Health Maintenance and Principal Microbial Diseases of Cultured Fishes [3 ed.] 0813816939, 9780813816937, 9780470958407

Table of contents :
Health Maintenance and Principal Microbial Diseases
of Cultured Fishes......Page 5
Contents......Page 7
About the Authors......Page 9
Preface......Page 11
Acknowledgments......Page 13
Part I: Health Maintenance......Page 15
1 Principles of Health Maintenance......Page 17
2 Epizootiology of Fish Diseases......Page 45
3 Pathology and Disease Diagnosis......Page 53
4 Disease Management......Page 71
Part II: Viral Diseases......Page 105
5 Catfish Viruses......Page 109
6 Carp and Minnow Viruses......Page 123
7 Eel Viruses......Page 149
8 Trout and Salmon Viruses......Page 161
9 Sturgeon Viruses......Page 233
10 Other Viral Diseases of Fish......Page 241
Part III: Bacterial Diseases......Page 287
11 Catfish Bacterial Diseases......Page 289
12 Carp and Minnow Bacterial Diseases......Page 329
13 Eel Bacterial Diseases......Page 341
14 Salmonid Bacterial Diseases......Page 359
15 Striped Bass Bacterial Diseases......Page 433
16 Tilapia Bacterial Diseases......Page 459
17 Other Bacterial Diseases......Page 479
Part IV: Appendices......Page 485
Appendix I. List of Common and Scientific Names of Fishes......Page 487
Appendix II. Table of Conversion Factors......Page 491
Appendix III. List of Cell Lines Commonly Used for Diagnostics......Page 493
Index......Page 497

Citation preview

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Health Maintenance and Principal Microbial Diseases of Cultured Fishes Third Edition

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Health Maintenance and Principal Microbial Diseases of Cultured Fishes Third Edition John A. Plumb and Larry A. Hanson

A John Wiley & Sons, Ltd., Publication

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Edition first published 2011
C 2011 Blackwell Publishing Ltd. Originally published as Health Maintenance of Cultured Fishes: Principal Microbial Diseases, CRC Press, Boca Raton, Florida 1994; the second edition as Health Maintenance and Principal Microbial Diseases of Cultured Fishes, Iowa State University Press, Ames, Iowa 1999. Blackwell Publishing was acquired by John Wiley & Sons in February 2007. Blackwell’s publishing program has been merged with Wiley’s global Scientific, Technical, and Medical business to form Wiley-Blackwell. Editorial Office 2121 State Avenue, Ames, Iowa 50014-8300, USA For details of our global editorial offices, for customer services, and for information about how to apply for permission to reuse the copyright material in this book, please see our Website at www.wiley.com/wiley-blackwell. Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Blackwell Publishing, provided that the base fee is paid directly to the Copyright Clearance Center, 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license by CCC, a separate system of payments has been arranged. The fee code for users of the Transactional Reporting Service is ISBN-13: 978-0-8138-1693-7/2011. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. Library of Congress Cataloging-in-Publication Data Plumb, John A. Health maintenance and principal : microbial diseases of cultured fishes / John A. Plumb and Larry A. Hanson. – 3rd ed. p. cm. Rev. ed. of: Health Maintenance of cultured fishes, originally published in 1994. Includes bibliographical references and index. ISBN 978-0-8138-1693-7 (hardback : alk. paper) 1. Fishes–Infections. 2. Fish-culture. I. Hanson, Larry A. II. Plumb, John A. Health Maintenance of cultured fishes. III. Title. SH171.P66 2011 639.3–dc22 2010020441 A catalog record for this book is available from the U.S. Library of Congress. R Inc., New Delhi, India Set in 10/12.5 pt Sabon by Aptara
Printed in Singapore

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2011

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Contents About the Authors Preface Acknowledgments

vii ix xi

Part I: Health Maintenance 1

Principles of Health Maintenance

3

2

Epizootiology of Fish Diseases

31

3

Pathology and Disease Diagnosis

39

4

Disease Management

57

Part II: Viral Diseases 5

Catfish Viruses

6

Carp and Minnow Viruses

109

7

Eel Viruses

135

8

Trout and Salmon Viruses

147

9

Sturgeon Viruses

219

Other Viral Diseases of Fish

227

10

95

Part III: Bacterial Diseases 11

Catfish Bacterial Diseases

275

12

Carp and Minnow Bacterial Diseases

315

13

Eel Bacterial Diseases

327

14

Salmonid Bacterial Diseases

345

15

Striped Bass Bacterial Diseases

419

16

Tilapia Bacterial Diseases

445

17

Other Bacterial Diseases

465

Part IV: Appendices Appendix I. List of Common and Scientific Names of Fishes Appendix II. Table of Conversion Factors Appendix III. List of Cell Lines Commonly Used for Diagnostics Index

473 477 479 483

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About the Authors John A. Plumb, Professor Emeritus, Department of Fisheries and Allied Aquacultures, Auburn University, Alabama, taught graduate courses in microbial disease and disease diagnosis of fish. His research included investigations of viral and bacterial diseases of fish and the effects of environmental stress on disease susceptibility. Plumb is a past president of the Fish Health Section of the American Fisheries Society. Widely published, he led the Southeastern Cooperative Fish Disease Project and has advised international fishery agencies.

Larry A. Hanson, Professor in the Department of Basic Sciences, College of Veterinary Medicine, Mississippi State University, Mississippi, teaches fish virology. His research includes molecular virology and application of molecular biology to investigate fish health problems associated with aquaculture. Activities include fish diagnostician and fish virology; he is an AFS/FHS Certified Fish Pathologist, and OIE reference expert for channel catfish virus, and enteric septicemia of catfish.

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Preface Infectious diseases of cultured fish pose significant constraints to expansion and realization of aquaculture’s full potential. Viral, bacterial, and parasitic agents infect many wild and all cultured fish species. Most pathogenic agents are endemic to natural waters where, under normal conditions, they cause no great problem. However, when these same diseases occur in an aquacultural environment they may cause significant disease and mortality. Cultured fish are often confined to an environment to which they are not biologically accustomed, a circumstance that often increases susceptibility to infectious disease. It is virtually impossible to separate the relationship of infectious disease from problems associated with environmental quality. The objective of Health Maintenance and Principal Microbial Diseases of Cultured Fishes is to emphasize salient points of host–pathogen–environment relationships, elucidate important aspects of infectious diseases, and explore how management can be used to prevent and reduce their effects on aquaculture. The revision was undertaken to update the diseases in earlier editions as well as to include those diseases that were previously poorly understood or have more recently emerged. Chemotherapy and vaccination sections have been updated as well as procedures and surveillance and detection of infectious agents in pathogen carrier populations. It is emphasized that isolation and detection of pathogens is only part of the infectious disease picture of fish. However, detail molecular methods of pathogen detection and identification are beyond the scope of this work.

The text is divided into three parts: Part I emphasizes the principles of fish health maintenance, recognition, diagnosis, and control of infectious fish diseases. Parts II and III concentrate on viral and bacterial diseases, respectively, that are important to aquaculture and wild fish populations where applicable. Emphasis is placed on geographical distribution, species susceptibility, clinical signs, etiological agents, their descriptions and methods of detection, epizootiology, pathology, and significance of a specific disease. We have tried to create a balance between diseases of warmwater, coolwater, and coldwater fishes. Although much of the information has been derived from North America, important disease problems from other parts of the world are included. Diseases are organized into fish groups or families that are most extensively cultured. Where a disease affects members of more than one fish family emphasis is placed on the family most commonly or severely affected. Although some viral and bacterial diseases occur in both marine and freshwater fish, no specific distinction is made between the two environments. It is not our intent to list every reported disease or all published papers on each disease or subject matter mentioned. Only those publications that we feel are pertinent to the discussion have been cited. This book is intended for students; scientists interested in health maintenance of fish and their pathobiology, and infectious fish diseases; as well as aquaculturists, fishery managers, fishery biologists, fish pathologists, and aquatic veterinarians.

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Acknowledgments The authors wish to thank the following individuals who reviewed portions of this book and/or provided valuable information: John Grizzle, John Hawke, Paul Bowser, Andy Goodwin, Rocco Cipriano, and Rosalee Schnick. A special thank you is extended to all who provided valuable photographs: W. Ahne, S. Bastien-Daigle, P. Bowser, R. Bootsma, G.

Camenisch, M. Chen, P. de Kinklin, D. Earlix, J. Ferguson, N. Fijan, P. Ghitino, J. Grizzle, J. Hawke, R. Hedrick, B. Hjeltness, T. Jones, S. LaPatra, S. Leek, T. Miyazaki, E. Morrison, B. Nicholson, M. Okihiro, D. Powell, J. Rohovec, T. Sano, E. Shotts, P. Williams, J. Winton and M. Yoshimizu.

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Part I

Health Maintenance

Fish health maintenance emphasizes many areas that affect the health of cultured fishes. It requires continuous efforts, which include the following: the location and construction of a culture facility; selection and introduction of culture species; and reproduction, culture, and harvesting of the final product. The aquatic habitat—a dynamic and continuously changing environment—is affected by structural material, facility design, soil quality and type, volume, and quality of water, fish species present, amount and quality of nutrients introduced into the system, climate, and daily human activities. Health maintenance involves a series of principles that apply to most farm-raised animals. However, fish tend to react more quickly to environmental change than terrestrial farm animals. Because of their homeothermic nature, most terrestrial farm animals respond comparatively slowly to unfavorable environmental conditions, whereas fish—being poikilothermic—respond quickly and often fatally to handling, temperature change, excessive or insufficient dissolved gasses in the water, metabolites, or chemical additives, and so forth, to which they are un-

able to adapt. These factors also increase fish susceptibility to infectious agents and compromise their immune response. Specific areas of concern addressed in this book include principles of health or health maintenance, epizootiology and pathology of fish diseases, disease recognition, basic concepts in disease diagnosis, and prevention and control of infectious fish diseases. Aquatic animal health management encompasses the entire production process, including disease diagnosis and treatment. The objective of health maintenance is to help control environmental fluctuations through management practices, thus reducing the magnitude of change and producing a more economical, healthier, and better quality product. The ultimate goals of health management are (1) disease prevention, (2) reduction of infectious disease incidence, and (3) reduction of disease severity when it occurs. Successful health maintenance and disease prevention or control do not depend on any single procedure but are the culmination of the application of integrated concepts and exercising management options.

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Chapter 1

Principles of health maintenance

“An ounce of prevention is worth a pound of cure” is a familiar phrase that describes one approach to the culture of food animal resources. Health maintenance is a concept in which animals are reared under conditions that optimize the growth rate, feed conversion efficiency, reproduction, and survival while minimizing problems related to infectious, nutritional, and environmental diseases, all within an economical context. “Health maintenance” encompasses the entire production management plan for food animals, whether they are swine, cattle, poultry, or fish. Aquaculture involves man’s intervention in the growth process of fish and other organisms in an aquatic environment. The degree of intervention is progressive, ranging from extensive (few fish per unit of water volume) to increasingly intensive (comparatively greater numbers/weight of fish per unit of water volume) in ponds, raceways, cages, and recirculating systems where higher fish densities are maintained. As culture becomes more intensive, need for intervention increases accordingly, and principles of health maintenance become of greater importance. These principles apply to aquaculture around the world, regardless of fish species, culture method, or climate. Health Maintenance and Principal Microbial Diseases of Cultured Fishes, 3rd edition. By J. A. Plumb and L. A. Hanson. Published 2011 by Blackwell Publishing Ltd.

Fish health management is not a new approach to aquaculture. Snieszko (1958) recognized the need for health maintenance in fish culture when he stated, “We are beginning to realize that among animals (including fish) there are populations, strains, or individuals that are not susceptible all of the time, or even temporarily, to some of the infectious diseases.” He theorized that fish possess a certain level of natural resistance to infectious diseases that can be enhanced through proper management, and that environmental stressors and/or fish cultural practices can adversely affect that natural resistance. Another contributor to a health maintenance concept for aquatic animals is Klontz (1973), who established a fish health management course at Texas A & M University that combined fish culture and infectious diseases into health management. The Great Lakes Fishery Commission published a Guide to Integrated Fish Health Management in the Great Lakes Basin, which was a regional concept for fish health management (Meyer et al. 1983). These references deal with the improvement of aquatic animal health through management. The most in-depth contribution to maintaining health of domestic (cultured) animals was Schnurrenberger and Sharman (1983), who set forth a series of principles for animal health maintenance, which apply in a general sense to all domesticated food animals. 3

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Health Maintenance

In the following pages these principles are applied to aquaculture. Theoretically, if these principles are utilized in daily, monthly, yearly, and long-term management of an aquatic culture facility, there will be fewer environmental and disease problems and optimum production will be more readily obtained. Biosecurity is the term recently applied to fish health management (Bebak-Williams et al. 2007) in which biosecurity is aimed at reducing the risk of pathogens being introduced to a facility, reducing the risk of pathogens being spread throughout the facility, and alleviating conditions that increase susceptibility to infections. It is emphasized that biosecurity cannot completely prevent entry of or eliminate all pathogens from the culture facility but emphasizes reduction of pathogens rather than their complete elimination. Biosecurity begins with selection of the aquaculture site and continues throughout production with complete control of water and human access.

sizes interruption of a disease cycle, deals with multiple segments of health maintenance, and results in more efficient production. Health maintenance does not simply target infectious diseases, but emphasizes proper utilization of physical facilities; use of genetically improved fish and certified “specific pathogen free” (SPF) stocks whenever available and/or feasible; environmental control; prophylactic therapy; feed quality and quantity, pond, cage, raceway, tank, or recirculating system management; control of vegetation; aeration and use of other water quality maintenance practices; and a management commitment to provide an optimum habitat in terms of water quality for fish being cultured. Its goal is to improve the health and well-being of animals that appear to be generally healthy. If sound health maintenance principles are followed, production will be more efficient and result in a healthier product. Obviously, all activities, policies, and improvements must be based on sound economic criteria.

Health maintenance Stress In an aquatic environment, there is a profound and inverse relationship between environmental quality and disease status of fish. As environmental conditions deteriorate, severity of infectious diseases increases; therefore, sound health maintenance practices can play a major role in maintaining a suitable environment where healthy fish can be grown. The aquatic environment is a dynamic ecosystem that changes over a 24-hour period and seasonally, particularly in ponds with limited water exchange. Tucker and Van der Pflog (1993) noted that in static catfish ponds, periods of poorest water quality occurred during summer months when feeding, temperature, and standing crops were at a maximum, but rainfall and available water were at a minimum, thus producing a higher potential for stressful conditions requiring health management. Fish health management is a positive concept that aids in disease prevention, empha-

“Stress” is difficult to define because it is used to describe many adverse situations that affect the well being of individuals, but generally it is the reaction of an animal to a physical, physiological, or chemical insult (Barton 1997). Stress may also produce a nonspecific response to factors that are perceived as harmful; however, stress in fish is usually related to handling, transport, environmental quality, or fright. For clarification in this text, “stressors” are factors that cause a “stress response,” which is the sum of physiological changes that occur as fish react to physical, chemical, or biological stressors as the fish attempt to compensate for changes that result from these stressors (Wedemeyer 1996). The corticosteroid level in plasma is the usual quantitative measure for stress; however, amounts of glucose, lactic acid, and ions will also increase during stressful conditions (McDonald and Milligan 1997).

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Principles of Health Maintenance

The aquatic environment is in a continuous state of flux, and because fish are poikilotherms and body functions are controlled by temperature, oxygen concentration, and many other water quality parameters, they must continually adapt physiologically to environmental changes. An inability to adjust to these changes may be manifested in lower productivity, reduced weight gain, poor feed conversion, decreased immunity, reduced natural disease resistance, increase in infectious disease, lowered hardiness in general, death, reduced profits for the commercial fish farmer, and reduced production. Some commonly known stressors in the aquatic environment are unionized ammonia, nitrite, chronic exposure to low concentrations of pesticides or heavy metals, insufficient oxygen, high concentrations of carbon dioxide (CO2 ), rapidly changing or extremes in pH or water temperature, external salinities, nutrition, and fish density (Barton 1997). Low alkalinity and hardness are also not conducive to good fish health or performance (Boyd 1990). Many of these factors are exacerbated by type, quality, and quantity of feed put in a pond, and by waste accumulation. Sensitivity to these conditions will vary with fish species. Suc-

cessful and efficient health maintenance programs for aquaculture facilities will include measures to reduce and modify stressful conditions that may be present in a fish population.

Hazard reduction by management Experience has shown that a wide variety of viral, bacterial, parasitic, and other fish diseases will cause mortality if cultured fish are held in unfavorable environmental conditions (Wedemeyer 1996). Health and environmental management decisions are not independent and a change in one area should not be made without evaluating its effect in other areas. Notable stressor-related fish diseases that result from a culmination of management and biological factors are furunculosis, enteric redmouth, motile Aeromonas septicemia, columnaris, vibriosis, bacterial gill disease, streptococcus, external fungal infections, and some protozoan parasites (Table 1.1). Stress on fish increases when environmental conditions approach the host’s limit of tolerance (Snieszko 1973). For example, if water temperature is critically high and oxygen

Table 1.1 Microbial diseases of fish commonly considered stress mediated. Disease

Predisposing Environmental Factor

Spring viremia of carp Bacterial gill disease

Handling after over wintering Crowding, poor water quality, elevated presence of causative bacteria Crowding, poor water quality, handling, seining, adverse temperature, physical injury Temperature decrease from >10◦ C to 250,000 fish annual production; (3) significantly higher BGD outbreaks in the hatchery house with presence of fish in the water supply; (4) significantly more outbreaks associated with previous exposure; and

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Salmonid Bacterial Diseases 373

(5) significantly higher outbreaks if production was >500,000 lb/year. It has been suggested that BGD may be incorrectly named because its development is closely related to conditions in the culture environment and injuries to gill epithelium. It has been widely accepted that gills are usually injured by chemical or physical irritants in the water prior to bacterial colonization. Gill injuries will occur if water exchange is inadequate, ammonia levels are elevated, oxygen concentrations are decreased or if water contains silt or excess feed. These injuries will result in epithelial hyperplasia, which makes gills more susceptible to microbial invasion by Flavobacterium and occasionally other bacteria. Studies by Ferguson et al. (1991) and Lumsden et al. (1994) showed that these injuries are not absolutely necessary for BGD to occur, and Speare et al. (1991a) presented a noninjurious scenario for BGD development. After an extensive study of 23 separate BGD outbreaks in rainbow trout, they concluded that “no other disease conditions, no gross errors in management or recent exposure to chemotherapeutics” preceded BGD. Although the causative organism was not definitively identified, it was thought to have been F. branchiophilum. During a 5-month monitoring regime prior to onset of a natural disease outbreak, gill morphology of examined fish remained unaltered and it was proposed that F. branchiophilum could cause BGD without environmental stress or gill injury. Furthermore, MacPhee et al. (1995b) showed that BGD of trout is linked to the consumption of feed by the fish, rather than to environmental changes arising from feeding or water quality. They also suggested that alterations in the undisturbed layer on the gill may aid bacterial colonization, although this is secondary to feed consumption and waste excretion. Nevertheless, most BGD outbreaks are associated with some management factor such as excessive feeding, poor feed quality, poor water quality, poor water circulation, or in-

adequate water flow and association with a history of BGD is significant. Wakabayashi and Iwado (1985) reported that fish infected with F. branchiophilum were more susceptible to hypoxia because the bacterium impaired respiratory functions. Noninfected fish consumed 251–289 mL of O2 /kg/hour while oxygen consumption rates for infected fish at 2 and 5 days postinfection were 183–229 and 155–167 mL O2 /kg/hour, respectively. In comparing virulence of seven wild strain isolates and two ATCC strains of F. branchiophilum, Ostland et al. (1995) found that one strain was pathogenic to five species of salmonids and one species of shiner, but that some isolates were avirulent. All isolates that colonize the gills have fimbria, but some are unable to produce pathology. Historically, salmonid fry and fingerlings less than 5 cm in length are particularly susceptible to BGD, but larger fish can occasionally become infected. Although BGD is usually associated with juvenile fish, adult rainbow and cutthroat trout and chinook salmon suffer BGD outbreaks. Ferguson et al. (1991) successfully infected rainbow trout up to 3 years of age with F. branchiophilum. In view of these reports BGD may be more of a problem in larger trout than previously realized. BGD mortality among small fish has the potential to become subacute. In experimental BGD infection studies, mortality reached 39–80% after 13 days (Bullock 1972). Speare et al. (1991b) found that morbidity could increase from approximately 5% to over 80% within 24–48 hours when disease is first detected. During this time, mortality rates rose to 20% per day, diminished by day 7 and only a few fish showed any clinical signs by day 10–14. Once juvenile trout have overcome an F. branchiophilum infection, disease may reoccur, particularly if environmental conditions are favorable. BGD generally occurs at 12–19◦ C (Heo et al. 1990) but ABGD occurred at 90% to 10◦ C. Protection against R. salmoninarum lasted for about 3 months at 10◦ C and 2 months at 14◦ C before BKD mortality reoccurred. Experiments to treat R. salmoninarum with enrofloxacin (fluoroquinolone) by Hsu et al. (1994) showed that feeding the drug at 1.25–2.5 mg/kg of body weight per day for 10 days reduced mortality, and when fed up to 20 mg/kg/day mortality was further reduced but palatability problems occurred with 100 mg/kg of fish. Overall, chemotherapy of clinically infected BKD fish is not overly successful because it does not totally eliminate R. salmoninarum from all treated fish and relapses can be expected following medication (Austin 1985). Among drugs experimentally used for P. salmonis, oral application of the quinolones, oxolinic acid, and flumequine appear to be

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398 Bacterial Diseases

the most effective (Almendras and Fuentealba 1997). However, losses have gradually increased, which is in part due to antibiotic resistance of the pathogen. Injection of enrofloxacin has shown some efficacy.

Vaccination Vaccination of salmonids has made major inroads in controlling and preventing several diseases of these fish (Table 4.4–4.5). However, all salmonid diseases do not respond favorably to vaccination. In a treatise on vaccinations in Europe, Press and Lillehaug (1995) pointed out that application of molecular technology to fish vaccines has produced products that include purified virulence factors that have increased protection against certain pathogens and are being used in situations where large numbers of fish are involved. Commercial vaccines now available for A. salmonicida, V. anguillarum, V. salmonicida, Yersinia ruckeri, P. salmonis and Renibacterium salmoninarum (Table 4.4) are being used extensively throughout the world where salmonids are cultured with excellent cost:benefit ratios. Research is also being carried out to develop vaccines for other bacterial diseases of salmonids. The highly successful sea-cage salmonid culture in Europe, the United Kingdom, the United States, and other locations can in large part be credited to vaccination. Generally, injectable furunculosis vaccines are most effective and their use has become cost beneficial through the use of more efficient application methods. Rodgers (1990) used a vaccine composed of whole cells and extracellular products to significantly enhance protection from a natural challenge of A. salmonicida in juvenile rainbow trout. Mortality of vaccinated fish was about 11% compared to 37% for unvaccinated controls. Vaccinated fish also grew more rapidly than did nonvaccinates. Paterson et al. (1992), in laboratory and field trials, used a pelletized diet containing a dried, coated A. salmonicida culture preparation to

orally vaccinate Atlantic salmon against furunculosis. Press et al. (1996) compared IP injection, immersion, and oral application of monovalent and trivalent vaccines and found the trivalent preparation to be the only one that lead to high levels of specific antibody and possible immunological enhancement. The efficacy of a commercially available vaccine for furunculosis was evaluated in two strains of Arctic char (Bebak-Williams (2002). The vaccine (Aqua Health Furogen 2), administered via injection, provided significant protection at 87 and 108 days postvaccination in both strains of char. When Atlantic salmon were vaccinated by injection for furunculosis, they experienced a temporary immunosupression, which resulted in subclinical carrier fish (Inglis et al. 1992). The researchers found that feeding amoxycillin following vaccination improved survival to over 80% compared to 0% survival for nonmedicated fish. As a result of vaccination followed by drug treatment, a relative percent survival of 86% was achieved in these fish when challenged 4 months postvaccination. Vaccination against vibriosis, particularly V. anguillarum and V. ordalii, has become an accepted practice for cultured salmon. Studies have also demonstrated its positive effect against vibriosis in other species as well; ayu, eels, milkfish, and striped bass (Kawano et al. 1984; Tiecco et al. 1988; Rogers and Xu 1992). Initially, vaccination was accomplished by injection of a formalin-killed bacterin into salmon smolts prior to seawater transfer (Rohovec et al. 1975). However, vaccination by immersion and/or spraying has proved to be more efficient and cost effective when done on a large scale (Amend and Johnson 1981). Bivalent vibrio vaccines containing V. anguillarum (Type I and II) and V. ordalii (Type III) (Table 4.5) are commercially available. Vaccinating salmon at appropriate times against vibriosis can improve survival as much as 90%. In some vaccinations of coho salmon mortalities were reduced from 52% in unvaccinated controls to 4% in fish vaccinated for 20 to

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30 seconds by immersion in a preparation containing 1% vaccine. Horne et al. (1982) reduced V. anguillarum mortalities from 100% in unvaccinated rainbow trout to 53% in immersion vaccinated fish. Ayu and rainbow trout were also successfully vaccinated against V. anguillarum by immersion and intraperitoneal injection by Muroga et al. (1995). Humoral agglutinating antibody resulted from both vaccination methods and significant protection was demonstrated by immersion challenge. Evidence exists that salmonid vaccination against V. anguillarum and V. ordalii not only significantly reduces mortality, but vaccinated fish often have increased growth and lower feed conversion ratios (Hastein et al. 1980). ¨ Thorburn et al. (1987) noted that on Swedish marine net-cage farms, the decision to vaccinate is dependent upon farm size and anticipated vibriosis risk. Vaccination against vibriosis is an appropriate management practice when fish will be exposed to an environment where V. anguillarum or V. ordalii are indigenous. Fish that are transferred to saltwater should be vaccinated against vibriosis 2–4 weeks prior to release. Salmonids can be vaccinated at any size over 2 g but for best and most economical results, it should be done when fish are less than 200/kg. Vaccination of Atlantic salmon with formalin-killed whole cell bacterins has shown promise and is used as the primary means for preventing V. salmonicida in the species. Prior to stocking Atlantic salmon into sea net cages, immersion vaccination reduced V. salmonicida mortality from 7.8–0.4% (Holm and Jørgensen 1987). When Lillehaug et al. (1990) vaccinated Atlantic salmon on Norwegian fish farms, CV mortality was reduced from 24.9% in nonvaccinate fish to 1.87% in vaccinated groups. Hjeltnes et al. (1989) showed that vaccination by injection afforded the most dependable protection against V. salmonicida but double immersion was probably more practical and economical. Schroder et al. (1992) pointed out that vaccination of

Atlantic salmon against V. salmonicida is a practical and beneficial management tool, even though protective immunity breaks down in 1.5–2 years. Enteric redmouth was one of the first fish diseases to be managed with a vaccine (Ross and Klontz 1965). A commercial ERM vaccine was introduced in 1976 and has since become an integral part of disease control in cultured salmonids in the United States, Great Britain, Scotland, Scandinavia, and other parts of Europe (Horne and Robertson 1987). To protect against ERM, trout are vaccinated by immersion in a killed bacterin (Johnson et al. 1982a). The most cost-effective fish size for vaccination is 4–4.5 g, but fish up to 200 g can be vaccinated. Trout smaller than 4 g can be vaccinated but protection will not be as lasting; 1.0 g (4 months), 2.0 g (6 months), and 4.0 g fish (12 months) (Johnson et al. 1982b). Fish should be either immersed for 30 seconds or sprayed with vaccine. A secondary immune response serves as a booster vaccination following exposure to living Y. ruckeri for up to 7 months after initial vaccination (Lamers and Muiswinkel 1984). In an attempt to develop a vaccine against Y. ruckeri Temprano et al. (2005), the aroA gene of the pathogen was inactivated with a DNA fragment containing a kanamycin-resistant determinant. The DNA fragment was reintroduced by allelic exchange into the chromosome of Y. ruckeri by means of a suicide vector and the mutant was injected into rainbow trout. The mutant was not recoverable from internal organs of fish vaccinated with the aroA mutant, but the vaccine conferred significant protection against the pathogenic wild-type relative percent survival of 90%. In a study of Y. ruckeri vaccinates, Tebbit et al. (1981) showed an 84% reduction in ERM mortalities, a 77% reduction in need for medication, and a 13.7% lower food conversion rate. These added values resulting from ERM vaccinations were confirmed by Horne and Robertson (1987). In a trout farm survey in the United Kingdom, approximately half the

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farms that vaccinated against ERM felt it failed to protect fish against disease; however, failure was usually attributed to poor condition of the fish and low water temperatures at time of vaccination (Rodgers 1991). A possible detrimental side effect of trout vaccination against ERM is that subclinical IPN or IHN virus infections can be exacerbated into a clinical state with potential for mortality (Busch 1983). Until recently there was little effort to develop a vaccine for F. branchiophilum. It was noted by Heo et al. (1990) that survivors of BGD episodes were susceptible to subsequent pathogen challenge. On the other hand, Lumsden et al. (1994) found that fish previously exposed to live F. branchiophilum, had received F. branchiophilum-specific serum intravenously, or those that had been bathvaccinated, experienced declining mortality in subsequent exposure compared to previously unexposed controls. Also, fish that had been bath-vaccinated three times by immersion were almost completely protected from experimental challenge. These researchers recognize the vaccination potential for BGD but feel that additional research is needed. Trout and salmon vaccination against bacterial cold-water disease has shown promise. Holt (1993) reported successful vaccination with a formalin-killed bacterin of F. psychrophilum by immersion and injection. Injection (with Freund’s adjuvant) produced complete protection against challenge compared to 43% mortality in nonvaccinated controls; however, immersion resulted in only 11% improved survival. Obach and Laurenci (1991) reported that 40-day-old post-hatch rainbow trout were not protected by immersion vaccination with a heat-inactivated preparation of F. psychrophilum. Practical vaccination against BCWD is very difficult because yolksac fry frequently contract the disease before immunity is possible unless it has been passively acquired from brood stock. LaFrentz et al. (2003) studied the influence of antibody in protecting rainbow trout and the effect of passive immunization on BCWD and de-

termined that antibody alone does not confer protection against the disease but does play a role in protection. However, Kondo et al. (2003) developed a practical vaccination method against BCWD in ayu in Japan. They applied the formalin killed bacterin orally and challenged the fish by immersion at 3 and 7 weeks postvaccination. Results showed that ayu were protected against BCWD. Since the first experimental vaccination for BKD was reported by Evelyn (1971b), there have been attempts to examine vaccines potential for controlling this disease (Paterson et al. 1981a; McCarthy et al. 1984). Some successful vaccinations have been reported, and results of some studies have been encouraging. It was found by Wood and Kaattari (1996) that the removal of p57 from R. salmoninarum cells enhanced its immunogenicity and resulted in a 20-fold increase in detectable antibody titers. Tests indicated that the antibody almost exclusively reacted with carbohydrate moieties on p57 negative cells leading to the conclusion that removal of the virulence factors from R. salmoninarum enhances antibody response in fish and is another step toward vaccine development. Two nutritionally mutant attenuated strains of R. salmoninarum (Strain Rs TSA1 and Rs-BHI1) that grow on BHI media were used as live vaccines via intraperitoneal injection of in Atlantic salmon (Daly et al. 2001). The best protection was achieved with the Rs TSA1 strain with RPS of 50 and 76 in two trials compared to 100% mortality in nonvaccinated controls. Results suggest that the nutritionally mutant strain could be used as a live vaccine by injection. There is evidence that vaccination of salmonids against R. salmoninarum can be successful. In contrast Alcorn et al. (2005) evaluated one commercial vaccine and five experimental vaccines against the pathogen. They found that none of the vaccines induced protective immunity against R. salmoninarum from infection in cohabitation with infected fish that were held in cages in tanks in which the vaccinates were held.

