Health Maintenance and Principal Microbial Diseases of Cultured Fishes

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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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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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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 <10◦ C High stocking density, elevated water temperature, handling, transport, poor water quality Low oxygen, handling, environmental stress Injury to skin, transport, improper handling, temperature stress, poor water quality, other parasites Handling and stocking in late early spring

Columnaris Cold-water disease Enteric redmouth Furunculosis Motile Aeromonas septicemia Ulcer disease of winter or goldfish and carp erythrodermatitis Vibriosis Streptococcicosis

5

Handling, poor environmental conditions, moving from freshwater to salt water Handling, poor water quality, parasites

Source: Walters and Plumb (1980), Piper et al. (1982), Roberts (1989), Wedemeyer (1996).

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concentration is adequate, fish may survive, and if oxygen is critically low and water temperature is normal, fish may also adjust and survive. Multiple stressful parameters have a compounding synergistic effect on the fish, and even though the fish may be able to handle the stressors individually, the combined effect can be lethal. An example of environmental stressors and their synergistic effect on fish involves dissolved oxygen (DO) and CO2 concentrations. Channel catfish can adapt to an elevated level of CO2 (20–30 mg/L) if the DO concentration is optimal (Boyd 1990). However, if the CO2 level is critically high and DO is critically low, fish cannot eliminate CO2 and will become listless (narcotized) and may die. If fish do adapt to these environmental stressors and survive, pathogen resistance is often compromised. Although salmonids require cooler water and higher oxygen concentrations, these factors are often offset in salmonid culture because in most instances the fish are reared in flowing water from springs or streams. These waters have more consistent temperatures and higher oxygen concentrations and lack high organic loads. A theory of host/pathogen/environment relationship was applied to fish with regard to development of infectious diseases by Snieszko (1973) (Figure 1.1). This theory is based on the premise that to have an infectious disease, a host and pathogen are required but an unfavorable environmental condition often acts as a trigger for disease to develop. Potential pathogens are often endemic in surface waters, especially in warm-water fish culture, and only environmental conditions and/or the hosts natural resistance can dictate onset of the disease process. The interaction of these factors is expressed in the equation: H(A + S2 ) = D, where: H = Species or strain of host (natural resistance)

Pathogen

Host Species

Infectivity

Age Virulence

Strain

Pathogenicity

Nutritional status

Viability

Population density

Strain

Disease potential

Environment Temperature Oxygen concentration Water alkalinity Water hardness Toxicants Season Figure 1.1 Variables of infectious agents, host, and environment that influences disease occurrence. (Adapted from Snieszko (1973) and Schnurrenberger (1983a).)

A = Etiological agent S = Environmental stressors D = Disease Environmental stressors are squared because as fish approach adaptation limits, stressors increase accumulatively rather than additively. Also, when more than one stressor is involved (oxygen, ammonia, CO2 , temperature, etc.), detrimental factors act synergistically. There is a relationship between water quality deterioration and bacterial infection. A sudden die-off of cyanobacteria (blue-green algae) in a channel catfish pond was followed by reduced DO production, decreased pH, and increased CO2 and NH4 (Plumb et al. 1976). These water quality changes in the pond resulted in “oxygen depletion” and

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7

10

10

8

9

6

8

4

pH

Dissolved oxygen (mg/liter)

12

7 Oxygen

2 0

Skin lesions on fish A. hydrophila isolated from fish

pH

Phytoplankton died

18 22 APRIL

26

30

4 MAY

8

12

6 5

16

(a) 1.8 Ammonia nitrogen Carbon dioxide

1.4

14

1.2

12

1.0

10

.8

8

.6

6

.4

4

.2

Skin lesions on fish

0

Phytoplankton died

26 28 APRIL

30

2 MAY

4

A. hydrophila isolated from fish

Carbon dioxide (mg/liter)

Ammonia nitrogen (mg/liter)

1.6

2 0

6

8

10

12

14

(b) Figure 1.2 Water quality parameters before, during, and after a channel catfish mortality and subsequent Aeromonas hydrophila infection (Plumb et al. 1976). (Printed with permission of Journal of Wildlife Diseases.)

a fish kill (Figure 1.2). This phenomenon has since been described by R. Schmittou (Department of Fisheries and Allied Aquacultures, Auburn University, Alabama, personal communication) as “low dissolved oxygen syndrome” (LODOS), which refers to the fact that low DO is part of an environmental condition that includes a variety of separate but interrelated elements. When fish first

began to die as DO dropped below 1 mg/L, no bacteria or other significant pathogens were found during necropsy. However, 4 days after oxygen depletion, channel catfish were found with hemorrhaged and depigmented skin and muscle lesions (Figure 1.3). When first observed, no bacteria were isolated from internal organs or skin–muscle lesions of these fish, but 2 days later and for several days thereafter,

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Figure 1.3 Channel catfish with depigmented, hemorrhaged, and necrotic lesions that appeared 6 days after oxygen depletion (Plumb et al. 1976). (Printed with permission of Journal of Wildlife Diseases.)

109 Number of bacteria/g of tissue (log 10)

Aeromonas hydrophila was isolated from both. When freshwater was added to the pond and remedial aeration provided, mortality ceased and clinical signs of infectious disease abated. It was theorized that while the water had low oxygen concentrations, some muscle areas in the fish became hypoxic, which led to tissue necrosis, hemorrhaging, and skin depigmentation. When protective epithelium integrity was lost, naturally occurring opportunistic A. hydrophila invaded the muscle beneath areas where skin had been injured and focal infections were established, which progressed into septicemia. Walters and Plumb (1980) demonstrated that low oxygen, low pH, high ammonia, or high CO2 alone was not particularly stressful and did not lead to bacterial disease. However, if two or more of these adverse environmental conditions occurred simultaneously, infection was much more likely to occur (Figure 1.4). Intensive fish culture causes unique but manageable environmental problems for the aquaculturist (Piper et al. 1982; Boyd 1990; Wedemeyer 1996). All fish require adequate water maintained at a suitable temperature and oxygen concentration level for proper growth and reproduction. Water temperature

108

a

a

(8)

(5) a

107

(8)

106 105 104 103

(4) (12)

(10)

VI

VII

(24)

102 101

I

II

III

IV Group

V

Figure 1.4 Number (CFU) of bacteria per gram of trunk kidney from channel catfish held in various environmental conditions. (i) Low dissolved oxygen (DO) only; (ii) Low DO, fish injected with A. hydrophila; (iii) Low DO, fish injected with A. hydrophila, NH3 added; (iv) Low DO, fish injected with A. hydrophila, NH3 , and CO2 added; (v) Low DO, fish injected with A. hydrophila, CO2 added; (vi) Aeration, fish injected with A. hydrophila; (vii) Noninjected fish in aerated water. Number of fish samples in parentheses; significantly higher numbers of bacteria are designated by “a” (Walters and Plumb 1980). (Printed with permission of the Fisheries Society of the British Isles.)

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Table 1.2 Range of temperature tolerance, optimum temperature for growth, and spawning temperature of selected cultured fishes. Temperature Commmon Name

Range

Optimum

Spawning

Atlantic salmon Brook trout Brown trout Channel catfish Chinook salmon Coho salmon Common carp Eel Grass carp Lake trout Largemouth bass Milkfish Northern pike Rainbow trout Sockeye salmon Striped bass Tilapias Walleye Walking catfish

1–24 1–22 1–25 4–35 1–25 1–25 4–35 4–35 4–35 1–21 4–35 10–35 1–27 1–25 1–21 2–32 15–35 1–27 13–38

10–17 8–13 9–17 28–30 10–14 9–14 23–30 25–28 22–30 7–14 13–27 25–35 4–18 10–17 10–15 13–24 23–32 8–16 20–30

7–10 8–13 9–13 25–27 8–13 8–13 13–27 16–17 22–27 9–11 16–20 23–32 4–9 10–13 8–12 13–22 23–32 9–13 20–30

Source: Ney (1978), Piper et al. (1982).

requirements will vary from species to species (Table 1.2). For example, channel catfish require a water temperature of 20–30◦ C for growth (optimum of 28◦ C), and they spawn at 27◦ C. They prefer oxygen concentrations above 5 mg/L. They will survive at 1–5 mg/L of oxygen, but prolonged exposure below 1 mg/L is lethal. If oxygen levels drop below a critical concentration, supplemental aeration is necessary as either a routine practice or as emergency management (Figure 1.5). For comparison, rainbow trout require water temperatures of 8–18◦ C for growth, spawn at temperatures as low as 6◦ C, and require oxygen concentrations above 5 mg/L. At the other extreme, walking catfish of Southeast Asia can survive water temperatures above 35◦ C and very low oxygen concentrations. However, these fish have an auxiliary gill that allows them to extract oxygen directly from the air. Extreme high or low water temperatures may result in greater fish susceptibility to viral, bacterial, and parasitic diseases (Austin and Austin 1987).

For a fisheries manager, biologist, or diagnostician to understand infectious fish diseases, he/she must also understand how and to what extent the environment affects the host and disease. It is not enough to simply identify a pathogen or parasite; it is equally important to identify and understand environmental stressors that predispose fish to disease. When stressors are known, corrective and preventive measures can often be initiated to help prevent or minimize a reoccurrence of the condition. In the aquatic world, maintaining an optimal environment is essential to good animal health. Extreme deviations will result in stress manifested by reduced feeding, higher feed conversions, poor growth, disease, or death.

Location, soil, and water Choosing a proper site for an aquaculture facility is paramount to its success. Land topography, soil quality, water quality and abundance, and proximity to market are primary

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(a)

(c)

(b)

Figure 1.5 Routine and emergency aeration with (a) water pump attached to tractor power takeoff, (b) electric paddle wheel in a catfish pond (could also be powered by tractor power takeoff), and (c) aeration by flowing water over a column of perforated plates.

factors to be considered when choosing a location. To maintain healthy fish populations, soil, water, and fish species to be grown must be compatible. Sometimes environmental modifications can be made to accommodate a species not indigenous to a specific area but these modifications must be cost effective. For example, channel catfish are not normally grown in northern latitudes of the United States because of a short growing season and lack of warm water. However, if artificially heated water from power plants or geothermal supplies are available, channel catfish or other warm-water species can be grown successfully, although seldom economically. Another example of culturing fish species out-

side of their normal geographical range are tilapia, which require water temperatures of over 18◦ C for survival and over 24◦ C for optimum growth. However where adequate volumes of artificially heated water is available these fish can be successfully grown in temperate and cool regions. Some soil requirements for culture ponds are very specific. Soil must have a high clay content to prevent ponds from leaking; leaky ponds are unstable, do not retain a suitable plankton bloom to shade out rooted vegetation, and require continuous water replenishment. Conversely, in an analysis of chemicals naturally present in soil, Boyd (1990) stated that it is possible to rear fish in ponds built

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Table 1.3 Water quality criteria for optimum fish health management of warm water and cold water species of fish (mg/L except for pH). Characteristic

Cold Water

Warm Water

Oxygen pH Ammonia (un-ionized) Calcium Carbon dioxide Hydrogen sulfide Iron (total) Manganese Nitrate Phosphorus Zinc Total hardness (CaCO3 ) Total alkalinity (CaCO3 ) Nitrogen (gas saturation) Total solids

5-saturation 6.5–8 0–0.0125 4–160 0–10 0–0.002 0–0.15 0–0.01 0–3.0 0.01–3.0 0–0.05 10–400 10–400 <100% 0–80

5-saturation 6.5–9 0–0.02 10–160 0–15 0–0.002 0–0.5 0–0.01 0–3.0 0.01–3.0 0–0.05 10–200 10–400 <100% 50–500

Source: Piper et al. (1982), Boyd (1990), Hajek and Boyd (1994).

on soils that have wide ranges of chemical properties. Certain regions have acid-sulfate soils containing high levels of iron pyrite (Boyd 1995). As long as these soils are submerged, the iron pyrite is stable and usually causes no problem. However, when the pond is drained and the bottom exposed to air, iron pyrite is oxidized and upon dehydration sulfuric acid is produced. Upon refilling the pond, the water becomes highly acid (pH may be as low as 3.5), rendering an unproductive environment. Generally, such areas should be avoided for aquaculture sites, but the acidity level can be corrected to some degree by addition of huge quantities of lime. It is important that pond soil is free of chemical and pesticide residues. If toxicants, usually of anthropogenic origin, are present, they can leach into the water and kill fish. Concern is not only with soil used in pond construction but also with soil in the watershed; therefore, prior to construction of a culture facility, watersheds should be inspected for toxicants that could contaminate the water supply. The most important ingredient in successful fish health management is water quality

and its availability. Water quality as described by Boyd (1990) varies, but most water can be made suitable for aquaculture, except under unusual circumstances. Water hardness, alkalinity, pH, presence of toxicants, or dissolved gases are important in aquaculture (Table 1.3). Before construction of a fish farm, water quality, volume, and reliability of the water source must be determined. Water sources for aquaculture may be lakes, rivers, springs, pumped or artesian wells, surface runoff, or irrigation canals. From a fish disease management standpoint, springs or wells are preferred because these sources are free of wild fish that may be carriers of infectious disease agents. However, from a water quality standpoint, these sources may not be usable without modification because of acidity, low DO, and/or high concentrations of CO2 or nitrogen gases, which require removal. These waters may be very soft (less than 50 mg/L CaCO3 ) or contain high concentrations of iron, sulfur, or manganese (Table 1.3). Although well and spring water may have a constant temperature, it may not be optimum for the aquaculture species to be cultured, thus requiring heating or cooling in the case of

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geothermal sources. If pumping is necessary to utilize a water source, it can be expensive and prone to mechanical breakdowns and interrupted water flow. However, most water quality problems can be overcome by proper planning and management. When streams or reservoirs are used as a water supply, there are inherent problems involved. Wild fish, including fry and eggs that harbor parasites, pathogenic bacteria, or viruses may be indigenous to these sources. These waters can be disinfected with ozone or ultraviolet light, but the efficacy of treatment depends on the physical nature of the water. For example, ultraviolet treatment is ineffective in silt or particulate-laden water. Also, surface waters including streams, reservoirs, and irrigation canals may be prone to wide seasonal temperature fluctuations, variable oxygen levels, silt loads, increased organic loads, volume fluctuations, or contamination with pollutants from municipal, industrial, or agricultural sources. Installation of sand and gravel filters or use of porous (saran) socks on inlet pipes are some options to prevent fish contamination.

Avoiding exposure The ideal way to control infectious fish disease is to prevent exposure to pathogenic agents whenever possible, thus avoiding most devastating health problems through biosecurity (Bebak-Williams et al. 2007). However, when dealing with the aquatic environment, it is virtually impossible to define all disease-causing agents and to keep them isolated from the fish host. Water provides an excellent medium for transfer of many communicable agents from fish to fish or from locality to locality. Moreover, many disease-causing organisms are endemic to the aquatic environment and are opportunistic, facultative pathogens that remain viable under various conditions. Since the mid-1960s some governments, fish hatcheries, and privately owned fish farms

have made great strides in avoiding fish exposure to certain infectious diseases by using fish certified as specific pathogen free (SPF), quarantine, routine water disinfection, and by destroying populations that have become infected with specific pathogens. The pros and cons of the latter approach should be carefully weighed before making a decision to destroy or treat the fish. Paraphrasing a statement by Snieszko, “A disease management practice should not destroy more than it saves.” The earliest attempt to prevent fish exposure to infectious pathogens was when trout stocks known to be positive for infectious pancreatic necrosis virus (IPNV) were not used as egg sources; a practice that is now applied to other diseases. It is impossible to declare a fish group, or population, simply “disease free,” implying they are free of all disease agents; therefore, the term “disease free” is limited to a specific pathogen. Currently, Specific Pathogen Free (SPF) certification is the best way to prevent introduction of unwanted pathogens into a “clean” facility, but it must be understood that testing procedures are not infallible because they are based on a statistically determined sample number of individuals taken from a larger population (AFS-FHS 2007). To detect a given pathogen with 95% confidence, the appropriate sample number for a 2% prevalence is 120 fish per 100,000 and for a 5% prevalence, 60 fish per 100,000 (Simon and Schill 1984). However, Thorburn (1996) indicated that these numbers may not be uniform for all diseases and all fish species. In some instances a larger number than recommended should be sampled to improve accuracy. Fish pathogen detection methods for determining SPF populations are discussed further in Chapter 3. Many states have now established fish health protection programs that specify that incoming fish must be accompanied by a document certifying them SPF. California has one of the oldest and most rigorous fish health regulatory programs in the United States. Regional fish health plans have also been

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established to prevent introduction of unwanted pathogens into geographically defined fish populations. The Colorado River Basin Council has a strong inspection program to prevent introduction of fish that are infected with certain disease agents into natural and/or hatchery waters. The Great Lakes Fisheries Commission initiated a regional fish health protection program for the states and Canadian provinces contiguous to the Great Lakes. The major problem with state and/or regional fish health regulations is statutory inconsistency and degree of implementation. However, they are a step in the right direction to prevent the spread of some fish diseases. All Nordic countries have established coordinated surveillance and monitoring systems for fish diseases of concern to the fish farming industry (Hastein et al. 2001). The ultimate goal of the program is to eradicate or keep the level of disease to a minimum. According to the authors, due to these surveillance systems coupled with good management practices the fish disease situation in the Nordic countries is generally good compared to other countries of the world. Diseases included in the surveillance are viral hemorrhagic septicemia, infectious hematopoietic necrosis, infectious pancreatic necrosis, viral nervous necrosis, bacterial kidney disease, and furunculosis. However, remedial measures to be taken when a disease is found depend on the country in which it is found. International fish disease control concepts are gaining support to keep pace with a growing, world-wide aquaculture industry. An increasing number of countries now require health certificates of some type verifying that certain imported fish products (alive or dead) are free of specific diseases. While not fool proof, these methods have generally been successful in inhibiting the spread of many infectious disease agents. Great Britain, the European Economic Union, Canada, and the United States all have regulations limiting fish movement with emphasis on fish health. Great Britain probably introduced the first fish health

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regulation by passing the Diseases of Fish Act 1937 (Hill 1996) with several subsequent amendments. It is believed that it’s continuous implementation has had a positive effect on the United Kingdom’s fish industry. In the United States, Title 50 applies to Injurious Wildlife and requires that fish and/or fish eggs imported into the United States be certified free of certain pathogens. The European Union has directives that provide guidelines that must be met before aquaculture products are shipped into their sovereign territories to insure protection against introduction of exotic diseases (Daelman 1996). The European Union also has an operative fish disease control service that requires that all aquaculture facilities be inspected and registered. However, some countries such as Japan, consider aquaculture practices to be too diverse and extensive for implementation of an effective national disease control program (Wakabayashi 1996). Quarantine is an approach to disease avoidance when fish are moved from one area to another. Fish should be isolated for a specific period of time before contact with a resident population. If disease develops in newly arrived animals, it can be dealt with more effectively and without exposing resident stocks. Drastic measures such as eradication of an entire fish population from an aquaculture facility are sometimes necessary to avoid exposing healthy fish to highly infectious, nonendemic disease agents. When contemplating the eradication of a fish population, the economic, environmental, and biological significance of the disease should be considered. Is the disease indigenous to the area? Can it be adequately controlled through management or chemotherapeutics? In other words, is eradication worth the long-term potential savings in fish and can the facility be maintained free of the disease organism? In 1989, an attempt was made to prevent establishment of viral hemorrhagic septicemia virus (VHSV) in the United States by destroying adult salmonids as they returned to spawn at two sites in the Pacific Northwest where

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the virus had been found (Hooper 1989). This constituted the first confirmed VHSV occurrence outside Europe and so these drastic measures appeared to be justified. However, in subsequent years, the virus was found in different areas of northwestern United States, so eradication did not solve the problem (Winton et al. 1991). Viral hemorrhagic septicemia was detected in the Great Lakes region of Canada and the United States in 2005 (Paul R. Bowser, Cornell University, personal communication). Destruction of lake trout and disinfection of contaminated hatcheries in the Great Lakes region of the United States and Canada in the late 1980s was apparently successful in eliminating epidermal epitheliotropic disease, a severe herpesvirus infection of juvenile lake trout (R. Horner, Illinois Department of Natural resources, personal communication); however, the disease eventually showed up in other locations. Also, a second fish eradication occurred in 1990 when salmonid brood stock at Jackson National Fish Hatchery (Wyoming) (Anderson 1991) and White Sulfur Springs National Fish Hatchery (West Virginia) (Cipriano et al. 1991) were killed because they were subclinically infected with Renibacterium salmoninarum (bacterial kidney disease, BKD). These two facilities were major egg suppliers for many federal and state agencies and could potentially contribute to the spread of BKD by egg distribution. Therefore, an administrative decision was made to destroy large numbers of valuable rainbow, lake, cutthroat, and brook trout. This decision was based on a highly sensitive ELISA method for detecting very low numbers of R. salmoninarum. The fish were killed in spite of the fact that neither facility demonstrated any evidence of clinical BKD. Snieszko’s theory on the application of fish health management practices comes to mind: “Was the cost and consequence of implementation greater than the value of what was saved?” Although infectious disease prevention is best accomplished through avoidance whenever possible, there are times when one must

look beyond the host for a disease source. Some disease agents (helminths, for example) have complex life cycles that involve fish-eating birds, snails, or copepods, and the pathogen life cycle is broken by controlling non-fish vectors, a nearly impossible task. Another example of a disease agent that may have a non-fish vector is IPNV. This virus has been shown to be infectious after passing through the intestines of fish-eating birds (Peters and Neukirch 1986). It would be extremely difficult to prevent exposure of fish to these contamination sources in an endemic area. Each disease outbreak must be considered individually and rarely will a general policy be applicable; therefore, “avoiding exposure” decisions must be made using biological facts and common sense, with an eye to the overall good of the aquatic environment.

Exposing dose As previously noted, total avoidance of infectious agents is the desirable approach to disease prevention; however, this is not always possible or practical because many organisms that infect fish are normally free-living, facultative, and opportunistic pathogens. Fish and some pathogens can coexist without disease unless the fish’s immune system or other defensive mechanisms are compromised. Aeromonas hydrophila, the bacterial agent of motile Aeromonas septicemia (MAS), is a ubiquitous organism that occurs naturally in most freshwaters of the world. It is capable of living and proliferating in any water containing organic enrichment. The bacterium usually causes disease only after a fish’s resistance has been compromised by environmental stressors, or when fish have suffered some mechanical or biophysical injury. Thus, the best approach to prevent MAS is to reduce organic load and bacterial numbers in the water and to keep fish at a high state of resistance by environmental management. Prophylactic treatments after fish are handled will aid in

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healing superficial wounds and in reducing bacterial populations on the skin. In a normal, healthy fish population, a certain percentage of fish may possess systemic A. hydrophila; however, disease occurs only when bacterial numbers overwhelm the fish’s resistance. Many fish, wild or cultured, normally have some parasites present on their gills or skin. As long as these parasites, either protozoa or monogenetic trematodes, are present in low numbers, there generally is no health problem; however, if water quality deteriorates to a point that fish are stressed, parasite numbers will increase. Parasite populations can be reduced with prophylactic chemotherapy and good health reestablished with restoration of environmental quality. Culling old brood stock and using younger fish for reproduction before they become heavily parasitized can reduce the effects of metazoan parasites. Using young largemouth bass as brood stock has proved successful in the southeastern United States where bass tapeworm (Proteocephalus ambloplitis) larvae can be a problem. This parasite can damage the ovaries of adult fish if the parasite is present in large numbers. The damage can cause a reduction in egg production that is cumulative with age (W. A. Rogers, Auburn University, Alabama, personal communication). Most infectious agents require specific levels of infective units before adversely affecting a host’s health status. Therefore, if infectious organisms can be maintained at levels below disease threshold, losses will be reduced.

Extent of contact Diseases that involve opportunistic (facultative) organisms are not germane to this particular discussion; therefore, the following remarks pertain to obligate pathogen-induced fish diseases (viruses, some bacteria, and most parasites). In managing such diseases, initial consideration is given to the pathogen source in the culture system.

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Portal of entry

Route of transmission

Reservoir (fish)

Direct, vehicle vector

Portal of exit Direct, vehicle vector Route of transmission

Portal of exit Susceptible host (fish) Portal of entry

Figure 1.6 Typical transmission cycle for communicable disease agents. The direct vehicle vector may be water, food, contaminated equipment, eggs, etc. (Adapted from Swan (1983).)

Pathogens must have a route of transmission from source to susceptible host, which may be by direct contact from fish to fish, via the food chain, through water, on nets, buckets, and other gear, or via reproductive products (Figure 1.6). Infected fish that survive can become pathogen carriers and a reservoir for disease agents. There are numerous examples of fish diseases (particularly viruses) where epizootics are associated with a reservoir host. A pathogen must have a way to escape the host; in fish this can be accomplished by shedding virus or bacteria across the gill membranes, through skin mucus, from skin lesions, in feces, urine, or via reproductive products. Internal metazoan parasites often must wait until the host fish is eaten by an intermediate or final host to complete its life cycle. Nematode and trematode parasite larvae that live in the eyes of some fish are released when the eye is destroyed. Some pathogens, especially viruses, are released from a reservoir host during spawning via eggs, ovarian fluid, or milt. In some viral diseases, the number of virus-shedding fish actually increases during spawning time (infectious hematopoietic necrosis virus, IHNV),

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which aids transmission (Mulcahy and Pascho 1985). Bacteria and parasites can be transmitted to cultured fish populations through feed that contains raw wild fish flesh. Marine fish are an especially good source for these pathogens. Mycobacteria (acid fast staining bacteria) are transmitted to uninfected fish when uncooked fish flesh or viscera is incorporated into their feed. Transmission of mycobacteria was common in Pacific salmon hatcheries in the 1950s and 1960s when ground raw marine fish was incorporated into hatchery feeds. Another example of a parasite being transmitted by feeding contaminated fish flesh occurred in Florida when chopped Atlantic menhaden, infected with Goezia (a nematode that inhabits the stomach and intestinal wall), was fed to cultured striped bass fingerlings (W. A. Rogers, Auburn University, personal communication). The infected striped bass were then stocked into Florida’s inland lakes where the parasite was transmitted to resident tilapia and largemouth bass. Although virus transmission resulting from feeding raw fish has not been reported, most virulent fish viruses have been transmitted orally under laboratory conditions. While infectious disease-induced mortalities in wild fish populations are rare, high losses are much more common in cultured fish populations. Like other animals, fish are more susceptible to infection when crowded. As fish density increases, rate of contact and pathogen transmission increases accordingly. Therefore, in high-density fish populations, many epizootics have a potential for reaching catastrophic proportions. Many fish disease organisms do not cause clinical disease until infected fish are placed in stressful conditions. Pathogens can be introduced into an aquaculture facility by humans, either staff or visitors. In order to prevent this mode of transmission fish hauling trucks should be disinfected before entering a facility (Bebak-Williams et al. 2007). Also footbaths containing disinfectant

placed between fish holding areas should be available to kill any pathogens on boots, etc.

Protection by segregation Segregating fish according to species and age can reduce transmission of disease-producing agents because some of these organisms are not equally pathogenic to all fish species, or all strains or ages within a susceptible species. Some fish disease-producing organisms lack host specificity, while others are more host specific. Nonhost-specific, opportunistic freeliving bacteria are generally facultative. Some diseases, such as furunculosis (Aeromonas salmonicida) and most viral agents are generally more host specific. Brook trout are considered highly susceptible to A. salmonicida and should be reared completely segregated from carrier populations. Rainbow trout and brown trout are also susceptible to A. salmonicida, but to a lesser degree than brook trout, and may be considered for culture where this disease is endemic. Rainbow trout and some species of Pacific salmon are highly susceptible to IHNV; therefore, culture of a less susceptible salmonid species should be considered where this virus occurs. Channel catfish are the most susceptible species to channel catfish virus disease (CCVD). However, within channel catfish stocks, inbred strains are more susceptible to CCVD than are “out-bred” strains (Plumb et al. 1975). In areas where CCVD is endemic, less susceptible catfish species, blue catfish for example, can be cultured even though they are not totally CCV resistant. The same catfish susceptibility pattern exists for Edwardsiella ictaluri, the bacterial causative agent of enteric septicemia of catfish (ESC) (Wolters et al. 1996). Generally, young fish have greater pathogen susceptibility than do older fish, particularly with regard to viruses and some parasites. In the case of IPNV and IHNV of salmonids and CCV of catfish, fish less than 3 months old

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are highly susceptible, but disease effects diminish with increasing age. If these fish species can be reared in noncontaminated water until past the critical age of susceptibility, severe losses can be avoided. The same is true with Myxobolus cerebralis (whirling disease), a parasite that infects young trout before the cartilage has matured into bone. Susceptible fish should be reared in well or spring water free of wild fish, but after fish have grown past a susceptible size or age, they can be transferred to less protected waters. Segregation for disease avoidance may also take into account water temperature because it affects the disease-causing ability of some fish pathogens. Infectious hematopoietic necrosis virus normally occurs at a temperature of 10–17◦ C; thus, if fish can be reared in water above 17◦ C, effects of disease is minimized. However, at elevated temperatures, trout may be more susceptible to other disease organisms and growth rate will be reduced. Channel catfish virus is most severe at temperatures above 25◦ C, therefore, if catfish are held at cooler temperatures, CCV can be avoided but growth rate will be reduced because of the suboptimal temperature. Unfortunately in most instances, optimum temperature for pathogens is also optimum for hosts. However, a combination of susceptible age and optimum temperature may be factors that are used to determine when strict segregation of young fish from the rest of the stock can be modified. Temporal segregation can also be an important factor in reducing disease-associated losses. This involves depopulating fish holding facilities between batches to prevent the transmission of pathogens carried by one crop to the next. Current channel catfish culture involves a continuous production system where fish are partially harvested for market (graded according to size) and then fingerlings are added to the pond to replace the number of fish harvested. This type of system perpetuates endemic pathogens and the newly arriving fish will bring their own pathogens (and strains) that may exacerbate disease problems. The use

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of batch systems (all in from one source and all out at one time) greatly reduces the opportunity for pathogen accumulation in a production system. To be effective, all fish must be removed from the production system and water source and the system disinfected before restocking.

Problem of new arrivals Newly arrived fish may bring pathogens with them, either in an active or carrier state, to which the resident population has not been exposed. Therefore, no new animals should be introduced into an existing population until reasonably certain they will not be detrimental. This rule applies to home aquaria, movement of fish from farm to farm, and to interstate and international live fish shipments. Even if the new arrivals carry the same pathogens as the farm population, there can be substantial differences in strains of the pathogen. The new strains may have different ability to cause disease or different host ranges, allowing more severe disease outbreaks to occur or they may have different antibiotic-resistance profiles, making treatment more difficult. Furthermore, new pathogen strains may have different antigen profiles that can allow the pathogen to infect fish that have developed immunity to the endemic strains of pathogen or to infect and cause more severe disease in fish vaccinated against the endemic strain. Several precautions to reduce potential for introducing disease with new arrivals are use of SPF eggs or be familiar with health history of fish to be introduced, know if any serious disease problems occurred, and if so what type of treatment, if any, was given. If not treated before transport, newly arrived fish or eggs should be given a prophylactic treatment with appropriate drugs to remove any external pathogens. When possible, new fish should be segregated (quarantined) from the resident population until shown to be disease free. Fish farmers must be extremely careful

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when replenishing brood stocks with outside sources to insure that no new pathogens are introduced into their facility. This danger is especially prevalent if fish from wild populations with unknown disease histories are used to replace brood stocks. Certain fish production facilities produce certified disease-free fish or eggs (certified to be free of known obligate pathogens). Disease free salmonid eggs have been extensively marketed (certified free of IPNV, IHNV, VHSV, and some salmonid bacterial diseases), but this is not 100% reliable. Because of the production systems, lack of legal restraints to shipping and limited number of pathogens to target, no facilities currently market diseasefree eggs or fry for warm-water aquaculture systems. Specific disease-free certification requires certification by an external authority. Sampling and diagnostic tests used are specified by the most current edition of “FHS Blue Book: Suggested Procedures for the Detection and Identification of Certain Finfish and Shellfish Pathogens” (AFS-FHS 2007), the “OIE Manual of Diagnostic Tests for Aquatic Animals” (OIE 2009) or must conform to “Code of Federal Regulations—Title 50: Wildlife and Fisheries”. In order to stop movement of disease agents, one must stop movement of infected animals. The prudent fish farmer must take every precaution available when acquiring new stock.

Breeding and culling Domestication of wild animals has been the foundation of successful animal husbandry throughout the ages. Selection, culling, and crossbreeding has been vital in developing today’s herds and flocks of domesticated animals. Culturing fish for food has been practiced for thousands of years in some areas of the world, especially in China. However, comparatively few studies have dealt with development of disease-resistant fish through selective

breeding and genetic manipulation. In most genetic fish improvement experiments, objectives have been to increase growth rates and fecundity or to improve feed conversions; therefore, any disease-resistance traits have resulted from happenstance. Different strains of channel catfish express different susceptibility to channel catfish virus disease and ESC (Plumb et al. 1975; Wolters and Johnson 1994; Bilodeau-Bourgeois et al. 2007). But few studies have investigated genetic selection or genetic manipulation in order to develop strains of fish with improved disease resistance. Peterson et al. (2008) demonstrated that certain selected strains of channel catfish not only had superior growth performance but also were more resistant to Edwardsiella ictaluri (ESC). Recently, transgenics has shown the potential to improve disease resistance. Dunham et al. (2002) enhanced disease resistance of transgenic channel catfish into which cecropin B gene from the cercropia moth had been inserted. In a pond epizootic of Flavobacterium columnare (columnaris) infection, 40.7% of the transgenic fish expressing the cecropin B gene survived compared to only 14.8% of the controls. Furthermore, 100% of the transgenic survivors possessed the cecropin B gene. Bilodeau-Bourgeois et al. (2007) points out that there is a variation in ESC susceptibility between strains and results from experiments in aquaria may not hold true in the pond environment. Lack of available scientifically developed disease-resistant brood fish should not prevent the aquaculturist from improving his stocks by gradually selecting fish that are less affected by specific disease outbreaks. If a particular lot of brood fish routinely produces offspring that develop a specific disease year after year (i.e., CCVD), then replacing those brood fish should be strongly considered; however, antidotal evidence of this practice suggests that success has been mixed. Routine culling of inferior individual brood fish will also improve performance.

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Nutritional basis of health maintenance Proper nutrition is essential for survival, growth, and reproduction of any animal species, and nutrient requirements of fish are similar to those of other animals (Lovell 1989). This applies to wild or cultured fish populations although nutritional management in the two systems is entirely different. In aquaculture, the fish’s nutrition is generally dependent on the quality and quantity of feed provide to the fish, but in wild fish populations, proper nutrition is dependent upon the natural food chain and availability of primary nutrients. Nutrients can be supplemented in these populations by addition of organic or inorganic fertilizers to stimulate growth of macroflora (vascular plants), microflora (phytoplankton), zooplankton, and other invertebrate fish food organisms. In some parts of the world, traditional aquaculturists continue to use manure and/or ingredient-based nutrient sources to feed cultured fish. This practice causes increased levels of organic matter in semiliquid layers of unoxygenated sediments, greater water quality problems, poor fish growth, higher feed conversion ratios, and higher incidences of infectious disease, especially stress-induced epizootics. More advanced aquaculture management practices advocate the use of manufactured feeds that contain proteins, fats, carbohydrates, fiber, vitamins, amino acids, and minerals. A deficiency in any one of these nutrients can result in nutritionally related disease or lowered disease resistance. The amount of each nutrient needed for proper growth depends upon species and age of fish being fed and intensity of the culture system. Fresh feed of the highest quality available should always be used. It has been shown that certain nutritional deficiencies can result in specific pathological conditions (Halver 1972). Injury to the spinal column of channel catfish (broken-back syndrome) results from a vitamin C deficiency.

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Lack of sufficient riboflavin can cause eye cataracts in rainbow trout and possibly in channel catfish. A niacin deficiency increases sensitivity to ultraviolet irradiation and “sun burn” on fish in clear water. Nutritional gill disease of trout is caused by a pantothenic acid deficiency and can progress into bacterial gill disease if the deficiency is not corrected. Also, severe anemia in channel catfish is linked to dietary deficiency of folic acid or presence of the folic acid antagonist pteroic acid (Butterworth et al. 1986). Several researchers have shown that a deficiency of specific elements in a fish’s diet can increase disease susceptibility and megadoses may increase immune response. Channel catfish fed a vitamin C-deficient diet were more susceptible to Edwardsiella tarda and E. ictaluri (Durve and Lovell 1982) and megadoses of vitamin C (five times the recommended 60 mg/kg of diet per day) enhanced immune response (Li and Lovell 1985). However, these studies were conducted in aquaria and when the same principles were applied to pondreared channel catfish, vitamin C-deficient diets did not affect susceptibility nor did megadoses of the vitamin enhance immunity (Liu et al. 1989). The attributes of organic zinc sulfate vs. inorganic zinc sulfate in a fish diet has received significant attention. Paripatananont and Lovell (1995) reported that channel catfish required less organic than inorganic zinc sulfate and that resistance to Edwardsiella ictaluri increased with addition of organic zinc sulfate in the diet. Contrastingly Lim et al. (1996) reported that the zinc source was unimportant and did not change disease resistance of channel catfish. Fish in poor nutritional condition (starved or lightly fed) may be more resistant to some infectious agents than are well-fed fish. Antidotal observations indicate that channel catfish susceptibility to CCV is decreased following starvation for 2 weeks (unpublished). More recently Kim and Lovell (1995) showed that

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channel catfish that were not fed at all during winter were significantly less susceptible to E. ictaluri in the spring than fish fed everyday or less frequently during the winter. Taking channel catfish off feed when ESC occurs has been practiced throughout the catfish industry. Fish that continue to be fed suffer higher mortality than do either medicated or nonfed populations (Wise and Johnson 1998). To what degree nutrition affects disease resistance in fish remains unclear. Subtle effects of a nutritionally deficient diet are reduced growth and poorer feed conversion; reduced fecundity and all dietary factors have the potential to affect health status and disease susceptibility of fish (Lovell 1989). When water quality becomes critical with high organic fertility and low DO, reduced feeding can hasten a reversal of this trend; therefore, flexibility in feeding can be a valuable tool in aquaculture management.

Eradication, prevention, control The terms eradication, prevention, or control are defined as follows: Eradication is the complete elimination of a disease-causing agent from a facility or specific geographical region, while practical eradication is elimination of an agent from its reservoir of practical importance. Prevention is avoiding introduction of a disease-producing agent into a region or facility and/or stopping a disease process before it becomes a problem. Control involves reduction of a problem to a level that is economically and biologically manageable, and/or confinement of a problem to a defined area. Eradication of a fish disease from a facility, watershed, or region is desirable but difficult to accomplish. To date, there are no reported examples of a fish disease agent being totally eradicated from a large geographical region. Practical eradication of IPNV, furunculosis, and bacterial kidney disease from individual fish hatcheries and farms in the United States

has been accomplished by removal of all fish and sterilization of facilities with chlorine or formaldehyde gas. These select facilities had closed water supplies with no indigenous fish populations to serve as disease reservoirs and precautions were taken to prevent reintroduction of disease. Only SPF eggs were used to repopulate the facilities and strict sanitation was established, which included disinfection of equipment and fish transport trucks that accessed the culture area. A commitment to eradicate a specific disease must be balanced with an equal commitment to keep the facility disease free. Eradication of a disease is much more difficult in warm water than cold-water aquaculture because most warm-water facilities do not have closed water supplies and use large ponds, which are more difficult to sterilize. They are more susceptible to animals (birds, snakes, muskrats, etc.) that may mechanically carry pathogens from one pond or farm to another. Additionally, there are no applicable SPF certification procedures that apply to warm-water fish pathogens. Disease prevention is primarily a farm management approach and should begin with construction of a culture facility. Disease prevention considerations should include site selection, development of water supply, and selection of fish to be stocked. Original fish stocks, as well as any subsequent fish brought onto the facility, should come from populations known to be free of obligate pathogens. When available, vaccination should be considered as a disease prevention tool. Sanitation practices are extremely important and disinfection of nets, buckets, boots, and other equipment should be a routine practice whenever used in different culture units. These precautions will also help prevent spread of disease within a facility once it occurs. Control is appropriate when obligate or nonobligate pathogens are involved or when effective chemotherapeutic and management practices are available to combat the disease. The objective of control is to reduce pathogen

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levels in an environment or host so that overt disease will not occur. This approach dictates that some fish will die while attempting to keep losses at a minimum. Chemotherapeutics are most successful when used in conjunction with elimination of environmental stressors and applying good health management practices. Biological and economic constraints usually dictate what approach will be employed when dealing with fish disease problems. Eradication may be the best approach when isolated cases of obligate pathogen infections are involved; however, destruction of entire animal populations remains controversial. Eradication vs. control should be determined on a case-by-case basis with strong consideration being given to economic consequences. Eradication is only feasible if a facility can be maintained free of a specific disease agent after sterilization.

Variable causes require variable solutions Epizootiology is the study of disease patterns in animals other than man and implies that the presence or absence of overt disease is not necessarily indicative of the presence or absence of a disease-causing agent. As previously discussed, many infectious fish diseases are caused by the complex interaction between host, pathogen, and environment. Although a host and pathogen must be present to cause infectious disease, environmental influences are often the trigger that elevates a subclinical infection to disease. Most pathogenic organisms can be identified, but environmental influences that precipitate disease are far more complex and difficult to detect. Each disease situation is different and must be evaluated and dealt with on an individual basis. When a fish disease problem arises, an environmental evaluation prior to and during time of disease outbreak must be part of the diagnostic process (Schnurrenberger 1983a). Facts pertinent to establishing an accurate diagnosis are as follows: had there been a recent

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oxygen depletion or any other water quality problem noted, any color change in the water, had heavy rains occurred recently, did mortality occur suddenly or increase gradually, were other fish recently introduced, and what are current water quality characteristics? Answers to these questions will influence type of management procedures or treatments to be used. At the time of disease outbreak, it is imperative that a complete disease examination (necropsy) be performed to determine if a single species of bacterium, parasite, or virus is involved or if multiple pathogens are present. It is emphasized that even after a pathogen is found, necropsy must be completed or other contributing pathogens may go undetected. Different disease organisms, requiring different treatments, can produce very similar clinical signs. Therefore, an incomplete necropsy may result in a premature and incorrect diagnosis and improper treatment. The key to treating multiple infections is to first identify and treat the primary disease-causing agent and then deal with lesser disease agents.

Don’t just cure, prevent A health maintenance program should be in place when an aquaculture facility begins production, and health management issues should be addressed on a daily basis, not just when disease outbreaks occur. When disease does occur, managers often expect an immediate diagnosis and treatment prescription from a fish disease specialist. This can put pressure on a diagnostician to recommend a treatment that will correct the immediate problem without addressing long-range problems. Often a chemical can be added to water and/or an antibiotic incorporated into feed, which will result in a positive response and cessation of mortality. However, the effect of these treatments may be temporary and disease may reoccur unless a commitment is made to seek and eliminate all predisposing disease factors. Even though a fish farmer is more likely to

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accept chronic disease losses because they are not dramatic and occur over an extended period of time, these losses can be minimized through proper health maintenance. If a subacute disease occurs and large numbers of fish die in a few days, the farmer will be more inclined to implement preventive steps. While preventive health maintenance is important in controlling fish diseases, it is not a “cure all.” Some fish diseases are less preventable through environmental management than are others and some have no known prevention or treatment. No health maintenance program is perfect, but one that emphasizes biosecurity and disease prevention is well worth the effort economically, productively, and for the pure satisfaction of knowing that the best possible aquaculture techniques are being used.

Law of limiting factors The law of limiting factors plays an important role in our ecology as well as in health management of fish. Schnurrenberger (1983b) compared this law to the concept that a “chain is no stronger than its weakest link.” Also, each major link consists of its own set of limiting factors and a weak spot within a subset of factors can cause failure in the overall chain. If one considers the fish production cycle a chain, with one end being spawning adults and the other harvestable-size fish, many potential weak links exist, any of which could be considered a limiting factor. Components present in a given body of water or water supply make it either suitable or unsuitable to support a healthy fish population. Volume of water, temperature, pH, and other parameters must all be compatible with the fish species being cultured. If adequate water is available but its temperature is not suitable, temperature then becomes a weak link and water becomes a limiting factor in the production cycle. Food quantity and quality is often a limiting factor in unmanaged fish ponds. With

fertilization, the base of the food chain in these ponds can be extended to support an increased standing crop, but if culture procedures become too intensive, other limiting factors may become involved. As a standing crop increases, supplemental feeding is required so that food will not again become a limiting factor. Supplemental feeding will increase the organic water load, and pond fertility will increase as a result of uneaten feed and the accumulation of metabolic wastes (feces, urine, ammonia, etc.). Decomposition of dead plants and uneaten feed remove oxygen from the water, and this oxygen loss may become a limiting factor. Accumulation of ammonia, nitrite, and CO2 may also become limiting factors. The law of limiting factors becomes acute as a fish culture system becomes more intensive. As fish density increases more feed is required; more water is necessary to remove, dilute, and neutralize metabolic wastes; more efficient supplemental aeration is required; and fish stressors become more critical. At what point these limiting factors become detrimental to effective production must be determined and appropriate corrective measures taken. A limiting factor does not always have to be biological. It can be financial or dependant on a manager’s ability to properly and effectively operate a facility. Each fish species has its own set of factors that are necessary for survival, growth, and reproduction. Factors for fish survival lie between minimum requirements and maximum tolerances. Consideration must also be given to the synergistic effect one limiting factor has on another and how altering one factor to improve health may illuminate a previously unrecognized factor.

Staying on top of the operation Many people go into aquaculture without fully understanding what is required to initiate and maintain a successful operation. Fish farming,

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especially of warm-water species, is possibly the most demanding of all agricultural enterprises during certain seasons of the year. During critical periods in the production cycle, a 24-hour schedule must be maintained to stay on top of the operation. It is as important in aquaculture for a fish farmer to routinely observe and evaluate his culture procedures and facility as it is for cattle or swine farmers to inspect herds daily. Although casual scrutiny of ponds, tanks, or raceways can be helpful in detecting possible health changes, specific aspects must also be considered. Operational procedures should be observed in such detail that any departure from good health management is immediately detected. The most obvious indicator of deteriorating fish health is a change (reduction) in feeding behavior, which can be affected by adverse environmental conditions and/or by infectious disease. If on a given day, fish are eating actively and the next day there is a noticeable reduction in feed consumption, the cause should be investigated. Water quality should be checked to determine DO, nitrite, or CO2 concentrations and a sample of fish should be examined for presence of infectious or parasitic disease. Some fish, channel catfish for example, normally reduce feed consumption in autumn and throughout the winter as water temperatures drop, but when this happens at other times of the year, it may indicate a health problem. Color changes in the water may indicate a potential water quality problem. When a pond turns from green (indicating a viable phytoplankton bloom) to dingy brown or suddenly clarifies, it could indicate algae are dying, or a pond “turn over” (good upper water mixes with unoxygenated deep water) is occurring and oxygen depletion is likely to ensue. If the cause can be detected and corrected while occurring, stress on fish can be minimized. DO concentrations in ponds should be measured on a daily and nightly basis during critical periods, either by an individual or automatic DO recording device. Computerized

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oxygen monitoring systems will register oxygen concentrations 24 hours a day and automatically turn on aeration when needed. Monitoring oxygen concentration is particularly critical in heavily stocked ponds that are receiving large amounts of feed. If over a period of days and nights a downward trend in oxygen concentration becomes evident, the DO may be ready to “crash” as a result of pond respiration and nightly aeration should be initiated. Also, when fish swim erratically or lethargically just beneath the surface, gasp (pipe) at the surface, or move into shallow water, one should look for causes for this abnormal behavior, which could be attributed to poor water quality, toxicants, infectious disease, or parasitic infestation. When fish surface, aeration should be initiated or freshwater added as soon as possible because a delay may cause fish to become so stressed they will not move to the aerated water. “Staying on top” can prevent catastrophes from occurring in a fish population, but it requires diligence and regularly scheduled visits to the entire culture facility. During critical times, if several days or nights are allowed to pass without checking water quality, high fish losses can occur in a very short period of time. Also, sublethal stressful conditions may lead to infectious disease.

Early diagnosis An early, accurate diagnosis when disease occurs is important in maintaining healthy fish; therefore, it is imperative that an aquaculturist know what is normal. It is important to be able to recognize clinical signs of disease and to have affected fish examined as soon as possible. It is also important to be able to distinguish signs of oxygen depletion and chemical toxicants from those of infectious disease. Few specific infectious fish diseases can be diagnosed solely upon clinical signs; therefore, whenever possible fish should be examined by a competent fish pathologist. In most cases,

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only a few fish die during early stages of a disease outbreak followed by a gradual increase in the daily mortality rate. When the first sick or dead fish appear, suitable specimens consisting of moribund fish showing clinical signs typifying the condition should be examined. Although an early and accurate diagnosis is important in nearly all infectious fish diseases it is absolutely crucial in bacterial infections when use of medicated feed is indicated or in parasitic infections requiring drug treatment. If infection progresses to a degree that fish are no longer feeding, oral treatment with antibiotics is no longer an option. In any disease situation, the earlier the diagnosis the sooner corrective measures or treatment can begin, thus reducing losses.

A dynamic team effort A dynamic fish health maintenance program must be flexible and coordinated, not a mixture of unrelated, unchanging procedures. While objectives and principles for fish health maintenance programs can follow a general outline, each facility needs to design and implement a health plan that best meets its needs. There are three principal times when an animal food resource producer is ready to discuss the establishment of a health maintenance program (Hudson 1983): (1) when experiencing a disease crisis, (2) when starting a new unit, and (3) when considering expansion with borrowed capital. The broad scope of basic knowledge and technology that applies to health maintenance of aquatic animals is usually beyond the expertise of any one individual. Therefore input from a variety of specialists is often indicated. In most geographical areas, State Cooperative Extension Services, or state, federal or university laboratories have qualified fishery specialists available to assist farm managers in developing a good health maintenance program that meets their specific needs.

Keeping current Health management decisions should be based on the most current and accurate information available. Shell (1993) quoted an old folk saying “It’s not what I don’t know that hurts me. It’s what I know that’s not true that hurts me.” In the past 40 years, major advances in aquaculture have been made in breeding, genetics, nutrition, disease identification, treatment, and processing. However, during this time, few new drugs or chemicals have been developed to aid in disease treatment and/or prevention. Because of constraints imposed by the U.S. Food and Drug Administration, there has actually been a reduction in number of drugs that can be used on food fish in the United States; a trend throughout many regions of the world. The reduction in approved chemotherapeutics is in part due to technological advances that have been made in identifying hazardous characteristics of drugs previously thought to be safe and their potential negative effects on efficacy for human medicine. It is, therefore, imperative that a producer stay current on which drugs and chemicals are approved for use on food fish. A fish farmer can stay current in aquaculture management techniques by being active in professional or trade organizations on a local, regional, state, national, and international level and by reading literature published by these organizations. Extension fishery specialists and agricultural agents can provide pamphlets, brochures, and newsletters with accurate, up-to-date information. If modern fishery managers are to maximize production facilities, they must keep pace with technological advances being made in the field.

Maintaining a clean environment Everyone recognizes the fact that an accumulation of trash, excrement, decaying animals, etc. in a terrestrial environment is associated

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with unhealthy conditions. Why should we expect it to be different in the aquatic environment? Clearly it is not! Generally, when visiting private and governmental aquaculture facilities around the world, it is often possible to visually identify facilities that have the most disease problems based on appearance. This is not to say that a “spit and polish” facility will never have disease problems, but they tend to have fewer and less severe disease outbreaks. Water is an excellent medium for transmitting disease-producing agents; therefore, sick or dead fish should be removed to reduce pathogen reservoirs. For example, in E. ictaluri-infected catfish ponds, areas where dead fish were allowed to accumulate had significantly higher pathogen concentrations in the water than where carcasses had not accumulated (Earlix 1995). Maintaining clean areas around ponds and controlling vegetation at water’s edge will not allow sick or dead fish to go unnoticed. If dead fish are allowed to remain in a culture unit, there is also the possibility that scavengers will carry infected carcasses to other ponds, thus spreading the disease. Accumulation of feces, uneaten feed, and other organic detritus is a major problem in ponds, raceways, and recirculating systems and contributes significantly to water quality degradation and can serve as a substrate for facultative pathogens. Removal of excreta and bottom detritus from the culture unit will help improve and maintain water quality. These problems can be avoided and/or corrected by improving feed quality, regular cleaning and periodic draining, drying, and when necessary disinfection of holding facilities. Removal of accumulated sediment is accelerated by drying the pond bottom, adding lime to the surface and tilling (Boyd and Pippopinyo 1994). Increasing soil pH to 7.5–8.0 with calcium hydroxide or calcium carbonate will enhance pond respiration and reduce pond sediment. Cleaning and disinfection of seines, nets, buckets, tanks, and other equipment will reduce transmission of many pathogens. Maintaining a clean aquaculture facility will pay

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dividends by providing a safer work place, reduced disease incidence, increased production, and generally healthier fish, all of which translates into more efficient production.

High-risk concept Aquaculture is a high-risk endeavor because of its dynamic relationship to environmental change. Since some aspects of fish culture present higher risk factors than others, management efforts should concentrate on areas that include production and financial losses, namely disease, growth rate, feed quality and conversion, water quality, and product marketability. Water quality is a major factor in rearing healthy fish and if not suitable, fish will not feed or grow well regardless of feed quality, and potential for disease becomes more prevalent. Consequently, major emphasis must be placed on water quality in any aquaculture health maintenance program. Infectious disease can also be classified as a major risk factor, but risks will differ from farm to farm depending on water type and quality, species and strain of fish being reared, general management practices, etc. Disease problems also change from year to year. The aquaculturist should be aware of high-risk factors, practice preventive health maintenance that addresses these areas, and at the same time not lose sight of lesser risk factors and their potential for causing problems.

Record keeping and cost analysis Well-organized records are basic to a successful aquaculture business. In addition to production records that include stocking and feeding rates, feed conversion, and harvest totals, each fish production facility should maintain a set of health records that include mortality patterns and dates, infectious disease incidences and treatments, and treatment results.

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Age

Natural resistance

Strain

Species

Acquired immunity

Host

Fish density Handling

Husbandry

Physiological status

Facilities

Fish health status

Type

Pathogen

Organic load

Environment

Species

Temperature Nitrite

pH

Strain

Nutrition Schedule Quantity

CO2

O2

Quality

Figure 1.7 The relationship of environmental conditions, biological factors, and management practices in aquaculture that influence health and infectious diseases of fish.

Size, water volume, and water quality characteristics of each production unit should also be included in health records. After several growing seasons, a manager may be able to ascertain from these records if a certain disease is likely to occur and what conditions predispose fish to a specific disease. This information can help prevent disease outbreaks by timely prophylactic treatments or other remedial actions. Detail and extent of record keeping may vary with size and intensity of an operation; however, records should be kept by every facility regardless of size. Accurate records are important when evaluating health maintenance procedures or a biosecurity program because without records, an economic feasibility and cost-effective evaluation is impossible. When considering the economic feasibility of a procedure cost, efficacy and benefits are considered and the most cost-effective (not necessarily the cheapest) procedure selected. Having to deal with greater pressure from imported fish products, domestic aquaculture must consider all aspects of the operation to maximize efficiency production and profit. When initiating fish health management practices, a long range, cost-effective analysis

should be made. Initial financial outlay may be high, but over a period of time, good health management will pay for itself in reduced mortality and optimal growth. A cost/benefit ratio should be calculated for each health management procedure and benefits must outweigh cost except under unusual circumstances.

Fish health status Health of fish depends on the interrelationship of six major components of the fish and the environment in which they live (Figure 1.7). The fish health matrix resembles a spider web in which if one satellite factor (host, husbandry, environment, etc.) is disrupted or diverge from the optimum, each of the other factors are also affected, thus impacting the health status of the fish. These components are such that one cannot be discussed without considering its relationship with or effect on each of the other components because of their interaction and their collective influence on fish health status. Overall physiological status of the fish host is determined by the husbandry practice, environmental quality, the fish’s nutritional well being and the pathogen, all of which influence

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the natural resistance and acquired immunity of the host. It is common knowledge that fish stressed by one of these factors are more susceptible to infection. Maintaining good health of the fish is the primary concern because it is necessary for survival and good growth. Disease susceptibility depends on the culture species because all fish species are not equally susceptible to a specific pathogen and strains within a fish species may vary in their susceptibility. Age of the fish is important because many high impact diseases (i.e., channel catfish virus, infectious pancreatic necrosis virus of trout, and the parasite Ichthyophthirius multifiliis) affect juvenile fish more severely than older and adult fish. Husbandry practices depend on the fish species being cultured. Stocking density of juvenile and grow out fish will vary because some species are more conducive to higher density than others; some species (salmonids) are adaptable to raceway culture but others (channel catfish) perform better in ponds. Generally, fish stocked at higher density are more prone to serious infectious disease than fish stocked at low or moderate density. Therefore higher stocking density requires more intense management. When and how fish are handled is an important consideration in disease susceptibility because the protective mucus layer that covers the skin serves as a barrier to ubiquitous invasive pathogens and if it is disturbed by improper handling, the fish becomes more susceptible to these pathogens. Also, moving fish when water temperatures are at the upper level of their tolerance is often stressful. The environment may be the most critical component of the fish health matrix because environmental quality influences the fish’s physiological well being, species cultured, feeding regimes, rate of growth, and ability to maintain natural and acquired resistance and immunity. Major environmental factors are temperature, organic load in the water, alkalinity and hardness, nitrite level, pH, oxygen and CO2 concentration as well as other water quality parameters. Tolerance of these

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various factors is dependent on the host and in many cases the husbandry practices. Nutrition is vital to health management, but feed quality and amount fed is related to husbandry practices. Good growth and maximum disease resistance is also dictated by the species of fish cultured. Some cultured species (i.e., salmonids and channel catfish) require complete manufactured feeds that contain complete vitamin packs, essential amino acids, and high protein content because they do not receive nutrients from their culture environment. In contrast, other species (i.e., tilapia or carp) feed extensively on water micro- and macroflora from which they receive essential nutrients. Quantity and quality of feed as well as feeding schedules are dictated by the fish species and age/size of the fish. The type of environment, whether it is a static water pond or flowing raceway, must be considered in the type of feeding program. The type of pathogen is dependent on all previously discussed factors in the fish health status matrix. Within a specific pathogen, there may be virulent or avirulent strains. While many pathogens are not host specific, others are either host specific or fish group (i.e., salmonids, ictalurids, etc.) specific. Whether or not a particular pathogen is present in an environment may depend on whether it is an obligate or nonobligate (facultative) pathogen. All fish viruses are obligate pathogens, while bacteria and parasites may be either obligate or nonobligate pathogens. Whether or not a pathogen causes an overt disease is often dictated by the host, husbandry practice, environmental quality, nutritional status, and the physiological status of the fish.

