Genetic Implications of Aquaculture Activities (Reviews: Methods and Technologies in Fish Biology and Fisheries)

Striped

Striped

Striped

Managing Sea Cucumber Fisheries With an Ecosystem Approach (Fao Fisheries and Aquaculture Technical Paper)

Recommend Documents