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Piscirickettsiosis (P. salmonis) is a relatively new salmonid disease and has received scant vaccination attention. Birkbeck et al. (2004) vaccinated Atlantic salmon with formalin or head-inactivated preparations via intraperitoneal injection. They found that the relative percent survival 6 months postvaccination was 71% for heat-inactivated and 50% for formalin inactivated preparations. This compared to 82% mortality in control fish. In view of these results, there is good potential for development of a vaccine against P. salmonis.

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Bravo, S. 1994b. First report of Piscirickettsia salmonis in freshwater. FHS/AFS Newsletter 22:6. Bricknell, I. R., D. W. Bruno, and J. Stone. 1996. Aeromonas salmonicida infectivity studies in gold sing Wrasse, Ctenolabrus rupestris (L.). Journal of Fish Diseases 19:469–474. Brocklebank, J. R., T. P. Evelyn, et al. 1993. Rickettsial septicemia in farmed Atlantic and chinook salmon in British Columbia: clinical presentation and experimental transmission. Canadian Veterinary Journal 34:745–748. Brown, L. L., T. P. T. Evelyn, et al. 1995. Bacterial species other than Renibacterium salmoninarum cross–react with antisera against R. salmoninarum but are negative for the p57 gene of R. salmoninarum as detected by the polymerase chain reaction (PCR). Diseases of Aquatic Organisms 21:227–231. Bruno, D. W. 1986. Histopathology of bacterial kidney disease in laboratory infected rainbow trout, Salmo gairdneri, Richardson, and Atlantic salmon, Salmo salar L. with reference to naturally infected fish. Journal of Fish Diseases 9:523–537. Bruno, D. W. 1990. Presence of a saline extractable protein associated with virulent strains of the fish pathogen Renibacterium salmoninarum. Bulletin of the European Association of Fish Pathologist 10(2):8–10. Bruno, D. W. 2004. Prevalence and diagnosis of bacterial kidney disease (BKD) in Scotland between 1990 and 2002. Diseases of Aquatic Organisms 59:125–130. Bruno, D. W., T. S. Hastings, and A. E. Ellis. 1986. ˚ Histopathology, bacteriology and experimental transmission of cold-water vibriosis in Atlantic salmon Salmo salar. Diseases of Aquatic Organisms 1:163–168. Bruno, D. W., and A. L. S. Munro. 1986. Observations on Renibacterium salmoninarum and the salmonid egg. Diseases of Aquatic Organisms 1:83–87. Brunt, J., A. Newaj-Fyzul, and B. Austin. 2007. The development of probiotics for the control of multiple bacterial diseases of rainbow trout, Oncorhynchus mykiss (Walbaum). Journal of Fish Diseases 30:573–579. Bullock, G. L. 1972. Studies on selected myxobacteria pathogens for fishes and on bacterial disease in hatchery-reared salmonids. Bureau of Sport

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marine environment. Journal of Fish Diseases 17:375–385. Egidius, E., K. Andersen, et al. 1981. Cold-water vibriosis or “Hitra disease” in Norwegian salmonid farming. Journal of Fish Diseases 4:353–354. Egidius, E., O. Soleim, and K. Andersen. 1984. Further observations on cold-water vibriosis of Hitra disease. Bulletin European Association of Fish Pathologist 4 (3): 50-51. Egidius, E., R. Wiik, et al. 1986. Vibrio salmonicida sp. nov., a new fish pathogen. International Journal of Systematic Bacteriology 36:518–520. Ehlinger, N. F. 1977. Selective breeding of trout for resistance to furunculosis. New York Fish and Game Journal 24:26–36. ´ Ekman, E., G. Åkerman, et al. 2003. Nanoinjection as a tool to mimic vertical transmission of Flavobacterium psychrophilum in rainbow trout Oncrohynchus mykiss. Diseases of Aquatic Organisms 55:93–99. ´ Ekman, E., G. Åkerman, et al. 2004. Impact of PCB on resistance to Flavobacterium psychrophilum after experimental infection of rainbow trout Oncorhynchus mykiss eggs by nanoinjection. Diseases of Aquatic Organisms 60:31–39. Ekman, E., H. Borjeson, and N. Johansson. 1999. ¨ Flavobacterium psychrophilum in Baltic salmon Salmo salar brood fish and their offspring. Diseases of Aquatic Organisms 37:159–163. Elliott, D. G., and T. Y. Barila. 1987. Membrane filtration fluorescent antibody staining procedure for detecting and quantifying Renibacterium salmoninarum in coelomic fluid of chinook salmon (Oncorhynchus tshawytscha). Canadian Journal of Fisheries and Aquatic Sciences 44:206–210. Elliott, D. G., and R. J. Pascho. 2001. Evidence that coded-wire-tagging procedures can enhance transmission of Renibacterium salmoninarum in Chinook salmon. Journal of Aquatic Animal Health 13:181–193. Elliott, D. G., R. J. Pascho, and G. L. Bullock. 1989. Developments in the control of bacterial kidney disease of salmonid fishes. Diseases of Aquatic Organisms 6:201–215. Elston, R., A. S. Drum, and P. R. Bunnell. 1995. Efficacy of orally adminstered difloxacin for the treatment of Furunculosis in Atlantic salmon held in seawater. Journal of Aquatic Animal Health 7:22–228.

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Wakabayashi, H., S. Egusa, and J. L. Fryer. 1980. Characteristics of filamentous bacteria isolated from a gill disease of salmonids. Canadian Journal of Fisheries and Aquatic Sciences 37:1499–1504. Wakabayashi, H., G. J. Huh, and N. Kimura. 1989. Flavobacterium branchiophila sp. nov., a causative agent of bacterial gill disease of freshwater fishes. International Journal of Systematic Bacteriology 39:213–216. Wakabayashi, H., M. Horiuchi, et al. 1991. Outbreaks of cold-water disease in coho salmon in Japan. Fish Pathology 26; 211–212. Wakabayashi, H., and T. Iwado. 1985. Changes in glycogen, pyruvate and lactate in rainbow trout with bacterial gill disease. Fish Pathology 20; 161–165. Wallbanks, S., A. J. Martinez-Murcia, et al., 1990. 16S rRNA sequence determination for members of the genus Carnobacterium and related lactic acid bacteria and description of Vagococcus salmoninarum sp. Nov. International Journal of Systematc Bacteriology 40:224–230. Waltman, W. D., and E. B. Shotts, Jr. 1984. A medium for the isolation and differentiation of Yersinia ruckeri. Canadian Journal of Fisheries and Aquatic Science 41:804–806. Warren, J. W. 1963. Kidney disease of salmonid fishes and the analysis of hatchery waters. The Progressive Fish-Culturist 25:121–131. Watkins, W. D., R. E. Wolke, and V. J. Cabelli. 1981. Pathogenicity of Vibrio anguillarum for juvenile winter flounder, Pseudopleuronectes americanus. Canadian Journal of Fisheries and Aquatic Sciences 38:1045–1051 Wichardt, U.-P., N. Johansson, and O. Ljunberg. 1989. Occurrence and distribution of Aeromonas salmonicida infections on Swedish fish farms, 1951–1987. Journal of Aquatic Animal Health 1:187–196. Wiens, G. D., and S. L. Kaattari. 1991. Monoclonal antibody characterization of a leukoagglutinin produced by Renibacterium salmoninarum. Infection and Immununity 59:631–637. Wiik, R., K. Anderson, et al. 1989. Virulence studies based on plasmid profiles of the fish pathogen Vibrio salmonicida. Applied Environmental Microbiology 55:819–825. Wiklund, T., K. Kaas, et al. 1994. Isolation of Cytophaga psychrophila (Flexibacter psy-

chrophilus) from wild and farmed rainbow trout (Oncorhynchus mykiss) in Finland. Bulletin of the European Association of Fish Pathologist 14:44–46. Willumsen, B. 1989. Birds and wild fish as potential vectors of Yersinia ruckeri. Journal of Fish Diseases 12:275–277. Wood, J. W. 1974. Diseases of Pacific salmon: their prevention and treatment, Second edition. Olympia, State of Washington, Dept. of Fisheries, Hathery Division, 22pp. Wood, P. A. and S. L. Kaattari. 1996. Enhanced immunogeniocity of Renibacterium salmoninarum in chinook salmon after removal of the bacterial cell surface-associated 57 kDa protein. Diseases of Aquatic Organism 25:71–79. Wood, P. A., G. D. Wiens, et al. 1995. Identification of an immunologically cross-reactive 60kilodalton Renibacterium salmoninarum protein distinct from p57: Implications for immunodiagnostics. Journal of Aquatic Animal Health 7:95–103. Wood, J. W., and J. Wallis. 1955. Kidney disease in adult chinook salmon and its transmission by feeding young chinook salmon. Fisheries Commission of Oreqon, Research Brochure 6: 32pp. Wood, E. M., and W. T. Yasutake. 1956. Histopathology of fish III: Peduncle (“coldwater”) disease. The Progressive Fish-Culturist 18:58–61. Woodall, A. N., and G. Laroche. 1964. Nutrition of salmonid fishes, XI. Iodide requirements of chinook salmon. Journal of Nutrition 824:475–482. Wooster, G. A., and P. R. Bowser. 1996. The aerobiological pathway of a fish pathogen: survival and dissemination of Aeromonas salmonicida in aerosols and its implications in fish health management. Journal of the World Aquaculture Society 27:7–14. Young, C. L. and G. B. Chapman. 1978. Ultrastructural aspects of the causative agent and renal histopathology of bacterial kidney disease in brook trout (Salvelinus fontinalis). Journal of the Fisheries research Board of Canada 35:1234–1248. Zhang, X.-H., and B. Austin. 2003. Pathogenicity of Vibrio harveyi to salmonids. Journal of Fish Diseases 23:93–102.

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Chapter 15

Striped bass bacterial diseases

Most of the bacterial disease organisms associated with other warm and cool-water aquaculture fish species also affect striped bass and hybrid striped bass. In fact, striped bass seem to have a higher level of susceptibility to a greater range of pathogens than most warmwater fish species. There appears to be no difference in disease susceptibility between genetically pure striped bass and their various hybrids with white bass and yellow bass. Expansion of striped bass and striped bass × white bass hybrid culture and intensification of rearing methods have not generated an increased diversity of diseases that affect these fish but may exacerbate many diseases causing them to become more acute (Harrell 1997; Plumb 1997). In some instances, bacterial diseases have limited and severely affected striped bass culture, causing the closure of farms (Hawke 1996). The bacteria that infect striped bass, with several exceptions, are saprophytic, facultative, and opportunistic organisms that often cause debilitating infections following exogenous, inanimate predisposing factors. Species in several genera of bacteria, namely

Health Maintenance and Principal Microbial Diseases of Cultured Fishes, 3rd edition. By J. A. Plumb and L. A. Hanson. Published 2011 by Blackwell Publishing Ltd.

Mycobacterium, Nocardia, Photobacterium, and Streptococcus cause the most problems. Other bacterial pathogens of striped bass include species of Aeromonas, Pseudomonas, Vibrio, Edwardsiella, Francisella, and Flavobacterium.

Mycobacteriosis and nocardiosis Mycobacterium spp. and Nocardia spp. cause chronic granulomatous diseases in fish, which may be difficult to distinguish from each other grossly and will be considered together in this section. Both are generally considered facultative pathogens that require a living host in order to replicate and survive in nature for extended periods. However, some species of Mycobacterium and Nocardia are considered environmental organisms that can be found in soil or water (Gordon 1985; Kirschner et al. 1992; Kamala et al. 1994). These two groups of acid-fast staining bacteria were first discovered as pathogens of common carp in Europe during the latter part of the nineteenth century. Mycobacteriosis was originally known as “fish tuberculosis” or “piscine tuberculosis” because the causative organism is taxonomically similar to Mycobacterium tuberculosis, which causes tuberculosis in humans. It was later suggested that the disease in fish should 419

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more correctly be called fish “mycobacteriosis” since other than the organism’s acid-fast staining characteristic and taxonomic classification, very little similarity exists pathologically between the fish infection and human tuberculosis. A review of mycobacteriosis in marine fish was recently published (Jacobs 2009), and Decostere et al. (2004) provided a review of piscine mycobacteriosis and its relationship to human disease. Recently, members of the M. tuberculosis clade (genotypes) associated with fish were reviewed (Kaattari et al. 2006). Included in this clade is M. marinum, a fish disease and zoonotic agent, and M. ulcerans, the causative agent of Buruli ulcer in humans in West and Central Africa that has been associated with aquatic environments and fish. Mycobacterium spp. historically associated with disease in fish are M. marinum, M. fortuitum (Van Duijn 1981), or M. chelonae (Arakawa and Fryer 1984), now M. abscessus (Kaattari et al. 2006). However, three additional species, M. shottsii (Rhodes et al. 2003), Mycobacterium pseudoshottsii (Rhodes et al. 2005), and Mycobacterium chesapeaki (Heckert et al. 2001), were recently described from striped bass in the Chesapeake Bay (Virginia and Maryland). Austin and Austin (2007) listed at least 11 species of Mycobacterium that have been reported in fish. In a survey of striped bass in Chesapeake Bay, Rhodes et al. (2004) found M. shottsii in 25% of the fish, but six additional species of Mycobacterium were identified in 3% of the fish examined, and a total of 57% of the fish were infected with one or more species. Modern molecular techniques applied to taxonomy have resulted in a name change for M. chelonae from M. chelonae subsp. abscessus to M. abscessus and delineation of different genotypes of M. marinum, causing disease in fish. Important species involved in the syndrome referred to as fish nocardiosis are Nocardia asteroides and Nocardia seriolae, formerly N. kampachi. These acid-fast bacilli also cause granulomatous disease in fish, which grossly

may be difficult to distinguish from mycobacteriosis.

Geographical range and species susceptibility Mycobacteriosis occurs in wild and cultured marine and freshwater fish throughout the world but more frequently in saltwater environments. While most serious in cultured fish, mycobacteriosis does occur in wild populations of striped bass (Gauthier et al. 2008). Sakanari et al. (1983) reported a high prevalence of mycobacteriosis in wild striped bass in California and Oregon, and in the late 1990s, the disease appeared in striped bass in Chesapeake Bay of Maryland and Virginia (Heckert et al. 2001; Overton et al. 2003)). Nigrelli and Vogel (1963) listed over 150 marine and freshwater fish species, including salmonids and ornamentals, from which M. fortuitum and/or M. marinum had been documented. Heckert et al. (2001) stated that 160 fish species from freshwater and saltwater were susceptible. This broad spectrum of species susceptibility indicates that all teleosts should be considered possible hosts. Under certain conditions, cultured and wild striped bass and their hybrids are particularly susceptible to M. marinum, M. shottsii, M. pseudoshottsii, and M. chesapeaki, although the latter three species do not seem to have been reported outside of the Chesapeake Bay drainage. Mycobacterium abscessus, formerly M. chelonae, occurs in salmonids in Japan (Arakawa and Fryer 1984). Mycobacterium spp., particularly M. marinum, is also pathogenic to other poikilotherms such as frogs, snakes, and lizards as well as humans and other homeotherms. Nocardia sp. infections in fish have been reported from the United States, Argentina, Germany, Spain, Japan, China, and Taiwan (Post 1987; Chen et al. 1988, 2000; dos Santos et al. 2002; Lan et al. 2008; Shimahara et al. 2008). It is likely that the occurrence of nocardiosis is worldwide in a variety of

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freshwater and saltwater fish but occurs less frequently than mycobacteriosis. In fact, dos Santos et al. (2002), studying mycobacteriosis in turbot in Spain, found M. marinum and Nocardia sp. in the same fish, but the latter was of lower frequency. Nocardia infections have also been documented in rainbow trout, brook trout, yellowtail, pompano, sea bass, Formosa snakehead, giant gourami, largemouth bass, neon tetra, and ornamental fish (Snieszko et al. 1964a; Kitao et al. 1989; Chen 1992; Chen et al. 2000; Lan et al. 2008).

Clinical signs The clinical signs in fish infected with Mycobacterium spp. or Nocardia sp. are similar regardless of fish species. Mycobacteriosis in striped bass is sometimes called “wasting disease” because of its manifesting muscular emaciation. Mycobacterium marinum-infected striped bass are lethargic, darkly pigmented, progressively emaciated with muscle loss and sunken abdomens, and have occasional ulceration and hemorrhaging in the skin; however, these gross external clinical signs vary with other fish species. Infected fish may also be anorexic, have grayish and irregular skin ulcerations (Figure 15.1), deformed vertebrae and mandibles, and exophthalmia, resulting in loss of one or both eyes. Externally, nodular granulomas in the muscle appear as diffuse, light brown spots, or swollen areas that can rupture into ulcers. Fish develop lepidorthosis before scales are lost. White streaks (granulomas) also occur parallel to cartilaginous gill filament supports. Secondary sexual characteristics (hooked jaw and color changes) do not develop in infected adult Pacific salmon. These fish may be smaller than normal, more darkly colored, and have undeveloped gonads. Diseased ornamental fish usually lose their bright coloration. Internal gross pathology is more consistent among fish species, the most notable being the occurrence of granulomas of varying size in

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the liver, spleen, and head and trunk kidney. The surface of the liver may be pale and rough, with a sandpaper-like, granular texture (Figure 15.1). Granulomas in hybrid striped bass are most numerous in the spleen and head kidney, resulting in enlargement of the organs (Figure 15.1). Granulomas also develop in the heart and mesenteries. Clinical signs of Nocardia infections are similar to those of mycobacteriosis but are generally less acute (Austin and Austin 2007). Fish do not actively feed, swim in a rapid tail-chasing mode or are sluggish and emaciated with abdominal distension. Fish may also lose scales, frequently exhibit exophthalmia with opaque eyes. Blood pools under the epithelium of the oral and operculum cavities and multiple yellowish white nodules varying in size from 0.5 to 2.0 cm in diameter are scattered throughout the muscle, gill, heart, liver, spleen, ovary, and mesenteries.

Diagnosis Mycobacteriosis is routinely diagnosed by detecting strongly acid-fast (Ziehl-Neelsen) staining (red), rod-shaped bacteria in smears from nodules or histological sections of granulomas (Figure 15.1) and isolation on suitable agar. Lowenstein-Jensen, Middlebrook (7H10) or Petragnani media provide the best growth for Mycobacterium spp., but M. marinum, M. fortuitum, and M. abscessus can be isolated on blood agar. Heckert et al. (2001) found M. chesapeaki difficult to isolate on typical mycobacteria isolation agar, but after inoculation in fish cell lines, the bacteria could be maintained on solid Middlebrook (7H10) media. The addition of 5% NaCl to the medium facilitates growth of some mycobacteria. M. chesapeaki failed to grow on media with 5% NaCl but did grow on media containing 0%, 0.5%, 1%, and 3% NaCl with optimum growth on 0% and 0.5% NaCl. All inoculated plates should be sealed to retain moisture because of lengthy incubation times. Rhodes et al. (2004) isolated

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(a)

(c)

(b)

(d)

Figure 15.1 Bacterial diseases of striped bass. (a) Mycobacterium marinum infection. Gills have pale areas at base as a result of granuloma. The liver and spleen (arrows) have white granulomatous, rough surfaces. (b) Acid-fast staining Mycobacterium cells (arrow) in a granuloma. (c) Striped bass infected with Photobacterium damsela. Note the pale, slightly mottled liver with granulomas (large arrows) and the pale, granulomas in spleen (small arrows). (Photos a, b, and c courtesy of J. Hawke.) (d) Aeromonas hydrophila infection in the skin of striped bass (arrow).

Mycobacterium spp. on Middlebrook (7H10) agar with incubation for 3 months at 23◦ C. Mycobacterium marinum is a photochromogen that forms yellow to orange, rough colonies in 1–2 weeks on Lowenstein-Jensen media when incubated in the presence of light at 25–30◦ C but does not produce pigment when incubated in the dark. M. marinum typically will grow over the range of 20–35◦ C, but its optimum is 28–33◦ C (Hedrick et al. 1987). Some strains from the Mediterranean fail to grow above 30◦ C (Ranger et al. 2006). Mycobacterium fortuitum forms colonies in about 7 days when incubated at 25◦ C and grows at 19–42◦ C with an optimum range of 30–37◦ C. Colonies may be smooth, rough, moist, dry, raised, or flat, depending upon

the media and age of culture. Material from skin and other tissues containing mixed bacteR rial species are treated with 0.3% Zepheran prior to primary culture, after which pure cultures can be maintained on Lowenstein-Jensen media. Colonies of M. shottsii on Middlebrook agar are rough, nonpigmented, and flat with irregular margins that become umbonate upon aging. Visible colonies are observed at 4–6 weeks incubation at 23◦ C with little growth at 30◦ C. Mycobacterium shottsii does not grow on Lowenstein-Jensen media with 5% NaCl (Rhodes et al. 2003). M. chesapeaki forms nonchromogenic, smooth colonies on Middlebrook (7H10) in about 45 days at 28◦ C with little growth at 37◦ C (Heckert et al. 2001).

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Nocardia spp. and Mycobacterium spp. cause similar gross pathology; therefore, they can best be distinguished by culture or molecular diagnostics. Nocardia spp. is more easily isolated than Mycobacterium spp. and typically grows faster with visible colonies occurring in 4–10 days. Nocardia spp. is Grampositive, weakly acid-fast, nonmotile, long and branching rods detected either in tissue sections or smears from granulomatous nodules (Chen et al. 1988). Nocardia spp. colonies are irregular and rough; white, pinkish, orange, or yellow in color; and some may require up to 21 days at 18–37◦ C for growth (Frerichs 1993). The cells isolated on Lowenstein-Jensen medium and trypticase soy agar are beadlike or long slender filamentous rods (Wang et al. 2005). Nocardia spp. is also differentiated from Mycobacterium spp. by the production of aerial hyphae when cultured on solid media. For rapid diagnosis, monoclonal antibodies were developed by Adams et al. (1996) for identification of M. marinum, M. fortuitum, and M. chelonae; however, these MABs were not strain specific when used in a sandwich ELISA system. It is possible that these reagents could, however, be used to detect Mycobacterium spp. in tissues of infected fish, thus providing a rapid diagnostic method. Because Mycobacterium spp. and Nocardia spp. are slow growing and difficult to isolate in pure culture, isolation of bacterial DNA from tissues and subsequent PCR using universal 16S rRNA primers and sequencing analysis has been used with success in several laboratories for identification. Specific PCR primers have been developed for a few of the common fish pathogens. Due to the extremely slow growth of M. shottsii, a nested PCR assay was developed by Gauthier et al. (2008) for its identification in wild striped bass from Chesapeake Bay. A similar test using different primers was also designed to detect M. marinum. Comparison of these molecular assays to culture-based techniques usually yielded similar results and demonstrated their applicability to rapid di-

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agnosis in wild and cultured fish. A diagnostic PCR was developed to detect and identify N. seriolae using primers to species-specific sequences in the 16S rRNA gene (Shimahara et al. 2008).

Bacterial characteristics All Mycobacterium spp. that cause diseases of fish possess the same basic characteristics of the genus; they are Gram-positive, strongly acid-fast, nonmotile bacilli that are nonspore forming and possess mycolic acids. Typically Mycobacterium spp. are pleomorphic rods measuring 0.25–0.35 × 1.5–2.0 µm that may appear filamentous (Wayne and Kubica 1986). M. marinum, M. fortuitum, M. abscessus, M. chesapeaki, M. shottsii, and M. pseudoshottsii can be differentiated based on several biophysical and biochemical properties (Table 15.1). M. marinum produces nicotinamidase and pyrazinamidase, but M. fortuitum does not; M. fortuitum is positive for nitrate reductase, while M. marinum and M. chelonae are negative. M. marinum produces a yellow orange pigment when exposed to light, which M. fortuitum lacks (Wheeler and Graham 1989). Both M. marinum and M. pseudoshottsii produce mycolactone toxin that results in apoptosis and necrosis in cultured cells, but the structure of the molecule is slightly different and less potent than the mycolactone produced by M. ulcerans (Ranger et al. 2006). Optimum growth temperature for all Mycobacterium spp. fish pathogens is 25–33◦ C. Some have relatively short incubation periods of 5–7 days (M. marinum, M. fortuitum, and M. abscessus), and others require longer incubation periods of up to 4 weeks (M. chesapeaki, M. shottsii, and M. pseudoshottsii) (M. marinum, M. abscessus, and M. chesapeaki are positive for urease, but M. shottsii is negative (Table 15.1). M. shottsii is positive for β-galactosidase, while all others are either negative or questionable. M. shottsii and

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Table 15.1 Selected biophysical and biochemical characteristics of Mycobacterium marinum, M. chelonei, and Nocardia asteroides. Characteristic

M. marinum

M. chelonei

N. asteroides

Possess mycolic acid Colony morphology Cell morphology Cell size (µm) Acid-fast intensity Growth at: 25◦ C 37◦ C Urease Nitrate reduction Acid phosphatase β-Galactosidase Acid phosphatase Catalase Peroxidase Degrades Tween 80∗ δ-Esterase Photochromogenic Growth on 5% NaCl Glucose sole source of carbon Pyruvate sole source of carbon Nicotinamidase production∗ Pyrazinamidase production∗ Mol% G + C of DNA

+ Yellow to orange Long, branching rods 0.25 − 35 × 2 Strong + May adapt + − + − + + + + + + + + + + + 62–70

+ Off-white Long, rods 0.3 − 0.6 × 1 − 4 Strong + − + + + ? + + ? ± ? ? − + ? − − 61–65

+ ? Polymorphic ? Weak + + + + ? ? ? V ? ? ? − ? ? ? ? ? 64–72

Source: Arakawa and Fryer (1984); Lechevalier (1986); Wayne and Kubica (1986). +, positive; –, negative; V, variable; ?, unknown. ∗ Characteristics to separate M. marinum from M. chelonei.

M. chesapeaki are negative for degrading Tween 80, while M. marinum and M. chelonae are positive. Also M. chesapeaki is questionable for catalase, while the other three are positive. M. pseudoshottsii fails to grow at 30◦ C and is photochromogenic whereas M. shottsii is nonchromogenic. Based on 16S rRNA sequence analysis using PCR, presence of a unique insertional sequence, and differences in the biochemical profile, it is believed that M. chesapeaki represents a new species of the genus Mycobacterium (Heckert et al. 2001); however, the name has not officially been recognized. Nocardia are weakly acid-fast, Grampositive, filamentous rods that may appear beaded and show branching. Nocardia spp. may be cultured on a variety of media including TSA with 5% blood or Lowenstein-Jensen; however, isolation in pure culture may be

problematic due to the growth of other faster growing contaminating or secondary bacteria. Colonies appear in 4–10 days at 25◦ C, and colonies may be pigmented and appear dry and wrinkled.