References AFS-FHS. 2007. FHS Blue Book: Suggested Procedures for the Detection and Identification of Certain Finfish and Shellfish Pathogens, 2007 edition. American Fisheries Society—Fish Health Section, Bethesda, Maryland.

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Anderson, D. E. 1991. BK episode (epitaph), Jackson National Fish Hatchery, a 1988 case study. In: 16th Annual Eastern Fish Health Workshop. Martinsburg, West Virginia, June 11–13 (Abstract). Austin, B., and D. A. Austin. 1987. Bacterial Fish Pathogens; Diseases in Farmed and Wild Fish. Chichester, Ellis Horwood Limited. Barton, B. A. 1997. Stress in finfish: past, present and future, A historical perspective. In: G. K. Iwama, A. D. Pickering, J. P. Sumpter, and C. B. Schreck (eds) Fish Stress and Health in A quaculture. Society for Experimental Biology Seminar Series 62. Cambridge, Cambridge University Press, pp. 1–33. Bebak-Williams, J., A. Noble, et al. 2007. Fish health management, Ch. 16. In: M. B. Timmons and J. M. Ebeling (eds) Recirculation Aquaculture. Cayuga Aqua Ventures, Ithaca, NY, and Northeastern Regional Aquaculture Center. Publication No. 01-007, pp. 619–664. Bilodeau-Bourgeois, A. L., F. G. Bosworth, and W. R. Wolters. 2007. Reduction in susceptibility of channel catfish, Ictalurus punctatus, to enteric septicemia of catfish through two generations of selection. Journal of the World Aquaculture Society 38:450–453. Boyd, C. E. 1990. Water Quality in Ponds for Aquaculture. Alabama Agricultural Experiment Station, Auburn University, Alabama. Boyd, C. E. 1995. Bottom Soils, Sediment, and Pond Aquaculture. New York, Chapman & Hall. Boyd, C. E., and S. Pippopinyo. 1994. Factors affecting respiration in dry pond bottom soils. Aquaculture 120:283–293. Butterworth, C. E., Jr., J. A. Plumb, and J. M. Grizzle. 1986. Abnormal folate metabolism in feed related anemia of cultured channel catfish. Proceedings of the Society for Experimental Biology and Medicine 181:49–58. Cipriano, R. C., C. E. Starliper, and J. D. Teska. 1991. Case study on the investigation of salmonids at the White Sulfur Springs National Fish Hatchery for Renibacterium salmoninarum. In: 16th Annual Eastern Fish Health Workshop, Martinsburg, West Virginia, June 11–13 (Abstract). Daelman, W. 1996. Animal health and the trade in aquatic animals within and to the European Union. Review in Scientific Technologies, Office International des Epizooties 15(2):711–722.

Dunham R. A., G. W. Warr, et al. 2002. Enhanced bacterial disease resistance of transgenic channel catfish Ictalurus punctatus possessing cecropin genes. Marine Biotechnology 4:338–344. Durve, V. S., and R. T. Lovell. 1982. Vitamin C and disease resistance in channel catfish (Ictalurus punctatus). Canadian Journal of Fisheries and Aquatic Science 39:948–951. Earlix, D. 1995. Host, pathogen, and environmental interactions of enteric septicemia of catfish. Doctoral Dissertation, Auburn University, Alabama. Hajek, B. F., and C. E. Boyd. 1994. Rating soil and water information for aquaculture. Aquacultural Engineering 13:115–128. Halver, J. 1972. Fish Nutrition. New York, Academic Press. Hastein, T, A. Hellstrom, et al. 2001. Surveillance of fish diseases in Nordic countries. Acta Veterinaria Scandinavica 42(Suppl. 1):S43–S50. Hill, B. J. 1996. National legislation in Great Britain for the control of fish diseases. Reviews in Scientific Technology, Office International des Epizooties, W. Daelman (ed.). 15(2):633– 645. Hooper, K. 1989. The isolation of VHSV from chinook salmon at Glennwood Springs, Orcas Island, Washington. Fish Health Section/American Fisheries Society News Letter 17(2):1. Hudson, R. S. 1983. A dynamic, coordinated entity. In: P. R. Schnurrenberger, and R. S. Sharman (eds) Principles of Health Maintenance. New York, Praeger Press, pp. 12–20. Kim, M. K., and R. T. Lovell. 1995. Effect of overwinter feeding regimen on body weight, body composition and resistance to Edwardsiella ictaluri in channel catfish, Ictalurus punctatus. Aquaculture 38:237–246. Klontz, G. W. 1973. Fish Health Management. Texas A & M Sea Grant Program, Texas A & M University, College Station, Texas. Li, Y., and R. T. Lovell. 1985. Elevated levels of dietary ascorbic acid increase immune response in channel catfish. Journal of Nutrition 115: 123–131. Lim, C., P. H. Klesius, and P. L. Duncan. 1996. Immune response and resistance of channel catfish to Edwardsiella ictaluri challenge when fed various dietary levels of zinc methionine and zinc sulfate. Journal of Aquatic Animal Health 8:302–307.

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Plumb, J. A., O. L. Green, et al. 1975. Channel catfish virus experiments with different strains of channel catfish. Transactions of the American Fisheries Society 104:140–143. Plumb, J. A., J. M. Grizzle, and J. deFigueiredo. 1976. Necrosis and bacterial infection in channel catfish (Ictalurus punctatus) following hypoxia. Journal of Wildlife Diseases 12:247–253. Roberts, R. J. 1989. Fish Pathology. London, Bailli`ere Tindall. Schnurrenberger, P. R. 1983a. Variable causes require variable solutions. In: P. R. Schnurrenberger and R. S. Sharman (eds) Principles of Health Maintenance. New York, Praeger Publishers, pp. 54–63. Schnurrenberger, P. R. 1983b. The law of limiting factors. In: P. R. Schnurrenberger and R. S. Sharman (eds) Principles of Health Maintenance. New York, Praeger Publishers, pp. 23–34. Schnurrenberger, P. R., and R. S. Sharman (eds). 1983. Principles of Health Maintenance. New York, Praeger Publishers. Shell, E. W. 1993. The Development of Aquaculture: An Ecosystems Perspective. Alabama Agricultural Experiment Station, Auburn University, Alabama. Simon, R. C., and W. B. Schill. 1984. Tables of sample size requirements for detection of fish infected by pathogens: three confidence levels for different infection prevalence and various population sizes. Journal of Fish Diseases 7:515–520. Snieszko, S. F. 1958. Natural resistance and susceptibility to infections. The Progressive FishCulturists 20:133–136. Snieszko, S. F. 1973. Recent advances of scientific knowledge and development pertaining to diseases of fishes. Advances in Veterinary Science and Comparative Medicine 17:291–314. Swan, A. I. 1983. The extent of contact. In: P. R. Schnurrenberger and R. S. Sharman (eds) Principles of Health Maintenance. New York, Praeger Publishers, pp. 35–42. Thorburn, M. A. 1996. Apparent prevalence of fish pathogens in asymptomatic salmonid populations and its effect on misclassifying population infection status. Journal of Aquatic Animal Health 8:271–277. Tucker, C. S., and M. Van der Pflog. 1993. Seasonal changes in water quality in commercial channel catfish ponds in Mississippi. Journal of the World Aquaculture Society 24:473–481.

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Walters, G. R., and J. A. Plumb. 1980. Environmental stress and bacterial infection in channel catfish, Ictalurus punctatus Rafinesque. Journal of Fish Diseases 17:177–185. Wakabayashi, H. 1996. Importation of aquaculture seedlings to Japan. Reviews in Scientific Techniques, Office International des Epizooties 15:409–422. Wedemeyer, G. A. 1996. Physiology of Fish in Intensive Culture Systems. New York, Chapman & Hall. Winton, J. R., W. N. Batts, et al. 1991. Characteristics of the first North American isolates of viral hemorrhagic septicemia virus. Proceedings Second International Symposium on Viruses of Lower Vertebrates, Corvallis, Oregon (Abstract).

Wise, D. J., and M. R. Johnson 1998. Effect of feeding frequency and Romet-medicated feed on survival, antibody response and weight gain of fingerling channel catfish (Ictalurus punctatus) after natural exposure to Edwardsiella ictaluri. Journal of the World Aquaculture Society 29:170– 176. Wolters, W. R., and M. R. Johnson. 1994. Enteric septicemia in blue catfish and three channel catfish strains. Journal of Aquatic Animal Health 6:329–334. Wolters, W. R., D. J. Wise, and P. H. Klesius. 1996. Survival and antibody response of channel catfish, blue catfish, and channel catfish female X blue catfish male hybrids after exposure to Edwardsiella ictaluri. Journal of Aquatic Animal Health 8:249–254.

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Chapter 2

Epizootiology of fish diseases

Understanding the dynamic aquatic environment and its role in fish health is imperative to management of infectious diseases. Fish respond in concert with, and quickly to, environmental changes that affect disease susceptibility and their overall general health. An effective fish health maintenance program continuously takes into account interaction between a fish population and its environment.

Communicable disease A communicable disease occurs as a result of multiplication, replication, or reproduction of the causative organism in a host and the likelihood that the organism can be transmitted (communicated) to other hosts either directly or indirectly.

Epizootic (epidemic) Epizootiology In a broad sense, epizootiology refers to the study of infectious diseases of animals, including spread of pathogens, mode of infection, prevention, and control. Relative to epizootiology, the following terms are important.

An epizootic is a disease outbreak among animals, other than humans (epidemic), in a specified and localized area. To be classified as an epizootic an abnormal number of animals must be infected.

Enzootic (endemic) Disease Disease is a deviation from normal or good health, not necessarily implying cause of deviation. Disease may be a condition resulting from infectious agents, nutritional deficiencies, toxicants, environmental factors, or genetics.

Enzootic indicates the continuing presence of a causative disease organism in a localized area, such as a pond, raceway, hatchery, river, or any other fish population but is not necessarily obvious as clinical disease.

Infection 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.

Infection is the presence of a pathogen in a host that may or may not be diseased. Fish may be 31

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A. hydrophila, F. columnare, and Saprolegnia spp., and protozoan parasites such as Trichodina sp. It should also be emphasized that multiple infections in which the presence of primary and secondary pathogens are frequent.

Seasonal trends of fish diseases Fish diseases generally occur seasonally and tend to fluctuate with temperature changes, presence of young susceptible fish, and environmental conditions that affect immunity and natural resistance (Meyer 1970; Plumb 1976; Warren 1991). In tropical climates, only minor differences are evident in numbers of fish disease outbreaks throughout the year; however, in temperate and colder climates, seasonal fluctuations can be noted, particularly in warmwater species (Figure 2.1). Disease incidence in cultured and wild fish populations in the southern United States is normally low from November through February, increases 20

Percentage of total diseases

infected, thus carriers of a pathogen, without becoming diseased. Many fish disease organisms are enzootic in the aquatic environment and will only cause disease when conditions are favorable. There are two basic types of organisms involved in communicable disease in terms of host relationship: (1) obligate pathogens and (2) nonobligate (facultative) pathogens. Obligate pathogens must have a definitive host or intermediate host to multiply and survive indefinitely in nature. Examples of fish obligate pathogens are all viruses, some bacterial pathogens, i.e., Mycobacterium marinum (Mycobacteriosis) and Renibacterium salmoninarum, and some parasitic protozoa (i.e., Ichthyophthirius multifiliis) and monogenetic trematodes. Facultative or nonobligate pathogens can live and multiply in a host or live freely deriving nutrients from organic matter in water, mud, or living host. Aeromonas hydrophila (motile Aeromonas septicemia) is an example of a facultative pathogen. The fungus Saprolegnia spp. is saprophytic and derives nutrients from living or dead organic material. Infections are also classified as primary or secondary infections in nature. Primary infections are those usually caused by obligate pathogens and can result in morbidity or death without the presence of additional factors. Primary pathogens are capable of causing disease in healthy fish even under ideal environmental conditions; however, any adverse environmental factor can synergize infection to a more serious level (Wedemeyer 1996, 1997). Generally, a secondary pathogen is caused by free-living, facultative, saprophytic, opportunistic organisms that infect a host when its defenses have been compromised by other pathogens or environmental or handling stressors. These infections often result from mechanical or physiological injury, disease that renders the host weakened, poor management practices, or environmental stressors such as low oxygen levels, temperature shock, and excessive or improper handling. There are many secondary pathogens that affect fish, including

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Figure 2.1 Monthly occurrence of 6,698 disease outbreaks in Alabama from 1991 through 1996. Data compiled from diagnostic records at the Southeastern Cooperative Fish Disease Laboratory, Auburn University, Alabama, and the Alabama Fish Farming Center, Greensboro, Alabama. (Data from Alabama Fish Farming Center courtesy of William Hemstreet.)

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Epizootiology of Fish Diseases 33

during March through June as waters begin to warm, decreases during the warm summer months, and incidence rises again in autumn. A similar disease pattern can be noted more northward but an increase in disease occurs later in the spring and earlier in the fall. Fish that live in waters with more constant temperatures, such as trout in flow through systems receiving ground water, temperature-controlled recirculating systems, or tropical fish, appear to have seasonal diseases when in reality disease is more related to age class or stage of development. Incidence of disease rises in spring because natural resistance and fish immunity are lower in late winter and early spring. Also, naturally occurring antimicrobial substances in the blood are low in the spring, making fish more susceptible to infection by facultative diseasecausing organisms (Snieszko 1958). Bly and Clem (1991) demonstrated that catfish immunity may be compromised during winter and fish will not respond positively until water temperatures rise to above 18◦ C in late spring or early summer. However, the immunologically favorable water temperature also coincides with periods of optimum growth for many pathogens. Changes in most water quality variables appear to be related to seasonal periodicity of phytoplankton abundance in channel catfish ponds (Tucker and van der Pfloeg 1993). Most water quality problems occur during summer. This is when water temperature and solar radiation are high, and fertility increases because ponds are being fed at a high rate. These factors tend to reduce oxygen availability. Stressful conditions precipitated by these events during summer months may actually be partially responsible for a delayed disease increase in autumn.

Factors in disease development When an infectious outbreak occurs, factors that dictate disease severity are source of infec-

tion and mode of transmission, portal of entry, virulence of the pathogenic organism, and resistance of the host. If one of these factors can be altered to favor the fish host through management, disease may be eliminated or its impact greatly reduced.

Source of infection and mode of transmission An infectious disease requires a pathogen source, which may include dead or moribund fish, carrier fish that show no clinical signs of disease, contaminated eggs from infected brood stock, infected water supplies, on contaminated equipment (seines, net, buckets, etc. or on humans (Bebak-Williams et al. 2007). Contaminated feed can be a source of bacterial and parasitic agents, especially if it contains uncooked fish flesh or viscera. The mode by which fish disease is transmitted is closely related to the source of infection. Water, because it contains fish metabolites and waste products, often provides an ideal high nutrient environment in which potential fish pathogens can grow and proliferate, thus serving as a pathogen source as well as a primary mode of transmission. Transmission of many pathogens can be prevented by disinfection of equipment, filtration of water, adjustment of stocking densities in infected waters, removal of dead and moribund fish (Earlix 1995), utilization of seed fish from specific pathogen-free (SPF) brood stocks, and use of proper feed by establishment of a biosecurity program (BebakWilliams et al. 2007). Eggs produced by disease-carrying brood stock are often a source for vertical transmission of viruses and some bacteria from generation to another generation. This mode of transmission can be disrupted in some instances by using SPF fish sources of eggs and fingerlings. Also intermediate hosts such as birds, snails, and crustaceans are important in the life cycles of some fish parasites and can serve as

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34 Health Maintenance

vehicles for disease organisms. Man often aids in the transmission of disease organisms by mechanically transporting infective agents in water, fish transport equipment, and on nets, boots, and other utensils and their clothing. Management practices that can help reduce these sources include the use of prophylactic treatments, control of intermediate hosts, disinfection of equipment, and use of a fish-free water source or water filtration treatment with ozone or ultraviolet light.

Portal of entry Each disease organism has an optimal point of entry to the host and if entry points are identified, management may help prevent infection. Common entry points are the intestine, gills, and skin. In fish, the mucous layer covering the epithelium provides protection against some pathogens. If this mucous layer is compromised, underlying epithelium is exposed to bacteria or parasites present in the water and as a result the skin, gills, or intestines become portals of entry for the pathogen. Also, parasites may penetrate the mucous layer and epithelium making fish vulnerable to invasion by viruses and bacteria. Proper handling and use of prophylactic treatments will often reduce or prevent epithelial infections.

Virulence of the pathogenic organisms Virulence is a measure of an organism’s ability to cause disease and may range from low to high. Because of varying antigenic traits, facultative bacteria can differ greatly in virulence from location to location. Aeromonas hydrophila, a facultative bacterium in water that causes motile Aeromonas septicemia, can be highly virulent to fish in one area, while an isolate from another body of water can be avirulent (not pathogenic). When A. hydrophila is isolated from diseased fish, it is

usually more virulent than when isolated from water (de Figueiredo and Plumb 1977). It is more likely that a severe disease outbreak will occur when highly virulent pathogen strains are present. Although it is difficult to manipulate virulence of pathogenic organisms in nature, management can help prevent severe disease outbreaks by ensuring that environmental conditions are more favorable to the host than to the pathogen. Furthermore, some pathogens can evolve or mutate rapidly. Management schemes that select for more virulent pathogens include conditions where the opportunity for transmission is optimized if the pathogen debilitates its host. This can occur in systems where cannibalism of diseased or dead fish is common, which can be minimized using optimum feeding regimes and vigilant removal of dead and diseased fish.

Natural resistance of the host Natural resistance occurs when a host has an inherent ability to subdue a pathogen to the point that clinical disease will not occur. A fish’s natural resistance to disease is extremely important in deterring infection (Chevassus and Dorson 1990). Intrinsic factors that influence a fish’s natural resistance to disease are natural killer cells, nonspecific phagocyte activity of neutrophils and macrophages, tissue integrity, and nonspecific serum components (interferon, complement, etc.). Many of these factors are related to genetic characteristics. However, extrinsic influences such as nutritional well-being and environmental conditions can adversely affect these traits. Also, natural resistance is related to species, strain, and age of fish. Natural resistance of adult fish is often reduced during, or prior to, the spawning season when some fish stop feeding and energy is diverted into reproductive activities. In temperate zones, older fish tend to have reduced resistance in spring due to over wintering and presence of young fish that have not yet acquired natural resistance. Taking advantage

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Epizootiology of Fish Diseases 35

of natural resistance is an integral part of fish health management and can be enhanced by culturing more disease resistant strains.

Host/pathogen relationship That there is a basic host–pathogen relationship is a fundamental concept in epizootiology of fish diseases. A pathogen may be present in the environment or host but as long as the host remains in good physical condition and its natural resistance remains high, clinical disease is less likely to occur. Disease organisms tend to cause infections in fish when there is an environmentally induced host/pathogen imbalance. This imbalance can be manifested by long-term nutritional problems and/or deterioration of the aquatic environment to the degree that a fish’s natural resistance is compromised and “infection” progresses into

“disease.” Through good management, a proper host/pathogen balance is better maintained and clinical disease outbreaks kept to a minimum resulting in better growth and higher survival. However, there are some pathogens that cause fish disease even though there is a balance in the host/pathogen relationship.

Degree of infection Fish diseases are categorized according to disease severity, mortality and morbidity rate, and clinical signs as acute, sub acute, chronic, or latent (Figure 2.2). Mortality pattern is related to severity of the causative agent or degree of infection and is helpful in determining whether cause of death is due to toxicants, adverse water quality, infectious agents, or nutritional deficiencies.

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Figure 2.2 Theoretical cumulative mortality patterns in fish populations. Acute patterns are usually associated with environmental problems and rarely infectious agents. Subacute mortalities may be associated with environmental problems or infectious agents. Chronic mortalities are frequently associated with infectious agents.

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36 Health Maintenance

Acute Acute mortality is marked by a high death rate in a short period of time (hours to 1–2 days). In most instances, truly acute mortalities are usually associated with water quality, oxygen depletions, or toxicants. In rare instances, infectious agents may cause acute mortality, but overt clinical signs (ulcerative lesions, severe hemorrhage, etc.) may be minimal or totally lacking. Examples of pathogens of warmwater fish that cause acute disease are channel catfish virus (CCV) and Flavobacterium columnare (columnaris); examples of acute infections in coldwater fish are bacterial gill disease and infectious pancreatic necrosis virus (IPNV). These are usually in densely stocked juvenile populations.

Subacute A subacute infection is less severe than an acute infection and mortality will accelerate over a 3–4 day period, lasting for several weeks with clinical signs present. Some highly virulent organisms that are capable of causing acute or subacute mortality include CCV, infectious hematopoietic necrosis virus (IHNV), and IPNV. On rare occasions F. columnare, bacterial gill disease of salmonids (F. branchiophilum), and some parasites (I. multifiliis) cause subacute mortality. Environmental conditions may also be associated with subacute mortality.

Chronic Many infectious fish diseases exhibit chronic characteristics that include maximum clinical signs that persist for weeks, during which time increased mortality rate is gradual with an undetectable peak, but cumulative mortality can be high. Examples of pathogens that establish chronic fish infections are motile Aeromonas septicemia (Aeromonas spp.), columnaris, furunculosis (A. salmonicida), bacterial kidney

disease (R. salmoninarum), and most protozoan parasites.

Covert In covert infections, a disease organism is present, but its host exhibits no overt clinical signs of disease. Covert infections are generally the mechanism by which obligate pathogens maintain themselves in endemic populations. Covertly infected fish are often called unapparent carriers where pathogens can be transmitted from these carriers to na¨ıve fish where they cause disease. Also, when environmental conditions are optimal for disease or when the carriers become stressed, overt disease may occur in the carrier population, a phenomenon called “recrudescent” infection. Diseases from some recrudescent infections display different clinical signs than the initial infection, such as tumors in cyprinid herpesvirus 1 (carp pox), salmonid herpesvirus 2, and walleye dermal sarcoma virus infections. There are two types of covert infections: persistent infections and latent infections. Persistence is a low level infection of a pathogen where the infectious agent can be isolated from the host and propagated ex vivo (in vitro). Some pathogens establish persistent infections in which the carrier fish continuously or frequently shed the infectious agent. Latency is where the pathogen cannot be detected (no infectious agent is present). This occurs in viral infections where the pathogen is present in a noninfectious form, usually as the nucleic acid genome within a host cell. The important difference between the two types of covert infections is the way in which the carrier state can be detected. Persistent infections are generally detectable using culture methods whereas latent infections require nucleic acid-based detection such as polymerase chain reaction (PCR) or RT PCR (discussed in Chapter 3). It cannot be overemphasized that a minor disease can progress into a severe condition if good management procedures are not

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Epizootiology of Fish Diseases 37

practiced prior to or during disease outbreaks. For example, if diseased fish in a pond are moved to a confined holding facility for treatment, the act of moving the fish will probably be more detrimental and stressful than any benefit derived from treatment.

References Bebak-Williams, J., A. Noble, et al. 2007. Fish health management, Chapter 16. In: M. B. Timmons and J. M. Ebeling (eds) Recirculation Aquaculture. Ithaca, NY, Cayuga Aqua Ventures and Northeastern Regional Aquaculture Center. Publication No. 01-007, pp. 69– 664. Bly, J. E., and L. W. Clem. 1991. Temperaturemediated processes in teleost immunity: in vitro immunosuppression induced by in vivo low temperature in channel catfish. Veterinary Immunology and Immunopathology 28:365–377. Chevassus, B., and M. Dorson. 1990. Genetics of resistance to disease in fishes. Aquaculture 85:83–107. de Figueiredo, J., and J. A. Plumb. 1977. Virulence of different isolates of Aeromonas hydrophila in channel catfish. Aquaculture 11:349–354. Earlix, D. 1995. Host, pathogen, and environmental interactions of enteric septicemia of catfish.

Auburn, AL, Auburn University. Doctoral dissertation. Meyer, F. P. 1970. Seasonal fluctuations in the incidence of disease in fish farms. In: S. F. Snieszko (ed.) A Symposium on Diseases of Fishes and Shellfishes. Bethesda, Maryland, American Fisheries Society Publication No. 5, pp. 21–29. Plumb, J. A. 1976. An 11-year summary of fish disease cases at the Southeastern Cooperative Fish Disease Laboratory. Proceedings Annual Conference of the Southeastern Association of Game and Fish Commissioners 29:254–260. Snieszko, S. F. 1958. Natural resistance and susceptibility to infections. The Progressive FishCulturist 20:133–136. Tucker, C. S., and M. van der Pfloeg. 1993. Seasonal changes in water quality in commercial channel catfish ponds in Mississippi. Journal of the World Aquaculture Society 24:473–481. Warren, J. W. 1991. Diseases of Hatchery Fish, Sixth Edition. U.S. Fish and Wildlife Service. Wedemeyer, G. A. 1996. Physiology of Fish in Intensive Culture Systems. New York, Chapman & Hall. Wedemeyer, G. A. 1997. Effects of rearing conditions on the health and physiological quality of fish in intensive culture. In: G. K. Iwama, A. D. Pickering, J. P. Sumpter and C. B. Shreck (eds) Fish Stress and Health in Aquaculture. Society for Experimental Biology Seminar, Series 62. Cambridge, Cambridge Press, pp. 35–71.

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Chapter 3

Pathology and disease diagnosis

When fish are dying or become diseased in an aquaculture system, an accurate and timely identification of the cause of the deaths is critical for managing the disease. Furthermore, a complete evaluation of the production system may be warranted to determine underlying causes (predisposing factors) so losses can be avoided in the future. Disease diagnosis is an investigative science that incorporates findings from multiple disciplines (epidemiology, pathology, toxicology, parasitology, bacteriology, and virology) to find the most likely cause of losses. This process must also include an assessment of environmental quality and related management practices. Chapter 2 broadly reviewed epidemiology of fish diseases. This chapter presents concepts of pathology, pathogen identification, and disease diagnosis.

Pathology Pathology, the study of disease and the bodily changes produced by them, includes functional and morphological alterations and reactions that develop in a living organism as a 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.

result of injurious pathogens, nutritional deficiencies, inherited characteristics, or those associated with inanimate causes. Pathology and epidemiology are important aspects of disease diagnosis. They indicate the importance of the problem at the individual and population level, respectively. Often several possible factors can be identified that may cause disease. The information provided by pathology indicates the disease process that is occurring and gives a better picture of the cause of the disease problem. Pathology terms used in description and interpretation are essentially the same for fish as for higher vertebrates (Cotran et al. 1989; Ferguson 1989; Roberts 1989). Fish pathology is too vast a subject to thoroughly cover in this section; therefore, the objective is to introduce basic terms and present examples of the most common pathological changes that occur in fish.

Pathology terms A unique vocabulary is used for the precise description of the physical condition of an organism or a tissue (Cotran et al. 1989; Ferguson 1989; Roberts 1989). Below are the terms that are often used when describing a disease or considering the disease process. In addition, the name of an affected tissue or 39

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40 Health Maintenance

organ may be conjugated with a suffix –osis that indicates a condition, -opathy that indicates a pathological change is occurring in this tissue, and –itis that indicates inflammation is occurring in this tissue. In these descriptions, specific tissues are indicated in the first part of the word: hepa- or hepatic indicates liver, branch- or branchial indicates gill, card- or cardiac indicates heart, gastr- or gastric indicates stomach, encephal- indicates brain, renal indicates kidney, occul- or ocular indicates eye, myo- indicates muscle, and derm- indicates skin.

Etiology

Lesion Any gross, microscopic, or biochemical tissue abnormality. The lesion can be either functional or morphological.

Clinical signs Characteristics or conditions associated with disease, which can be seen during a macroscopic external examination of the affected animal. Clinical signs include behavioral and gross morphological changes but do not include microscopic, biochemical, or internal changes.

The cause of or origin of a specific disease.

Histology Etiological agent The pathogen, nutritional deficiency, etc., which causes a particular disease; for example, channel catfish virus (CCV) is the etiological agent of channel catfish virus disease (CCVD).

Pathogenesis The sequence of events by which a disease develops, including method of infection, establishment of the agent in the host, and injury inflicted on the host.

The study of normal tissue on a microscopic level. Representative species from several important fish groups have been studied, and their normal histology has been described. These include channel catfish (Grizzle and Rogers 1976), striped bass (Groman 1982), salmonids (Yasutake and Wales 1983), Atlantic cod (Morrison 1987–1993), and walking catfish (Chinabut et al. 1991), fathead minnow (Yonkos et al. 2000), normal trout (Horsch 2008), and others including an atlas of fish histology (Genten et al. 2009).

Pathogenicity This refers to a pathogen’s characteristic ability to gain access to susceptible tissue, become established, proliferate, avoid host defense mechanisms, and cause injury and disease to the host.

Virulence Degree of pathogenicity of a disease agent: avirulent (not virulent), low, medium, or highly virulent. These terms are usually related to a specific group, species, or strain of pathogenic organism.

Histopathology The microscopic study of injuries to tissue and cells.

Cause of disease Disease can be caused by an etiological (specific cause) or a nonetiological (contributing cause) agent. Etiologies can be classified as either inanimate or animate. Inanimate etiologies are factors without life of their own and

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Pathology and Disease Diagnosis

can originate within a host (endogenous) or outside of a host (exogenous). Endogenous, inanimate factors are those associated with genetic and/or metabolic disorders of the host. Exogenous, inanimate agents include trauma, temperature shock, electrical shock, chemical toxicity, and dietary deficiencies. These etiologies may serve as sublethal stressors that predispose fish to infectious disease. Animate etiologies are living communicable infectious agents, which include viruses, bacteria, fungi, protozoa, helminths, and copepods. Nonetiological causes of disease are characterized as extrinsic (from outside the body) or intrinsic (within the body). Extrinsic factors are usually associated with environmental conditions or dietary problems while intrinsic factors include age, sex, heredity, and fish species. Both species and strain of fish are important because all are not equally susceptible to a specific disease organism. Feed quality, water quality factors, and water temperature extremes can be classified as either etiological or nonetiological extrinsic factors and can contribute to the onset of infectious disease.

Pathological change Pathological changes in individual fish can aid in recognition and identification of disease. Using definitions from Roberts (1989) and Ferguson (1989), the following are brief descriptions of some pathological changes that can occur in diseased fish. While the effects of many of these changes are detectable with the unaided eye, confirmation of the type of pathological change is provided by microscopy.

Circulatory disturbances Circulatory disturbances are abnormalities (lesions) that reflect injury to the vascular system. They may be anemia, hemorrhage, edema, hyperemia, congestion, emboli, or telangiectasis.

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Anemia Anemia is a reduction of red blood cells that results from cell lyses during infection, presence of toxicants, or lack of erythrocyte production. Anemia may result in red blood cell volumes falling from a normal range of 25–40%, depending upon fish species, to less than 15%. Hemorrhage Hemorrhage is the escape of blood from vessels and can occur in the skin, mucous membranes, within serous cavities, or between cells of any tissue or organ. Hemorrhages are generally classified according to size or degree of area affected. Petechiae are hemorrhages that measure less than 2 mm in diameter. They are common in bacterial and viral infections and typically occur in the skin or on the surface of visceral organs. Ecchymosis is larger, more diffuse hemorrhage than petechiae and measure 3 mm or larger in diameter. Paintbrush hemorrhage refers to large affected areas that appear as though they were splashed with red paint. Hematomas (seldom seen in fish) are localized hemorrhages that result in a bloodfilled swelling. Hemorrhages can be caused by ruptured blood vessels as a result of trauma or increased vessel porosity resulting from the presence of infectious bacteria, viruses, or toxicants. In fish, overall effects of hemorrhaging are mild if the process is gradual because hematopoiesis is generally able to compensate for a mild loss of blood cells. However, if hemorrhaging is acute, as often is the case in infectious disease, anemia can occur and will be characterized by pale gills and internal organs and a low hematocrit (percent packed red blood cells). Edema Edema is an abnormal accumulation of fluids (not whole blood) in body cavities, interstitial spaces of tissues, and organs, which results in swelling. Four categories of edema occur in fish: (1) Ascites is fluid accumulation in the body cavity caused by dysfunction of the heart, liver, or kidney and gives affected fish a

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42 Health Maintenance

“pot-bellied” appearance. This fluid may range from colorless to pale yellow, cloudy, or bloody depending on the etiology. Turbidity is caused by high numbers of bacteria. (2) Hydropsy results from fluid in muscle, interstitial, and connective tissues and gives fish a swollen appearance. When pressure is applied to the skin, an indentation will remain (pitting on pressure). (3) Lepidorthosis is the accumulation of fluid in scale pockets, causing scales to protrude from the body surface making the surface rough. (4) Exophthalmia is the accumulation of fluid, either behind or in the eye, causing a protrusion of the eyeball, which gives affected fish a “popeyed” appearance. Edema indicates an imbalance of hydrostatic pressure or improper osmolarity of the blood, increased capillary permeability, vascular obstruction, or organ dysfunction. These conditions are normally associated with chemical toxicants; viral, bacterial, or parasitic agents; or mechanical injury. Edema caused by either injury or disease may predispose fish to further infection, as fluids provide a medium for bacterial growth. The effects of edema can be harmful and long lasting. Hyperemia and congestion Hyperemia and congestion involve an excessive volume of blood within the vessels; however, mechanisms of development differ. Hyperemia is an active engorgement of blood in the vascular bed and is often associated with inflammation. By contrast, congestion is a passive accumulation of blood in vessels caused by an impairment of blood flow. Hyperemia and congestion cause affected tissue to appear red with possible swelling, and blood vessels in the fins and mesenteries become more prominent. Telangiectasis Telangiectasis is the bulging of a blood vessel in fish gills that is equivalent to an aneurysm in higher vertebrates. However, telangiectasis is passive and reversible, whereas an aneurysm is the permanent swelling of an artery. Telangiectasis may be caused by mechanical in-

jury, toxicants, viruses, bacteria, bacterial toxins, parasites, and, in some cases, nutritional deficiencies. Embolism Embolism is an obstruction, often gas bubbles, in the vascular system that causes an interruption of blood flow. Gas bubbles in blood vessels caused by a supersaturation of gas (usually oxygen or nitrogen) in the water are common in fish (Bouck 1980). When dissolved gas in water exceeds 100% saturation (supersaturation) for an extended period of time, gas bubbles may form in the blood producing “gas bubble disease.” Fish affected by this disease may have microscopic or macroscopic gas bubbles in the operculum, gills, eyes, fins, and around the head. Gas bubble disease in cultured fish occurs when deep wells are used as a water supply, when air gets trapped in a pressurized water supply, or when cool water is heated just before it flows to a culture unit. Cool water has a higher gas saturation point, and when water is warmed, it will become supersaturated. Wild fish populations living just below high dams may be affected by gas bubble disease as a result of water tumbling over spillways and entrapment of gas in the plunge pool.

Cellular degeneration Degeneration is a broad term referring to a process in which cells or tissues deteriorate, usually with a corresponding degree of functional inhibition. In the process of cellular degeneration, cells go through biochemical alterations and functional abnormalities that finally result in morphological change. The degree and progress of degeneration varies with type and number of cells affected, nature of the injurious agent, intensity, and duration of injury. Degeneration of cells and tissues can be reversible, changing from a regressive to a progressive process and affected cells restored to normal if the injurious cause is removed before cell death occurs.

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Pathology and Disease Diagnosis

Degeneration can result from the following: (1) deficiency of a critical material (oxygen or a vital nutrient); (2) lack of an energy source that affects metabolism; (3) mechanical, thermal, or electrical injury; or (4) accumulation of an abnormal substance in cells caused by viral, bacterial, or parasitic pathogens and their toxins, toxic chemicals, nutritional imbalance, mild irritants, and others. Types of cellular degeneration found in fish include cloudy swelling, fatty change, and mucoid degeneration. Cloudy swelling Cloudy swelling (or cellular swelling) often results from bacterial toxemia and is the earliest microscopic indication of cellular degeneration. Cells affected with cloudy swelling are enlarged, and cytoplasm has a homogeneous, hazy appearance identified by microscopic examination of the tissue. Fatty degeneration Fatty degeneration (or fatty change) is the result of an accumulation of lipids, most commonly in the liver. This change is usually caused by infectious disease, nutritional imbalance, hypoxia, anemia, and some toxicants. The liver is characterized by a pale yellowish to gray appearance. Although macroscopic evidence of fatty degeneration may eventually be noticed, early indications are microscopic. Mucoid degeneration Mucoid degeneration is the deposition of mucus or mucoid substances in the connective tissue of organs. Necrosis Necrosis is the death of cells following cell degeneration in a living animal and is the final stage in an irreversible process. Characteristics of necrotic tissue can include the following: (1) paler than normal color, (2) loss of tensile strength (tissue becomes friable and easily torn), (3) may have a cheesy or pasty consistency, and (4) an unpleasant odor. Necro-

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sis should not be confused with postmortem change, which is cellular deterioration after death of an animal. Necrosis can be caused by trauma, biological agents (viruses, bacteria, fungi, and parasites), chemicals, or an interrupted blood supply to a specific area. Three types of necrosis are as follows: (1) liquefactive, (2) coagulation, and (3) caseous. When a cell dies, enzymes within the cell cause liquefactive necrosis—appears as a semisolid or fluid mass—which is the most common type of fish necrosis. When this occurs in the epidermis or exposed muscle, the necrotized tissue is sloughed off, leaving an open ulcerative lesion that can be invaded by facultative pathogens present in the water. Coagulation necrosis refers to an area in which the gross and microscopic nature of affected tissue is still recognizable and is associated with injury caused by several types of toxicants. Caseous necrosis, uncommon in fish, results when a pathogenic organism produces cheesy, whitish material in a lesion.

Disturbances of development and growth Several growth changes (excessive, deficient, or abnormal growth in cells, tissues, or organs) can occur in fish in response to an infection, toxicant, or other irritant. Generally, growth disturbances involve either an abnormal number of cells, cells of abnormal size in an organ or tissue, or a combination of the two. Atrophy Atrophy refers to a decrease in size of a mature body part or organ due to decreased size or number of cells. Atrophy is a slow process and results from starvation or malnutrition (most common cause), lack of adequate blood supply, or chronic infection. Hypertrophy Hypertrophy is an increase in size or volume of a body part or organ due to an increase in

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size of individual cells. Hypertrophy usually not only results from an increased demand for function but can also be initiated by an infectious agent such as lymphocystis virus. Hyperplasia Hyperplasia relates to an increase in size of a body part or organ due to an increase in the number of cells. One form of hyperplasia is characterized by an increased thickness of gill lamellae epithelium due to infection or exposure to a continuous mild irritant. Chemical water pollutants, as well as some fish viruses or bacteria, cause formation of hyperplastic lesions, particularly of the integument. Enlargement of a thyroid gland to form a goiter in fish is an example of hyperplasia. Neoplasia Neoplasia refers to new growth, often a process that can result in tumor formation. The cells are induced to replicate beyond normal responses to a stimulus. Benign tumors are noninvasive, generally contained within the originating organ. Malignant tumors invade other tissue where cells may change in their morphology, break loose (metastasize), and start tumors in other locations in the body. Tumor pathology nomenclature can be very involved, but it generally indicates the cell type with a suffix indicating the severity. The suffix –oma indicates a neoplasia. Metaplasia Metaplasia indicates a change in the function of a cell in a tissue to a form that is not normal for that tissue.

Inflammation Inflammation, which can be acute or chronic, is an aggressive vascular and cellular defensive response of living animal tissue to a sublethal injury that helps to minimize the effect of an irritant or pathogen on tissue. When injury or infection occurs, fluids from the vascular system and lymphocytes, neutrophils, macrophages,

and other blood components migrate to the injured area. Accumulated fluids and cells attempt to dilute, localize, destroy, and remove any irritant or infectious agent and to stimulate replacement and repair of injured tissue. Inflammation is caused by viruses, bacteria, parasites, trauma, heat, irradiation, or toxicants. The “cardinal signs” of inflammation in higher vertebrates are as follows: (1) redness (hyperemia), (2) swelling, (3) heat, (4) pain, and (5) loss of function. How exactly these apply to fish is not clearly understood, but in most instances, the process is similar to that of other animals. Acute inflammation is classified according to type of exudate involved: serous, fibrinous, hemorrhagic, catarrhal, purulent, or granulomatous. Exudates are named according to histological appearance and all types are seen in fish to varying degrees. Serous Serous inflammation is exudation of clear fluid from the vascular system in response to a mild irritant or infection. A blister is a good example of serous inflammation although they are seldom seen in fish. Fibrinous Fibrinous inflammation occurs when large amounts of fibrin escapes from blood vessels and forms a clear clot when exposed to air. This type of inflammation usually occurs in the peritoneal cavity of fish and is associated with hyperemia. Purulent Purulent inflammation typically contains a combination of necrotic cells, neutrophils, and proteolytic enzymes in the exudate. Generally, fish do not produce pus, although in some bacterial diseases, focal infections do produce purulent exudate. Purulent inflammation occurs in furunculosis of trout and has occasionally been found in motile Aeromonas septicemia infections.

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Catarrhal Catarrhal inflammation is the excessive production of mucus on the epithelium of the skin, gills, and digestive tract. The exudate can be clear, cloudy, or pink with a consistency of fluid to mucoid. The fecal cast of trout infected with infectious hematopoietic necrosis virus contains catarrhal exudate. Catarrhal exudate also forms on the gills and skin of fish infected with Ichthyophthirius multifiliis.

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ulomatous inflammation will have small white spots, and the surface may have a rough sandpaper texture. In some cases, several granulomas may be connected by fibrous connective tissue to form a large encapsulated hard nodule. If the host survives the causative infection, after inflammation has disappeared, a central necrotic area remains that may retain the viable causative agent.

Disease recognition and diagnosis Hemorrhagic Hemorrhagic inflammation is usually characterized by the presence of large numbers of erythrocytes and other blood components on organ surfaces or in exudate. Hemorrhagic inflammation, caused by viruses, bacteria, parasites, or toxicants, will generally be diffused on serous or mucous membranes. Erythema, a reddish skin condition, is usually associated with hemorrhagic inflammation. Granulomatous Granulomatous inflammation is usually a chronic condition associated with several microbial fish infections and is characterized by a predominance of macrophages or related cell types. Edwardsiella ictaluri causes a condition similar to granulomatous inflammation in the skull of channel catfish, where the bone tissue becomes soft and spongy, but it is not typical of this pathology. Other pathogens, Mycobacterium spp., Photobacterium damsella subsp. piscicida (bacteria), and Ichthyophonus hoferi (fungus), for example, cause granulomatous inflammation often progressing into macroscopic granulomas. Acute granulomatous inflammation occurs initially in internal organs and muscle tissue. Macrophages and other inflammatory cells form a layer resembling epithelium around the irritant, and as the lesion ages increased, numbers of fibroblasts and lymphocytes appear. The resulting granulomas are white to yellow in color and have a cheesy to hard consistency. Tissue affected with gran-

An accurate fish disease diagnosis involves recognizing the presence of an infectious agent or other etiology, its role in the disease process, environmental factors that may have influenced or even precipitated the process, and how the problem can be corrected. Aquaculturists may be capable of recognizing some infectious diseases, but usually it is necessary to consult a fish health professional for positive identification. Diagnostic procedures discussed here are basic techniques employed in fish disease diagnosis. It is beyond the scope of this work to delve into molecular diagnostic procedures in great detail.

Disease recognition Although infectious agents play a major role in fish health, other important factors must be identified and considered when a disease diagnosis is made (Meyer and Barclay 1990; Noga 1996). Actual identification of a parasitic, bacterial, or viral agent causing disease may not be difficult; however, knowing how to deal with the problem and preventing it from reoccurring is often more difficult. History, mortality pattern, and water quality characteristics provide vital information for an accurate diagnosis and treatment regimen (Lasee 1995). Once an etiological agent is identified and a treatment strategy is established, steps must be taken to prevent a reoccurrence of disease.

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History History of a fish population exhibiting morbidity or mortality should include origin, size, age, number, and species of fish involved, previous disease history, type of holding facility, how long fish with clinical signs of illness have been noted, behavior and feeding patterns, and type of treatment administered. It is also essential to collect as much environmental and water quality data as possible, which should include water temperature; dissolved oxygen, carbon dioxide, ammonia, and nitrite concentrations; any recent change in water color, clarity, or flow; stocking density and weather conditions just prior to mortality; and agricultural activities in the area and size of culture unit. All of these data can be vital in diagnosing and treating disease, and more importantly, preventing reoccurrence.

Mortality pattern A mortality pattern may indicate cause of death due to an infectious agent, poor water quality, or presence of a toxicant. Generally, infectious disease outbreaks are marked by a gradual acceleration in morbidity and mortality unless an extremely virulent pathogen is involved (Figure 2.2). Infectious disease agents seldom, if ever, result in “overnight” mass mortality but most often produce subacute or chronic mortality patterns, resulting in cumulative losses of 30–40% or less. Mortality seldom reaches 100%; however, when highly virulent agents (some viruses and bacteria) infect highly susceptible fish, 90–l00% cumulative mortality may occasionally occur. When the majority of a fish population dies overnight or in a 24-hour period, oxygen depletion and other water quality changes, chemical toxicants, or other environmental causes should be strongly considered. Mortalities due to nutritional deficiencies are usually chronic and protracted.

Clinical signs “Clinical signs” is a term used to describe clinical findings, which include behavioral, physical, or other pathological changes in affected animals; when combined with gross internal pathology changes, these can be helpful in diagnosing fish diseases. However, in most cases, it is not possible to diagnose a specific disease or determine a specific etiological agent based solely on clinical signs because very few are specific (pathognomonic) for a particular disease (Table 3.1). In this text, emphasis is placed on recognition of clinical signs that are readily apparent by gross inspection. Behavior Initial reaction of fish to many diseases is a cessation of feeding activity; therefore, any sudden drop in feed consumption should be investigated because stress of any type can cause fish to “go off feed.” Diseased fish may lose their equilibrium, swim lethargically into shallow water or at the surface, lie listlessly on the pond or tank bottom, float down stream, swim erratically, or rub against underwater structures as if trying to eliminate a skin irritant. Some pathogens cause afflicted fish to swim in a longitudinal spiral or “tail chase” in a circle. Crowding around a water inlet to take advantage of oxygenated water may be a sign of diseased gills or low dissolved oxygen. Sick fish may gather in a tight school or ride high in the water. External External signs of infectious fish diseases are varied and generally not distinctive for a specific disease; however, in a few instances, infection is characterized by specific types of lesions (Table 3.1). Most obvious external clinical signs are inflammation and/or hemorrhage of fins, skin, or head; frayed fins; open necrotic and ulcerative lesions at any location on the body; and lepidorthosis, and excess or total lack of mucus. Excessive mucus on the

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Table 3.1 Summary of behavioral and external clinical signs and internal lesions of fish and types of diseases that could be associated with them. Clinical Sign or Lesion

Behavior Reduce or stop feeding Lethargic swimming Crowding to freshwater Spinning, erratic swimming External Gills necrotic Gills with excess mucus Gills are pale Skin with excess mucus Depigmented areas in skin Dark skin pigmentation Hemorrhage, erythema Frayed, eroded, erythema in fins Exophthalmia, hemorrhaged opaque eyes Ulcerative, necrotic lesions Hydropsy Lepidorthosis Enlarged abdomen (fluid) Growths, nodules, raised white spots on skin Internal gross pathology Clear, yellow fluid in peritoneal cavity Cloudy, bloody fluid in peritoneal cavity Viscera generally hyperemic Liver pale, friable Liver mottled, hyperemic, petechiae present Liver, kidney, spleen with white nodules Spleen dark red, swollen Kidney dark red, pale, soft Intestine, erythemic, flaccid, void of food Kidney with large white, irregular nodules Muscle with petechiae, necrotic pustules Muscle with white, yellow nodules

skin gives fish a grayish appearance. Edema, an enlarged abdomen, growths on the body surface, presence of yellow or black spots on the skin, prolapsed anus, and exophthalmia are all clinical signs of disease. Varying degrees of erythema, hyperemia, or hemorrhage may also be observed on the body and fins. Gills are adversely affected by viruses, bacteria, and parasites as well as toxins in the water or possibly dietary imbalance. They can become frayed and necrotic (gray or white), pale (anemic), swollen as a result of hyperplasia, or can produce excessive mucus that

Type of Disease

Viral, bacterial, parasitic, environmental factor Viral, bacterial, parasitic, fungal, water quality Bacteria or parasites on gill, low oxygen Viral, parasitic, toxicants Bacterial, parasitic, fungal Bacterial, parasitic, environmental, nutritional Viral, bacterial, nutritional Parasitic, environmental Bacterial, parasitic Viral, bacterial, nutritional, eye parasites Viral, bacterial, parasitic, toxicants Bacterial, parasitic, mechanical or physiological disorder Viral, bacterial, parasitic, gas supersaturation Viral, bacterial, parasitic Bacterial, viral, metazoan parasites Bacterial, viral, parasitic Viral, bacterial, parasitic Viral, parasitic, neoplasms, fungal Viral, bacterial rarely Bacterial Viral, bacterial Viral, bacterial, nutritional Viral, bacterial Parasitic, bacterial, neoplasms Viral, bacterial Viral, bacterial Viral, bacterial Bacterial, parasitic, fungal Viral, bacterial Metazoan parasites

interferes with oxygen absorption and causes affected fish to crowd around water inflow. Very pale gills (and internal organs) of channel catfish has been associated with a folate deficiency caused by a bacterium in the feed (Plumb et al. 1991). Skin lesions of diseased fish are diverse and usually nonspecific. Initially, they are often slightly raised or swollen, depigmented areas, where the epithelium has not been totally necrotized or sloughed. In advanced stages of disease, or when pathogens severely affect the skin, the epithelium and dermis can be totally

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missing (necrotized and sloughed into the water), leaving exposed musculature. Necrotic lesions accompanied by areas of deep ulceration can occur in muscle, or on opercles, head, or mouth. Margins of ulcerative lesions are often white (necrotic) and surrounded by inflammation, hemorrhage, or erythema in the epithelium. In the center of such lesions, muscle bundles (myomeres) are often distinguishable. Lesions caused by infectious agents often appear first on fins that may become extensively necrotic (frayed), hyperemic, or pale. Injury may progress until soft tissue of the fins is destroyed, leaving only hard spines, or fins can be completely missing. Scale pockets may show lepidorthosis (raised scales), or the body may appear swollen as a result of muscle edema (hydropsy) and presence of fluid in the abdominal cavity (ascites). Growths on the body and fins may appear as cottony-like material (fungus); solid tumors of various size, color, and texture; or as parasites embedded in/or beneath the skin. Embedded helminths will be yellow, white, or black spots in the skin, muscle, or organs. In fish that have systemic bacterial infections, the anus is often prolapsed, swollen, and red; however, this condition should not be confused with the swollen vent of a gravid female ready to spawn. Protruding (exophthalmia), hemorrhaged, or opaque eyes may be associated with an accumulation of fluid (edema) or inflammatory exudate behind the eye caused by bacterial infection, helminth parasites, or supersaturation of gas in the water. Opaqueness of the eye may also be due to a trematode or nematode infestation, dietary deficiency, or bacterial infection. Mortalities caused by most acute water quality changes or toxicants generally affect all species of fish present and will occur over a short period of time. These fish generally show no external lesions or clinical signs other than flared gill covers and gaping mouth at death. However, lesions such as brown blood (methemoglobin anemia) caused by elevated nitrite can occur. Some chemical toxicants will cause

hemorrhagic inflammation of the skin and fins. Fish suffering from oxygen depletion gasp at the surface and have dark red gills. Fish deformities are common and usually involve the head and/or spine. Juvenile trout infected with the parasite Myxobolus cerebralis (whirling disease) can develop head deformities and/or curvature of the spine. Spinal deformities may result in scoliosis (lateral curvature) or lordosis (forward or vertical curvature), which may be congenital, a result of dietary-related (i.e., vitamin C deficiency) factors, or infections (mycobacteriosis and IHNV).

Gross internal lesions After observing and recording behavioral and clinical signs of affected fish, a necropsy (postmortem examination) is required. Internal gross lesions may help to determine whether an infection is viral, bacterial, or parasitic (Table 3.1). Clear, straw-colored fluid in the abdominal cavity may indicate a viral infection. Bloody and cloudy fluid and/or large white pustule or variable size granulomas on visceral organs may indicate a bacterial infection. Some bacterial infections will cause the visceral cavity to have an intense putrid odor even in fresh fish. Small, white, or yellow uniform-sized cysts in internal organs are indicative of metazoan parasites. A uniform hyperemia or hemorrhage in the viscera is indicative of a viremia or bacterial septicemia in which case the spleen will usually be dark red and enlarged and the liver may be friable, soft and pale, or mottled with petechiae. The kidney is often swollen and soft in systemic bacterial or viral infections. In diseased fish, intestines are usually devoid of food, may contain white or bloody mucoid material, and the intestinal wall is often flaccid and erythemic. Internal organs may appear brown due to methemoglobinemia caused by nitrite toxicity. Some toxicants cause necrosis; chronic exposure to toxicants can result in several types of lesions.

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Disease diagnosis In diagnosing fish diseases, a small number of animals are examined, and information derived from these individuals is used to determine the health status of an entire population; therefore, it is essential that fish being examined are representative of those that are affected. Moribund specimens exhibiting clinical signs are best for examination and yield most dependable results. Fish that have been dead more than a few minutes should not be used for diagnosis unless put on ice immediately upon death, especially when water temperatures are warm. Dead fish that have lost normal skin color, have pale and soft gills, cloudy eyes, and an offensive odor have most likely been dead too long for proper necropsy. When a fish dies, parasites tend to leave the gills and skin, bacteria escapes the gut and invades visceral organs and body cavity, and waterborne bacteria may invade the skin. Also, apparently healthy hook- and line-caught fish are not suitable for necropsy because they may or may not be infected. Etiologies of fish diseases should not be determined from clinical signs alone because many viral, bacterial, or parasitic diseases have similar clinical signs. Paradoxically, clinical signs of ichthyophthiriasis (parasitic) and columnaris (bacteria) can provide reasonably accurate clues to the cause. In either disease scenario, a complete laboratory examination (necropsy) of affected fish should be carried out. A diagnosis without proper laboratory examination often leads to an erroneous diagnosis, improper treatment, and a waste of time, money, and fish. Diagnostic procedures should include examination for external parasites, attempted isolation of possible viruses and/or bacteria pathogen identification, and histopathology. If a pathogen is found on initial examination, all phases of the necropsy should be completed because multiple infections and successful therapy will require that all pathogens be addressed. Also, the presence of a particular pathogen does

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not necessarily imply that it is the etiological agent. A fish may host multiple pathogens even when it is healthy. When they become diseased, pathogen numbers often increase and may contribute to disease but may not be the primary cause of death. An accurate diagnosis requires integration of clinical signs, pathology, epidemiology, pathogen identification, and pathogen quantification. If the disease pattern and clinical signs are typical of the disease caused by the identified pathogen and pathogen numbers support this, the diagnosis can be made with a high level of confidence. Advanced molecular diagnostic methods of disease detection and viral, bacterial, and parasitic pathogen identification are becoming more routing in many fish disease diagnostic laboratories. These procedures provide rapid, sensitive diagnostic results.

Pathogen identification The science of pathogen identification is multifaceted. The identification of a pathogen as the cause of a disease requires identification of the pathogens on/in the diseased fish that are submitted for necropsy and an evaluation of their importance. There are several good references on methods for pathogen identification including the AFS Blue Book (AFS-FHS 2007) and the OIE Diagnostic Manual (OIE 2009). Often these manuals will emphasize specificity and sensitivity because the objective is to look for the presence of a defined pathogen in unapparent carriers, often having very low pathogen loads. In disease diagnosis, the focus is identifying the causative agent in affected fish. Therefore, the desire is for speed and breadth of coverage of potential pathogens. Generally the agent causing disease will be in sufficient numbers for easy detection and identification. The methods used for identifying pathogens include morphology, biochemical processes and growth characteristics in culture, and the presence of unique identifiable molecules (usually proteins or nucleic acids).

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Parasitic diseases This book does not directly address parasitic fish diseases, but a necropsy is not complete without considering these pathogens (Thoesen 1994; Lasee 1995; Mitchum 1995; Hoffman 1999; Noga 1996). Because alcohol or some other disinfectant is used on skin when isolating bacteria, an external parasitic examination should be made prior to aseptic necropsy. When examining fish for external parasites, material, taken from gills and skin including fins before disinfection (usually alcohol), is placed on a glass slide, a drop of water added, and a coverslip applied. The slide is examined first under light microscopy at low magnification (40–100×), and then at higher magnification (400–440×). Most protozoa, myxosporidians, helminths (monogenetic trematodes), and parasitic copepods are identifiable within these magnifications. Phase contrast is beneficial in detecting and identifying parasites. Internal fish parasites are detected by dissection and examination of internal organs and tissues including the muscle, gut, liver, spleen, and kidney. Many metazoan parasites are visible as small white, yellow, or black cysts and their presence is verified in wet mounts. Tissues may contain larval or adult nematodes, trematodes, or cestodes. Muscle-dwelling parasites, primarily larval trematodes, are detected by dissection of cysts, which are examined in wet mounts. Histopathological examination is valuable in detecting many tissue-dwelling parasites. Infestation severity and extent of injury to the host can be ascertained by this method. Viral diseases Diagnosis and detection of virus diseases of fish requires special expertise (Wolf 1988; Sanz and Coll 1992; Thoesen 1994). Detection methods include isolation of the virus in tissue culture, electron microscopic examination of tissue, and utilization of serological and molecular procedures. In spite of available

biotechnology and the fact that some viruses are known only via electron microscopy, isolation of viral pathogens in tissue culture continues to be the usual method for fish virus diagnosis. Type of tissue used for virus assay is based on size and age of fish; when assaying fry the entire fish is homogenized; only visceral organs are excised for virus assay of fingerlings, and parts or whole organs are used from larger fish. When checking brood fish for virus during egg collection, ovarian fluids are assayed. Collected tissue is homogenized in a stomacher or tissue grinder, diluted in physiological saline at 1:5 or 1:10, centrifuged, and the supernatants decontaminated by passing them through a 0.22–0.45-µm porosity filter. An alternative method for sanitizing homogenates is to incubate them in a solution of physiological saline containing a high concentration of antibiotics for 24 hours (LaPatra 2003). Viruses cannot be detected by light microscopy because they are generally less than 300 nm in size. Many are less than 100 nm; therefore, living cells grown in vitro are used to detect their presence. Viruses infect cultured cells and cause cell injury known as cytopathic effect (CPE), which can be seen with light microscopy. Many permanent fish cell lines have been developed for virus diagnosis and research. Appendix III lists the common cell lines used for fish diagnostics. A cell line derived from the same species or one phylogenetically close to the fish species being assayed for virus should be used when available. However, many known fish viruses replicate in a variety of fish cell lines. Once a cell line is chosen, filtrates or antibiotic sanitized materials are inoculated onto cell cultures. Depending upon cell line and virus assay samples, inoculated cells are incubated at 15–30◦ C for a few days to several weeks. If a virus is present and infectious to the inoculated cells, CPE will usually occur in 1–5 days, depending on virus and incubation temperature. A few fish viruses will require longer incubation to develop CPE. Type of CPE is

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often specific for a certain virus group, but differences are usually too subtle for positive identification. Once viruses are isolated, they can be positively identified by a variety of procedures that include serum neutralization, fluorescent antibody and enzyme-linked immunosorbent assay (ELISA) techniques, polymerase chain reaction (PCR) techniques (Schill et al. 1989), and other molecular procedures. Some of these techniques are also used to detect viral antigen or nucleic acids in infected fish tissue, which provides a more rapid diagnosis.

Bacterial diseases Diagnosis of most bacterial fish diseases is accomplished by culturing and isolating the pathogen on laboratory media and then identifying the organism biochemically, serologically, or molecularly. However, some bacterial species are fastidious and difficult to isolate and grow on culture media. Basic equipment needed for isolating bacteria includes dissecting kit, method for sterilizing instruments to be used, inoculating loop, sterilized bacterial culture agar, and incubation temperatures of 15–30◦ C (Plumb and Bowser 1983; Austin and Austin 1989; Lasee 1995; Noga 1996). Scrapings taken from body, fin, or gill lesions before disinfection should be examined in wet mounts for external bacteria and fungi. Skin and muscle lesions are disinfected with alcohol or hot scalpel before an incision is made and an inoculum is taken. Internal organs can be accessed through the abdominal region by disinfecting the skin surface and removing the muscle flap from the left side of the body cavity via three cuts ((1) medial from anterior of the rectum to the isthmus, (2) from anterior of the rectum along the dorsal line of the coelomic cavity, and (3) from isthmus to above the pectoral fin) to yield access to the peritoneal cavity, liver, spleen, or kidney. A second approach to accessing the kidney is to disinfect the dorsal area and cut through the back and spinal col-

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umn just posterior to the dorsal fin, thus penetrating the dorsal musculature to reach the kidney. Most bacteria that are known disease agents in fish grow on general laboratory media such as brain heart infusion (BHI), trypticase soy agar (TSA), nutrient, or blood agar; however, some specialized media are required for isolating mycoplasma, some flavobacteria, Renibacterium, some vibrios, Francisella, and mycobacterial organisms. Incubation temperatures are not critical for most bacterial pathogens of fish, but generally those from warm water fish are incubated at 25–30◦ C and those from salmonids at 15–20◦ C. Initial criterion for making a presumptive identification of fish pathogenic bacteria is that bacteria be isolated from clinically diseased fish. The presumptive bacterial identification is based on relatively few biophysical and biochemical tests using conventional bacteriological procedures and tube media (Plumb and Bowser 1983; Shotts and Teska 1989; Shotts 1994; Elliot 2003; Noga 1996). Criteria for presumptive bacterial identification include: cell morphology, motility, Gram stain reaction, production of cytochrome oxidase; reaction on carbohydrates (primarily glucose); indole, gas, and hydrogen sulfide production; temperature sensitivity; and sensitivity to certain growth inhibitors (0/129 or Novobiocin). Dichotomous keys are helpful in identifying many fish pathogens and can be used to classify most to genus and by using a variety of additional tests can be further classified to species (Shotts and Teska 1989; Shotts 1994; Shotts 2001). Additionally, when bacteria are cultured, antibiotic sensitivity tests are used to determine if available treatments are effective. The most common of these tests is the Kerby–Bower test that involves spreading a suspension of the bacteria on Mueller–Hinton agar and adding antibiotic sensitivity disks, and then measuring the zone of bacterial growth inhibition after 24- or 48-hour incubation.

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A battery of conventional tube media tests are used to identify bacteria but are labor intensive, and final identity may take several days. Numerous commercially prepared systems are available to shorten identification time, but these systems are not always accurate for fish pathogens. The most popuR R lar of these systems are API , Minitek , BiR TM olog GN Microplate , Abbott Diagnostics ,  R and Vitek Systems , all of which were originally developed for human or environmental microbiological application, and attempts to adapt them to fish pathogens have not been completely successful (Shotts and Teska 1989; Teska et al. 1989; Taylor 1995; Robohm 1997). These systems were developed for identifying pathogens of mammals; therefore, most fish pathogens are not in their numerical identification database. Also, the systems are designed for 35–37◦ C incubation temperatures when establishing the numerical codes. Identifying codes provided for bacterial pathogens of fish that may also occur in mammals can be different from those found in a laboratory that uses lower temperatures more appropriate for fish. Several diagnostic laboratories have found that these systems can be adapted for fish pathogens, but unique code identifiers must be established using known bacterial isolates; for example, E. ictaluri produces an API 20E code of 4004000 (Hawke et al. 1998). The codes for Yersinia ruckeri, Aeromonas salmonicida, and Vibrio anguillarum can be found in USFWS and AFS (2007). To speed up identification and increase accuracy, molecular-based methods are gaining popularity.

Molecular diagnostics Molecular methods are used for identifying parasite, bacterium, and virus pathogens in vivo and in vitro. These methods are based on identifying macromolecules that are common within the species yet unique to that species. Molecular methods can be very specific and are often used for confirming a presumptive diagnosis but may also be applied to tissues of in-

fected fish. However the narrow species specificity for most of these assays limits their utility during initial phases of a diagnostic procedure. In other words, the method can indicate the presence or absence of the pathogen for which the test was developed, but it provides no information about other pathogens. There are several nucleic acid-based methods that incorporate an amplification step. These methods are very sensitive and can detect the nucleic acid genome representing less than ten organisms. The molecular method can be based on fatty acids, complex carbohydrates, proteins, and/or nucleic acids. Fatty acids Fatty acids can be quickly and reliably identified by gas chromatography. This method is used to specifically identify many species of bacteria from cultures. It is particularly useful for mycobacteria because this group of bacteria is not conducive to traditional biochemical diagnostic tests (Mosca et al. 2007). The primary advantage of these systems is that they are more broadly applicable, and less presumptive, than other systems. The disadvantages are that they generally require isolated organism, and the equipment required to perform the assay is expensive. Antigen identification This can be a very sensitive method of identifying specific pathogens in infected tissue or for confirming the identification of a cultured pathogen. Antigens are generally large complex carbohydrates (e.g., lipopolysaccharides) or proteins. They are detected by producing antibodies by vaccinating an animal (usually a mouse, rat, rabbit, goat, or chicken). The serum from vaccinated animals can then be isolated from clotted blood and used as antiserum. If the animal is vaccinated with a whole organism, it will contain antibodies to many different antigens of the pathogen. If the animal is vaccinated with a purified antigen, it will develop antibodies, termed polyclonal antibody, to several portions (epitopes) of the

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antigen. An even more specific antibody preparation, monoclonal antibody, is specific to just one epitope of the antigen. Monoclonal antibodies are made by isolating spleen cells from an immunized mouse and fusing them with a tumor cell line, thus producing a hybridoma (clone) that generates one specific antibody for each hybridoma clone. The most simple of the methods that use antibodies are agglutination assays where antibodies clump particulate matter (usually bacterial cells) in a suspension. The BIONOR Mono-kit R, based on latex agglutination of bacteria, has proved to be quick and accurate in detecting most strains of Vibrio anguillarum, Yersinia ruckeri, and Renibacterium salmoninarum (Romalde et al. 1995). However, reagents used to identify V. splendidus and motile Aeromonas isolates have shown some cross-reactivity and false positives. The kits are considered cost effective and suitable for screening large numbers of samples when used in fish health laboratories but may not be universally applicable. With viruses, serum neutralization can be used in which antibodies inactivate the virus or interfere with its ability to infect cells; therefore, no CPE occurs in cell cultures inoculated with neutralized virus. The more common, more sensitive methods detect antibody binding to a substrate by attaching fluorescent dyes, enzymes, or marker molecules, such as biotin, to the antibodies. These systems can be used to detect antigens in tissues using Fluorescent Antibody Technique (FAT), indirect FAT (IFAT), and immunohistochemistry. Similar methods can be used on infected cell cultures. Also the presence of antigen in a tissue extract can be detected using ELISA or dot blots.

Nucleic acid identification Nucleic acid-based methods rely on the ability of complementary nucleotides to bind to each other. The sequence of nucleotides in a nucleic acid provides the specificity; only a complementary sequence can bind to the nucleic acid. Probes for detecting a genome of a pathogen

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are synthesized or produced from cloned plasmids in bacteria. These probes can be labeled with molecules similar to those used for labeling antibodies, or they can be used as primers for amplification reactions such as PCR. Several different amplification reaction systems are used for diagnostics, but variations of PCR or reverse transcriptase PCR (RT-PCR) are the most common. PCR uses a DNA substrate to generate 106 (or more) copies of a specific fragment of DNA between two DNA primers (that bind to complementary strands). For RT-PCR, an RNA target is first converted to DNA (cDNA) using reverse transcriptase, and then the cDNA target region is amplified by PCR. PCR products can be detected using gel or capillary electrophoresis or by performing a separate DNA–DNA hybridization assay. A recent modification of PCR that has gained much popularity in diagnostic laboratories is the TaqMan real-time PCR assay. In this system, a probe is made to the internal region of the PCR target. This probe is labeled with two fluorescent dyes, one of which quenches the other. When the Taq polymerase used for PCR encounters this probe, it breaks the probe thus releasing the marker dye from the quenching activity. This allows the PCR assay to be monitored as the reaction progresses, which is detected using simple fluorometry. TaqMan assays not only provide accurate, rapid methods for detecting pathogens, but they also provide reliable quantitative information; the time point in the assay when that product is first detected is inversely correlated to the amount of template (Holland et al. 1991). The price of equipment for performing real-time PCR assays has become reasonable for laboratories that would perform many PCR assays. The faster results and reduced labor have made these assays very popular in larger diagnostic laboratories.

Conclusion To perform reliable reputable diagnostics, you must use all available background

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information, pathology, and agent identification information that is practical and compare these findings to determine the most likely cause of the disease and the underlying factors. The combined information requires a subjective interpretation that is best done by individuals trained in aquatic animal health and familiar with the fish species, aquaculture system, and region where the disease problem occurs.

References AFS-FHS (American Fisheries Society-Fish Health Section). 2007. FHS blue book: suggested procedures for the detection and identification of certain finfish and shellfish pathogens, 2007 edition. American Fisheries Society-Fish Health Section, Bethesda, Maryland. Austin, B., and D. A. Austin. 1989. Methods for the Microbiological Examination of Fish and Shellfish. Chichester, England, Ellis Horwood Limited. Bouck, G. R. 1980. Etiology of gas bubble disease. Transactions of the American Fisheries Society 109:703–707. Chinabut, S., C. Limsuwan, and P. Kitsawat. 1991. Histology of the Walking Catfish, Clarias batrachus. International Development Research Center, Canada. Cotran, R. S., V. Kumar, and S. L. Robbins. 1989. Robbins Pathologic Basis of Disease, Fourth Edition. Philadelphia, W. B. Saunders Company. Elliot, D. 2003. General procedures for bacteriology. In: Suggested Procedures for the Detection and Identification of Certain Finfish and Shellfish Pathogens, Blue Book Fifth Edition. Bethesda, Maryland, Fish Health Section, American Fisheries Society. Ferguson, H. W. 1989. Systematic Pathology of Fish: A Text and Atlas of Comparative Tissue Responses in Diseases of Teleosts. Ames, Iowa, Iowa State University Press. Genten, F., E. Terwinghe, and A. Danguy. 2009. Atlas of Fish Histology. Enfield, Enfield Publishers, 215pp. Grizzle, J. M., and W. A. Rogers. 1976. Anatomy and Histology of the Channel Catfish. Auburn

University, Alabama, Alabama Agricultural Experiment Station. Groman, D. B. 1982. Histology of the Striped Bass. Bethesda, Maryland, American Fisheries Society. Hawke, J. P., R. M. Durborow, et al. 1998. ESCEnteric Septicemia of Catfish. Stoneville, MS, USDA-CSREES Southern Regional Aquaculture Center Publication 447. Hoffman, G. L. 1999. Parasites of North American Freshwater Fishes. Ithica, New York, Cornell University Press, 537pp. Holland, P. M., R. D. Abramson, et al. 1991. Detection of specific polymerase chain reaction prod  uct by utilizing the 5 –>3 exonuclease activity of Thermus aquaticus DNA polymerase. Proceedings The National Academy of Science. USA 88:7276–7280. Horsch, C. 2008. Normal Trout Histology. National Conservation Training Center. Shepherdstown, WV. LaPatra, S. E. 2003. General Procedures for Virology. In: Suggested Procedures for the Detection and Identification of Certain Finfish and Shellfish Pathogens, Blue Book Fifth Edition. Bethesda, Maryland, Fish Health Section, American Fisheries Society. Lasee, B. A. (ed). 1995. Introduction to Fish Health Management. U.S. Fish and Wildlife Service, Department of the Interior, Washington, D. C. Meyer, F. P., and L. A. Barclay (eds). 1990. Field Manual for the Investigation of Fish Kills. U.S. Fish and Wildlife Service, Resource Publication 177, Washington, D. C. Mitchum, D. L. 1995. Parasites of Fishes in Wyoming. Cheyenne, Wyoming, Wyoming Game and Fish Department. Morrison, C. E. 1987–1993. Histology of the Atlantic cod, Gadus morhua: An Atlas, Volume I–IV. Ottawa, Canada, Department of Fisheries and Oceans. Mosca, A., F. Russo, et al. 2007. Utility of gas chromatography for rapid identification of mycobacterial species frequently encountered in clinical laboratory. Journal of Microbiological Methods 68:392–395. Noga, E. J. 1996. Fish Disease Diagnosis and Treatment. Wiley-Blackwell Press, 488pp. OIE 2009. Manual of Diagnostic Tests for Aquatic Animals, Sixth edition. Paris, France, World Organization for Animal Health.

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Pathology and Disease Diagnosis

Plumb, J. A., and P. R. Bowser. 1983. Microbial Fish Disease Laboratory Manual. Auburn University, Alabama, Alabama Agricultural Experiment Station. Plumb, J. A., P. R. Liu, and C. E. Butterworth, Jr. 1991. Folate-degrading bacteria in channel catfish feeds. Journal of Applied Aquaculture. 1:33–43. Roberts, R. J. 1989. Fish Pathology, Second edition. London, Bailliere Tindall. Robohm, R. A. 1997. An evaluation of the use of Biolog GN MicroplateTM reactions in constructing taxonomic trees for classification of bacterial fish pathogens. 22nd Annual Eastern Fish Health Workshop. Atlantic Beach, North Carolina, March 18–20 (Abstract). Romalde, J. L. B. Magarinos, et al. 1995. Evalua˜ tion of BIONOR Mono-kits for rapid detection of bacterial fish pathogens. Diseases of Aquatic Organisms 21:25–34. Sanz, F., and J. Coll. 1992. Techniques for diagnosing viral diseases of salmonid fish. Diseases of Aquatic Organisms 13:211–223. Schill, W. B., G. L. Bullock, and D. P, Anderson. 1989. Serology. In: B. Austin and D. A. Austin (eds) Methods for the Microbiological Examination of Fish and Shellfish, 98–140. Chichester, England, Ellis Horwood Limited. Shotts, E. B., Jr. 1994. Flow chart for the presumptive identification of selected bacteria from fish. In: J. C. Thoesen (ed.) Bacterial Diseases of Fish. Bluebook: Suggested Procedures for the Detection and Identification of Certain Finfish and Shellfish Pathogens, Fourth Edition. Bethesda, Maryland, Fish Health Section/American Fisheries Society. Shotts, E.B., Jr. 2001. Flow chart for the presumptive identification of selected Bacteria from fishes. In: Suggested Procedures for the Detection and Identification of Certain Finfish and Shellfish Pathogens, Blue Book Fifth Edition. Bethesda,

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Maryland, Fish Health Section, American Fisheries Society. Shotts, E. B., Jr., and J. D. Teska. 1989. Bacterial pathogens of aquatic vertebrates. In: B. Austin and D. A. Austin (eds) Methods for the Microbiological Examination of Fish and Shellfish. Chichester, England, Ellis Horwood Limited, pp. 167–186. Taylor, P. W., J. E. Crawford, and E. B. Shotts, Jr. 1995. Comparison of two biochemical test systems with conventional methods for the identification of bacteria pathogenic to warmwater fish. Journal of Aquatic Animal Health 7:312– 317. Teska, J. H., E. B. Shotts, and T. C. Hsu. 1989. Automated biochemical identification of bacterial fish pathogens using the Abbott Quantum II. Journal of Wildlife Diseases 25:103–107. Thoesen, J. C. (ed.). 1994. Bluebook: Suggested Procedures for the Detection and Identification of Certain Finfish and Shellfish Pathogens, Fourth edition. Bethesda, Maryland, Fish Health Section/American Fisheries Society. USFWS and AFS-FHS. 2007. Standard procedures for aquatic animal health inspections. In: AFSFHS. FHS blue book: suggested procedures for the detection and identification of certain finfish and shellfish pathogens, 2007 edition. Bethesda, Maryland, AFS-FHS. Wolf, K. 1988. Fish Viruses and Fish Viral Diseases. Ithaca, NewYork, Cornell University Press. Yasutake, W. T., and J. H. Wales. 1983. Microscopic Anatomy of Salmonids: An Atlas. Resource Publication 150. U.S. Fish and Wildlife Service, Department of the Interior. Washington, D. C. Yonkos, L. T., D. J. Fisher, et al. 2000. Atlas of fathead minnow normal histology. An online publication of the University Of Maryland Aquatic Pathology Center. http://aquaticpath. umd.edu/fhm. Accessed 01/15/10.

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Chapter 4

Disease management

In addition to practicing good fish husbandry, a fish disease control plan includes proactive health management, judicial use of appropriate and approved chemotherapeutics when disease occur or the application of selected chemicals after handling, and the use of vaccines when available. A “team effort” is important to good fish health management and the central figure of that team should be a fish health professional. The individual can be a trained fish health specialist or veterinarian as long as he/she is familiar with good fish husbandry practices and understands the aquatic environment and its relationship to fish health.

Fish health management Fish health management can be classified as “applied art” or a “science” depending on one’s perspective; in reality it is probably a combination of the two where science is combined with “good ol’ common sense.” A number of encompassing fish health management principles was outlined earlier; therefore, this section emphasizes aspects that relate more directly to disease prevention with speHealth Maintenance and Principal Microbial Diseases of Cultured Fishes, 3rd edition. By J. A. Plumb and L. A. Hanson. Published 2011 by Blackwell Publishing Ltd.

cific management tools and control/prevention with drugs and vaccination. Fish health management through environmental manipulation may not totally eliminate infectious disease in cultured fish, but when disease does occur, its impact will be minimized. There are numerous examples of fish diseases that are directly associated with poor environmental conditions (Table 1.1), many of which can be addressed and mitigated by environmental management. Fish physiology is important in health management because as intensity of the culture system increases, physiological response increases proportionally. Wedemeyer (1996) discussed in detail the relationship between an aquatic environment and homeostasis of fish, particularly in intensive systems, and the practical application of water management blended with science and how these affect disease susceptibility. Deviations from optimal environmental conditions are stressful and may result in reduced feed consumption, higher feed conversion ratios, poor growth, disease, and/or death. Management procedures that help reduce stress as it relates to increased disease susceptibility are (1) fish handling and stocking, (2) feed management, (3) water flow and temperature management, (4) aeration management, (5) controlling other environmental problems, and (6) waste management. 57

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However, when discussing these aspects, it is difficult to consider them individually because they do not function independently.

Fish handling and stocking As one old saying goes “fish are not potatoes” and should not be handled as such. Fish should be handled gently whether as individuals or groups. Careful handling during transport, sorting, spawning, and stocking is critical to disease susceptibility as the protective mucus layer covering the skin can be disrupted and injury to the skin and gills will provide an opportunity for pathogens, some of which occur naturally in most waters, to become established. A good time to apply sodium chloride or potassium permanganate (KMnO4 ) as a prophylactic to help prevent secondary bacterial infections is while fish are still in hauling or holding tanks following handling. It is usually counterproductive to move fish strictly to treat them. Slowly changing water temperature from one environment into another (tempering) is advisable for most fish. Tempering allows the fish’s physiology to adapt to its new environment; time required varies depending upon fish species and temperature differential. Some fish species (i.e., salmonids) require several hours, over which time temperature is slowly changed, to be properly tempered, while other species (i.e., carp) require less time for this process. The greater the temperature differential, the longer tempering time required. Fish generally can adapt to lower temperatures more quickly than to higher temperatures; therefore, fish handling should be minimized during hot weather. Crowding is a common factor that adversely affects the general health of fish (Wedemeyer 1996). Carrying capacity in raceways can be elevated by increasing water flow rates, but according to R. Schmittou (Department of Fisheries and Allied Aquacultures, Auburn University, personal communication), over-

crowding itself is seldom a limiting factor in pond or cage culture. However, increased fish density is dependent on quality and quantity of feed required and can be adversely affected by any water quality problems associated with feed management. Carrying capacity of fertilized bodies of water is usually 25–150 kg of fish per hectare (about 30–175 lb/acre) without supplemental feeding. Under these conditions, there is sufficient natural food, oxygen, and other lifesustaining substances to adequately support a fish population, but this is not profitable. Therefore, an aquaculturist in the business of rearing fish for profit must maintain a high fish population density requiring supplemental feed. When nutrients in the form of feed are added to support higher stocking densities (fish numbers and weight per unit volume), water quality can be adversely affected and potential for disease increase. In an effort to improve productivity, some catfish farmers elevate fish densities from approximately 10,000 fish per hectare (4,500/acre) to 20,000 fish per hectare (9,000/acre). A catfish pond stocked with fingerlings to yield as much as 10,000 kg of food size fish per hectare often results in environmental deterioration because of the greater quantity of feed required to sustain growth for that biomass. Uneaten feed, fecal, and metabolic waste contribute an enormous organic load to the water providing an abundance of nutrients to support plant and bacterial growth, which in time consume oxygen through organic decomposition. As stocking densities increase from low to moderate, high and super intensive aquaculture, disease becomes more critical, but not necessarily as a direct result of increased numbers of fish per volume of water. However, increased population density enhances pathogen transmission from fish to fish. More importantly supplemental feeding is required to support the greater numbers and weight of fish and this additional nutrient input proportionally increases environmental instability and can cause extreme fluctuations in critical water

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Table 4.1 Projected production in a 131 ha (288 acre) catfish farm stocked at two fish densities and harvested annually for 3 years (4 ponds per treatment) based on experimental data from smaller scale studies. Criteria

Treatment A∗

Treatment B∗

Stocking density (no./ha) Average No harvested/year Mean survival Average net harvest (kg/yr) Average wt./fish (kg) Feed conversion ratio Aeration hours Net Revenue/ha ($US)

11,120 8,800 79% 5,200 0.59 1.37 1,606 $3,560

19,770 13,500 68% 6,600 0.49 1.56 2,120 $3,550

Source: Tucker et al. (1992). ∗ Each pond harvested annually and number of fish removed were replaced with equal number of seed stock.

quality factors (oxygen, ammonia, carbon dioxide concentrations, etc.). Consequently, a change in one management technique (addition of feed) often must be matched by another management procedure to offset any detrimental effects of additional feed (addition of aeration or increased water flow). Tucker et al. (1992) reported that catfish ponds stocked at 11,120 fish per hectare (4,448 fish/acre) produced 5,200 kg/ha (2,080 lb/acre) per year and ponds stocked at 19,500 fish per hectare (7,800 fish/acre) produced 6,600 kg/ha (2,640 lb/acre) per year (Table 4.1). Although the higher stocking density resulted in a slightly greater weight of harvested fish, individuals did not grow as well, had greater size variation, reduced survival, higher feed conversion ratios, increased need for aeration, and increased waste, thus poorer overall fish performance. Actual net profit in each culture system was about the same. Another factor that increases the correlation of disease-related losses with stocking density is the ability of the pathogen to spread in a population. When a single species of fish is cultured at a high density, an obligate pathogen can replicate and spread more quickly in the population because there are more susceptible fish per unit volume. This not only increases the efficiency of transmission, but in the progress of a disease out-

break, higher concentrations of pathogens can be found in the water. In this situation, naive fish become exposed to higher numbers of the pathogen. The higher initial dose of pathogens can more readily overwhelm the fish’s defenses against infection and cause disease. The correlation of transmission with density has been modeled and experimentally tested with furunculosis in Chinook salmon (Ogut et al. 2005).

Feed management Quality of feed should be the primary consideration in feed management. Feed should always be fresh, of the highest quality, and consistent with nutrient requirements of a particular species, size of fish, and type of culture unit involved (Lovell 1989). A second consideration in feed management, as it relates to disease, is the amount and frequency of feed applied per unit of surface area. Accumulated metabolic waste in water creates a demand for oxygen as it decomposes and serves as a source of nutrients for phytoplankton, which in turn exert an oxygen demand. As feed rates increase, phytoplankton growth also increases. The phytoplankton can produce compounds that impart an earthy or musty flavor, “off flavor,” to the fish, affecting marketability and

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also, during the night and low light conditions, phytoplankton use oxygen. Therefore high feeding rates cause a decrease in water quality and a need for additional management procedures (i.e., aeration, reduction in feeding rates, phytoplankton bloom control, etc.). Furthermore, phytoplankton species compete for light and nutrients. The phytoplankton populations are always in flux; when blooms become too dense, they may produce toxic compounds that may kill or stress fish. Also, if the dense bloom suddenly dies (algae die-off), there is extremely high oxygen consumption with no oxygen production, creating a low dissolved oxygen crisis. These die-offs are often associated with a change in weather or water temperature or a chemical treatment and are characterized by a sudden clearing of water or change in bloom color from green to brown. These events are often catastrophic in nutrientrich ponds (that receive high amounts of feed) because the oxygen demand can exceed available oxygen without significant supplemental aeration. Detailed information about fertility and water quality management in warm water ponds can be found in Boyd (1990) and Boyd and Tucker (1998). Much of the aforementioned feed management pertains to warmwater and coolwater aquaculture, but in coldwater aquaculture, flow through systems or pen culture for salmonids over feeding can be a problem if water flow or exchange is insufficient to remove excess food, excreta, etc. A recently investigated approach to disease prevention is the practice of feeding “probiotics” to fish (Irianto and Austin 2002). A wide variety of potential probiotics including algae, yeasts, and selected Gram-positive and Gram-negative bacteria have been evaluated. They concluded in their literature review that by feeding probiotics, growth of fish pathogens were inhibited; however, the reasons for this phenomenon were not stated. It is theorized that the probiotics inhibit colonization of potential pathogens or compete for nutrients and/or space, thus altering microbial

metabolism and stimulating the host immune response. Bacillus sp. and Aeromonas sobria recovered from the digestive tract of rainbow trout and carp, respectively, demonstrated that they were effective probiotics when fed to juvenile rainbow trout (Brunt et al. 2007). Those fish fed the probiotics exhibited reduced susceptibility to A. salmonicida, Lactococcus garvieae, Streptococcus iniae, Vibrio anguillarum, V. ordalli, and Yersinia ruckeri. Control groups not receiving the probiotics suffered 80–100% mortality compared with 0–13% mortality of groups fed the probiotics.

Water flow management In flow through culture systems (i.e., trout raceways), water quality problems associated with high fish densities are overcome by increasing water flow volume in conjunction with maintaining adequate dissolved oxygen levels. However, in large volume static culture ponds, routine continuous freshwater exchange is usually limited or nonexistent and actually may be harmful (Lorio 1994). Recirculating water for 8 hours at night in catfish ponds increased yield by a factor of 2.5 but these data may be misleading because of a lack of adequate controls in the study. Indications are that excessive water flow in ponds can adversely affect the environment and has little overall effect on pond water quality and continuous water flow in ponds has not proved to be a deterrent to infectious diseases. However, during periods of poor water quality, particularly oxygen depletion, addition of good quality water can be beneficial by providing a zone of safety and may prevent severe stress and reduce disease susceptibility. In all culture systems, caution should be used when increasing water flow to improve poor water quality because a water source may be low in dissolved oxygen or contain high amounts of carbon dioxide or nitrogen gas.

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Aeration management Natural oxygenation of static water in ponds is a result of photosynthesis driven by sunlight, consequently oxygen levels fluctuate diurnally, being highest in the afternoon and lowest during the early morning hours before daylight; therefore, oxygen should be monitored 24 hours a day during warm weather. To calculate a pond’s oxygen concentration at dawn, measure the oxygen concentration at dark and 3 hours later and extrapolate a straight line to predict the oxygen concentration at dawn (Figure 4.1) (Boyd et al. 1978). If the predicted oxygen concentration at dawn is below minimum levels (i.e., 3 mg/L) mechanical night aeration should be initiated during night time hours. Mechanical aeration technology is a major reason for the rapid growth of aquaculture. A variety of aerators (paddle wheels, agitators, or sprayers) (Figure 1.5) are available; each having unique advantages (Boyd and Wattson 1989). Energy sources for mechanical aerators are tractor power-take-off, propane gas, diesel fuel, or electricity.

Dissolved oxygen (mg/liter)

10 Measured values

Measured value

5

Projected value 0 2000

2400 Hours

400

Figure 4.1 Projection technique for estimating nightime dissolved oxygen (DO). By measuring oxygen concentrations at 8:00 PM and 12:00 midnight, the DO at daylight can be estimated by straight line projection. (Boyd et al. 1978; printed with permission of the American Fisheries Society.)

Nightly aeration during warm weather prevents stressful oxygen depletion that can lead to increased disease incidence in pond cultured fish. During the growing season, 6 hours of aeration at night can almost double channel catfish production and improve feed conversion ratios compared to ponds that are not aerated (Lai-Fa and Boyd 1988). Successive cloudy days have a significant detrimental effect on water quality in aquaculture ponds. Piedrahita (1991) found by computer modeling that short-term management schemes (lowering pH, raising alkalinity, and increasing nitrogen or phosphorus) were not effective for alleviating oxygen loss or depletion. The conclusion was that the best way to counter cloudy-day water quality syndrome is to reduce water depth and flush freshwater through the pond at a rate of at least 20% of the volume per 24 hour period. This of course is only practical in small ponds or where there is a substantial amount of water available. It is essential, however, that oxygen concentrations in the incoming water be higher than that of the pond water. Aeration in closed or recirculating systems is achieved by using compressed air, liquid oxygen, pure oxygen, or by various mechanical agitators. Bubbling air or oxygen directly into the culture system or pipeline manifold is probably not as efficient as introducing oxygen or air by injection into a sealed column of water (Doulos et al. 1994).

Other environmental problems Oxygen concentration is not the only environmental water quality parameter important to health of cultured fish. Maintaining low levels of carbon dioxide, nitrite, and ammonia, as well as proper pH, alkalinity, and water hardness are also important (Table 1.3). Carbon dioxide is normally elevated during pond respiration and decomposition of organic matter, but seldom poses a problem because healthy

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algae and plants utilize it almost as fast as it is produced. However, high CO2 can be a problem in closed systems. Water acidity, which is dependent upon its buffering capacity (alkalinity), should be pH 6.5 to 9. For aquaculture purposes, total alkalinity and total hardness should be above 20 mg/L (CaCO3 ) with values over 50 mg/L preferred (Boyd 1990). Low or widely fluctuating pH, low alkalinity, or hardness in ponds can be partially corrected by adding agricultural lime and in recirculating or flow through systems by dripping lime into the water or passing water through a bed of crushed oyster shells. Ammonia is seldom a problem in open ponds, but does accumulate in closed recirculating systems and can be controlled with biological filters. Some of these water quality problems result from high fish density and can be partially corrected using manipulative management; reducing the standing crop, limiting quantity of feed, or increasing aeration. In the long term, routine but judicial application of Environmental Protection Agency (EPA) registered herbicides, such as copper sulfate or aquazine, can control massive blooms of phytoplankton or filamentous algae. Care must be taken in the application of such chemicals so that a sudden, massive die-off of algae is not created, which will adversely affect dissolved oxygen concentration (oxygen depletion). When natural waters are used to fill ponds, raceways or house cages in saltwater or freshwater, the water source must be free of chemical pollutants or contaminants that may affect fish performance. Arkoosh et al. (2001) reported that juvenile Chinook salmon exposed to estuary water contaminated with chlorinated and aromatic compounds in Puget Sound, Washington, were more susceptible to Vibrio anguillarum. In geographical areas where water temperatures normally fall below optimal levels for growth and health of cultured fish, water from warm underground aquifers or heated water from power plants can be used to increase water temperatures in rearing systems. Heat

exchange units or solar heating can also be used to warm a relatively small amount of water.

Waste management Uneaten feed, accumulation of fecal waste, and dead plant matter in the culture system will elevate the water’s organic load. In trout raceways with high water exchange rates, most of this waste material is flushed out, whereas in ponds it settles on the bottom. Contrary to popular belief, old pond soils do not necessarily contain high quantities of organic matter (Boyd et al. 1994). Bottom soil of new shrimp ponds in southern Thailand contained 1.1% organic matter, while sediment in recently drained established shrimp ponds contained only 1.9%. Optimum respiration and oxidation of organic material occurred when soil moisture was 12–20% and at pH 7.5–8.0 (Boyd and Pippopinyo 1994). Addition of either calcium hydroxide or calcium carbonate enhances respiration of acidic soils. Pulverizing the hard surface crust of dried pond soil by tilling will also accelerate respiration. In recirculating closed culture systems, organic material must be removed by mechanical scrubbers and biological filters. Herbivorous fish can help control vegetation in environments with high fertility and an over abundance of phytoplankton or vascular plants. Grass carp eat vascular plants while silver carp and other filter feeders consume phytoplankton. Controlling aquatic vegetation with herbivorous fish is preferred to chemicals when practical. Benefits are usually longer lasting, more economical, and more user friendly. Precautions must be taken to avoid escape of exotic species into natural waters where they could be ecologically detrimental. Because of this possibility, exotic carp are prohibited by law in some states; however, in some instances stocking sterile triploid grass carp is allowed.

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Drugs and chemicals Aquaculture has expanded globally during the past 50 years aided by the use of drugs for combating infectious fish diseases. However, chemotherapeutics are only a small part of a comprehensive management plan and should not be relied upon exclusively to solve all health problems in aquaculture. Rather than being proactive, it is reactive. Use of drugs has in part allowed farmers to increase carrying capacity for greater food production levels, but in some instances, drugs have been detrimental. An over dependency on drugs, their indiscriminant, and improper use can be harmful to the animal being produced as well as the consumer and is ill-advised in today’s environmentally conscious society. As previously noted, any drug or chemical applied to fish, regulatory, public health, and biological constraints must be clarified and any specific drug/chemical use in the United States must be approved as a drug by the U.S. Food and Drug Administration Center for Veterinary Medicine (FDA CVM) or registered as a pesticide by the U.S. EPA. Proper aquaculture management will reduce the need for drug or chemical use to prevent or alleviate infectious diseases in fish, but despite appropriate management measures drugs will continue to be a necessary tool in aquaculture. Objectives of drug applications are: 1. Restore fish populations to health. 2. Protect economic livelihood of fish producers. 3. Ensure the continued production of animal origin. 4. Minimize the shedding of zoonotic pathogens into the environment and potentially the human food chain. Drugs are used to treat an existing infection (therapeutic) or prevent a disease from occurring (prophylactic); however, the prophylactic use of many drugs is prohibited in the United

States but is allowed in some other countries. Advantages of chemotherapy include (1) immediate availability, (2) advance purchase for some drugs and chemicals for future need, (3) different modes of administration (i.e., oral, immersion, and injection) under a variety of conditions, and (4) ability to affect multiple pathogens. Drug effectiveness is often short term and once withdrawn disease may reoccur, especially if all pathogens are not eliminated or predisposing disease factors are not corrected. For example, bacterial kidney disease may not develop in trout or salmon as long as erythromycin (currently not FDA approved but under investigation) is incorporated into feed, but when treatment ceases, clinical infection reappears (Groman and Klontz 1983). Enteric septicemia of catfish often behaves in a similar manner when infected fish are given drugs, particularly when water temperatures remain optimum for the pathogen for an extended period of time. By the time chemotherapy is applied, the pathogen being treated has often taken a toll on the fish population either in mortalities, growth reduction, and/or productivity. Also, chemotherapy presents a variable cost problem because it cannot be predetermined how often treatments will be required. Bacteria may develop resistance to antimicrobials if used too often, over an extended period of time, or if applied improperly (ChaslusDancla 1992; Dixon 1994). Some drugs and chemicals used on fish may be environmentally hazardous or toxic to treated animals, the applicator, or consumer. Accumulation of drug residues in aquaculture products is a concern and one reason why drugs are closely controlled in most countries. In spite of aquaculture’s huge global expansion, numbers of approved chemotherapeutics available for food fishes is limited, and the market potential for sale of pharmaceutical products in aquaculture is small when compared to beef, swine, or poultry. Schnick (1999) and Stoffregen et al. (1996) emphasized a lack of drug availability for aquaculture in

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the United States, but Schnick (1999) also emphasized that progress toward approval was being made on a number of drugs through partnerships and new leadership from FDA CVM even though the approval process governed by the FDA can be arduous, time consuming, expensive, and the outcome uncertain making it difficult to persuade pharmaceutical companies to invest in drug research and development for aquaculture. Also, regulatory constraints further limit drugs from being used or developed in many other countries. Drug regulations vary among countries, but with increasing international trade of aquaculture products, regulations between countries have become more interrelated.

Treatment process Infectious diseases will be minimized if the health maintenance principles outlined in Chapter 1 are incorporated into good fish health management programs. However, even the highest level of health maintenance will not completely eliminate infectious diseases; therefore, the question of control will eventually arise. Treatment of most aquaculture diseases is considered on a population basis because it is impractical, often impossible, to treat individually diseased fish unless only small numbers (i.e., brood stock) are involved. In a confined aquaculture environment, if one fish in a population has an infectious disease, it is assumed that the entire population has at least been exposed and all fish must be treated. Disease treatments and procedures described here are designed for cultured fish populations. In most wild populations, diseases are not treatable at this time for either practical or economic reasons. Since drugs seldom completely eradicate all pathogens, their application is often simply “buying time” until fish can overcome infection through their own defensive mechanism. A therapeutic agent can reduce infection, temporarily prevent pathogen reproduction, or retard pathogen growth, thus giving the host’s natural or acquired defen-

sive mechanisms time to develop and overcome disease. Wellborn (1985) proposed a critical question to be considered before treatment is applied: What is the prognosis if treatment is applied or withheld and does potential mortality justify treatment? If treatment is still indicated after answering this question, four additional criteria should be considered: (1) know the water, (2) know the fish, (3) know the drug, and (4) know the disease. Ignoring any one of these factors can result in an ineffective treatment and over the long term may prove detrimental to affected fish.

Know the water Before applying a treatment, water volume to be treated must be accurately calculated to prevent a lethal or ineffective treatment. Total alkalinity and hardness, pH, organic load, and temperature of the water are factors that influence efficacy and toxicity of some drugs.

Know the fish Toxicity of some drugs may vary between fish species, strains within species, and between different age groups. If the drug of choice has never been used in a particular water supply to treat a specific fish species or age class of that species, toxicity should be tested by treating a few fish in a small vessel containing a sample of water to be treated and observing the drug’s effect on the fish.

Know the drug Toxicity and percent active ingredient of a drug must be known to correctly calculate the proper amount required. Some drugs are affected by sunlight, pH, temperature, organic water load, and alkalinity and all drug formulations may not have the same level of active ingredient. Some drugs are toxic to plants that oxygenate the water; therefore, decomposition of plants upon death contributes to reduced oxygen.

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Know the disease The disease must be accurately diagnosed by a complete necropsy of infected fish. Multiple infections involving different pathogens often occur and will require multiple treatments beginning with the most serious pathogen. An incorrect diagnosis and treatment can prove disastrous to the fish.

Therapeutic applications The most effective and economical method of drug application is determined on a case-bycase basis and is dependent upon the disease, drugs prescribed, type of unit to be treated, and age and species of fish. Six basic methods for treating fish diseases are (1) dip, (2) flush, (3) prolonged bath, (4) indefinite, (5) orally in the feed, and (6) injection.

Dip The dip method is usually used to treat small numbers of easily confined fish infected with external parasites or bacteria. Fish to be treated are immersed in a strong drug solution for a short time, usually 15 to 60 seconds. Drug concentrations can be expressed as percentage of material, milligrams per liter (mg/L), microliters per liter (µL/L) [both equivalent of parts per million (ppm)], or a ratio of chemical to volume of water (i.e., 1:5,000). Dipping fish requires caution because of strong drug concentrations used and the potential for toxic response. Time of immersion is critical; therefore, it is best to treat a few fish to determine their reaction before treating an entire population. A disadvantage of this procedure is the necessity to handle fish.

Flush Flushing is a method popular at trout and salmon hatcheries for treating eggs and fish in raceways for external parasites and bacteria. A flush treatment involves adding specific

amounts of drug to a trough, tank, or raceway and allowing it to flush through without interrupting water flow. Drug concentrations are usually expressed as mg/L, µL/L, or as a ratio of drug to volume of water and can last from a few minutes up to 1 hour. In the latter, a stock solution of concentrated drug is introduced by a continuous flow delivery system, which assures a desired concentration throughout the entire treatment period. The flush treatment is seldom used on warm water farms or hatcheries except for egg treatment.

Prolonged bath Prolonged treatment can be employed for either external parasitic or bacterial infections utilizing high drug concentrations; caution is required to insure equal distribution to avoid “hot spots” that can be toxic to fish. A prolonged bath treatment exposes fish to a drug for a specified period of time. Concentrations are normally in mg/L or µL/L, or as a ratio of drug to volume of water. The drug is added to a trough or holding unit at the desired concentration and left for a predetermined time, usually 1 hour, under static conditions with aeration. When treatment is terminated, water flow is resumed and the drug is flushed out as quickly as possible. Fish should be observed continuously during treatment and if any signs of discomfort, such as gasping, "flashing," or loss of equilibrium are noted, the drug must be flushed immediately regardless of exposure time. It is essential that an adequate amount of water is available at all times for rapid flushing if needed.

Indefinite Indefinite treatments are used for parasitic or bacterial infections. Indefinite treatment means that a drug is introduced into a pond or tank at comparatively low concentrations for an undetermined length of time and allowed to break down and dissipate naturally. Drug concentrations are usually mg/L or µL/L (ppm); therefore, this method is generally considered

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safe for treating fish. Indefinite treatments often require large quantities of drug that are difficult to apply, especially in large ponds requiring application from a boat, but can be applied to small bodies of water by hand or sprayers. Easily dissolved dry chemicals, such as copper sulfate or KMnO4 , are dissolved in water and then dispensed with a boat bailer or siphoned depending upon kind and quantity of drug being applied. It may be necessary to apply the drug uniformly to avoid “hot spots” and supplemental aeration should always be available because some drugs will cause an algae die off, resulting in loss of dissolved oxygen.

Oral Treatment of systemic bacterial infections and intestinal parasites requires incorporation of antimicrobial or antihelminthic drugs into feed (no antihelminthic drugs are currently approved for fish in the United States). The only legal way to use medicated feed in the United States is to purchase commercially prepared feed containing the drug. Drugs are either incorporated into heat-extruded floating feeds or for heat-sensitive drugs into sinking pellets. Medicated feed is administered to fish based on body weight to be treated. Standard units of treatment are given in grams of active ingredient per 45 kg (100 lb) of fish or in milligrams active ingredient per kilogram (mg/kg) of body weight per day for a defined number of days. Drugs are incorporated into feed at a concentration that delivers a desired dose per unit of fish weight per day and fish are fed at a specific feeding rate (percentage of body weight divided into a defined number of feedings per day). Prophylactic use of drugs in feed for short periods of time or continuous feeding at low dosage is not advised because these practices are not favored by FDA and they enhance drug resistance of bacteria. A “withdrawal time”, defined as the period between the last day of drug exposure and day of slaughter (or in the case of fish released into public waters, the time when

they can be legally caught, is required when drugs are fed to food animals to ensure that no drug residues of concern are present in the flesh when eaten. Withdrawal time varies with drug, fish species, and water temperature when administered.

Injection Broodfish, or small numbers of other valuable fishes, can be treated for some diseases by drug injection. Currently, no antimicrobial drugs are approved for injection in aquaculture in the United States; however, some of these drugs can be legally administered through extra label use on the orders of a veterinarian. Dosages to be injected are measured in either international units (IU) or milligrams (mg) of active drug per kilogram (or pound) of fish and are administered intraperitoneal (IP) or intramuscular (IM). IP injections are given in the posterior area of the body cavity by inserting the needle through the peritoneal wall at a 45◦ angle at the base of the pelvic fin, taking care not to puncture internal organs. IM injections are given slowly in the thick dorsal musculature near the dorsal fin. The IM method is slower to administer and quantity of injection material is reduced because of potential drug loss through the puncture wound. The injection application is usually used to treat bacterial infections and administer vaccines or spawning enhancers, but the need to handle each fish may increase stress if fish are already in poor health.

Calculations There are two basic methods of mass drug application for fish: (1) bath/immersion and (2) medicated feed. Once the drug of choice is established for bath/immersion, it is essential to know the volume of water to be treated, concentration of drug to be used, and percent active ingredient to calculate how much drug to apply. Formula 4.1 is used to calculate the

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proper amount of drug required to treat static tanks, raceways, or ponds. Appendix I provide a conversion table from Arabic to metric measurements and other information to aid in calculating volume or weights of drug or chemical required.

Formula 4.1 Wt. or vol. 100 Desired × × (V.1) water % Activity conc. =

Wt. or volume of drug to use

Example No. 1: To treat 1000 L of water at 2 mg/L (ppm) with a drug that is 100% active the calculation is as follows: 100 × 2 (mg/L) 100 = Wt./vol. of drug required

1000 (litres) ×

Therefore: 1000 × 1 × 2 = 2000 mg (2 g) of drug required. Example No. 2: To treat a 5 acre (2.02 ha) pond that averages 4 ft. (1.22 m) at 4 mg/L (ppm) with a drug that is 50% active. One acre foot is 1 surface acre (43,560 ft2 ) that is 1 foot deep; therefore, the pond contains 20 acre feet of water. Also, 1 acre ft. weighs 2.7 million pounds; therefore, 2.7 pounds of anything in 1 acre foot = 1 mg/L (ppm), and 1 acre foot contains 1,238,230 litres of water. Using the above formula, the calculation is 100 × 4 (mg/L) 50 = mg of drug required per acre ft. of water 1,238,230 (litres) ×

Therefore; 1,238,230 × 2 × 4 = 9, 905, 840 mg of drug = 9.9 kg/acre ft. of water × 20 acre ft. 198.0 kg (89.8 lb) of drug required to treat the 5 acre pond.

When drugs are fed to fish it is essential to know the weight of fish to be treated, treatment rate (weight of drug fed per unit weight per day), and concentration of active ingredient in the drug. Formula 4.2 is used to calculate how much drug to feed per day.

Formula 4.2 Wt. of Fish ×

100 % Activity

× Treat. Level = Wt. of Drug (mg/kg/day) Example No. 3: To treat 100 kg of fish with a drug that is 25% active at a rate of 100 mg/kg of fish per day, the calculation is 100 × 100 (mg/kg) of Fish 25 = Wt. of drug/day of Fish 100 (kg) ×

Therefore: 100 × 4 × 100 = 40,000 mg (40 g) of drug mixed in a 1 day ration.

Drugs for fish Drugs were used with little or no restrictions to prevent or treat fish diseases in the United States until the mid-1960s when a more cautious attitude emerged concerning their indiscriminate abusive use necessitated due to increased oversight by FDA. Herwig (1979) listed approximately 275 different drugs and chemicals that had at one time been used in aquaculture, some of which were ineffective, uneconomical, known carcinogens, remained in fish flesh for long periods of time or were dangerous to the applicator or harmful to the environment. However, as the potential for drug residue contaminating fish flesh, residue being passed on to consumers or drug use in fish enhancing drug-resistant human pathogens became apparent, steps were taken worldwide to restrict and eliminate the use of unsafe drugs for treating fish diseases.

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FDA CVM has primary responsibility for approving drugs used for disease control or production efficiency in animals, including fish, while EPA has jurisdiction over chemicals applied to water to control pests. Since fish are a “minor” agricultural product, they fall under the FDA CVM definition of a minor species (Anonymous 2007). Drugs to treat fish must meet certain data requirements before approval. Drugs for fish must go through a rigorous Investigational New Animal Drug (INAD) application process during which requisite data are developed. Following critical review of submitted data, CVM determines whether or not a drug can be used on the target animal or if more data are necessary. Once such data as product chemistry, effectiveness, and safety to target animals, humans, and environment are acceptable to FDA CVM, the sponsor (usually the manufacturer of the drug) may then file a New Animal Drug Approval (NADA) application with FDA CVM. As a result of this lengthy process, only a few drugs were approved until recently for use on fish in the United States. The FDA CVM allows for the legal use of drugs under the following classifications: (1) Approved drugs for fish that are the subject of an NADA, (2) unapproved drugs of Low Regulatory Priority (LRP), (3) unapproved drugs with deferred regulatory status, (4) unapproved drugs for which data are being generated under an INAD, and (5) approved drugs that are the subject of case-specific regulatory discretion through extra label use (Bowker 2006; Anonymous 2008; Johnson 2008). Information that will be discussed in this chapter includes drugs currently being used in aquaculture for disease treatment, including status with the FDA, methods of application, concentration and dosage, length of application, limitations, and diseases for which they are used. All drugs should be handled with care; therefore, safety precautions including wearing protective clothing, rubber gloves, breathing mask, and safety glasses should be taken when weighing, measuring, and apply-

ing them. If the skin comes into contact with any drug, it should be thoroughly washed immediately.

Approved drugs for fish In the United States, sponsors currently have received FDA CVM approval of four feed additive antibacterials and two immersion drugs for parasitic and fungal control, one each as a sedative, marking aid, and spawning aid for food fish in aquaculture (Table 4.2). Some drugs may have more than one product approved for a specified application method for a specified disease of usually a specific fish species. Technically, their use for other diseases or other fish species is illegal unless used under an INAD or via extra label use. Currently, approved drugs are administered to treat diseased fish in one of two ways: as feed additives or by immersion.

Feed additive antibacterial Most antibacterials are incorporated into the feed at the feed mill at a concentration to provide the recommended dosage per kilogram (0.45 lb) of fish when fed at a specific percentage of body weight per day. All feed antibacterial substances require a minimum withdrawal time after the last day the drug is given before the fish are slaughtered or stocked into waters where they would be susceptible to angling. There are four drugs currently approved for treating bacterial diseases of fish: (1) florfenicol, (2) oxytetracycline dihydrate, (3) sulfadimethoxine and ormetoprim, and (4) sulfamerazine.
R

Florfenicol (Aquaflor ) Florfenicol (Aquaflor; Intervet/ScheringPlough Animal Health) is the latest antibacterial approved by FDA CVM for aquaculture. Aquaflor has broad spectrum efficacy for Gram-negative and Gram-positive bacteria

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Table 4.2 Drugs approved by the U.S. Food and Drug Administration, Center for Veterinary Medicine for disease control in and handling foodfish. Withdrawal (days)

Drug/Chemical

Species

Indication

Dosage

Feed addatives  Aquaflor R ∗ (Florfenicol)

Channel catfish,

Enteric septicemia, columnaris, Coldwater disease

10 mg/kg fish/days/ 10–12 days 10 mg/kg fish/days/ 10–12 days 50–75 mg/kg fish/days/ 10 days

12

50–75 mg/kg fish/days/ 10 days

21

220 mg/kg fish/days/ 10 days

21

Salmonids Terramycin  200 R For Fish†

Sulfamerazine Fish Grade‡  Romet 30 R § (Romet TC) Immersion/bath  Formalin F R ¶

 Parasite-S R ¶ Perox-Aid∗∗

Salmonids

Furunculosis, ulcer disease, bacterial hemorrhagic septicemia, pseudomonas disease

Catfish

Salmonids

Bacterial hemorrhagic Septicemia, pseudomonas disease Furunculosis

Salmonids

Furunculosis

50 mg/kg fish/d/5 days

42

Catfish

Enteric septicemia

50 mg/kg fish/d/5 days

3

All species Fish eggs

External protozoa Raceways, tanks for monogenetic trematodes,

Penaeid shrimp Salmonid eggs, Juveniles Channel catfish, Warmwater and coolwater sp., all stages

 Finquel R ††  Tricaine R ††

15

All species All species

Raceways, tanks: >10◦ C 170 µL/L for 1 hour <10◦ C 250 µL/L for 1 hour Fungus on eggs 1000–2000 µl/L for 15 minutes Ponds: 15–25 µL/L indef. Parasites, fungus on eggs 50–100 µL/L for 4 hour Ponds: 15 to 25 µL/L indef. Fungus 500–1000 mg/L, 15 minutes Bacterial gill disease 100 mg/L, 30 minutes Fungus on eggs, and 50–100 mg/L 60 minutes Columnaris alternate days for 3 days 50–75 mg/L, 60 minutes Anesthetic 50–100 µL/L Anesthetic 50–100 µL/L

Source: JSA Working Group on Aquaculture Drugs, Biologics, and Pesticides 2007. ∗ Florfenicol requires a veterinary feed directive. † Oxytetracycline dehydrate. ‡ Not currently available. § Sulfadimethoxine-ormetoprim in 5:1 ratio. ¶ About 37–36% formaldehyde gas. ∗∗ 35% hydrogen peroxide. †† Tricaine Methanesulfonate, “tricaine,” and MS-222.

21

NA

NA

21 21

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but is currently approved only for control of mortality due to Gram-negative bacteria (Table 4.2). There is no indication that Aquaflor leads to reduction in feed consumption of fish and the drug is safe for human consumption when administered to fish according to label directions. R Aquaflor is approved for control of mortality due to enteric septicemia of catfish (Edwardsiella ictaluri) and conditionally approved for control of mortality due to columnaris disease (Flavobacterium columnare) in channel catfish. It is also approved for control of mortality due to coldwater disease (Flavobacterium psychrophilum) and furunculosis (Aeromonas salmonicida) in freshwaterR reared salmonids. Aquaflor is the first drug to be classified as Veterinary Feed Directive (VFD) requiring a veterinarian prescription. R Oxytetracycline dihydrate (Terramycin
200 for Fish) Oxytetracycline dihydrate (Terramycin 200 for Fish; Phibro Animal Health) is a bacteriostatic antibiotic powder added to feed to control bacterial hemorrhagic septicemia (motile Aeromonas spp.) and pseudomonas disease in catfish, ulcer disease, furunculosis, bacterial hemorrhagic septicemia, and pseudomonas in salmonids, the control of mortality due to coldwater disease in freshwaterreared salmonids, and columnaris disease in rainbow trout (Table 4.2). In addition, TerR ramycin 200 for Fish is approved as a feed additive for control of gaffkemia caused by Aerococcus viridans in lobster and as a marking aid for Pacific salmon. Oxytetracycline has been used so extensively in aquaculture (sometimes improperly) that a high percentage of target organisms, especially motile Aeromonas spp. and typical Aeromonas salmonicida, have become resistant to it. A disadvantage is that Terramycin 200 for Fish is only available in sinking pellets because the heat required to extrude floating pellets during manufacture destroys the drug.

Sulfadimethoxine and ormetoprim R R (Romet-30
, Romet TC
) Romet-30 (Pharmaq AS) is a bactericidal potentiated sulfonamide in powder form in a combination of five parts sulfadimethoxine and one part ormetoprim. The drug is approved for controlling enteric septicemia in catfish and furunculosis in salmonids (Table 4.2). Romet-30 is incorporated into feed to provide 50 mg/kg of fish per day for 5 days. At higher drug concentrations fish tend to refuse feed; therefore, it is prudent to use a reduced drug concentration and increase feeding rate accordingly. Romet-30 is heat resistant; therefore, can be incorporated into extruded floating pellets. Mortalities are reduced quickly when fish consume feed containing Romet-30; however, a low incidence of drug resistance has been encountered with frequent infection of enteric septicemia of catfish recrudescence (Plumb et al. 1995). Romet TC is a Type B medicated feed that was developed to improve the palatability of the feed to fish. Romet TC can be mixed onsite without a feed mill license. Sulfamerazine (Sulfamerazine R in Fish Grade
) Sulfamerazine in Fish Grade (Pharmaq AS) is approved for treating furunculosis in rainbow, brook, and brown trout (Table 4.2). However this drug is no longer manufactured or marketed in the United States, but it may be available elsewhere.

Immersion Immersion treatments are either as short term (bath or flush), where the drug/chemical is flushed after a specified time, or long term, where the drug is allowed to dissipate naturally. Two immersion drugs approved for control of diseases in fish are formalin (four trade names supplied by several manufacturR ers) and hydrogen peroxide (35% Perox-Aid supplied by one manufacturer).

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Disease Management 71 R R Formalin (Formalin-F
, Paracide-F
,

R R Parasite-S , and Formacide-B ) The four currently FDA CVM-approved formalin products for controlling certain external parasites on fish and penaeid shrimp and fungus on fish eggs and their sources are Formalin F (Natchez Animal Supply), Paracide-F (Argent Chemical Laboratories Inc.), ParasiteS (Western Chemical Inc.), and Formacide-B (B. L. Mitchell, Inc.). Formalin is a clear liquid containing 37–40% formaldehyde gas, but for treatment calculations, it is considered 100% active. Formalin-F, Parasite-S, and FormacideB formalin products are approved as a parasiticide in tanks, raceways, and ponds for all species of fishes (Table 4.2). Care must be taken with formalin treatments during warm weather when oxygen depletion may occur within several days of application, thus requiring aeration for fish survival. Formalin-F, Parasite-S, and Formacide-B formalin products are also approved for application as a flush treatment for fungus on all fish eggs and penaeid shrimp, and as an external protozoan treatment in tanks.