Epizootiology Mycobacteriosis in fish has been known for years, but in the past two decades, the disease has become one of the most serious infections to occur in hybrid striped bass reared in intensive, recirculating culture systems, and in wild populations of striped bass. The disease, typically caused by M. marinum, is usually chronic in intensively cultured hybrid striped bass, a condition that may progress to subacute infection, but seldom becomes acute with high daily mortality. Recent surveys of wild

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striped bass populations in the Chesapeake Bay of Maryland and Virginia showed a high infection incidence of Mycobacterium spp. In a prevalence study in Chesapeake Bay, Overton et al. (2003) found of 217 striped bass assayed, 38% had clinical signs of mycobacteriosis, had a poor condition factor, and decreased overall health. Sakanari et al. (1983) found the prevalence of mycobacteriosis in wild striped bass to be 25–68% in California and 46% in Oregon. Gauthier et al. (2008) reported that M. shottsii and M. pseudoshottsii are the dominant species from diseased striped bass in Chesapeake Bay. However, Rhodes et al. (2004) suggested that because eight species of Mycobacterium were isolated from striped bass, a variety of species could be causative agents of the disease. It is likely that low-level Mycobacteria infections in wild fish populations serve as the pathogen reservoir for cultured fish. However, months, or possibly years, may pass between natural exposure to the bacterium and appearance of clinical disease. The bacterium source in semi-closed and closed recirculating culture systems is unknown, but no mycobacteriosis problems have been reported in pond-cultured striped bass. Experimentally infected fish shed the acidfast bacteria into water, facilitating horizontal transmission. It is assumed that a similar route of infection occurs in wild populations where the incidence increases with age. In Chesapeake Bay disease surveys, it was determined that 11% of 1-year-old striped bass were infected with M. shottsii or M. chesapeaki, and the incidence increased to 60% in 3- to 5-yearold fish (Heckert et al. 2001). During an epidemiological study of Atlantic mackerel from the northeastern Atlantic Ocean, fish over 2 years of age showed evidence of increased presumptive mycobacterial infection and suggested that it affected growth and retarded spawning (MacKenzie 1988). This greater incidence of infection in older fish indicates a chronic condition; however, mycobacteriosis may occur in any age or size fish.

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Diamant et al. (2000) found that the prevalence of M. marinum in net cages of cultured rabbitfish in the Gulf of Eilat, Red Sea, Israel was 50%, and that the prevalence in this species in the area surrounding the cages was 39%. Furthermore, they found 21–42% of all individuals of different fish species surrounding the cages were also infected with M. marinum; these fish could serve as a continuous reservoir of the pathogen. In the 1950s, ingestion of raw contaminated fish viscera was the probable source of mycobacteriosis in cultured salmon in Northwestern United States because at that time raw fish was common in their diet. Mycobacteria free fish were found only in hatcheries where raw fish was not fed and the bacteria disappeared from all hatcheries upon discontinuing raw fish diets and removal of infected fish (Ross 1970). Transmission via oral ingestion was further substantiated when Chinabut et al. (1990) successfully transmitted mycobacteria to snakehead in Thailand by feeding raw offal to naive fish. Conversely, in Australia, Ashburner (1977) reported M. marinum in freshwater-cultured chinook salmon but eliminated feed as a pathogen source and presented evidence that the pathogen was vertically passed to the F1 generation during spawning. It was shown by dos Santos et al. (2002) that because M. marinum and Nocardia sp. were isolated from inlet water of culture units of turbot that the water source was the probable and most plausible source of infection. Juvenile rainbow trout and juvenile chinook salmon were experimentally infected with M. chelonae at 18◦ C; the trout suffered 20–52% mortality, whereas 98% of the salmon died within 10 days postinfection, suggesting a difference in species susceptibility (Arakawa and Fryer 1984). They also found the prevalence of Mycobacterium to be 0–26% in wild juvenile coho salmon compared to 1.4–4.0% in chinook salmon. Mycobacteriosis was not diagnosed in Oregon salmon hatcheries between 1964 and 1981 (Fryer and Sanders 1981), but studies suggest it was still present in wild

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salmon and would continue to occur throughout the anadromous salmon’s life cycle. Morbidity in cultured striped bass populations may be low at any given time, but cumulative mortality can be high. Hedrick et al. (1987) reported that 50% of a M. marinuminfected yearling striped bass population died within months of being stocked into an intensive culture system; the greater the intensification, the more serious the disease became, and 80% of survivors were carriers. In closed recirculating systems where hybrid striped bass were experiencing chronic mortalities, 30–50% of randomly sampled fish had characteristic mycobacteria granulomas in their internal organs (J. Newton, Auburn University, personal communication). Mortality in other farmed fish species in Columbia varied from 35% in naturally infected populations of three-spot gourami to 100% in experimentally infected pejerrey (Hatai et al. 1988). In Israel, cultured European sea bass suffered 50% mortality from mycobacteriosis, but pathogen prevalence was 100% of the fish (Colorni 1992). In a case in Spain where cultured turbot were infected with M. marinum and Nocardia sp., the water supply was probably the source of the pathogen and the mortality was 2% per month (dos Santos et al. 2002). In aquarium fish, the incidence of mycobacteriosis may be high. Beran et al. (2006) found that using Ziehl-Neelsen staining or conventional bacterial culturing that from 14% to 75% of the fish in aquaria were infected with multiple species of Mycobacterium with M. marinum being the predominant species. They also concluded that other constituents of the aquarium environment including snails and crustaceans used for fish feeding had high incidence of Mycobacterium spp. M. marinum and possibly other mycobacteria from fish can potentially cause infections of human extremities, which come in contact with infected fish or contaminated water. Individuals who clean marine fish for a living, handle certain cultured fish, or work with

saltwater ornamental fish are at risk of contracting the disease known as “fish handler’s disease.” The presence of open scratches or wounds on the skin most likely enhances infection. Therefore, caution should be exercised by fish culturists as well as sport fishermen when handling striped bass or any other fish from sources that may harbor the pathogen. Rubber gloves should be worn and hands and forearms washed thoroughly after contact with fish. Skin lesions on humans caused by M. marinum and other species are usually confined to cutaneous lesions on the hands, wrist, and forearms where hard, raised, calcified, granulomas develop. Occasionally, M. marinum is more invasive in humans, causing infection of tendon sheaths, joints, bone, and lymph nodes. About 4 weeks after the bacterium enters the skin, a swelling develops over the bony prominence or of an abrasion. A cyst or abscess develops that may be filled with pus which then ulcerates, leaving a scar. The disease is found in saltwater aquaria where it poses a threat to humans causing a disease called “fish tank granuloma.” M. marinum is also the causative agent of “swimming pool granuloma” (swimmer’s itch), a disease contracted by swimmers. Although Mycobacterium spp. fish pathogens grow best at 25–33◦ C, they may adapt somewhat to the higher body temperatures of humans. It was suggested that the lower skin temperature of extremities is more conducive to infection; therefore, infections seldom become systemic unless an individual is otherwise debilitated (Frerichs 1993). Immunocompromised individuals, such as those suffering from human immune deficiency virus, or undergoing cancer treatments may be particularly susceptible to the so called “atypical mycobacterial infections” (Frerichs and Roberts 1989). Nocardia spp. may be normal inhabitants of soil or water, and fish may serve as pathogen reservoirs. Whatever the source, Nocardia spp. produce a slowly developing chronic infection in fish. There is little available information on Nocardia-caused fish mortalities, but a

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documented case involving N. asteroidesinfected Formosa snakehead cultured in a freshwater pond in Taiwan revealed that 20% of 30,000, 8–9 month old (20–30 cm) fish were killed in 2 weeks (Kariya et al. 1968). Successful experimental transmission of nocardiosis has been inconsistent. Snieszko et al. (1964a) were unable to orally transmit N. asteroides from rainbow trout to other trout, but were somewhat more successful with transmission by injection, requiring 1–3 months for disease to develop. Chen (1992) transmitted N. asteroides from largemouth bass to other individuals of the same species by intramuscular injection, which resulted in characteristic granular nodules in the visceral organs and 100% mortality. N. seriolae is a pathogen of marine fish species worldwide, but most accounts have been described from Japan, China, and Taiwan in yellowtail, amberjack, Japanese flounder, sea bass, striped mullet, and yellow croaker (Shimahara et al. 2008). The disease begins as a “silent infection” as the organism can multiply in the tissues with no visible outward clinical signs and can go undetected in fingerling fish for months (Sheppard 2005). It is believed that the practice of feeding raw fish to fry and fingerlings may be a source of the pathogen. In marine finfish culture, it is believed that nocardia infections progress rapidly in the summer months when water temperatures are 24◦ C or greater, but mortalities do not occur until water temperatures begin to decline in the fall and early winter perhaps due to declining immune responses of the host. Kusuda and Nakagawa (1978) reported successful transmission of the disease in yellowtail by injection or by smearing surface wounds with N. seriolae (formerly N. kampachi), suggesting that the route of infection is more likely through epidermal injury than orally. Nocardia seriolae caused over 17% mortality in cultured sea bass in Taiwan (Chen et al. 2000), while Wang et al. (2005) isolated N. seriolae from cultured yellow croaker in China where it caused 15% losses during August to October. N. seriolae has caused chronic mortali-

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ties in pompano reared in sea cages in China. The disease became more acute, and mortalities increased as water temperatures cooled in the late fall (Lan et al. 2008; J. P. Hawke College of Veterinary Medicine, Louisiana State University, Baton Rouge, Louisiana, personal communication).

Pathology In fish, mycobacterial infection results in proliferation of connective tissue but stimulates little inflammatory response other than granulomatous inflammation (Van Duijn 1981). Mycobacteriosis in teleosts is considered less cellular than is tuberculosis in mammals, and some refute the presence of Langerhans giant cells in fish, which characterize human tuberculosis (Nigrelli and Vogel 1963; Giavenni 1979). However, Timure et al. (1977) showed caseation, typical Langerhans cell production, and cell-mediated immunity in M. marinum infections in plaice. Large masses of bacteria were found in the visceral adipose tissue and hematopoietic tissue of the kidneys, spleens, and livers of young fish as well as in adult fish. Foci of bacteria that surrounded the intestine of young fish disappeared in older fish, leaving large areas of caseous necrosis. The spleen, liver, and kidney had severe lesions with massive concentrations of acid-fast bacteria. Caseous necrosis also formed in the kidney. Gauthier et al. (2004) reported that the primary host response to injected M. marinum was formation of large macrophage aggregations containing phagocytosed bacteria and that the bacteria were always contained in phagosomes. Development of granulomas involved epithelial transformation of macrophages, followed by appearance of central necrosis. In advanced recrudescent lesions, normal tissue was replaced by macrophages, fibroblasts, and other inflammatory leukocytes.

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Gauthier et al. (2003) compared the pathology of M. marinum, M. shottsii, and M gordonae in striped bass. M. marinum produced distinct, severe progression of morphological stages culminating in well-circumscribed lesions surrounded by normal or healing tissues. Large numbers of acid-fast rods were present in granulomas beginning 8 weeks postinfection. Between 25 and 45 weeks postinfection, reactivation of M. marinum infection in some fish was characterized by disintegrating granulomas, revealing inflammation and elevated bacterial densities approaching 107 cfu/g−1 of spleen tissue. Fish infected with M. shottsii or M. gordonae failed to produce severe pathology, but both established persistent infections in the spleen with less dense bacteria than in the case of M. marinum. Granulomas of M. shottsii- and M gordonae-infected fish resolved over time with no reactivation. In these comparative experimental infections, the pathology was more severe in experimentally infected fish with M. marinum than with M. shottsii, or M. gordonae. Wolf and Smith (1999) compared pathology of M. marinum in striped bass and Nile tilapia × blue tilapia hybrids and found that by comparing mortality rates, clinical signs, and histopathology, the bacterium was four times more pathogenic to the striped bass than the hybrid tilapia. They suggested that the difference in pathogenicity may be attributed to intrinsic functional differences in the immunologic systems for the striped bass and hybrid tilapia. Infected fish combat mycobacteriosis by surrounding the bacteria with connective tissue that kills the bacteria by inhibiting metabolism (Van Duijn 1981). The resulting tubercle, which may contain black pigmentation, becomes necrotic, and mineralization takes place leading to cavitation. Acid-fast bacteria are usually present in early developing nodules but are absent in older granulomas. In early stages of Nocardia infection in largemouth bass, acute, serous inflammation occurs and results in the production of ex-

udate containing cellular and bacterial debris, which eventually becomes granulomatous (Chen 1992; Chen et al. 2000). The nodules consist of necrotizing foci surrounded by epithelial cells, fibroblasts, or fibrous encapsulation. The most characteristic tubercular nodule structures are bacillary masses within small cavities surrounded by concentric layers of fibrous tissue. Long, branching, filamentous, weakly acid-fast bacteria lay within these nodules.

Significance The impact of mycobacteriosis on wild fish populations is poorly understood, but in some regions of the world, its influence is significant (Rhodes et al. 2004; Jacobs et al. 2009). In Southern and Eastern United States where M. marinum has become established, its impact is greatest in hybrid striped bass × white bass populations reared in recirculating culture systems and in some wild striped bass populations, particularly in saltwater. Because raw fish is no longer fed to salmon, mycobacteriosis is seldom a problem in today’s cultured salmon but occurs in other species of mariculture fish. Mycobacteriosis is a major disease problem in Japanese mariculture and is a chronic disease problem in ornamental fish in home aquaria and/or in large public aquaria. When a pathogen, as is the case with Mycobacterium spp., can be transmitted from a lower vertebrate to humans, the significance of that disease is elevated. Nocardiosis due to N. asteroides does not appear to be a particularly significant disease because outbreaks have been sporadic and infectious incidence is low. N. seriolae is a serious, chronic problem in marine fish culture worldwide and particularly in China, Japan, and Taiwan. The control and prevention of Nocardiosis is through improved husbandry and management practices, avoidance of the use of raw fish in feeds, and reducing shellfish fouling of cages.

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Photobacteriosis (Pasteurellosis) Pasteurellosis, caused by the halophilic bacterium Pasteurella piscicida, was first described in white perch and striped bass in Chesapeake Bay, Virginia, and Maryland (Snieszko et al. 1964b). The organism was reclassified and renamed Photobacterium damsela subsp. piscicida (Gauthier et al. 1995), and the disease it causes referred to as “photobacteriosis.” The name was later corrected to Photobacterium damselae subsp. piscicida (Truper and DeClari 1997). The disease manifests as an acute septicemia in wild striped bass and cultured hybrid striped bass in the United States and as a subacute to chronic disease in other marine species such as the Japanese yellowtail Seriola quinqueradiata. The systemic infection of wild and cultured yellowtail was at one time known as “pseudotuberculosis” due to the prominent white nodules seen in the internal organs.

Geographical range and species susceptibility In the United States, P. damselae subsp. piscicida has been reported from the Chesapeake Bay area, Long Island Sound, New York, and the Gulf of Mexico (Snieszko 1964b; Robohm 1983; Hawke et al. 1987). Its range outside of the United States now includes Japan, Taiwan, Mediterranean Sea region (Spain, Greece, and Malta), France, Italy, Norway, Denmark, Portugal, Croatia, Egypt, Turkey, and Israel (Kimura and Kitao 1971; Tung et al. 1985; Baudin-Laurencin et al. 1991; Toranzo et al. 1991; Bakopoulos et al. 1997; Zorrilla et al. 1999). Photobacteriosis is primarily a disease of marine fish but infrequently occurs in freshwater. There has been one report of the disease in freshwater snakehead in Taiwan; however, these fish had been fed raw marine

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fish products prior to infection (Tung et al. 1985). In addition to striped bass and white perch, fish known to be susceptible to P. damselae subsp. piscicida include yellowtail, Red Sea bream, black sea bream, striped jack, and gilthead sea bream, and a wide variety of other marine fishes (Toranzo et al. 1991; Nakai et al. 1992; Kitao 1993). P. damselae subsp. piscicida was recently associated with high mortalities in cage-reared Atlantic bluefin tuna in Croatia and cobia in Taiwan (Liu et al. 2003; Mladineo et al. 2006), and Pedersen et al. (2009) identified the bacterium in rainbow trout in Denmark.

Clinical signs and findings Photobacteriosis may manifest as an acute or chronic disease but is always a septicemia (Thune et al. 1993). In the acute form, clinical signs are subtle but may include anorexia, loss of mobility, and sinking in the water column, pale gills, dark pigmentation, and presence of discrete petechia at the base of fins and on the operculum (Robohm 1983; Hawke 1996). Internally, an enlarged spleen and mottled liver are evident with other organs appearing normal (Figure 15.1). In chronic infections in yellowtail and other marine species, small, white, nodular lesions grossly similar to mycobacteriosis may be present in the swollen spleen and kidney thus the name “pseudotuberculosis.” The disease is very acute in wild and cultured striped bass and hybrid striped bass in the United States, and the so-called pseudotubercles may not be visible unless antibiotic therapy has slowed the progress of infection.

Diagnosis Photobacterium damselae subsp. piscicida is isolated on ordinary bacteriological media (BHI) or blood agar containing 0.5–4.0% NaCl (2% is optimum) (Robohm 1983;

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Hawke 1996); it does not grow on saltfree media. Colonies of P. damselae subsp. piscicida are 1–2 mm in diameter, entire, convex, viscous, and opaque to translucent when incubated at 25◦ C for 48–72 hours. Definitive identification of the pathogen is by conventional biochemical reactions (API-20E; P. damselae subsp. piscicida is not in the API index, but all strains exhibit the same API 20E profile code no.: 2005004), molecular characteristics, or slide agglutination; however, some genetic variation between strains may be detected by plasmid profiles and/or RAPD analysis (Magarinos ˜ 2000; Hawke et al. 2003). Several other studies emphasized the use of rapid detection and identification methods for P. damselae subsp. piscicida using molecular technology. Jung et al. (2001) distinguished between P. damselae subsp. piscicida and P. damselae subsp. damselae using monoclonal antibody to detect bacteria in kidney, spleen, liver, and red blood cells of infected sea bass. Kvitt et al. (2002) used direct amplification of 16S rRNA gene sequences and genotypic variation as determined by amplified fragment length polymorphism (AFLP) to identify the pathogen. The method is highly sensitive and has immediate practical consequences by providing diagnosticians with rapid, accurate diagnosis. A rapid detection method developed by Rajan et al. (2003) using a combination of PCR and plating on thiosulfate citrate bile salts-sucrose agar (TCBS-1) distinguished between P. damselae subsp. piscicida and P. damselae subsp. damselae. The later subspecies grows on TCBS-1 producing green colonies while the former does not. It was concluded that TCBS-1 was cost and labor effective compared to other more traditional methods of identification. A rapid PCRRFLP method, developed by Zappulli et al. (2005) for identification of P. damselae subsp. piscicida, eliminates isolation and laborious biochemical techniques because of similarities of this species with P. damselae subsp. damselae.

Bacterial characteristics Photobacterium damselae subsp. piscicida is a Gram-negative, bipolar staining, nonmotile bacillus that is oxidase and catalase positive (Snieszko et al. 1964b; Hawke 1996). The pleomorphic rods measure 0.5–0.8 µm by 0.7–2.6 µm. The organism has an optimum growth temperature of 23–27◦ C and grows at 10–30◦ C but not at 37◦ C. P. damselae subsp. piscicida is generally considered a homogeneous species because isolates from various geographical locations are biochemically, morphologically, and physiologically similar (Magarinos et al. 1992; Bakopoulos et al. 1995; ˜ Hawke 1996; Hawke et al. 2003) (Table 15.2). However, strains from Louisiana, Chesapeake Bay, Greece, and Japan differ in their plasmid profiles (Hawke et al. 2003). All strains of P. damselae subsp. piscicida tested serologically by Robohm (1983) and Magarinos ˜ et al. (1992) were similar. Kimura and Kitao (1971) were unable to serologically distinguish Japanese from American isolates. Also, Kitao and Kimura (1974) identified the pathogen l00% of the time by using a direct FAT on impression smears from organs that exhibited characteristic white lesions. Mori et al. (1976) detected incipient infection in spleens and/or kidneys of yellowtail by using fluorescent antibody and suggested that this technique could be used for detection of subclinical infections. The Aquarapid-Pp ELISA kit, developed to detect European isolates, was cross-reactive with the P. damselae subsp. piscicida isolate from Louisiana, providing further evidence of homogeneity (Hawke 1996). However, Kusuda et al. (1978) could not differentiate P. damselae subsp. piscicida isolates by immunoelectrophoresis, implying that more than one serological strain may be involved. Strains of P. damselae subsp. piscicida from Louisiana, Chesapeake Bay, Greece, Japan, and Israel were similar biochemically, phenotypically, and in enzyme activity (Hawke et al. 2003). Random amplified polymorphic DNA

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Table 15.2 Biophysical and biochemical characteristics of Photobacterium damsela subsp. Piscicida. Characteristic∗

Reaction

Characteristic

Reaction

Cell morphology Gram stain Motility Growth at: 10◦ C 15◦ C 25◦ C 35◦ C Production of: Catalase Oxidase Phenylalanine deaminase Gluconidase Arginine dihydrolase 2–3, butanediol dehydroginase Nitrate reduction Methyl red Voges–Proskauer Oxidation/fermentation glucose Gas from glucose

Short rod – (bipolar) − − + + −

Growth in 0.0% NaCl 0.5% NaCl 3.0% NaCl 5.0% NaCl Growth on: TSA + 2% NaCl Arginine dihydrolase Phospholipase Acid from: Glucose Mannose Galactose Fructose Maltose Sensitivity to: Vibriostat 0/129 Novobiiocin Degradation of: Arginine Starch Tween 80

− + + − + + +

+ + − − + + − + + +/+ −

+ + + + (+) + + + + −

Source: Gauthier et al. (1995); Hawke (1996). ∗ Negative for indole production, citrate utilization, β-galactosidase (ONPG), elastase, amylase, urease, tryptphane deaminase, lysine and ornithine decarboxylase; negative for acid from sucrose, lactose, rhamnose, arabinose, amygdalin, melibiose, mannitol, inositol, sorbitol, and glycerol.

(RAPD) analysis has been used to place strains from different geographic regions into two different clone groups (Magarinos et al. 2000; ˜ Hawke et al. 2003). Photobacterium damselae subsp. piscicida possess transferable multiple-drug resistance plasmids (Kim et al. 2008). Plasmid pP91278 carries resistance to tetracycline, trimethoprim, and sulfonamide, and pP99-018 carries resistance to kanamycin, chloramphenicol, tetracycline, and sulfonamides; these plasmids were isolated in the United States (pP91278) and Japan (pP99-018). These transferable plasmids could present important drugresistant problems in treating photobacteriosis.

Epizootiology The initial epizootic of P. damselae subsp. piscicida, originally called Pasteurella piscicida, occurred in the Chesapeake Bay involving a

massive white perch and striped bass die-off (Snieszko et al. 1964b). The epizootic began in the lower Potomac River in June and spread throughout the Chesapeake Bay during July. At the time of the epizootic, the white perch population was high and the bay and its tributaries were heavily polluted with organic material, conditions that were thought to have contributed to the disease outbreak (Sinderman 1970). During the year following the epizootic, commercial harvest of white perch in the bay was reduced by almost one–half, leading to a speculation that P. damselae subsp. piscicida had killed up to 50% of the population. No other wild fish kill episode associated with the pathogen has been as devastating as was the Chesapeake Bay epizootic. P. damselae subsp. piscicida is the most serious infectious disease problem of the striped bass industry along the Gulf of Mexico coast especially in Louisiana (Hawke 1996). Hawke et al. (1987) reported mortality of about 80% in juvenile striped bass cultured in brackish

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water ponds in Alabama. From 1990 to 2000, the Louisiana Aquatic Diagnostic Laboratory identified 50 cases of photobacteriosis in cultured hybrid striped bass with mortality of 5–90% (Hawke et al. 2003). Nakai et al. (1992) reported that in Japan, photobacteriosis was responsible for the loss of 34% of 10,000 sea-cage-reared juvenile striped jack. Mortality from P. damselae subsp. piscicida varies in other cultured fish but is generally high in young yellowtail and black sea bream. Photobacterium damselae subsp. piscicida appears to be a normal inhabitant of the estuarine environment, where fish are most likely the natural host; however, the pathogen’s survival in water is short lived outside of the host. The pathogen survives for less than 2 days in freshwater, less than 3 days in sterile brackish water, and less than 5 days in saltwater. In view of these data, it was suggested that fish other than clinically ill individuals may be the pathogen reservoir (Toranzo et al. 1982; 1991). Route of P. damselae subsp. piscicida transmission is unknown, but its short-lived nature in brackish water, and inability to survive in freshwater, has led to speculation that transmission is probably fish to fish, even though a carrier or latent condition has not yet been proven. In support of the theory that fish are the pathogen reservoir, dead fish are definitely a reservoir of the pathogen because Matsudka and Kamada (1995) showed that yellowtail shed 107 to 109 CFU/fish 10 minutes after death and for 5 days thereafter during decomposition. Environmental conditions probably play a major role in determining the severity of photobacteriosis (Sinderman 1970). Matsusato (1975) reported that disease incidence in yellowtail rose during the rainy season when salinity dropped below 30 ppt and water temperatures were optimum (25◦ C) for the pathogen. Photobacteriosis generally occurs in U.S. striped bass populations during optimum water temperatures of spring and autumn.

Photobacterium damselae subsp. piscicida can be highly pathogenic, but susceptibility varies with fish species. In experimental transmission studies with an immersion LD50 of about 687 CFU/mL, hybrid striped bass began to die 5 days postexposure, and deaths continued through 10-day postexposure (Hawke 1996). An LD50 of about 100 CFU/mL by injection was established for Formosa snakehead (Tung et al. 1985), but in experiments with other species, greater numbers of bacteria were required to kill fish. Nakai et al. (1992) established an injection LD50 of 1,000 CFU/fish in striped jack and an LD50 of 10 million CFU/fish in red Sea bream.

Pathology Pathology of photobacteriosis varies to some degree between fish species. According to Hawke (1996), experimental photobacteriosis in hybrid striped bass is characterized by a generalized septicemia. Severe necrosis occurs in the spleen, kidney, and gills; the liver is generally affected to a lesser degree. Microscopically macrophages laden with bacteria are seen in each of these tissues, but as noted by Wolke (1975), little inflammation is present in infected tissue. The “pseudotubercles” reported in some species of fish are actually large accumulations (colonies) of bacteria forming white nodules in the tissues. Bacterial counts in the blood 6 days postinfection range from 107.7 to 109.6 CFU/g of tissue or milliliter of blood. No histopathology was seen in the olfactory lamellae, brain, intestine, heart, or skin of infected fish. In wild populations, few pathological changes are noted in fish with acute photobacteriosis, whereas a chronic infection is characterized by miliary lesions in the kidney and spleen (Wolke 1975; Robohm 1983). Also in the chronic form, necrotic lymphoid and peripheral blood cells collect in the spleen, and focal hepatocyte necrosis occurs in the liver. No inflammatory response occurs in infected

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white perch, but spleens of experimentally infected juvenile striped bass develop multiple foci of bacterial colonies and reduced densities of cells. In infected yellowtail, live bacteria are present in phagocytes, which become swollen to the point that capillary blood flow is blocked (Kubota et al. 1970). In chronically infected yellowtail, bacteria are localized in nodules in the spleen and kidney. In striped bass naturally infected with P. damselae subsp. piscicida, extensive necrosis of spleen lymphoid tissue, coagulation necrosis, and karyorrhexis occurred (Hawke et al. 1987). As in experimental infections similar but less severe, histopathology was seen in the liver. Rod-shaped bacteria inhabit sinusoids and hepatic vessels of livers, and large areas exhibit hyperplasia of reticuloendothelial cells lining the hepatic sinusoids. Inflammatory cellular accumulations are absent in striped bass. Pathogenesis of P. damselae subsp. piscicida was clarified by Elkamel (2002), who found that the bacterium replicates within striped bass macrophages where the organism increases for up to 18 hours. The bacterium also replicates in EPC, CCO, and FHM cells in vitro (Elkamel and Thune 2003). Bacteria are found in small close-fitting vacuoles that develop into large clear spacious vacuoles; however, bacteria are not found in the cell cytoplasm. In addition, Elkamel et al. (2003) reported that P. damselae subsp. piscicida replicates within macrophages obtained from hybrid striped bass, indicating that the bacterium is a facultative intracellular pathogen that can survive and multiply within hybrid striped bass macrophages. Nakai et al. (1992) found that extracellular products (ECPs) of P. damselae subsp. piscicida were as pathogenic to striped jack and Red Sea bream as was the bacterium, suggesting that pathology is the result of ECPs. In support of this, Noya et al. (1995) found reduction of circulating red blood cells in gilthead sea bream injected with ECPs from P. damselae subsp. piscicida. Severe lesions in the liver and gills further suggested the importance of tox-

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ins in the pathogenesis of P. damselae subsp. piscicida. Injection of ECPs also produced an inflammatory response similar to that associated with live bacteria; including lymphopenia, granulocytosis, an increase in peritoneal exudate cells, and degranulation of eosinophil granular cells. As pathology stimulated by live bacteria progressed, it was noted that degenerating macrophages contained intact bacteria causing the investigators to postulate that these macrophages played a major role in the dissemination of the organism. Death of infected hybrid striped bass is likely attributed to respiratory failure resulting from a combination of gill epithelium and support tissue necrosis and congestion of the sinusoids and capillaries of secondary gill lamellae (Hawke 1996).

Significance Photobacteriosis is a major disease problem in some mariculture communities in the United States where it is a major threat to the striped bass cage culture industry in Louisiana and in other mariculture operations around the globe. The disease has caused closure of some operations, threatened the demise of others, and continues to be a serious problem (Hawke 1996; Hawke 2003 et al.). In some wild, striped bass populations, it is one of the most serious infectious diseases. Photobacteriosis is also a serious threat to yellowtail culture in Japan (Egusa 1983) and to the culture of several fish species in the Mediterranean region. Wherever the disease occurs, there is potential for high mortality and significant economic loss.