R Hydrogen peroxide (35% Perox-Aid
) Hydrogen peroxide (35% active H2 O2 ) (Eka Chemicals, Inc.), trade name 35% PEROXAID, is approved for control (1) saprolegniasis (fungi of the family Saprolegniaceae) on all freshwater-reared finfish eggs, (2) bacterial gill disease (F. branchiophilum) in all freshwaterreared salmonids, and (3) external columnaris (Flavobacterium columnare) in all freshwaterreared coolwater fish and channel catfish (Table 4.2). Hydrogen peroxide is to be used only in tanks and raceways, not in earthen ponds with no water exchange. Hydrogen peroxide should not be used on any life stages of northern pike (except eggs) or pallid sturgeon fry and applied on walleye with caution. Walleye are sensitive to the drug; they developed extensive epithelial lifting and necrosis on gill of fish treated with high doses (Tort et al. 2002).

Other approved drugs Three additional FDA CVM-approved drugs labeled for application in aquaculture for purposes other than disease treatment are tricaine R methanesulfonate (Finquel and Tricaine R R S ), chorionic gonadotropin (Chorulon ), and oxytetracycline hydrochloride (various sponsors) (Table 4.2). Tricaine methanesulfonate (Finquel and Tricaine-S) Tricaine methanesulfonate (Finquel; Argent Chemical Laboratories Inc.; Tricaine-S, Western Chemical Inc.), also known as MS-222 and tricaine, is an anesthetic sedative used in a variety of aquaculture procedures that require tranquilizing and handling individual fish (Table 4.2). It is approved by the FDA for use in fish intended for food but should be restricted to Ictaluridae, Salmonidae, Esocidae, and Percidae. Chorionic gonadotropin (Chorulon) Chorionic gonadotropin (Chorulon, Intervet, Inc.) is a reproductive hormone for male and female fish to induce spawning of some species of fish (Table 4.2). Multiple IM injections (1 to 3) may be required to stimulate spawning, but the total Chorulon administered per fish intended for human consumption should not exceed 25,000 IU per fish of chorionic gonadotropin (25 mL). A prescription is required from a licensed veterinarian for use of this drug. Oxytetracycline hydrochloride (OxyMarineTM , Oxytetracycline HCL R R Soluble Powder-343
, Terramycin-343
,
R and TETROXY Aquatic ) Oxytetracycline hydrochloride soluble powder is approved for marking skeletal tissue (otoliths) of all finfish. The manufacturers of these products are OxyMarine, Alpharma; Oxytetracycline HCL Soluble Powder-343, IVX Animal Health; Terramycin-343, Pfizer

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Table 4.3 Unapproved drugs used in fish health but classified by the U.S. FDA as low regulatory priority. Drug

Indication

Dosage/Treatment

Acetic acid (vinegar)

External parasites

Calcium chloride

Potassium chloride Povodine iodine

Egg hardening Osmotic balance External parasites Anesthetic Helminths, sea lice Reduce adhesiveness of eggs Reduce metabolic rate Monogenetic trematodes external crustaceans Crustacean parasites Remove gelatinous Matrix on eggs Osmoregulator Egg disinfection

Dip at 1,000–2,000 mg/L for 1–10 minutes 10–20 mg/L for eggs; 150 mg/L fish transport Dip at 2,000 mg/L for 5 seconds Until fish loose equilibrium Unknown

Sodium Bicarbonate Sodium chloride (NaCl)

C02 into water as fish anesthetic Osmoregulatory, prophylaxis

Sodium sulfite Thiamine Hydrochloride

Improve hatchability of eggs Prevent thiamine deficiency

Calcium oxide Carbon dioxide gas Garlic (whole) Fullers earth Ice Magnesium sulfate (epsom salts) Onion (whole) Papin

(Do not use chlorinated ice) Dip 3,000 mg/L plus 7,000 mg/L NaCl for 5–10 minutes. Unknown 0.2% solution 10–20 mg/L 50 mg/L, 30 minutes during water hardening 142–642 mg/L for 5 minutes 0.5% indefinitely; 1–3% for second to 30 min or until loose equilibrium 15% for 5–8 minutes 100 mg/L during water hardening eggs

Source: Guide to Drug, Vaccine, and Pesticide Use in Aquaculture (1994).

Animal Health; and TETROXY Aquatic, Cross Vetpharm Group Ltd.).

Drugs of low regulatory priority (LRP) LRP Drugs are unapproved animal drugs for which sponsors are not likely to seek approval, especially for compounds like sodium chloride, garlic, or ice; however, the FDA recognizes that drugs classified as LRP have physiological impact on fish (MacMillan 2003). Sixteen compounds are listed by the FDA as LRP aquaculture drugs (Table 4.3). These drugs, including chemicals, and other household compounds are substances used in our everyday lives and the FDA is unlikely to object to their use if the following conditions are met (Anonymous 2007): (1) used for prescribed indications, including species and life stage where specified, (2) used at prescribed dosages, (3) used according to good management practices, (4) the product is of an appropriate grade for use in food animals, and (5) an adverse

effect on the environment is unlikely. These include acetic acid (vinegar), calcium chloride, calcium oxide, carbon dioxide gas, Fullers earth, garlic, and others (Table 4.3). The position of the FDA on the use of these substances should not be considered an approval or endorsement of their safety or efficacy. All of these compounds do not necessarily have fish disease treatment implications.

Deferred regulatory status Deferred regulatory status applies to two drugs, i.e., copper sulfate (CuSO4 ) and KMnO4 , that were registered in the past by the EPA for use as either a herbicide (CuSO4 ) or an oxidizing agent (KMnO4 ) and have shown some propensity as therapeutic in fish (Schnick et al. 1997). Regulation has been deferred while data are generated on their safety and effectiveness. The FDA has indicated that as long as these chemicals are used per EPA registered labels, no objections will be made if the products have an incidental effect on fish diseases.

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Copper sulfate Copper sulfate (CuSO4 ), known as bluestone, comes in two formulations, “crystal” or “snow,” both of which are considered 100% active. Copper sulfate, registered by the EPA as a herbicide, can be used in water containing food fish. The chemical also is a moderately effective drug for external bacterial and/or protozoan infections, particularly Ichthyophthirius multifiliis “Ich.” Because of its availability and price, copper sulfate is one of the most widely used chemicals in the fish husbandry industry worldwide. Copper sulfate has a narrow margin of safety between an effective treatment concentration and toxicity to fish; therefore, it is most often used in low concentrations as an indefinite treatment. Toxicity of copper sulfate varies with the copper formulation, water pH, total alkalinity, and water hardness (Straus and Tucker 1993). They also determined that chelated copper is significantly less toxic than copper in sulfate salt. These data, and general literature, support the position that copper sulfate should never be used unless water hardness and alkalinity are known. Generally 1.0 mg/L of copper sulfate is applied in water for each 100 mg/L total alkalinity (TA). More specifically, when TA is 0 to 49 mg/L, a toxicity test should be run before use; when TA is 50–99 mg/L, use 0.5–1 mg/L of CuSO4 ; 100–149 mg/L, use 1–2 mg/L of CuSO4 ; and 150–300 mg/L, use 2–3 mg/L of CuSO4 . In saltwater or if TA of freshwater is above 300 mg/L, copper sulfate precipitates rapidly and will most likely be ineffective; therefore, it is not used in saltwater or in highly alkaline waters.

Potassium permanganate KMnO4 , considered 100% active, is a purple crystalline material previously registered by the EPA for use in oxidizing organic matter in ponds. The EPA required all registrants of registered pesticides and water treatment compounds to resubmit data for re-registering their products; however, the registrant of KMnO4

(Carus Chemical Company) chose not to submit re-registration. However, KMnO4 is effective for some external protozoa (Epistylis spp.) infestations and bacterial infections, especially columnaris disease and can be applied in tanks at 5– 10 mg/L for 1 hour as a prolonged bath. Fish should be observed continuously and at the first hint of discomfort, freshwater introduced. Pond application rate is 2–4 mg/L, indefinitely depending on organic content of the water. In order to be effective, the KMnO4 concentration must be 2 mg/L over the oxidizing demand of water to be treated. The oxidizing demand is determined by the concentration of KMnO4 that is necessary to turn water from reddishpurple (active) to brown (inactive) within 15 minutes (Tucker and Boyd 1977). The burgundy color should remain for 12 hours after application to be therapeutically effective. KMnO4 reduces phytoplankton and may temporarily have a negative effect on oxygen levels. Cost is a disadvantage of KMnO4 if treating large volumes of water. The toxicity margin of KMnO4 is narrow (1–3 mg/L); therefore, caution is required at higher application rates. If fish are exposed to KMnO4 too often or the concentration is excessively high, severe gill injury will occur; therefore, a minimum of 5 days should elapse between treatments. Darwish et al. (2002) showed that in channel catfish, the gill tissue only developed mild hypertrophy during exposure to recommended doses of KMnO4 for 36 hours. When exposed to 3–5 times the rate of therapeutic doses the gill recovered and appeared normal at 8 days postexposure. Internal organ tissues were not affected.

Investigative new animal drugs (INAD) Several drugs have been investigated for potential application in treating fish diseases and are currently in some stage of consideration for INAD exemption. This category allows scientists to be involved in a specific drug approval

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74 Health Maintenance

process to test the efficacy of a drug in a commercial setting and test for safety in the laboratory. INAD exemptions must be obtained from FDA and involves scrutiny to assure testing is valid and that human, animal, and environmental safety is protected. If approved by the FDA, oral drugs would likely be available only through prescription by veterinarians. Some of the following drugs are or have been under INAD exemptions in the United States. The information presented below should not be construed as an endorsement or recommendation of any of these drugs.

Amoxicillin Amoxicillin is a penicillin-type antimicrobial that has shown promise for treating Streptococcus spp. infections in tilapia and hybrid stripe bass. It is also efficacious against Aeromonas salmonicida in salmonids for which it is approved in the United Kingdom.

Chloramine-T Chloramine-T is an antimicrobial that has potential for the control of mortality due to bacterial gill disease and external columnaris disease. This drug as HALAMID AQUA (sponsor: Acentive SARL) is nearing approval for these two diseases in a variety of finfish species in the United States.

Copper sulfate Copper sulfate as Triangle Brand Copper R Sulfate (sponsor: McMoRan Copper and Gold) is being pursued for approval for the control of mortality due to ichthyophthiriasis in channel catfish and saprolegniasis in channel catfish eggs.

Emamectin benzoate R Emamectin benzoate (Slice ; Intervet/ Schering-Plough animal Health) is approved in European countries for control of sea lice

on saltwater-reared salmonids and is being pursued for the same use in the United States

Erythromycin thiocyanate Erythromycin thiocyante is active against Gram-positive and some Gram-negative bacteria. Erythromycin has been administered experimentally by oral application or injection to control the mortality of bacterial kidney disease in Pacific salmon (Groman and Klontz 1983; Moffitt 1992). The oral application of erythromycin for the above-mentioned indiR cation is being pursued under Aquamycin (sponsor: Bimeda). Erythromycin may also be effective against Streptococcus spp. in tilapia and hybrid striped bass.

17 α-methyltestosterone 17 α-methyltestosterone is being pursued for approval as a gender manipulation aid for tilapia and ornamental fish in the United States under the auspices of the sponsor, Rangen, Inc. for its product Masculinizing Feed for R Tilapia .

Potassium permanganate R KMnO4 as Cairox (sponsor: Carus Chemical Company) is being pursued for approval for the control of mortality due to bacterial gill disease and coldwater disease in freshwaterreared salmonids and external columnaris disease in all freshwater-reared finfish.

Extra label drugs Extra label use drugs provides for drugs legally administered to one species for specific indications to be applied to another species for indications not listed on the label. These require prescription from a licensed veterinarian as defined under the Animal Use Clarification Act of 1994 (AMDUCA) (Anonymous 2007). This includes, but is not limited to, use in species or for indications not listed on the label. For

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Disease Management 75

example Terramycin- or Romet-30-medicated feed can be used in cultured aquatic animals other than salmonids and channel catfish if used as directed in designated target species. Criteria for implementing AMDUCA are:

r r r r

r r

Medicated feed or drug is already approved for use in aquatic species. A written recommendation is obtained from an attending veterinarian. The drug is used for therapeutic purposes only. The aquaculturist has accurate feed records, a current copy of veterinarian’s recommendation and production records that assure that the withdrawal time was met. The drug is used in accordance with federal, state, and local environmental laws and regulations. Application follows established safety provisions.

International use of drugs in aquaculture The past two decades has seen a significant increase in international aquaculture and an influx of international movement of finfish and shellfish food products to satisfy a growing demand for fish and shellfish to replace diminishing wild caught supplies. Concomitant with this increase is a growing concern over the possibility of fish products transmitting infectious diseases from one geographical region to another, and whether or not these food products contain drug residues that could be harmful to the consumer. The FDA assays products imported to the United States through international trade. According to Acheson (2007), during 2006 and 2007, the FDA identified residues of unapproved drugs in seafood imported from China including nitrofurans, malachite green, and gentian violet; none of these are legal in the United States. Many countries including Canada, United Kingdom, Taiwan, Germany and other European Economic

Community countries, and others also assay imported fish products for unauthorized drugs. In view of these international problems, the National Coordinator for Aquaculture New Animal Drug Applications (http:/aquanic. org/aquadrugs) has been working for international harmonization to establish tolerance levels of drugs, data requirements, and study protocols needed for aquaculture drug approvals. Each country in which aquaculture is practiced has different regulations concerning what drugs and/or chemicals can be used to combat infectious diseases. Whereas the United States has a limited number of legal drugs for aquaculture, other countries have greater availability of these drugs. For example, in 1992, Japan had 29 individual combinations of antibiotics for aquaculture (Okamoto 1992) at a time when the United States had two legal antibiotics. Schnick et al. (1999) listed 46 different drugs available in Japan, Australia, Europe, Canada and the United States. Some reports indicate that up to 50 different drugs are used in finfish and shrimp aquaculture in Asia alone (Arthur 1996; Johnston and Santillo 2002). The number of drugs available varies from country to country and region to region around the world and in some instances their regulation or control is rather lax. Young (2002) provided a partial list of drugs available to aquaculture outside the United States at that time:

• Acriflavin • Amoxicillin • Ampicillin • Benzocaine • Bicozamycin • Chloramphenicol • Colistin sulfate • Doxycyline • Erythromycin • Florfenicol • Flumequine • Fosfomycin • Fruluphenicol • Furnace • Furazolidone • Josamycin

• Kanamyacin • Kitasamycin • Lincomycin • Malachite green • Methyldihydrotestosterone • Methylene blue • Miroxisacin • Nalidixic acid • Nitrofurantoin • Novobiocin • Nifurstyrenate • Oleandomycin • Oxolinic acid • Spiramycin • Thimphenico

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Antibiotics are allowed to be used as a prophylactic in some countries, thus enhancing antibiotic resistance in fish pathogens and potentially in human pathogens as well as accumulation of drug residue in the edible flesh. In addition some of these compounds may be carcinogens; i.e., chloramphenicol. Indiscriminate use of drugs exacerbates all of these potential problems; therefore, it is essential to protect international food supplies. Fish raised in pens in open marine waters present a unique problem. When medicated feed is fed to caged fish, all of the pellets do not remain in the pen; thus, target fish do not consume all of it. Therefore, some of the medicated feed is eaten by wild fish in the environment. Fortt and Buschmann (2007) reported that wild fish outside of salmon cages in Chile that consume this feed accumulated antibiotics in the flesh, therefore posing an environmental issue and possible residue problem in wild fish flesh.

Future outlook With many aquaculture endeavors around the world adopting a holistic approach to fish production, thus eliminating many of the environmental and stress producing conditions that contribute to infectious diseases, the need for drugs has and will continue to decrease. Evidence currently indicates the benefit of better health management. The catfish industry in the United States is one example and the salmonid industry in Scandinavia is another. These fish health management approaches should be encouraged to reduce the dependency on drugs and chemicals for disease control. Although many countries have a more diverse list of “legal drugs” for use in fish than does the United States, before any of the aforementioned compounds are used, they should be approved by the governing agency of the country in which they are to be used. Also, if aquaculture products are destined for international trade, any compound being

used must be acceptable to the country of destination. In the European Economic Community (EEC), Asia and other regions where drugs and chemicals are used in aquaculture, the regulations controlling them are diverse. The subject of chemotherapy is dynamic, and most regulatory changes have resulted in more restrictive drug use and/or decline in drug availability for aquaculture. As the world population becomes more environmentally conscious and concerned with the role of drugs in human health problems, regulation and use of chemotherapeutics will become even more restrictive. In the final analysis, these restrictions will result in a more complicated and expensive drug approval process that will not encourage pharmaceutical companies to invest in research and development of new aquaculture drugs. On the positive side, these restrictions have brought about a significant shift in attitude away from reactive chemotherapeutic fish disease control to one of proactive prevention by management. Nevertheless, chemotherapy will continue to be important in aquaculture because chemicals and drugs are key components in health management, disease prevention, and control in cultured animals.

Vaccination The first experimental fish vaccination was reported by Duff (1942) when he fed a killed Aeromonas salmonicida preparation to cutthroat trout to prevent furunculosis. From the early 1940s to the mid-1960s, little effort was made to advance the potential for vaccine use in preventing and/or controlling fish diseases; this period might be characterized as the “golden age of antibiotics” because it was widely thought that chemotherapy could correct all ills, but it became obvious that drugs were not a panacea; therefore, alternative methods of disease control were sought. A renewed interest in development of fish vaccines occurred in the early 1970s and vaccination

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Disease Management 77

of cultured aquatic animals has now become a viable, proactive fish health management tool for many diseases. Like drugs, vaccines cannot solve all fish disease problems but when available can significantly aid the industry. Examples where antibiotics have failed to control infectious microbial diseases in fish adequately but proactive vaccination can be successfully implemented of fish were pointed out by Klesius et al. (2000). They state that “the best alternative method to chemical control is to prevent communicable diseases by stimulation of fish immune system by vaccination and use of immunostimulants.” An acceptable vaccine for aquaculture must have the following attributes and characteristics (Leong and Fryer 1993): 1. Provide adequate immunoprotection for a specific disease in intensive rearing conditions. 2. Provide protection when the animal is most susceptible to disease. 3. Provide protection of long duration. 4. Protect against all serotypic variants of the disease agent. 5. Easily administered; preferably orally, immersion, or spray and application should minimally disrupt the normal management routine but injection is also popular. 6. Safe for the vaccinated animal and not result in undesirable residue in the vaccinates. 7. Cost effective to produce, license, and apply. Implementing a vaccination program to prevent infectious fish disease is a “proactive” approach as opposed to a “reactive” approach that depends on drugs after the disease appears. Vaccines present distinct advantages over chemotherapy; they reduce disease impact, decrease the need for drugs, provide long-term protection that often continues until slaughter, provide more fixed-cost disease prevention expenditure, and in many instances vaccinated fish grow better than unvaccinated fish. Perhaps the greatest value of vaccines is

the fact that no residue is left in edible flesh of vaccinated animals, except possibly in the case of some adjuvants. Vaccines cannot completely eliminate disease organisms or prevent the target organism from being present in vaccinated populations because of individual immunological variances. Vaccinated fish can become carriers and consequently reservoirs of the disease organism. Vaccines are expensive to make and apply but in some instances a low percentage of improved survival justifies vaccination cost. For example, a 5% increase in survival in vaccinated channel catfish for ESC can economically justify vaccination. The first commercially licensed fish vaccine became available in 1976 to treat salmonids for enteric redmouth (ERM) (Yersinia ruckeri) and was soon followed by a vaccine for vibriosis (Vibrio anguillarum and V. ordalii). Since the introduction of these vaccines, they have been used internationally wherever ERM and vibriosis are a problem and are also occasionally used in treatment of nonsalmonids. Ellis (1988b) reported that at that time three bacterial vaccines were available for aquaculture and an additional five bacterial and five viral vaccines were being researched. In 1994, there were 15 licensed fish vaccines available in the United States for use against 5 fish pathogens, where today there are over 30 univalent vaccines (Table 4.4) and over 2 dozen multivalent vaccines (Table 4.5) now available globally for a variety of fish species. These vaccines provide protection against viral and bacterial pathogens but none are available for parasites. Autogenous bacterins made from specific isolates from individual aquaculture sites are also available from some biological companies. Fish vaccines are used extensively in Norway where at least six commercial biological manufacturers have marketed up to 24 different products to vaccinate salmon and trout against IPNV, V. anguillarum, V. ordalii, V. salmonicida, A. salmonicida, and Y. ruckeri. Vaccines available in the United States are

78 Bacterin Bacterin Bacterin Bacterin

Bacterin

AQUAVAC VIBRIO MICROVIB MICROViBoral MICROSALwVoral

CYPROVAC MICROVIRIPNV MICROVIRIHNV Kv3

Coldwater vibrosis: Injection Vibrio salmonicida Autogenous vaccines

Virus vaccines Infectious pancreatic necrosis Infectious hematopoietic necrosis Koi herpes virus

Adjuvanted Adjuvanted Live/attenuated

Bacterin Bacterin Bacterin Bacterin Bacterin

MICROPaS MICRO-SrS Norvax Strep Si NORVAX VIBRIOSIS VIBROGEN

Salmonids Salmonids Koi carp

Cultured Fish injection, oral

Seabream/bass Salmonids Warmwater fish Salmonids Salmonids, sea bream, sea bass Salmonids Salmonids Salmonids Salmonids

Channel catfish Channel catfish Channel catfish Channel catfish Salmonids Salmonids Salminids Salmonids Trout/salmon, koi carp Trout/salmon Salmonids Salmonids Salmonids Salmonids Koi carp Salmonids Salmonids Salmonids

Fish Species

Injection Injection Immersion,

Immersion,

Immersion Immersion, injection Oral Oral

Immersion Immersion Immersion/injection Immersion, Injection

Immersion Immersion Immersion Oral Oral Immersion Injection Immersion Oral Immersion injection Injection Oral Immersion Injection Injection Immersion Immersion, injection Oral Immersion

Application

Microtek Microtek KoVax Ltd.

Alpharma

Schering-Plough Microtek Microtek Microtek

Microtek Microtek Intrervet Intervet Intervet

Schering-Plough Aqua Health Schering-Plough Aqua Health Schering-Plough Schering-Plough Alpharma Microtek Aqua Health Aqua Health Schering-Plough Schering-Plough Alpharma Alpharma Schering-Plough Microtek Microtek Microtek

Company†

†

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Photobacterosis: Photobacterium damsela P. salmonis Streptococcus: Streptococcus iniae Vibriosis: V. anguillarum

Furunculosis: Aeromonas salmonicida

Live/attenuated Live/attenuated Live/attenuated Encapsulated Bacterin Bacterin Bacterin Bacterin Bacterin Bacterin, Bacterin Bacterin Bacterin Bacterin Bacterin Bacterin Bacterin Live/attenuated

Type of Vaccine

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Information compiled from literature supplied by the manufacturers and commercial sources. Alpharma, Redmond, Washington, USA; Intervet Norbio (Schering-Plough), Bergen, Norway; Aqua Health Ltd. Charllottetown, Prince Edward Island, Canada; AVL, Aquaculture Vaccine Ltd. Essex, United Kingdom; Microtek International Limited, Saanichton, British Columbia, Canada.

∗

AQUAVAC-COL FRY VAC AQUAC ESC ESCOGEN Aqua Vac∗ ERM Oral Aaua Vac∗ ERM ALPHAA JECT 1200 MICROPaM FUROGEN DIP FUROGEN 2 Aqua Vac∗ Furovac 5 Aqua Vac∗ Furovac 5 BIOVAX 1500 BIOVAX 1800 AQUAVAC MICROSaLimm,inj MICROSALoral MICRO-SaLAtten

Columnaris: Flavobacterium columnare

Enteric septicemia of Catfish: Edwardsiella ictaluri Enteric redmouth: Yersinia ruckeri

Vaccine Trade name

Disease (Pathogen)

Table 4.4 A partial list of monovalent vaccines for bacterial and viral diseases currently available for finfish aquaculture.∗

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Disease Management 79

for V. anguillarum, V. ordalii, V. salmonicida, A. salmonicida, and Y. ruckeri in salmonids and Edwardsiella ictaluri and Flavobacterium columnare in channel catfish. Vaccines are available as univalent or multivalent preparations and are applied by immersion, injection, spray, and orally. Vaccines have improved survival and production of salmon and rainbow trout populations in many areas where used. According to Press and Lillehaug (1995), in 1986, 3,500 litres of fish vaccine were used in Norway, 76,250 litres in 1988, and in 1992 vaccine quantity had decreased to 24,780 litres. A decrease in vaccine quantity may have been due to use of multivalent vaccines. In fact some reports claim that vaccines prevented catastrophe in the salmon farming in Norway from severe Vibrio spp. infections. The efficacy of vaccination of fish against infectious diseases has been shown in a number of studies in other regions of the world. Intervet reports that survival of yellow tail and seabass vaccinated against Vibriosis and Photobacterosis (Pasteurellosis) in Asia improved survival from 25–30% to 60–80% with relative percent survival (RPS) of 40–82% (Anonymous 2006). The monetary value of vaccination soared from $40,000 to $1.14 US per 100,000 fish in each group. Generally, the benefits of vaccination are reduction in disease and contribution to profitability if incorporated into a good disease monitoring and fish health management program. It improves survivability, secures availability of juveniles, and reduces feed conversion ratio, predictable production, and fish for export and trade. The cost/benefit ratio of vaccination far exceeds that of antibiotics and drugs for disease control. Another benefit exR ample of vaccination is the use of Norvax Strep Si for Streptococcus iniae in seabass. Vaccination by immersion improved survival from 22% in controls to 75% in vaccinates and by injection from 37% in controls to 100% in vaccinates. Much of the fish vaccine production technology has been adapted from that of pro-

ducing vaccines for homoeothermic animals. Early vaccines were monovalent and effective against only a single disease organism, but multivalent vaccines containing antigens of two or more pathogens (viral and bacterial) or strains have been developed, resulting in reduced volumes of vaccine being used.

Antigens Most of the early bacterial vaccine research, development, and application targeted salmonid diseases (A. salmonicida, Y. ruckeri, V. anguillarum, and V. salmonicida) but there is a growing effort to develop vaccines for cool and warm water fishes, some of which are now available. Research and development of immunological products for nonsalmonids include vaccines for the most serious bacterial pathogens that affect them: Flavobacterium columnare, Edwardsiella ictaluri, E. tarda, Photobacterium damsela subsp. piscicida (Pasteurella piscicida), and Streptococcus iniae. Many of these vaccines are multivalent, therefore, provide protection to vaccinated fish for multiple genetic types within a species of pathogen (Table 4.5). A vaccine developed to protect against a specific pathogen for a specific species or fish group (multivalent vibriosis and furunculosis vaccines for trout and salmonids) can also be used for nontarget species, although USDA approval is essential for extra label use. Rogers and Xu (1992) experimentally used a commercial V. anguillarum-V. ordalii bacterin developed for salmonids to protect striped bass against vibriosis. Fish virus vaccine research for aquaculture lagged behind that for bacterial pathogens; however, viral vaccines include attenuated, chemically inactivated, and subunit (recombinant DNA technology) preparations have been formulated (Leong and Fryer 1993). These include vaccines for IPNV, VHSV, IHNV, CCV, and spring viremia of carp virus (SVCV). Intervet Norbio (Bergen, Norway) developed a

80 Salmonids

Bacterin Bacterin Bacterin Bacterin Bacterin Bacterin, oil adjuvant Recombinant subunit of SRS and IPNV, killed V. ordalii Bacterin + recombinant virus protein

ALPHA JECT 5200 BIOVAX 1150 BIOVAX 1300 BIOJEC AQUAVAC FHV LIPOGEN TRIPPLE; LIPOGEN FORTE Bayovac 3.1

Salmonids

Salmonids Salmonids Salmonids

Salmonids

Salmonids

Immersion Iinjection

Injection

Injection

Injection

Immersion Immersion Injection

Injection

Injection

Immersion, injection

Intervet Intervet

Microtek

Aqua Health

Scherring-Plough

Alpharma Alpharma Alpharma

Alpharma

Alpharma

Intervet

†

Information compiled from literature supplied by the manufacturers and commercial sources. Alpharma, Redmond, Washington, USA; Intervet Norbio (Schering-Plough), Bergen, Norway; Aqua Health Ltd. Charllottetown, Prince Edward Island, Canada; AVL, Aquaculture Vaccine Ltd. Essex, United Kingdom; Microtek International Limited, Saanichton, British Columbia, Canada.

∗

Salmonids

Bacterin

ALPHA JECT4000

Salmonids

Alpharma Microtek Microtek

Company†

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NORVEX PROTECT-IPN

Salmonids

Bacterin with 2X adjuvant

NORVEX PROTECT

A. salmonicida, V. anguillarum, V. salmonicida, A. salmonicida, V. anguillarum, V. salmonicida A. salmonicida, V. anguillarum (serotypes 01 and 02), V. salmonicida A. salmonicida, V. anguillarum, V. salmonicida, V. viscosus V. anguillarum, V. ordalii V. anguillarum, V. ordalii A. salmonicida, V anguillarum, V. salmonicida A. salmonicida, V. anguillarum, V. salmonicida A. salmonicida, V. anguillarum V. ordalii, V. salmonicida Salmon Rickettsial septicemia, Infectious pancreatic necrosis virus, atypical Vibrio ordalii Vibrio anguillarum, Infectious pancreatic necrosis virus

Injection Immersion, Injection Injection

Application

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Salmonids Salmonids Salmonids

Bacterin Bacterin Bacterin

ALPHA JECT 3000 MICROTIVaCC3 MULTIVaCC4

Fish Species

A. salmonicida, V. anguillarum

Type of Vaccine

Vaccine Trade Name

Pathogens

Table 4.5 A partial list of multivalent vaccines for bacterial, ricketsial, and viral diseases currently available for finfish aquaculture.∗

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Disease Management 81

recombinant vaccine against IPN virus, which is marketed in a tetravalent preparation that also includes antigens for furunculosis, vibriosis, and coldwater vibriosis. According to some reports, millions of Atlantic salmon smolts have been vaccinated with multivalent vaccines, resulting in a 96% relative survival.

Vaccine preparation Various types of vaccines exist: inactivated vaccines that are essentially bacteria grown by conventional fermentation or virus’ grown in tissue culture and then inactivated with formalin, and live attenuated bacteria or viral preparations that are generally nonpathogenic. Also, using molecular biology, new types of vaccines using recombinant subunit and DNA are under development. At least one subunit DNA vaccine is now available that incorporates antigens of salmon ricketsia syndrome, Vibrio ordalii, and infectious pancreatic necrosis virus. Most early fish vaccines, as well as many currently available are simply killed pathogens. Although these bacterins are relatively easy to make, formalin or other additives may alter the organisms’ antigenic quality, thus reducing its efficacy. It has been shown that in some channel catfish diseases (i.e., E. ictaluri), and possibly others, live bacterial cells may be essential to stimulate a cell-mediated response and elicit protection (Shoemaker and Klesius 1997; Shoemaker et al. 1997). Vaccines containing live bacteria, usually in an attenuated form, also elicit a more rapid immune response than killed bacterins. However, it is possible that conventionally attenuated live pathogens can revert to a pathogenic form when released into an aquatic environment where water disposition cannot be controlled, a condition that concerns regulatory agencies and aquaculturists. Inducing attenuation of bacterial pathogens by altering their molecular structure or other

means of attenuation will likely eliminate reversion to a pathogenic form. To avoid pathogen virulence, Vallejo et al. (1992) proposed that synthetic peptides be used as fish vaccines. Vaughan et al. (1993) described a nonpathogenic aromatic-dependent preparation of A. salmonicida (aeroA) that protects trout against furunculosis. The advantage of aromatic-dependent mutants is that they will not revert to pathogenic form but retain their ability to produce antigenic components such as the A-layer. Attenuation can also be linked to a bacterium’s inability to synthesize p-aminobenzoic acid, which is important in pathogenesis. Cooper et al. (1996) used a nonpathogenic chondroitinase negative E. ictaluri to protect channel catfish against ESC. Cloning, mapping, and sequencing genes for individual proteins of A. salmonicida may also yield mechanisms by which immunogenic protective products will be pathogen specific (Bennett et al. 1992). Biotechnology used in vaccine production for humans and veterinary animals is now being applied to vaccines for aquaculture. Genetic recombination, attenuation, protein engineering, and subunit vaccines have proven to be promising and the possibility for reversion to pathogenicity is eliminated. According to reports from Norway, recombinant viral vaccine preparations are economically feasible, especially when combined with bacterins (Press and Lillehaug 1995). Development strategies for fish virus vaccines emphasize genetic engineering and molecular biology. Most of this work has targeted IPNV, VHSV, and IHNV of salmonids (Leong and Fryer 1993) but more recently has expanded to other pathogens. As already noted, a commercial DNA-recombinant vaccine for IPNV is available in Norway. The recombinant subunit vaccine, in the form of a crude bacterial lysate, protected fish against IPNV via immersion vaccination. A subunit VHSV vaccine was produced by employing the virus’s surface glycoprotein and inserting the gene into the bacterium Escherichia coli, thus

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82 Health Maintenance

making it possible for large quantities of viral glycoprotein to be inexpensively produced (Thiery et al. 1990). An experimental IHNV vaccine was constructed using this same technology; a crude lysate of E. coli containing fusion IHNV protein was used to immunize several salmonid species from challenge with a lethal dose of IHNV (Xu et al. 1991). In a field trial, fish were immersed in IHNV subunit vaccine and mortality to virus was reduced from 27% to about 4% (Leong et al. 1992). In order for virus vaccines to provide protection against most serotypes of a specific target virus, it is important to identify different strains of the particular virus (Novoa et al. 1995). Some research has been directed at producing a vaccine for the protozoan I. multifiliis (Goven et al. 1981; Dickerson et al. 1984). However, there are no vaccines available for parasites of fish.

Adjuvants Immunostimulants or adjuvants in fish vaccine preparations have received some attention. These substances include glucans and other yeast extracts, lipopolysaccharides (LPS), extracts of abalone, rough mutants, natural or synthetic oil-based materials, and prevaccination salt baths. Adjuvants are added to oral, injection, or immersion vaccines to enhance immune response, increase longevity, and/or broaden effects of the vaccines, while decreasing pathogen specificity. Anderson (1992) listed 19 different immunostimulants, adjuvants, or vaccine carriers but not all were equally suitable or effective for fish. Press and Lillehaug (1995) identified bovine lactoferrin given orally, or levamisole in an immersion preparation, as showing promise for fish vaccination. Saponin (Quil-A) administered orally or anally increases absorption of human gamma globulin in tilapia.

Methods of vaccine application Fish offer a unique group of animals for vaccination. They can be vaccinated orally, by immersion/dipping in a vaccine or they can be injected with the vaccine (Figure 4.2). Each of these offers advantages and disadvantages and each method are suitable for aquaculture under certain conditions.

Oral Oral vaccination is logistically easy to apply because fish are not confined or handled, therefore eliminating stress. Disadvantages include variable protection, a large quantity of vaccine is required, and the possible need for encapsulation of the antigen. In oral vaccination, the antigen must be incorporated into the feed at manufacturing or top-coated onto the pellets and/or encapsulated. When incorporated into feed the heat to which the antigen is exposed is critical because the antigen may be denatured. However, oral vaccines have very short-term stability once mixed with the feed and in many cases protection is of short duration. Although oral vaccination would appear to be the optimum way to vaccinate fish, there are few such vaccines now available for fish. Oral vaccination may best be suited for secondary or booster vaccinations. The efficacy of orally delivered fish vaccines is hindered by the potential for proteins (antigens) to be denatured in the fish’s acidic stomach before the antigen can be absorbed by the intestine and gain access to immunologically competent tissue. The problem can be circumvented by encapsulating, or coating the antigen; however, this process is expensive. Plumb et al. (1994) demonstrated that protective immunity was obtained in channel catfish when a coated vaccine preparation of E. ictaluri was incorporated into their diet. A titration of the oral vaccine indicated that 0.5, 1.0, and 10% of vaccine (w/w) in the diet for two 5 day treatments with a 5 day window between

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(a)

(b)

(c)

Figure 4.2 Vaccinating fish. (a) Vaccinating trout by intraperitoneal injection. (b) Vaccination by immersion for specified period of time. (c) Vaccination by spraying the vaccine onto fish as they pass through the shower apparatus. Unvaccinated fish netted into top (small arrow) and vaccinated fish coming out of tube (large arrow). (Photographs courtesy of David Powell, ALPHARMA, Bellvue, Washington.)

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significantly increased survival of the vaccinates. Vaccines can also be encapsulated in fish food organisms such as Artemia nauplii, which are then fed to fish (Joosten et al. 1995).

Immersion Immersion vaccination constitutes immersing or dipping fish in a solution of diluted vaccine for seconds to minutes and allowing them to absorb the antigen through the skin, across the gill membrane, and/or by ingestion (Figure 4.2). Immersion is widely used for vaccinating fish and can be easily incorporated into the normal culture routine. Immersion often produces an acceptable level of protection and fish are less stressed than when injected. The skin and gills have specialized cells, such as antibody-secreting cells that are activated when exposed to a specific antigen (pathogen), that will respond by producing antibody when the fish is subsequently exposed to the pathogen. Also, macrophages absorb the vaccine and transport it to the antibody producing organs. Vaccination by dipping is carried out by placing a specified number or weight of fish in a net and dipping them into the vaccine solution for a short period of time—usually 30 seconds. The advantage of dipping is that a solution of vaccine can be used multiple times. When fish are immunized in a diluted vaccine, the fish are usually exposed for a longer period of time. While both the methods provide protection, it is of less duration than the injection vaccination. Spray or shower is a modification of immersion vaccination that provides a high level of protection and can accommodate a higher weight of fish per unit of vaccine volume (Figure 4.2). Disadvantages include a need to handle fish, it is labor intensive, and specialized equipment is required. Semi-automated systems expedite spray vaccination.

Injection Injection is the most common and widely used method of vaccinating fish (Figure 4.2). Injecting a vaccine (intraperitoneally, intramuscularly, or occasionally subcutaneously) provides the highest level of protection and requires relatively small amounts of vaccine. They are economical for use with larger and highly valuable fish, and multivalent vaccine preparations and adjuvants can be conveniently included as needed. Vaccination by injection against multiple disease organisms or different serological strains of the same species simultaneously is economically superior to other procedures and also provides a higher degree of protection (Press and Lillehaug 1995). Other advantages are protection of long duration, assure correct dosage, and incorporation of multiple antigens. To facilitate handling, ideally fish that are to be injected should weigh at least 10 g each. Manual injection vaccination is used extensively using a spring-loaded or air-powered syringe, but this method is more labor intensive, requires each fish to be lightly anesthetized and handled; therefore, there is risk of operator injection. Because of the need to handle each fish, some mortality, particularly of weak fish, does ensue but it is normally only about 0.25%. Evolution of automatic vaccination machines via injection has improved the cost effectiveness of vaccinating salmonids, they do not require handling individual fish, and the danger of self injection is eliminated. These machines require fish to be a more uniform size and generally larger than for hand injection. Other automated vaccination machine advantages include a vaccination rate efficiency of 7,000–9,000 fish per hour compared to about 2,500 per hour (depending on the technician) for hand vaccination. Injection is impractical for devastating diseases of juvenile fish in their first few months of life, and there is also a potential safety hazard to persons administering the vaccine.

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Klesius et al. (2006) describes a vaccination and challenge model using noninvasively fluorescent chromophore calcein marked tilapia to identify S. iniae-vaccinated fish in a cohabitation study. The fish were then challenged by IP injection with virulent S. iniae. In this cohabitation study, not only were the calcein vaccinated fish identified with a portable hand held UV lamp, but vaccinated fish experienced significantly lower mortality than nonvaccinated fish. This is a valuable procedure to identify experimentally vaccinated/challenged fish in cohabitation studies.

Problems When contemplating fish vaccination, several criteria must be considered. Newman (1993) noted 17 factors that influence protective immunity in fish, including vaccine characteristics, biological nature of fish and their nutritional well being, and environmental conditions. Three important aspects of vaccination are age and/or size of fish, effect of temperature on immunity, and level and duration of protection. Most fish have a minimum age or size at which they become immunocompetent. Rainbow trout are immunocompetent at 0.5 g, but when vaccinated at a larger size, they will develop a stronger immunity (Johnson et al. 1982a). Duration of immunity is also dependent upon fish size when vaccinated. Rainbow trout vaccinated at 1 g, will retain immunity for about 120 days, for 180 days when vaccinated at 2 g, and for 1 year if immunized at 4 g (Johnson et al. 1982b). Joosten et al. (1995) showed that efficacy of oral vaccination with encapsulated vaccine preparations can also depend on age of fish when immunized. Vibrio anguillarum vaccine encapsulated into Artemia nauplii was used to orally vaccinate common carp when 15, 29, and 58 days old and gilthead seabream when they were 57 and 69 days old. Carp immunized at 58–69 days developed an insignificantly higher immune response, but seabream

immunized at the same age produced a significantly higher response. Environmental conditions under which fish are vaccinated influence stress factors that adversely affect immune response (Ellis 1981). The immune systems ability to respond to antigens is compromised by poor water quality (low oxygen, high ammonia, etc.), and during any stressful condition, fish produce corticosteroids that suppress immune response (Pickering and Pottinger 1985). Sublethal concentrations of toxicants, particularly phenols, will severely reduce immunity and render aquatic animals more susceptible to infectious disease (Ellis 1988a). High population density will also reduce immune response. Channel catfish vaccinated against E. ictaluri and placed in high-stocking densities had poorer survival than vaccinated fish stocked at lower densities (Plumb et al. 1993). In the United Kingdom, ERM vaccinations of trout failed when environmental conditions were less than favorable or when fish were in poor condition (Rogers 1991). Water temperature is the principal immunomodulator in fish (Avtalion 1981; Johnson et al. 1982b; Bly and Clem 1991a). Temperature does not determine if an immune response occurs, but it regulates the rapidity and degree to which it develops. The optimum temperature for immunization corresponds to the optimum temperature in which the fish normally lives. Generally, salmonids respond better when vaccinated at 15–18◦ C, while warmwater fish respond better when vaccinated above 20◦ C. Plumb et al. (1986) showed that channel catfish immunized and held at 12–15◦ C developed a slow but longer lasting humoral antibody response than did fish immunized and maintained at 28◦ C. However, when warmwater fish are vaccinated and held at an optimum temperature for a few days to 2 weeks before temperature is reduced, the immune system is primed and immunological memory may occur, which will later become activated. Although Bly and Clem (1991b) demonstrated temperature-dependent

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immunity, they proposed that channel catfish may be immunocompromized during winter. It appears that immunization at low temperatures may actually suppress immune response, but a secondary response may occur when fish are again exposed to that antigen when water temperatures warm. It should be noted that low water temperatures suppress T-dependent lymphocyte antibody functions in channel catfish and possibly other fishes as well (Bly et al. 1986). Seasonal modulation of immune response may also occur in salmonids (Zeeman 1986). Rainbow trout vaccinated and held at 18◦ C prior to winter, when water temperatures would be declining, produced lower antibody titers than fish vaccinated at the same temperature just prior to spring, when water temperatures would be rising. In the last two decades, vaccination has become an integral part of salmonid, warm water and marine aquaculture. When taking into consideration the current attitude toward drugs, time, expense, and legal questions involved in new drug approval, low drug approval rate, and current effort being expended in vaccine research and development, one can be assured that vaccination will become an even more significant and routine practice in aquaculture fish health management.

References Acheson, D. 2007. Testimony before Subcommittee on Agriculture, Rural Development, Food and Drug Administration, U.S. House of Representatives, Washington, D.C. Anderson, D. P. 1992. Immunostimulants, adjuvants, and vaccine carriers in fish: applications to aquaculture. Annual Reviews of Fish Diseases 2:281–307. Anonymous. 2006. Effects and benefits of fish vaccination. Intervet. http://www.thefishsite.com/ articles. Anonymous. 2007. Minor use guidance document: a guide to the approval of animal drugs for minor uses and for minor species. FDA Center for Veterinary Medicine, Food and Drug Administration, Rockville, Maryland, USA.

Anonymous. 2008. Legal and judicious use of drugs and therapeutants in aquaculture. Coldwater Fish Culture Course, Bozeman, Montana, March 18, 2008. http://www.fws.gov/fisheries/ aadap/PDF/Fish/20He. Arkoosh, M. R., E. Clemons, et al. 2001. Increased susceptibility of juvenile chinook salmon to Vibriosis after exposure to chlorinated and aromatic compounds found in contaminated urban estuaries. Journal of Aquatic Animal Health 13:257–268. Arthur, J. R., Cr. Lavilla-Pitogo, and R. P. Subasinghe. 1996. Use of chemicals in aquaculture in Asia. Proceedings of the meeting on the use of chemicals in aquaculture in Asia. Tigbauan, Iloilo, Philippines. http://www.seafdec.org.ph/. Avtalion, R. R. 1981. Environmental control of the immune response in fish. CRC Critical Reviews in Environmental Contamination 11:163–188. Bennett, A. J., P. W. Whitby, and G. Coleman. 1992. Retention of antigenicity by a fragment of Aeromonas salmonicida 70 kDa serine protease which includes the primary substrate binding site expressed as ß-galactosidase hybrid proteins. Journal of Fish Diseases 15:473–484. Bly, J. E., T. M. Buttke, et al.. 1986. The effects of in vivo acclimation temperature on the fatty acid composition of channel catfish (Ictalurus punctatus) peripheral blood cells. Comparative Biochemistry and Physiology 83:791–795. Bly, J. E., and L. W. Clem. 1991a. Temperaturemediated processes in teleost immunity: in vitro immunosupression induced by in vivo low temperature in channel catfish. Veterinary Immunology and Immunopathology 28:365–377. Bly, J. E., and L. W. Clem. 1991b. Temperaturemediated processes in teleost immunity: in vivo low temperature immunisation does not induce tolerance in channel catfish. Fish & Shellfish Immunology 1991:229–231. Bowker, J. 2006. Legal use of drugs and therapeutants in aquaculture. Southern DivisionAFS Aquatic Animal Drug Approval Workshop. San Antonio, TX. http://www.fws.gov/fisheries/ aadp//pdf/legal%20us. Accessed 03/15/09. Boyd, C. E. 1990. Water Quality in Ponds for Aquaculture. Alabama Agricultural Experiment Station, Auburn University, Auburn, Alabama. Boyd, C. E., P. Munsiri, and B. J. Hajek. 1994. Composition of Sediment from intensive shrimp ponds in Thailand. World Aquaculture 25:53–55.

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Boyd, C. E., and S. Pippopinyo. 1994. Factors affecting respiration in dry pond bottom soils. Aquaculture 120:283–293. Boyd, C. E., R. P. Romaire, and E. Johnston. 1978. Predicting early morning dissolved oxygen concentrations in channel catfish ponds. Transactions of the American Fisheries Society 107:484–492. Boyd, C. E. and C. S. Tucker 1998. Pond Aquaculture Water Quality Management. Boston, Kluwer Academic. Boyd, C. E., and B. J. Wattson. 1989. Aeration systems in Aquaculture. CRC Critical Reviews in Aquatic Sciences 1:425–472. Brunt, J., A. Newaj-Fyzul, and B. Austin. 2007. The development of probiotcs for control of multiple bacterial diseases of rainbow trout, Oncorhynchus mykiss (Walbaum). Journal of Fish Diseases 30:573–579. Chaslus-Dancla, E. 1992. Les probleme d’antibioresistancechez Lesespeces animales autres que les poissons. In: C. Michel and D. J. Alderman (eds) Chemotherapy in Aquaculture. Paris, Office International des Epizooties, pp. 243–253. Cooper, R. K., II, E. B. Shotts, Jr., and L. K. Nolan. 1996. Use of a mini-transposon to study chondroitinase activity associatedwith Edwardsiella ictaluri. Journal of Aquatic Animal Health 8:319–324. Darwish, A. M., B. G. Griffin, et al. 2002. Histological and hematological evaluation of potassium permanganate exposure in channel catfish. Journal of Aquatic Animal Health 14:134–144. Dickerson, H. W., J. Brown, et al. 1984. Tetrahymena pyriformes as a protective antigen against Ichthyophthirius multifiliis infections: comparison between isolates and ciliary preparations. Journal of Fish Biology 24:423–528. Dixon, B. A. 1994. Antibiotic resistance of bacterial fish pathogens. Journal of the World Aquaculture Society 25:60–63. Doulos, S. K., A. J. Garland, et al. 1994. Comparison of two methods of oxygen supplementation for enhancing water quality in fish culture. The Progressive Fish Culturist 56:130–134. Duff, D. C. B. 1942. The oral immunization of trout against Bacterium salmonicida. Journal of Immunology 44:87–94. Ellis, A. E. 1981. Stress and the modulation of defense mechanisms in fish. In: A. D. Pickering

(ed.) Stress and Fish. Academic Press, pp. 147– 169. Ellis, A. E. 1988a. Fish Vaccination. London, Academic Press. Ellis, A. E. 1988b. Current aspects of fish vaccination. Diseases of Aquatic Organisms 4:159–164. Fortt, A. Z. and B. R. Buschmann. 2007. Residues of tetracycline and quinolones in wild fish living around a salmon aquaculture center in Chile. Revista Chilena de Infectologia 24:8–12. Goven, B. A., D. L. Dawe, and J. B. Gratzek. 1981. Protection of channel catfish (Ictalurus punctatus) against Ichthyophthirius multifiliis (Fouget) by immunization with varying doses of Tetrahymena pyriformes (Lwoff) cilia. Aquaculture 23:269–273. Groman, D. B., and B. W. Klontz. 1983. Chemotherapy and prophylaxis of bacterial kidney disease with erythromycin. Journal of the World Mariculture Society 14:226–235. Herwig, N. 1979. Handbook of Drugs and Chemicals Used in the Treatment of Fish Diseases. Springfield, Charles C. Thomas. Irianto, A., and B. Austin. 2002. Probiotics in aquaculture. Journal of Fish Disases 25:633– 642. Johnson, B. 2008. Legal use of drugs and therapeutants I aquaculture. Alaska Hatchery Managers Meeting, January 15–17, 2008. Seward, AK. hhtp://NNW./gov/fisheries/aadap/home.htm. Johnson, K. A., J. K. Flynn, and D. F. Amend. 1982a. Onset of immunity in salmonid fry vaccinated by direct immersion in Vibrio anguillarum and Yersinia ruckeri bacterins. Journal of Fish Diseases 5:197–205. Johnson, K. A., J. K. Flynn, and D. F. Amend. 1982b. Duration of immunity in salmonids vaccinated by direct immersion in Yersinia ruckeri and Vibrio anguillarum. Journal of Fish Diseases 5:207–213. Johnston, P., and D. Santillo. 2002. Chemical usage in aquaculture: implications for residues in market products. Greenpeace Research Laboratories, Department of Biological Sciences, University of Exeter EX4 4PS, United Kingdom. Joosten, P. H. M., M. Avil´es-Trigueros, et al. 1995. Oral vaccination of juvenile carp (Cyprinus carpio) and gilthead seabream (Sparus aurata) with bioencapsulated Vibrio anguillarum bacterin. Fish & Shellfish Immunology 5:289–299. Klesius, P., J. Evans, and C. Shoemaker. 2000. Communicable bacterial diseases in fish farming:

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prevention by vaccination and immunostimulation. Proceedings Pan American Congress of Veterinarian Science. http://www.ars.usda.gov/ research/publications.htm?seq no 115=162981. Accessed 03/26/2009. Klesius, P. H., J. J. Evans, et al. 2006. A vaccination and challenge model using calcein marked fish. Fish & Shellfish Immunology 20:20–28. Lai-Fa, Z., and C. E. Boyd. 1988. Nightly aeration to increase the efficiency of channel catfish production. The Progressive Fish-Culturist 50:237–242. Leong, J. C., E. D. Anderson, et al. 1992. Biotechnological approaches to the development of salmonid fish vaccines. In: T. Kimura (ed.) Proceedings of the OJI International Symposium on Salmonid Diseases. Sapporo, Hokkjaido University Press, pp. 250–255. Leong, J. C., and J. L. Fryer. 1993. Viral vaccines for aquaculture. Annual Reviews of Fish Diseases 3:225–240. Lorio, W. J. 1994. Production of channel catfish in ponds with water recirculation. The Progressive Fish-Culturist 56:202–206. Lovell, T. 1989. Nutrition and Feeding of Fish. New York, Van Nostrand Reinhold. MacMillan, R. 2003. Drugs used in the U.S. aquaculture industry. Information available on world wide web, accessed 02/01/2009. Moffitt, C. M. 1992. Survival of juvenile chinook salmon challenged with Renibacterium salmoninarum and administered oral doses of erythromycin thiocyanate for different durations. Journal of Aquatic Animal Health 4:119–125. Newman, S. G. 1993. Bacterial vaccines for fish. Annual Reviews of Fish Diseases 3:145–185. Novoa, B., S. Blake, et al. 1995. Comparison of different procedures for serotyping aquatic birnavirus. Applied and Environmental Microbiology 61:1925–1929. Okamoto, A. 1992. Restrictions on the use of drugs in aquaculture in Japan. In: C. Michel and D. J. Alderman (eds) Chemotherapy I Aquaculture: From Theory to Reality. Paris, Office International de Epizooties, pp. 109–114. Ogut, H., S. E. LaPatra, and P. W. Reno. 2005. Effects of host density on furunculosis epidemics determined by the simple SIR model. Preventive Veterinary Medicine 71:83–90. Pickering, A. D., and T. G. Pottinger. 1985. Cortisol can increase the susceptibility of brown trout

Salmo trutta L., to disease without reducing the white blood cell count. Journal of Fish Biology 27:611–619. Piedrahita, R. L. 1991. Simulation of short-term management actions to prevent oxygen depletion in ponds. Journal of the World Aquaculture Society 22(3):157–166. Plumb, J. A., C. C. Sheifinger, et al. 1995. Susceptibility of six bacterial pathogens of channel catfish to six antibiotics. Journal of Aquatic Animal Health 7:211–217. Plumb, J. A., S. Vinitnantharat, et al. 1993. Densitydependent effect on oral vaccination of channel catfish against Edwardsiella ictaluri. Aquaculture 122:91–96. Plumb, J. A., S. Vinitnantharat, and W. D. Paterson. 1994. Optimum concentration of Edwardsiella ictaluri vaccine in feed for oral vaccination of channel catfish. Journal of Aquatic Animal Health 6:118–121. Plumb, J. A., M. L. Wise, and W. A. Rogers. 1986. Modulary effects of temperature on antibody response and specific resistance to challenge of channel catfish, Ictalurus punctatus, immunized against Edwardsiella ictaluri. Veterinary Immunology and Immunopathology 12:297–304. Press, C. M., and A. Lillehaug. 1995. Vaccination in European salmonid aquaculture: a review of practices and prospects. British Veterinary Journal 151:45–69. Rogers, C. J. 1991. The use of vaccination and antimicrobial agents for control of Yersinia ruckeri. Journal of Fish Diseases 14:291–301. Rogers, W. A., and D. Xu. 1992. Protective immunity induced by a commercial Vibrio vaccine in hybrid striped bass. Journal of Aquatic Animal Health 4:303–305. Schnick, R. A. 1999. Use of chemicals in fish management and fish culture: past and future. In: D. J. Smith, W. H. Gingerich, and M. G. BeconiBarker (eds) Xenobiotics in Fish. New York, Kluwer Academic/Plenum Publishers, pp. 1–14. Schnick, R. A., D. J. Alderman, et al. 1997. Worldwide Aquaculture drugs and vaccine registration progress. http://www.aquanic.org/awuadrugs/ publications/world drug progress 9-20-99.htm. Accessed 3/10/2009. Shoemaker, C. A., and P. H. Klesius. 1997. Protective immunity against enteric septicaemia in channel catfish. Ictalurus punctatus (Rafinesque), following controlled exposure to

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Edwardsiella ictaluri. Journal of Fish Diseases 20:101–108. Shoemaker, C. A., P. H. Klesius, and J. A. Plumb. 1997. Killing of Edwardsiella ictaluri by macrophages from channel catfish immune and susceptible to enteric septicemia of catfish. Veterinary Immunology and Immunopathology 58:181–190. Stoffregen, D. A., P. R. Bowser, and J. G. Babish. 1996. Antibacterial chemotherapeutants for finfish aquaculture: a synopsis of laboratory and field efficacy and safety studies. Journal of Aquatic Animal Health 8:181–207. Straus, D. L., and C. S. Tucker. 1993. Acute toxicity of copper sulfate and chelated copper to channel catfish Ictalurus punctatus. Journal World Aquaculture Society 24:390–395. Thiery, M., F. Lecocq-Xhonneux, et al. 1990. Molecular cloning of the mRNA coding for the G protein of the viral haemorrhagic septicaemia (VHS) of salmonids. Veterinary Microbiology 23:221–226. Tort, M. J., D. J. Pasnik, et al. 2002. Quantitative scoring of gill pathology of walleyes exposed to hydrogen peroxide. Journal of Aquatic Animal Health 14:154–159. Tucker, C. S., and C. E. Boyd. 1977. Relationships between potassium permanganate treatment and water quality. Transactions of the American Fisheries Society 106:481–488. Tucker, C. S., J. A. Steeby, et al. 1992. Production characteristics and economic performance for four channel catfish, Ictalurus punctatus, pond

stocking density-cropping system combinations. In: D. Tave and C. S. Tucker (eds) Recent Developments in Catfish Aquaculture. Binghamton, New York, Food Products Press, pp. 333–352. Vallejo, A. N., N. W. Miller, and L. W. Clem. 1992. Antigen processing and presentation in teleost immune response. Annual Review of Fish Diseases 2:73–90. Vaughan, L. M., P. R. Smith, and T. J. Foster. 1993. An aromatic-dependent mutant of the fish pathogen Aeromonas salmonicida is attenuated in fish and is effective as a live vaccine against the salmonid disease furunculosis. Infection and Immunity 61:2172–2181. Wedemeyer, J. W. 1996. Physiology of Fish in Intensive Culture Systems. New York, Chapman & Hall. Wellborn, T. L., Jr. 1985. Control and therapy. In: J. A. Plumb (ed.) Principal Diseases of Farm Raised Catfish. Alabama Agricultural Experiment Station, Auburn University, Alabama, Southeastern Cooperative Series Bulletin No. 225, pp. 50–67. Xu, L., D. V. Mourich, et al. 1991. Epitope mapping and characterization of the infectious hematopoietic necrosis virus glycoprotein, using fusion proteins synthesized in Escherichia coli. Journal of Virology 65:1611–1615. Young, K. 2002. Aquaculture: A need for import tolerances. http://www.fda.gov/cvm/index/ vmac/Young. Zeeman, M. G. 1986. Modulation of the immune response in fish. Veterinary Immunology and Immunopathology 12:235–241.

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Part II

Viral Diseases

Compared with research history of bacterial and parasitic diseases of aquatic animals, the study of fish viral diseases is relatively new. Infectious pancreatic necrosis virus (IPNV) of salmonids was the first proven viral fish disease, even though some might argue that infectious hematopoietic necrosis virus (IHNV) was demonstrated earlier. Infectious pancreatic necrosis virus was described by Wood et al. (1955) and viral etiology demonstrated by Wolf et al. (1959, 1960). The first major literary compilation on fish viral diseases was Viral Diseases of Poikilothermic Vertebrates, published by the New York Academy of Sciences (Whipple 1965) and included contributions by fisheries scientists from the United States, Europe, and Asia. The most extensive work published on fish viruses to date is Fish Viruses and Fish Viral Diseases (Wolf 1988); a detailed scholarly documentation of all fish viruses and viral diseases known at that time. The study of fish virology has grown rapidly since its inception. McAllister (1979) discussed about 30 virus diseases that affect fish. Wolf (1988) listed 59 fish diseases that were in some way associated with viral agents, 35 of which had been isolated in tissue culture. In 1993 the number of fish virus diseases was updated to approximately 95 (Hetrick and Hedrick 1993). However, Ahne (1994) indicated that only about 60 different viruses were known

(isolated?) from finfish in the first edition of this book (Plumb 1999) and about 40 virus diseases, mostly of cultured fish, were discussed. Increased fish virus detection may be attributed to more extensive surveillance, a greater number of fish health oriented laboratories that have virus detection and research capabilities, and the fact that intensive aquaculture is conducive to expression of viral infections and improved and more sensitive virus detection methods and reagents have been developed. Research of viral fish diseases remains a high priority because of their potential to cause catastrophic losses and the tumor producing ability of some of these agents. New fish viruses are being discovered every year. Although comparatively few fish viruses cause severe disease in aquaculture, when they do occur results can be devastating. Most known fish viruses have been reported in freshwater cultured species of high economic value, some occur in marine fish only, and others are found in both environments. Until fish are exposed to environmental stressors, the devastating potential of some viruses may not be realized; therefore, stressors often stimulate discovery and identification of previously unknown viruses that affect wild freshwater and/or marine fish populations. The emphasis of this book is on viral and bacterial diseases of cultured fishes, but it is 91

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becoming increasingly clear that viruses are more wide spread in natural fish populations than was once suspected. Evidence also indicates that some viruses are transmitted directly from wild populations to cultured fishes and vise versa. An important virus characteristic is the fact that it must invade a living cell in order to replicate. Viruses cannot reproduce themselves, but must rely on synthetic cell machinery to produce a new virus. A virus may manifest itself in several ways, the most drastic being production of hemorrhagic inflammation and primary necrosis, which often results in high mortality particularly of juvenile fishes. Tumors grow on the skin and fins and can be caused by a virus infection, or there may be total absence of pathology. Viruses vary in virulence; several are capable of killing a high percentage of infected fish (i.e., IPNV of trout and CCV of channel catfish), some have relatively low virulence (i.e., golden shiner virus), while others are avirulent (i.e., catfish reovirus). Also, virulent viruses can be carried for extended periods of time by healthy fish without the host showing any overt signs of infection. Viral infections are confirmed by their isolation in cell culture, producing an identical disease in susceptible animals by injection of cellfree filtrates from diseased animals, or electron microscopically observing the virus in tissues. The first isolated fish virus in cell culture was reported by Wolf and Quimby (1962). This pioneering technology led to major advances in the field of fish virology during the last three decades. Once a virus can be cultured in cell lines, it is amenable to detailed molecular studies, vaccine research, and pathogenesis research. Other research tools such as serological, radiological, polymerase chain reaction, and other technological procedures are used in fish virus detection. Viruses often affect in vivo cell cultures in a particular way. Cell injury (cytopathic effect—CPE) can be detected with relatively low magnification light microscopy. The CPE produced by a virus in cell culture is commonly

characteristic of a particular type or group of viruses but is seldom specific enough for definitive identification. Multiple, interrelated factors determine to what extent a virus will affect a fish population; however, water temperature is among the most important factors. Typically, fish viruses have a temperature range in which they are most pathogenic. Channel catfish virus is a good example; usually severe mortality occurs in young-of-the-year fish when water temperatures are above 25◦ C, moderate mortality occurs at 21–24◦ C, and little, if any, mortality occurs below 18◦ C. Age also affects a fish’s susceptibility to virus infection with younger fish generally suffering greatest mortality. Although severe mortality is rare among fish older than 1 year, they often become carriers and serve as virus reservoirs for as long as they remain alive. Stressful effects of transport, handling, poor water quality, high stocking densities, and inadequate nutrition can also affect severity of a virus infection. Viral agents with the greatest potential to cause disease in cultured fish are emphasized in this text and are organized according to the primary family or group of fish they infect. Only minor attention is given to nonvirulent viruses or those that have been detected only by electron microscopy. No attempt has been made to include all viruses that affect fish.

References Ahne, W. 1994. Viral infections of aquatic animals with special reference to Asian aquaculture. Annual Review of Fish Diseases 4:357–388. Hetrick, F. M. and R. P. Hedrick. 1993. New viruses described in finfish from 1988–1992. Annual Review of Fish Diseases 3:187–207. McAllister, P. E. 1979. Fish viruses and viral infections. In: H. Fraenkel-Conrat and R. R. Wagnor (eds) Comprehensive Virology. Plenum Press, New York, pp. 401–470. Plumb, J. A. 1999. Health Maintenance and Principal Microbial Diseases of Cultured Fishes. First edition. Iowa State University Press, Ames, IA.

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Whipple, H. E. 1965. Viral diseases of poikilothermic vertebrates. Annals of the New York Academy of Science 126:1– 608. Wolf, K. 1988. Fish Viruses and Fish Viral Diseases. Cornell University Press, Ithaca, NY. Wolf, K., C. E. Dunbar, and S. F. Snieszko. 1960. Infectious pancreatic necrosis of trout, a tissue-culture study. Progressive Fish Culturist 22:64–68.

Wolf, K, and M. C. Quimby. 1962. Established erythermic line of fish cells in vitro. Science 135:1065. Wolf, K., S. F. Snieszko, and C. E. Dunbar. 1959. Infectious pancreatic necrosis, a virus-caused disease of fish. Excerptas Medica 13:228. Wood, E. M., S. F. Snieszko, and W. T. Yasutake. 1955. Infectious pancreatic necrosis in trout. American Medical Association Archives in Pathology 60:26–28.

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Chapter 5

Catfish viruses

The most serious virus disease of cultured catfishes is channel catfish virus disease (CCVD). There is also a catfish reovirus (CRV) isolated from channel catfish, a herpesvirus isolated from black bullhead, and an iridovirus have been found in black bullhead and European catfish (sheatfish).

Channel catfish virus disease CCVD, an acute, highly communicable infection of cultured juvenile channel catfish was discovered in 1968 (Fijan et al. 1970). The etiological agent of CCVD is channel catfish virus (CCV), a member of the family Herpesviridae with Herpesvirus ictaluri suggested as the specific epithet (Wolf and Darlington 1971). The virus was subsequently renamed Ictalurid herpesvirus 1 (IcHV-1)to follow International Committee on Taxonomy of Viruses (ICTV) conventions for herpesviruses, but it is most commonly known by its original name. The second herpesvirus from black bullhead was designated I. melas herpesvirus (IcmHV) subsequently Ictalurid herpesvirus 2 and most recently the name Ameiurine herpesvirus 1 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.

(AmHV-1) have been proposed (Hedrick et al. 2003; Droszpoly et al. 2008).

Geographical range and species susceptibility CCV has been isolated from channel catfish in most southern states and several other areas of the United States. The virus was also isolated from a group of channel catfish fry shipped from the United States to Honduras in 1972 (J. A. Plumb, unpublished). These fish were destroyed and the virus apparently failed to become established in that country. There have also been undocumented reports of a disease resembling CCVD in other countries where channel catfish have been introduced. Channel catfish appear to be the most susceptible catfish species to CCV. Experimental infections have been induced in fingerling blue catfish and channel catfish X blue catfish hybrids by virus injection, but no infection occurred when CCV was introduced orally or by cohabitation of naive nonchannel catfish ictalurids with virus-infected channel catfish (Plumb and Chappel 1978). Contrary to being experimentally refractive to CCV, the virus was isolated from diseased fingerling blue catfish in Missouri in 1988 (Anonymous 1988) and Mississippi (Hanson et al. 2006). Brown 95

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and yellow bullheads cannot be experimentally infected with the virus by injection or feeding. European catfish are also resistant to the virus (Plumb et al. 1985). When CCV was fed to different strains of channel catfish fry, a variation in susceptibility was noted among strains of the species (Plumb et al. 1975). Young fish that were the product of cross breeding between different channel catfish strains were more resistant to CCV than were inbred parental strains that would suggest a strain hybrid vigor.

Clinical signs Clinical signs of CCV vary with some or all of the following being present (Figure 5.1): distension of the abdomen, exophthalmia, pale or hemorrhagic gills, and hemorrhage at the base of fins and throughout the skin on the ventral surface (Fijan et al. 1970). Infected fish swim erratically, sometimes rotating about their longitudinal axis. Moribund fish hang head up at the surface or sink to the bottom, become

quiescent, and respire weakly but rapidly just prior to death. Internal signs include presence of a clear straw-colored fluid in the peritoneal cavity and a general hyperemia throughout the visceral cavity although the liver and kidneys can be pale. The spleen is generally dark red and enlarged, and the stomach and intestine are void of food but contain a mucoid secretion.

Diagnosis When a sudden increase in morbidity occurs among young channel catfish during the summer, CCV should be suspected and fish necropsied for virus. Virus is detected by inoculating filtrates made from whole fry or visceral tissue into appropriate cell cultures. The cell line of choice is channel catfish ovary (CCO) because of its high sensitivity to CCV and viral productivity (Bowser and Plumb 1980). Intranuclear CCV replication occurs in CCO cells at 15–35◦ C with 30–35◦ C being optimum. Cytopathic effect (CPE) may be visible in 12–24 hours

Figure 5.1 A 10-cm channel catfish naturally infected with channel catfish virus. Note the abdominal distension and exophthalmia. The upper lobe of the caudal fin (arrow) has a columnaris lesion.

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at 30◦ C if an active CCV infection is in progress. Initial CPE is indicated by foci of pyknotic cells, which coalesce into multinucleated syncytia that connect to surrounding normal cells by strands of protoplasm resembling irregular wheel spokes (Figure 5.2). As CPE progresses, cells separate from the culture vessel surface and form a loose network of Syncytia in susceptible cell cultures, which is presumptive identification of CCV; positive identification is made by serum neutralization using CCV-specific antiserum, fluorescent antibody staining, or PCR. Gray et al. (1999b) utilized PCR to identify CCV DNA in acutely infected channel catfish; the method proved to be rapid, sensitive, and specific. Virus was detected in blood, intestines, and liver. CCV was also detected by PCR in channel catfish fingerlings that had survived infection 140 days postimmersion infection (Gray et al. 1999a). Virus DNA was detected in blood, brain, intestines, kidney, and peripheral blood leukocytes of these latently infected fish, thus confirming that CCV establishes a latent infection in channel catfish. Subsequently, Stingley et al. (2003), using reverse transcriptase PCR, showed that 24 days postinfection, CCV was not replicating in tissues in latently infected channel catfish. Ictalurus melas herpesvirus from black bullhead replicates in CCO cells where it produces similar CPE to that of CCV. However, IcmHV does not react to CCV-specific polyclonal or monoclonal antibody to CCV in neutralization or to indirect immunofluorescence assays (Hedrick et al. 2003). Brown bullhead (BB) cells and cell cultures derived from walking catfish kidney (K1K) are also susceptible to CCV, but are not as sensitive as CCO cells (Bowser and Plumb 1980; Noga and Hartman 1981). Cell lines developed from Japanese striped knife jaw and amberjack and the hybrid of kelp and spotted grouper showed CPE when infected with CCV. Other poikilothermic and mammalian cell culture systems are refractory to the virus (Fernandez et al. 1993).

Virus characteristics CCV has been recognized as a herpesvirus since its characterization. On the basis of its rapid replication and destruction of tissue culture cells, it was categorized as an alphaherpesvirus (Roizman 1990). However, nucleotide sequence comparisons demonstrate very little relationship to herpesviruses of birds, reptiles, and mammals; therefore, CCV has been placed in a separate group of herpesviruses (Davison 1992). Recent molecular data on other fish and amphibian herpesviruses suggests these viruses are more closely related to each other than to herpesviruses of homeotherms and reptiles. A new group of herpesviruses, the Ictalurivirus genus, has been designated by the ICTV with CCV being the type species. Biological characteristics and molecular analyses indicate that CCV functions much like other herpesvirus. Hanson and Thune (1993) demonstrated that CCV encoded a thymidine kinase, the genome was shown to be infectious, allowing functional gene evaluation by marker rescue (Hanson et al. 1994) and by using homologous recombination to produce gene knock-outs (Zhang and Hanson 1995, 1996). Booy et al. (1996), using cryoelectron microscopy and three-dimensional image reconstruction, demonstrated that the capsid structure of CCV was very similar to herpesviruses of mammals even though the proteins had very little sequence homology. Gene expression studies by Silverstein et al. (1998), Huang and Hanson (1998), and Stingley and Gray (2000) demonstrated that CCV gene expression was coordinately regulated and sequentially ordered such that the genes can be classified into three broad temporal groups: immediate-early, early, and late genes. The recent establishment of bacterial artificial chromosomes containing the CCV genome allows researchers to manipulate the CCV sequence using powerful tools developed for prokaryotes (Kunec et al. 2008). This is an exciting development for studying the

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(a)

(b)

(c)

Figure 5.2 Channel catfish ovary cells infected with channel catfish virus and incubated at 30◦ C. (a) Focal cytopathic effect (CPE) 19 hours after inoculation surrounded by normal cells. (b) Total CPE 24 hours after inoculation. (c) Channel catfish virus (arrow) in the nucleus of a spleen cell from experimentally infected channel catfish. Note the hyaline-like inclusion bodies (I). (From Plumb et al. (1974); reprinted with permission of Blackwell Scientific Publications Ltd.)

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molecular basis of the pathogenesis of CCV because it allows production and testing of sophisticated recombinants relatively easily. CCV is enveloped and icosahedral with a DNA genome and nucleocapsid diameter of 95–105 nm. Enveloped virions have a diameter of 175–200 nm. Infectivity is inactivated by either 20% or 5% chloroform. CCV is heatlabile at 60◦ C for 1 hour, unstable in seawater, and sensitive to ultraviolet light, requiring 20–40 minutes for inactivation (Robin and Rodrique 1980). CCV survives for less than 24 hours on dried fish netting or glass cover slips; it retains infectivity in pond water for about 2 days at 25◦ C, and for 28 days at 4◦ C. In dechlorinated tap water, infectivity is retained for 11 days at 25◦ C and for over 2 months at 4◦ C. Under experimental conditions, CCV infectivity is almost immediately neutralized when introduced into pond mud. Infectious CCV cannot be isolated from decomposing infected fish at 22◦ C 48 hours after death but virus is recoverable for up to 14 days from viscera of whole iced fish, for 162 days from fish frozen at −20◦ C, and for 210 days from fish frozen at −80◦ C (Plumb et al. 1973).

Epizootiology Outbreaks of CCVD in channel catfish have been diagnosed during May through October when water temperatures are above 25◦ C. In Alabama, 94% of 53 confirmed CCVD cases reported between 1991 and 1996 occurred during June through September, while none were diagnosed from November through April. Epizootics on catfish farms occur most frequently during years of high water temperature coupled with heavily stocked fingerling ponds. Many outbreaks are preceded by handling and transport of fingerlings. As long as active infection is present, fry and fingerling channel catfish transmit CCV horizontally by contact and through the water. Under experimental conditions, it is also pos-

sible to transmit virus from infected moribund or dead fish to healthy fish through water. Other CCV transmission modes are intramuscular or intraperitoneal injection, incorporation of virus into feed, or swabbing gills with a saline solution containing virus. Infectivity can be experimentally induced in fish weighing up to 10 g by waterborne exposure, but injection of CCV is required to infect and cause overt disease in larger fish. Occasionally, 1-year-old fish will suffer a CCVD outbreak that is generally associated with low mortality and usually follows stressful transport or handling. Often CCVD occurs in conjunction with a secondary Flavobacterium columnare infection that prolongs mortality. Incubation time between fish exposure to virus and appearance of clinical signs and morbidity is related to water temperature (Plumb 1972). Experimental infection at 30◦ C was followed by clinical signs in 32–42 hours with death several hours later (Plumb and Gaines 1975). At 20◦ C incubation was 10 days. Under field conditions at 25–30◦ C, healthy channel catfish fingerlings develop disease within 72–78 hours postexposure, and up to 100% may die within 6 days. In a documented case of CCVD, a group of fry held at 28◦ C in troughs receiving well water developed clinical CCVD when 21 days old and 72 hours later 100% were dead. These fish had been negative for CCV in cell culture when assayed 7 days prior to disease outbreak. Disease developed rapidly even though infected fish were not exposed to a known source of virus between day 14 and 21; therefore, it must be assumed that the fry were infected by brood stock via egg production. Young-of-the-year fish less than 4 months old are most susceptible to CCV and case data suggest that the younger and/or smaller the fish, the higher the mortality. However, experimental data using immersion challenges demonstrate that very young fry (1–2 weeks of age) are more resistant to CCV and they become more susceptible as they age (Hanson et al. 2004). This phenomenon was correlated with the CCV neutralizing antibody levels of

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the female parent suggesting potential maternal transfer. CCV is communicable from the time clinical signs appear until soon after death. In experimentally infected juveniles, virus began to decline in surviving fish 120 hours postinfection (Plumb 1971). Virus was not isolated from surviving fish by cell culture once clinical signs disappear. This observation was substantiated by the PCR research by Gray et al. (1999b), who did not detect newly replicated virus in the latent stage following clinical disease. Kancharla and Hanson (1996) found that CCV was shed by infected juveniles and could be detected in water for up to 12 days postinfection. Virus was also found in internal organs, skin, and gills. Fish infected with thymidine kinase (TK)-negative CCV shed less virus than do fish infected with unaltered virus. Recombinant TK-negative CCV replicates in cell cultures; however, 100 times more mutant virus is required to kill fish than the wild-type virus (Zhang and Hanson 1995). Adult fish are considered likely sources of CCV by vertical transmission. CCV genomic DNA has been found in brood fish and in the young offspring of the positive broodstock, while Gray et al. (1999b) did so in postinfection juveniles using PCR. Although active CCV infections are relatively easy to diagnose by virus isolation in cell culture, detection of covert CCV in a carrier population is difficult. Plumb (1973) detected CCV antibodies in channel catfish that had been exposed to virus and suggested that this may be a method for separating adult fish that had been exposed to CCV from those that had not. Amend and McDowell (1984) used serum neutralization tests on adult catfish to separate possible carrier from noncarrier populations. They later demonstrated that 7 of 17 major brood populations tested showed positive CCV serum neutralization. Virus was subsequently isolated from young-of-the-year fish from 2 of the 7 populations in which CCV antibody positive fish were detected. CCV neutralizing antibodies were found in adult channel catfish for 2

years following last known exposure to virus (Hedrick et al. 1987). In this same study, adults not exposed to CCV as juveniles were experimentally infected and one died apparently as a result of the virus. Surviving adults produced CCV antibody for 6 months although titers were low. It was suggested that CCV does become latent in epizootic survivors and that expression of certain viral antigens, or periodic virus reactivation, may be a stimulus for continued anti-CCV antibody production. Studies strongly support the idea that adult fish carry CCV and can possibly develop active infection. Plumb et al. (1981) demonstrated that CCV antigens could be detected in gonads of adult fish by immunofluorescence. Bowser et al. (1985) isolated CCV from adult channel catfish in winter when water temperatures were 6–8◦ C and later from individuals of the same population when fish were stressed by steroid injection. Wise et al. (1985) utilized a CCV-specific DNA probe to clearly demonstrate the presence of CCV nucleic acid in various tissues of adult channel catfish, including adults from which virus had been isolated. It was further shown by a CCV-nucleic acid probe that CCV genetic material was present in adults as well as their offspring, indicating vertical transmission (Wise et al. 1988). Bird et al. (1988) used molecular cloning of CCV genome fragments to suggest that CCV can persist in channel catfish in a dormant or transcriptional active state without causing clinical signs. Polymerase chain reaction was used by Boyle and Blackwell (1991) to detect latent CCV, and Baek and Boyle (1996) refined the procedure with nested primers that allowed detection of CCV DNA in the blood of brood stock. Latent CCV is apparently prevalent in the commercial catfish industry with a relatively low number of overt CCVD out breaks. Thompson et al. (2005), using PCR, demonstrate a carrier state in fry populations from all seven of evaluated commercial channel catfish hatcheries and showed an increase in prevalence of latent CCV in three fingerling ponds that were monitored through the first growing

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season. Throughout this study, none of the populations broke with CCVD.

Pathology A generalized viremia is established within 24 hours post-experimental infection. The kidneys, liver, spleen, and intestines become active sites of virus replication 24–48 hours after infection and virus may be isolated from brain tissue after 48 hours (Plumb 1971). Peak virus titers occur in the kidney and intestine 72 hours after infection and in the spleen, brain, and liver at 96 hours. Virus titers in muscle tissue are comparatively low. Histopathology changes are similar in both natural and experimental CCV infections (Wolf et al. 1972; Plumb et al. 1974). Renal hematopoietic tissue is edematous and has extensive necrosis and cellular dissolution coupled with increased numbers of macrophages. The liver develops edema, necrosis, and hemorrhage; hepatic cells have eosinophilic intracytoplasmic inclusions. Pancreatic acinar cells are necrotic. The submucosa of the digestive tract is edematous with focal areas of macrophage concentrations and hemorrhage. The spleen shows extensive reduction of lymphoid tissue and becomes congested with erythrocytes. Virus particles have been seen in electron micrographs of the liver, kidney, and spleen of infected fish (Figure 5.2).

Significance The effects of a CCVD on individual channel catfish farms can be great because of the potential for exceptionally high mortality among young-of-the-year fish. However, CCVD comprises only 1–2% of total disease cases recorded at fish disease diagnostic laboratories in geographical areas where channel catfish are cultured. While individual outbreaks of CCVD can be very significant, its overall effect on the channel catfish industry has not been great.

Other viruses of catfish Several other viruses have been isolated from catfish; among them are a reovirus from channel catfish, a herpesvirus, and an iridovirus from black bullhead and an iridovirus from sheatfish.

Catfish reovirus A reovirus was isolated from overtly healthy juvenile channel catfish during a CCV assay project in California (Amend et al. 1984). The virus, named CRV, is a nonpathogenic aquareovirus when injected into naive channel catfish. The virus replicates in CCO, BB, and CHSE214 cells; FHM cells are refractory. Optimum incubation temperature is 25–30◦ C. Characterization of CRV showed that the icosahedral virions have a double capsid and a diameter of 75 nm (Hedrick et al. 1984). Nucleic acid staining suggests a double-stranded RNA with 11 segments. Serum neutralization tests made with CRV, chum salmon (CSV), and golden shiner virus (GSV) indicate that these reoviruses are distinctly different, yet some cross-reactivity occurs. Using morphological and biochemical properties of four aquareoviruses (CRV, GSV, CSV, and 13p2 from American oyster), Winton et al. (1987) showed that these viruses were unlike other genera of Reoviridae and formed the basis for a new genus of aquareoviruses. Since CRV causes no pathology in injected channel catfish, it is considered to be insignificant.

Black bullhead herpesvirus Black bullhead herpesvirus was associated with acute losses of adult black bullheads in 1994 in northern Italy at water above 20◦ C (Alborali et al. 1996). The virus was originally designated Ictalurus melas herpesvirus, later, Ictalurid herpesvirus 2 and most recently the name Ameiurine herpesvirus 1 (AmHV-1) has been proposed (Hedrick et al. 2003; Droszpoly

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et al. 2008). This virus has only been reported in Italy but its origin remains a mystery since black bullheads are not native to Europe. The black bullhead herpesvirus in black bullhead in Italy caused clinical signs similar to those of CCV in channel catfish, which include spiral swimming and vertical positioning in the water column prior to death. Hemorrhages develop in abdominal skin and at the base of fins; internally the kidney and other major organs also show hemorrhaging (Alborali et al. 1996). Upon isolation in cell cultures, the virus produces CPE consisting of syncytia similar to that of CCV, but it has a wider host range including BF-2, EPC, and CCO cells (Hedrick et al. 2003). Negative stain-electron microscopy revealed a typical herpesvirus with a nucleocapsid measuring 107 nm in diameter and an envelope measuring 277 nm (Hedrick et al. 2003). These authors also show that black bullhead herpesvirus is antigenicly and molecularly distinct from CCV. Droszpoly et al. (2008) used partial DNA sequencing to demonstrate that the black bullhead herpesvirus was genetically similar to CCV but was a distinct virus species. The black bullhead herpesvirus was shown to be pathogenic to fry and fingerling channel catfish at 25◦ C, precipitating 78% to 96% cumulative mortality, respectively, following immersion or injection and this level of mortality was higher than groups exposed to CCV (Hedrick et al. 2003). The susceptibility of channel catfish to black bullhead herpesvirus and its ability to cause high losses in adult black bullheads during the natural outbreaks in Italy suggests that this agent could be potentially devastating if it became established in an area of intensive channel catfish culture. In view of this it is strongly recommended that the introduction of this virus into geographical regions where channel catfish are produced should be prevented. Histologically, the posterior kidney was most severely affected with diffuse necrosis of the hematopoietic organs. Microscopic lesions in infected black bullheads are most severe in the kidney; lesions include hemorrhage and necrosis of hematopoietic cells (Alborali et al.

1996). Focal necrosis, congestion, and hemorrhages also occur in the spleen and liver.

European catfish virus European catfish virus (ECV) was isolated from cultured black bullhead in France (Pozet et al. 1992). Diseased fish showed body and muscle edema and petechiae around pectoral and abdominal girdles and viscera. Gills were pale, and ascites was present in the body cavity. European catfish virus was isolated in EPC cells but also grew well in BF-2 and CCO cells at temperatures of 15–25◦ C. The virus forms very small round plaques in BF-2 cells under agar overlays. The hexagonal virions measure 150–160 nm in diameter. Biochemical tests and inhibition tests with 5-iodo-2deoxyuridine indicate a DNA genome; these results combined with virus morphology are compatible with iridovirus characteristics. Sequence analysis of genes in ECV demonstrated that it is in the ranavirus genus (Mao et al. 1997; Hyatt et al. 2000). The ECV was isolated from dead fish in a population in which 100% succumbed to infection over a 1-week period. Experimental infection trials via intraperitoneal injection and waterborne exposure of subadult and adult black bullheads held at 18◦ C resulted in high mortality. High levels of virus were found in internal organs (kidney and spleen) of infected fish and necrosis occurred in the spleen and kidney. Subsequently most virus-exposed fish died. The full significance of ECV is not yet known, but it could be important where black bullheads are cultured.

European sheatfish virus The European sheatfish virus (ESV) was isolated in 1988 from cultured sheatfish in a recirculating system in Germany (Ahne et al. 1990). Moribund sheatfish (European catfish) fry infected with this virus show spiral swimming and hemorrhage in the skin and internal organs. The virus can be isolated in BF-2 cells when incubated at 25◦ C.

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Data evaluating genome restriction enzyme profiles and gene sequences homologies demonstrate that ESV is a ranavirus and suggest that ECV and ESV are closely related and may be strains of the same virus species (Mao et al. 1997; Hyatt et al. 2000; Pallister et al. 2007). Experimental infection of sheatfish fry by waterborne exposure resulted in 100% mortality within 8 days (Ogawa et al. 1990). Horizontal transmission from infected fish to naive fry resulted in 100% mortality in 11 days at 25◦ C. Pathology of infected fish included a generalized destruction of hematopoietic tissue of the spleen and kidney. The most common lesion of the gill was hyperplasia and edema of the epithelium in primary and secondary lamellae. Less severe pathology was seen in the skin, heart, eye, liver, pancreas, brain, and digestive tract. Indications are ESV could be a serious problem in culture of European catfish.

Management of catfish viruses Practical measures for controlling catfish viruses are management, avoidance, quarantine of infected stocks, and proper sanitation. Since mortality of CCV-infected fish is closely correlated to water temperature, in certain circumstances a reduction in temperature from an optimum of greater than 25◦ C to <19◦ C is beneficial. Also, accelerating hatching of eggs and fry growth by increasing water temperature should be avoided. During periods when CCVD is most likely to occur (June through September), efforts should be made to eliminate adverse environmental conditions where susceptible channel catfish are reared. Excessively high fingerling stocking rates should be avoided, water temperatures should be kept at moderate levels if possible, transportation of susceptible fish during summer should be minimized, and bacterial infections, especially “columnaris,” should be treated. Additional precautions to avoid CCV include surveying brood stock for CCV neu-

tralizing antibodies or use of other sensitive molecular virus detection methods. As CCVspecific nucleic acid probes are perfected, detection of carrier fish may become more reliable. Application of PCR techniques may provide an accurate and sensitive method for detecting CCV carrier fish. Baek and Boyle (1996) used a nested PCR procedure to detect fewer than 10 copies of CCV DNA (about 1 fg) in asymptomatic adult channel catfish. The PCR technique may be applicable to sensitive CCV detection and provide a means of avoiding this virus. Suhalim (1997) employed FAST-ELISA to measure CCV antibody in serum of juvenile and adult channel catfish that had been exposed to CCV or had produced CCV-infected offspring. There was no correlation between the FAST-ELISA optical density levels and serum neutralization titers. However, the FAST-ELISA distinctly separated sera from experimentally CCV-infected and noninfected channel catfish. Similarly, Crawford et al. (1999) used an ELISA to evaluate antiCCV antibodies and demonstrated higher sensitivity of the ELISA procedure than serum neutralization. Vaccination is being considered as a means to combat CCVD. The virus becomes attenuated (Strain V60) when passed frequently and rapidly in KlK cells (Noga and Hartman 1981; Walczak et al. 1981). When channel catfish are injected with, or bathed in, the attenuated virus they become immune to wild type (virulent) CCV. The potential for developing a CCV subunit vaccine was explored by Awad et al. (1989) who demonstrated that the envelope fraction induced protective immunity. Hanson et al. (1994) identified the thymidine kinase gene of CCV followed by a genetic engineering approach to delete this gene for vaccine development (Zhang and Hanson 1995). The thymidine kinase deletant CCV provided immunological protection of juvenile channel catfish when challenged with a lethal dose of wild type CCV. The degree of protection provided by TK-negative CCV was related to mutant dose. Later, genetic and protein analysis of the attenuated virus V60

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demonstrated a major deletion in gene 50 which encodes a secreted glycoprotein (Vanderheijden et al. 1996, 1999) and when this gene was deleted using recombinant methodology it was shown to be even more highly attenuated than the TK mutant yet induced protective immunity against CCVD (Vanderheijden 2001). A type of vaccine that uses the expression of cloned genes from DNA that is injected in fish (DNA vaccinations) was shown to have potential for protecting catfish by Nusbaum et al. (2002). They found that the product of gene 59 (the major glycoprotein) and gene 6 (a membrane protein) induced protective immunity against CCVD when expressed from intramuscularly injected plasmids. However, others have found that the DNA vaccines were not effective (Harbottle et al. 2005). Ponds from which CCV-diseased fish have been removed should be disinfected with 40 mg/L of chlorine and then drained. Although evidence suggests that epizootic survivors may be stunted (McGlamery and Gratzek 1974), they can be grown to a marketable size, providing affected fish are segregated from ponds containing healthy, susceptible channel catfish. Under no circumstances should CCV epizootic survivors be stocked in noninfected waters, nor should they knowingly be used as brood stock. No management strategy for control of black bullhead herpesvirus (IcHV-2), European catfish virus, or European sheatfish viruses has been established; however, it would be a good practice to limit the dissemination of these viruses.

References Ahne, W., M. Ogawa, and J. J. Schlotfeldt. 1990. Fish viruses: Transmission and pathogenicity of an icoshedral cytoplasmic deoxyribovirus isolated from sheatfish (Silurus glanis). Journal of Veterinary Medicine B37:187–190. Alborali, L., G. Bovo, et al. 1996. Isolation of an herpesvirus in breeding catfish (Ictalurus melas).

Bulletin of the European Association of Fish Pathologists 16:134–137. Amend, D. F., and T. McDowell. 1984. Comparison of various procedures to detect neutralizing antibody to the channel catfish virus in California brood channel catfish. The Progressive Fish Culturist 46:6–12. Amend, D. F., T. McDowell, and R. P. Hedrick. 1984. Characteristics of a previously unidentified virus from channel catfish (Ictalurus punctatus). Canadian Journal of Aquatic Sciences 41:807–811. Anonymous. 1988. Southeastern Cooperative Fish Disease Project. 24th Annual Report. Auburn University, Alabama. Awad, M. A., K. E. Nusbaum, and Y. J. Brady. 1989. Preliminary studies of a newly developed subunit vaccine for channel catfish virus disease. Journal of Aquatic Animal Health 1:233–237. Baek, Y.-S., and J. A. Boyle. 1996. Detection of channel catfish virus in adult channel catfish by use of a nested polymerase chain reaction. Journal of Aquatic Animal Health 8:97–103. Bird, R. C., K. E. Nusbaum, et al. 1988. Molecular cloning of fragments of the channel catfish virus (Herpesviridae) genome and expression of the encoded mRNA during infection. American Journal of Veterinary Research 49:1850–1855. Booy, F. P., B. L. Trus, et al. 1996. The capsid architecture of channel catfish virus, an evolutionarily distant herpesvirus, is largely conserved in the absence of discernible sequence homology with herpes simplex virus. Virology 215:134–141. Bowser, P. R., and J. A. Plumb. 1980. Channel catfish virus: Comparative replication and sensitivity of cell lines from catfish ovary and the brown bullhead. Journal of Wildlife Diseases 16:451–454. Bowser, P. R., A. D. Munson, et al. 1985. Isolation of channel catfish virus from channel catfish Ictalurus punctatus (Rafinesque) broodstock. Journal of Fish Diseases 8:557–561. Boyle, J., and J. Blackwell. 1991. Use of polymerase chain reaction to detect latent channel catfish virus. American Journal of Veterinary Research 52:1965–1968. Crawford, S. A., I. A. Gardner, et al. 1999. An enzyme linked immunosorbent assay (ELISA) for detection of antibodies to channel catfish virus (CCV) in channel catfish. Journal of Aquatic Animal Health 11:148–153.

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Davison, A. J. 1992. Channel catfish virus: a new type of herpesvirus. Virology 186:9–14. Doszpoly, A., E. R. Kovacs, et al. 2008. Molecular confirmation of a new herpesvirus from catfish (Ameiurus melas) by testing the performance of a novel PCR method, designed to target the DNA polymerase gene of alloherpesviruses. Archives of Virology 153:2123–2127. Fernandez, R. D., M. Yoshimizu, et al. 1993. Characterization of three continuous cell lines from marine fish. Journal of Aquatic Animal Health 5:127–136. Fijan, N. N., T. L. Wellborn, Jr., and J. P. Naftel. 1970. An acute viral disease of channel catfish. U. S. Fish Wildlife Service Technical Paper No. 43, 11p. Gray, W. L., R. J. Williams, et al. 1999a. Detection of channel catfish virus DNA in latently infected catfish. Journal of General Virology 80:1817–1822. Gray, W. L., R. J. Williams, and B. R. Griffin 1999b. Detection of channel catfish virus DNA in acutely infected channel catfish, Ictalurus punctatus (Rafinesque), using the polymerase chain reaction. Journal of Fish Diseases 22:111–116. Hanson, L., and R. L. Thune. 1993. Characterization of thymidine kinase encoded by channel catfish virus. Journal of Aquatic Animal Health 5:199–204. Hanson, L. A., K. G. Kousoulas, and R. L. Thune. 1994. Channel catfish herpesvirus (CCV) encodes a functional thymidine kinase gene: elucidation of a point mutation that confers resistance to Ara-T. Virology 202:659–664. Hanson, L. A., M. R. Rudis, and L. PetrieHanson. 2004. Susceptibility of channel catfish fry to channel catfish virus (CCV) challenge increases with age. Diseases of Aquatic Organisms 62:27–34. Hanson, L. A., M. R. Rudis, et al. 2006. A broadly applicable method to characterize large DNA viruses and adenoviruses based on the DNA polymerase gene. Virology Journal 3:28. Harbottle, H., K. P. Plant, and R. L. Thune. 2005. DNA vaccination against channel catfish virus results in minimal immune response and is not efficacious against challenge. Journal of Aquatic Animal Health 17:251–262. Hedrick, R. P., J. M. Groff, and T. McDowell. 1987. Response of adult channel catfish to waterborne

exposures of channel catfish virus. The Progressive Fish-Culturist 49:181–187. Hedrick, R. P., T. S. McDowell, et al. 2003. Systemic herpes-like virus in catfish Ictalurus melas (Italy) differes from Ictalurid herpesvirus 1 (North America). Diseases of Aquatic Organisms 55:85–92. Hedrick, R. P., R. Rosemark, et al. 1984. Characteristics of a new reovirus from channel catfish (Ictalurus punctatus). Journal of General Virology 65:1527–1534. Huang, S., and L. A. Hanson. 1998. Temporal gene regulation of the channel catfish virus (Ictalurid herpesvirus 1). Journal of Virology 72:1910–1917. Hyatt, A. D., A. R. Gould, et al. 2000. Comparative studies of piscine and amphibian iridoviruses. Archives of Virology 145:301–31. Kancharla, S. R., and L. A. Hanson. 1996. Production and shedding of channel catfish virus (CCV) and thymidine kinase negative CCV in immersion exposed channel catfish fingerlings. Diseases of Aquatic Organisms 27:25–34. Kunec, D., L. A. Hanson, et al. 2008. An overlapping bacterial artificial chromosome system that generates vectorless progeny for Channel catfish herpesvirus. Journal of Virology 82:3872–3881. Mao, J., R. P. Hedrick, and V. G. Chinchar. 1997. Molecular characterization, sequence analysis, and taxonomic position of newly isolated fish iridoviruses. Virology 229:212–220. McGlamery, M. H., Jr., and J. B. Gratzek. 1974. Stunting syndrome associated with young channel catfish that survived exposure to channel catfish virus. The Progressive Fish-Culturist 36:38–41 Noga, E., and J. X. Hartmann. 1981. Establishment of walking catfish (Clarias batrachus) cell lines and development of a channel catfish (Ictalurus punctatus) virus vaccine. Canadian Journal of Fisheries and Aquatic Sciences 38:925–930. Nusbaum, K. E., B. F. Smith, et al. 2002. Protective immunity induced by DNA vaccination of channel catfish with early and late transcripts of the channel catfish herpesvirus (IHV-1). Veterinary Immunology and Immunopathology 84:151–68. Ogawa, M., W. Ahne, et al. 1990. Pathomorphological alterations in sheatfish fry Silurus glanis experimentally infected with an

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iridovirus-like agent. Diseases of Aquatic Organisms 9:187–191. Pallister, J., A. Gould, et al. 2007. Development of real-time PCR assays for the detection and differentiation of Australian and European ranaviruses. Journal of Fish Diseases 30:427–38. Plumb, J. A. 1971. Tissue distribution of channel catfish virus. Journal of Wildlife Diseases 7:213–216. Plumb, J. A. 1972. Effects of temperature on mortality of fingerling channel catfish (Ictalurus punctatus) experimentally infected with channel catfish virus. Journal of the Fisheries Research Board of Canada 30:568–570. Plumb, J. A. 1973. Neutralization of channel catfish virus by serum of channel catfish. Journal of Wildlife Diseases 9:324–330. Plumb, J. A., and J. Chappel. 1978. Susceptibility of blue catfish to channel catfish virus. Proceedings of the Annual Conference Southeastern Association of Fish Wildlife Agencies 32:680–685. Plumb, J. A., and J. L. Gaines, Jr. 1975. Channel catfish virus disease. In: R. E. Ribelin and G. Migaki (eds) The Pathology of Fishes. Madison, Wisconsin, University of Wisconsin Press, pp. 287– 302. Plumb, J. A., J. L. Gaines, et al. 1974. Histopathology and electron microscopy of channel catfish virus in infected channel catfish, Ictalurus punctatus (Rafinesque). Journal of Fish Biology 6:661–664. Plumb, J. A., O. L. Green, et al. 1975. Channel catfish virus experiments with different strains of channel catfish. Transactions of the American Fisheries Society 104:140–143. Plumb, J. A., V. Hilge, and E. F. Quinlan. 1985. Resistance of the European catfish (Silurus glanis) to channel catfish virus. Journal of Applied Ichthyology 1:87–89. Plumb, J. A., R. L. Thune, and P. H. Klesius. 1981. Detection of channel catfish virus in adult fish. Developments in Biological Standardization 49:29–34. Plumb, J. A., L. D. Wright, and V. L. Jones. 1973. Survival of channel catfish virus in chilled, frozen and decomposing channel catfish. The Progressive Fish-Culturist 35:170–172. Pozet, F., M. Morand, et al. 1992. Isolation and preliminary characterization of a pathogenic icosahedral deoxyribovirus from the catfish (Ictalurus melas). Diseases of Aquatic Organisms 14:35–42.

Robin, J., and A. Rodrique. 1980. Resistance of herpes channel catfish virus (HCCV) to temperature, pH, salinity and ultraviolet irradiation. Reviews in Canadian Biology 39:153–156. Roizman, B. 1990. Herpesviridae: a brief introduction. In: B. N. Fields et al. (eds) Virology, Second Edition. New York, Raven Press, pp. 841– 847. Silverstein, P. S., V. L. van Santen, et al. 1998. Expression kinetics and mapping of the thymidine kinase transcript and an immediate-early transcript from channel catfish virus. Journal of Virology 72:3900–3906. Stingley, R. L., and W. L. Gray. 2000. Transcriptional regulation of the channel catfish virus genome direct repeat region. Journal of General Virology 81:2005–2010. Stingley, R. L., B. R. Griffin, and W. L. Gray 2003. Channel catfish virus gene expression in experimentally infected channel catfish, Ictalurus punctatus (Rafinesque). Journal of Fish Diseases 26:487–493. Suhalim, R. 1997. Detection of channel catfish virus antibody using Falcon assay screening testenzyme linked immunosorbent assay. M.S. Thesis, Auburn University, Auburn, Alabama. Thompson, D. J., L. H. Khoo, et al. 2005. Evaluation of channel catfish virus latency on fingerling production farms in Mississippi. Journal of Aquatic Animal Health 17:211–215. Walczak, E. M., E. J. Noga, and J. X. Hartmann. 1981. Properties of a vaccine for channel catfish virus disease and a method of administration. Developments in Biological Standardization 49:419–429. Winton, J. R., C. N. Lannan, et al. 1987. Morphological and biochemical properties of four members of a novel group of reoviruses isolated from aquatic animals. Journal of Virology 68:353–364. Vanderheijden, N., P. Alard, et al. 1996. The attenuated V60 strain of channel catfish virus possesses a deletion in ORF50 coding for a potentially secreted glycoprotein. Virology 218:422–426. Vanderheijden, N., L. A. Hanson, et al. 1999. Channel catfish virus gene 50 encodes a secreted, mucin-like glycoprotein. Virology 257:220–227 Vanderheijden, N., J. A. Martial, and L. A. Hanson. 2001. Channel Catfish Virus Vaccine. U.S. Patent 6,322,793. Wise, D. A., P. R. Bowser, and J. A. Boyle. 1985. Detection of channel catfish virus in asymptomatic

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adult catfish, Ictalurus punctatus (Rafinesque). Journal of Fish Diseases 8:485–493. Wise, D. A., S. F. Harrell, et al. 1988. Vertical transmission of channel catfish virus. American Journal of Veterinary Research 49:1506–1509. Wolf, K., and R. W. Darlington. 1971. Channel catfish virus: a new herpesvirus of ictalurid fish. Journal of Virology 8:525–533. Wolf, K., R. L. Herman, and C. Carlson. 1972. Fish viruses: histopathologic changes associated with experimental channel catfish virus disease. Jour-

nal of the Fisheries Research Board of Canada 29:149–150. Zhang, G. H., and L. A. Hanson. 1995. Deletion of thymidine kinase gene attenuates channel catfish herpesvirus while maintaining infectivity. Virology 209:658–663. Zhang, H. G., and L. A. Hanson. 1996. Recombinant channel catfish virus (Ictalurid herpesvirus 1) can express foreign genes and induce antibody production against the gene product. Journal of Fish Diseases 19:121–128.

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Chapter 6

Carp and minnow viruses

Cyprinidae, the family including carp and minnows, has worldwide distribution, and the highest numbers of species of any family including many that are economically important in aquaculture. As would be expected, this group has many virus pathogens. The most pathogenic of these are the rhabdoviruses (spring viremia of carp virus, SVCV), a reovirus and three herpesviruses. The three herpesviruses are closely related but cause distinctly different diseases: (1) fish pox (cyprinid herpesvirus 1, CyHV-1), (2) cyprinid herpesvirus 2 (CyHV-2), and (3) koi herpesvirus (cyprinid herpesvirus, 3 CyHV-3). Additionally, cyprinids may be infected by viruses that have a wide host range such as viral hemorrhagic septicemia virus (Chapter 8), and pike fry rhabdovirus (PFR) (Chapter 10).

Spring viremia of carp Spring viremia of carp is a subacute to chronic disease of subadult cultured common carp. The disease is important to common carp aquaculture and the causative agent, SVCV, is an OIE notifiable pathogen (OIE 2008). The 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.

virus was found in the former Yugoslavia and isolated in tissue culture from diseased fish with clinical signs of infectious dropsy (ID) (Fijan et al. 1971). Consequently, it was proposed that ID syndrome be divided into two etiologically and clinically independent diseases: (1) SVC, causative agent SVCV, and (2) carp erythrodermatitis (CE), a bacterial skin infection characterized by necrotic lesions surrounded by hemorrhagic areas (Fijan 1972). Bootsma et al. (1977) demonstrated that CE was associated with the bacterium Aeromonas salmonicida achromoqenes (atypical A. salmonicida), which was later shown to be the causative agent of goldfish ulcer disease. It should be noted that Aeromonas hydrophila, historically thought to be the causative agent of ID, can complicate both SVC and CE infections as a secondary infection, which is probably responsible for the dropsy that is associated with the SVCV. Ahne et al. (2002) provides a detailed review of SVCV.

Geographical range and species susceptibility For many years, spring viremia of carp had occurred primarily in Europe: the former Yugoslavia, Hungary, Czechoslovakia, Austria, Bulgaria, France, Germany, Romania, Spain, 109

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Great Britain, and the former Soviet Union (Fijan et al. 1971; Wolf 1988) where it is widespread in carp aquaculture. In 1998, SVC was reported in goldfish in Brazil (Alexandrino et al. 1998). In 2002, SVCV was first identified in United States on a koi carp aquaculture facility in North Carolina (Goodwin 2002). Subsequently it was discovered in common carp as the cause of wild carp die-off that same year in northwestern Wisconsin (Dikkeboom et al. 2004). The following year it was identified in apparently healthy wild common carp in Illinois, and in 2004, it was identified in disease outbreaks in koi in a private pond in Washington and in commercial production facility in Missouri (Warg 2007). In 2007, wild common carp with SVCV were found in the upper Mississippi river (USFWS 2007). In 2006, SVCV was identified during an inspection of adult common carp from Lake Ontario in Canada (Garver et al. 2007). During this period there were SVC outbreaks in China (Liu et al. 2004). Essentially SVCV may be found throughout the world. Fish species naturally infected with SVCV are common carp, crucian carp (goldfish), bighead carp, silver carp, grass carp (Shchelkunov and Shchelkunova 1989), koi carp (color carp) (Warg 2007), and tench and sheatfish (Fijan et al. 1981). Guppies, northern pike fry, pumpkinseed, roach, and zebrafish have been experimentally infected (Ahne 1985a; Haenen and Davidse 1993; Sanders et al. 2003). Sequence analysis of the glycoprotein gene of rhabdovirus of penaeid shrimp (RPS) isolated from diseased shrimp in Hawaii (Lu et al. 1991) was almost identical to SVCV, suggesting that the host range of SVCV may include invertebrates (Johnson et al. 1999).

Clinical signs At the onset of SVC, fish are attracted to the water inlet and seriously affected fish become moribund, respire slowly, and lie on their side (Fijan 1972). Fish experimentally infected by

cranial injection express similar behavior. External clinical signs include dark pigmentation, a pronounced enlargement of the abdominal area, exophthalmia, a prolapsed and inflamed anus, and pale gills with distinct petechiae (Figure 6.1). Internally, a generalized hyperemia is apparent with peritonitis, enteritis, and hemorrhages in the kidney, liver, and air bladder. Also, the liver is edematous and adhesions and hemorrhages occur in muscle tissue. In fry and small carp, the swim bladder becomes inflamed and shows focal hemorrhaging or petechiae.

Diagnosis Spring viremia of carp is diagnosed by combination of recognizing clinical signs, isolation of SVCV from diseased fish in cell culture, and molecular methods. The virus replicates between 10 and 30◦ C with an optimum of 21◦ C. Under normal conditions, the virus can be isolated only from fish during clinical infection. Fathead minnow (FHM) and epithelioma papillosum cyprini (EPC) cell lines, as well as primary carp ovary cells are susceptible to SVCV and virus yields of 108 pfu/mL occur. The BB, RTG-2 (rainbow trout gonad), and BF-2 (bluegill fry) cells are also susceptible to SVCV, but CPE develops more slowly and virus yield is lower than in other cell lines (Fijan et al. 1971; de Kinkelen and Le Berre 1974). Focal CPE includes granulation of chromatin in the nucleus with thickening or lysis of the nuclear membrane and cytoplasmic degeneration. Advanced CPE consists of degeneration and rounding of cells, which then detach from the culture vessel surface (Figure 6.2). SVCV is positively identified by neutralization tests using specific SVCV antiserum, but their utility is reduced because some cross reactive neutralization may occur with other fish rhabdoviruses, i.e., pike fry virus (Table 6.1). The antigen can be detected in frozen liver, kidney, and spleen tissues by either immunoperoxidase (ELISA) or fluorescent antibody

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(a)

(b)

(c)

Figure 6.1 Common carp infected with spring viremia of carp virus. (a) Petechiae in the skin (arrow). (b) Upper fish with pale but hemorrhaged (arrow) gills and lower fish with normal gills. (c) Petechiae in the muscle and swim bladder (arrow) and pale liver (L). (Photos (a) and (b) courtesy of N. Fijan; Photo C by P. Ghitino.)

(Faisal and Ahne 1984). These serological techniques identify SVCV in FHM cell cultures 12 hours after inoculation as compared to 48 hours required for CPE appearance in cell cultures. Way (1991) used an ELISA system to

detect SVCV antigen in inoculated cell cultures and in tissue extracts from clinically infected fish within 1 hour. This system also detected virus in subclinical fish but with less accuracy and some false positives were noted. The

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(a)

(b)

Figure 6.2 (a) Focal CPE of spring viremia of carp virus in epithelioma papillosum cyprini (EPC) cells. Infected cells are rounded and detached (×230) (Photo by N. Fijan). (b) Massive spring viremia of carp virus CPE in fathead minnow cells 72 hours after infection (Photos courtesy of W. Ahne).

antibody in SVCV–ELISA reagents also reacted with PFR but at a very low level. Commercial kits using polyclonal or monoclonal SVCV antibodies showed too much cross reaction or false positives for use in confirmation of SVCV for certification purposes (Dixon and

Longshaw 2005; OIE 2009). Because SVCV is an OIE reportable disease, definitive confirmation may be needed such as reverse transcription polymerase chain reaction (RT-PCR) amplification of the glycoprotein gene followed by sequencing (Stone et al. 2003; OIE 2006).

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Table 6.1 Cross neutralization tests with five rhabdoviruses from fish against homologous and heterologous antisera. 50% Plaque Reduction Titer of Virus† Antisera∗

SVC SBI PFV IHN VHS

SVC

SBI

PFV

IHN

VHS

6,500 980 33 25 10

6,200 980 28 25 10

20 68 200 27 10

20 50 27 1,700 10

10 10 10 10 2,300

Source: Hill et al. (1975). ∗ Acronym of viral antigen: SVC, spring viremia of carp virus; SBI, swim bladder infection of carp; PFV, pike fry rhabdovirus, IHN, infectious hematopoietic necrosis virus; VHS, viral hemorrhagic septicemia virus. † Reciprocal of antiserum dilution giving 50% plaque reduction against the specified virus.

Koutina´ et al. (2003) utilized a nested RTPCR to identify SVCV in infected cell culture and fish tissues. Identity of cDNA was confirmed by direct sequencing. The PCR sensitivity in cell-culture systems was assessed at about 10-1 TCID50 /mL. Accuracy of the method was confirmed by observing virions in electron micrographs of clinical samples, and positive ELISA; this lead to the suggestion that the RTPCR method has a potential for an accurate diagnostic method. Shivappa et al. (2008) developed a loop-mediated isothermal amplification (LAMP) procedure for detecting SVCV in koi carp. The procedure is specific, sensitive, and can be done in a short time.

Virus characteristics SVCV is a bullet-shaped RNA virus measuring approximately 70 × 180 nm (Ahne 1976). Virus activity is destroyed when exposed to 45◦ C, ether, or pH 3 (de Kinkelen and Le Berre 1974). In tissue culture medium containing 5% serum, the virus survives at a low level for 28 days at 23◦ C and for over 6 months at 4◦ C and −20◦ C. Infectivity is reduced if serum is not included in the medium. Most disinfectants (formalin, sodium hydroxide, and chlorine) kill the virus in minutes, but it remains infective for up to 42 days in water and mud. The genome is a nonsegmented, negative sense, single-stranded RNA 11,019 nucleotides in length and encodes

five proteins. Genome structure and sequence similarity studies confirm earlier indications that SVCV is a member of the Vesiculovirus genus (Ahne et al. 2002). Virus isolates from carp with swim bladder inflammation (SBI), named SBI virus, appear to have the same characteristics as SVCV (Bachmann and Ahne 1973). Hill et al. (1975) showed that virus isolated from SBI-infected carp was serologically identical to SVCV. Biochemical properties of SVC and PFR are similar and there may be a serological relationship between these two rhabodviruses of the vesiculovirus group (Enzmann et al. 1981). Immunochemical comparisons indicate that SVCV and PFR share common antigenic determinants on the G, N, and M proteins, which may be a problem in distinguishing them by indirect fluorescent antibody technique. However, these viruses can be differentiated by ELISA and serum neutralization tests while being aware of possible cross-reactivity. SVCV is distinctly different from IHNV and VHSV of salmonids (novirhabdovirus group) (Table 6.1). The best measure of relationships between rhabdoviruses is by RNAse protection assays (Ahne et al. 1998) or by genomic RNA sequence analysis. Several rhabdoviruses isolated from cyprinids proved to be PFR. When Stone et al. (2003) compared the sequences of a 550 nucleotide fragment of the glycoprotein gene of 36 SVCV and PFR isolates from several host species and geographical locations, they

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found that these viruses were related but four distinct genogroups existed. All SVCV isolates were in genogroup 1 and within this group there were four subgroups. Subsequent additional evaluations found all Asian and North American isolates were genogroup 1a and all 1a genotypes identified were from these regions (Garver et al. 2007).

Epizootiology Spring viremia of carp occurs primarily in populations of carp 1 year old or older. Epizootics of SVC were initially confirmed during the spring of 1969 and 1970 in Yugoslavia when water temperatures ranged from 12 to 22◦ C (Fijan et al. 1971). The optimum temperature for this disease is 16–17◦ C. Following waterborne virus exposure, 50 g carp were evaluated to clarify what role water temperature played in SVC development (Baudouy et al. 1980). Clinical signs appeared between 11 and 18◦ C. Some fish survived infection and became immune to the disease while others died. Below 10◦ C, no immunity developed and disease was always fatal. The researchers suggested that elevated water temperatures in the spring are not necessarily a factor in the disease process. Severity of SVC varies from farm to farm, pond to pond, and year to year on a given farm (Fijan 1972). After consecutive years with no SVC outbreaks, heavy losses can be experienced for several years. What causes these severity variations in SVC outbreaks is unknown but probably results from a combination of susceptibility of fish, a virulent virus strain, and environmental conditions. Carp that survived an SVCV infection in the laboratory were resistant to repeated challenges with virulent virus, but remained susceptible to CE. This further supports different etiologies for SVC and CE, the latter probably having a bacterial etiology. Spring viremia of carp is transmitted by waterborne exposure, cohabitation of infected

and healthy fish, contaminated food, and by intraperitoneal, intramuscular, or intracranial injection. The virus is shed from infected fish via feces and urine; therefore, waterborne transmission is the primary route of infection. Fish parasites such as leaches or fish louse (Argulus foliaceus and Pisciocola geometra) can also transmit SVC to uninfected fish (Ahne 1985b). Incubation time for SVC varies from 6 to 60 days depending upon water temperature and exposure method. Mean survival of carp experimentally infected with virus by injection in the swim bladder was 8.9 days compared to 12.8 days for intraperitoneal injection and 12.9 days for intracranial injection (Varovic and Fijan 1973). Mortality of common carp infected with SVCV varies but Haenen and Davidse (1993) reported 97.7% mortality over 35 days following waterborne exposure to 105.8 TCID50 /mL in water. Shedding of SVCV by adult carp during spawning was reported by B´ek´esi and Csontos (1985). They isolated SVCV from 3 of 491 ovarian fluid samples assayed but none from 211 seminal fluid samples. Whether or not virus was actually inside the eggs was not determined. However, it does indicate that some adult survivors of SVCV infections can be carriers, even though survivors generally do not shed infectious virus after clinical disease dissipates.

Pathology During waterborne exposure, SVCV enters the host through the gills where virus replicates and then spreads via the blood stream. Primary viremia becomes evident at 6 days postinfection (Ahne 1978). Target organs are kidney, liver, spleen, heart, and alimentary canal. Clinical disease is evident 7 days postexposure, and virus is shed by intestinal mucus casts and feces. Nagele (1977) described the histopathology of experimentally infected carp fingerlings.

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Carp and Minnow Viruses 115

The liver has edema, necrotic blood vessels, and focal necrosis in the parenchyma. Vessels in the peritoneum are hyperemic and intestines become inflamed. The spleen is hyperemic and hyperplastic. Renal tubules are clogged with casts and tubule cells develop cytoplasmic inclusions. The swim bladder has extensive focal areas of hemorrhage.

Significance Spring viremia of carp is an extremely important disease of cultured carp in Europe where it is most prevalent in yearling fish with up to 70% mortality of infected populations. Adult fish are also susceptible but to a lesser degree. The overall impact of SVC has been somewhat offset by the fact that in years following an epizootic, mortalities are greatly reduced in a given year class of carp, a likely result of increased immunity among epizootic survivors. Because SVCV is an OIE-reportable disease, its presence in a region will impact trade. In the USA, USDA APHIS has an active eradication program to limit the spread of this virus.