Other bacterial diseases of striped bass Other bacteria that cause mild-to-severe disease in cultured and wild striped bass discussed in detail in other chapters are Aeromonas hydrophila complex (motile

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Aeromonas septicemia—MAS) and Flavobacterium columnare (see Chapter 11), Vibrio anguillarum (see Chapter 14), Edwardsiella tarda (see Chapter 13), Francisella sp. (see Chapter 16), and Streptococcus sp. (see Chapter 16). While infections associated with MAS are generally not as severe in striped bass as mycobacteriosis and photobacteriosis, A. hydrophila is a frequently detected pathogen in these fish. Nedoluha and Westhoff (1997) monitored striped bass hybrids in three types of freshwater aquaculture: earthen ponds, flow-through tanks, and recirculating tanks. They found A. hydrophila (MAS) in intestines of 19% of fish sampled, which was the highest incidence of all fish pathogens. There was no statistical difference between the types of culture units; however, no overt MAS infections were noted. Striped bass afflicted with MAS show reduced feeding response or completely stop feeding, and the fish swim lethargically at the surface. While clinical signs are not specific in striped bass, fish may develop mild-to-severe hemorrhage in the skin and fins, and the fins will also have pale, frayed margins (Figure 15.1). As a result of fluid accumulation (edema) in scale pockets, scales may protrude (lepidorthosis) and slough, and the eyes may protrude (exophthalmia). A cloudy, bloody fluid is often present in the body cavity, and internal organs may be hyperemic or pale, depending upon stage of infection. Vibriosis can be a significant problem in saltwater-cage-reared striped bass and wild populations. Although several species of Vibrio are implicated in the disease, the most common species is V. anguillarum. These infections are usually related to stress, high stocking densities, handling, temperature shock, and/or poor water quality. In addition to lethargy, clinical signs of vibriosis in striped bass are hyperemia of the fins and skin, resulting in scale loss and ulcerated epidermal lesions at any location on the body including head and gill covers. Gills are often pale, and eyes may be hemorrhaged with exophthalmia. Internally, the body cavity may contain bloody fluid, the

liver 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. E. tarda is a common pathogen in some cultured fish species and occasionally infects cultured striped bass. Herman and Bullock (1986) reported that 4–5 cm cultured juvenile striped bass in freshwater were infected with E. tarda and was transmitted to naive juveniles by waterborne exposure. As fish became moribund, they swam lethargically at the surface, displayed pale gills, and had a slightly discolored area in the cranium. Epithelium of experimentally infected fish was necrotic, and fins were frayed. Numerous abscesses were present in the kidney accompanied by necrosis of the trunk kidney hematopoietic tissue. In a significant E. tarda infection in wild adult striped bass in the Chesapeake Bay, Baya et al. (1997) indicated that the most notable clinical signs were numerous irregular, coalescing, hemorrhagic ulcers on the body and fins that emitted a distinct odor. Internally the body cavity contained abundant yellowish or bloody mucoid fluid, visceral organs had multiple tiny white foci, and the intestines contained thick white opaque mucus. Histopathology revealed ulcerative dermatitis, cardiac endothelial hyperplasia, and necrotic foci and granulomas in multiple organs. In infectivity trials, the isolate was pathogenic for striped bass, gilthead sea bream, and turbot. The causative agent of columnaris (F. columnare) in freshwater fishes occurs in both cultured and wild striped bass, but degree of severity is greater in cultured populations. However, Flavobacterium spp. was found in 16% of cultured hybrid striped bass intestines (Nedoluha and Westhoff 1997). Columnaris is usually confined to the body, fins, and/or gills, but will occasionally occur systemically. Whitish areas appear on the skin, and lost scales often expose underlying musculature. Fins are usually white and in various stages of fraying; necrotic gill lesions are pale. Lesion margins may be yellowish because of the

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presence of large numbers of bacteria. Although there is no published record of Tenacibaculum maritimum (formerly Flexibacter maritimus), the marine equivalent of “columnaris” in freshwater, infecting striped bass in saltwater, there is no reason to believe it cannot do so. Streptococcus septicemia affects a variety of fish species but is particularly serious in some striped bass culture operations (J. P. Hawke, Louisiana State University, personal communication). The most common species affecting cultured hybrid striped bass are Streptococcus iniae and Streptococcus agalactiae (group B streptococcus). Although Streptococcus spp. have been reported in wild striped bass populations inhabiting brackish water, the infection is more significant in cultured fish reared in brackish water ponds and net cages, where the organism is apparently endemic. The disease is usually chronic, but occasionally subacute. Clinical signs of Streptococcus spp. infection are not particularly specific in striped bass, but fish are generally darker than normal, exhibit erratic, spiral swimming, and often display body curvature. They often have either bilateral or unilateral exophthalmia with hemorrhage in the iris. Hemorrhages develop at the base of fins, in scale pockets, and on the operculum and mouth; ulcerative lesions occasionally occur on the body. A bloody fluid is present in the intestine that is also hyperemic, the liver is pale, and the spleen is dark and greatly enlarged. Mortality of intensively cultured striped bass can be high, especially when water temperatures are 25–30◦ C. Handling and moving fish seems to trigger overt disease where Streptococcus is endemic. Abrasions, loss of scales, and other skin injuries and environmental stressors are also important precursors to Streptococcus spp. infections in some fish species (Chang and Plumb 1996). The presence of gill and skin parasites can also exacerbate infections with Streptococcus spp. The genus Enterococcus is a taxonomic group that was previously included in Streptococcus (Mundt 1986). Enterococcus infections

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have been noted in a few intensive freshwater striped bass units when water temperatures were 28–32◦ C (J. P. Hawke, personal communication). The causative organism is Enterococcus faecium (formerly Streptococcus faecium). When infected with Enterococcus, adult and juvenile striped bass develop a septicemia with hemorrhages in the scale pockets, and the eyes become swollen, hemorrhaged, or cloudy. Enterococcus is a Gram-positive, nonmotile, more ovoid than round cocci. The organisms are beta-hemolytic (Lancefield’s Group D) and grow at 45◦ C, in 40% bile, and in 6.5% NaCl—characteristics that separate them from Streptococcus spp. The full impact of E. faecium on striped bass is unknown, but judging from reported epizootics, the potential is notable. Several additional bacteria have been known to cause disease in striped bass but may not be serious pathogens. Baya et al. (1990) isolated one such bacterium belonging to the genus Moraxella. The organism is a short, Gram-negative rod often appearing in pairs; it exhibits bipolar bodies, is cytochrome oxidase positive, nonfermentative in glucose, and does not produce acid from most carbohydrates. Fish affected with the bacterium had large hemorrhagic lesions on the dorsolateral body surface accompanied by scale loss. Internally, the liver was enlarged and pale or mottled. The swim bladder was inflamed and adhesions connected the liver to the body wall. The role of Moraxella in the disease process was somewhat speculative because of the presence of a viral agent tentatively named striped bass reovirus that was isolated from these fish. Also gills of afflicted fish were heavily parasitized with Trichodina and Ergasilus. Baya et al. (1991) isolated a Carnobacterium-like organism from moribund and dead striped bass and other fish in the Chesapeake Bay of Maryland, but no clinical signs were described. However, the fish had been stressed prior to infection. The organism was similar to Carnobacterium piscicola, which is a Grampositive bacillus that tolerates salinities from

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0 to 6%, has a growth temperature range of 10–37◦ C, and is easily isolated on BHI media or TSA. Toranzo et al. (1993) failed to kill striped bass with C. piscicola by injection, but the organism did cause mild infiltration of the liver, hemorrhage in the kidney, and inflammation of the meninges. The investigators did however kill about 35% of rainbow trout injected with 4.5 × 106 cells. These observations led to the conclusion that C. piscicola possesses low virulence to striped bass, producing only moderate injury to internal organs. However, infected striped bass can be carriers of C. piscicola for at least two months, which suggests that infected fish could be more susceptible to secondary pathogens or environmental stress. Baya et al. (1992) isolated a bacterium, identified as Corynebacterium aquaticum, from the brain of striped bass that were exhibiting exophthalmia. This is the first report suggesting that C. aquaticum, a normal waterborne organism, is pathogenic to fish. The LD50 of the organism in striped bass was 1.0 × 105 colony-forming units. Experimentally infected fish developed hemorrhaging in most internal organs with it being most severe in the brain and eyes. It should also be noted that that C. aquaticum may be infectious to homeothermic animals. Francisella sp. was identified as the cause of chronic mortalities in commercially raised hybrid striped bass in semi-closed recirculating systems in California (Ostland et al. 2006). The organism from hybrid striped bass and tilapia was identified by isolation of bacterial DNA from tissue of diseased fish and amplification of 16S rRNA gene and comparing the sequences to a comparative database (GenBank).

Management of striped bass bacterial disease Chemical and drug availability for controlling infectious diseases of striped bass or their hybrids in aquaculture is limited; therefore, man-

agement, good health maintenance, and disease prevention are keys to successful culture.

Management To reduce potential for catastrophic disease outbreaks in striped bass populations, the highest water quality possible should be maintained. This is best accomplished by using prudent stocking densities, adequate flow of freshwater through intensive culture units, removal of metabolites from recirculating water, supplemental aeration in flow-through and earthen pond culture systems, and providing a high-quality feed. Striped bass are sensitive to improper handling; therefore, gentle netting and handling is essential. Prophylactic chemotherapy of acceptable drugs, i.e., salt or potassium permanganate, during or following handling and/or moving of fish will reduce external parasite loads and reduce the possibility of secondary bacterial infections. In cultured striped bass, the absence of raw fish products in diets will usually disrupt the cycle of mycobacteriosis because it can be a source of the pathogen. Once fish become infected with the organism, it is extremely difficult to treat, and it has been suggested by some that infected fish populations be destroyed, buried, and the facility dewatered and sterilized. Fish should definitely be removed from contaminated facilities, the system cleaned thoroughly with an acid wash to remove as much organic material as possible from water lines, and the facility disinfected with 200 mg/L of HTH, chlorine dioxide, or phenolic compounds. In a study to determine efficacy of common disinfectants to kill M. marinum, Mainous and Smith (2005) found that the most effective disinfectants tested were ethyl alcohol (50 and 70%), benzyl-4-chlorophenol (1%), and sodium chlorite (mixed as 1:5:1 or 1:18:1 (base water:activator)). Each of these reduced or eliminated the number of detectable M. marinum within 1 minute of contact. Recirculating aquaculture systems should be

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equipped with UV and/or ozone water sterilization systems; however, water treatment rates and actual value of these units when acidfast bacteria are present has not been studied in detail. According to Kitao (1993), avoidance of overcrowding and good management may help prevent photobacteriosis and other bacterial diseases. Most other diseases that affect cultured striped bass or their hybrids respond to normal health management practices.

Chemotherapy It must be emphasized that no drugs are approved by the USFDA for striped bass in the United States, and any use of drugs must be approved by the regulating agency. However, some drugs and chemicals have been successfully used prophylactically or in chemotherapy to treat clinical bacterial infections. Prophylactics used to prevent some skin diseases include bath in NaCl (0.5–3% for various periods of time) and/or potassium permanganate (2–5 mg/L for 1 hour to indefinitely). Clinical, systemic bacterial infections are usually treated with medicated feed containing oxytetracycline at a rate of 2.5–3.5 g/45 k of fish per day for 10 days. Xu and Rogers (1993) determined that oxytetracycline injected IP into striped bass cleared from the muscle in 32 days, and when applied in feed, clearance time was reduced. Romet-30 fed at a rate of 50 mg/kg of fish per day is also effective against most systemic bacterial infections. Romet-30 may or may not be effective against Streptococcus spp. Some isolates of M. marinum are sensitive to minocycline, doxycycline, and tetracycline (Contorer and Jones 1979) while others are resistant to most antibiotics (J. Hawke, personal communication). The long-term antibiotic therapy required to keep the disease in remission in fish populations is probably not economically or biologically feasible nor is the practice condoned by regulating agencies.

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Application of medicated feed after disease occurs has successful in controlling photobacteriosis. Ampicillin fed at 10 and 100 mg/kg of body weight per day reduced mortality in yellowtails to 30 and 0%, respectively, compared to a 100% mortality of nonmedicated controls in Japan (Kusuda and Inoue 1976). In Taiwan when an antibiotic was fed to cultured snakehead infected with P. damselae subsp. Piscicida, only a 30% loss was noted (Tung et al. 1985). Hawke et al. (1987) fed oxytetracycline at 50–150 mg/kg of body weight of striped bass per day with only a slight reduction in mortality. However, Nakai et al. (1992) controlled a P. damselae subsp. piscicida infection in striped jack by including oxytetracycline and oxolinic acid in the feed. The fact that transferable multiple-drug resistance plasmids in P. damselae subsp. piscicida may complicate future use of chemotherapy in some instances (Kim et al. 2008). This is problematic particularly in situations where tetracycline and/or sulfonamide are the only available drugs available for treating the disease.

Vaccination Fish vaccination is becoming an important tool in combating bacterial infection in fish including striped bass. Vaccination was effective in prevention of photobacteriosis, especially in Japan, and this method of treatment is expected to become an effective management tool in the future for control of this disease in striped bass. Photobacterium vaccines that contain killed whole-cell bacterins can be effectively delivered by intraperitoneal injection, immersion, spray, or fed orally (Fukuda and Kusuda 1981). Their research reported survival rates of 60–88% with oral and immersion treatments and 100% when using injection and spray vaccination. Fukuda and Kusuda (1985) also found that LPS preparations delivered by immersion or spray provided better protection than did whole-cell

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preparations. Early vaccinations of yellowtail showed a response to an IP injection of formalin-killed P. damselae subsp. piscicida in Freund’s complete adjuvant. Kusuda et al. (1988) demonstrated a high degree of protection with an injected ribosomal vaccine prepared from P. damselae subsp. piscicida. Vaccination by immersion with a live attenuated bacterial preparation has been attempted in striped bass, but to date, no commercial immunogenic preparations are on the market. Hawke (1996) and Thune et al. (2003) developed a live attenuated strains of P. damselae subsp. piscicida by mutation of the specific genes, the siderobiosysnthesis gene (LSU-P1) and the aroA gene (LSU-P2). Hybrid striped bass were then vaccinated by immersion for 15 minutes. Challenge of the fish with wild-type P. damselae subsp. piscicida LADL 91-197 six weeks after vaccination showed a high level of protection by both vaccine strains and more than 90% mortality in nonvaccinated controls. Rogers and Xu (1992) reported successful vaccination of striped bass against vibriosis; therefore, vaccination shows some promise as a preventive treatment for V. anguillarum. A novel vaccine preparation consisting of deactivated whole P. damselae subsp. piscicida cells and extracellular products were used to vaccinate sea bass by intraperitoneal injection, immersion, and incorporation into feed (Bakopoulos et al. 2003). When challenged by immersion 6 and 12 weeks postvaccination small fish (1.5–2 g) had higher survival in all vaccine treatments compared to the commercial product; however, neither group was protected against IP injection challenge. Larger fish (20g) also benefited from vaccination with the novel preparation. A number of factors, including salinity of the medium used to produce the bacteria in the vaccine preparation, can affect the efficacy of a vaccine. Nitzan et al. (2003) demonstrated that the salinity in which P. damselae subsp. piscicida was grown affected immune stimulation in hybrid striped bass. Bacteria grown in

a 2.5% NaCl medium at 25◦ C provided the best protection in hybrid striped bass. No correlation was found between antibody response and protection. In summary, due to the rapid onset and acute nature of many of the bacterial diseases of striped bass and hybrid striped bass, the ineffectiveness of antibiotic therapy due to poor timing or inadequate dose and/or selection of antibiotic resistant strains, vaccines may be the best tool to combat diseases in this fish in the future.

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Chapter 16

Tilapia bacterial diseases

Tilapias (Oreochromis spp.) are the most extensively cultured fish in the world; however, no one bacterial infectious agent is specific for this group. Tilapia are hardy fish with good resistance to bacterial infections as long as they are kept under good water quality conditions, in the proper temperature range, and using proper husbandry practices. Wild or extensively cultured tilapia reared in a normal static warm-water habitat seldom contract serious infectious disease. However, as culture systems intensify (cages, raceways, or recirculating systems), stress due to improper handling, exposure to poor water quality, high fish density requiring high feeding rates, or low water temperatures can exacerbate the impact of some bacterial pathogens on tilapia. Tilapia culture has expanded into temperate and colder climates, where it is more difficult to maintain an ideal environment; consequently, infectious diseases in these environments have become more serious. By some accounts, bacterial diseases may be the number one threat to the future of the tilapia aquaculture industry. The most serious bacterial diseases in cultured tilapia are caused by the Streptococcus spp. (Anonymous 2006) and Francisella asiatica, Health Maintenance and Principal Microbial Diseases of Cultured Fishes, 3rd edition. By J. A. Plumb and L. A. Hanson. Published 2011 by Blackwell Publishing Ltd.

a rickettsia-like organism (RLO), originally noted in tilapia in Taiwan (Chen et al. 1994). The latter has recently caused serious disease problems in cultured tilapia in other areas including Central America, Canada, and Hawaii (Mauel et al. 2007; Soto et al. 2009). Recently Lactococcus garvieae was isolated from Nile tilapia in Brazil (Evans et al. 2009). Other bacterial diseases affecting tilapia include motile Aeromonas septicemia (MAS), Pseudomonas septicemia, and columnaris (Chapter 12), vibriosis (Chapter 15), Edwardsiellosis (Chapter 14), and Plesiomonas shigelloides (Faisal and Popp 1987); however, these probably represent opportunistic pathogens of tilapia, requiring environmental stress for them to manifest disease.

Streptococcosis Reports of Streptococcus spp. infections in fish date back to the mid-1950s (Hoshina et al. 1958), but the first case involving tilapia was a decade later (Wu 1970). There is little question that streptococcosis represents a real danger to tilapia in warm-water aquaculture, especially in intensive systems where it has emerged as a serious problem. The disease may be subacute but is often chronic. Although multiple species of Streptococcus, including 445

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S. agalactiae, S. iniae, S. dysqalactiae subsp. dysgalactiae, S. dysgalactiae subsp. equisimilis, S. pyogenes, S. parauberis, S. equi subsp. equi, and S. equi subsp. Zooepidemicus, have at various times been reported from fish, S. iniae and S. agalactiae are the two that most frequently cause serious disease in tilapia.

Geographical range and species susceptibility Tilapia and a variety of other fishes are affected by streptococci; Kitao (1993) listed 22 fish species that are naturally susceptible to Streptococcus spp., and the list has expanded since then. Streptococcus spp. infections have been reported in tilapia from North America (Canada, Mexico, and the United States), South America (Brazil and Columbia), Central America (Honduras), Australia, throughout Asia (Japan, China, Taiwan, The Philippines, India, and others), South Africa and other African countries, Great Britain, Norway, Israel, Saudi Arabia and Kuwait, and other Middle Eastern countries. Tilapia-associated streptococci is essentially worldwide in distribution (Klesius et al. 2008). Streptococcus iniae and S. agalactiae are the most commonly encountered species in tilapia and other fish species in most countries and occurs in both freshwater and marine environments. Perera et al. (1994) first reported that S. iniae was responsible for chronic mortality of hybrid tilapia (Nile × blue tilapia) on a Texas fish farm. Two species of Streptococcus (S. shiloi and S. difficile) causing meningoencephalitis in tilapia resulted in great economic losses in Israel during the mid to late 1980s (Eldar et al. 1994). These organisms were later shown to be synonymous with S. iniae and S. agalactiae, respectively (Eldar et al. 1995; Vandamme et al. 1997). Evans et al. (2002) isolated S. agalactiae from diseased wild Klunsinger’s mullet and cage-cultured gilthead seabream in Kuwait Bay, Kuwait. In Brazil, Salvador et al. (2005)

identified S. agalactiae from Nile tilapia in addition to other isolates classified as Lancefield group B streptococci. Plumb et al. (1974) isolated a group B-type 1B Streptococcus spp. from ten wild marine species, including Gulf menhaden, silver seatrout, pinfish, Atlantic croaker, hardhead catfish, and others in estuaries of the northern Gulf of Mexico. This organism has also caused mortality in cultured tilapia on Texas and Central American fish farms, where the fish are reared in brackish water (J. P. Hawke, personal communication; LADL case records). Streptococcus dysgalactiae, a group C Streptococcus, has been isolated from diseased cultured tilapia in Louisiana, Mississippi, and Colorado (J. P. Hawke, personal communication) and appears to be similar to the group C Streptococcus isolated from amberjack and yellowtail on marine fish farms in Japan (Nomoto et al. 2004). Group C Streptococcus infections in tilapia are chronic in nature and mortality is low; however, abscesses that form in the muscle may render the fish unmarketable. It is suspected that handling stress and injury to the fish may be a predisposing factor to this type of infection. Clearly the streptococcal pathogens of tilapia are cosmopolitan in distribution. L. garvieae previously known as Enterococcus seriolicida is an important pathogen in Japan and the Mediterranean (Kitao 1993; Kusuda and Salati 1993; Klesius et al. 2008); however, this species has only recently been described as a pathogen of tilapia and pintado in Brazil (Evans et al. 2009).

Clinical signs Clinical signs of streptococcosis in tilapia are not always specific, but in most species of fish, eye disease and meningoencephalitis are common. Affected fish generally are darkly pigmented, lethargic, exhibit erratic and spiral swimming, and body curvature, indicating central nervous system impairment. (Figure 16.1) (Plumb et al. 1974; Kitao

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(a)

(b)

Figure 16.1 Streptococcus iniae-infected tilapia. (a) Tilapia with S. iniae in eye. Note the swelling and hemorrhage and (b) Visceral organs of tilapia with S. iniae infection. Note the opaque cornea, pericarditis (arrows), ascites, and swollen darkened spleen. (Photos courtesy of John P. Hawke.)

1993; Chang and Plumb 1996). Infected fish exhibit abdominal distension, exophthalmia with hemorrhage and opaque corneas, and diffuse hemorrhage in the operculum, skin, and at the base of fins. Epidermal lesions are usually more superficial than those associated with MAS or vibriosis but will occasionally become necrotic, bloody ulcers. Bloody mucoid fluid in the lower intestine may be discharged from the anus. Internal findings include a bloody, sometimes gelatinous, exudate in the abdominal cavity, pale livers, hyperemic digestive tract, and a greatly enlarged, nearly black spleen. The lower gut is flaccid and hyperemic. The streptococci show a pronounced neurotropism, and the bacteria are often isolated from the brain and eye of sick fish.

Diagnosis While recognition of clinical signs is important, isolation of Streptococcus spp. on agar media is essential when diagnosing the disease. Presumptive diagnosis is made by detection of Gram-positive cocci (sometimes ovoid) in histological sections or smears from infected tissues. In Gram-stained smears from infected fish, bacteria may be arranged singly, paired, or as short chains of two to six cells. Streptococci are isolated on Todd-Hewitt, BHI, or TSA agar plates to which 5% sheep blood has been added. For selective isolation, Columbia CNA agar (containing colistin and nalidixic acid) will enhance and expedite isolation from contaminated sites such as the skin, gills, or

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intestine or from mixed infections (Kitao et al. 1981; Hawke 2000). Inoculated media is incubated at 28–37◦ C where small yellowish to gray, translucent, rounded, slightly raised, pinpoint, or pinhead (punctuate) size colonies appear in 24–48 hours. When streptococci are grown in broth, the cells have a greater tendency to form longer chains of up to seven or eight cells.

Bacterial characteristics The most common species of Streptococcus isolated from tilapia are S. iniae and S. agalactiae, while S. dysgalactiae is less commonly seen. Streptococcus iniae is typically betahemolytic and does not belong to a recognized Lancefield serological group. S. agalactiae is a member of Lancefield group B and may be beta-hemolytic or nonhemolytic, depending on the strain and geographic location. S. agalactiae isolates from Kuwait are typically betahemolytic while isolates from the U.S. Gulf Coast are nonhemolytic. Isolates previously identified as Streptococcus spp. in Japan were originally renamed Enterococcus seriolicida (Kusuda et al. 1991) and were more recently renamed Lactococcus garvieae (Teixeira et al. 1996). All of the streptococcal pathogens of tilapia are nonmotile, fermentative in glucose, catalase negative, non-sporeforming, Grampositive cocci (Table 16.1). Streptococcus agalactiae isolates are presumptively identified by positive agglutination with Lancefield typing antisera and a positive CAMP test. S. agalactiae isolates from fish can be placed in two groups: those that are betahemolytic and grow well at 37◦ C and those that are nonhemolytic and have a lower optimum incubation temperature and are biochemically less reactive. The Enterococcus spp. are members of Lancefield group D and are identified by hemolysis and biochemical characteristics: growth at 10 and 45◦ C, in 6.5% salt, in 40% bile, at pH 9.6, and positive bile esculin reaction (Kusuda et al. 1991; Murray

et al. 2007). L. garvieae possesses similar characteristics to Enterococcus spp. but does not belong to Lancefield group D or any other recognized group. Also direct fluorescent antibody can be used to identify S. iniae in culture and tissues of infected fish (Klesius et al. 2006). Most studies of S. iniae indicate little morphological or biochemical diversity in the species; therefore, all appear closely related regardless of habitat or fish origin. However, the question remains as to whether or not all S. iniae isolates, regardless of geographical location or environment from which they come, are the same strain. Kvitt and Colorne (2004) compared 26 Israeli isolates to nine non-Israeli isolates using PCR. While there are many molecular, antigenic, and biochemical similarities in most of these, they found that generally S. iniae afflicting fish in Israel originating in fresh and brackish water fell into homogeneous clusters that could be distinguished from marine (Mediterranean Sea or Red Sea) isolates. The isolates from the marine environment were a distinct cluster with few differences between the Israeli clusters and isolates from the United States, Canada, Australia, and Barbados. In addition, OlivaresFuster et al. (2008) reported that there are 13 genotypes of S. agalactiae from Kuwait, the United States, Brazil, and Honduras using amplified fragment length polymorphism fingerprinting. They concluded that there was good correlation between geographical origin and genotypes. Taken in whole, these reports indicate that there is biochemical and molecular diversity among the various streptococci isolated from fish. Olivares-Fuster et al. (2008) determined genetic variability among isolates of S. agalactiae and identified five distinct genotypes. All isolates from Kuwait were genotype 1 while the other four genotypes included species from the United States, Brazil, and Honduras. These molecular tests provided a good correlation between geographical origin and genotypes. Barnes et al. (2003) determined that serological differences

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Table 16.1 Characteristics of most frequently isolated Streptococcus spp. and Enterococcus sp. from fish. Test/Characteristic

S. iniae

S. faecalis

S. faecium

S. agalactiae

Group B

E. seriolicida

Cell morphology

Spherical envelope long chain −

Ovoid pairs or short chains −

Spherical pairs or short chains −

?

Spherical short chains

Ovoid

−

−

−

+ −

+ +

+ +

? −

− −

+

− ? − ? ? + ? + ? + + + − ? − −

+ ? + ? ? + ? ? ? + − + − + + ?

+ ? + + ? + + ? ? + ? − − + ? ?

d ? − ? ? + ? ? ? − − + − + + +

− ? − ? − + ? d ? ? − − − + + ?

− + + ? + − − + − +

? ? ? − ? − ? ? − + + None 32.9

? ? ? ? ? ? ? ? ? + ? D 33.5–38

? ? ? ? + ? ? ? − + − D 38.3–39

? ? ? d ? − ? ? − + ? B 34

? ? + − ? ? ? ? − + ? B ?

− − − − − − − − + + + Not D 44

Motility Growth at: 10◦ C 45 ◦ C Growth in: 6.5% NaCl 0.4% tellurite pH 9.6 Tetrazolium 0.1% methylene blue 40% bileArabinose Salicin Trehalose Esculin hydrolysis δ-Hemolysis β-Hemolysis H2 S Arginine hydrolysis Hippurate hydrolysis Voges–Proskauer Acid production from: D-xylose L-Rhamnose Sucrose Lactose Melibiose Raffinose Glycerol Adonitol Sorbitol Glucose Mannitol Lancefield grouping Mol% G + C DNA

+ − + + +

Source: Kitao (1982); Rotta (1984); Kusuda et al. (1991).

of S. iniae are due to the presence of capsule antigens in the arginine dehydrolase (+ or −) strains. Results suggested that cross-reactive antigens of the capsule are effectively hidden by the specific capsule, while they are partially exposed on the AD+ isolate. Two species of Streptococcus (S. shiloi and S. difficilis were reported to cause meningoencephalitis in tilapia, which resulted in great

economic losses in Israel during the late 1980s (Eldar et al. 1994).