Fish pox (CyHV-1) Fish pox, also known as carp pox, epithelioma papillosum, or carp herpesvirus, is one of the oldest known fish diseases, being recorded as early as 1563 (Hedrick and Sano 1989). It takes the form of a benign hyperplastic, epidermal papilloma on common carp. On the basis of electron microscopy, Schubert (1966) proposed that fish pox was caused by a virus and Sonstegard and Sonstegard (1978) further suggested that it was caused by a herpesvirus. Sano et al. (1985a, b) confirmed the previous findings when they for the first time isolated the virus from epithelioma growths on Japanese ornamental carp. Because the disease is not a true “pox,” they proposed that it be called papillosum cyprini and the virus desig-

nated CyHV-1. The virus is also called Herpesvirus cyprini, or carp herpesvirus (CHV).

Geographical range and species susceptibility Fish pox is present in most European countries, China, Japan, Russia, Israel, Korea, Malaysia, and the United States. Common carp and ornamental carp (Agasi, koi, color) are its primary hosts. Other cyprinids crucian carp, willow shiner, and grass carp are not susceptible (Sano et al. 1985b; 1991a). Fish pox was reported for the first time in North America in ide imported from Europe one year prior to the disease’s discovery in the United States (McAllister et al. 1985) and later in koi carp in California (Hedrick et al. 1990). According to Jiang (1995), fish pox is a major disease in cultured common carp in northern and eastern China.

Clinical signs Fish pox is benign, forming hyperplastic, papillomatous growths in the epithelium of carp (Sano et al. 1985b). The tumors are milkywhite to gray and are raised 1–3 mm above the skin on the head, fins, or any body surface (Figure 6.3). The generally small, smooth, raised growths may increase in size and cover large areas of the body surface before they eventually regress and disappear. Infected fish show no distinct behavioral signs and there is usually no morbidity among infected adult fish but an exception was reported in China (Jiang 1995). Juvenile carp are, however, adversely affected by CyHV-1 and can suffer high mortality (Sano et al. 1991a). Two-week-old common carp fry infected with the virus developed a loss of appetite, swam in rigid lines with intermittent immobility, had distended abdomens, exophthalmia, darkened skin pigmentation, and hemorrhages on the operculum and abdomen.

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(a)

(b)

Figure 6.3 (a) Massive papilloma on common carp naturally infected with cyprinid herpesvirus 1 (fish pox, herpesvirus of carp). (b) Papilloma on juvenile carp that survived experimental infection by immersion. (From Sano et al. 1991b; reprinted by permission of Blackwell Scientific Publications Ltd.)

Diagnosis When clinical signs of fish pox appear, virus isolation should be attempted in FHM or EPC cells (Sano et al. 1991a) where virus titers in fish tissues range from 102.5 TCID50 /mL in EPC cells to 105.8 TCID50 /mL in FHM cells. Optimum cell culture incubation temperature is 20◦ C; replication also occurs at 15◦ C, but not at 10 or 25◦ C. CyHV-1-infected FHM cells become vacuolated, rounded, with slight focal pyknosis 5 days after inoculation. Cellular disruption occurs slowly but will expand to 60 or 70% of the cell sheet with occasional cellular recovery. Syncytia formation, characteristic of cell lines infected with most herpesviruses, does not oc-

cur. However, intranuclear, Cowdry type A inclusion bodies develop in infected FHM cells (Figure 6.4). Sano et al. (1985b) found CyHV1 antigen in neoplastic tissues at all stages of tumor development, especially in older and recurrent tumors. Identification of CyHV-1 is made by serum neutralization, however, Sano et al. (1992) also detected CyHV-1 in infected carp fry by in situ hybridization with biotinylated probes.

Virus characteristics CyHV-1 is a DNA virus with an icosahedral capsid measuring 110–113 nm in diameter (Figure 6.4) and an envelope measuring

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Epizootiology

(a)

(b)

Figure 6.4 (a) Fathead minnow cells infected with herpesvirus of carp showing Cowdry Type A inclusion bodies (arrow). (b) Cyprinid herpesvirus 1 with electron dense nucleocapsid and envelope. (Photos courtesy of T. Sano.)

190 nm. The virus is sensitive to ether, pH 3, 50◦ C for 1 hour, and replication is inhibited by iododeoxyuridine (IUdR) (Sano et al. 1985b). Basically only one CyHV-1 strain is recognized but Sano et al. (1991b) noted slight differences in the viral genome of two different isolates by restriction endonuclease cleavage profiles. Sequence analysis of portions of the genes encoding helicase, DNA polymerase, an intercapsomeric triplex protein and major capsid protein indicates that CyHV-1 is closely related to CyHV-3 (KHV) and cyprinid herpesvirus 2 (goldfish herpesvirus) (Waltzek et al. 2005) and can be broadly grouped within the herpesviruses of fish and amphibians. This places it within the family Alloherpesviridae (ICTV 2008).

Most data concerning CyHV-1 has been generated from cultured fish populations; its incidence in wild common carp is not known but is assumed to be low. Jiang (1995) reported death of 200 kg of CyHV-1 infected caged common carp of unspecified fish size in a large reservoir in China. It has been suggested that young fish are infected from carrier brood fish, but whether this occurs vertically and/or horizontally is unknown. Released virus from ruptured cells sloughed from the skin is probably instrumental in transmission. Evidence indicates that the disease can be transmitted by cohabitation or experimentally by intramuscular and intraperitoneal injection of media from infected cell cultures. Time between exposure and tumor development is 5 months to a year at 15◦ C (McAllister et al. 1985; Sano et al. 1991b). Sano et al. (1985a) found that CyHV-1 can be transmitted to 30-day-old carp and manifests itself in papillomas 6 months later. Epizootiology confusion surrounding fish pox was clarified by Sano et al. (1991b), who infected 2-week-old common carp fry by immersion at 20◦ C, resulting in viremia and clinical disease. From 86 to 97% of virus-exposed fish died compared to 3% mortality in controls. CHV was isolated from infected fish for 3 weeks postexposure. When 4- and 8-weekold common carp were infected with CyHV-1, mortality was 20 and 0% respectively; however, neoplasia developed in 55% of survivors 6 months postexposure to CyHV-1 and virus was isolated from tumors. A second set of neoplasia developed in 83% of the fish 7.5 months after initial tumors had sloughed; CyHV-1 was again isolated from all necropsied moribund fish and survivors. Papillomas also developed in 13% of adult mirror carp and 10% of adult fancy carp 5 months after intraperitoneal virus injection. The foregoing studies suggest that CyHV-1 replicates in epidermal tissue and can be suppressed or stimulated by water temperature.

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Although CyHV-1-induced papillomas occur at an optimum temperature of 15◦ C, lesion regression begins when water temperature rises to 20◦ C and accelerates as temperature is increased to 30◦ C (Sano et al. 1993a). The regression mechanism was not understood until Morita and Sano (1990) showed that peripheral blood lymphocytes (PBL) play an important role in this process. When carp with CyHV-1 papillomas were injected with anticarp PBL serum, regression was delayed, suggesting that spontaneous cytoxicity of carp PBL contributes to carp papilloma regression. This would explain the cycle of neoplasia with an induction phase, desquamation, and recurrence being related to environmental and hormonal cycles and PBL. The CyHV-1 genome was traced in carp by in situ hybridization with a biotinylated probe following acute infection (Sano et al. 1993b). The viral genome was detected in the cranial nerve ganglia, subcutaneous tissue, and in spinal nerves. Viral antigen was not detected nor was virus isolated from these fish. However, when papillomas did appear, the viral genome was found in the same tissue as before, and virus antigen and infective virus particles were present in the papillomas. The viral genome was still present in the spinal nerves, cranial nerve ganglia, and subcutaneous tissue after papilloma regression. It was suggested that CyHV-1 becomes latent in these tissues and is associated with lesion recurrence.

Pathology Histopathology of fish pox tumors was described by Sano et al. (1985a, b) and McAllister et al. (1985). Scaled areas of the skin show hyperplastic epidermal cells and similar lesions develop on fins and where no epidermal mucus or alarm substance cells are present. The compact dermis layer is thickened where eosinophils and lymphocytes in loose connective tissue or hypodermis suggest an in-

flammatory response. The oncogenic nature of CyHV-1 in carp was demonstrated by Sano et al. (1985a) when they experimentally produced tumors on fish skin and fins. The juvenile common carp developed a viremia and had necrosis in the liver parenchyma, kidneys, and lamina propria of the intestinal mucosa. Moribund fish had severe hepatitis and extensive necrosis in the liver, limited necrosis in renal tissue, and epithelial erosion and edema of gills.

Significance When very young carp are infected with CyHV-1, high mortality can occur and survivors may develop tumors within a year. Fish pox, generally regarded benign except in juveniles, however, from an aesthetic point of view, CyHV-1-infected adult fish, particularly “fancy carp” are unsightly and undesirable for food or display. Therefore, because mortality can be high in juveniles and tumors lowers market value of adults, one must conclude that CyHV-1 has a significant impact in carp culture.

Cyprinid herpesvirus 2 (CyHV-2) A previously unknown herpesvirus was isolated from cultured goldfish in Japan (Jung and Miyazaki 1995). The disease, named herpesviral hematopoietic necrosis (HVHN), caused a severe epizootic but no external clinical signs were apparent on affected fish. The virus has been designated CyHV-2.

Geographical range and species susceptibility Following the initial detection and isolation of HVHN in Japan (Jung and Miyazaki 1995), CyHV-2 was detected in diseased goldfish in Taiwan (Chang et al. 1999), at several

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locations in the USA (Groff et al. 1998, Goodwin et al. 2006a), Australia (Stephens et al. 2004), and the UK (Jeffery et al. 2007). Given the distance between diagnosed outbreaks of HVHN and the difficulty in culturing CyHV-2, it can be assumed that this virus has been widely dispersed with the international goldfish trade. The agent has only been identified in goldfish; challenge trials in koi carp caused no disease, suggesting that CyHV2 has a limited host range (Jung and Miyazaki 1995).

Clinical signs Goldfish populations with HVHN can experience high losses; however, no distinct clinical signs are associated with infection. Affected individuals are generally lethargic and inappetent and display pale patches on the gills but no other notable external clinical signs. Internally, the spleen and kidney of HVHN-affected fish are swollen and discolored (Jung and Miyazaki 1995; Groff et al. 1998; Stephens et al. 2004; Goodwin et al. 2006a; Jeffery et al. 2007). In some cases, granular nodules were found in the spleen of affected fish (Jung and Miyazaki 1995; Jeffery et al. 2007).

Diagnosis A presumptive diagnosis of HVHN is based on high mortality, presence of limited clinical signs; and hyperplasia and necrosis of gill epithelium and hematopoietic tissue with nuclei of some cells displaying chromatin along the margins (Jung and Miyazaki 1995; Groff 1998). Jung and Miyazaki (1995) reported isolating the virus in both FHM and EPC cells, but isolation is not dependable. Other investigators cultured the virus with limited success in koi fin cells (KF-1) cells (Jeffery et al. 2007) with CPE occurring after 10 days of incubation at 20◦ C; CPE consisted of granulation, vacuolation, and rounding of the cells. Because of

the difficulty in culturing CyHV-2, PCR and sequencing or CyHV-2-specific PCR was advocated and described (Goodwin et al. 2006b; Jeffery et al. 2007). Another PCR procedure was reported by Waltzek et al. (2009) using the sequence found in the helicase gene of CyHV2 that was specific for this virus; this distinguishes CyHV-2 from CyHV-1, CyHV-3, or the genomic DNA of ictalurid herpesvirus-1 (IcHv-1, CCV) because it does not amplify DNA from these viruses. The helicase CyHV2 PCR, therefore, complements real-time PCR test as a conventional method of detection and preventing the spread of CyHV-2 by serving as a tool to determine CyHV-2 free stocks of goldfish.

Virus characteristics CyHV-2 undergoes replication in the nucleus and produces an enveloped virion that measures 170–220 nm. Its replication is sensitive to IudR and virus infectivity is inactivated by pH3 and ether (Jung and Miyazaki 1995). Sequence analysis of portions of the genes encoding helicase, DNA polymerase, an intercapsomeric triplex protein, and major capsid protein indicates that cyprinid herpesvirus 2 is closely related to CyHV-3 (KHV) and CyHV1 (CHV) (Waltzek et al. 2005) and can be broadly grouped within the herpesviruses of fish and amphibians. This places it within the family Alloherpesviridae (ICTV 2008).

Epizootiology Outbreaks of HVHN generally occur when temperatures are between 15◦ C and 25◦ C, affecting all sizes of goldfish in aquaculture facilities and ornamental pools. Losses are species specific with cohabitating koi showing no signs of disease. The virus is widespread in goldfish farms in the United States; Goodwin et al. (2009) tested fish from 18 farms, 14 of which were positive for CyHV-2 by PCR. Also 42

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samples of brood goldfish were positive from 118 pools tested. The use of sensitive PCR demonstrates that goldfish that show no clinical signs of HVHN may harbor CyHV-2, and given the nature of other herpesvirus, this likely represents a latent carrier state (Goodwin et al. 2006b). More recently Goodwin et al. (2009) subjected apparently healthy CyHV-2 carrier goldfish to cold shock (rapid change from 22◦ C to 10◦ C), stimulating them to test positive for the virus by PCR; heat shock, high ammonia, or high pH did not affect the virus positive status. This trend also has been observed on commercial goldfish farms where outbreaks occur naturally when the fish are subjected to a sharp temperature drop followed by holding them at the low permissive temperature for the disease. Apparent wide distribution of CyHV-2, the recalcitrant nature to cell-culture-based diagnostics and the prevalent covert carrier state, suggests that PCR-based screening of populations may be required to avoid CyHV-2 introduction when bringing fish into a goldfish production facility.

Pathology The most prominent changes associated with HVHN are necrotic foci in the hematopoietic tissue, gills, spleen pulp, pancreatic cells, and lamina propria of the intestine. The nuclei of some cells in the affected tissue display margination of the chromatin and pale basophilic intranuclear inclusions (Jung and Miyazaki 1995; Groff et al. 1998; Goodwin et al. 2006a; Jeffery et al. 2007).

Significance The number of confirmed cases of HVHN is limited but testing by Goodwin et al. (2009) indicates the virus is widespread in the goldfish culture industry in the United States. Losses in some confirmed cases were substan-

tial. The use of PCR methods will provide easier diagnosis of this disease and may result in a better evaluation of the disease’s importance. The apparent worldwide distribution of CyHV-2 makes this agent a potentially important emerging pathogen to the goldfish industry.

Koi herpesvirus (Cyprinid Herpesvirus 3) Since 1998, disease causing severe gill necrosis and high mortality in koi carp has been identified in USA, Europe and Asia (Bretzinger et al. 1999, Hedrick et al. 1999, 2000). After isolating the virus in cell culture, Hedrick et al. (2000) classified it as a herpesvirus and designated it as koi herpesvirus (KHV). Several unique characteristics of KHV lead to the suggestion that KHV was not a herpesvirus and referred to it as carp nephritis and gill necrosis virus (CNGV) (Ronen et al. 2003; Hutoran et al. 2005). However analysis of specific genes (Waltzek et al. 2005) and whole genome sequencing (Aoki et al. 2007) confirm that the virus is a herpesvirus and it was given the designation cyprinid herpesvirus 3 (CyHV-3).

Geographical range and species susceptibility Since its first recognition in United States, Germany, and Israel, KHV has been found throughout Europe, South Africa, Japan, Indonesia, Thailand, Taiwan, China, and Malaysia (Liu et al. 2002, Haenen et al. 2004; Sano et al. 2004, Tu et al. 2004, Hedrick et al. 2005). The disease can cause devastating losses in both wild and cultured common carp, including the ornamental variety (koi) and as such KHV is an OIE notifiable pathogen (OIE 2008). The disease appears to be restricted to Cyprinus carpio but other species including goldfish may become infected by KHV and, thus, may function as unapparent carriers

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(a)

(b)

(c)

(d)

Figure 6.5 Lesions and cytopathic effect caused by koi herpesvirus. (a) Pale areas on the skin (arrows) of koi with koi herpesvirus disease. (b) Mottled gills due to koi herpesvirus induced lesions. (c) Histopathology of the gill of a carp with koi herpesvirus disease demonstrating the hyperplasia and an affected cell with margination of the chromatin (arrow). (d) Vacuolation and syncytium formation (arrows) in KF-2 cells caused by koi herpesvirus after 5 days. (Photos courtesy of R. Hedrick, photo B by M. Okihiro.)

(El-Matbouli et al. 2007a; Sadler et al. 2008). Experimental challenge demonstrated that goldfish, grass carp, Nile tilapia, and silver perch are resistant to KHV disease (Perelberg et al. 2003).

irregular necrotic discolored areas on the skin; gills also have patchy, irregular, discolored areas (Figure 6.5). Some fish may display hemorrhages at the base of fins and the eyes may be sunken or show exophthalmia. Internally adhesions may occur in the body cavity (Hedrick et al. 2000).

Clinical signs Populations of carp with KHV disease experience high losses at temperatures of 18–26◦ C. Affected fish are lethargic accompanied by disoriented swimming; in some cases, fish may become hyperactive before death. The predominant external signs are excessive mucus and

Diagnosis KHV disease is presumptively diagnosed by the presence of clinical signs in koi carp in the 18–26◦ C temperature range, the presence of characteristic epithelial proliferation and

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lamellar fusion of gills and the production of typical cytopathic effect on a susceptible cell line. KHV can be isolated from gill tissue and most internal organs of infected fish, primarily kidney and spleen, but the skin, brain, gut, and liver may also yield virus. Confirmation requires the use of specific antibodies or the use of KHV-specific PCR to identify the virus; PCR has not identified KHV in fish tissue (Gilad et al. 2002; Gray et al. 2002; Bercovier et al. 2005; El-Matbouli et al. 2007a). Koi fin (KF-1) (Hedrick et al. 2000) and common carp brain (CCB) (Neukirch et al. 1999a) cell lines are most sensitive for KHV isolation. Silver carp fin cells and goldfish fin cells are less susceptible to KHV, while EPC and FHM cells are resistant to KHV (Davidovich et al. 2007). In all susceptible cell lines KHV causes extensive vacuolization and moderate syncytium formation of the cells 4–10 days postinoculation (Figure 6.5). In contrast, CyHV-1 and SVCV cause a more rapid CPE characterized by rounding and detachment of the cells. Optimum KHV cell culture replication temperature is 20–25◦ C; no replication occurs at 4◦ C or 30◦ C (Gilad et al. 2003).

Virus characteristics KHV is a DNA virus with an icosahedral capsid measuring 108 nm in diameter and an envelope measuring 180–230 nm (Hedrick et al. 2000). Thin section EM demonstrates nuclear replication and virus assembly, budding from the inner nuclear membrane and transport similar to other herpesviruses (Miwa et al. 2007). The genome is linear 295 kb in length, making it the largest genome among characterized herpesviruses (Hutoran et al. 2005; Aoki et al. 2007). Sequence analysis demonstrates that it is distinct but closely related to CyHV1 (CHV) and CyHV-2 (Goldfish herpesvirus) and can be broadly grouped within the herpesvirus’ of fish and amphibians (Waltzek et al. 2005; Aoki et al. 2007). This places it within

the family Alloherpesviridae (ICTV 2008). Aoki et al. (2007) sequenced the genomes of three isolates, one each from Japan, United States and Israel. They found only slight differences in the sequences with the United States and Israeli isolates being most similar. Like other herpesvirus’, KHV genome is infectious and an infectious bacterial artificial chromosome has been produced, facilitating the production of recombinant KHV (Costes et al. 2008).

Epizootiology Temperature is the dominant environmental factor affecting onset and severity of KHV (Hedrick et al. 2005). Outbreaks of KHV disease can occur in wild or cultured populations in the spring and fall when temperatures are 18–28◦ C with losses sometimes exceeding 90% (Ishioka et al. 2005; Grimmett et al. 2006). Carp exposed to KHV by bath challenge or cohabitation exposure and held at 23◦ C develop clinical signs of KHV disease and begin dying at 7–9 days postexposure (Gilad et al. 2003, Pikarsky et al. 2004). Fish infected at 13◦ C did not experience clinical signs of KHV, but these fish harbored viral DNA detected by PCR (Gilad et al. 2004). When temperature on these fish shifted to 23◦ C 30 days postinfection, they developed disease clinical signs and mortality. If the temperature is shifted down from 21◦ C to 12◦ C shortly after exposure, then shifted up to 23◦ C several weeks later, KHV disease breaks with high losses (St-Hillaire et al. 2005). If the temperature is raised to 30◦ C, the fish survive the infection and become resistant to rechallenge (Ronen et al. 2003). In cell culture, the virus becomes latent at 30◦ C and reexpresses itself at 20◦ C (Dishon et al. 2007), suggesting that a similar mechanism may be occurring in the fish. Survivors of KHV disease retain the KHV genome presumably in a latent state (Gilad et al. 2004). Furthermore, cyprinid

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species that are not susceptible to KHV disease can carry KHV genome (El-Matbouli et al. 2007b; Sadler et al. 2008). Therefore cyprinids that have been previously exposed to KHV may be a reservoir for later KHV outbreaks. The mode of transmission has not been defined but water-borne exposure is suspected. Dishon et al. (2005) demonstrated the presence of viable KHV in the feces from infected carp, suggesting the potential for fecal shedding as a virus source for horizontal transmission. It is not known if KHV is vertically transmitted.

Pathology The most obvious pathology is proliferative inflammation of the gills in all ages of fish (Figure 6.5). Epithelial hyperplasia and hypertrophy causes fusion of secondary lamellae and is often accompanied with secondary infection by F. columnare. There may be hyperplasia and necrosis of the oral epithelium. Internally KHV disease causes inflammation and necrosis of the liver, spleen, intestinal tract, and kidney also in all ages of fish (Hedrick et al. 2000; Pilarsky et al. 2004). Necrosis within these organs occurred primarily in the hepatocytes, pancreatic acinar cells, spleen parenchyma, interstitial tissue of the kidney, and lamina propria of the intestine. The nuclei of some affected cells may display thickening of the nuclear membrane and the chromatin margin (Figure 6.5) (Hedrick et al. 2000).

Significance Unregulated international transport of koi carp probably facilitated worldwide distribution of KHV in the late 1990s. This made KHV disease one of the most devastating diseases to common and koi carp aquaculture. Recently OIE has listed KHV as a reportable disease

and many countries require carp, especially koi carp, to be certified free of KHV before importation.

Golden shiner virus An increasing number of viruses belonging to the family Reoviridae have been isolated from fish (Hetrick and Hedrick 1993). Consequently, an Aquareovirus genus was formed to include reoviruses from fish and shellfish (Francki et al. 1991). Generally aquareoviruses exhibit low or no pathogenicity for fish; however, GSV and several others have the capacity to cause significant mortality under certain circumstances. Golden shiner virus was first isolated in 1977 from cultured golden shiners in which it produces a mild viremia (Plumb et al. 1979).

Geographical range and species susceptibility Golden shiner virus appears to be confined to the southeastern region of the United States and California where golden shiners are grown commercially (Plumb et al. 1979; Hedrick et al. 1988). The virus has also been isolated from grass carp but, other fish species are apparently refractive.

Clinical signs Shiners infected with GSV may swim lethargically at the surface, dive when they are disturbed, hold their fins close to the body, and lie on the bottom of aquaria or tanks (Plumb et al. 1979). External clinical signs are not obvious but the back and head of infected fish are reddish (erythema) as a result of hemorrhage in the skin and dorsal musculature. Petechiae are present in visceral fat and intestinal mucosa. Occasionally, open ulcerative lesions will

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develop in the skin; however, these lesions may be due to secondary bacterial infections.

Diagnosis Golden shiner virus is most readily isolated from the kidney, liver, and spleen in FHM cells where replication occurs at 20–30◦ C, with an optimum temperature of 28–30◦ C. No replication occurs at 15 or 35◦ C (Brady and Plumb 1991). Some virus replication accompanied by CPE also occurs in chinook salmon embryo cells (CHSE-214) and BB cells, but virus yield is less than in FHM. At approximately 24 hours postinoculation, focal CPE occurs in FHM monolayers incubated at 30◦ C. By 48 hours, CPE envelops the entire cell sheet (Figure 6.6). Infected cells become large and rounded, detach from the culture vessel surface, and float in the media as spherical, highly vacuolated debris. If very low levels of virus (less than 101 TCID50 /mL) are inoculated onto a cell sheet, initial foci may be overgrown by apparently healthy cells, leaving no evidence of infection other than a few large spherical cells floating in the media.

(a)

(b)

Virus characteristics The aquareoviruses are icosahedral, possess a double capsid that measures 70–75 nm in diameter and a double stranded RNA genome (Plumb et al. 1979; Schwedler and Plumb 1980; Winton et al. 1987). Several studies comparing GSV, catfish reovirus (CRV), chum salmon reovirus (CBSV), and 13p 2 reovirus (of American oysters) indicate a serological relationship (Winton et al. 1987; Brady and Plumb 1988). Changes in virus nomenclature based on complete genome sequence analysis places GSV as a strain of Aquareovirus C. It is very closely related to Chinese grass carp reovirus and are considered virus strains of the same species (Attoui et al. 2002).

Epizootiology Although not a major disease, under certain conditions GSV can cause significant losses. Normally, GSV causes a chronic mortality of less than 10%; however, if fish are stressed in holding tanks, mortality can increase to 75%

(c)

Figure 6.6 (a) Non-infected control fathead minnow (FHM) cell cultures. (b) Focal infection of golden shiner virus CPE in FHM with multinucleated cells. (c) Massive CPE with rounded, vacuolated FHM cells that have released from the substrate. (From Plumb et al. 1979; reprinted with permission of Canadian Journal of Fisheries and Aquatic Sciences.)

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isolated in only about 25% of the cases. Therefore, other contributing etiologies may also be involved where GSV is suspect.

60

Percent GSV infectivity

50 12.5 g/L

Pathology

6.25 g/L

40

In addition to petechiae in the skin and occasionally the internal organs, the most notable histopathology of GSV disease is invasion of lymphocytes into the liver and intestinal hemorrhage.

30

20

10

Significance 0

0

48

96 Hours

144

196

Figure 6.7 Effect of crowding on golden shiners infected with golden shiner virus and stocked in troughs at two different densities. (From Schwedler and Plumb 1982; reprinted with permission of the Journal of Wildlife Diseases.)

and disease may be compounded by secondary columnaris. Crowding will increase numbers of GSV positive fish from very low to as high as 50% (Figure 6.7) (Schwedler and Plumb 1982). Intermediate size (7–10 cm) fish appear to be more susceptible than smaller fish. Limited experimental GSV transmission studies indicate that virus is not easily transmitted by cohabitation but is readily transmitted by injection. Possible vertical transmission has not been investigated. Early documented cases of GSV disease occurred during summer when water temperatures exceeded 25◦ C; however, A. J. Mitchell (Stuttgart National Aquaculture Research Center, Stuttgart, Arkansas, personal communication) reported that high losses to GSV have occurred from March to November. He also noted that the virus impact of GSV on the golden shiner industry is unclear because when clinical signs and rate of mortality indicate a possible GSV infection, virus can be

The disease has little impact on the aquaculture industry based on its modest pathogenicity and low frequency of occurrence. Mortality is generally low in culture ponds, but potential for high losses exist when golden shiners are stressed in holding tanks. This is especially true if a secondary infection of columnaris occurs. Increasing demand for larger golden shiners as bait for the sport fishing industry may cause GSV to become more significant because the virus seems to target older fish.

Other viruses of carp and minnows Several other viruses that have been reported in different species of cyprinids (goldfish, common carp, colored (koi) carp, grass carp, and black carp) are also noteworthy.

Goldfish iridoviruses Two tentatively identified iridoviruses were isolated from swim bladders of healthy goldfish by Berry et al. (1983). Isolates were designated goldfish virus 1 (GFV-1) and goldfish virus 2 (GFV-2). These enveloped, DNA, icosahedral viruses measure 180 nm in diameter.

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Grass carp hemorrhagic virus The disease, known as grass carp hemorrhagic virus disease (GCHV), was initially reported in China in 1972 (Nie and Pan 1985); however, it was not until the virus was isolated by Chen and Jiang (1984) that its causative agent was classified as an aquareovirus. The disease occurs in China where it primarily infects grass carp. Other susceptible species include black carp, gudgeon (topmouth), and rare minnow (Ding et al. 1991; Wang et al. 1994). The virus replicates in silver carp but causes no clinical signs. Infected grass carp develop hemorrhage in the skin, fin base, opercula, and mouth cavity; infected fish show exophthalmia of the eye and pale gills (Shen 1978). The most dramatic internal sign of GCHV is a severely hemorrhagic intestinal tract with some hemorrhaging in the liver, spleen, kidney, and musculature. Grass carp hemorrhagic virus can be isolated in a cell line developed from gonad tissue (Shen 1978) or cells derived from the snout of grass carp. Cytopathic effect appears in 3–4 days when incubated at 28◦ C. The icosahedral virus, measuring 60–78 nm in diameter, contains an RNA genome composed of 11 segments along with other reovirus characteristics (Ke et al. 1990). According to Jun et al. (1997), methods for detection and identification of GCHV include immunoenzyme staining, staphylococcal coagglutination test, immunofluorescence, and enzyme-linked immunosorbent assays. These authors also described a method for detecting GCHV based on a reverse transcription-polymerase chain reaction that detected GCHV in infected cell culture fluids and infected grass carp tissue whether or not fish exhibited hemorrhagic clinical signs. Sequence analysis of the grass carp hemorrhagic virus indicate a high degree of similarity to GSV and has resulted in the two viruses being classified as strains of Aquareovirus C (Attoui et al. 2002) The virus infects primarily fry and yearling grass carp, causing very high losses. Morbid-

ity in these fish can be over 65% following experimental infection (Shen 1978; Shao et al. 1990). Mortality is less in yearling fish than in fry where up to 80% mortality has been noted. Most epidemics occur during warm summer months as water temperatures rise to 25–30◦ C. Under these conditions GCHV becomes one of the most serious diseases affecting grass carp in China.

American grass carp reovirus A second reovirus was isolated from diseased grass carp fingerlings in Arkansas, USA, in the winter of 2005 (Mohd Jaafar et al. 2008). This virus was distinct form GCHV and GSV and was similar to golden ide reovirus identified in Germany (Neukirch et al. 1999b). The two are considered strains of Aquareovirus G. The clinical importance of this virus is not known; it was associated with disease in grass carp fingerlings, but it was also isolated from apparently healthy golden shiners. Golden ide reovirus was isolated from fish with epidermal hyperplasia.

Carp edema virus, koi sleepy disease virus Carp edema virus causes mortality of juvenile carp in earthen ponds. Affected fish are lethargic at the pond surface and have swollen gill filaments and sunken eyes (Ono et al. 1986). The causative agent was identified by electron microscopy as a Pox-like virus. The disease was transmissible using filtrates from diseased fish but was not successfully propagated in cell culture (Oyamatsu et al. 1997a). The agent was designated carp edema virus (CEV) and a PCR assay was developed (Oyamatsu et al. 1997b). Koi sleepy disease is a proliferative gill condition that occurs in Japanese koi operations after being brought in to concrete holding tanks from nursery ponds. Miyazaki et al.

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(2005) describe this disease and use CEVspecific PCR to demonstrate that CEV is the causative agent of koi sleepy disease. Affected fish have gill epithelial cell hyperplasia and hypertrophy, cloudy swelling and hyaline droplet degeneration of hepatocytes, and renal tubule cells. The skin was characterized by epidermal erosions (Miyazaki et al. 2005).

Carp cornavirus Miyazaki et al. (2000) reported on an ulcerative dermatitis in koi caused by a previously undescribed virus. The pathology associated with this disease includes necrosis and ulceration of the epidermis, dermis and underlying musculature, and necrosis in hematopoietic tissue, spleen, intestine, and heart. Electron microscopy of affected cells revealed cornaviruslike particles. The virus produce CPE on EPC cells when cultured at 25◦ C, but no CPE on EK-1 (eel kidney), FHM, CHSE-214, or RTG2 cells. Cell cultured virus was ether sensitive, replicated in the presence of IUdR (suggesting that it was enveloped and contained an RNA genome), and viral particles appeared in cytoplasm vacuoles with corona virus morphology. Cell culture propagated virus caused ulcerative dermatitis in challenge trials with common carp. The geographic range, management and importance of this disease, termed carp viremia associated ana-aki-byo (CVA), was not expounded upon.

Fathead minnow rhabdovirus Iwanowicz and Goodwin (2002) reported on the isolation of a rhabdovirus from diseased fathead minnows. This agent, termed fathead minnow rhabdovirus (FHMRV), produced extensive CPE on EPC and RTG-2 cells at 10–25◦ C. Of the other cell lines evaluated, FHM, CCO, and BB cells were susceptible, but CHSE-214 and BF-2 cells were resistant. Immunodot evaluation indicated that FHMRV was not recognized by SVCV-, PFR-,

or VHSV-specific antibodies. In challenge trials, FHMs were susceptible to FHMRV but channel catfish, carp, goldfish, and golden shiners were resistant. Infected fathead minnows swam erratically in circles early in the infection, then later became listless but would swim nervously when startled. Lesions included petechial hemorrhages in the eyes, skin, muscle, and necrosis in the liver spleen and anterior kidney.

Management of cyprinid viruses Management of viral diseases of carp and minnows include best aquaculture practices, avoidance of the viruses, and vaccination. Proactive aquaculture practices include maintaining environment free of stressful water quality conditions, avoiding adverse temperature where possible, maintaining appropriate population densities, and providing appropriate feeds. Spring viremia of carp is best prevented and controlled by quarantining infected fish from healthy populations. Disinfection of holding facilities between crops of fish prevents transmission to naive fish later placed in those units. It has been a goal of fish farmers to prevent the spread of SVCV by detecting virus carriers and eliminating them as brood stock. Since isolation of SVCV from carriers is not possible, serological detection to indicate prior exposure has been pursued. A competitive immunoassay to detect SVCV antibodies in fish was developed by Dixon et al. (1994), who detected SVCV-positive antibody in 88.5% of a carp population known to have been exposed to the virus compared to 34.6% positive detection by serum neutralization. In testing sera from three other groups, 8–12% were positive by the competitive immunoassay method verses 0% positive by serum neutralization. The competitive assay has potential for use in large scale screening of fish populations. Also, Koutina´ et al. (2003) described a PCR procedure for detection of SVCV in carp.

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Proactive vaccination for SVCV may be beneficial when practical. Fish exposed to inactivated SVCV produce antibodies that confer varying degrees of resistance during subsequent virus exposures. Carp were first experimentally vaccinated by immersion and hyperimmunization for SVCV by Fijan et al. (1977a); however, immune response was measured by serum neutralization only. Fijan et al. (1977b) reported that protective immunity in carp was obtained for 9 months after vaccination by intraperitoneal injection with a viable SVCV isolate produced on human diploid and FHM cells. Vaccines made of viral DNA were shown to be efficacious against SVC (Kanellos et al. 2006). This vaccine consisted of a purified plasmid DNA containing the SVC glycoprotein gene driven by the human cytomegalovirus immediate early promoter and was delivered by intramuscular injection. While being safe and efficacious, the DNA vaccines have not been applied in an extensive commercial aquaculture setting because of the requirement for individual fish application. Immunity against spring viremia of carp is temperature dependent. Fish vaccinated by intraperitoneal injection develop protective immunity at 20–22◦ C but not at 10◦ C. Successful mass carp vaccinations by injection have been reported; however, vaccination must be done when water temperatures are 18◦ C or above. Mass vaccination for SVC remains problematical. The herpesvirus diseases present a little different scenario in that carrier fish are often difficult to identify. Avoidance of the herpesvirus status of carp is the best approach by knowing the disease history of the population from which one obtains fish. This is especially true with carp pox (CyHV-1) because of the long incubation period between exposure to the virus and appearance of the tumors. Much of the effort in controlling KHV outbreaks has focused on avoidance. Most production facilities have opted to use a closed system of common carp brooder fish with a

known history of being reared in KHV-free systems. Koi breeders and hobbyists are encouraged to obtain fish from reputable sources and avoid situations where koi from different locations are mixed. The development of virusspecific PCR and other molecular procedures may be used to detect carrier fish of some carp viruses to aid avoiding their spread and should be employed whenever practical. Waltzek et al. (2009) described an accurate method of detecting HVHN (CyHV-2) that can be used to identify populations of goldfish that are positive for this virus. There are three reports of potential vaccines for KHV. The first was essentially exposing carp to wild type virus at permissive temperature followed by raising the temperature to nonpermissive temperature. The second employees an attenuated KHV strain that was produced by cell culture passage and UV exposure (Ronen et al. 2003). The third is a recombinant live virus modified as a bacterial artificial chromosome by deleting the thymidine kinase gene (Costes et al. 2008). Of these, only the naturally attenuated vaccine is patented and has been marketed. There is no known control for CHV (CyHV-1) and the only management practice is avoidance. Carp shipped from CHV-infected areas into regions where it has not been reported should be carefully selected from stocks historically free of the disease because virus is likely transferred from one geographical area to another in covertly infected fish. Since tumor development follows long incubation, it is advisable to quarantine newly transported carp for 1 year before they are released into communal waters. Proper management of golden shiner virus is based on avoidance of overcrowding in conjunction with treatment of external bacterial infections. Vaccination against grass carp reovirus by injection has been practiced in China using two vaccine preparations; one is made by extracting virus antigen from visceral organs of moribund and dead fish and the second is virus

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grown in tissue culture. Both are formalininactivated and juvenile fish are injected intraperitoneal with these preparations. In laboratory studies, Zhang et al. (1990) achieved an average of 68% survival in fish vaccinated by immersion with virus plus adjuvant (scopolamine) verses 0% survival of nonvaccinated controls following challenge. They also acquired 85% survival verses 40% survival in vaccinated and control fish groups, respectively, under field conditions. Use of GCHV vaccine, along with improved fish culture environmental conditions, has been accredited with reduced GCHV severity in cultured grass carp in China.

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colorcarp juveniles showing edemas. Fish Pathology 32:81–88. Oyamatsu, T., H. Matoyaka, et al. 1997b. A trial for the detection of carp edema virus by using polymerase chain reaction. Suisanzoshoku 45:247–251. Perelberg, A., M. Smirnov, et al. 2003. Epidemiological description of a new viral disease afflicting cultured Cyprinus carpio in Israel. Israeli Journal of Aquaculture (Bamidge) 55:5–12. Pikarsky, E., A. Ronen, et al. 2004. Pathogenesis of acute viral disease induced in fish by carp interstitial nephritis and gill necrosis virus. Journal of Virology 78:9544–9551. Plumb, J. A., P. R. Bowser, et al. 1979. Fish viruses: a double-stranded RNA icosahedral virus from a North American cyprinid. Journal of the Fisheries Research Board of Canada 36:1390–1394. Ronen, A., A. Perelberg, et al. 2003. Efficient vaccine against the virus causing a lethal disease in cultured Cyprinus carpio. Vaccine 21(32):4677–4684. Sadler, J., E. Marecaux, and A. E. Goodwin. 2008. Detection of koi herpes virus (CyHV-3) in goldfish, Carassius auratus (L.), exposed to infected koi. Journal of Fish Diseases 31:71–72. Sanders, G. E., W. N. Batts and J. R. Winton. 2003. Susceptibility of zebrafish (Danio rerio) to a model pathogen, spring viremia of carp virus. Comparative Medicine 53(5):514–521. Sano, T., H. Fukuda, and M. Furukawa. 1985a. Herpesvirus cyprini: Biological and oncogenic properties. Fish Pathology 20:381–388. Sano, T., H. Fukuda, et al. 1985b. A herpesvirus isolated from carp papilloma in Japan. In: A. E. Ellis (ed.) Fish and Shellfish Pathology. Orlando. Academic Press, pp. 307–311. Sano, N., R. Honda, et al. 1991a. Herpesvirus cyprini: restriction endonuclease cleavage profiles of the viral DNA. Fish Pathology 26:207–212. Sano, T., N. Morita, et al. 1991b. Herpesvirus cyprini: lethality and oncogenicity. Journal of Fish Diseases 14:533–543. Sano, M., T. Ito, et al. 2004. First detection of koi herpesvirus in cultured common carp Cyprinus carpio in Japan. Fish Pathology 39:165–168. Sano, N., M. Moriwake, and T. Sano. 1993a. Herpesvirus cyprinid: thermal effects on pathogenicity and oncogenicity. Fish Pathology 28, 171–175.

Sano, N., M. Moriwake, et al. 1993b. Herpesvirus cyprini: a search for viral genome in infected fish by in situ hybridization. Journal of Fish Diseases 16:495–499. Sano, N., M. Sano, et al. 1992. Herpesvirus cyprini: detection of the viral genome in situ hybridization, Journal of Fish Diseases 15:153– 162. Schubert, G. 1966. The infective agent in carp pox. Bulletin Office International des Epizooties 65:1011–1022. Schwedler, T. E., and J. A. Plumb. 1980. Fish viruses: serologic comparison of the golden shiner and infectious pancreatic necrosis viruses. Journal of Wildlife Diseases 16:597–599. Schwedler, T. E., and J. A. Plumb. 1982. Golden shiner virus: effects of stocking density on incidence of viral infection. The Progressive FishCulturist 44:151–152. Shao, J., S. Mao, and Y. Shen. 1990. Studies on the isolation and pathogenecity of two kinds of hemorrhagic virus of grass carp, Ctenopharynogodon idellus. Journal of Hangzhou University 17:74–79 (Chinese with English abstract). Shchelkunov, I. S., and T. I. Shchelkunova. 1989. Rhabdovirus carpio in herbivorous fishes: Isolation, pathology, and comparative susceptibility of fishes. In: W. Ahne and E. Kurstake (eds) Viruses of Lower Vertebrates. Berlin, SpringerVerlag, pp. 333–348. Shen, L. Z. 1978. Studies on the causative agent of hemorrhage of the grass carp (Ctenopharyngdon idellus). Acta Hydrobiologica Sinica 6:322–329 (Chinese with English summary). Shivappa, R. B., R. Savan, et al. 2008. Detection of spring viraemia of carp virus (SVCV) by loopmediated isothermal amplification (LAMP) in koi carp, Cyprinus carpio L. Journal of Fish Diseases 31:249–258. Sonstegard, R. A., and K. S. Sonstegard. 1978. Herpesvirus-associated epidermal hyperplasia in fish (carp). In: G. de The, W. Henle and F. Rapp (eds) Proceedings International Symposium on Oncogenic Herpesvirus. Lyon, France International Aqency Research for Cancer Science, Scientific Publication No. 24, pp. 863–868. Stephens, F. J., S. R. Raidal, et al. 2004. Haematopoietic necrosis in a goldfish (Carassius auratus) associated with an agent morphologically similar to herpesvirus. Australian Veterinary Journal 82:167–169.

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Stone, D. M., W. Ahne, et al. 2003. Nucleotide sequence analysis of the glycoprotein gene of putative spring viraemia of carp virus and pike fry rhabdovirus isolates reveals four genogroups. Diseases of Aquatic Organisms 53(3):203–210. St-Hilaire, S., N. Beevers, et al. 2005. Reactivation of koi herpesvirus infections in common carp Cyprinus carpio. Diseases of Aquatic Organisms 67:15–23. Tu, C., M. C. Weng, et al. 2004. Detection of koi herpesvirus in koi Cyprinus carpio in Taiwan. Fish Pathology 39:109–110. USFWS (U.S. Fish and Wildlife Service). 2007. Carp virus discovered in Upper Mississippi River. Available at: http://www.fws.gov/news/ newsreleases/showNews.cfm?newsId=792D5D09ACC1-10D1-06E4CCB7618DD470. Varovic, K., and N. Fijan. 1973. Osjetljivost sarana prema rhabdovirus carpio pri raznim nacinima inokulacije. Veterinary Arhives 43:271–276. (English summary). Waltzek, T. B., G. O. Kelley, et al. 2005. Koi herpesvirus represents a third cyprinid herpesvirus (CyHV-3) in the family Herpesviridae. Journal of General Virology 86:1659–1667. Waltzek, T. B., T. Kurobe, et al. 2009. Development of a polymerase chain reaction assay to

detect cyprinid herpesvirus 2 in goldfish. Journal of Aquatic Animal Health 21:60–67. Wang, T., H. Chen, et al. 1994. Preliminary studies on the susceptibility of Gobiocypris rarus to hemorrhagic virus of grass carp. Acta Hydrobiologia Sinica 18:144–149 (Chinese with English abstract) Warg, J. V., A. L. Dikkeboom, et al. 2007. Comparison of multiple genes of spring viremia of carp viruses isolated in the United States. Virus Genes 35:87–95. Way, K. 1991. Rapid detection of SVC virus antigen in infected cell cultures and clinically diseased carp by enzyme-linked immunosorbent assay (ELISA). Journal of Applied Ichthyology. 7:95–107. Winton, J. R., C. N. Lannan, et al. 1987. Morphological and biochemical properties of four members of a novel group of reoviruses isolated from aquatic animals. Journal of General Virology 68:353–364. Wolf, K. 1988. Fish Viruses and Fish Viral Diseases. Ithaca, Cornell University Press. Zhang, N., G. Yang, et al. 1990. Scopolamine enhances the immunity of carp using immersion vaccine. Journal of Zhejiang College of Fisheries 9:79–83 (Chinese).

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Chapter 7

Eel viruses

Viruses have been reported in true eels (genus Anguilla; European eel, American eel, Japanese eel, and New Zealand longfin eel). Isolates are mainly from Europe and Japan and occasionally from North America and elsewhere. Virus impact on cultured eel populations is difficult to assess because many isolates have come from apparently normal fish. Also numerous reported viruses from eels are poorly classified (Ahne et al. 1987). Six viruses affect eels are discussed: eel virus European (EVE), eel rhabdovirus (multiple designations), anguillid herpesvirus, stomatopapilloma, eel iridovirus (EV-102), and viral endothelial cell necrosis.

Eel virus European EVE was isolated from cultured European eels in Japan (Sano 1976). Later it was suggested that the disease should be called viral kidney disease because of how severely the virus affected that particular organ (Sano et al. 1981). Also, a gill necrosis disease in Japanese eels

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.

was associated with a birnavirus that was designated pillar cell necrosis Virus (PCNV) (Lee et al. 1999a). Both isolates were shown to be related to infectious pancreatic necrosis virus Genogroup II (Lee et al. 2001; Nishizawa et al. 2005).

Geographical range and species susceptibility While EVE was initially isolated from European eels in Japan, it was also found in Japanese eels and tilapia in Taiwan (Sano 1976; Chen et al. 1985). Viruses similar to EVE, characterized as IPNV strain Ab, were isolated from normal young eels in England (Hudson et al. 1981) and from moribund cultured American eels in Florida (USA) in 1988 (J. A. Plumb, unpublished, Serotyped by B. Hill, Disease Laboratory, Weymouth, England, personal communication). Ueno et al. (1983) determined that an EVE isolate from Japanese eels was not infectious to common carp at 20◦ –25◦ C, but tilapia were severely affected at 10–16◦ C, temperatures that approach being stressful or lethal limits for tilapia. However, EVE has low virulence for experimentally infected young rainbow trout (Sano et al. 1981). 135

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Clinical signs Eels experimentally infected with EVE become rigid with muscle spasms and congested anal fins (Sano 1976; Ueno et al. 1983). A microscopic gill examination show marked hyperplasia of the lamellar epithelium, and lamellae fusing and clubbing of filaments, giving them a swollen appearance. The underside of affected fish show some petechia. Ascitic fluid can be present in the abdominal cavity, the alimentary tract is void of food, and the kidney hypertrophied. Japanese eels with PCNV infections demonstrated high numbers of telangiatasis (aneurisms) of secondary lamellae in the gills caused by necrosis of pillar cells and proliferation of intralamellar epithelium. Also many fish display necrosis of gastric gland cells in the stomach (Lee et al. 1999a).

Diagnosis EVE is isolated from internal organs of affected fish in BF-2, CHSE-214 or RTG-2 cells incubated at 10 to 20◦ C, however, cell lines from eels should be equally satisfactory (Chen and Kou 1981). Sano (1976) noted that CPE of EVE is characterized by nuclear pyknosis and cells retain an elongated shape almost identical to CPE caused by IPN virus. PCNV also showed a wide range of cell susceptibility with FHM, eel kidney EK-1 and eel ovary (EO-2) cells being most susceptible and CHSE-214, RTG-2,and EPC being less susceptible (Lee et al. 1999a).

Virus characteristics The EVE and PCNV are birnaviruses with biophysical characteristics (size, morphology, etc.) similar to IPNV-strain Ab. They have a genome composed of two dsRNA fragments and small naked icosahedral capsids. EVE is closely related serologically to the IPNVAb strain (Hedrick et al. 1983). Okomoto

et al. (1983) compared ten strains of IPNV to EVE and found the eel virus to be more closely related to the IPNV-Ab serotype than to any other viral strain. The similarity of EVE to other aquatic birnavirus isolates from Japanese eels affected with branchionephritis using monoclonal antibody (MAB) was made by Chi et al. (1991). Three of six MABs identified epitopes possessed by IPN-Ab virus only; one recognized an epitope on both the Ab and Sp serotypes and two MABs exhibited epitopes on Ab, Sp, and VR-299 IPNV serotypes. PCNV was serologically most similar to IPNV SP (Lee et al. 1999a) but sequence analysis of the VP2-NS junction of the genomic RNA demonstrated that it was in genogroup II and most similar to IPNV strain Ab.

Epizootiology Cultured young European and Japanese eels appear to be most vulnerable to EVE. The virus has a propensity for producing disease at temperatures from 8 to 14◦ C, especially in small eels (Sano 1976). Mortality due to EVE may be as high as 60% in 4–26 g eels and Sano et al. (1981) reported 55–75% mortality of experimentally infected 9–12 g Japanese eels. EVE can be transmitted by waterborne exposure and fish to fish transmission seems to be a likely possibility. It was postulated that virus was transmitted from Europe to Japan in elvers and then to Taiwan in eyed trout eggs (Wolf 1988). Japanese eels are also affected by a gill disease commonly called branchionephritis, from which EVE was isolated; however, the virus did not cause this condition in experimentally infected fish (Ueno et al. 1983). But in 1987, a severe gill disease characterized by lamellar aneurisms (telangiatasis) caused high losses in Japanese eels in warm-water ponds in western Japan; this agent, designated PCNV, reproduced the disease when injected intramuscularly just posterior to the head when the fish were held at 25◦ C.

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Pathology

Geographical range and species susceptibility

European eels with EVE develop congestion in the fins, gills and occasionally the abdomen (Sano 1976). Gills display marked hyperplasia of the lamellar epithelium and lamellar fusion. Histopathological findings show proliferative glomerulonephritis and necrosis in the renal interstitial nephrons, hyaline droplet degeneration, and cloudy swelling in the tubular epithelial cells. Focal necrosis occurs in the liver and extensive necrosis occurs in the spleen of some eels (Egusa et al. 1971).

Significance Significance of EVE and PCNV in cultured eels is not clear; however, both have been associated with high mortality in cultured eels in Japan and Taiwan.

Eel Rhabdoviruses A large number of rhabdoviruses in the Novirhabdovirus and Vesiculovirus genera have been isolated from cultured eels (Castric et al. 1984) but seldom caused significant disease (Table 7.1).

The eel vesiculoviruses appear to be widely distributed. A rhabdovirus, designated eel virus America (EVA), was isolated from American eels imported into Japan from Cuba (Sano 1976). A rhabdovirus isolated from European Eels in Japan was designated eel virus European - X (EVEX) (Sano et al. 1977). Castric et al. (1984) identified five rhabdoviruses in eels in Europe including two novirhabdoviruses. Later a third novirhabdovirus and viral hemorrhagic septicemia virus (VHSV), normally associated with salmonids but also isolated from a variety of other fish species, was found in European eels in France (Castric et al. 1992; Thiery et al. 2002). In a survey of eels from various geographic regions, vesiculovirus (EVEX) was isolated from European eels in Italy, The Netherlands, and Morocco and from New Zealand longfin eel (A. dieffenbachia) in New Zealand (van Ginneken et al. 2004). The host ranges of these viruses have been poorly defined. In experimental challenges, E6, EVA, and EVEX were shown to be pathogenic to rainbow trout (Castric and Chastel 1980; Nishimura et al. 1981). The VHS isolate from eel was shown to be pathogenic to

Table 7.1 Rhabdoviruses isolated from eels (Anguilla spp.). Rhabdovirus Group

Isolate Designation

Eel Species

Geographical Origin

Novirhabdovirus∗

B12 C26 VHSV L59X Eel virus American (EVA) Eel virus European – X (EVEX) C30 B44 D13

A. anguilla A. anguilla A. anguilla A. rostrata†

Europe Europe Europe Japan

A. anguilla‡

Japan

A. anguilla A. anguilla A. anguilla

Europe Europe Europe

Vesiculovirus

Source: Castrick and Chastil (1980) and Castric et al. (1984). ∗ Originally designated by Lyssavirus group. † From elvers from Cuba. ‡ From elvers from Europe.

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rainbow trout by injection challenge (Castric et al. 1992).

Clinical signs The eel novirhaboviruses were isolated from normal, apparently healthy elvers; there are few clinical signs associated with infection. There are several accounts of disease outbreaks with the eel vesiculoviruses. Sano (1976) reported that naturally EVA-infected American eel elvers imported to Japan from Cuba displayed hemorrhages and congestion on the body’s underside and on anal and pectoral fins. Fish also turned their heads downward and gills showed hemorrhage and degradation of bone or cartilage. A disease called rhabdoviral dermatitis, described by Kobayashi and Miyazaki (1996), produces cutaneous lesions and mild histopathology of internal organs.

Diagnosis For isolation of eel rhabdoviruses, filtrates from homogenates of brain, kidney, liver, and spleen tissue are inoculated onto BF-2, eel kidney (EK-1), eel ovary (EO-2), FHM, or RTG-2 cells and incubated at 15 or 20◦ C depending upon cell line (Wolf 1988). Cytopathic effect consists of pyknosis and cytoplasmic granulation followed by cell lysis. In a comparison of CPE of EVA and eel virus European (unknown)-X, Nishimura et al. (1981) showed that effects were similar on RTG-2 cells and CPE was also similar to that of IHNV.

Virus characteristics At least eight different isolates of rhabdoviruses have been made from eels and are designated as eel virus American (EVA), eel virus European – X (EVEX), B12 , B44 , C26 , C30 , and D13 (Table 7.1) (Wolf 1988). These

viruses, several of which came from apparently healthy eels, were analyzed by Castric et al. (1984); two of the viruses belonged to the Lyssavirus genus (later reclassified to the Novirhabovirus genus) (B12 and C26 ) and five were members of the Vesiculovirus genus (EVA, EVEX, C30 , B44 , and D13 ). The rhabdovirus reported by Kobayashi and Miyazaki (1996) was similar to EVA and EVEX. Later VHSV was isolated from wild eels in Loire estuary in France and was determined to belong to genotype II typical of marine VHSV isolates (Castric et al. 1992; Thiery et al. 2002)

Epizootiology Of all the viruses isolated from eel, the rhabdoviruses may be of greatest interest to fish virologists because they seem to be very prevalent in some established wild populations of European eels but do not appear to cause any significant problem among cultured populations. However, Castric and Chastel (1980) showed that eel rhabdovirus strain B6 was experimentally infectious to rainbow trout fry by bath exposure but was not pathogenic for elvers (young eels). Strain B12 , C30 , and EVEX produced no significant mortality in either fish species. The American eel from Cuba infected with EVA suffered high mortality at 20–27◦ C (Sano 1976). Infectivity trials of EVA and EVEX in rainbow trout, Japanese eel, common carp, and ayu resulted in the death of only rainbow trout (Nishimura et al. 1981). Infectivity in trout was more severe at 15 and 20◦ C than 10◦ C, which led investigators to be concerned with the possibility of introducing another trout pathogen into Japan. Natural outbreaks of rhabdoviral dermatitis in Japanese aquaculture facilities occurred primarily when Japanese eels were held at cool temperatures prior to shipment. In challenge trials, Kobayashi et al. (1999) demonstrated temperature dependence for disease development. Although the virus replicated well in fish

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Eel Viruses 139

at 25◦ C, when fish were challenged by subcutaneous inoculation at 15, 20, and 25◦ C, only the fish held at 15◦ C showed substantial morbidity and cutaneous lesions. Similarly, Shchelkunov et al. (1989) demonstrated that an eel vesiculovirus isolate was pathogenic to 4-year-old European eels when held at 10–13.5◦ C. The infected fish developed hemorrhages on the fins, eyes, and mouth.

mia, hemorrhages, bloody ascites, and stopped swimming before 1,500 km. Uninfected fish developed increased hematocrits, no-lesions, and swam over 5,500 km (van Ginneken et al. 2005). They indicated that eel vesiculovirus was the most prevalent virus in wild populations, postulating that the effect on endurance in migrating adults could be at least partially responsible for declining eel populations.

Pathology

Anguillid herpesvirus

American eels with EVA from Cuba had hemorrhaging and necrosis of the branchial vessels, skeletal muscle, and kidneys (Sano 1976). European eel infected with rhabdovirus showed no pathology. Experimental EVA and EVEX infections in rainbow trout produced hemorrhagic skeletal muscle, intensive necrosis, and hemorrhage of the kidneys hematopoietic tissue, and focal necrosis of the liver, spleen, and pancreas (Nishimura et al. 1981). Histopathology was comparable to that associated with IHNV infections in salmonids. The rhabdoviral dermatitis-infected eel demonstrated dermal lesions with necrosis of the dermal fibrocytes, hemorrhage, and inflammatory cellular infiltration. They also displayed diffuse necrosis of the spleen liver and hematopoietic tissue (Kobayashi and Miyazaki 1996; Kobayashi et al. 1999).

Sano et al. (1990) reported a herpesvirus isolated from diseased cultured Japanese and European eels at two locations in Japan. They designated the virus Anguillid herpesvirus 1 and Herpesvirus anguillae (HVA) as the Latinized name; however, the common name is “eel herpesvirus”. Ueno et al. (1992) isolated a herpesvirus, designated eel herpesvirus in Formosa (EHVF), from cultured Japanese eel in Taiwan. Herpesvirus anguillae and EHVF are structurally and serologically very similar (Ueno et al. 1996). A herpesvirus was also found in skin lesions of European eels in Europe by electron microscopy (B´ek´esi et al. 1986) and a herpesvirus was isolated from healthy eel in France (Jørgensen et al. 1994). Eventually the herpesvirus was isolated in cell culture from a disease outbreak and was characterized (Davidse et al. 1999); the virus was implicated in at least 19 outbreaks on Dutch European eel aquaculture facilities (Haenen et al. 2002). DNA analysis of the European and Japanese isolates indicate they are very similar and are likely to be the same species (Haenen et al. 2002; Rijsewijk et al. 2005).

Significance In aquaculture, eel dermatitis is primarily a problem when eels are being prepared for shipping and held at suboptimum temperatures (Kobayashi et al. 1999). The effect of eel vesiculoviruses on the health of wild eels may be significant. In an experiment comparing survival and endurance of a population of eels naturally infected with eel vesiculovirus to specific pathogen-free eels in a marine swim tunnel (to simulate a 5,500 km spawning migration), all of the infected fish developed ane-

Geographical range and species susceptibility Anguillid herpesvirus has been confirmed in cultured Japanese and European eels in Japan, Taiwan, Italy, France, and The Netherlands and in wild eels from The Netherlands; the

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virus is thought to be widespread in Europe (van Ginneken et al. 2004). In challenge trials of Japanese eels with HVA, Kobayashi and Myazaki (1997) and Lee et al. (1999b) demonstrated reproduction of the disease in natural outbreaks of HVA, while Hangalapura et al. (2007) fulfilled Koch’s postulates using a European isolate in immersion challenges of European eels. Natural outbreaks of HVA have not been reported in other species of fish. However, the finding of Ueno et al. (1992) that common carp were sensitive to the Taiwanese isolate in injection trials and the ability to infect non-eel cell lines suggest that the virus may not be restricted to Anguillids.

for HVA is 20–25◦ C, but replication occurs from 10 to 37◦ C (Sano et al. 1990). Presumptive diagnosis can be made using clinical signs and the virus producing characteristic CPE on EK-1 cells. Confirmation requires a positive reaction with HVA-specific antisera or HVAspecific PCR (Ueno et al. 1996; Davidse et al. 1999; Lee et al. 2001; Rijsewijk et al. 2005).

Virus characteristics

Affected eels are generally lethargic and demonstrate petechial diffuse hemorrhages on the skin and fins, ulcerative skin and fin lesions, hemorrhages in the gills, congested gills with necrosis of the connective tissue, and a pale liver. However the clinical signs may vary, with outbreaks being characterized predominantly by skin and gill lesions (Sano et al. 1990; Ueno et al. 1992; Lee et al. 1999b; Haenen et al. 2002).

Anguillid herpesvirus clearly conforms to the herpesvirus criteria while being serologically distinct from channel catfish virus, salmonid herpesvirus type 2, and cyprinid herpesvirus type 1 (Sano et al. 1990). Pulse field electrophoresis reveals a DNA genome of about 245 kb. The DNA sequence analysis of a fragment of the DNA polymerase gene reveals HVA is identical to EHVF in this region and that it may be more similar to cyprinid herpesvirus 1 than to channel catfish virus (Rijsewijk et al. 2005). The icosahedral nucleocapsid measures 110 nm and the mean enveloped virion diameter is 200 nm. Twentyfive polypeptides were identified in the virion with molecular weights of 19.5–320 k (Sano et al. 1990).

Diagnosis

Epizootiology

Anguillid herpesvirus is diagnosed by virus isolation in cell culture. Although HVA produces CPE in BF-2, FHM, and RTG-2 cells, optimum virus replication is in eel kidney cells (EK-l) and eel ovary cells (EO-2) (Sano et al. 1990). Ueno et al. (1992) reported successful culture of the EHVF in black grouper kidney (BGK), colored carp fin (CCF), colored carp fin testis (CCT), tilapia ovary (TO-2), EK-1, and EO-2 cells but no replication in RTG-2, FHM, or BF-2 cells. Cytopathic effect consists of syncytia formation and rounded cells with Cowdry type A intranuclear inclusions. Optimum cell culture incubation temperature

Anguillid herpesvirus was initially isolated from Japanese eels that suffered l% mortality at one aquaculture facility and European eels that suffered 6.8% mortality at a second facility (Sano et al. 1990) where fish were being cultured at high density in recirculating systems at 30◦ C. It was speculated that unfavorable environmental conditions may have caused stress and onset of the herpesvirus infection. Virus was isolated 2 weeks after initial detection but not 1 month later, suggesting progression into a dormant viral state. More extensive losses were reported from 1993 to 1995 in Japan (Lee et al. 1999b) and after 1996 on eel farms in The

Clinical signs

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Eel Viruses 141

Netherlands (Davidse et al. 1999) where the virus persisted in apparently healthy stocks of fish. Virus isolation was improved from carrier fish after dexamethasone treatment and HVAspecific antibodies could be detected in these fish using ELISA (van Nieuwstadt et al. 2001). Using the serological test for prior exposure detection, the researchers suggested “tracing sero-positive eel stock, which had previous contact with HVA and of HVA carrier fish can be useful to control disease outbreaks due to HVA infection.”

Pathology Pathology may vary according to disease manifestation and has been addressed by several researchers (Sano et al. 1990, Kobayashi and Miyazaki 1997; Lee et al. 1999b; Hangalapura et al. 2007). In general the virus appears to have a tropism for fibroblast with focal areas of necrosis occurring in the dermis and connective tissue of the gills. In the skin, pathology was followed by infiltration of the affected regions by inflammatory cells, more extensive necrosis, hemorrhage, and tissue sloughing. Gills displayed extensive lamellar hyperplasia hemorrhaging, congestion of the filaments, and necrosis of the tips of affected filaments. Lesions in the internal organs include mild diffuse necrosis of the liver, spleen, and kidney tissue and hemorrhages in the adipose tissue.

Significance Eel herpesvirus is a moderately important pathogen in eel aquaculture where numerous HVA outbreaks have occurred in Japan, Taiwan, and Europe. Furthermore, the presence of this virus in wild stocks and its ability to remain unapparent or latent in carrier populations suggest that it may be difficult to control the spread of this pathogen.

Stomatopapilloma Stomatopapilloma, also known as cauliflower disease because of lesion morphology, is a tumorous disease of eels that has been recognized since 1910 (Wolf 1988). Several viruses have been associated with these lesions, but their role in tumor production is unclear.

Geographical range and species susceptibility Stomatopapilloma affects only European eel and occurs mainly in tributaries and brackish waters of the Baltic and North Seas of northern Europe (Wolf 1988) but has also been reported in Great Britain (Delves-Broughton et al. 1980).

Clinical Signs In stomatopapilloma chronic, benign epidermal, neoplasms usually develop on the mouth, nares, and head and occasionally on the fins of eels. Lesions may cover the entire head and occlude the oral cavity and occasionally on the fins and other body locations (Peters 1975). The lesions are amorphous, irregular in shape, have a pebbly surface, and are usually light in color but can be pigmented.

Diagnosis Diagnosis of stomatopapilloma is based on morphology of head lesions, histopathology, and electron microscopy. Virus has been isolated from tumors and visceral organ tissue in RTG-2 and FHM cells.

Virus characteristics Ahne et al. (1987) reported that nine virus isolates have been made from European eels

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with stomatopapilloma. The first virus isolation was from blood of affected fish and was named “eel virus (Berlin)” (Pfitzner and Schubert 1969). Characterization of the virus indicates an icosahedral virion with an RNA genome that is morphologically similar to IPNV (Schwanz-Pfitzner et al. 1984). This isolate was most likely IPNV serotype Ab, therefore would be classified as a birnavirus. Wolf and Quimby (1970) isolated a virus from European eel with stomatopapilloma and designated it EV-1. This virus was plaque purified and then designated EV-2 for characterization and was classified as an orthomyxo-like virus (Nagabayashi and Wolf 1979). The 80–140 nm diameter pleomorphic virion has 10 nm surface projections and an RNA genome. The virus replicates in FHM cells at 16–18◦ C, where CPE appears after 10 hours, which consists of cell fusion, syncytia, and irregularly rounded cell masses. Three viruses were isolated from a single European eel with stomatopapilloma by Ahne and Thomsen (1985). These viruses were identified as IPNV-Ab from the blood and Rhabdovirus anguilla-subtype EVEX, and an unclassified agent from the tumor. The viruses produced CPE in FHM cells in 3 days. Ahne et al. (1987) determined that of nine virus isolates taken from one eel and identified by serum neutralization, five from blood were IPNV-subtype Ab, and four from tumors were Rhabdovirus anguilla-subtype EVEX.

pilloma lesions with these viruses were unsuccessful. Stomatopapilloma appears to be a disease primarily of wild eel populations with no specificity for age or size. The number of affected eels nearly doubled between 1957 and 1967 from 5.6% to 11.9% in the Elbe River in Germany (Koops et al. 1970). Tumors were found on 15–35 cm eels in the III–V year age group in fresh water in the Lower Elbe River (Peters 1975). Incidence of tumors was low (2–3%) in spring and autumn compared to summer months when incidence increased to about 28%. A similar seasonal incidence was found in Scottish rivers, where the disease was not known until 1980 (Hussein and Mills 1982). While not totally eliminating a viral etiology, Peters (1975) speculated that seasonal fluctuations in tumor growth were due to water temperature and oxygen content, but conceded that chemical pollutants could also be a factor. No such water quality correlations were found by Hussein and Mills (1982) in comparing incidence of stomatopapilloma in four tributaries of the River Tweed in Scotland. Stomatopapilloma probably does not kill eels directly, but massive head tumors obstructing the oral cavity can reduce feeding, thus causing as much as a 50% loss in their fat content (Koops et al. 1970).

Pathology Epizootiology Stomatopapilloma of European eel has been of interest to fishery scientists for years; however, the disease is still not fully understood and its etiology remains unresolved. The fact that at least nine different viruses isolated from one infected fish does not necessarily indicate that any were responsible for the tumors (Ahne et al. 1987). As noted, viruses have been isolated from blood, visceral organs, and tumor tissue, but experiments to induce stomatopa-

Stomatopapilloma lesions are hyperplastic skin epithelium that may or may not be pigmented. The resultant tumor consists of fibroepithelium most commonly seen on the upper and lower jaw and occasionally on fins and lateral areas (Koops et al. 1970). Pathological changes begin at the basal layer of the epidermis followed by transformation of cuboidal cells into elongate cells with spindleshaped nuclei. Papillomas are smaller (up to 300 mm3 ) in cooler weather and larger in August/September when they may reach 900 mm3 and show signs of invasiveness (Peters 1975).

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Significance Currently stomatopapilloma is of little concern to the eel culture industry. Its presence does not cause large mortality but could have an adverse economic effect on marketability of cultured eels.

Other virus diseases of eels Two other virus diseases have been reported from cultured eel. One is an iridovirus (eel iridovirus, EV-102) that has been reported once (Sorimachi 1984) and the other is an adenovirus named viral endothelial cell necrosis of eel (VECNE) (Ono et al. 2007).

Eel Iridovirus The Japanese eel iridovirus (EV-102) was isolated from cultured eels in Japan during a routine farm survey (Sorimachi 1984). Japanese eel iridovirus is known only in Japan, where it is pathogenic to Japanese eel, but not European eel. Experimental infection of Japanese eels with EV-102, produced skin depigmentation, fin congestion, and increased skin mucus in 3–5 days postinfection (Wolf 1988). Japanese eel iridovirus replicates in EPC and RTG-2 cells as well as EK-l and EO-2 cells when incubated at 15–20◦ C (Wolf 1988). Biophysical and biochemical characteristics of eel iridovirus are typical of the group. It is an icosahedral virus that measures 100–200 nm in diameter containing a DNA genome (Sorimachi and Egusa 1982). After experimentally infecting eels up to 120 g with EV-102 by immersion or injection, clinical signs appeared in 3–5 days at 15–23◦ C (Sorimachi 1984). Mortalities began shortly after onset of clinical signs at about 1 week postexposure and continued for 2 weeks. Deaths stopped and clinical signs abated when water temperature reached 23◦ C, but mortal-

ity was 70% at that point. Pathology associated with EV-102 is unknown and the importance of this virus appears to be minimal.

Eel adenovirus An adenovirus in cultured Japanese eel in Japan produced a disease named viral endothelial cell necrosis virus (VECNE) (Ono et al. 2007). Clinical signs are reddening of the fins and swollen abdomen. Gills are congested in the central venous sinus of gill lamellae. Internally the liver and intestine are congested with hemorrhaged areas. The virus was isolated in a newly established vascular endothelial cell line (Japanese eel endothelial cells, JEEC), where it produced hypertrophied nuclei in 7 days when incubated at 25o C. The virus can be isolated from gills, liver, and kidney. Icosahedral virus particles of about 75 nm in diameter with no envelope were observed. The virus was classified as an adenovirus that was tolerant to chloroform and ph 3.0. In experimental infections, cumulative mortality was about 60%. Little is known about the epizootiology of VECNE.

Management of Eel Viruses There is no known management procedure specifically for prevention and/or control of eel viruses; however, avoidance whenever possible is advisable. Also, for viruses that are more virulent at low temperatures increasing water temperature to >23◦ C may be beneficial.

References Ahne, W., I. Schwanz-Pfitzner, and I. Thomsen. 1987. Serological identification of 9 viral isolates from European eels (Anguilla anguilla) with stomatopapilloma by means of neutralization tests. Journal of Applied Ichthyology 3:30–32. Ahne, W., and I. Thomson. 1985. The existence of three different viral agents in a tumor

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bearing European eel (Anguilla anguilla). Journal of Veterinary Medicine 32:228–235. B´ek´esi, L. I., I. Horvath, et al. 1986. Demonstration of herpesvirus-like particles in skin lesions of the European eel (Anguilla anguilla). Ichthyology 4:190–192. Castric, J., and C. Chastel. 1980. Isolation and characterization attempts of three viruses from European eel, Anguilla anguilla; preliminary results. Annals in Virology 13E:435–448. Castric, J., J. Jeffroy, et al. 1992. Isolation of viral haemorrhagic septicaemia virus (VHSV) from wild elvers Anguilla anguilla. Bulletin of the European Association of Fish Pathologists 12:21–23. Castric, J., D. Rasschaert, and J. Bernard. 1984. Evidence of Lyssaviruses among rhabdovirus isolates from the European eel Anguilla anguilla. Annals in Virology 135E:35–55. 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., G. H. Kou, et al. 1985. The occurrence of viral infections of fish in Taiwan. In: A. Ellis (ed.) Fish and Shellfish Pathology. London, Academic Press, pp. 313–319. Chi, S. C., S. N. Chen, and G. H. Kou. 1991. Establishment, characterization and application of monoclonal antibodies against eel virus European (EVE). Fish Pathology 26:1–7. Davidse A., O. L. M. Haenen, et al. 1999. First isolation of herpesvirus of eel (Herpesvirus anguillae) in diseased European eel (Anguilla anguilla) in Europe. Bulletin of the European Association of Fish Pathologists 19(4):137–141. Delves-Broughton, J., J. K. Fawell, and D. Woods. 1980. The first occurrence of “cauliflower disease” of eels Anguilla anguilla L. in the British Isles. Journal of Fish Diseases 3:255–256. Egusa, S., H. Hirose, and H. Wakabayashi. 1971. A report of investigations on branchionephritis of cultured eel II. Conditions of the gills and serum ion concentrations. Fish Pathology 6:57–61. Haenen, O., S. Dijkstra, et al. 2002. Herpesvirus anguillae (HVA) isolations from disease outbreaks in cultured European eel, Anguilla anguilla in The Netherlands since 1996. Bulletin of the European Association of Fish Pathologists 22(4):247–257. Hangalapura, B. N., R. Zwart, et al. 2007. Pathogenesis of Herpesvirus anguillae (HVA) in juvenile European eel Anguilla anguilla after in-

fection by bath immersion. Diseases of Aquatic Organisms 78(1):13–22. Hedrick, R. P., J. L. Fryer, et al. 1983. Characteristics of four birnaviruses isolated from fish in Taiwan. Fish Pathology 18:91–97. Hudson, E. B., D. Bucke, and A. Forrest. 1981. Isolation of infectious pancreatic necrosis virus from eels, Anguilla anguilla in the United Kingdom. Journal of Fish Diseases 4:429–431. Hussein, S. A., and D. H. Mills. 1982. The prevalence of “cauliflower disease” of eel, Anguilla anguilla L., in tributaries of the River Tweed, Scotland. Journal of Fish Diseases 5:161– 165. Jørgensen, P., J. Castric, et al. 1994. The occurrence of virus-infections in elvers and eels (Anguilla anguilla) in Europe with particular reference to VHSV and IHNV. Aquaculture 123:11–19. Kobayashi, T., and T. Miyazaki. 1997. Characterization and Pathogenicity of a herpesvirus isolated from cutaneous lesion in Japanese eel, Anguilla japonica. Fish Pathology 32(2):89–95. Kobayashi, T., and T. Miyazaki. 1996. Rhabdoviral dermatitis in Japanese eel, Anguilla japonica. Fish Pathology 31:183–190. Kobayashi, T., T. Shiino, and T. Miyazaki. 1999. The effect of water temperature on rhabdoviral dermatitis in the Japanese eel, Anguilla japonica Temminck and Schlegel. Aquaculture 170(1):7–15. Koops, H., H. Mann, et al. 1970. The cauliflower disease of eels. In: S. F. Snieszko (ed.) Diseases of Fishes and Shellfishes. American Fisheries Society, Special Publication No. 5:, pp. 91–295. Lee, N. S., Y. Nomura, and T. Miyazaki. 1999a. Gill lamellar pillar cell necrosis, a new birnavirus disease in Japanese eels. Disease of Aquatic Organisms 37(1):13–21. Lee, N-S., J. Kobayashi, and T. Miyazaki, 1999b. Gill filament necrosis in farmed Japanese eels, Anguilla japonica (Temminck & Schlegel), infected with Herpesvirus anguillae. Journal of Fish Diseases 22(6):457–463. Lee, N-S., M. Miyata, and T. Miyazaki. 2001. Genome sequence of a VP2/NS junction region of pillar cell necrosis virus (PCNV) in cultured Japanese eel Anguilla japonica. Diseases of Aquatic Organisms 44(3):179–182. Nagabayashi, T., and K. Wolf. 1979. Characterization of EV-2, a virus isolated from European eels (Anguilla anguilla) with stomatopapilloma. Journal of Virology 30:358–364.

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Nieuwstadt, A. P., S. G. Dijkstra, and O. L. M. Haenen. 2001. Persistence of herpesvirus of eel Herpesvirus anguillae in farmed European eel Anguilla anguilla. Diseases of Aquatic Organisms 45:103–107. Nishimura, T., M. Toba, et al. 1981. Eel rhabdovirus, EVA, EVEX and their infectivity to fishes. Fish Pathology 15:173–184. Nishizawa, T., S. Kinoshita, and M. Yoshimizu. 2005. An approach for genogrouping of Japanese isolates of aquabirnaviruses in a new genogroup, VII, based on the VP2/NS junction region. Journal of General Virology 86(7):1973–1978. Okomoto, N., T. Sano, et al. 1983. Antigenic relationships of selected strains of infectious pancreatic necrosis and European eel virus. Journal of Fish Diseases 6:19–25. Ono, S., K. Wakabayashi, and A. Nagai. 2007. Isolation of the virus causing viral endothelial cell necrosis of eel from cultured Japanese eel Anguilla japonica. Fish Pathology 42:191–200 (English abstract). Peters, G. 1975. Seasonal fluctuations in the incidence of epidermal papillomas of the European eel Anguilla anguilla L. Journal of Fish Biology 7:415–422. Pfitzner, I., and G. Schubert. 1969. Ein virus aus dem blut mit blumendohlkrankheit behalfteter ale. Zeitschrift fuer Naturforschung 24b: 790. Rijsewijk, F., S. Pritz-Verschuren, et al. 2005. Development of a polymerase chain reaction for the detection of Anguillid herpesvirus DNA in eels based on the herpesvirus DNA polymerase gene. Journal of Virological Methods 124(1–2):87– 94. Sano, T. 1976. Viral diseases of cultured fishes in Japan. Fish Pathology 19:221–226. Sano, M., H. Fukuda, and T. Sano. 1990. Isolation and characterization of a new herpesvirus from eel. In: F. O. Perkins, and C. T. Cheng (eds) Pathology in Marine Science. New York, Academic Press, pp. 15–31. Sano T., T. Nishimura, et al. 1977. Studies on viral diseases of Japanese fishes. VII. A rhabdovirus isolated from European eel (Anguilla anguilla). Fish Pathology 10:221–226. Sano, T., N. Okamoto, and T. Nishimura. 1981. A new viral epizootic of Anguilla japonica Temminck and Schlegel. Journal of Fish Diseases 4:127–139.

¨ Schwans-Pfitzner, I., M. Ozel, et al. 1984. Morphogenesis and fine structure of eel virus (Berlin), a member of the proposed birnavirus group. Archives of Virology 81:151–162. Shchelkunov, I. S., E. K. Skurat, et al. 1989. Rhabdovirus anguilla in eels in the USSR and its pathogenicity for fish. Voprosy Virusologii 34(1):81–4. Sorimachi, M. 1984. Pathogenicity of ICD virus isolated from Japanese eel. Bulletin National Research Institute of Aquaculture 6:71–75 (English abstract). Sorimachi, M., and S. Egusa. 1982. Characteristics and distribution of viruses isolated from pondcultured eels. Bulletin National Research Institute of Aquaculture 6:71–75. Thiery, R., C. de Boisseson, et al. 2002. Phylogenetic analysis of viral haemorrhagic septicaemia virus (VHSV) isolates from France (1971–1999). Diseases of Aquatic Organisms 52(1):29–37. Ueno, Y., S. Chen, et al. 1983. Characterization of a virus isolated from Japanese eels (Anguilla japonica) with nephroblastoma. Bulletin of the Institute Zoological Academy of Sinica (Taipei) 23:47–55. Ueno, Y., T. Kitao, et al. 1992. Characterization of a herpes like virus isolated from cultured Japanese eels in Taiwan. Fish Pathology 27(1):7–17. Ueno Y., J. W. Shi, et al. 1996. Biological and serological comparisons of eel herpesvirus in Formosa (EHVF) and Herpesvirus anguillae(HVA). Journal of Applied Ichthyology 12:49–51. van Ginneken, V., B. Ballieux, et al. 2005. Hematology patterns of migrating European eels and the role of EVEX virus. Comparative Biochemistry and Physiology, Part C: Toxicology & Pharmacology 140(1):97–102. van Ginneken, V., O. Haenen, et al. 2004. Presence of virus infections in eel populations from various geographic areas. Bulletin of the European Association of Fish Pathologists. 24:270–274. van Nieuwstadt, A. P., S. G. Dijkstra, and O. L. Haenen. 2001. Persistence of herpesvirus of eel Herpesvirus anguillae in farmed European eel Anguilla anguilla. Diseases of Aquatic Organisms 45(2):103–107. Wolf, K. 1988. Fish Viruses and Fish Viral Diseases. Ithaca, Cornell University Press, 476pp. Wolf, K., and M. Quimby. 1970. Virology of eel stomatopapilloma. In: Progress in Sport Fishery Research 1970. U.S. Fish and Wildlife Service Resource Publication No. 106, pp. 94–95.

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Chapter 8

Trout and salmon viruses

Viral diseases of salmonids are among the most studied infectious fish diseases. In almost all cool and cold-water regions of the world, trout and salmon are important to popular culture and sport fishing. Therefore, virus diseases of salmonids receive much attention from fish pathologists, aquaculturists, and the public. Furthermore, international interest in salmonids and the trade of live fish and eggs has facilitated the spread of pathogens and is the basis of many national and regional disease surveillance and avoidance programs. Four of the seven OIE listed virus diseases of fish affect salmonids (OIE 2008). Wolf (1988) cited nine viruses of salmonids. Sano (1995) discussed 20 viruses that infect salmonids, including those that are highly or intermediately pathogenic and those that apparently cause few problems. Some of these viruses are readily isolated in cell cultures while others have not been successfully cultured in cells but can be propagated in susceptible fish and re-isolated; several have only been detected by electron microscopy.

Infectious hematopoietic necrosis virus Infectious hematopoietic necrosis (IHN) is a highly contagious virus disease primarily of rainbow trout and several species of Pacific salmon. The etiological agent of IHN is a rhabdovirus, infectious hematopoietic necrosis virus (IHNV) (Amend et al. 1969). Episodes of IHN date back to the 1940s and 50s when unexplained juvenile salmon mortalities were reported on the west coast from California to Washington. Rucker et al. (1953) suggested a viral etiology for these deaths. Related disease outbreaks in the Pacific Northwest have also been known as Columbia River sockeye salmon disease, Oregon sockeye virus disease, sockeye salmon virus disease, and Sacramento River chinook disease (Amend et al. 1969). Comparative studies of histopathology and viral biochemical, biophysical, and serological properties determined that the viruses that caused these diseases were the same (Amend et al. 1969; Amend and Chambers 1970).

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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Geographical range and species susceptibility Originally, IHNV was confined to anadromous salmon in tributaries that flowed directly into the Pacific Ocean from northern California to Alaska. But by the late 1960s, IHNV epizootics were confirmed in Minnesota (Plumb 1972), South Dakota, Montana, and West Virginia. The virus was later reported in rainbow trout in a closed culture system in New York (Wolf 1988). Each of these epizootics involved fish that had originated in northwestern United States and fortunately the virus has not become established in eastern United States. In addition to being enzootic on the west coast of North America, IHNV has now become established in the Hagerman Valley of Idaho and has been confirmed in China, France, Germany, Italy, Japan, Korea, and Taiwan (Sano 1976; Bovo et al. 1987; Hattenberger-Baudouy et al. 1989; LaPatra et al. 1989a; Park et al. 1993; Enzman et al. 2005). In Russia, IHN was found in Rainbow trout near Moscow in 2000 and has been shown to be endemic in some wild and hatchery populations of sockeye salmon on the Pacific coastal streams since 2001 (Rudakova et al. 2007). Species most susceptible to IHNV are rainbow trout (steelhead), chinook, sockeye (kokanee) (Rucker et al. 1953; Amend et al. 1969; St-Hillaire et al. 2001), and chum salmon (K. Amos, Washington Department of Fish and Wildlife, personal communication). Wingfield and Chan (1970) listed coho salmon as being refractive to IHNV; however, LaPatra and Fryer (1990) isolated the virus from adult coho, but alevin coho were resistant. Sano (1976) reported that amago and masu (yamame) were also susceptible. IHNV will infect brook trout and brown trout but to a much lesser degree than rainbow trout. According to Follett et al. (1997) Arctic char, Arctic grayling, and pink salmon are refractory to experimental infection with IHNV, but lake trout will support virus

replication with concomitant clinical signs. Mulcahy (1984) reported Atlantic salmon mortalities from a natural IHNV infection. Experimentally IHNV-infected larval white sturgeon can possibly become vectors of the virus, but fingerlings and adults are probably refractive (LaPatra et al. 1995).

Clinical signs and findings Clinical signs of IHN may vary somewhat from species to species (Amend et al. 1969; Mulcahy 1984). Larger fish in an infected fingerling population usually exhibit clinical signs before smaller fish of the same age. IHNV-infected fry and fingerlings are lethargic, avoid currents, move to the edge of ponds or raceways, or lie on the bottom respiring weakly. In final stages of disease, they swim in circles and hang vertically or flash followed quickly by death. Externally, affected fish are dark, exophthalmic, have swollen abdomens, and pale gills. The base of fins may be hemorrhagic, petechiae are present on mouth and body surface, and chevron-like hemorrhages occur along the lateral musculature (Figure 8.1). An opaque, mucoid fecal cast trails from the vent. An abnormally dark (red) area can develop behind the head, in the abdominal region, or on the caudal peduncle (Burke and Grischkowsky 1984). Internal organs appear anemic with petechia in the mesenteries, peritoneum, air sac, liver, and kidney. The digestive tract is void of food but contains mucoid fluid and the body cavity may contain pale, yellowish, clear fluid. Several deformities associated with IHNV are malformed heads, scoliosis, or lordosis in 5–60% of survivors (Rucker et al. 1953; Amend et al. 1969) (Figure 8.1).

Diagnosis Clinical IHNV is diagnosed by recognition of nearly pathognomonic clinical signs and conventional isolation of virus in cell culture.

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(a)

(b)

(c)

Figure 8.1 (a) Coho salmon infected with infectious hematopoietic necrosis virus (IHNV). Note the chevron shaped hemorrhages in the musculature (small arrow), the hemorrhage behind the head and on the abdomen and swollen abdomen (large arrow) (Photograph (a) courtesy of J. Rohovec). (b) Exophthalmia in hybrid rainbow trout with IHNV. (c) Petechial hemorrhage in liver (large arrow) and enlarged, dark spleen (small arrow) in hybrid rainbow trout with IHNV. (Photographs (b) and (c) courtesy of S. LaPatra, Clear Springs Food Incorporated.)

Whole fry or viscera of fingerlings are suitable material for virus isolation. Rainbow trout infected by immersion produce IHNV titers as high as 109 PFU/g of body tissue within 5 days (Yamamoto et al. 1990). Significant viral infectivity remains in tissue samples for up to 2 weeks at 4◦ C and IHNV in some preparations of ovarian fluid, eggs, serum, and brain can retain infectivity for 5 weeks (Burke and Mulcahy 1983). However, Hostnik et al. (2002) showed that rapid laboratory process and proper storage of potential IHNV positive tissue samples as well as different detection

methods are important in efficient and accurate diagnosis. They isolated IHNV from samples for 3 days at 4◦ C but only during day 1 at 25◦ C. Also viral RNA could be amplified and identified during incubation for 35 days at 4◦ C but only for 8 days at 25◦ C. Frozen samples stored at −20◦ C retain infectivity for 3–5 months. The EPC cell line is the cell line of choice for isolation, but CHSE-214, RTG-2, FHM, BF2 cells and others are susceptible and are commonly used (Lorenzen et al. 1999). It was shown by Ristow and Avila (1994) that a

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(a)

(b)

Figure 8.2 Cytopathic effect (CPE) of infectious hematopoietic necrosis virus in CHSE-214 cells incubated at 15◦ C. (a) Focal CPE at 48 hours and (b) total CPE at 60 hours after inoculation (×500).

cell line from rainbow trout (RBTE 45) was susceptible to IHNV as were three cell lines from coho salmon. All produced better viral replication than CHSE-214, but EPC cells remained the most sensitive. Yoshimizu et al. (1988a) indicated that IHNV would replicate in all 17 cell lines derived from salmonids and in 12 of 15 nonsalmonid fish cell lines tested. Lorenzen et al. (1999) did an interlaboratory evaluation of IHNV replication in EPC, CHSE-214, RTG-2, FHM, and BF2 cells and

found the highest and most consistent replication in EPC cells. Cultures are incubated at 15◦ C (never above 20◦ C) and maintained at a pH of 7.3-7.6 for up to 14 days. If IHNV is present in the sampled tissue, CPE may appear in 2–3 days at 15◦ C, but usually appears in 4–5 days. The CPE is initially characterized by pyknotic cells followed by rounding and sloughing of these cells from the surface (Figure 8.2). Treatment of monolayers of IHNV-susceptible cells with 7% polyethylene

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glycol prior to inoculation with virus suspect materials enhances plaque-forming unit (PFU) titers by 4–17 fold (Batts and Winton 1989). Blind passage is recommended for samples that are negative on primary isolation attempts. Coinfection with infectious pancreatic necrosis virus (IPNV) strongly inhibits IHNV replication (Alonso et al 1999). Therefore, if a culture is infected with IPNV, the sample should be reinoculated onto IHNV-susceptible cells after pretreatment with IPNV neutralizing antisera. Once IHNV is isolated, positive identification is made by conventional serum neutralization, ELISA, or other serological methods or by RT-PCR. Alkaline phosphatase immunocytochemical procedures detected and identified IHNV in dried tissue cultures more than 1 year after drying (Drolet et al. 1993). The system was sensitive for five known IHNV types and no cross-reactivity was noted with six other fish rhabdoviruses or with IPNV. The test was specific, sensitive, and was not affected by age of fixed or dried cell-culture plates. LaPatra et al. (1989b) found that polyclonal and monoclonal antibody can be used to detect IHNV in blood and organ smears of juvenile salmonids and in ovarian fluid of adults. Virus was also detectable in cell cultures 48 hours after inoculation and was sensitive for all virus isolates. According to Arnzen et al. (1991) the fluorescent antibody technique (FAT) is as sensitive as plaque assay and requires less time to obtain a confirmed diagnosis; however, monoclonal antibody against IHNV nucleoprotein and glycoprotein detected stages of virus replication in cell cultures as early as 6–8 hours postinoculation. Dixon and Hill (1984) developed an ELISA technique for rapid IHNV identification in infected tissues with results being obtained in 2 hours. An ELISA assay detected as few as 70 PFU viruses in 100 µg of homogenized Atlantic salmon tissue and virus in cell culture media (Medina et al. 1992). Results were obtained in one day and cross-reacted with other types of IHNV, thus enabling detection of all tested types. The immunodot blot

method may also be used in detecting IHNV protein (McAllister and Schill 1986). Molecular methods has been used extensively in detecting IHNV in fish tissue and cell cultures. Arakawa et al. (1990) utilized RT-PCR in combination with a nucleic acid probe to develop a rapid, highly specific, and sensitive IHNV detection method. A nonradioactive DNA probe developed by Deering et al. (1991) detected and identified IHNV using a dot blot format in virus-infected tissue cultures. This technique proved to be sensitive in detecting very small concentrations of target viral mRNA. There are several RT-PCR assays in use, but the standard method advocated by OIE (OIE 2008) and the AFS Bluebook (USFWLS and AFS-FHS 2007) use a modification of the assay targeting the G gene by Emmenegger et al. (2000). Furthermore, products can be sequenced for genotyping (Emmenegger et al. 2000; Kurath et al 2003). The molecular assays are commonly used as confirmation of presumptive diagnosis based on cell culture and they can be used directly on tissues from affected fish. They are more rapid than standard tissue culture methods and some are just as sensitive (Knusel et al. 2007).

Virus characteristics IHNV is a rhabdovirus in the Novirhabdovirus genus. The virion is bullet shaped, enveloped, and has a mean diameter of about 70 nm and a length of 170 nm (Amend and Chambers 1970). The genome is a nonsegmented negative-sense single-stranded RNA, 11,131–11,137 nucleotides in length and encodes six genes: nucleocapsid (N), polymerase-associated phosphoprotein (P), matrix protein (M), surface glycoprotein (G), nonvirion protein (NV), and virus polymerase (L) (Morzunov et al. 1995; Schutze et al. 1995). The virus is heat, acid, and ether labile. Stability studies of IHNV are contradictory. Pietsch et al. (1977) reported that virus survived for 5 days or less in seawater and Hank’s

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balanced salt solution (HBSS) but with addition of l0% fetal calf serum, viability was extended to 12 days. Barja et al. (1983) demonstrated that IHNV survived for about 30 days in freshwater at 15◦ C and 17 days at 20◦ C. Survival was 17 days at 20◦ C and 22 days at 15◦ C in seawater. Infectivity of IHNV is reduced in distilled water at pH 5 and 9, but at pH 6–8, it remains infective for over 10 days. IHNV does not survive well under dry conditions at any temperature but freezing has little adverse effect. Virus titers in cell culture supernatant during cyclic freezing and thawing remained constant throughout 33 cycles, whereas the same treatment of visceral organ homogenates resulted in reduction of IHNV titers (Hostnik et al. 2002). McCain et al. (1971) demonstrated by plaque reduction that there was only one IHNV serotype and this was confirmed by Engelking et al. (1991). On the basis of the molecular weights of nucleocapsid proteins and glycoproteins, Hsu et al. (1986) classified the virus into five different electropherotypes: Type 1, 2, 3, 4, and 5. Using monoclonal antibodies, Winton et al. (1988) separated IHNV isolates from Alaska, California, Idaho, Oregon, Washington, and Japan into four separate groups. Although only one IHNV serological strain has been identified, Arkush et al. (1989) showed that the United States isolate has a different polypeptide profile and is antigenically different from European isolates. Isolates of IHNV from France were typed by Danton et al. (1994) and only 5 of 27 isolates were recognized by monoclonal antisera Mab1NDW14D, previously considered to be a universal immunofluorescent reagent for IHNV. All 27 isolates did share the nucleocapsid epitope identified by Mab2NH105B, thought to be specific for IHNV electropherotype 2; however, these isolates did not belong to this electropherotype. It was suggested that IHNV had been in Europe for a sufficient amount of time to evolve separately from North American strains.

Nucleic acid analysis has been used to group IHNV isolates. Genetic comparisons are made by RT-PCR amplifying two fragments of the genome, a 303 nt fragment midG from the glycoprotein gene and a 412 nt frag ment 5 N from the nucleocapsid gene, and sequencing them (Garver et al. 2003). Using this system Kurath et al. (2003) identified three major genogroups U, M, and L that were geographically distributed in the upper (northern), middle, and lower (southern) portions of the IHNV range, respectively. European isolates were found to be within the M group (Enzmann et al. 2005; Kolodziejek et al. 2008). All Russian isolates were in the U group (Rudakova et al. 2007). Japanese isolates were in U and a new group designated JRt (for Japanese rainbow trout) (Nishizawa et al. 2006). Biacchesi et al. (2000) established a system to produce IHNV recombinants and allowing the evaluation of gene functions. They found that IHNV recombinants lacking the NV gene are viable but produced less progeny. In a later study, the NV deletant was shown to be attenuated and induce protective immunity to wild type IHNV challenge (Thoulouze 2004). They also produced a recombinant containing the virulent VHSV NV gene. Viable recombinants containing VHSV and SVCV G genes in place of the IHNV G gene and VHSV M gene replacements have also been produced, showing that the gene product functions of these fish rhabdoviruses are very similar (Biacchesi et al. 2002).

Epizootiology IHN is generally considered to cause high mortality among young cultured salmonids. Actual percentage of deaths can vary from low to nearly 100% depending upon fish species, age, and size, environmental conditions, particularly water temperature and virus strain. The mortality pattern of IHNV is typical of highly virulent viruses that affect

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young fish. Under optimum conditions, death rate accelerates rapidly and often 70% or greater mortality will accrue within 10 days. Amend and Nelson (1977) demonstrated that sockeye salmon, which had undergone selective breeding, suffered 52–98% mortality after artificial infection with IHNV; concluding that these variances were due to genetic differences among the host. A 30% heritability of resistance to IHNV resulted from selective breeding, suggesting that more IHNV-resistant fish strains could be developed (McIntyre and Amend 1978). Using rainbow trout as the host, Idaho, Oregon, and California IHNV strains were compared. At 10◦ C, 62% mortality occurred in groups infected with the Idaho strain, 4% in groups infected with the Oregon strain, and 67% in groups infected with the California strain. Age of fish influences mortality due to IHN and most high IHN-related mortalities involve fry or small fingerlings (Amend et al. 1969; Plumb 1972; LaPatra et al. 1990a). Mortalities of fry up to 2 months of age exceeded 90% at 10◦ C (Pilcher and Fryer 1980). In 2–6 month old fish, mortality is often more than 50% but can be as low as 10% in yearlings; sockeye salmon were susceptible to IHNV for up to 210 days of age (7.2 g) and rainbow trout up to 170 days (13.1 g) (LaPatra et al. 1990a). However, outbreaks have occurred in 2-yearold sockeye salmon, but mortality has usually been comparatively low. The virulence of five European and one North American IHNV isolates was tested in 2, 20, and 50g rainbow trout by immersion (Bergman et al. 2003). The smallest fish were most susceptible to any isolate with 18–100% mortality, leading to the conclusion that the virulence of different IHNV isolates was related to the age and weight of the fish. Chinook salmon, generally considered not susceptible to IHNV, can be experimentally infected, become virus carriers and can transmit the disease to other species during cohabitation (St-Hillare et al. 2001). During cohabitation, the virus was transmitted from chinook

Table 8.1 Percent mortality of rainbow trout (0.2 to 0.3 g) not infected and infected with infectious hematopoietic necrosis virus by waterborne exposure and held at seven temperatures. Water Temperature (◦ C)

Control Infected

3

6

9

12

15

18

21

0 83

0 81

0 76

1 68

6 66

30 80

67 63

Source: Hetrick et al. (1979).

to Atlantic salmon with no serious effects on chinook unless they had a concomitant bacterial kidney infection. During the dual infection, IHNV could be isolated from both species. Water temperature also affects mortality rates due to IHNV seldom occurring at temperatures above 15◦ C (Amend 1970; LaPatra et al. 1989a). However, Hetrick et al. (1979) showed that IHNV could cause 72–88% mortality in 0.2–0.3 g rainbow trout at 18◦ C and 72–94% at 3◦ C (Table 8.1). The mean day to death was 15–18 days at 3◦ C and 7–10 at 18◦ C. IHNV was isolated from infected dead fish at 21◦ C, but mortality among groups of infected fish and noninfected controls at the higher temperature were similar. Concurrent infection with other viruses such as IPNV can also influence disease and reduce losses due to IHNV (de Kinkelin et al. 1992; Alonso et al. 2003). The actual cause of the reduced pathogenesis is not known, but IPNV interferes with IHNV replication in cell cultures as well (Alonso et al. 1999). This interference may at least be partially due to inhibition of virus uptake. Using green fluorescent protein labeled IHNV, VHSV and IPNV in flow cytometry experiments on BF-2 cells, de las Heras et al. (2008) demonstrated a significant inhibition of IHNV uptake when coinfected with IPNV but no inhibition of VHSV uptake in IPNV coinfections. Also, IPNV induces interferon activity (de Kinkelin et al. 1992), which may suppress IHNV replication. Coinfection of trout with the nonpathogenic cutthroat trout reovirus also induces an IHNVresistant state (Hedrick et al. 1995).

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Survivors of IHNV infections are thought to become virus reservoirs, but it is not clear if these fish actually continue to carry and shed virus after surviving an epizootic as juveniles or if they become reinfected horizontally at a later time. Amos et al. (1989) found that adult sockeye salmon captured as they left salt water during spawning migration were free of IHNV, and if allowed to become sexually mature in IHNV-free freshwater, they remained uninfected until spawning. Adults allowed to migrate naturally to sea and return to spawning grounds developed an IHNV positive prevalence of 90–100%. These data strongly suggest that returning adults become infected from other sources rather than by reappearance of latent virus. In a study by Wingfield and Chan (1970), 34% of females and 5% of males assayed were IHNV carriers. Mulcahy et al. (1983a) sampled females from seven adult salmon populations and found that active infections ranged from 39–100% (average 85%). The samples were taken from ovarian fluid or postspawning tissue and many virus titers exceeded 105 TCID50 /mL. Prevalence of IHNV positive adult female Alaskan sockeye salmon varied from 7–94% (average 44%) and positive adult males varied 0–48% (average 13%) (Grischkowsky and Amend 1976). Mulcahy et al. (1984) studied the IHNV carrier rate of female sockeye salmon from August (prespawning) through October (peak spawning). In August, no assayed fish were positive for IHNV but in October, 100% were positive (Table 8.2). As females progress from a prespawning to spawning condition they shed greater numbers of virus and potential for vertical transmission increases. Sperm could also be a vehicle for virus to enter the egg during fertilization because experimentally sperm adsorbed IHNV (Mulcahy and Pascho 1984). These researchers subsequently reported two instances of vertical IHNV transmission (Mulcahy and Pascho 1985). Contrastingly, Yoshimizu et al. (1989) showed that IHNV does not survive in fertil-

Table 8.2 Incidence of infectious hematopoietic necrosis virus in prespawning female sockeye salmon in 1980.

Date

August 17-–30 September 11-–25 October 6-–7 October 15 October 20-–29

Number Assayed

Percent Positive

18 31

0 0

87 69 65

21.5 96.5 100

Condition

Prespawning Prespawning Spawning Spawning Spawning

Source: Mulcahy et al. (1984).

ized egg yolks prior to the eyed stage and that no virus was present on the egg surface. However, when eyed eggs were injected with IHNV, virus concentrations increased and 90% of the eggs died. Also, 90% of hatched masu salmon and 20–30% of hatched chum salmon from injected IHNV eyed eggs died. These authors doubt that vertical IHNV transmission occurs because of the virus’s inability to survive in eggs before they become eyed. The first IHNV isolation from adult sockeye salmon while at sea was reported by Traxler et al. (1997). They investigated the possibility of vertical transmission by these virus positive fish but found that IHNV was neither transmitted during spawning nor could they infect ova by hardening eggs in water containing high concentrations of IHNV. These data would tend to contradict earlier circumstantial evidence, indicating that vertical transmission does occur and leaves this mode of transmission open to further study. The virus can be transmitted horizontally via waterborne exposure, feeding, or injection (Mulcahy et al. 1983b). IHNV can be isolated from coelomic fluid. However, Mulcahy and Batts (1987) showed that if nonerythrocytic cells are separated from coelomic fluid and incubated at 15◦ C for 7 days and then assayed for IHNV, the number of IHNV positive fish will be higher than if only the coelomic fluid is assayed. Detection of specific antibody in fish serum may be used to determine prior exposure. Jørgensen et al.

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(1991) screened serum from rainbow trout for IHNV antibody by plaque neutralization test (PNT), ELISA, and FAT. The three methods detected antibody in 9 to 18 of 20 serum samples; ELISA proved to be the most sensitive and PNT the least. These assays could be useful in epizootiological IHNV studies, but because some sera cross-reacts with VHSV, the procedure probably could not be used when specific pathogen-free determinations are required, especially since IHNV and VHSV have been isolated from the same fish (Eaton et al. 1991a). A nonsalmonid IHNV reservoir is uncertain, but Mulcahy et al. (1990) isolated high concentrations of virus from a leech (Piscicola salmositica) and copepod (Salmincola sp.) parasitizing sockeye salmon. High IHNV concentrations were also found in detached leeches recovered from bottom gravel of a spawning area. Though the role, if any, these parasites play in IHNV transmission is not known, the possibility that skin parasites facilitate infection during spawning must be considered. IHNV can cause significant losses in wild juvenile salmon. Williams and Amend (1976) confirmed an IHNV epizootic in naturally spawned sockeye salmon fry in British Columbia and speculated that abnormally low egg survival in Chilko Lake was due to IHNV. The virus also caused significant mortality in the Weaver Creek spawning channel, where an estimated 50% of 16.8 million migrating sockeye salmon fry succumbed to IHNV (Traxler and Rankin 1989).

Pathology Hematopoietic tissue of the kidney and spleen is the most extensively IHNV-affected tissue where degeneration and necrosis takes place, especially in the anterior kidney (Yasutake 1975). Pleomorphic intracytoplasmic inclusions frequently occur in pancreatic cells along with focal necrosis in the liver, granular cells of the lamina propria, the stratum

compactum, and stratum granulosum of the intestine. A sloughing of mucous membrane (possibly catarrhal inflammation) in the intestine may be the origin of fecal cast observed in moribund fish (Amend et al. 1969). The pathogenesis of three IHNV strains were compared by Lapatra et al. (1990b); the most virulent strain produced severe tissue injury in 3–4 days, while other strains required 8–10 days to cause similar pathology. The pathology of IHNV was also compared in selected tissues of rainbow trout and coho salmon by Helmick et al. (1995a) who found morphological differences in histopathology of the gills, esophagus/cardiac stomach region, anterior intestine, and pyloric caeca in the two species. Gills exhibited differences in distribution of mucous and chloride cells and size of pillar cells. The esophageal/cardiac regions differed with respect to epithelial cell architecture of mucus-secreting glands. In a companion study, Helmick et al. (1995b) used strept-avidin gold labeled monoclonal antibody to follow pathogenesis and replication of IHNV from esophageal mucosa to the mucussecreting serous cardiac glands in rainbow trout and coho salmon. Glandular cystic degeneration occurred as early as 1 hour postinfection and progressed into severe cystic degeneration in 24 hours. This process was more severe in rainbow trout than in coho salmon, leading to the conclusion that IHNV replicated more efficiently in mucus-secreting serous cardiac glands of rainbow trout than in coho salmon. Pathogenesis studies by Yasutake (1975) indicate that IHNV target tissue is the hematopoietic tissue of the kidney with a close correlation between histopathological progression and virus concentration. About 3 days postinfection, focal concentrations of macrophages were observed in the kidney followed by involvement of the pancreas, liver, and presence of granular cells in the intestinal wall. Amend and Smith (1975) concluded that death probably resulted from a severe electrolyte and fluid imbalance caused by renal

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failure. The role of leukocytes in IHNV replication was shown in vitro by Chilmonczyk and Winton (1994). LaPatra et al. (1989a) suggested that infection from waterborne exposure resulted from virus invading the integument and gills because an increase in detectable virus developed in these tissues for 3 days postexposure. They also postulated that detectable virus in the mucus resulted from replication in the integument. Subsequent studies by Cain et al. (1996) found IHNV in cutaneous and intestinal mucus following injection or waterborne virus exposure. Some antiviral activity was confirmed in intestinal tract mucus and the authors suggested that mucus from these areas could be an innate mechanism of viral resistance which may be important as a first line of defense against IHNV. Yamamoto and Clermont (1990) described the sequential spread of IHNV to tissues of rainbow trout infected by immersion. Gill tissues had virus titers 16–20 hours postexposure and virus spread rapidly to the kidney and spleen. In fact, Zhang and Congleton (1994) showed that IHNV could be isolated from water that contained infected juvenile steelhead trout several days prior to onset of clinical signs and mortality. Using whole-body assay and immunohistochemistry, Yamamoto et al. (1990) determined that initial lesions following infection occurred in the epidermis of pectoral fins, opercula, and on the ventral body surface. LaPatra et al. (1989c) also found virus in mucus of juvenile and adult fish infected with IHNV and postulated that the epithelium is a portal of entry for virus and a site of replication. Lesions did not become apparent in the kidneys until the third day. These findings were essentially confirmed by Drolet et al. (1994), who followed IHNV progression to various organs and tissues of steelhead trout fry after immersion and oral infection. The virus passed directly from the gills into the circulatory system where virus was detected 2 days later. Virus was found in the brain, pancreas, and heart 5 days after exposure with detection

in other organs being intermediate. After gastrointestinal exposure, virus passed through the epithelium into the circulatory system and then to all organs and tissues. These experiments indicate that the epidermis and gills are important inital sites of IHNV replication. A recent study using luciferase expressing IHNV recombinants demonstrated that fin bases are the first sites of viral replication when challenged by immersion (Harmache et al. 2006).

Significance IHNV is the most serious viral disease that affects trout and salmon in the Pacific Northwest of the United States. It has also become an important disease in Europe and Asia, particularly Japan and China (Sano et al. 1977; Jiang 1993). Seventy million young salmon succumbed to IHNV infections in Japan in 1975. Clearly, the virus has been responsible for the death of millions of juvenile hatchery reared Pacific salmon and rainbow trout and likely large numbers of wild salmonids as well.

Infectious pancreatic necrosis virus Infectious pancreatic necrosis (IPN) is generally characterized as a subacute disease of fry and fingerling salmonids, but IPN- or IPN-like viruses have also been isolated from a variety of marine and freshwater fish and invertebrates. These isolates are combined into the aquatic birnavirus group (Hill and Way 1995). IPNV was likely the etiological agent of “acute catarrhal enteritis” described by M’Gonigle (1941) in Canada. Wood et al. (1955) were the first to describe the disease as “infectious pancreatic necrosis” and to propose its possible viral etiology. Wolf et al. (1960) demonstrated the viral etiology by isolating and propagating the virus in cell culture, thus making IPNV the first proven viral disease of fish.

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Geographical range and species susceptibility IPNV and IPNV-like agents are some of the most widely distributed aquatic viruses, essentially occurring worldwide. Hill and Way (1995) listed 17 countries in which IPNV or aquatic birnaviruses have been found including North, Central and South America, Europe, and Asia (Table 8.3). IPNV has been isolated from 16 species of salmonids (Wolf 1988). Brook and rainbow trout are the most susceptible, but virtually all salmonids are susceptible to some degree, including Pacific and Atlantic salmon (McAllister et al. 1984; Christie and Haverstein 1989). Hill and Way (1995) listed 31 families of nonsalmonid fish, ranging from primitive Petromyzonidae (lampreys) to phylogenetically advanced Cichlidae (cichlids), nine

species of mollusks, four species of crustaceans, and two gastropods from which aquatic birnaviruses have been recovered. These animals come from freshwater and marine habitats in subarctic to tropical climates. While all aquatic birnaviruses are not overtly detrimental to the health of affected animals, clearly they are not host specific and appear to affect salmonids more severely than nonsalmonids. In a survey of fish farms in England and Wales, Buck et al. (1979) found that of 29 salmonid farms surveyed, 17% were rearing fish infected with IPNV, but there was no evidence that nonsalmonid fishes living in the effluent from these farms were infected with virus. However, Wallace et al. (2008) isolated IPNV from 9 of 17 species of fish collected from Scottish waters in the vicinity of Atlantic salmon farms.

Clinical signs Table 8.3 Country (year reported) in which infectious pancreatic necrosis virus and other aquatic birnaviruses have been isolated.

Country

Year Reported

Australia Canada Chile China Denmark France Germany Greece Finland Italy Japan Mexico New Zealand Norway Scotland South Africa Spain Sweden Taiwan Thailand United Kingdom United States

2000 1982 1984 1989 1969 1965 1975 1975 ? 1972 1971 2002 1987 1976 1971 ? 1988 1973 1983 ? ? 1955

Source: McAllister and Reyes (1984); Wolf (1988); McAllister (1993); Hill and Way (1995); Crane et al. 2000, Ortega et al. 2002.

Larger, more robust fry and small fingerlings of a population of susceptible salmonids are usually the first to develop clinical signs of IPNV associated disease. Externally infected trout have an overall darker pigmentation, abdominal distention, exophthalmia, hemorrhages on the ventral surface and fins, and pale gills (Figure 8.3) (Snieszko et al. 1959; Wolf et al. 1960). Rapid whirling on the long axis followed by quiescence is a typical behavior of IPNV-infected trout. Internally, a general

Figure 8.3 Rainbow trout infected with infectious pancreatic necrosis virus. Note the enlarged abdomen, exophthalmia and very dark skin pigmentation. (Reprinted with permission of CRC Press.)

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hemorrhagic appearance is obvious with petechia throughout the viscera, especially in the pyloric caeca and adipose tissue. The spleen, heart, liver, and kidneys are unusually pale in advanced cases. The body cavity is filled with clear yellow fluid, the digestive tract is void of food, and the posterior stomach contains a gelatinous, mucoid (clear or milky) plug, which is pathognomonic for IPNV. The gelatinous plug remains intact in 10% formalin; therefore, it is of diagnostic value in examining preserved specimens. Clinical signs described above are present primarily in IPNV-infected salmonids and are not always seen in other birnavirus-infected fish. Sano et al. (1981) reported that Japanese eels infected with IPNV had muscle spasms, retracted abdomens, and congestion of the anal fin, abdomen, and gills. These fish also had ascites and mild hypertrophy of the kidneys. IPNV-infected larval seabass displayed spiral swimming, swim bladder distention, fecal casts, exophthalmia, and sloughing of intestine epithelium (Bonami et al. 1983). Infected striped bass showed no overt clinical signs of disease other than darker skin pigmentation (Schutz et al. 1984). Atlantic menhaden from the Chesapeake Bay exhibited spinning (Stephens et al. 1980), and gizzard shad, from which an IPN-like virus was isolated, showed only dark skin pigmentation (Southeastern Cooperative Fish Disease Laboratory, Auburn University, Alabama, USA, unpublished). IPNV-infected juvenile Atlantic halibut (0.1–3.5 g) had distended abdomens, exhibited uncoordinated swimming, the livers, kidneys, and intestines became necrotic but the pancreatic tissue was unaffected (Biering et al. 1994).

Diagnosis In trout, presumptive diagnosis of IPNV is based on mortality patterns and clinical signs combined with virus isolation in tissue culture. However, in non-salmonids, IPNV or aquatic birnavirus infections may not be ex-

pressed in clinical disease or mortality, and IPN diagnosis is confirmed by virus isolation and serological identification. Histological examination can be used to support a clinical diagnosis by detecting typical IPNV-associated histopathology (Roberts and McKnight 1976). Virus isolation is obtained by homogenizing whole fish (fry) or visceral organs from IPNV suspect fish. Homogenates decontaminated by filtration or antibiotics and diluted to at least 1:100 are inoculated onto susceptible cells (RTG-2, CHSE-214, BF-2, etc.). Yoshimizu et al. (1988a) tested 32 salmonid and nonsalmonid cell lines and found 26 to be susceptible to IPN virus. Inoculated cells incubated at about 20◦ C develop CPE consisting of cellular pyknosis, elongation, and lysis in 18–36 hours (Figure 8.4) (Wolf et al. 1960). Temperature range for IPNV replication is 4–27.5◦ C, but incubation temperature should be appropriate to cell line used, i.e., neither CHSE-214 nor RTG-2 should be incubated above 20◦ C. A blind passage at dilutions greater than 1:10 should be made in the absence of CPE after 7–14 days of incubation. Clinically, IPNV-infected fish will have titers of 106 –109 TCID50 /g of tissue, but in the carrier state titers can be as low as 101 TCID50 /g of tissue. Serum neutralization is usually used to positively identify IPNV and other aquatic birnaviruses. Other types of identification techniques used in identifying IPNV in cell cultures, cell culture fluids, and infected tissues are fluorescent antibody, ELISA, and immunoperoxidase systems (Hattore et al. 1984; Shankar and Yamamoto 1994; Hill and Way 1995). Nucleic acid probes and monoclonal antibodies have made rapid and accurate detection and identification of IPNV possible (Dominquez et al. 1991). Babin et al. (1990) described a capture immuno-dot procedure that uses monoclonal antibodies to detect IPNV in field identification studies. A purified reference IPNV strain in conjunction with immunogold and electron microscopy was used to distinguish between the Sp, Ab, and West

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(a)

(b)

Figure 8.4 CHSE-214 cells infected with infectious pancreatic necrosis virus and incubated at 15◦ C. (a) Non-infected controls. (b) Extensive viral CPE 48 hours after inoculation (×500). (Photo by J. Grizzle.)

Buxton (WB) strains of IPNV (Novoa 1996). A dot-blot hybridization test was developed to detect IPNV-RNA in fish cells and cell cultures, a procedure that was 100% effective in detecting IPN-RNA in inoculated cell cultures when applied 12–24 hours after inoculation, but less sensitive in fish tissue (Dopazo et al. 1994). These researchers concluded that the hybridization test is appropriate to identify IPNV when combined with laboratory diagnostic cell culture.