Epizootiology Streptococcus spp. infections occur in both fresh- and saltwater grown tilapia. S. iniae is more common in freshwater systems and

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S. agalactiae more common in brackish water. Streptococcus salinity tolerance in vitro supports this phenomenon, but most isolates from freshwater fish do not grow at salinity above 30 ppt (Kitao et al. 1981). It appears that Streptococcus isolates adapt to salinity levels in various ecosystems, and that they are generally less virulent in an ecosystem that differs from their origin. Tilapia in water with 15–30 ppt salinity at 25◦ C and 30◦ C are more susceptible to Streptococcus isolated from fish in saltwater than when in freshwater at the same temperature (Chang and Plumb 1996). Little difference in susceptibility is noted at 20◦ C. In the United States and Japan, most epizootics occur during late summer through autumn (Kitao 1993). J. P. Hawke (personal communication) observed outbreaks of S. iniae in closed recirculating indoor farms when water temperatures are allowed to fluctuate. Mortality of tilapia infected with Streptococcus varies from low to high, depending on other circumstances. Under culture conditions, mortality is as high as 75% in naturally infected tilapia. In experimental infections with S. agalactiae in tilapia, mortality can reach as high as 90%, but mortality in natural infections is generally lower (Evans et al. 2002). In a study by Colorni et al. (2002) of S. iniae infections in Red Sea cage-cultured red drum and wild fish, it was found that cage-cultured fish were probably the source of pathogen for wild fish. They also suggest that because the area is essentially landlocked, the original source of S. iniae in the region was from imported seed fish for mariculture into Israel. In Japan, Streptococcus spp. remains in seawater and mud year-round near rearing facilities, but numbers of bacteria are elevated during the summer. However, according to Kitao et al. (1979), isolation of Streptococcus from mud was more consistent during autumn and winter. In naturally occurring infections, Streptococcus transmission is thought to be by contact or even cannibalism. Experimentally, transmission occurs by immersion in wa-

ter containing the pathogen, by injection, or cohabitation of infected fish with naive fish. Immersion transmission is enhanced by epithelium injury or stressful environmental conditions. Chang and Plumb (1996) had difficulty establishing Streptococcus spp. infections in Nile tilapia or channel catfish unless skin was scarified prior to exposure. Contrastingly, Ferguson et al. (1994) easily produced infection and high mortality in unstressed ornamental fish and rainbow trout that were exposed to Streptococcus spp. Tilapia susceptibility to Streptococcus is usually associated with environmental stress, skin injury, scale loss, and other factors associated with intensive aquaculture (Chang and Plumb 1996). Bowser et al. (1998) found S. iniae in Nile tilapia in a recirculation system in New York and demonstrated transmission of the pathogen from infected fish to naive fish in the culture units. However, they could not correlate the infection to any specific stressful condition. In a study of S. agalactiae in Columbia, the pathogen was found in intensive culture conditions of broodstock, grow out, and market size tilapia, but it was not isolated from wild fish (Hernandez et al. 2009). The ¨ protozoan Trichodina spp. on gills of tilapia also increases infection of Streptococcus (J. A. Plumb, unpublished). It was shown by Xu et al. (2007) that tilapia with a gill infestation by Gyrodactylus niloticus were significantly more susceptible to S. iniae infection than nonparasitized individuals. Furthermore, the bacterium could be isolated from up to 60% of the trematodes 24-hour postbacterial challenge. There is some question concerning the after effects of an S. iniae infection in tilapia. Shoemaker et al. (2006a) showed that tilapia survivors of an S. iniae infection without overt disease grew as well as control fish, which had not been exposed to the bacterium. These fish also developed acquired resistance (immunity) to subsequent exposure to S. iniae. The data indicate that tilapia survivors of an S. iniae epizootic were not impaired and were immune to subsequent exposure to the pathogen.

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Streptococcus spp. can occur with other opportunistic bacterial infections. Streptococcus spp. and A. sobria were implicated in mortality of adult Nile tilapia in The Philippines where about 1,200, 800, and 500 individuals died during successive weeks (Yambot 1996). Also dual infections of E. tarda and Streptococcus spp. have been noted in intensive tilapia culture systems. From late 1995 through mid-1996, reports surfaced of Streptococcus spp. infections being transmitted to humans from infected tilapia as a result of puncture wounds or cuts acquired while cleaning farm-cultured tilapia purchased live from a fish market (Anonymous 1996). The bacteria in these human infections were identified as S. iniae and caused major concern for the commercial tilapia industry, particularly in North America (Weinstein et al. 1997). However, there is little conclusive evidence that the organism isolated from affected humans was the same in all respects as isolates taken from fish. In fact, biochemical profiles differentiated human from fish isolates, suggesting that fish and human S. iniae isolates were genetically distinct from each other (Dodson et al. 1999). But DNA analysis and PCR are not sufficiently sensitive to differentiate between these streptococcus isolates. Moreover, because of a low prevalence of S. iniae in the flesh of healthy, commercially grown tilapia and hybrid striped bass, the pathogen poses only a limited risk for older or immunocompromised humans handling or cleaning fish (Shoemaker et al. 2001) unless of course the fish have overt infection.

Pathology Pathologic lesions appear to be similar with different streptococcus species even in different hosts. Group B Streptococcus spp. in Gulf killifish affects the spleen, liver, eye, and in some cases the kidneys, typical of general bacteremia (Rasheed and Plumb 1984; Rasheed et al. 1985). The exophthalmic eye

may display severe granulomatous inflammation (Miyazaki et al. 1984). Also infiltration of bacterial-laden macrophages and granulomas in infected lesions of the pericardium, peritoneum, stomach, intestine, brain, ovary, testes, and capsules of the liver and spleen were noted. Spleens of infected Gulf killifish were about ten times larger than normal and splenic pulp was severely congested. Some tissues were necrotic, and it appeared that phagocytosed bacteria multiplied in the cells. Livers of these fish had edema and sinusoids dilated while hepatocytes were atrophied and focally necrotic. The comparative pathology of S. iniae and S. agalactiae indicated that the gross pathology and histopathology in tilapia resulting from infection of both bacterial species are similar (Chen et al. 2007). Tilapia infected by injection displayed pericarditis, epicarditis, myocarditis, endocarditis, and meningitis. However, large numbers of bacteria were present in tissues and in the circulatory system of S. agalactiae-infected fish but not in those infected with S. iniae. It was also concluded that intraperitoneal injection of a relatively high dose of S. iniae reproduced large numbers of bacteria in lesions, thus suggesting that the pathogenesis of different streptococcal species in tilapia may be similar, but tilapia may combat natural S. iniae infections more effectively resulting in more chronic forms of the disease compared to S. agalactiae. Pathogenesis of Streptococcus spp. is thought to be facilitated by exotoxins. Kimura and Kusuda (1979) reported that when yellowtail were injected with a cell-free culture media in which Streptococcus spp. was grown, susceptibility increased upon subsequent exposure of the fish to the bacterium. The kidney and spleen contained higher numbers of bacteria than the blood, liver, or intestines. The route of experimental infection of S. iniae can be via the nares or the gills. Infection via the gills of hybrid striped bass resulted in the pathogen being isolated from optic and cerebellum regions of the brain, eye, head, trunk kidney, spleen, and liver 12–48 hours

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after exposure, depending on the infectious dose (McNulty et al. 2003); however, mortality of fish infected across the gill was lower than the mortality in hybrid striped bass resulting from infection via the nares (Evans et al. 2001). Whether or not this same route of infection is true for tilapia is unknown.

Significance Streptococcosis caused by S. iniae, S. agalactiae, and S. dysgalactiae has become a major disease of cultured tilapia in North America, South and Central America, Asia, and the Middle East and could be a limiting factor in the expansion and intensification of tilapia aquaculture. Shoemaker et al. (2001), in a survey of cultured tilapia and striped bass, found that the prevalence in tilapia was 3.81 and 7.23%, respectively. However, prevalence by farm was 27.4% for tilapia and 21.6% for striped bass. In the same survey, the prevalence in market size and nursery tilapia was higher than in grow-out size fish. Its potential for human infections, although limited and uncertain, is worthy of concern as noted earlier; however, Shoemaker et al. (2001) indicated that the potential for fish-born streptococci infections in humans is limited unless those handling infected fish have open wounds on their hands, are older adults, or are immunocompromised.

Francisellosis An RLO, a relatively new and emergent pathogen of fish, was first detected in diseased Nile tilapia in Taiwan (Chen et al. 1994). This intracellular bacterium from tilapia and a limited number of other warm-water fish species were initially referred to as a piscirickettsialike organism (PLO) or Francisella-like organism (FLO) and was believed to be nonculturable on cell-free media. Isolates from Taiwan were cultured by tissue culture in CHSE-214 cells, and the DNA recovered from the cultured bacteria was identified by comparing

nucleotide sequences of whole 16S rRNA gene to reference organisms. The results of these studies indicated the bacterium belonged to the genus Francisella (Hsieh et al. 2006). The organism from hybrid striped bass and tilapia was identified by isolation of bacterial DNA from tissue of diseased fish and amplification of 16S rRNA gene and comparing the sequences to a comparative database (GenBank) (Ostland et al. 2006; Mauel et al. 2007). The bacterium was later cultured on agar media and subjected to additional molecular studies where it was determined that warm-water strains of Francisella from tilapia were distinct from cold-water isolates from cod (Soto et al. 2009). A detailed taxonomic study concluded that the two organisms belong to two different subspecies. The warm-water strains from tilapia were identified as F. noatunensis subsp. orientalis and the cold-water strains as F. noatunensis subsp. noatunensis (Ottem et al. 2009). The two strains were given species status with the warm-water isolates forming F. asiatica and the cold-water strains F. noatunensis (Mikalsen and Colquhoun 2009).

Geographical range and species susceptibility Members of the genus Francisella affect a wide range of animals including humans (F. tularenses), and francisellosis has become a serious health problem for cultured tilapia in many geographical regions. The species of Francisella associated with disease in warmwater fish, F. asiatica, has been found in Taiwan, the United States (Hawaii and the continental USA), and Latin America (Costa Rica) (Chen et al. 1994; Mauel et al. 2007; Soto et al. 2009), where it affects both freshwater and saltwater fish species. The pathogen has been implicated in disease of several species of tilapia (O. mossambicus, O. niloticus, and Sarotherodon melanotheron) (Mauel et al. 2003). Francisella spp. has also been

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found in three-line grunt (Fukuda et al. 2002) and grouper (Chen et al. 2000). F. noatunensis affects cold-water fish species such as the Atlantic cod (Nyland et al. 2006). These reports indicate a wide geographical range and fish species susceptibility to Francisella spp.

Clinical signs and findings Francisellosis may be either an acute, subacute, or chronic disease, depending on culture conditions and water temperatures. Affected fish are dark, swim lethargically and erratically, and have a loss of appetite; they display petechiae or hemorrhages and ulcers on the skin; they have exophthalmia and varying degrees of ascites; the spleen and kidneys are enlarged and contain distinct white nodules of varying sizes (Chen et al. 1994; Mauel et al. 2007; Soto et al. 2009). The gills exhibit epithelial hyperplasia with multifocal consolidation of secondary lamellae. Also multiple white granulomas occur in the gills, spleen, kidney, choroid gland, and testes but seldom in the liver; occasionally black granulomas are seen internally. It should be pointed out that some fish may show no clinical signs (Mauel et al. 2003).

Diagnosis The fish pathogenic Francisella were slow to be characterized due to the fastidious nature of the bacteria and the resulting difficulties in culturing the organism from fish tissues. In addition to the previously mentioned clinical signs, Francisella spp. can be diagnosed by detection of tiny Gram-negative, intracellular, coccobacilli in inclusions, cytoplasmic vacuoles, or free in the cytoplasm of host cells (Mauel et al. 2003; Mauel et al. 2007; Soto et al. 2009). Bacterial cells are also visible in Giemsa-stained blood smears or spleen imprints. Francisella spp. cannot be cultured on general bacterial media. However, Soto

et al. (2009) found that cystine heart agar supplemented with bovine hemoglobin solution (CHH) alone or as a selective medium containing the antibiotics colistin and ampicillin (SCHH) are useful for the primary isolation of F. asiatica from spleen and kidney of infected fish. Growth on CHH media can be seen in 36–48 hours at which time grey, smooth, and convex colonies are visible at optimum incubation temperature of 28–30◦ C; however, at 28◦ C visible growth may require 4 days. The bacterium can also be cultured in cation-adjusted Mueller Hinton II broth supplemented with 2% IsoVitalex and 0.1% glucose. The pathogen has been isolated in CHSE214 cell cultures from which subculture and isolation on bacteriological media can be accomplished. A quantitative real-time PCR was developed for diagnosis of F. asiatica by Soto et al. (2009).

Bacterial characteristics The Gram-negative F. asiatica is a nonmotile, coccobacillus that measures about 0.56 × 0.7 µm with a double cell wall. Initial work by Hsieh et al. (2006) showed ten strains of RLOs isolated from tilapia in Taiwan had nucleotide sequences for whole 16S rRNA gene that are highly similar to reference strains of Francisella spp.; Mauel et al. (2007) confirmed these molecular similarities.

Epizootiology Francisellosis affects all ages and sizes of tilapia from small fingerlings to adults. Mortality of tilapia infected with Francisella spp. ranged from 5 to 80% with an average of 50% in Nile tilapia in Latin America (Mauel et al. 2007). The disease has a propensity to occur during cooler months of the year, and infections are stimulated by cold stress on farms in which it is endemic (Mauel et al. 2003). In temperature studies, tilapia maintained

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between 21.5 and 26.5◦ C, initial mortalities occurred on day 15 and mortality doubled almost daily thereafter (Mauel et al. 2003). Tilapia maintained between 26.5 and 29.5◦ C showed no sign of disease or mortality. Mauel et al. (2005) also described a PLO in cultured tilapia in continental United States, where it was associated with cold stress and poor water quality. The disease in Hawaii occurred during the cooler months of October to April and did not appear in warmer months. Koch’s postulates were fulfilled with Francisella by Soto et al. (2009) by exposing tilapia via intraperitoneal injection, gill exposure, and immersion. Injection resulted in 100% mortality in 72 hours. Fish exposed by gill immersion exhibited 80% mortality but occurred more gradually over a 10-day period. The LD50 was calculated by immersion exposure and injection was 1152 CFU/fish by injection and 2.3 × 107 CFU/ml by immersion (Soto et al. 2009) Francisella is also transmissible horizontally from infected fish to naive fish by cohabitation (Chen et al. 1994; Mauel et al. 2003). The potential for vertical transmission is unknown.

Pathology Francisella causes acute to chronic disease in tilapia cultured under certain environmental conditions. Fish infected with Francisella show nonspecific clinical signs such as lethargy, erratic swimming behavior, anorexia, anemia, and exophthalmia. Gross pathology of tilapia indicates that the gills and most internal organs are affected by Francisella spp. Microscopic evaluation of thin sections of tissue reveal granulomatous lesions in the gills, spleen, and kidneys with some pathologic changes present in the liver, heart, eye, central nervous system, and gastrointestinal tract (Soto et al. 2009). Most organs contain granulomatous inflammation, and distinct granulomas are usually visible in the spleen, head, and trunk kidney. The gills exhibit primary and secondary

lamellar fusion because of epithelial hyperplasia. The pericardium and myocardium display widespread cellular infiltration and presence of granulomas in severe cases. Special stains such as Giemsa revealed small, pleomorphic coccobacilli inside and outside the cells.

Significance Although Francisella spp. is a relatively recently recognized disease in fish, its impact has been significant in tilapia aquaculture. The effect on susceptible populations under temperature stress is high, and this disease poses a potentially serious health problem in the future.

Other bacterial tilapia diseases In tilapia, MAS is associated with several different species of bacteria, the most common of which is Aeromonas hydrophila (liquifaciens, punctata); however, A. sobria and other related species do occasionally occur. Other opportunistic pathogens of tilapia are P. shigelloides, Edwardsiella tarda, Flavobacterium columnare, Photobacterium damselae, Vibrio spp., and Pseudomonas spp. Mycobacteria have been reported on rare occasions. Although MAS is not uncommon in cultured tilapia, there are few published “case reports” of its occurrence, but when it occurs, it is often a secondary problem. Tilapia with MAS lose their equilibrium, swim lethargically, gasp at the surface, and generally display the same clinical signs as other fish species (Figure 16.2). A. hydrophila was shown to cause “eye disease” in cage-cultured Nile tilapia in The Philippines (Yambot and Inglis 1994). Pseudomonas fluorescens occasionally produces clinical signs and pathology similar to A. hydrophila and can cause significant mortality in tilapia (Duremdez and Lio-Po 1985). In Japan, this bacterium caused a disease in Nile tilapia characterized by hemorrhagic

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Figure 16.2 Motile Aeromonas septicemia (Aeromonas hydrophila) in tilapia. Note the white lesions from which scales have fallen.

lesions in gonads and ovaries (Miyashita 1984). The infection occurred during winter and spring when water temperatures were 15–20◦ C resulting in mortalities of 0.2–0.3% per day. Plesiomonas shigelloides is becoming more frequently reported as an opportunistic fish pathogen, but little is known about its pathological capability. However, Faisal and Popp (1987) reported that P. shigelloides was responsible for 30–60% mortality among overcrowded 4-week-old Nile tilapia. The disease was transmitted only by injection to 6-weekold tilapia but not to 9-month-old fish. Epizootiology of vibriosis in tilapia is similar to that of MAS in the respect that both diseases are usually secondary infections (Sakata 1988). In saltwater, V. anguillarum or V. vulnificus are involved, while in freshwater, V. mimicus or V. cholerae are found, and V. parahaemolyticus can occur in either environment. Vibriosis in tilapia is often mild to chronic, and clinical signs do not differ significantly from those for MAS. Mortality of infected tilapia is usually chronic with relatively low daily losses, but cumulative mortality can be significant. Hubert (1989) stated that V. parahaemolyticus infections were not spontaneous, but were fulminating septicemias that occurred in market-size

tilapia 2–3 days following handling and transfer to wintering ponds. Sakata (1988) reported that Nile tilapia suffered 10–20% mortality due to a vibriosis infection following transfer from freshwater to saltwater pens at 18–20◦ C. Decreasing water temperatures, coupled with high salinity, are considered compounding stressors on tilapia populations. Nonspecies-specific columnaris disease has not been frequently reported in cultured tilapia, even though the pathogen is ubiquitous in freshwater. When columnaris occurs in tilapia, pale areas form on the body and frayed fins (Figure 16.3) are the most frequently observed clinical sign of disease, and infected fish will swim lethargically or float at the surface. Primary cause of death is attributed to injury of skin, fins, and gills. Any physical injury or environmentally induced stress can precipitate these infections. Marzouk and Bakeer (1991) showed that Nile tilapia are more susceptible to columnaris infections and a higher mortality when pH is either very acidic or alkaline. Amin et al. (1988) isolated F. columnare from cultured Nile tilapia, which had gill lesions. In this study, pathogenicity varied between seven different isolates, but infection could not be established with any isolate unless fish were stressed, had skin injury, and/or the water contained ammonia.

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Figure 16.3 Columnaris (Flavobacterium columnare) infection on tilapia. Note the pale area (excess mucus) on the lateral side and the frayed fins.

Edwardsiellosis (E. tarda) affects tilapia cultured under high density and other stressful conditions in either freshwater or marine environments. Tilapia infected with E. tarda swim lethargically on their sides at the surface; have an enlarged abdominal area, swollen, opaque, and hemorrhaged eyes (Figure 16.1); and possibly some discrete inflammation or skin discoloration. Internally, focal areas of necrosis are seen, hemorrhage and gas-filled cavities occur in the muscle, bloody fluid accumulates in the body cavity, the liver is often pale and mottled, spleen is dark red and swollen, kidney swollen and soft, and intestine is inflamed and usually void of food. In Japan, an E. tarda infection was reported in Nile tilapia with mortalities of 0.2–0.3% per day when water temperature was 20–30◦ C (Miyashita 1984). Infected fish had small white nodules in the spleen and abscesses in the swim bladder. Some E. tarda-infected tilapia display severe exophthalmia and eye opaqueness. E. tarda occurs most often in tilapia in intensive culture systems with marginal water quality, high organic load, or high fish density; it can occur in conjunction with A. hydrophila or Streptococcus spp. In Egypt, an E. tarda infection was identified in Nile tilapia reared in ponds that received domestic waste water (Badran 1993). Photobacterium damselae subsp. damselae was shown to cause disease in freshwater wild and cultured fish in Egypt including Nile

tilapia (Khalil and Aly 2008). Infected tilapia swam lethargically and had dark skin. Internally they had swollen kidney, spleen, and liver with bloody fluid in the abdominal cavity. Also pinhead nodules were present on the liver, spleen, and kidney. Mortality of experimentally infected fish was 20–40%. Mycobacteriosis (fish tuberculosis) (Chapter 16) was detected in tilapia in Central Africa (Roberts and Sommerville 1982). This pathogen presents problems in tilapia populations similar to those in intensively reared striped bass.

Health maintenance of tilapia As tilapia culture has become more intensive, the need for environmental control, water quality stability through management, stress reduction, and other sound health maintenance procedures are more essential. Three approaches to health maintenance include management, chemotherapy, and vaccination.

Management When starting a tilapia culture operation, great care should be taken to make certain that the facility is “clean” of wild fish that can harbor pathogens, although in some instances,

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i.e., MAS, this is not feasible. Sound management procedures should be implemented to regularly eliminate accumulation of detritus, waste, and dead fish. When fish are introduced, only specific pathogen-free stocks should be used if available. To reduce potential for catastrophic disease outbreaks, water quality should be maintained at the highest possible level and reduction of environmentally induced stress is a top priority. Prudent stocking densities, adequate water exchange in intensive culture units, removal of metabolites and fecal waste from recirculating water, supplemental aeration, and feeding high-quality diets are a means to that end. Although manufactured tilapia feed does not generally contain raw fish products, diets of other fish species sometimes do and should be avoided because it is possible to contract Streptococcus spp., Mycobacterium spp., or other pathogenic bacteria (Minami 1979). If disease occurs, infected fish and dead carcasses should be promptly removed. Prophylactic chemotherapy such as salt (NaCl or CaCl2 ) or potassium permanganate, during or after handling, aids in healing minor skin abrasions and reduces external parasites and the possibility of contracting secondary bacterial infections. Disinfection of water with ultraviolet (UV) light and ozone will help reduce bacterial populations in recirculating water or open water supplies. Sanitation by routine sterilization of nets, buckets, and other utensils reduces accidental cross contamination of culture units. Sterilization is accomplished by dipping utensils and boots into 200 mg/L of chlorine or in 100 mg/L of a quaternary ammonium compound (Roccal). Iodine at 1,000 mg/L is a disinfectant that is safer than chlorine or the quaternary ammonia compounds. Thoroughly rinsing nets and utensils in freshwater is essential before disinfected items come into contact with fish. Nets, seines, etc., should be thoroughly dried, preferably in the sun to kill most pathogens. Antidotal data indicate that reducing fish density, changing handling practices, improving water quality, maintaining

stable water temperatures in the optimal range, eliminating gill parasites, and adopting other health maintenance procedures reduce effects of Streptococcus spp. infections. Also maintaining broodstock on the premises will provide seed fish with a known disease history.

Chemotherapeutics Currently there are no chemotherapeutics with approved labels for treating any tilapia disease in the United States; however, it is encouraging that some chemicals and drugs are being considered and may be approved by the FDA for use on cultured tilapia in the future. Although not FDA approved, some drugs and chemicals have been successfully used prophylactically or in chemotherapy for bacterial infections of tilapia (Table 4.2). Prophylaxis includes salt baths (NaCl or CaCl2 at 0.5–3%) for dip or prolonged treatments and/or potassium permanganate (5–10 mg/L) for 1 hour or 2–5 mg/L indefinitely. The concentration of KmNO4 depends on water quality. Systemic clinical bacterial infections are usually treated with medicated feed, but any unapproved antibiotic require an “extra-label use” (AMDUCA) permit, which in some instances can be provided by a veterinarian. Terramycin (oxytetracycline) incorporated into feed to provide 2.5–3.5 g/45 kg (50–75 mg/kg) of fish per day for 14 days is effective in treating most systemic infections. R Sulfadimethoxine-ormetoprim (Romet -30) fed at a rate of 2.5–3.5 g/45 kg (50–75 mg/kg) of fish per day for 5 days is also effective against most systemic bacterial infections but may present a palatability or toxicity problem. Erythromycin is effective against Gram-positive bacteria, and its use is currently under FDA consideration for use in Streptococcus spp.-infected tilapia. If approved, medication level would probably be 50 or 100 mg/kg of fish per day for 10 days. Oxytetracycline fed to S. iniae-infected blue tilapia at rates of 75 and 100 mg per kg of fish per day for

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14 days increased survival to 85 and 96%, respectively, compared to 7% survival for nonmedicated control fish (Darwish et al. 2002). Survivors of 100-mg treated fish were not carriers of the pathogen after treatment but control fish were. Experimentally fed amoxicillin successfully controlled S. iniae in blue tilapia (Darwish and Hobbs 2005). The drug was fed at 0, 5, 10, 30, and 80 mg/kg of fish per day for 12 days and then challenged by waterborne exposure to the pathogen 22–24 hours post final drug feeding. Survival in nonmedicated controls, and 10, 30, and 80 mg/kg were 3.8, 45, 75, and 94%, respectively. At the conclusion of the study, the bacterium was recovered from nonmedicated challenged fish but not the medicated fish. When bacterial infections are confined to the skin (columnaris or external motile Aeromonas infections), potassium permanganate at previously noted rates is effective. Aquaflor (florfenicol) is being evaluated for a feed additive treatment of streptococcal infections in tilapia at 10–15 mg/kg/day (Gaunt 2004). Either whole leaf or extract from the plant Rosmarinus officianalis (rosemary) has a therapeutic effect when incorporated into feed of tilapia infected with S. iniae or S. agalactiae (Abutbul et al. 2004). Initially the extract was bacteriostatic in in vitro laboratory tests. Dried leaves (whole or ground) or the dried acetate extract in the feed in a ratio of 1:17 or 1:24, respectively, significantly reduced mortality rate following infection with S. iniae. Ground rosemary leaves in feed is most practical because of easy preparation (D. Zilberg, unpublished). Oxytetracycline in the feed successfully reduced mortality in tilapia infected with Francisella (Mauel et al. 2003).

Vaccination Few immunological or vaccination experiments involved tilapia until the last decade when significant strides were made in vaccinating them against S. iniae and S. agalac-

tiae (Klesius et al. 2008). In Nile tilapia, protective immunity to A. hydrophila was demonstrated by intraperitoneal injection with vaccine containing killed bacteria (Rungpan et al. 1986). They reported 53–61% protection within 1 week postvaccination and 100% protection in 2 weeks. The first vaccines used experimentally to prevent streptococcus infections in tilapia were in Japan (Sakai et al. 1987), but since then, numerous vaccination studies have been carried out. A formalinkilled vaccine of two S. iniae isolates (ARS10 and ARS-60), injected IM and IP individually and in combination, protected against the respective antigens (Klesius et al. 2000). Protection was best demonstrated when vaccinates challenged with the heterologous antigen; an RPS of 93.7% compared to nonvaccinated controls (RPS-17.7%) was achieved. Using OraljectTM (PerOs Systems Technology, Inc.), Shoemaker et al. (2006a), using modified and lyophilized S. iniae incorporated into feed and applied to tilapia orally and by injection, protected the fish against challenge. Both provided significant protection; however, injection was more protective (RPS 100%) compared to an RPS 63% resulting from fed vaccine. Furthermore, tilapia survivors of an S. iniae infection are significantly more immune to subsequent challenge with the homologous antigen (Shoemaker et al. 2006b). Also their studies suggest that tilapia surviving S. iniae challenge without overt disease signs performed as well as noninfected tilapia and the challenge survivors developed acquired resistance to S. iniae as shown in subsequent challenges. Vaccination by IP injection with S. agalactiae elicited a high degree of protection with an 83.4% RPS, and the fish had only mild short-term stress (Evans et al. 2004a). Studies have shown promise in application for protecting tilapia against S. agalactiae (Pasnik et al. 2005a) in which a vaccine of injected intracellular products stimulated antibody response conferring a significant degree of protection to a homologous challenge. In a study to

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determine whether or not tilapia vaccinated against S. agalactiae were protected against S. iniae, it was found that they were not (Evans et al. 2004b). The duration of vaccinating Nile tilapia using extracellular product fraction and formalin-killed S. agalactiae showed that survival was 67, 62, and 49% at 47, 90, and 180 days postvaccination, respectively (Pasnik et al. 2005b). These survivals compared to 16, 16, and 4% for the three challenge periods in nonvaccinated control fish. Vaccination of tilapia is not confined to Streptococcus spp. Kwon et al. (2006) showed that O. mossambicus positively responded to injection with a formalin-killed E. tarda as well as E. tarda ghosts (envelope from nonliving whole cell) preparations. Vaccinated fish had elevated E. tarda agglutinating antibody from both preparations, but upon challenge, the E. tarda ghost preparation resulted in significantly higher survival than the formalin-killed preparation and controls. These data as a whole indicate that vaccines against S. iniae, S. agalactiae, or E. tarda provide a feasible and effective proactive approach to prevent serious infections in tilapia by these pathogenic bacteria. An experimental live attenuated vaccine, utilizing an attenuated mutant of F. notunensis subsp. orientalis, has shown efficacy in trials using the immersion challenge model (Hawke and Soto et al. manuscript in preparation), and this type of health management procedure should be established to avoid dependence on antibiotics.

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yellowtail (Seriola guingueradiata) – Distribution of Streptococcus sp. in seawater and muds around yellowtail farms. Bulletin of the Japanese Society of Scientific Fisheries 45:567–572. Kitao, T., T. Aoki, and R. Sakoh. 1981. Epizootic caused by ß-haemolytic Streptococcus species in cultured freshwater fish. Fish Pathology 15:301–307. Klesius, P. H., J. J. Evans, et al. 2006. Rapid detection and identification of Streptococcus iniae using a monoclonal antibody based indirect fluorescent antibody technique. Aquaculture 258:180–186. Klesius, P. H., C. A. Shoemaker, and J. J. Evans. 2000. Efficacy of single and combined Streptococcus iniae isolate vaccine administered by intraperitoneal and intramuscular routes in tilapia (Oreochromis niloticus). Aquaculture 188:2378–246. Klesius, P. H., C. A. Shoemaker, and J. J. Evans. 2008. Streptococcus: A worldwide fish health problem. International Symposium on Tilapia Aquaculture, Cairo, 1:83–107. Kusuda, R., K. Kawai, et al. 1991. Enterococcus seriolicida sp. Nov., a fish pathogen. International Journal of Systematic Bacteriology 41:406–409. Kusuda, R. and F. Salati. 1993. Streptococcal disease problems and control: a review. Annual Review of Fish Diseases. Oxford, Pergamon Press, pp. 69–85. Kvitt, H., and A. Colorni. 2004. Strain variation and geographic endemism in Streptococcus iniae. Diseases of Aquatic Organisms 61:67–73. Kwon, S. R., Y. K. Nam, et al. 2006. Protection of tilapia (Oreochromis mosambicus) from edwardsiellosis by vaccination with Edwardsiella ghosts. Fish and Shellfish Immunology. 20(4):621–625. Marzouk, M. S. M., and A. Bakeer. 1991. Effect of water pH on Flexibacter columnaris infection in Nile tilapia. Egyptian Journal of Comparative Pathology and Clinical Pathology 4:227–235. Mauel, M. J., D. L. Miller, et al. 2003. Characterization of a piscirickettsiosis-like disease in Hawaiian tilapia. Diseases of Aquatic Organisms 53:249–255. Mauel, M. J., E. Soto, et al. 2007. A piscirickettsiosis-like syndrome in cultured Nile tilapia in Latin America with Francisella spp. as the pathogenic agent. Journal of Aquatic Animal Health 19:27–34.

Mauel, M. J., D. L. Miller, et al. 2005. Occurrence of piscirickettsiosis-like syndrome in tilapia in continental United States. Journal of Veterinary Diagnostic Investigation 17:601–605. Mauel, M. J., E. Soto, et al. 2007. A piscirickettsiosis-like syndrome in cultured Nile tilapia in Latin America with Fracisella spp. as the pathogenic agent. Journal of Aquatic Animal Health 19:27–34. McNulty, S. T., P. H. Klesius, et al. 2003. Streptococcus iniae infection and tissue distribution in hybrid striped bass (Morone chrysops X Morone saxatilis) following inoculation of the gills. Aquaculture 220:165–173. Mikalsen, J., and D.J. Colquhoun. 2009. Francisella asiatica sp. nov. isolated from farmed tilapia (Oreochromis sp.) and elevation of Francisella philomiragia subsp. noatunensis to species rank as Francisella noatunensis comb. nov., sp. nov. International Journal of Systematic and Evolutionary Microbiology Epub ahead of print. Minami, T. 1979. Streptococcus sp., pathogenic to cultured yellowtail, isolated from fishes for diets. Fish Pathology 14:15–19. Miyashita, T. 1984. Pseudomonas fluorescens and Edwardsiella tarda isolated from diseased tilapia. Fish Pathology 19:45–50. Miyazaki, T., S. S. Kubota, et al. 1984. A histopathological study of streptococcal disease in tilapia. Fish Pathology 19:167–172. Murray, P. R., E. J. Baron, et al. 2007. Manual of clinical microbiology, Ninth Edition. Washington, D.C., American Society of Microbiology Press. Nomoto, R., L. I. Munasinghe, et al. 2004. Lancefield group C Streptococcus dysgalactiae infection responsible for fish mortalities in Japan. Journal of Fish Diseases 27:679–686. Nyland, A., K. F. Ottem, et al. 2006. Francisella sp. (Family Francisellaceae) causing mortality in Norweigian cod (Gadus morhua) farming. Archives in Microbiology 185:383–392. Olivares-Fuster, O., P. H. Klesius, et al. 2008. Molecular typing of Streptococcus agalactiae isolated from fish. Journal of Fish Diseases 31:277–283. Ostland, V. E., J. A. Stannard, et al. 2006. Aquatic Francisella-like bacterium associated with mortality of intensively cultured hybrid striped bass

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Morone chrysops X M. saxatilis. Diseases of Aquatic Organisms 72:135–145. Ottem, K. F., A. Nylund, et al. 2009. Elevation of Francisella philomiragia subsp. noatunensis Mikalsen, et al. (2007) to Francisella noatunensis comb. nov. [syn. Francisella piscicida Ottem, et al. 2008 syn. nov.] and characterization of Francisella noatunensis subsp. orientalis subsp. nov., two important fish pathogens. Journal of Applied Microbiology 106:1231–1243. Pasnik, D. J., J. J. Evans, et al. 2005a. Antigenicity of Streptococcus agalactiae extracellular products and vaccine efficacy. Journal of Fish Diseases 28:205–212. Pasnik, D. J., J. J. Evans, and P. H. Klesius. 2005b. Duration of protective antibodies and correlation with survival in Nile tilapia Oreochromis niloticus following Streptococcus agalactiae vaccination. Diseases of Aquatic Organisms 66:129–134. Perera, R. P., S. K. Johnson, et al. 1994. Streptococcus iniae associated with mortality of Tilapia nilotica x T. aurea hybrids. Journal of Aquatic Animal Health 6:335–340. Plumb, J. A., J. H. Schachte, et al. 1974. Streptococcus sp. from marine fishes along the Alabama and northwest Florida coast of the Gulf of Mexico. Transactions of the American Fisheries Society 103:358–361. Rasheed, V. M., and J. A. Plumb. 1984. Pathogenicity of a non-haemolytic group B Streptococcus sp. in Gulf killifish (Fundulus grandis Baird and Girard). Aquaculture 37:97–105. Rasheed, V. M., C, Limsuwan, and J. A. Plumb. 1985. Histopathology of bullminnows, Fundulus grandis Baird and Girard, infected with a non-haemolytic group B Streptococcus sp. Journal of Fish Diseases 8:65–74. Roberts, R. J., and C. Sommerville. 1982. Diseases of tilapia. In: S. V. Pullin, and R. H. Lowe-McConnel (eds) The biology and Culture of Tilapias. . Manila, Philippines, International Center for Living Aquatic Resource Management, pp. 246–263. Rotta, J. 1984. Pyogenic hemolytic streptococci. In Bergey’s Manual of Systematic Bacteriology, vol. 2. Edited by P. H. A. Sneath, N. S. Mair, M. E. Sharpe, and J. G. Holt. Baltimore, MD: Williamd & Wilkins, pp. 1047–1065. Rungpan, L., T. Kitao, and Y. Yoshida. 1986. Protective efficacy of Aeromonas hydrophila vac-

cines in Nile tilapia. Veterinary Immunology and Immunopathology 12:345–350. Sakai, M., R. Kubota, et al. 1987. Vaccination of rainbow trout, Salmo gairdneri against beta-hemolytic streptococcal disease. Bulletin of the Japanese Society of Scientific Fisheries 53:1373–1376. Sakata, T. 1988. Characteristics of Vibrio vulnificus isolated from diseased tilapia (Saratherodon niloticus). Fish Pathology 23:33–40. Salvador, R., E. E. Muller, et al. 2005. Isolation and characterization of Streptococcus spp. group B in Nile tilapias (Oreochromis niloticus) reared in hopas net and earth nurseries in northern region of Parena State, Brazil. Santa Maria, Cinicia Rural, 35(6):1374–1378. Shoemaker, C. A., P. H. Klesius, and J. J. Evans. 2001. Prevalence of Streptococcus iniae in tilapia, hybrid striped bass, and channel catfish on commercial fish farms in the United States. American Journal of Veterinary Research 62:174–177. Shoemaker, C. A., L. Chorn, et al. 2006a. Growth response and acquired resistance of Nile tilapia, Oreochromis niloticus (L.) that survived Streptococcus iniae infection. Aquaculture Research 37:1238–1245. Shoemaker, C. A., S. W. Vandenberg, et al. 2006b. Efficacy of a Streptococcus iniae modified bacterin delivered using OraljectTM technology in Nile tilapia (Oreochromis niloticus). Aquaculture 255:151–156. Soto, E., J. P. Hawke, et al. 2009. Francisella sp., an emerging pathogen of tilapia, Oreochromis niloticus (L.) in Costa Rica. Journal of Fish Diseases 32:713–722. Teixeira, L.M., V.L.C. Merquior, et al. 1996. Phenotypic and genotypic characterization of atypical Lactococcus garvieae strains isolated from water buffalos with subclinical mastitis and confirmation of L. garvieae as a senior subjective synonym of Enterococcus seriolicida. International Journal of Systematic Bacteriology 46:664–668. Vandamme, P., L. A. Devriese, et al. 1997. Streptococcus difficile is a nonhemolytic Group B type Ib Streptococcus. International Journal of Systematic Bacteriology 47:81–85. Weinstein, M.R., M. Litt, et al. 1997. Invasive infections due to a fish pathogen, Streptococcus

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iniae. The New England Journal of Medicine 337(9):589–594. Wu, S. Y. 1970. New bacterial disease of Tilapia. FAO Fish Culture Bulletin 2:14. Xu, D-H., C. A. Shoemaker, and P. H. Klesius. 2007. Evaluation of the link between gyrodcactylosis and streptococcosis of Nile tilapia, Oreochromis niloticus (L.). Journal of Fish Diseases 30:233–238. Yambot, P. 1996. Streptococcus spp. and/or Aeromonas spp. associated with fish kill in the Nile tilapia (Oreochromis niloticus) breeders in the Philippines. World Aquaculture 96, Bangkok, Thailand (Abstract).

Yambot, A. V., and V. Inglis. 1994. Aeromonas hydrophila isolated from Nile tilapia (Orechromis niloticus L.) with “eye disease”. International Symposium on Aquatic Animal Health, Fish Health Section/American Fisheries Society, Seattle, Washington (Abstract). Zilberg, D. (Unpublished). Use of rosemary (Rosmarinus officianalis) as a treatment against Streptococcosis in tilapia (Oreochromis sp.). Department of Dryland Biotechnologies, The Jacob Blaustein Institutes for Desert Research, BenGurion University of the Negev, Sede Boger Campus, Israel.

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Chapter 17

Other bacterial diseases

Some bacterial fish diseases cannot be conveniently categorized according to host because they do not affect one fish species or group more than another. Arguably some of the bacterial diseases previously associated with certain groups of fishes may fall into this category but one that clearly falls into the “miscellaneous” category was formerly called “marine flexibacteriosis” caused by Flexibacter maritimus (Wakabayashi et al. 1986). The etiology is now named Tenacibaculum maritimum (Suzuki et al. 2001; Avenano˜ Herrera 2009) and the disease is referred to as “tenacibaculosis.”

Tenacibaculosis The marine fish disease tenacibaculosis, caused by Tenacibaculum maritimum (formerly Flexibacter maritimus), depending on location and species of fish infected, is also referred to as marine flexibacteriosis, saltwater columnaris, black patch necrosis (in Dover sole in Europe), or eroded mouth syndrome (Bernardet et al. 1990; Santos et al. 1999). Health Maintenance and Principal Microbial Diseases of Cultured Fishes, 3rd edition. By J. A. Plumb and L. A. Hanson. Published 2011 by Blackwell Publishing Ltd.

Tenacibaculum maritimum occurs exclusively in the marine environment in cultured and wild fishes. Labrie et al. (2005) established that T. maritimum is a primary pathogen by experimental transmission.

Geographical range and species susceptibility Tenacibaculum maritimum infects marine fish in Japan, the Pacific coast of North America, the United Kingdom, Spain, Italy, and other European countries, Japan, Australia (Tasmania), and several Asian countries (Santos et al. 1999; Labrie et al. 2005; Lopez et al. 2009). A ´ variety of susceptible fish species include white seabass, Pacific sardine, northern anchovy, Atlantic salmon, chinook salmon, rainbow trout, turbot, seabream, sea bass, several species of flounder, several species of sole, and striped trumpeter (Bernardet et al. 1990; Pazos et al. 1993; Frelier et al. 1994; Chen et al. 1995; Magarinos ˜ et al. 1995; Handlinger et al. 1997; Salati et al. 2005; Lopez et al. 2009). ´ However, it has been noted that many other species of marine fishes could be susceptible to T. maritimum. The disease has also been reported on larval shrimp in Brazil (Mourino ˜ et al. 2008) but whether the two pathogens are identical is unclear (Avendano-Herrera 2009). ˜ 465

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Clinical signs

Diagnosis

Affected fish become lethargic, white to pinkish ulcers develop on the skin, exposing necrotic underlying muscle, and variable scale loss (Figure 17.1). Fins become frayed and white necrotic lesions form on the gills (Chen et al. 1995; Salati et al. 2005). Clinical signs in Atlantic salmon, rainbow trout, and greenback flounder are consistent among the species with eroded skin lesions being the most prominent sign (Handlinger et al. 1997). No internal clinical signs have been reported.

Marine tenacibaculosis is diagnosed by detection of long, thin rod shaped flexing bacteria in wet mount scrapings from body, fin, or gill lesions at 400× and presence of long Gramnegative cells in stained smears. Phase contrast microscopy is advantageous in detecting the bacteria. Tenacibaculum maritimum is not easily isolated on conventional high nutrient bacterial media but can be isolated on low nutrient agar that contains sea salts (Labrie et al. 2005).

(a)

(c)

(b)

Figure 17.1 Marine Tenacibaculosis (Tenacibaculum maritimum). (a) Chinook salmon with a filamenous bacterial (T. maritimum) induced lesion on the gill (arrow). (b) Northern anchovy with hemorrhagic and necrotic lesions of T. maritimum on the body (arrows). (c) White seabass with ulcerative body lesions caused by T. maritimum. (Photos courtesy of M. Chen, California Department of Fish and Game.)

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Bacterial characteristics Tenacibaculum maritimum are Gram-negative bacteria ranging in size from 0.5 µm in diameter to 2–30 µm in length (Table 17.1). Tenacibaculum maritimum grow on Ordal Table 17.1 Biophysical and biochemical characteristics∗ of Tenacibaclum maritimum.

Characteristic

Cell morphology Temperature range Optimum temperature Growth in 0% SW-HS† 33% SW-HS 66% SW-HS 100% SW-HS Cell size (µm) Colony color Colony morphology Motility Flexirubin pigment Binds Congo red Resistant to neomycin sulfate, polymyxin B Chondroitin lyase O-nitrophenyl-β-Dgalactopymanoride Growth on peptone Glucose source of carbon Acid from carbohydrates Degredation of Gelatin Casein Starch Tyrosine Tween 80 Urease H2 S Nitrate reduced Catalase Cytochrome oxidase G + C content (mol%) Habitat

Tenacibaculum maritimum∗

Long, Gram-negative rods 15–34◦ C 30◦ C + + + + 0.5 × 2–30 White, pale yellow Flat, irregular edges Gliding − + + − − + − + + + − + + − + + + 33–42 Marine (saprophytic)

˜ Source: Wakabayashi et al. 1986; Suzuki et al. 2001; Avendano 2009. +, positive reaction or characteristic; −, negative. ∗ Negative for Sucrose, D-ribose, DL-apartate, L-leucine; Degradation of starch, agar, carboxymethyl cellulose, cellulose, chitin, esculin; H2 s production. † SW-HS is Hsu-Shotts media (Shotts 1991) made with 100% of sea water.

media supplemented with 40% NaCl or seawater-Hsu-Shotts (SW-HS) media (Shotts 1991) made with 50% seawater and supplemented with neomycin sulfate (4 µg/mL) and polymyxin B (200 IU/mL) (Baxa et al. 1986). The 50% salt water can be made by adding 18.7 g/L of sea salts to the media. The bacterium grows at temperatures from 13 to 34◦ C with an optimum incubation temperature of 30◦ C. Colonies are rhizoid, have uneven edges, and are pale yellow, white, or light tan in color. The bacterium requires at least 33% seawater in the media with maximum growth occurring in broth made with 66–100% seawater (Chen et al. 1995). The bacterium is intolerant to 0% salt. According to Chen et al. (1995), cysts or microcysts are not formed by cultured T. maritimum, but microcysts have been reported by Baxa et al. (1986) and Kent et al. (1988). Key biochemical characteristics of T. maritimum are negative for flexirubin pigments and starch hydrolysis and positive for Congo red, catalase, cytochrome oxidase, hydrolysis of gelatin, and tyrosine (Avendano-Herrera ˜ 2009) (Table 17.1). A nested polymerase (PCR) system was developed by Cepeda et al. (2003) for identification of F. maritimum in fish tissue. The process can be completed in less than 4 hours and can detect as few as 75 cfu/mg in fish tissue. The nested PCR method is rapid and very sensitive in detecting T. maritimum applicable to routine diagnosis. Tenacibaculosis can also be confirmed by a fluorescein-based technique applied to samples from skin lesions (Labrie et al. 2005). Bader and Shotts (1998) used RNA gene sequence amplified by PCR to show close molecular similarities of T. maritimum (F. maritimus), Flavobacterium columnare (columnaris), and Flavobacterium psychrophilum (cold water disease of salmonids). Avendano-Herrera et al. (2004) reported sim˜ ilarities among these bacterial groups. In studying 29 T. maritimum fish isolates and 3 reference stains, Avendano-Herrera et al. ˜ (2004) found that biochemically all strains

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were identical serologically (slide agglutination, dot-blot assay, and immuno-blotting of LPS), and membrane proteins detected two major serological groups: serotype 01 and serotype 02. Furthermore these two strains seem to be host specific.

Epizootiology Tenacibaculum maritimum affects only marine fish and appears capable of infecting a wide variety of cultured species. To date most epizootics have involved marine cage cultured fish or wild fish. Trauma and skin abrasions are thought to be important precursors to marine tenacibaculosis in anchovies and sardines (Chen et al. 1995). Labrie et al. (2005) established a primary infection by immersion for 1 hour with exposure to as low as 104 cfu/ml of water with the pathogen easily observed in material from skin ulcers by FAT. Net-cage reared white sea bass are normally aggressive feeders and develop the disease following an interruption of feeding or due to climatic conditions and/or mechanical failure. Tenacibaculosis is often associated with environmental stress at temperatures above 15◦ C, inappropriate handling, parasite infestations, and inadequate water conditions. A natural reservoir is unknown. Gill lesions were also reported in chinook salmon in cages down current from other cages of T. maritimum infected bait fish. The theory is that gill lesions in the salmon developed when pieces of infected tissue from the bait fish lodged in their gills. It was also speculated that a gill infestation of Trichodina spp. may have predisposed the salmon to T. maritimum. While salmon culture in some coastal areas of California has been discontinued for various reasons, T. maritimum being one of them; however, successful culture of the more resistant white sea bass continues (Chen et al. 1995). Experimental induction of T. maritimum on gill of Atlantic salmon

was established by first gently abrading the gill and then exposing the gill to the pathogen (Powell et al. 2004). Some strains were highly pathogenic, while others were less pathogenic. The highly pathogenic strain produced morbidity and mortality within 24 hours and caused acute focal bronchial necrosis associated with significant increases in plasma osmolarity. Gill abrasion resulted in acute telangiectasis and focal lamellar hyperplasia regardless of the strain used for inoculation. Fish appear to also become infected via colonization of T. maritimum in the skin mucus (Magarinos ˜ et al. 1995) as the bacterium adheres strongly to the mucus and skin of turbot, seabream, and seabass, concluding that skin is probably a significant portal of entry for T. maritimum.

Pathology There is consistency in pathology between salmonids and non-salmonid species infected with T. maritimum and little difference is seen between natural and experimentally infected fish. Long, thin, basophilic bacteria are seen in sectioned skin and muscle lesions taken from northern anchovy and white sea bass (Chen et al. 1995). The bacteria extend into subdermal connective tissue producing congestion and hemorrhage. A loss of epidermis is noted in the ulcerated tissue and mats of bacteria extended into the dermis and subdermal layers. Bacteria also colonize the scale pockets accompanied by variable degrees of scale loss, low level inflammation in scale pockets, plus variable small adherent bacterial mats before complete epithelial erosion. Infiltration and inflammatory cells in the affected area are mild or absent. The earliest lesions to develop in salmonids and non-salmonids are fragmentation and degeneration of epithelium with infiltration of amorphous protein-like materials, congestion, and hemorrhage of the superficial dermis (Handlinger et al. 1997).

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Significance Marine tenacibaculosis is emerging as a significant pathogen of a variety of cage-cultured marine fishes in many parts of the world. Labrie et al. (2005) stated that importance of T. maritimum is largely underestimated since the isolation procedure, its difficulty to culture, and timing of sampling are critical factors for successful identification. Over the past 10 years, there has been an increase in scientific interest in tenacibaculosis, reflecting a growing concern over its adverse affect on mariculture. In some instances, it has contributed to the discontinuance of culture of certain highly susceptible species.

Management of other marine bacterial diseases Considering the relationship of T. maritimum infection to environmental stressors (water quality, temperature, handling, etc.), good mariculture practices are essential in management of this disease. Management recommendations for rearing white sea bass include placement of culture cages as far from live bait pens as possible and provide frequent and sufficient feeding to avoid antagonism and cannibalism (Chen et al. 1995). It was suggested by Santos et al. (1999) that avoiding high stocking density, reducing general stress, and avoid overfeeding are the best management practices to prevent marine tenacibaculosis. Also, adding a layer of sand on the bottom of tanks reduces infection in some instances. Hydrogen peroxide (H2 O2 ) kills T. maritimum at 30–240 ppm in in vitro tests but only the higher dose (240 ppm) had a positive effect in treating infection on fish (turbot) (Avendano-Herrera et al. 2006). They recom˜ mended that tanks, nets, etc. be disinfected with the higher dose of H2 O2 be used between stocking fish in them. Antibiotics in the feed can be considered when marine tenacibaculosis occurs but none

are FDA approved in the United States for this purpose. Tenacibaculum maritimum isolates from white seabass were sensitive to Terramycin and Romet-30 as well as some other drugs. However, Avendano-Herrera et al. ˜ (2008) demonstrated that of 63 T. maritimum isolates from fish all strains were resistant to oxolinic acid and all susceptible to amoxicillin, nitrofuranton, florfinicols, oxytetracycline (Terramycin), and trimethoprimsulfamethoxazole. Some isolates were resistant to enrofloxacin and flumequine ranging from 10 to 30% and from 25 to 60%, respectively. Vaccination can also be used to reduce effects of T. maritimum infections in Red Sea bream and Japanese flounder in Japan (Kato et al. 2007). The vaccine was a formalin killed bacterin from two strains of the pathogen applied by immersion. At 10 days postvaccination survival in vaccinated red sea bream and Japanese flounder were 80% (RPS 75%) and 40% (RPS 25%), respectively.

References Avendano-Herrera, R. 2009. Identification of Flex˜ ibacter maritimus or Tenacibaculum maritimum from post-larvae of Litopenaeus vannamei. Comment on Mourino ˜ et al. (2008). Brazilian Journal of Biology 69:225–226. Avendano-Herrera, R., M. Beatriz, et al. 2004. ˜ Phenotypic characterization and description of two major O-serotypes in Tenacibaculum maritimum strains from marine fishes. Diseases of Aquatic Organisms 58:1–8. Avenano-Herrera, R., M. Beatriz, et al. 2006. Use ˜ of hydrogen peroxide against the fish pathogen Tenacibaculum maritimum and its effect on infected turbot (Scophthalmus maximus. Aquaculture 257:104–110. Avenano-Herrera, R., S. Nu´ nez, et al. 2008. Evolu˜ ˜ tion of drug resistance and minimum inhibitory concentration to enrofloxacin in Tenacibaculum maritimum strains isolated in fish farms. Aquaculture International 16:1–11. Bader, J. A., and E. B. Shotts. 1998. Determination of phylogenetic relationship of Flavobacterium psychrophilum (Flexibacter

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psychrophilus), Flavobacterium columnare (Flexibacter columnare) and Flexibacter maritimus by sequence analysis of 16S ribosomal RNA genes amplified by polymerace chain reaction. Journal of Aquatic Animal Health 10:320–327. Baxa, D. V., K. Kawai, and R. Kusuda. 1986. Characteristics of gliding bacteria isolated from diseased cultured flounder, Paralichthys olivaceus. Fish Pathology 21:251–258. Bernardet, J. F., A. C. Campbell, and J. A. Buswell. 1990. Flexibacter maritimus is the agent of “black patch necrosis” in Dover sole of Scotland. Diseases of Aquatic Organisms 8:233–237. Cepeda, C., S. Garcia-Marquez, and Y Santos. 2003. Detection of Flexibacter maritimus in fish tissue using nested PCR amplification. Journal of Fish Diseases 26:65–70. Chen, M. F., D. Henry-Ford, and J. M. Groff. 1995. Isolation and characterization of Flexibacter maritimus from marine fishes in California. Journal of Aquatic Animal Health 7:318–326. Frelier, P. F., R. A. Elston, et al. 1994. Macroscopic and microscopic features of ulcerative stomatitis in farmed Atlantic salmon Salmo salar. Diseases of Aquatic Organisms 18:227–231. Handlinger, J., M. Soltani, and S. Percival. 1997. The pathology of Flexibacter maritimus in aquaculture species in Tasmania, Australia. Journal of Fish Diseases 20:159–168. Kato, F., K. Ishimoru, et al. 2007. Comparison of immersion-vaccination against gliding bacterial disease in red sea bream Pagrus major and Japanese flounder Paralichthys olivaceus. Aquaculture Science 55(1):97–101. Kent, M. L., C. F. Dungan, et al. 1988. Cytophaga sp. (Cytophagales) infection in seawater penreared Atlantic salmon Salmo salar. Diseases of Aquatic Organisms 4:173–179. Labrie, L., L. Grisez, et al. 2005. Tenacibaculum maritimum and underestimated fish pathogen in Asian marine fish culture (Abstract). World Aquaculture Society, Bali, Indonesia, May 9–13. Lopez, J. R., S. Nu´ nez, et al. 2009. Tenacibacu´ ˜ lum maritimum from wedge sole, Dicolgoglossa cuneata (Moreau). Journal of Fish Diseases 32:603–610.

Magarinos, B., F. Pazos, et al. 1995. Response of ˜ Pasteurella piscicida and Flexibacter maritimus to skin mucus of marine fish. Diseases of Aquatic Organisms 21:103–108. Mourino, ˜ J. L. P. L. Vinatea, et al. (2008) Characterization and experimental infection of Flexibacter maritimus (Wakabayashi, et al. 1986) in hatcheries of post–larvae of Litopenaeus vannamei Boon, 1931. Brazilian Journal of Biology 68(1). Pazos, F., Y. Santos, et al. 1993. Characterization of nisolated in northwest of Spain. 6th Interna˜ tional Conference of the European Association of Fish Pathologist, Brest, France (Abstract). Powell, M., J. Carson, and R. van Gelderen. 2004. Experimental induction of gill disease in Atlantic salmon Salmo salar smolts with Tenacibaculum maritimum. Diseases of Aquatic Organisms 61:179–185. Salati, F., C. Cubadda, et al. 2005. Immune response of sea bass (Dicentrarchus labrax) to Tenacibaculum maritimum antigens. Fisheries Science 71:563–567. Santos, Y., F. Pazos, and J. L. Baria. 1999. Flexibacter maritimus causal agent of flexibacteriosis in marine fish. Leaflet No. 55. ICES Identification Leaflet and Parasites of Fish and Shellfish. ICES Working Group of Pathology and Disease of Marine Organisms. International Councel for the Exploratioin of the Sea. Shotts, E. B. 1991. Selective isolation mehtods for fish pathogens. Journal of Applied Microbiology 70:75s–80s. Suzuki, M., Y. Nakagawa, et al. 2001. Phylogenetic analysis and taxonomic study of marine Cytophaga-like bacteria: proposal for Tenacibaculum gen. nov. with Tenacibaculum maritimum comb. Nov. and Tenacibaculum ovolyticum comb. Nov., and description of Tenacibaculum mesophilum sp. nov. and Tencibaculum amylolyticum sp. nov. International Journal of Systematic and Evolutionary Microbiology 51:1639–1652. Wakabayashi, H., M. Hikida, and K. Masumura. 1986. Flexibacter maritimus sp. nov., a pathogen of marine fishes. International Journal of Systematic Bacteriology 36:396–398.

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Part IV

Appendices

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Appendix I List of common and scientific names of fishes Common and scientific names of fishes used in the text. Latin names are based on World Fishes Important to North Americans, Aamerican Fisheries Society, Special Publication 21, American Fisheries Society, Bethesda, Maryland, USA (1991).

Common Name

Scientific Name

Alewife Amberjack Amberjack (yellowtail) American eel Arctic char Arctic grayling Atlantic cod (Baltic) Atlantic croaker (Baltic) Atlantic herring Atlantic halibut

Alosa pseudoharengus Seriola dumerili Serola lalandi Anguilla rostrata Salvelinus alpinus Thymallus arcticus Gadus morhua Micropogonius undulatus Clupea harengus Hippoglossus hippoglossus Scomber scombrus Brevoortia tyrannus Liparis atlanticus Salmo salar Microgadus tomcod Sardinops sagax neopilchardus Plecoglossus altivelis

Atlantic mackerel Atlantic menhaden Atlantic seasnail Atlantic salmon Atlantic tomcod Australian pilchard Ayu

Health Maintenance and Principal Microbial Diseases of Cultured Fishes, 3rd edition. By J. A. Plumb and L. A. Hanson. Published 2011 by Blackwell Publishing Ltd.

Common Name

Scientific Name

Baramundi perch Bengal danio (sind) Betta Bighead carp Black bullhead Black carp

Lates calcarifer Devario devario Betta splendens Aristichthys nobilis Ameiurus melas Mylopharyngodon piceus Hypophthalmichthys schlegeli Alburnus alburnus Ictalurus furcatus Oreochromus aureus Lepomis macrochirus Abramis brama Salvelinus fontinalis Ameiurus nebulosus Epinephelus lovina Salmo trutta Ictalurus punctatus Oncorhynchus tshawytscha Oncorhynchus keta Oncorhynchus kisutch Cyprinus carpio Phoxinus phoxinus

Black seabream (porgy) Bleak Blue catfish Blue tilapia Bluegill Bream (common) Brook trout Brown bullhead Brown-spotted grouper Brown trout Channel catfish Chinook salmon Chum salmon Coho salmon Common carp Common minnow (Eurasian) Common shiner Cutthroat trout Dab (North Sea) Damsel fish Doctorfish Emerald shiner Estuarine grouper

Lulilus cornutus Oncorhynchus clarki Pleuronectus limanda Chrysiptera sp. Labroides dimidatus Notropis atherinoides Epinephelus tauvina

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Common Name

Scientific Name

Common Name

Scientific Name

European catfish (wels) European perch European flounder European seabass

Silurus glanis Perca fluviatilis Platichthys flesus Morone (Decentrarchus) labrax Osmerus eperlanus Anquilla anquilla Pimephales promelas Channa maculata Osphronemus goramy Sparus auratus Dorosoma cepedianum Eigenmannia virescens Ctenolabrus rupestris Notemigonus crysoleucas Carassius auratus

Pacific herring Pacific sardine Pacific white shrimp Pallid sturgeon Pearl danio

Clupea pallasi Sardinops sagax Penaeus vannamei Scaphirhynchus albus Brachydanio albolineatus Odonthestes banariensis Lagodon rhomboides Oncorhynchus gorbuscha Pleuronectes platessa Siganus canaliculatus

European smelt European eel Fathead minnow Formosa snakehead Gourami Gilthead seabream Gizzard shad Glass knifefish Gold sing wrasse Golden shiner Goldfish (Crucian carp) Grass carp Grayling Greenback flounder Gudgeon (topmouth) Gulf killifish Gulf menhaden Guppy Hardhead catfish Ide (Orfe) Indian glassfish Itipa mojarras Japanese catfish Japanese eel Japanese striped knife jaw Kelp (red) grouper Lake sturgeon Lake trout Macquarie perch Masu (yamame, cherry salmon) Mozambique tilapia Mosquitofish Mountain galaxias Muskellunge Neon tetra Nile tilapia Northern anchovy Northern pike Olive flounder Pacific cod Pacific halibut

Ctenopharyngodon idella Thymallus thymallus Rhombosolea tapirina Gobio gobio Fundulus grandis Brevoortia patronus Poecilia reticulata Arius felis Leuciscus idus Chanda ranga Diapterus rhombeus Silurus asotus Anguilla japonica Oplegnathus faciatus

Pejerrey Pinfish Pink salmon Plaice Whitespotted rabbitfish Rabbitfish Rainbow smelt Rainbow trout Rare minnow Red drum Red seabream (Asia) Red seabream (New Zealand) Redfin perch (European perch) Roach Rudd Sablefish Seabream Sea raven Shiner perch Shorthorn sculpin Shortnose sturgeon

Epinephelus moora Acinpenser fulvescens Salvelinus namaycush Macquaria australasica Oncorynchus masou Oreochromis mossambicus Gambusia affinis Galaxias olidus Esox masquinongy Paracheirodon innesi Oreochromis niloticus Engraulis mordax Esox lucius Paralichthys olivaceus Gadus macrocephalus Hippoglosus stenolepis

Shovelnose sturgeon Silver carp Silver perch (North America) Silver perch (Australia) Silver seatrout Sockeye (kokanee) Snakehead (Chevron) Sole (Dover) Spotted grouper Striped bass Striped jack (White trevaly) Striped mullet Striped snakehead

Siganus rivulatus Osmerus mordax Oncorhynchus mykiss Gobiocypris rarus Sciaenops ocellatus Pagrus major Chrysophrys major Perca fluviatilis Rutilus rutilus Scardinius erythrophthalmus Anoplopoma fimbria Sparus aurata Hemitripterus americanus Cymatogaster aggregata Myoxocephalus scorpius Acipenser brevirostrum Scaphirhynchus platorynchus Hypophthalmichthys molitrix Bairdiella chrysoura Bidyanus bidyanus. Cynoscion nothus Oncorhynchus nerka Channa striata Solea solea Epinephelus akaara Morone saxatilis Caranx dentex Mugil cephalus Channa striata

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Appendix I List of Common and Scientific Names of Fishes

475

Common Name

Scientific Name

Common Name

Scientific Name

Striped trumpeter Tench Tellina Threespot gourami

Latris lineata Tinca tinca Tellina tenuis Trichogaster trichopteru Scophthalmus maximus Clarias batrachus Stizostedion vitreum Penaeus stylirostris Morone americana Morone chrysops Blicca bioerkna Ameiurus catus

Whitefish (ciscos) White sturgeon

Coregonus spp. Acipenser transmontanus Atractoscion nobilis Catostoma commersoni Gnathopogon elongatus Pleuronectus americanus Ameiurus natalis Perca flavescens Seriola quinqueradiata Danio rerio

Turbot Walking catfish Walleye Western blue shrimp White perch White bass White bream (silver) White catfish

White seabass White sucker Willow shiner Winter flounder Yellow bullhead Yellow perch Yellowtail Zebra danio (zebrafish)

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Appendix II Table of conversion factors

Conversion tables of metric and US customary measurement units in length, area, weights, volume and capacity, temperatures and other data used in aquaculture health management.∗

1 centimeter = 0.3937 inch 1 meter = 3.281 feet 1 meter = 1.094 yards 1 sq. centimeter = 0.155 sq. inch 1 sq. meter = 10.76 sq. feet 1 sq. meter = 1.196 sq. yards 1 hectare = 2.47 acres 1 hectare = 107,593 sq. feet 1 cu. centimeter = 0.061 cu. inch 1 cu. meter = 35.3 cu. feet 1 cu. meter = 1.308 cu. yards 1 milliliter = 0.0338 liquid ounce 1 liter = 0.2642 gallons 1 liter = 1.057 quarts 1 liter = 61.03 cu. inches 1 cubic inch = 16.39 cu. centimeters 1 acre foot = 325,850 gallons 1 gram = 15.43 grains 1 gram = 0.0535 ounces 1 ounce = 28.35 grams

Length

Area

Volume and Capacity

Weights

1 inch = 2.540 centimeters 1 foot = 0.305 meter 1 yard = 0.914 meter 1 sq. inch = 6.45 sq. centimeters 1 sq. foot = 0.0929 sq. meter 1 sq yard = 0.836 sq. meter 1 acre = 0.405 hectare 1 acre = 43,560 sq. feet 1 cu. foot = 7.48 liquid gallons 1 cu. foot = 28.3 liters 1 cu. yard=0.765 cu. meter 1 liquid ounce = 29.57 milliliters 1 liquid quart = 0.946 liters 1 gallon = 3.785 liters 1 gallon = 231 cu. inches 1 gallon = 128 fluid ounces 1 acre slice = 24,175 cu. feet 1 gallon = 8.34 pounds 1 gallon = 3.785 kilograms 1 pint = 1.04 pounds (Cont.)

Health Maintenance and Principal Microbial Diseases of Cultured Fishes, 3rd edition. By J. A. Plumb and L. A. Hanson. Published 2011 by Blackwell Publishing Ltd.

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478 Appendices

1 kilogram (water) = 1 liter 1 kilogram = 2.205 pounds 1 pound = 453.6 grams 1 ppm = 1 mg/liter or 1 mg/kg 1 ppm = 2.72 lbs. per acre foot 1 ppm = 8.34 lbs. per million gallons

1 cubic foot water = 62.4 pounds 1 cubic foot water = 28.3 kilograms 1 acre foot water = 2,718,144 pounds 1 ppm = 3.8 mg per gallon 1 ppm = 1 gm in 264 gallons 1 ppm = 0.38 gm. per 100 gallons

Percent Solution For 1 percent add: 38 grams per gallon 38 cubic centimeter per gallon 1.3 ounces per gallon 10 grams per liter 10 cubic centimeters per liter

Feeding Drugs For 1 percent add: 4.5 grams per pound of food 0.16 ounces per pound of food 70 grains per pound of food

Temperature Conversions Degrees Centigrade to Fahrenheit = (C × 9/5) + 32 Degrees Fahrenheit to Centigrade = (F – 32) × 5/9 Or substitute either in the formula: 1.8 C = F – 32 Source: Bureau of Sport Fisheries and Wildlife, Branch of Fish Hatcheries, Atlanta, Georgia (no date), and J. Jensen and R. Durborow, Tables for Applying Common Fishpond Chemicals. Circular ANR-414, Alabama Cooperative Extension Services, Auburn University, Alabama (no date).

Appendix II: Table A.1 Measurement Conversion Table: Weight in grams for teaspoon (tsp), table spoon (Tbsp), and cup volume for six commonly used chemicals in aquaculture. Chemical

1 Tsp

1 Tbsp

1/ 2

Copper sulfate (snow) (CuSO4 ) Copper sulfate (powder) (CuSO4 ) Potassium permanganate (KmNO4 ) Coarse-grain salt (NaCl) Table salt (NaCl) Formalin (37% formaldehyde)

6.4 g 4g 8g 4.8 g 6.5 g 5.3 g

19.2 g 12 g 24 g 14.4 19.5 g 15.8 g

153.6 g 96 g 192 g 115.2 g 156 g 126.4 g

Cup

1 Cup

307.2 g 192 g 384 g 230.4 g 312 g 252.8 g

Source: J. Jensen and R. Durborow, Tables for applying common fishpond chemicals. Circular ANR-414, Alabama Cooperative Extension Services, Auburn University, Alabama (no date).

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Appendix III List of cell lines commonly used for diagnostics Fish cell lines and viruses to which they are susceptible and that are commonly used for fish diagnostics. The list is sorted by fish family. Family Cell Line

Abbreviation

Source

Diagnostic Use∗

Acipenseridae White sturgeon skin White sturgeon spleen-2 White sturgeon Gonad Anguillidae Eel ovary Eel kidney

WSSK-1

Hedrick et al. 1991a

WSHV-1, WSHV-2

WSS-2

Hedrick et al. 1991b

WSGO

Watson et al. 1995

WSAV, WSIV, WSHV-1, WSHV-2 WSHV-1, WSHV-2

EO-2

Centrarchid Bluegill fry-2

BF-2

Japanese eel ovary. Chen and Kou 1981 Japanese eel kidney. Chen et al. 1982 Bluegill larvae. Wolf et al. 1966

Channidae Striped snakehead Snakehead spleen

SSN-1

Cyprinidae Fathead minnow

FHM

Epithelioma papillosum cyprini

EPC

EK-1

SHS

Striped snakehead fry. Frerichs et al. 1991. Spleen from fingerling striped snakeheads. Lio-Po et al. 1999 Pooled posterior portion of adult fathead minnows. Gravell and Malsberger 1965 Originally derived from a papilloma on a carp. Fijan et al. 1983. DNA data suggests fathead minnow source†

EVE, PCNV, Eel rhabdoviruses, HVA, EV-102 EVE, PCNV, Eel rhabdoviruses, HVA EV-102 LMBV, IHNV, VHSV, ECV, AmHV-1, EHNV, LCDV, EUS-P, EVE, Eel rhabdoviruses NNV, SKRV EUS-P

IHNV, VHSV, LMBV, SVCV, CyHV-1, GSV, PFR, EVE, Eel rhabdoviruses IHNV, VHSV, LMBV, SVCV, AmHV-1, ECV, CyHV-1, PFR, EV-102

(Cont.)

Health Maintenance and Principal Microbial Diseases of Cultured Fishes, 3rd edition. By J. A. Plumb and L. A. Hanson. Published 2011 by Blackwell Publishing Ltd.

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480 Appendices

Abbreviation

Source

Diagnostic Use∗

Koi Fin-1

KF-1

CyHV-2, CyHV-3

Common carp brain Grass carp ovary

CCB

Ctenopharyngodon idellus kidney Haemulidae Grunt fin Ictaluridae Brown bullhead Channel catfish ovary Percidae Walleye ovary Walleye embryo Walleye fibroblasts

CIK

Common carp (Koi) fin. Hedrick et al. 2000 Common carp brain. Neukirch et al. 1999 Grass carp ovary Zhang et al. 2003. Grass carp kidney. Zuo et al. 1986 Blue-striped grunt fin tissue. Clem et al. 1961 Brown bullhead caudal trunk. Wolf and Quimby 1969 Juvenile channel catfish ovary, Bowser and Plumb 1980 Kelley et al. 1983 Kelley et al. 1983 Derived from walleye dermal tumor. Kelley et al. 1980 Walleye dermal sarcoma cells. Rovnak et al. 2007 Pooled embryos of Chinook salmon. Lannan et al. 1984

WaHV WaHV

Family Cell Line

Spring tumor explant cells Salmonidae Chinook salmon embryo 214 Rainbow trout gonad-2

GCO

GF BB CCO WO We-2 WC-1 STEC CHSE-214

RTG-2

Steelhead embryo

STE-137

Atlantic salmon kidney

ASK

Atlantic salmon head kidney-1

SHK-1

Sciaenidae Red drum dorsal fin cells Sparidae Red sea bream fibroblast

RDDF-1

CRF-1

Pooled ovary and testis from rainbow trout. Wolf and Quimby 1962 Pooled embryos of steelhead trout. Lannan et al. 1984 Head kidney from an adult female Atlantic salmon. Devold et al. 2000 Head kidney from presmolt Atlantic Salmon. Dannevig et al. 1995 Red drum dorsal fin. Bowden et al. 1995 Fibroblastic cells from the caudal fin of red sea bream. Imajoh et al. 2007.

CyHV-3 LCDV GCHV RSIV CCV, CRV CCV, CRV, AmHV-1, ECV, EUS-P WaHV

WDSV‡ IHNV, VHSV, ISAV (some strains), IPNV, CRV, SaHV-1, SalHV-2, SAV IHNV, IPNV, VHSV, EHNV, Eel rhabdoviruses, EV-102, SalHV-1, SalHV-2, SAV VHSV ISAV

ISAV

LCDV

RSIV

Notes: ∗ AmHV-1, Ameiurine herpesvirus 1 (Black bullhead herpesvirus); CCV, channel catfish virus (Ictalurid herpesvirus 1); CRV, catfish reovirus; CyHV-1, cyprinid herpesvirus 1 (carp pox, carp herpesvirus); CyHV-2, cyprinid herpesvirus 2 (goldfish herpesvirus); CyHV-3, cyprinid herpesvirus 3 (Koi herpesvirus); ECV, European catfish virus, European sheatfish virus; EHNV, epizootic hematopoietic necrosis virus; EUS-P, epizootic ulcerative syndrome associated rhabdovirus; EV-102, Japanese eel iridovirus; EVE, eel virus European; GCHV, grass carp hemorrhagic virus (grass carp reovirus); GSV, golden shiner virus; HVA, herpesvirus Anguillidae (Anguillid herpesvirus 1); IHNV, infectious hematopoietic necrosis virus; IPNV, infectious pancreatic necrosis virus; ISAV, infectious salmon anemia virus; LCDV, lymphocystis disease virus; LMBV, largemouth bass virus (Santee-Cooper Ranavirus); NNV, nervous necrosis virus (fish nodavirus); PCNV, pillar cell necrosis virus; PFR, pike fry rhabdovirus, RSIV-red sea bream iridovirus; SalHV-1, salmonid herpesvirus 1; SalHV-2, salmonid herpesvirus 2 (Oncorhynchus masou virus); SAV, salmon alphavirus (salmon pancreas disease virus, sleeping disease virus); SKRV, Snakehead reovirus; SVCV, spring viremia of carp virus; WaHV, walleye herpesvirus; WDSV, walleye dermal sarcoma virus. † Early stocks of EPC cells were likely displaced by contaminating FHM cells but EPC and FHM cells have distinct differences in cell morphology and culture characteristics (personal Communication James Winton, U.S. Geological Survey, Western Fisheries Research Center, Seattle, WA, USA). ‡ The cell line persistently sheds WDSV. It is not used for diagnostics.

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Appendix III List of Cell Lines Commonly Used for Diagnostics

References Bowden, R. A., D. J. Oestmann, et al. 1995. Lymphocystis in red drum. Journal of Aquatic Animal Health 7:231–235. Bowser, P. R., and J. A. Plumb 1980. Growth rates of a new cell line from channel catfish ovary and channel catfish virus replication at different temperatures. Canadian Journal of Fisheries and Aquatic Sciences 37:871–873. Chen, S. N., and G. H. Kou. 1981. A cell line of Japanese eel (Anguilla japonica) ovary. Fish Pathology 16:129–137. Chen, S.-N., Y. Ueno, and G.-H. Kou. 1982. A cell line derived from Japanese eel (Anguilla japonica) kidney. Proceedings of the National Science Council Republic of China 6:93–100. Clem, L. W., L. Moewus, and M. M. Sigel. 1961. Studies with cells from marine fish in tissue culture. Proceeding of the Society of Experimental Biology and Medicine 108:762–766. Dannevig, B. H., K. Falk, and E. Namork. 1995. Isolation of the causal virus of infectious salmon anaemia (ISA) in a long-term cell line from Atlantic salmon head kidney. Journal of General Virology 76:1353–1359. Devold, M., B. Krossoy, et al. 2000. Use of RTPCR for diagnosis of infectious salmon anaemia virus (ISAV) in carrier sea trout Salmo trutta after experimental infection. Diseases of Aquatic Organisms 40:9–18. Frerichs, G. N., D. Morgan, et al. 1991. Spontaneously productive C-type retrovirus infection of fish cell lines. Journal of General Virology 72:2537–2539. Fijan, N., D. Sulimanovic, et al. 1983. Some properties of the Epithelioma papulosum cyprini (EPC) cell line from carp cyprinus carpio. Annales de l’Institut Pasteur. Virologie 134:207–220. Gravell, M., and R. G. Malsberger. 1965. A permanent cell line from the fathead minnow (Pimephales promelas). Annals of the New York Academy of Sciences 126:555–565. Hedrick, R. P., O. Gilad, et al. 2000. A herpesvirus associated with mass mortality of juvenile and adult koi, a strain of common carp. Journal of Aquatic Animal Health 12:44–57. Hedrick, R. P., J. M. Groff, and T. S. McDowell. 1991a. Isolation of an epitheliotropic her-

481

pesvirus from white sturgeon (Acipenser transmontanus). Diseases of Aquatic Organisms 11:49–56. Hedrick, R. P., T. S. McDowell, et al. 1991b. Two cell lines from white sturgeon. Transactions of the American Fisheries Society 120:528–534. Imajoh, M., T. Ikawa, and S. Oshima. 2007. Characterization of a new fibroblast cell line from a tail fin of red sea bream, Pagrus major, and phylogenetic relationships of a recent RSIV isolate in Japan. Virus Research 126:45–52. Kelly, R. K., H. R. Miller, et al. 1980. Fish cell culture: characteristics of a continuous fibroblastic cell line from walleye (Stizostedion vitreum vitreum). Canadian Journal of Fisheries and Aquatic Sciences 37:1070–1075. Kelly, R. K., O. Nielsen, et al. 1983. Characterization of Herpes virus vitreum isolated from hyperplastic epidermal tissue of walleye, Stizostedion vitreum vitreum (Mitchill). Journal of Fish Diseases 6:249–260. Lannan, C. N., J. R. Winton, and J. L. Fryer 1984. Fish Cell Lines: Establishment and Characterization of Nine Cell Lines from Salmonids. In Vitro 20:671–676. Lio-Po, G.D., G.S. Traxler, and L.J. Albright. 1999. Establishment of cell lines from catfish (Clarias batrachus) and snakeheads (Ophicephalus striatus). Asian Fisheries Science 12:345–349. Neukirch, M., K. Bottcher, and S. Bunnajirakul. ¨ 1999. Isolation of a virus from Koi with altered gills. Bulletin of the European Association of Fish Pathologists 19:221–224. Rovnak, J., R. N. Casey, et al. 2007. Establishment of productively infected walleye dermal sarcoma explant cells. Journal of General Virology 88:2583–2589. Watson, L. R., S. C. Yun, et al. 1995. Characteristics and pathogenicity of a novel herpesvirus isolated from adult and subadult white sturgeon Acipenser transmontanus. Diseases of Aquatic Organisms 22:199–210. Wolf, K., M. Gravell, and R. G. Malsberger 1966. Lymphocystis virus: isolation and propagation in centrarchid fish cell lines. Science 151:1004–1005. Wolf, K. and M. C. Quimby 1962. Established eurythermic line of fish cells in vitro. Science 135:1065–1066.

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Wolf, K. and M. C. Quimby. 1969. Fish cell and tissue culture. In: W. Hoar and D. J. Randall (eds) Fish physiology, Vol. 3. New York, Academic Press, pp. 253–301. Zhang, Q.-Y., H.-M. Ruan, et al. 2003. Infection and propagation of lymphocystis virus isolated from the cultured floun-

der Paralichthys olivaceus in grass carp cell lines. Diseases of Aquatic Organisms 57:27–34. Zuo, W., H. Qian, et al. 1986. A cell Line derived from the kidney of grass carp. Journal of Fisheries of China 10:11–17. (Chinese with English abstract)

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Index Acetic acid (vinegar), 72t, 72 Acriflavin, 75 Acute infection, 36 Adenovirus eel, 143 white sturgeon, 219–220 Adjuvants, 82 Aeration, 9, 10f Aerococcus viridans, 70 Aeromonas hydrophila, 8, 14–15, 34, 321–322, 332, 337 in association with atypical Aeromonas salmonicida, 109, 318, 320 striped bass, 433–434 tilapia, 454 Aeromonas salmonicida, 16, 36, 52, 70, 74, 76, 315–321 bacterial characteristics, 318, 319t clinical signs, 316, 317f in cultured eels, 337 diagnosis, 316, 318 epizootiology, 319–321 furunculosis. See Furunculosis geographical range and species susceptibility, 315–316 management, 393–394 pathology, 321 probiotics and, 60, 395 significance, 321 vaccines, 77, 78t, 80t, 81, 398 Aeromonas salmonicida achromoguenes, 109, 315, 319t, 321 Aeromonas sobria, 60 AFS Blue Book, 49, 52 Alaskan sockeye salmon, 147–148, 153–155, 154t, 177, 185, 187, 194 Algae, 60, 62 Ameiurine herpesvirus 1 (AmHV-1), 95 American grass carp reovirus, 126 Amoxicillin, 74–75 Ampicillin, 75 Anemia, 41 Anguillid herpesvirus, 139–141 Animal Use Clarification Act of 1994 (AMDUCA), 74–75 Antigens, 53, 79, 81 Appendices cell lines commonly used for diagnostics, 479–480

conversion factors, 477–478 scientific names of fishes, 473–475 Approved drugs, 68, 69t, 70–72 Aquabirnaviruses, 247–248 Aquaculture, 3 Aquareovirus diseases, 123, 193–194 Aquatic birnaviruses, 160, 161t Argulus foliaceus, 114 Ascites, 41–42 Atlantic salmon, temperature requirements for, 9t Atlantic salmon paramyxovirus (ASPN), 195–196 Atrophy, 43 Bacillus mycoides, 300–301 Bacterial diseases bacterial cold-water disease, 375–380 bacterial gill disease (BGD), 5t, 36, 71, 369–375 bacterial kidney disease (BKD), 36, 74, 380–389 carp bacterial diseases. See Carp and minnow bacterial diseases catfish bacterial diseases. See Catfish bacterial diseases disease diagnosis, 51–52 eel bacterial diseases. See Eel, bacterial diseases minnow bacterial diseases. See Carp and minnow bacterial diseases miscellaneous, 465–469 salmonid bacterial diseases. See Salmonid bacterial diseases striped bass bacterial diseases. See Striped bass, bacterial diseases tenacibaculosis. See Tenacibaculosis tilapia bacterial diseases. See Tilapia, bacterial diseases Benzocaine, 75 Bicozamycin, 75 Biosecurity, 4 Black bullhead herpesvirus, 95, 97, 101–102 Black bullhead iridovirus, 95, 101 Blue spot disease of pike and muskellunge, 253, 253f Breeding and culling, 18 Brook trout, temperature requirements for, 9t Brown bullhead, 97 Brown trout, 168, 347f temperature requirements, 9t Calcium chloride, 72, 72t Calcium oxide, 72, 72t

483

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484 Index

Carbon dioxide gas, 72, 72t Carnobacterium piscicola, 392–393, 435–436 Carp and minnow bacterial diseases, 315–323 atypical nonmotile Aeromonas salmonicida infections, 315–321 management, 322–323 miscellaneous bacterial diseases of cyprinids, 321–322 overview, 315 Carp and minnow viruses, 109–129 American grass carp reovirus, 126 carp cornavirus, 127 carp edema virus, 126–127 cyprinid herpesvirus 2 (CyHV-2), 118–120 fathead minnow rhabdovirus, 127 fish pox (CyHV-1), 115–118 golden shiner virus, 123–125 goldfish iridoviruses, 125–126 grass carp hemorrhagic virus, 126 koi herpesvirus (cyprinid herpesvirus 3), 120–123 koi sleepy disease virus, 126–127 management, 127–129 overview, 109 spring viremia of carp, 109–115 Carp cornavirus, 127 Carp edema virus, 126–127 Carp erythrodermatitis, 315 Carrying capacity, 58 Catarrhal inflammation, 45 Catfish, drugs approved for, 69t Catfish bacterial diseases, 275–305 Bacillus mycoides, 300–301 columnaris, 275–283 Edwardsiellosis, 300 ESC (“hole-in-the-head” disease), 283–293 management, 301–305 chemotherapy, 302–304 vaccination, 304–305 MAS, 293–300 overview, 275 Pseudomonas septicemia, 300 Catfish viruses, 95–104 black bullfish herpesvirus, 101–102 catfish reovirus, 101 channel catfish virus disease, 95–101 European catfish virus (ECV), 102 European sheatfish virus (ESV), 102–103 management of, 103–104 overview, 95 Cauliflower disease, 141 Cell lines commonly used for diagnostics, 479–480 Cellular degeneration in fish, 42–43 Channel catfish drugs approved for, 69t temperature requirements, 9, 9t Channel catfish virus disease (CCVD), 16–17, 95–101 clinical signs, 96, 96f diagnosis, 96–97, 98f epizootiology, 99–101 geographical range and species susceptibility, 95–96 pathology, 101

significance, 101 virus characteristics, 97, 99 Chemicals. See Drugs and chemicals Chinook salmon epizootic epitheliotropic disease, 189 erythrocytic inclusion body syndrome, 180 infectious hematopoietic necrosis virus, 147–148, 153 infectious salmon anemia virus, 166 plasmacytoid leukemia virus, 194 salmonid herpesvirus 1, 183 temperature requirements, 9t viral hemorrhagic septicemia, 171, 177 Chlamydia, 195 Chloramine-T (Halamid Aqua), 74 Chloramphenicol, 75 R Chorionic gonadotropin (Chorulon
), 71 Chronic infection, 36 Chum salmon erythrocytic inclusion body syndrome, 180 infectious hematopoietic necrosis virus, 148, 154 infectious salmon anemia virus, 166 reovirus, 193 salmonid herpesvirus 1, 183 salmonid herpesvirus 2, 185, 186f, 187–188, 188f Circulatory disturbances in fish, 41–42 Clinical signs, 40, 47t Cloudy swelling, 43 Coho salmon erythrocytic inclusion body syndrome, 180, 182–183, 182t infectious hematopoietic necrosis virus, 148, 149f, 150, 155 infectious salmon anemia virus, 166 salmonid herpesviruses, 183, 185, 187–189 temperature requirements, 9t viral hemorrhagic septicemia, 171, 172t, 174, 177 Cold-water vibriosis, 359–363 bacterial characteristics, 361–362 clinical signs, 359, 360f–361f diagnosis, 360–361 epizootiology, 362–363 geographical range and species susceptibility, 359 management, 394 pathology, 363 significance, 363 vaccines, 78t Colistin sulfate, 75 Columnaris, 36, 275–283, 393 bacterial characteristics, 277, 279, 280t clinical signs, 276, 277f diagnosis, 276, 278f epizootiology, 280–282, 282f geographical range and species susceptibility, 275–276 pathology, 282 significance, 282–283 stress mediated, 5t tilapia, 445, 455, 456f vaccines, 78t Communicable disease, definition of, 31 Contact, extent of, 15–16, 15f

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Index 485

Conversion factors, 477–478 Copper sulfate, 72–74 Corynebacterium aquaticum, 436 Covert infection, 36–37 Crowding, 58 Cutthroat trout virus, 194 Cyprinidae, 109, 479 Cyprinid herpesvirus 2 (CyHV-2), 118–120 clinical signs, 119 diagnosis, 119 epizootiology, 119–120 geographical range and species susceptibility, 118–119 pathology, 120 significance, 120 virus characteristics, 119 Cytophaga psychrophila, 375 Deferred regulatory status drugs, 72–73 copper sulfate, 73 potassium permanganate, 72–73 Diagnostics, cell lines commonly used for, 479–480 Diffuse epidermal hyperplasia, 254, 255f, 256 Dip method, 65 Discrete epidermal hyperplasia, 256, 258 Disease, definition of, 31 Disease, epizootiology of. See Epizootiology of fish diseases Disease management, 57–86 drugs and chemicals. See Drugs and chemicals fish health management. See Fish health management overview, 57 vaccination. See Vaccination Disease recognition and diagnosis, 45–53 clinical signs, 46–48 behavior, 46 external, 46–48 gross internal lesions, 48 disease diagnosis, 49–52 bacterial diseases, 51–52 parasitic diseases, 50 pathogen identification, 49–52 viral diseases, 50–51 disease recognition, 45–46 history, 46 mortality pattern, 46 molecular diagnostics, 52–53 antigen identification, 53 fatty acids, 52 Disturbances of development and growth, 43–44 Doxycycline, 75 Drugs and chemicals, 63–76 approved drugs, 68, 69t, 70–72 R chorionic gonadotropin (Chorulon
), 71 TM oxytetracycline hydrochloride (OxyMarine , R Oxytetracycline HCl Soluble Powder-345
, R R

Terramycin-345 , TETROXY Aquatic ), 71–72 R tricaine methanesulfonate (Finquel
and R
Tricaine-S ), 69t, 71 calculations, 66–67

deferred regulatory status, 72–73 copper sulfate, 73 potassium permanganate, 72–73 drugs of low regulatory priority (LRP), 72, 72t extra label drugs, 74–75 feed additive antibacterial, 68, 69t, 70 R florfenicol (Aquaflor
), 68, 69t, 70 R oxytetracycline dihydrate (Terramycin 202
for fish), 69t, 70 R sulfadimethoxine and ormetoprim (Romet-30
, R Romet TC
), 69t, 70 R sulfamerazine (Sulfamerazine in Fish Grade
), 69t, 70 future outlook, 76 immersion, 69t, 70–71 R R formalin (Formalin-F
, Paracide-F
, R R Parasite-S
, Formacide-B
), 69t, 71 R hydrogen peroxide (35% PEROX-AID
), 69t, 71 international use of drugs in aquaculture, 75–76 investigative new animal drugs (INADs), 73–74 17 α-methyltestosterone, 74 amoxicillin, 74 chloramine-T, 74 copper sulfate, 74 emamectin benzoate, 74 erythromycin thiocyanate, 74 potassium permanganate, 74 overview, 63–64, 67–68 therapeutic applications, 65–66 dip method, 65 flushing, 65 indefinite treatments, 65–66 injection, 66 oral, 66 prolonged bath, 65 treatment process, 64–65 withdrawal time, 66 Drugs of low regulatory priority (LRP), 72, 72t Early diagnosis, 23–24 Edema, 41–42 Edwardsiella ictaluri, 16, 18–20, 25, 45, 70, 329–330 API 20E code, 52 vaccines, 78t, 79, 81–82, 85 Edwardsiella tarda, 19, 327–334, 329f, 338–339, 393 striped bass, 434 tilapia, 454 Edwardsiellosis, 300, 327–334 bacterial characteristics, 329–331 330t clinical signs and findings, 328, 329f diagnosis, 328–329 epizootiology, 331–333 geographical range and species susceptibility, 327–328 pathology, 333–334 significance, 334 tilapia, 445, 456 Eel bacterial diseases, 327–339 Edwardsiellosis, 327–334 management, 338–339

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486 Index

Eel (cont.) chemotherapeutics, 338–339 vaccination, 339 miscellaneous diseases, 337–338 overview, 327 red spot disease, 334–337 temperature requirements, 9t viruses, 135–143 anguillid herpesvirus, 139–141 eel adenovirus, 143 eel iridovirus, 143 eel rhabdoviruses, 137–139, 137t eel virus European (EVE), 135–137 management, 143 overview, 135 stomatopapilloma, 141–143 Emamectin benzoate, 74 Embolism, 42 Entamoeba histolytica, 333 Enteric redmouth (ERM), 363–369 bacterial characteristics, 365–367, 365t–366t clinical signs, 364 diagnosis, 364–365 epizootiology, 367–368 geographical range and species susceptibility, 363–364 management, 394 pathology, 368 significance, 368–369 stress mediated, 5t vaccines, 78t Enteric septicemia of catfish (ESC), 283–293 bacterial characteristics, 286–288, 287t clinical signs, 284 diagnosis, 284, 285f, 286 epizootiology, 288–291, 289f–290f geographical range and species susceptibility, 283–284 pathological manifestations, 291–292 significance, 292–293 vaccines for, 78t Enterococcus faecium, 435 Environment, clean, maintaining, 24–25 Enzootic (endemic), definition of Epistylis spp., 73 Epizootic (epidemic), definition of, 31 Epizootic epitheliotropic disease (EED), 189–191 Epizootic hematopoietic necrosis (EHN), 193, 227–230 Epizootic ulcerative syndrome (EUS), 227, 248 Epizootiology of fish diseases, 31–37 degree of infection, 35–37, 35f factors in disease development, 33–35 natural resistance of the host, 34–35 portal of entry, 34 source of infection and mode of transmission, 33–34 virulence of pathogenic organisms, 34 host/pathogen relationship, 35 overview, 31 seasonal trends, 32–33, 32f terms, 31–32

Epsilonretrovirus, 195 Epsom salts (magnesium sulfate), 72t Erythrocytic inclusion body syndrome (EIBS), 180–183 Erythromycin, 74–75 Esox lymphosarcoma, 260–261, 260f Etiological agent, definition of, 40 Etiology, definition of, 40 European catfish virus (ECV), 102 European eel virus (EVE), 135–137 European sheatfish virus (ESV), 102–103 Exophthalmia, 42, 96f Exposure, avoiding, 12–14 Extra label drugs, 74–75 Fathead minnow rhabdovirus, 127 Fatty acids (molecular diagnostics), 52 Fatty degeneration, 43 FDA CVM (U.S. Food and Drug Administration Center for Veterinary Medicine), 63, 68 Feed additive antibacterial, 68, 69t, 70 R florfenicol (Aquaflor
), 68, 69t, 70 R oxytetracycline dihydrate (Terramycin 202
for fish), 69t, 70 R sulfadimethoxine and ormetoprim (Romet-30
, R Romet TC
), 69t, 70 R sulfamerazine (Sulfamerazine in Fish Grade
), 69t, 70 Fibrinous inflammation, 44 Fish disease. See Disease entries Fish eggs, drugs for, 69t, 71, 72t Fish health management, 3, 57–62 aeration management, 61, 61f feed management, 59–60 fish handling and stocking, 58–59, 59t miscellaneous environmental problems, 61–62 overview, 57–58 waste management, 62 water flow management, 60 Fish health status, 26–27, 26f Fish lice, 114 Fish names, common and scientific, 473–475 Fish pox (CyHV-1), 115–118 Flavobacterium aquatile, 369–371, 372t Flavobacterium branchiophilum, 36, 400 Flavobacterium columnare, 18, 36, 70–71, 322, 393 bacterial gill disease, 369–375 eel, 327 infection in conjunction with atypical Aeromonas salmonicida, 320 infection in conjunction with CCVD, 99 striped bass, 434–435 vaccine, 18, 79 Flavobacterium psychrophilum, 18, 70, 375–380, 394–395, 400 Flexibacter maritimus, 465 Flexibacter psychrophila, 375 R Florfenicol (Aquaflor
), 68, 69t, 70, 75 Flumequine, 75 Flushing, 65 R R R Formalin (Formalin-F
, Paracide-F
, Parasite-S
, R Formacide-B
), 71

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Fosfomycin, 75 Francisella asiatica, 445, 452–454 Francisella spp., 436, 452–454 Francisellosis, 452–454 Fruluphenicol, 75 Fullers earth, 72, 72t Furazolidone, 75 Furnace, 75 Furunculosis, 16, 36, 76, 345–353 bacterial characteristics, 348 clinical signs, 346–347, 347f diagnosis, 347–348 epizootiology, 348–352 geographical range and species susceptibility, 346 pathology, 352–353 significance, 353 stress mediated, 5t vaccines, 78t Gaffkemia, 70 Garlic (whole), 72, 72t Goezia, 16 Golden shiner virus, 123–125 Goldfish, 315 cyprinid herpesvirus 2, 118–120 erythrodermatitis, stress mediated, 5t iridoviruses, 125–126 Granulomatous inflammation, 45 Grass carp hemorrhagic virus, 126 reovirus, vaccination against, 128–129 temperature requirements, 9t Health maintenance disease management. See Disease management epizootiology of fish diseases. See Epizootiology of fish diseases general principles. See Health maintenance, principles of pathology and disease diagnosis. See Pathology and disease diagnosis Health maintenance, principles of, 3–27 avoiding exposure, 12–14 breeding and culling, 18 clean environment, maintaining, 24–25 contact, extent of, 15–16, 15f dynamic team effort, 24 early diagnosis, 23–24 eradication, prevention, control, 20–21 exposing dose, 14–15 fish health status, 26–27, 26f hazard reduction by management, 5–9, 6f–8f, 9t, 10f health maintenance, 4 high-risk concept, 25 keeping current, 24 law of limiting factors, 22 location, soil, and water, 9–12, 11t new arrivals, 17–18 nutritional basis of health maintenance, 19–20

overview, 3–4 prevention rather than just cure, 21–22 record keeping and cost analysis, 25–26 segregation, protection by, 16–17 staying on top of operation, 22–23 stress, 4–5, 5t variable causes require variable solutions, 21 Hematomas, 41 Hemorrhage, 41 Hemorrhagic inflammation, 45 Herpesviral hematopoietic necrosis, 118 Herpesvirus black bullfish, 101–102 carp, 115 cyprinid, 118–123 eel, 139–141 koi (cyprinid herpesvirus 3), 120–123 miscellaneous, 251 salmonid. See Salmonid herpesviruses turbot, 252 white sturgeon, 221–223 Herpesvirus anguillae, 139–140 Herpesvirus ictaluri, 95 Histology, 40 Histopathology, 40 Hitra disease, 357. See also Cold-water vibriosis Hole-in-the-head disease. See Enteric septicemia of catfish (ESC) Host/pathogen relationship, 35 R Hydrogen peroxide (35% PEROX-AID
), 69t, 71 Hydropsy, 42 Hyperemia and congestion, 42 Hyperplasia, 44 Hypertrophy, 43–44 Ice, 72t Ichthyophonus hoferi, 45 Ichthyophthirius multifiliis, 27, 36, 45, 73 Ictalurid herpesvirus 1, 95 Ictalurid herpesvirus 2, 95 Ictalurid melas herpesvirus, 95, 97 Immersion treatments, 69t, 70–71, 83f, 84 R R R formalin (Formalin-F
, Paracide-F
, Parasite-S
, R
Formacide-B ), 69t, 71 R hydrogen peroxide (35% PEROX-AID
), 69t, 71 Indefinite treatments, 65–66 Infection, 35–37, 35f acute, 36 chronic, 36 covert, 36–37 definition of, 31–32 subacute, 36 Infectious hematopoietic necrosis virus (IHN), 147–156 clinical signs and findings, 148, 149f diagnosis, 148–151, 150f epizootiology, 152–155, 153t–154t geographic range and species susceptibility, 148 pathology, 155–156 significance, 156 vaccine, 78t virus characteristics, 151–152

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488 Index

Infectious pancreatic necrosis (IPN) virus, 12, 36, 156–166 clinical signs, 157–158, 157f diagnosis, 158–160, 159f epizootiology, 162–165 geographical range and species susceptibility, 157, 157t pathology, 165–166 significance, 166 vaccines, 77, 78t, 80t virus characteristics, 160–162, 161t Infectious salmon anemia (ISA) virus, 166–170 clinical signs, 166 diagnosis, 166–167 epizootiology, 168–169 pathology, 169–170 significance, 170 species susceptibility and geographical range, 166 virus characteristics, 167–168 Inflammation, 44–45 catarrhal, 45 fibrinous, 44 granulomatous, 45 hemorrhagic, 45 purulent, 44 serous, 44 Injection, 66, 83f, 84–85 Investigative new animal drugs (INADs), 73–74 17 α-methyltestosterone, 74 amoxicillin, 74 chloramine-T, 74 copper sulfate, 74 emamectin benzoate, 74 erythromycin thiocyanate, 74 potassium permanganate, 74 Iridoviruses eel, 143 miscellaneous, 251 Isavirus, 167 Josamycin, 75 Kanamyacin, 75 Kitasamycin, 75 Koi herpesvirus (cyprinid herpesvirus 3), 120–123 clinical signs, 121, 121f diagnosis, 121–122 epizootiology, 122–123 geographical range and species susceptibility, 120–121 pathology, 123 significance, 123 vaccine, 78t virus characteristics, 122 Koi sleepy disease virus, 126–127 Lactococcus garvieae, 60, 395, 445 Lake trout, temperature requirements for, 9t Largemouth bass, temperature requirements for, 9t Largemouth bass virus (LMBV), 248–251, 249f Law of limiting factors, 22 Leeches, 114, 155

Lepidorthosis, 42 Lesion, definition of, 40 Lincomycin, 75 Low dissolved oxygen syndrome, 7 Lymphocystis, 230–231, 233–235 clinical signs, 231, 232f–233f diagnosis, 231 epizootiology, 234–235 geographical range and species susceptibility, 230–231 pathology, 235 significance, 235 virus characteristics, 233–234 Magnesium sulfate (epsom salts), 72t Malachite green, 75 MAS. See Motile Aeromonas septicemia Metaplasia, 44 Methyldihydrotestosterone, 75 Methylene blue, 75 Milkfish, temperature requirements for, 9t Minnow bacterial diseases. See Carp and minnow bacterial diseases Minnow viruses. See Carp and minnow viruses Miroxisacin, 75 Molecular diagnostics, 52–53 antigen identification, 53 fatty acids, 52 Moraxella, 435 Motile Aeromonas septicemia (MAS), 36, 52, 70, 293–300 bacterial characteristics, 295, 296t clinical signs, 293, 294f, 295 diagnosis, 295 eel, 327, 337 epizootiology, 295–299, 297f geographical range and species susceptibility, 293 pathology, 299 significance, 299–300 stress mediated, 5, 5t tilapia, 445 Mucoid degeneration, 43 Mycobacteriosis and nocardiosis, 419–428 bacterial characteristics, 423–424, 424t clinical signs, 421 diagnosis, 421–423, 422f epizootiology, 424–427 geographical range and species susceptibility, 420–421 pathology, 427–428 significance, 428 tilapia, 456 Mycobacterium spp., 45, 419 mycobacteriosis. See Mycobacteriosis and nocardiosis Myxobolus cerebralis (whirling disease), 17, 48 Nalidixic acid, 75 Names of fish, common and scientific, 473–475 National Coordinator for Aquaculture New Animal Drug Applications, 75 Necrosis, 43

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Neoplasia, 44 New arrivals, danger of, 17–18 Nifurstyrenate, 75 Nitrofurantoin, 75 Nocardia spp. See Mycobacteriosis and nocardiosis Nocardiosis. See Mycobacteriosis and nocardiosis Northern pike, temperature requirements for, 9t Norwegian salmonid alphavirus, 193 Novobiocin, 75 Nucleospora salmonis, 194–195 Nutritional basis of health maintenance, 19–20 OIE Diagnostic Manual, 49 Oleandomycin, 75 Oncorhynchus masou virus, 185, 186f Onion (whole), 72t Oral administration of drugs, 66 Oral vaccination, 82, 84 Oxolinic acid, 75 R Oxytetracycline dihydrate (Terramycin 202
for fish), 69t, 70 Oxytetracycline hydrocholride TM (OxyMarine , Oxytetracycline HCl Soluble R
Powder-345 R , Terramycin-345
, TETROXY R Aquatic
), 71–72 Pacific salmon, 74 Papin, 72t Paracolobacterum anguillimortifera, 327 Parasitic diseases, diagnosis of, 50 Pasteurella piscicida, 429 Pasteurellosis. See Photobacteriosis (Pasteurellosis) Pathogenesis, definition of, 40 Pathogenicity, definition of, 40 Pathogen identification, 49–52 Pathological changes in fish, 41–45 cellular degeneration, 42–43 circulatory disturbances, 41–42 disturbances of development and growth, 43–44 inflammation, 44–45 overview, 41 Pathology and disease diagnosis, 39–54 cause of disease, 40–45 pathological change. See Pathological changes in fish disease recognition and diagnosis. See Disease recognition and diagnosis histology, 40 histopathology, 40 overview, 39, 54 terms, 39–40 Peduncle disease, 375 Penaeid shrimp, drugs approved for, 69t, 71 Perch rhabdoviruses, 247 Photobacteriosis (Pasteurellosis), 429–433 bacterial characteristics, 430–431, 431t clinical signs and findings, 429 diagnosis, 429–430 epizootiology, 431–432 geographical range and species susceptibility, 429 pathology, 432–433

significance, 433 vaccines, 78t Photobacterium damsella, 353, 454 vaccines, 78t Photobacterium damsella subsp. piscicida, 45, 429–433 Photobacterium salmonis, vaccines for, 78t Phytoplankton, 59–60, 62 Pike fry rhabdovirus disease (PFRD), 244–247, 245f Pilchard herpesvirus (PHV) disease, 251–253 blue spot disease of pike and muskellunge, 253, 253f herpesvirus of turbot, 252 Pisciocola geometra, 114 Piscirickettsia salmonis, 389–392, 398, 401 Piscirickettsiosis, 389–392, 401 clinical signs, 389–390 diagnosis, 390 epizootiology, 391–392 geographical range and species susceptibility, 389 pathogen characteristics, 390–391 pathology, 392 significance, 392 Plasmacytoid leukemia virus, 194–195 Plesiomonas shigelloides, 445, 454–455 Portal of entry, 34 Potassium chloride, 72t Potassium permanganate, 72–74 Povodine iodine, 72t Prevention, disease, 21–22 Probiotics, 60, 395 Prolonged bath, 65 Proteocephalus ambloplitis (bass tapeworm), 15 Protozoan parasites, 36 Pseudokidney disease, 392 Pseudomonas anguilliseptica, 327, 330t, 334–339 Pseudomonas fluorescens, 322, 328, 330t, 454 Pseudomonas septicemia, 300 tilapia, 445 Pseudomonas spp., 322 Purulent inflammation, 44 Quarantine, 13 Rainbow trout Atlantic salmon paramyxovirus, 195 bacterial cold-water disease, 375–380, 376f bacterial gill disease, 369, 370f, 371, 373–374 bacterial kidney disease, 381–382, 381f, 385–388 cold-water vibriosis, 359 enteric redmouth, 363–364 epizootic hematopoietic necrosis virus, 193 erythrocytic inclusion body syndrome, 180 furunculosis, 346, 350–352 infectious hematopoietic necrosis virus, 147–150, 149f, 152–153, 153f, 155–156, 198 infectious pancreatic necrosis virus, 157, 157f, 160, 161t, 162, 164–165, 198–199 infectious salmon anemia virus, 166 piscirickettsiosis, 389–391 probiotics, 60 pseudokidney disease, 392

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490 Index

Rainbow trout (cont.) rainbow trout fry syndrome, 375 salmonid herpesvirus 1, 183–184 salmonid herpesvirus 2, 185, 187–189 sleeping disease, 191–193 temperature requirements, 9t vibriosis, 357–358, 362–363 viral hemorrhagic septicemia, 170–171, 172t, 173f, 174–175, 177–179, 197–199 Yersinia ruckeri, 361f, 365, 367–369 Record keeping and cost analysis, 25–26 Recrudescent infection, 36 Redfin perch, 193 Red sea bream iridoviral disease (RSIV), 242–244 clinical signs, 243 diagnosis, 243 epizootiology, 243–244 pathology, 244 significance, 244 species susceptibility and geographical range, 243 virus characteristics, 243 Red spot disease, 334–337 bacterial characteristics, 335 clinical signs, 335 diagnosis, 335 epizootiology, 336 geographical range and species susceptibility, 334 pathology, 336–337 significance, 337 Renibacterium salmoninarum, 14, 36, 52, 380–389, 394–395, 398, 400 Reovirus American grass carp reovirus, 126 aquareovirus diseases, 193–194 catfish reovirus, 101 Resistance, host, 34–35 Rhabdoviruses, eel, 137–139, 137t, 142 clinical signs, 138 diagnosis, 138 epizootiology, 138–139 geographical range and species susceptibility, 137–138 pathology, 139 significance, 139 virus characteristics, 138 Rickettsia, 195 Salmincola sp., 155 Salmonid bacterial diseases, 345–401 bacterial cold-water disease, 375–380 bacterial gill disease (BGD), 369–375 bacterial kidney disease (BKD), 380–389 cold-water vibriosis, 359–363 enteric redmouth (ERM), 363–369 furunculosis, 345–353 management, 393–401 chemotherapy, 395–398 vaccination, 398–401 miscellaneous diseases, 392–393 overview, 345 piscirickettsiosis, 389–392 vibriosis, 353–359

Salmonid herpesviruses, 183–191 salmonid herpesvirus 1 (SalHV-1), 183–185 clinical signs, 184 diagnosis, 184 epizootiology, 184–185 geographical range and species susceptibility, 183 pathology, 185 significance, 185 virus characteristics, 184 salmonid herpesvirus 2 (SalHV-2), 185–189 clinical signs, 185, 186f diagnosis, 185, 187 epizootiology, 187–188, 188f geographical range and species susceptibility, 185 pathology, 188–189 significance, 189 virus characteristics, 187 Salmonid retroviruses, 194–196 Atlantic salmon paramyxovirus (ASPN), 195–196 plasmacytoid leukemia virus, 194–195 salmon swim bladder sarcoma virus, 195 viral erythrocytic necrosis, 196 Salmonids, drugs approved for, 69t Salmon leukemia virus, 194 Salmon pancreas disease/sleeping disease in rainbow trout, 191–193 clinical signs, 191–192 diagnosis, 192 epizootiology, 193 geographical range and species susceptibility, 191 pathology, 193 significance, 193 virus characteristics, 192–193 Salmon rickettsial septicemia, 80t Salmon swim bladder sarcoma virus, 195 Salmon viruses. See Trout and salmon viruses Saprolegniasis, 71 Scientific names of fishes, 473–475 Sea lice, 168 Seasonal trends, 32–33, 32f Segregation, protection by, 16–17 Sekiten-byo, 334 Serous inflammation, 44 17 α-methyltestosterone, 74 Sleeping disease, 191–193 Sockeye salmon, temperature requirements for, 9t Sodium bicarbonate, 72t Sodium chloride, 72t Sodium sulfite, 72t Specific pathogen free (SPF) stocks, 4, 12 Spiramycin, 75 Spring viremia of carp, 109–115 clinical signs, 110, 111f diagnosis, 110–113, 112f, 113t epizootiology, 114 geographical range and species susceptibility, 109–110 pathology, 114–115 significance, 115 stress mediated, 5t virus characteristics, 113–114

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Index 491

Stomatopapilloma, 141–143 clinical signs, 141 diagnosis, 141 epizootiology, 142 geographical range and species susceptibility, 141 pathology, 142 significance, 143 virus characteristics, 141–142 Streptococcosis, 445–452 bacterial characteristics, 448–449, 449t clinical signs, 446–447, 447f diagnosis, 447–448 epizootiology, 449–451 geographical range and species susceptibility, 446 pathology, 451–452 significance, 452 stress mediated, 5t Streptococcus agalactiae, 446–452 Streptococcus iniae, 60 probiotics and, 395 tilapias, 446–452 vaccines, 78t, 79, 85 Streptococcus spp., 74, 332, 435 Stress and disease, 4–5, 5t Striped bass, 74 bacterial diseases, 419–438 management, 436–438 chemotherapy, 437 vaccination, 437–438 miscellaneous diseases, 433–436 mycobacteriosis and nocardiosis, 419–428 overview, 419 photobacteriosis (Pasteurellosis), 429–433 temperature requirements, 9t Sturgeon viruses, 219–224 management, 223–224 miscellaneous virus diseases, 223 overview, 219 white sturgeon adenovirus (WSAV), 219–220 white sturgeon herpesvirus (WSHV-1, WSHV-2), 221–223 white sturgeon iridovirus (WSIV), 220–221 Subacute infection, 36 R Sulfadimethoxine and ormetoprim (Romet-30
, R Romet TC
), 69t, 70 R Sulfamerazine (Sulfamerazine in Fish Grade
), 69t, 70 Swim bladder inflammation (SBI), 113 Table of conversion factors, 477–478 Telangiectasis, 42 Tenacibaculosis, 465–469 bacterial characteristics, 467–468, 467t clinical signs, 466, 466f diagnosis, 466 epizootiology, 468 geographical range and species susceptibility, 465 management, 469 overview, 465 pathology, 468 significance, 469 Tenacibaculum maritimum, 465–469

Thiamine hydrochloride, 72t Thimphenico, 75 Tilapia, 74, 82 bacterial diseases, 445–459 francisellosis, 452–454 management, 456–459 chemotherapeutics, 457–458 vaccination, 458–459 miscellaneous diseases, 454–456, 455f–456f overview, 445 streptococcosis, 445–452 temperature requirements, 9t Title 50, 13 R Tricaine methanesulfonate (Finquel
and R Tricaine-S
), 69t, 71 Trout and salmon viruses, 147–199 aquareovirus diseases, 193–194 cutthroat trout virus, 194 epizootic epitheliotropic disease (EED), 189–191 epizootic hematopoietic necrosis (EHN) virus, 193 erythrocytic inclusion body syndrome (EIBS), 180–183 infectious hematopoietic necrosis virus (IHN), 147–156 infectious pancreatic necrosis (IPN) virus, 156–166 infectious salmon anemia (ISA) virus, 166–170 management, 196–199 avoidance, 196–197 prevention, 197–198 vaccination, 198–199 overview, 147 salmonid herpesviruses. See Salmonid herpesviruses salmonid retroviruses, 194–196 Atlantic salmon paramyxovirus (ASPN), 195–196 plasmacytoid leukemia virus, 194–195 salmon swim bladder sarcoma virus, 195 viral erythrocytic necrosis, 196 salmon pancreas disease/sleeping disease in rainbow trout, 191–193 viral hemorrhagic septicemia (VHS), 170–180 Tumor viruses, 253–254, 256, 258–262 Esox lymphosarcoma, 260–261, 260f factors that affect virus-induced tumors in fish, 262 miscellaneous, 261 tumors in walleye, 254, 254t, 256, 257f, 258–259 diffuse epidermal hyperplasia, 254, 255f, 256 discrete epidermal hyperplasia, 256, 258 walleye dermal sarcoma virus (WDSV), 258–259 Ulcer disease of goldfish, 315 Ulcer disease of winter, 5t USFWS, 52 Vaccination, 76–77, 78t, 79–86, 80t, 83f adjuvants, 82 antigens, 79, 81 methods of application, 82, 84–85 immersion. See Immersion treatments injection, 83f, 84–85 oral, 82, 84 overview, 76–77, 79

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492 Index

Vaccination (cont.) preparation of, 81–82 problems, 85–86 Vibrio alginolyticus, 353 Vibrio anguillarum, 52, 60, 62, 337, 353–358, 356t, 394 probiotics and, 395 vaccines, 77, 78t, 79, 80t, 85, 398–399 Vibrio carchariae, 353–354 Vibrio cholerae, 353 Vibrio damsella, 353 Vibrio harveyi, 353, 358 Vibrio ichthyoenteri, 353 Vibrio (Listonella) anguillarum in eels, 327 in marine fish in association with atypical Aeromonas salmonicida, 320 Vibrio mimicus, 353 Vibrio ordalii, 60, 353–355, 356t, 357–358 probiotics and, 395 vaccines, 77, 79, 80t, 81, 398–399 Vibrio parahaemolyticus, 353 Vibrio pelagiius, 353, 357 Vibrio salmonicida, 353, 356t, 359–361, 360f vaccines, 77, 78t, 79, 398–399 Vibriosis, 353–359 bacterial characteristics, 355–356, 356t clinical signs, 354, 355f diagnosis, 354–355 in eels, 337 epizootiology, 356–358 geographical range and species susceptibility, 353–354 pathology, 358 significance, 358–359 stress mediated, 5t tilapia, 445, 455 vaccines, 78t Vibrio splendidus, 52, 353, 357 Vibrio spp., 454 Vibrio vulnificus, 337–339, 353 Vinegar (acetic acid), 72t, 774 Viral diseases of fish carp viruses. See Carp and minnow viruses catfish viruses. See Catfish viruses diagnosis, 50–51 eel viruses. See Eel, viruses minnow viruses. See Carp and minnow viruses miscellaneous viral diseases of fish. See Viral diseases of fish, miscellaneous salmon viruses. See Trout and salmon viruses sturgeon viruses. See Sturgeon viruses trout viruses. See Trout and salmon viruses Viral diseases of fish, miscellaneous, 227–263 aquabirnaviruses, 247–248 epizootic hematopoietic necrosis (EHN), 227–230 EUS virus, 248 LMBV, 248–251, 249f lymphocystis, 230–231, 233–235 management, 262–263

miscellaneous herpesviruses, 251 miscellaneous iridoviruses, 251 overview, 227, 228t perch rhabdoviruses, 247 PHV disease, 251–253 blue spot disease of pike and muskellunge, 253, 253f herpesvirus of turbot, 252 pike fry rhabdovirus disease (PFRD), 244–247, 245f red sea bream iridoviral disease (RSIV), 242–244 tumor viruses. See Tumor viruses VEN, 235–240 viral nervous necrosis (VNN), 240–242 Viral erythrocytic necrosis (VEN), 196, 235–240 clinical signs, 236–237, 236f diagnosis, 237–238, 237f epizootiology, 238–239, 238t geographical range and species susceptibility, 236 pathology, 239–240 significance, 240 virus characteristics, 238 Viral hemorrhagic septicemia (VHS), 13–14, 170–180 clinical signs, 171–172, 172t, 173f, 174 detection, 174–176, 175f epizootiology, 177–179 geographic range and species susceptibility, 170–171, 171t pathology, 179 significance, 179–180 virus characteristics, 176–177, 176t Viral nervous necrosis (VNN), 240–242 clinical signs, 240–241 diagnosis, 241 epizootiology, 241–242 pathology, 242 significance, 242 species susceptibility and geographical range, 240 virus characteristics, 241 Virulence, definition of, 40 Walking catfish, temperature requirements for, 9, 9t Walleye temperature requirements, 9t tumors in, 254, 254t, 256, 257f, 258–259 diffuse epidermal hyperplasia, 195, 254, 255f, 256 discrete epidermal hyperplasia, 195, 256, 258 walleye dermal sarcoma virus (WDSV), 195, 258–259 Whirling disease, 17, 48 White sturgeon adenovirus (WSAV), 219–220 White sturgeon herpesvirus (WSHV-1, WSHV-2), 221–223 White sturgeon iridovirus (WSIV), 220–221 Yersinia ruckeri, 52, 60, 361f, 363–369, 365t–366t, 394 probiotics and, 395 vaccines, 77, 78t, 79, 398–399