A method for detecting IPNV by RTPCR was described by Lopez-Lastra et al. (1994). Wang et al. (1997) developed a singletube, noninterrupted RT-PCR procedure to detect IPNV in infected CHSE-214 cells. Amplified products were detected in 5–6 days postinfection with isolates of WB, Ja, Sp, Ab, VR299, 3372 MFK (isolated from eel in Taiwan), and CV-HB-1 (from hard clams in Taiwan) but not in control cultures nor cells infected with IHNV. Detection tests for

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IPNV are crucial for identifying possible virus carrier populations and properly managing the disease. When individual rainbow trout were checked for IPNV antibodies, about 30% were virus positive by isolation compared to twice that number being positive by ELISA (Dixon and de Groot 1996). These authors noted that not all virus positive fish by isolation were identified by ELISA as having been exposed however, the procedure can aid in detecting rainbow trout populations that have had previous IPNV exposure. Virus detection in carrier fish using a nondestructive procedure involves collecting ovarian and seminal fluids or fecal material (the proper amount of feces is “about as much as you can put on the tip of your tongue”) (Wolf et al. 1967). McAllister et al. (1987) found that ovarian fluid yields greater amounts of virus when the fluid is centrifuged into a pellet and sonicated before being inoculated into cell cultures. A more accurate and reliable method of detecting IPNV carrier fish is to process internal organ tissue from a population sample, but this is not always practical because it requires killing assayed animals. Munro et al. (2008) addressed the issue of a destructive procedure (visceral organ homogenates) verses a nondestructive procedure of assaying blood from Atlantic salmon at the time of spawning. Thirty fish that were positive by blood leukocyte assay prior to spawning were assayed again by destructive kidney homogenates and ovarian fluid assay at spawning. They concluded from a combination of studies that virus assay by destructive kidney homogenate is more sensitive than nondestructive blood sampling at time of spawning.

Virus characteristics IPNV is a member of the family Birnaviridae and is included in the group of viruses called “aquatic birnaviruses” (Hill and Way 1995). Other aquatic birnaviruses that are important pathogens include Eel virus European

(EVE) (Chapter 7) and yellowtail ascites virus (YTAV), which causes an acute infection of yellowtail fry in Japan (Sorimachi and Hara 1985). The virus is a nonenveloped icosahedral particle averaging 60 nm in diameter (55–75 nm) (Dominquez et al. 1991). It consists of a single capsid layer and a genome of bisegmented double-stranded RNA. The virion is stable in acid, ether, and glycerol, and is relatively heat stable. The virus is also stable at 4◦ C for 4 months in cell culture media containing serum, but a temperature of −20◦ C or lower is recommended for long term storage. At −70◦ C, about 1 log1O of infectivity is lost per year. Most IPNV infectivity is lost in freshwater within 10 weeks at 4◦ C, with residual infectivity remaining for 24 weeks. IPNV remains infective for up to 20 days in freshwater at 15◦ C and 15 days at 20◦ C compared to a 20-day survival time at both temperatures in seawater (Barja 1983). The virus will survive drying for more than 8 weeks (Desautels and Mackelvie 1975) and for up to 1 year following lyophilization in lactalbumin hydrolysate, lactose, powdered milk, or saline with 10% fetal calf serum (Wolf et al. 1969). Most aquatic birnaviruses are antigenically related, regardless of geographic origin or host species. These isolates were placed in two serogroups, a major Serogroup I, and a minor Serogroup II (Hill and Way 1988; CaswellReno et al. 1989). Serogroup I contains nine cross-reactive but distinct serotypes including WB (VR-299), Sp, Ab, He, Te, Canada 1 (C1), Canada 2 (C2), Canada 3 (C3), and Jasper (Ja) (Table 8.4). A tenth serotype (N1) was proposed by Christie and Haverstein (1989). Serogroup II contains Serotype TV-1 because it is serologically nonreactive to VR-299, Sp, Ab, and several other aquatic birnaviruses. Using 196 aquatic birnavirus isolates from 56 animal species in 17 countries, Hill and Way (1995) applied serum neutralizaton and monoclonal antibodies to reclassify this group of viruses (Table 8.4). They proposed Serogroup A (formerly Serogroup I) and Serogroup B (formerly Serogroup II).

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Table 8.4 Major serological and genogroups of infectious pancreatic necrosis virus (aquabirnaviruses).

Serogroup A (formerly I)

Serogroup B (formerly II) Serogroup C Serogroup D MABV

Serotype∗

Genogroup†

Strain

Location

Host

A1 A2 A3 A3 A4 A5 A6 A7 A8 A9 B1

I III II II VI IV IV V V I B‡

WB Sp Ab EVE He Te Can. 1 Can. 2 Can. 3 Ja TV-1

West Buxton, USA Spjarup, Denmark Abild, Denmark Japan W. Germany United Kingdom Canada Canada Canada Jasper, Canada United Kingdom

Rainbow trout Rainbow trout Rainbow trout European eel Northern pike Tellina (bivalve) Atlantic salmon Atlantic salmon Arctic char Rainbow trout Tellina

C1 § D1 ND

C§ D¶ VII†

S1 M2 Y6

Singapore Malaysia Kochi, Japan

Giant Snakehead Angelfish Yellowtail

Source: Hill and Way (1988); Lecomte et al. (1992); Hill and Way (1995). ∗ Hill and Way (1995); Lecomte et al. (1992); and Dixon et al. (2008). † Genogroups as designated by Nishizawa et al. (2005). ‡ Determined to be distinct from IPNV by Nobiron et al. (2008). § Determined to be distinct from IPNV by Da Costa et al. (2003). ¶ Sequence analysis of fragment of VP1 indicates D is distinct from A, B, and C (Dixon et al. 2008).

Serogroup A contains most of the isolates from a variety of locations and are designated Serotype A1 through A9 , whereas Serogroup B contains Serotype B1, which was isolated from a marine mollusk from the United Kingdom. Later two more serogroups were identified. Serogroup C was found in snakeheads and marble gobies in Southeast Asia, and Serogroup D was found in Angelfish, dwarf danio, and Ram Cichlids in Malaysia (Dixon et al. 2008). Nearly all aquatic birnaviruses isolated from salmonids in the United States belong to Serotype A1 (WB); however; examples of Serotype A3 (Ab) have also been isolated from other fish species, mollusks, and crustaceans (McAllister and Owens 1995). Serotypes A6 (C1), A7 (C2), A8 (C3), and A9 (Ja) have been found in Canadian fish populations (Lecomte et al. 1992), and Serotypes A2 (Sp), A3 (Ab), A4 (He), and A5 (Te) have been found in Europe. Representatives of Serotype A1 (WB), A2 (Sp), and A3 (Ab) have been detected in Asia. Biering et al. (1997) serotyped 12 birnavirus

isolates from Norway with monoclonal antibody and immunodot assay and found slight serological variations between them but concluded that the N1 and Sp/A1 (A2 ) strain were identical. Many nonsalmonid IPN isolates are of the A3 (Ab) serotype and tend to be of lower virulence for trout (McAllister and Owens 1995). In an IPNV study, Rodriguez et al. (1994) found that all IPNV strains in Spain were of the A2 (Sp) or A3 (Ab) serotype. Novoa and Fugueras (1996) detected the A1 (WB) strain in turbot in Europe; a strain previously thought to be confined to North America. They also isolated six aquatic birnaviruses from turbot that were heterogenous and could not be serotyped, so these isolates were placed in the IPNV group based on genomic segment mobility. Blake et al. (2001) used nucleotide sequencing the majority of genome segment A of 28 aquatic birnaviruses of serogroup A including all 9 serotypes and identified 6 genogroups which generally separated

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according to geographic origin and serological classification. Genome segment A sequencing of IPNV and marine aquatic birnaviruses demonstrated seven genogroups with the marine birnaviruses being very similar to each other (Zhang and Suzuki 2004). This genotype designation was confirmed by Nishizawa et al. (2005) using more isolates with an assay that evaluated sequences of the VP2/NS junction. Genome analysis of aquatic birnavirus serogroups B, C, and D indicate that they are distinct from IPNV (Da Costa et al. 2003, Dixon et al. 2008, Nobiron et al. 2008). The genomes of aquatic birnavirus serogroups B and C encode an additional protein when compared to IPNV (serogroup A) and sequence comparisons suggest that they may be more similar to avian or insect birnavirus genera (Da Costa et al. 2003, Nobiron et al. 2008).

Epizootiology IPNV is one of the more fascinating fish viruses because of its diverse host susceptibility and broad geographical range. Severity of IPNV is influenced by species, age, strain, and physiological condition of affected fish, virus strain, and environmental conditions. Manifestation of an IPNV infection varies from subclinical to acute with nearly l00% mortality possible in some affected trout populations (Wolf 1988). In Spain, mortality related to natural infections of IPNV A2 (Sp), and A3 (Ab) strains in rainbow trout farms ranged from 20–60% (Rodriguez et al. 1994). Experimental waterborne transmission of IPNV exposure was demonstrated in 13-weekold Atlantic salmon and 15-week-old brook trout (Taksdal et al. 1997). Cumulative mortality in two tanks of Atlantic salmon was 76 and 82% compared to 6 and 15% for control fish. Infected brook trout mortality was 42 and 43% compared to 8–10% for control fish. Onset of clinical infection occurred first in brook trout and virus was isolated from all moribund and dead fish assayed. The risk factors of con-

tracting IPNV by post-smolt Atlantic salmon after being moved from fresh-water to 124 marine culture sites were studied. Results showed 39.5% mortality attributable to IPNV and that risk factors were related to age, geographical location of the culture site, and method of fish transport (Jarp et al. 1995). The fact that water effluence from an IPNVcontaminated hatchery can cause problems downstream was emphasized by McAllister and Bebak (1997). They found no IPNV in streams or spring water above hatcheries, but virus was found in effluent streams 19.3 km below outfall of infected hatcheries. They also isolated IPNV from 3 of 11 brook trout but failed to isolate virus from 30 brown trout, 4 rainbow trout, or 61 nonsalmonid fish. These data emphasize the infective potential of water for susceptible salmonids below viruscontaminated facilities. The effect of continuous exposure of rainbow trout to different concentrations of IPNV was studied by Bebak and McAllister (2008). They determined that fish from eggs exposed to IPNV 6 days prior to hatching suffered higher mortality than fish from nonvirus exposed eggs. Their results also suggest that the natural aquatic environment where rainbow trout densities are lower than in their studies results in lower mortality when ambient virus concentrations are <103 IPNV/L. In aquaculture, higher fish densities result in IPN epizootics in response to continuous low exposure to IPNV. Optimum water temperature for IPNV development is 10–15◦ C. When trout fry are naturally exposed to IPNV, clinical signs can occur from 1 week to 6 months of age. Under experimental conditions, appearance of clinical signs can be as short as 3–5 days at 15◦ C. Brook trout reach peak IPNV susceptibility between 30 and 50 days of age, and mortality becomes less severe as fish age. At 10◦ C, l-month-old brook trout suffered 85% cumulative mortality, but by 6 months of age mortalities were less than 5% (McAllister and Owens 1986).

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McAllister and McAllister (1988) demonstrated IPN virus transmission from carrier striped bass to brook trout. Mortality of IPNV-infected nonsalmonids also varies from none to high. Bonami et al. (1983) reported up to 95% mortality in cultured seabass fry, and Schutz et al. (1984) reported that 2 million 4-week-old IPNV-infected striped bass died. Both of these reports carefully avoided claiming that IPNV actually killed the fish. An attempt to experimentally infect four strains of 1–120 day old striped bass with IPNV did not produce clinical disease and virus was subsequently isolated from only virus-infected fish (Wechsler et al. 1986). Although it is questionable if striped bass are killed by IPNV, McAllister and McAllister (1988) successfully transmitted virus from asymptomatic IPNVcarrier striped bass to juvenile brook trout and 8% of the surviving brook trout became longterm virus carriers. Because striped bass may carry IPNV and serve as reservoirs for more susceptible species, all striped bass or their hybrids stocked into IPNV-free waters with access to trout populations should be virus free. Numerous reports describe isolation of IPNV or IPN-like virus from asymptomatic fish during routine virus assays and any of these carriers can be a virus source for more susceptible species. The virus can also be transported by fish-eating birds because it survives in their digestive tracts and is released through excreta (Peters and Newkirch 1986). Gregory et al. (2007) isolated IPNV from mussels, bottom sediments, and surface water in the vicinity of clinically infected Atlantic salmon farms in the direction of current flow from infected fish. There was evidence of decreasing prevalence of IPNV in mussels once IPN outbreaks subsided and they concluded that mussels were an unlikely source of reinfection of maricultured Atlantic salmon. Biering et al. (1994) isolated IPNV from Atlantic halibut that was experimentally infectious to 0.1–3.5 g halibut fry at 12 and 15◦ C with significant mortality. An experimental IPNV infection was later established in

Atlantic halibut yolk-sac larvae and resulted in 100% mortality following waterborne exposure to 107 TCID50 /mL (Biering and Bergh 1996). Virulence of isolates from salmonids, nonsalmonids, and molluscs was compared in brook trout by McAllister and Owens (1995). The trout and salmon isolates were more pathogenic to brook trout than were those from nonsalmonids or molluscs. At least some virulence was found in 5 of 6 salmonid, 10 of 16 nonsalmonid, and 2 of 5 molluscan IPNV isolates. These authors concluded that IPNV is widespread and may not always result from introduction of carrier salmonids into an area. When an IPNV outbreak occurs in trout, some survivors probably become lifelong carriers but prevalence of carrier fish in a population can vary with the season. Mangunwiryo and Agius (1988) showed that percentage of fish shedding IPNV increased between March and June and declined during the latter part of the year. The heightened carrier state of these fish was inversely related to IPNV antibody levels, implying that high serum antibody titers reduce active virus production. Billi and Wolf (1969) found that when an IPNV-carrier population was sampled over time, more than 90% were carriers, but not all fish or even the same fish shed virus at each sampling. A model for an IPNV carrier state in trout and cell cultures by Hedrick and Fryer (1982) showed that virus was produced by less than 1% of the cell population of either persistently infected cell lines or kidney cells of carrier trout. In cell culture, neither antibody nor interferon affected replication of virus; however, defective virus did interfere with virus replication. A similar mechanism may function in IPNV-carrier fish. Replication of IPNV in leukocytes (in vivo) from the head kidney of Atlantic salmon suggests that these cells, especially the adherent macrophages, could play a role in maintaining a virus carrier state (Johansen and Sommer 1995). Although adverse environmental conditions are not necessarily associated with IPNV

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epizootics, Roberts and McKnight (1976) linked temperature and transport stress to recurrent disease outbreaks in yearling rainbow trout. Frantsi and Savan (1971) described a similar pattern in yearling IPNV carrier fish following exposure to low oxygen concentrations. These observations suggest that stressful environmental conditions may cause a reoccurrence of disease, especially in older fish populations. Using naturally infected juvenile brook trout, McAllister et al. (1994) demonstrated that when fish were fed immunosuppressant triamcinolone acetonide R (Kenolog ) and water temperatures were raised from 12 to 18◦ C, IPNV prevalence and virus titers increased significantly, thus showing that “stress” can affect viral infections. This procedure could also be used to increase the probability of isolating IPNV from asymptomatic fish. Post-smolt Atlantic salmon appear to be more susceptible to IPNV when transferred to seawater, which could be related to stress (Stangeland et al. 1996). Following an experimental injection of IPNV, 50% mortality was experienced under seawater conditions, while extended clinical disease and higher mortality were noted in stressed fish that was possibly induced by the injection process. Fish density probably plays a role in IPNV epizootics of rainbow trout (Bebak-Williams et al. 2002). They showed that when the number of infectious fish was low and density increased the peak death rate increased, time of peak death rate decreased and the probability of survival decreased. The effect of density diminished when number of infectious fish was high. As fish density increased, the death rate also increased, and as time of peak death rate decreased, the possibility of survival decreased. Multiple infections involving two or more pathogens in the same fish is not uncommon. Yamamoto (1975a) demonstrated that IPNV and R. salmoninarum (bacterial kidney disease) occurred in the same population of rainbow and brook trout and electron microscopy

indicated that both pathogens actually occupied the same cell. It was noted by Evensen and Lorenzen (1997) that IPNV and Flavobacterium psychrophilum, the causative agent of bacterial coldwater disease, can coinfect juvenile rainbow trout. These disease agents can occur in the same cells but generally the virus appears in the exocrine pancreatic cells and the bacteria in the interstitial tissue surrounding pancreatic islets. These pathogens were detected by immunohistochemistry in asymptomatic fish as well as those with clinical signs of disease. It has also been shown that IHNV and IPNV occurred in the same rainbow trout fry, but concentrations of IHNV (2.8 × 105 PFU/g) were greater than IPNV (1.2 × 104 PFU/g) (Mulcahy and Fryer 1976). In Norway, both IPNV and erythrocytic inclusion body syndrome (EIBS) virus were present in Atlantic salmon (Jarp et al. 1996). Fish that had experienced an IPNV epizootic as fry in freshwater could have a recrudescence of disease when moved into saltwater even though IPNV antibody may be present. However, fish with IPNV antibodies, but no clinical IPNV in freshwater were protected against IPNV in seawater. This immunity could be important in controlling IPNV in sea farmed salmon. There was no relationship between ability to hypo-osmoregulate, infection with EIBS, or risk of clinical IPN after transfer to seawater. Transmission of IPNV occurs horizontally from infected fish through water to susceptible fish, where it is absorbed by the gills and gut. During an epizootic, virus titers in water can be as high as 105 TCID50 /mL (Barja et al. 1983). Carrier trout shed IPNV in reproductive products and feces. Vertical transmission was strongly suggested by the early work of Wolf et al. (1963) and evidence suggests the potential for it to occur via the gametes (Bootland et al. 1991). Circumstantial evidence, and some experimental data, suggests that vertical transmission is a possible but unpredictable event. IPNV can be found in ovarian fluid and has been shown to be adsorbed onto sperm

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(Mulcahy and Pascho 1984). The presence of IPNV on fresh sperm of rainbow trout was confirmed by FAT and flow cytometer analysis in Spain (Saint-Jean et al. 1993). This would further substantiate a likely mode of transmission to progeny during spawning. An unusual means of IPNV dissemination could also occur when pituitary extracts from infected trout are injected into other fish species to stimulate ovulation (Ahne 1983). Immunization of trout against IPNV is a management tool but may not prevent vaccinated fish from becoming carriers and active virus shedders (Bootland et al. 1995). Brook trout vaccinated with inactivated IPNV developed a strong humoral immune response but after injection with virulent IPNV, immunized fish and nonvaccinated controls shed virus in their feces and both male and female reproductive products contained virus. The IPNV was carried at least 15 weeks, leading to the conclusion that even though trout develop an immune response, they can still transmit virus during spawning activities. The importance of IPNV as a pathogen of cultured trout, salmon, and occasionally other species is unquestioned, but its significance in wild populations is uncertain. Survivors of IPNV epizootics are often stocked into streams or lakes for sport fishing and at times these populations become reproductively self-sustaining. Yamamoto (1975b) found that a hatchery population of brook trout had an IPNV-carrier prevalence of 90% and 2.5 years after stocking the carrier prevalence was 69%. Shankar and Yamamoto (1994) determined that 44.4% of adult lake trout in Cornwall Lake, Alberta, Canada, were infected with IPNV and virus transmission to juvenile brook trout resulted in 74% mortality. The same virus produced 10% mortality in experimentally infected young rainbow trout but was avirulent to young lake trout, implying that lake trout IPNV was not pathogenic for that species. Even though these fish were not affected by the virus, they could become reservoirs of infection for other species.

Pathology Pathological changes resulting from IPNV are typical of fish virus infections (Yasutake 1975). The most profound histopathological change, especially with Serotype A1 (VR-299) in trout, is the obliteration of most acinar tissue of the pancreas and its replacement by necrotic detritus containing fragmented acinar cells, pyknotic nuclei, and zymogen granules. In advanced cases, polymorphonuclear lymphocytes and monocytes increase, indicating presence of some inflammation. Initially, foci of necrosis are scattered throughout the acinar tissue and then rapidly spread to surrounding tissue. At the periphery of necrotic areas, many acinar cells will contain intracytoplasmic inclusions. Fatty tissue surrounding pancreatic tissue often becomes necrotic, and necrosis frequently occurs in the islets of Langerhans. Smail et al. (1992) also noted extensive pathology in pancreatic acinar cells of IPNV Serotype Sp (A2 ) infected Atlantic salmon. Scattered to extensive necrosis occurs in voluntary muscles accompanied by mild inflammation. Excretory and hematopoietic kidney tissue also becomes necrotic to varying degrees. Congestion, hemorrhage, and edema occur in the epithelium of renal tubules. Sano (1971) found congestion and necrosis in liver tissue of rainbow trout. Using rainbow trout as an experimental model, McKnight and Roberts (1976) described the sequence of histopathological changes that occurred in the pancreas, renal tissue, and intestinal mucosa. Changes in the pancreatic tissue were characterized by destruction of acinar tissue, which degenerated into an amorphous eosinophilic mass. Contrary to earlier descriptions, the islets of Langerhans and fatty tissue were normal. Hematopoietic tissue in the kidney showed focal degeneration. The intestine was more severely affected than previously reported and exhibited necrosis of the intestine, sloughing of the epithelium, and accumulation of a pinkish or whitish catarrhal exudate in the lumen all

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of which persisted to a diminishing degree for over a year. These authors postulated that intestinal lesions contributed more to the death of infected fish than did pancreatic lesions.

Significance Because mortalities in susceptible fish populations are usually moderate to high, IPNV is a significant disease of cultured fishes. Despite extensive fish health inspections, IPNV has spread to most regions of the world where trout are cultured and its geographic and host range continues to expand. New IPNV incidences may be less due to movement of infected fish than the result of the ubiquitous nature of the virus. IPNV is one of the most significant diseases in Norwegian Atlantic salmon culture, where it may cost the industry $4.5–$5 million per year (Anonymous 1994).

Infectious salmon anemia virus Infectious salmon anemia (ISA) is a highly infectious disease that can cause high cumulative losses in farmed Atlantic salmon. The causative agent, infectious salmon anemia virus (ISAV), is an Orthomyxovirus and is an OIE notifiable pathogen (OIE 2008). This important disease, initially found in Norway, was described by Thorud and Djupvik (1988).

Species susceptibility and geographical range ISAV has been found in Atlantic salmon in Norway, the East Coast of Canada (Mullins et al. 1998), Maine (Bouchard et al. 2001), the Faroe Islands (Lyngøy 2003), Scotland (Rodger et al. 1998), and Chili (Kibenge et al. 2001, Godoy et al. 2008). In Chili, the disease occurred in coho salmon. The disease generally occurs in smolts after they are transferred from freshwater to seawater cages but outbreaks have been reported in fry in freshwater

(Kibenge et al. 2001) sea-run brown trout (sea trout), which are likely asymptomatic virus carriers (Devold et al. 2000, Raynard et al. 2001). Stealhead trout, chinook, coho, and chum salmon are resistant to ISAV (Rolland and Winton 2003); however, the virus did occur in coho salmon in Chili (Kibenge et al. 2001).

Clinical signs Atlantic salmon with ISA are lethargic, often remaining near the net pen sides. The population may experience low to moderate mortality over a long period, resulting in potentially high cumulative mortality. These fish are anemic (pale gills and internal organs), exophthalmic, develop ascites, and liver and spleen are congested and enlarged. The intestine is congested and petechiae are present in visceral fat (Nylund et al. 1994). Hematocrits of infected fish are reduced from a normal of 35–48% to 12–25% (Dannevig et al. 1993). Naturally infected coho salmon in Chile demonstrated similar clinical signs with the addition of jaundice and associated yellow coloration of fin bases and belly (Nylund et al. 1999). Experimentally infected brown trout developed no mortality or clinical signs other than significantly reduced hematocrits (Nylund and Jakobsen 1995). In contrast, experimentally challenged rainbow trout experienced moderate mortality with clinical signs similar to those of Altantic salmon except moribund rainbow trout displayed hemorrhagic fin bases and exophthalmia more often (MacWilliams et al. 2007).

Diagnosis Because ISAV was recalcitrant to tissue culture in common cells lines, early diagnostic methods relied on clinical signs, pathology, and electron microscopy. Virus presence was also demonstrated by injecting ascitic fluid from the body cavity or prepared organ filtrates from suspect infected fish into suitable size, naive

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Atlantic salmon (Nylund et al. 1995a). However, efficient propagation of the virus in an Atlantic salmon head kidney cell line SHK1 (Dannevig et al. 1995) opened the way for traditional and molecular-based diagnostic assays. Sommer and Menner (1996) cultivated the virus in primary in vitro leukocyte cultures from Atlantic salmon head kidney. Sommer and Mennen (1997) demonstrated that ISAV actually replicates on the common Atlantic salmon cell line AS. It fails to produce CPE on AS cells, but replicative centers can be detected using hemadsorption with Atlantic salmon erythrocytes. Since then ISAV has been cultured in two other Atlantic salmon kidney cell lines: ASK cells (Devold et al. 2000) and TO cells (Wergeland and Jakobsen 2001). Also, many strains of ISAV can replicate in CHSE 214 cells (Bouchard et al. 1999, Kibenge et al. 2001). Comparative analysis of SHK-1, ASK, and CHSE-214 cells by (Rolland et al. 2005) showed that SHK-1 and ASK cells were similar in sensitivity but ASK cells showed CPE more quickly. They also found that CHSE-214 were much less sensitive to ISAV infection than SHK-1 or ASK cells. For cell-culture-based diagnosis, filtered kidney and spleen homogenates are inoculated on to susceptible cells and incubated at 15–16◦ C for 14 days. After five days, cultures with ISAV may demonstrate CPE consisting of vacuolated cells that round up from the growth surface. Samples that have IPNV should be reanalyzed after incubation with IPNV neutralizing antibodies. Also, cultures that do not display CPE should be evaluated by IFAT (Falk et al. 1998), hemadsorption (Sommer and Menner 1996), or RT-PCR (Kibenge et al. 2000) because ISAV may replicate without inducing CPE. Additionally viral antigens can be detected in infected tissues using IFAT on cryosection or tissue imprints (Falk et al. 1998) and RT-PCR can be done on infected tissue to detect the presence of genomic RNA (Devold et al. 2000). Detection of ISAV carrier fish has been problematic because methods require killing

the fish for virus isolation or applying RTPCR or IFAT to visceral tissue. Giray et al. (2005) proposed a nonlethal procedure of applying PT-PCR to detect ISAV in blood drawn from asymptomatic fish or on tissue of moribund fish. Snow et al. (2003) evaluated four diagnostic tests for detection of ISAV in Atlantic salmon: RT-PCR, indirect fluorescent antibody (IFAT), virus isolation in cell culture and histopathology. Fish were infected and the tests applied before clinical disease appeared. The most sensitive technique was RT-PCR (45%), followed by IFAT (15%), isolation in cell culture (14%), and detection by histopathology (negligible). An in situ hybridization probe method developed by Gregory (2002) accurately detected ISAV, while the probe did not cross–react with IPNV, IHNV, SPDV (salmon pancreatic disease virus), or VHSV. Because of the difficulty in isolation and detection of ISAV, Mikalsen et al. (2001) suggested that for maximum detection capability, regardless of procedure to be used, whole fish should be transported to the laboratory for viral assay.

Virus characteristics Electron microscopy of tissues from Atlantic salmon with clinical ISA revealed enveloped virus budding from endothelial cells of the luminal side of the plasma membrane and some intracellular membranes (Koren and Nylund 1997). The 100 nm (80-120 nm) diameter pleomorphic virion has a regularly arranged filamentous nucleocapsid and a matrix protein-like structure. The morphology and morphogenesis of ISAV resembles the orthomyxoviruses and after cell culture propagation, extensive characterization confirmed that it was a unique aquatic orthomyxovirus (ICTVdB 2008). ISAV is the type species of the recently designated Isavirus genus. The genome consists of eight segments of negativesense, single-stranded RNA. The complete genome is approximately 13,500 nucleotides

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long. Gene segments 1-6 encode single proteins P1, PB1, nucleoprotein (NP), P2, fusion protein (F), and hemagglutinin-esterase (HE) protein respectively (Clouthier et al. 2002). Segment 7 encodes three proteins (Kibenge et al. 2007) and segment 8 encodes two proteins. Genetic comparison of segment 2 and 8 from different isolates demonstrate two distinct strains of ISA designated as European strain and the North American strain (Inglis et al. 2000, Krossoy et al. 2001a). The HE gene (segment 6) is more variable and can be used to distinguish closely related subtypes (Krossoy et al. 2001b, Mjaaland et al. 2002). More recent analysis of the fusion protein gene (segment 5) demonstrates similar variability as the HE gene and combined analysis suggests genetic reassortment occurs in ISAV as has been demonstrated in other orthomyxoviruses (Devold et al. 2006). This characteristic of being able to transfer genetic material between strains along with the ability for the virus to mutate readily provides strong potential to develop into new strains.

Epizootiology When Atlantic salmon smolts are transferred from freshwater to marine culture cages, they become infected with ISAV from natural sources. It appears that brown trout serve as natural virus reservoirs in the marine environment even though they show no clinical signs of disease (Nylund and Jakobsen 1995, Devold et al. 2000). ISAV is easily transmitted from infected brown trout or Atlantic salmon to healthy Atlantic salmon by injection of visceral tissue homogenates, peritoneal ascitic fluid, blood, or cohabitation (Olsen et al. 1992; Nylund et al. 1994; Nylund et al. 1995b). Experimentally infected brown trout remain infected for at least 135 days. Mortality generally occurs as early as 18 days postinjection. Nylund et al. (1994) also suggested that sea lice (Caligus elongatus and Lepeophtheirus salmonis) may be instrumental in ISAV transmission

in epizootic and enzootic phases and that passive transmission in seawater may be of lesser importance to the spread of virus than was originally thought. However, ISAV remained infectious after 20 hours in seawater and 4 days in blood and kidney tissue stored at 6◦ C. The virus shed from infected fish increased at 7 days postinfection and a large virus shedding event takes place just before death (Gregory et al. 2009). Vertical transmission of fish viruses is always a major concern but Melville and Griffiths (1995) presented evidence that this is an unlikely event for ISAV. The virus was not detected by RT-PCR in eyed Atlantic salmon eggs or par from infected fish nor did mortalities occur among young fish injected with egg homogenates of infected eggs. Mortalities due to ISAV can be high with up to 50% occurring in naturally infected culture populations. Nylund et al. (1994) reported 6–82% cumulative mortality 30 days postexperimental infection and 45–100% mortality in these same populations 70 days postinfection. In other experimental infections using infected leukocytes from Atlantic salmon, up to 50% mortality occurred in 22 days (Dannevig et al. 1993). During cohabitation experiments, 57–60% of Atlantic salmon exposed to infected brown trout died in 48 days (Nylund and Jakobsen 1995). One experimental infection in Atlantic salmon produced 98% mortality, whereas there was no significant mortality in other infected salmon species under identical conditions (Rolland and Winton 2003). Several risk factors influenced the incidence of ISA along the Norwegian coast (Jarp and Karlsen 1997). Disease incidence was significantly higher when farms were within 5 km of a salmonid slaughterhouse and risk of infection increased by 8:1 if the culture site was closer than 5 km to an ISAV-positive site. Generally, in areas where density of seacaught fish markets was high, there was an increased incidence of ISAV. Jarp and Karlsen (1997) concluded that ISAV is mainly transmitted from infected salmonid sources to clean

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sites through seawater; therefore, suggesting that the disease would lend itself to control by appropriate location of culture operations and slaughterhouses. An analysis of ISA out breaks in net pens in the New Brunswick indicated higher ISA risk was associated with boat travel between aquaculture facilities, the presence of high numbers of wild Pollock around the cage, the stocking of large smolts, and the presence of high poststocking mortality with smolts. McClure et al. (2005) reported that more frequent sea lice treatment reduced the risk of ISA. They suggested producers use more frequent sea lice treatments, use more care in saltwater adapting smolts, avoid the larger smolts, and establish boat travel protocols that avoid travel between sites. Another study in New Brunswick found that ISA was associated with numbers of smolts stocked into a pen and the presence of more than one year class and the use of frequent sea lice treatment and extended feeding of moist pellets reduced the risk (Hammel and Dohoo 2005). Despite the close proximity between culture sites, modification of management factors would probably favorably influence mortalities caused by ISAV. On the basis of existing information and new data, it was hypothesized by Nylund et al. (2003) that the following observations are true for ISAV: (1) ISAV is maintained in wild populations of trout and Atlantic salmon; (2) The virus is transmitted between wild hosts during freshwater spawning in freshwater rivers; (3) Wild salmonids carry benign wild-type ISAV; (4) Mutation in virulence occurs in wild type ISAV; (5) ISA emerges in farmed Atlantic salmon when mutated viruses are transmitted from wild salmon or following mutation of benign virus in farmed salmon; (6) Farming activity is an important factor in transmission of ISAV between farming sites in addition to transmission from wild to farmed salmon; (7) Transmission from farmed to wild salmon probably occurs less frequently than transmission from wild to farmed salmon; (8) Frequency of new outbreaks from wild to farmed salmon prob-

ably reflects natural variations in the presence of ISAV in wild populations of salmonids.

Pathology ISAV causes a viremia. By electron microscopy, Nylund et al. (1995b) found virus in a variety of organs and tissues including the liver, spleen, anterior and trunk kidney, urinary bladder, gut, somatic muscle, and in several glands including thyroid and gonad tissue. The gross lesions are exophthalmia, ascites, congestion and enlargement of liver and spleen, congestion of the foregut, and petechiae in the peritoneum. Histologically, congestion in the liver, spleen and kidney are the predominant early lesions that progresses to hepatic necrosis, necrosis in the kidney and congestion and hemorrhage in the lamina propria of the foregut (Moneke et al. 2005). Using in situ hybridization, Moneke et al. (2005) supported earlier findings that endothelial cells are the predominant cell type infected and destruction of these cells is most likely the cause of hepatitis, nephritis, and hemorrhages throughout the fish. They found the peak, earliest, and most prolonged signal in the cardiac endothelial tissue. It was also demonstrated viral expression in hematopoietic cells and blood cells but no indication of viral replication in hepatocytes or renal tubular epithelial cells. Although high levels of virus replication occurs in the heart, histopathology generally reveals only mild cardiac lesions, predominantly, congestion. However, Gattuso et al. (2002) demonstrated substantial, decreased cardiac function during loading. The most dramatic lesion associated with ISAV is anemia (Olsen et al. 1992). There are several factors that may contribute to the anemia, hemorrhages (blood loss), destruction of erythrocytes, and damage to hematopoietic tissue. Erythrophagocytosis by spleen and kidney macrophages is prevalent in ISA-infected fish (MacWilliams et al. 2007). This may be due to external influence of the virus,

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given the hemagglutating activity of ISAV or due to the influence of virus infection and replication in the erythrocyte. Uptake into Atlantic salmon erythrocytes has been demonstrated (Workenhe et al. 2007) and subsequently the erythrocytes were shown to support viral replication (Workenhe et al. 2008). Furthermore, this replicative ability in erythrocytes was found only in the most pathogenic of the isolates tested. Analyses of liver and the immune system also suggest impaired function in fish with ISA. Atlantic salmon infected with ISAV show a significant overall increase in plasma glutathione, which may be due to intracellular release of this substance; however, there is a significant decrease in its level in the liver, which may affect the organ’s ability to transform and excrete xenobiotics from the body (Hjeltnes et al. 1992). Dannevig et al. (1993) showed that leukocytes from ISAV-infected Atlantic salmon had reduced ability to respond to mitogens, which suggest a compromised immune system, and this event was independent of anemia developing. Histopathology showed a close relationship between ISAV and hemorrhagic kidney syndrome (HKS). Bouchard et al. (1999) isolated ISAV from Atlantic salmon in New Brunswick, Canada, in which the posterior kidney had interstitial hemorrhaging, tubular epithelium degeneration, and necrosis. Pleomorphic virus particles measuring 80–120 nm were seen in electron micrographs, which were confirmed as ISAV by RT-PCR. Simko et al. (2001) detected ISAV pathonomonic lesions in renal tissue of 91% of Atlantic salmon with HKS.

Significance ISAV is a serious disease in sea-pen reared Atlantic salmon in Norway, Scotland, Chili, northeastern cost of the United States, and east coast of Canada. Furthermore, the marine source and its ability to mutate and reassort genomic segments gives this pathogen substan-

tial potential to evolve into new antigenic and pathogenic types similar to orthomyxoviruses of mammals and birds. In the USA, USDA APHIS has an active eradication program to limit the spread of this virus.

Viral hemorrhagic septicemia Viral hemorrhagic septicemia (VHS) is a highly infectious viral disease that, in its acute form, can cause high losses in a short period. Traditionally VHS was considered a disease of cultured rainbow trout in Europe. However, this disease is now known to cause important losses in farmed turbot and Japanese flounder and many species of wild marine and freshwater fish. It has also become problematic among non-salmonid wild fish in North America. VHS has a long history in trout aquaculture in Europe with references to the disease in the mid-1930s (Wolf 1988). It was originally named Egtved disease after the village in Denmark from which it was first reported (Jensen 1965). There have been many common names associated with the disease, most of which reflect its viral nature or its kidney and liver tropism. Viral hemorrhagic septicemia virus (VHSV) is a rhabdovirus in the Novirhabodvirus genus and is an OIE notifiable pathogen (OIE 2008).

Geographical range and species susceptibility Following establishing its viral etiology in the early 1960s (Jensen 1965), clinical VHS was confined to the European continent until the virus was isolated from adult Pacific salmon returning to hatcheries in Washington state in 1988 (Hooper 1989; Winton et al. 1991). Wolf (1988) listed 13 European countries where VHSV had been found and the virus has since been found in the UK, Turkey, Spain, Canada, Japan, Korea, and the Great Lakes region and west coast of the United States and Canada.

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Table 8.5 Distribution of VHSV genogroups. Genogroup

Endemic Regions

Ia Ib Ic Id Ie II III IVa IVb

Freshwater—Continental Europe and UK Marine—Europe: Baltic Sea to the North Sea and English Channel, Japan Freshwater—Denmark Brackish water aquaculture, Gulf of Finland Black Sea—Turkey, Georgia Marine—Baltic Sea Marine—North Sea and coastal areas of UK and Ireland, N. Atlantic near Newfoundland Marine—Pacific Coast of North America, Japan and Korea Freshwater—Great Lakes, St. Lawrence River, Coastal rivers of Eastern Canada

Source: Snow et al. 1999; Nishizawa et al. 2002; Nishizawa et al. 2006; Einer-Jensen et al. 2004; Elsayed et al. 2006; Stone et al. 2008.

Although rainbow trout is the most susceptible species to the European freshwater strain of VHSV (genogroup 1a), the brown trout (sea trout) is considered a natural host. The virus has been isolated from at least 70 fish species and of those clinical signs of VHS were seen in 48 species (Skall et al. 2005) (Table 8.5). Dorson et al. (1991) demonstrated that rainbow X brook trout hybrids, and Arctic char are resistant to VHSV, but lake trout are susceptible and exhibit clinical disease and significant mortality. It was reported by Meier (1985) that under experimental conditions, juvenile northern pike were more susceptible to VHSV than were rainbow trout. At least 13 fish species, including some hybrid salmonids, are experimentally or naturally susceptible to the virus in which it causes overt disease and 9 additional species or hybrids that are refractive to the virus (Wolf 1988). The list of susceptible species includes eight salmonid and five non-salmonid species, including the marine seabass and turbot (Castric and de Kinkelin 1984). In 1990 and 1991, VHSV was isolated from Pacific cod with ulcerative lesions in Prince William Sound, Alaska, and infected Pacific herring showed similar lesions (Meyers et al. 1994). Wolf (1988) also listed nine fish species that are refractive to overt VHSV disease. Ironically, this list included chinook (Ord et al. 1976) and coho salmon (de Kinkelin et al. 1974); the two species in Washington from which VHSV was isolated in 1988. Of particular note are farmed Japanese

flounder in Japan (Muroga et al. 2004) and farmed turbot in Europe (King et al. 2001). Upon the discovery of VHSV in the Great Lakes region of North America (Elsayed et al. 2006; Lumsden et al. 2007), the number of fish species in which it causes disease increased as did the list of species that are susceptible without clinical disease (Table 8.5). Groocock et al. (2007) noted 15 fish species in the Great Lakes area from which VHSV was isolated; however, fish in many of these cases did not show overt disease. Clearly VHSV does not have a limited species susceptibility or geographical range.

Clinical signs VHS of salmonids occurs as an acute, chronic, or nervous form (Table 8.6). The description of each reflects behavior, overt pathological changes, and mortality pattern, which typify each particular form of the disease. Typically, infected salmonids do not feed and behavior ranges from lethargic to hyperactive. High mortality is associated with acute VHS. Affected rainbow trout swim erratically, have dark skin, exophthalmia, and pale gills may be flecked with petechia (Figure 8.5). Internally, there are multiple hemorrhages in the skeletal muscle and swim bladder, the kidney is swollen and hyperemic with liquefactive necrosis, and the liver is either hyperemic, grey, or yellowish. Subacute to chronic VHS is

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Table 8.6 Species of fish from which viral hemorrhagic septicemia virus has been isolated. Those marked with (∗) have exhibited clinical signs of disease. Virus isolated from others but not mortality. ∗

Atlantic cod Atlantic herring ∗ Atlantic salmon ∗ Black crappie ∗ Blackcod ∗ Brook trout ∗ Brown trout ∗ Burbot ∗ Emerald shiner ∗ Freshwater drum ∗ Gizzard shad ∗ Golden trout ∗ Grayling ∗ Haddock ∗ rainbow trout X Coho salmon ∗ Japanese flounder ∗ Lake trout ∗ Mummichog ∗ Muskellunge ∗ Northern pike ∗ Pacific cod ∗ Pacific hake ∗ Pacific herring ∗ Pacific sardine ∗ Pilchard ∗ Rainbow trout ∗ Round goby ∗ Sea bass ∗ Smallmouth bass ∗ Striped bass ∗ Surf smelt ∗ Three-spined stickleback ∗ Turbot ∗ Walleye ∗ Walleye pollock ∗ Whitefish ∗ Yellow perch ∗ Black rockfish ∗

Blue whiting Bluegill Bluntnose minnow Brown bullhead Channel catfish Chinook salmon Chub roach Coho salmon Common carp Common dab Emerald shiner Eurasian perch European eel European flounder European plaice European sprat Four bearded rockling Goldfish Greenland halibut Japanese sandlance Largemouth bass Lake whitefish Lesser argentine Northern Short nose redhorse Norway pout Pacific mackerel Pacific tomcod Poor cod Pumpkinseed Rock bass Smelt Sand eel Sand goby Silver redhorse Spottail shiner Tench White perch

Source: Wolf 1988; Groocock et al. (2007).

characterized by lower mortality. Affected fish have dark pigmentation (melanosis), exophthalmia, and swollen abdomens. Internally, the liver is pale. In the nervous stage (neurovegetative or N-form), there is very low mortality but fish exhibit poor balance, swim in a circle, and are somewhat anemic. Either environmental stress or handling can cause a less severe form of VHS to transition into a more severe one. Clinical signs of the VHSV-infected fishes in the Great Lakes region were similar

to those previously described for the disease in Europe. Non-salmonids with VHS show similar clinical signs. Naturally infected whitefish developed hemorrhages in the skeletal muscle and swim bladder (Ahne and Thomsen 1985). Naturally infected pike had exophthalmia, extensive hemorrhages in the skin, muscle, and kidney (Meier and Jørgensen 1980), and the body cavity contained ascitic fluid. Meier and Wahli (1988) found that

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(a)

(b)

(c)

Figure 8.5 Fish infected with viral hemorrhagic septicemia infection. (a) Hemorrhage (arrow) in skin of rainbow trout. (b) Petechia in muscle, peritoneum, and pale liver and gills in rainbow trout. (Photos (a) and (b) by P. Ghiteno; photo B reprinted with permission of CRC Press). (c) Hemorrhage in the muscle and pale gills of pike infected with VHSV. (Photo courtesy of W. Ahne.)

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in Switzerland, VHSV-infected grayling exhibited anemia, enteritis, and subcutaneous and intramuscular hemorrhage. Naturally infected freshwater drum from Lake Ontario displayed fin and skin hemorrhages, enteritis, exophthalmia, ascites, and diffuse vasculitis (Lumsden et al. 2007). Naturally infected round gobies from Lake Ontario and the St. Lawrence River displayed anemia and scattered hemorrhages in the skin, fins, viscera, and musculature (Groocock et al. 2007).

Detection During an epizootic, VHSV can be readily isolated in cell cultures from clinically diseased fish, but in later stages of infection, isolation becomes more difficult. Shortly after clinical disease disappears, virus can often no longer be isolated from survivors. Kidney and spleen are the organs of choice when attempting VHSV isolation (Meier and Jørgensen 1980; Meier and Wahli 1988). In Europe, the BF-2 cells are most susceptible to VHSV, where as FHM and EPC cells are more susceptible to the North American strains of VHSV (Winton et al. 2007), but CHSE-214, RTG-2, or STE137 cells may also be used. It is important to maintain media at pH 7.6–7.7 and an incubation temperature of 15◦ C or less (Jensen 1965). Virus adsorption to cells can be enhanced by pre-treating the cells with a 7% solution of 20,000 MW polyethylene glycol or adding it to the virus diluent (Batts and Winton 1989, 2007). Focal CPE consists of rounded cells followed by lysis and involvement of the entire cell sheet (Figure 8.6). Positive VHSV identification is obtained by plaque reduction, virus serum neutralization, ELISA, FAT, or RT-PCR in conjunction with virus-specific reagents. FATs may be used to identify VHSV in tissue culture or frozen tissue sections from infected animals. Ahne et al. (1986) reported a VHSV strain that could not be neutralized by VHSV antiserum but was positive by FAT.

In a given population, VHSV may coexist with IPNV or IHNV in the same fish; therefore, identification for all three viruses should be used when assaying trout and salmon populations, particularly when fish originate from an area where all three viruses are endemic. Furthermore, in a coinfection one virus can suppress the replication of other viruses. A study examining competition for receptors as possible cause of this inhibition was conducted by de las Heras et al. (2008). They found that uptake of VHSV by BF2 cells is inhibited by IHNV but not IPNV. Also, VHSV inhibits IHNV uptake. Because of this inhibitory effect, if a sample cultures positive for one virus, the sample should be pretreated with neutralizing antibody specific for that virus and then the sample should be recultured to be sure that another virus is not present. IHNV and VHSV were isolated from coho salmon in the same watershed in the Pacific Northwest of North America (Eaton et al. 1991b). Monoclonal antibodies, which do not bind with any other fish rhabdovirus and can identify a conserved epitope on VHSV or IHNV, can be used to easily distinguish the two, regardless of origin (North America or Europe), when employed in an immunodot assay (Ristow et al. 1991). Olesen and Jørgensen (1991) emphasized that the ELISA test for VHSV detection is a valuable tool when rapid diagnosis is essential but should be used only in conjunction with cell culture. These researchers found that 24 of 40 tested trout were VHSV positive by culture, but only 80% of the 24 culture positive fish were positive by the ELISA. Detection of VHSV in tissues with avidin–biotin alkaline phosphatase was compared to virus isolation in BF-2 cells by Evensen et al. (1994), who found that about 1/3 fewer positive fish were detected by immunohistochemistry than by virus isolation in asymptomatic experimentally infected rainbow trout. Some have proposed that immunohistochemistry and other rapid diagnostic procedures be used to replace more labor intensive virus isolation in

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(a)

(b)

Figure 8.6 Viral hemorrhagic septicemia virus growth in FHM cells. (a) Focal CPE consisting of rounding, and pyknotic cells. (b) Advanced CPE involving entire cell sheet (iodine stain). (Photos courtesy of P. de Kinkelin.)

tissue culture, but this has not been universally adopted. Detection of VHSV antibody by virus neutralization, FAT, or ELISA is used for determining possible virus carrier trout populations in Europe. Olesen et al. (1991a) noted that the ELISA technique was more accurate than serum neutralization or FAT, but less specific than serum neutralization. Like IHNV, VHSV neutralization is dependent on com-

plement. After IHNV was found in France, Hattenberger-Baudouy et al. (1989) serologically surveyed an IHNV-infected rainbow trout population, which also carried VHSV, and found that some fish had both neutralizing antibodies. Because of potential crossreactive antibody in the same fish Jørgensen et al. (1991) cautioned against using antibody detection in fish sera to indicate presence of either VHSV or IHNV in populations

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where both viruses may coexist. More recently, Knusel et al. (2007) found higher numbers of samples testing positive for VHSV neutralizing antibodies than by RT-PCR or cell culture. The use of RT-PCR directly on tissues has become popular for screening for the presence of VHSV in diagnostic cases and in surveys. The OIE and AFS bluebook accepted RT-PCR method uses primers that target a conserved portion of the nucleocapsid gene and this can be used for confirmation on cell culture isolations (OIE 2006; USFWS and AFS-FHS 2007). The direct use of this method on fish tissues was as sensitive as virus isolation on cell culture (Knusel et al. 2007). An even more sensitive realtime RT-PCR method has been developed and evaluated for European strains of VHSV (Matejusova et al. 2008). Currently, RT-PCR is the procedure of choice in terms of accuracy and sensitivity.

Virus characteristics VHSV, a member of the family Rhabdoviridae, is an enveloped, bullet-shaped particle measuring about 65 nm in diameter and 180 nm in length (Zwellenberg et al. 1965). The virus has a single stranded RNA genome that is about 11,000 bp in length and encodes 6 genes including the NV gene. This makes it a member of the Novirhabdovirus genus along with IHNV. Similar to other rhabdoviruses,

the VHSV virion is inactivated by glycerin, heat, ether, and acid, but is stable at pH 5–10 (Ahne 1982). The virus is completely inactivated by 3% formalin, 2% NaOH, and 0.01% iodine in 5 minutes and in 2 minutes by 500 mg/L of chlorine. VHSV is 90% inactivated after 14 days in stream or tap water at 10◦ C, 99% inactivated in mud at 4◦ C after 10 days, and drying at 15◦ C in 14 days. Gamma and UV irradiation completely inactivates the virus (Ahne 1982). Three serological VHSV strains were identified by Le Berre et al. (1977); however, Olesen et al. (1991b) identified four strains using monoclonal antibodies in neutralization tests. There are three recognized serotypes of VHSV (F1, F2, and 23.75) plus the North American isolate (Le Berre et al. 1977). Serological cross-reactivity occurs among the three serotypes in IFAT assays and in some neutralization tests (Meier and Jørgensen 1980). McAllister and Owens (1987) identified all three serotypes by immunoblot ELISA with antiserum to F1 serotype. Four VHSV strains are defined by genotyping appear to be of regional origin (Table 8.7) (Nishizawa et al. 2002; Thiery et al. 2002; Einer-Jensen et al. 2004, 2005; Skall et al. 2005 ): Genogroup I is from freshwater and marine sites in Europe; Genogroup II is from marine fish in the Baltic Sea; Genogroup III is from marine fish in the North Sea and North Atlantic as far west as the Flemish

Table 8.7 Characteristics of the three different disease forms of viral hemorrhagic septicemia in salmalmonids. Form

Behavior

External Signs

Internal Sesions

Acute

Poor feeding, erratic swimming, high mortality

Dark pigmentation, anemia, pale gills, exophthalmia

Multiple hemorrhages in skeletal muscle, eye, viscera; pale, gray liver; hyperemic kidney; swollen intestine, void of food

Chronic

Lower mortality, reduced activity

Dark pigmentation exophthalmia, some hemorrhage

Liver pale to grayish; kidney not smooth

Nervous

Negligible mortality, erratic swimming

Source: Ghittino (1965).

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Cap near Newfoundland (Dopazo et al. 2002; Lopez-Vazquez et al. 2006); Genogroup IVa is of marine origin in anadromous fish on the West Coast of North America, Korea, and Japan; and Geneogroup IVb is from freshwater fish in the Great Lakes region and the eastern coastal areas of Canada. An exception is the identification of Genogroup I in Japan, which appears to have been introduced from Europe (Nishizawa et al. 2002).

Epizootiology Originally VHS was thought to be a nutritional disease; however, this proved to be incorrect when Jensen (1965) isolated the VHSV agent and a more detailed description of its etiology was made by Zwellenberg et al. (1965). Winton et al. (1991) showed that VHSV isolates from chinook and coho salmon in Washington were similar serologically and biochemically to European isolates. It was later determined, however, that this strain was less virulent to salmonids than VHSV strains from Europe. The Washington VHSV strain did not produce significant mortality in any of eight different salmonid species (including rainbow trout) exposed to it by waterborne exposure or injection. In comparative susceptibility studies, Follett et al. (1997) found that coho, chinook, pink, and sockeye salmon were refractory to the North American VHSV strain following waterborne virus exposure, but rainbow trout developed clinical signs and disease with 12% virus specific mortality. These data indicate that the North American strain was different pathogenically from typical European strains. An additional difference in the two strains was noted by Bernard et al. (1991), who compared the sequenced nucleoprotein genes of the North American strain and a European strain. This difference seems to suggest that the North American strain is not of European origin. Attempts were made to eliminate VHSV from the continental United States in 1988 by destroying infected fish populations and disinfecting

facilities, but later it was determined that the virus was enzootic to the west coast of North America and no more eradication efforts were attempted. According to Meyers et al. (1994), VHSV is widely indigenous to Pacific herring populations in the Pacific Northwest of the United States. This virus has been associated with mass mortality event in 1998 of pacific herrings, pacific hake and walleye Pollock in Alaska (Meyers et al. 1999) and in two mortality events of Pacific sardines (pilchards) in British Columbia (Hedrick et al. 2003). In both cases, the herring and/or pilchard were subjected to unusual environmental stressors before the outbreaks occurred. These infected forage fish may have been the source for the VHS outbreaks in the other species. There is no conclusive evidence that VHSV is transmitted vertically. However, VHSV was isolated from both milt and ovarian fluid of spawning salmonids in North America as well as from reproductive products and kidneys of coho salmon in the same water shed in Washington (Eaton et al. 1991b). Attempts to infect eggs experimentally with the virus were unsuccessful. In a hatchery environment, VHSV can be transferred on surfaces of animate and inanimate objects and persists in water for several days. Kocan et al. (2001) studied VHSV survival in several media. The virus survived 72 hours and 96 hours with 0.01% and 1.0% salmonid ovarian fluid, respectively. These data suggest ovarian fluids prolong virus survival and enhance potential for vertical transmission. The virus also had 100% survival for 15 days in cell culture media supplemented with 10% fetal bovine serum. The virus is transmitted by injection, brushing the gills with infected material, cohabitation of naive fish with infected fish, and by waterborne exposure (de Kinkelin et al. 1979). Oral infection is difficult, although pike fry were infected when fed VHSV contaminated trout flesh (Ahne 1985). Mortality in infected fish varies with species, age, and size of fish, method of infection, and water temperature. In immersion challenge

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experiments, virus isolates from turbot did not cause mortality in rainbow trout but all isolates from farmed rainbow trout caused significant mortality (Skall et al. 2004). Incubation time between exposure and overt appearance of VHS is temperature dependent and can vary from 5 days to 4 weeks (Jørgensen 1973). VHS occurred 5 days postinjection at 9◦ C and 8–10 days following exposure by immersion at 12◦ C. According to de Kinkelin (1983), incubation time was 3–10 days at 7–14◦ C and a subclinical infection eventually occurred at 2◦ C. Rasmussen (1965) found that VHSV outbreaks in rainbow trout were associated with low water temperatures of 5–10◦ C and that water temperature directly affected duration and severity of disease. Optimum water temperature for VHSV is between 8 and 12◦ C and temperature fluctuations of a few degrees seem to trigger death. Clinical disease in hatchery populations normally does not occur at temperatures above 14◦ C. The younger the fish, the more susceptible they are to VHSV. In trout populations, rainbow trout that weigh 0.5–4 g are most susceptible (de Kinkelin et al. 1979). Although resistance to VHSV increases as fish become larger but trout up to 200 g may develop VHSV. Typical of several fish viral diseases, the healthier appearing individuals in a population are the first to show clinical disease. It was also noted that VHS-associated mortality was 80% in rainbow trout 1 month after they were moved from freshwater to full strength seawater. Also, in Danish mariculture facilities, VHS-related mortalities of 15–50% occurred in naturally infected rainbow trout 18 months after stocking (Horlyck et al. 1984). In a study in Switzerland, Meier and Wahli (1988) noted that mortality in both rainbow trout and grayling was 56–100% respectively, depending upon the virus, fish strain, and the environment. It has been postulated that up to 30% of cultured trout in Italy are killed by VHSV. Seabass and turbot injected with VHSV experienced over 90% mortality (Castric and de Kinkelin 1984). Meier et al. (1986) reported

lower mortalities in whitefish (28–50%) than in rainbow trout (40–80%) at temperatures of 11–14◦ C. Although mortalities from VHSV in nonsalmonid fish varies, Ahne (1985) induced 90% mortality in 0.2 g northern pike infected by waterborne exposure and when these virusinfected pike were fed to 1-year-old, 60 g pike, clinical disease and 30% mortality occurred. In the Pacific, VHSV was isolated from epidermal ulcerative lesions on Pacific herring, but its role in these lesions was unknown (Meyers et al. 1994). However, the herring population was diminished by 1/3 during the time disease was occurring. It was speculated that VHSV is simply an opportunistic pathogen that is indigenous to Pacific herring. Also, two mass mortality events of Pacific sardines (pilchards) were attributed to VHS in British Columbia (Hedrick et al. 2003). In these cases, the herring and/or pilchard were subjected to unusual environmental stressors before the outbreaks occurred. On the other hand, VHSV-induced mass mortality events in freshwater drum, yellow perch, emerald shiners, gizzard shad, round goby, and muskellunge in the Great Lakes region suggest that stain IVb is more virulent. These massive mortality events were not associated with obvious stressing events (Groocock et al. 2007; Lumsden et al. 2007). Furthermore, a molecular epidemiology study of isolates from these outbreaks showed very little genetic differences, suggesting that the losses in different species in different lakes are components of one large epizootic (Winton et al. 2008). VHSV was first discovered in the Great Lakes in 2005, and by the end of 2007, it had been found in all of the Great Lakes except Lake Superior (Winton et al. 2008). In 2009 it was found in Lake Superior (P. R. Bowser, Cornell University College of Veterinary Medicine, Ithaca, New York, personal communication). Over 800 individual fish of multiple species were collected during routine sampling from various sites on the lake and tested for the virus using the highly sensitive

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qRT-PCR. The results were confirmed by a second laboratory using a different PCR-based testing method followed by partial sequencing of the PCR product. There were no clinical signs associated with the presence of virus and no indication that VHSV caused mortality in Lake Superior fish but presence of the virus in a covert carrier state indicates that this could serve as a VHSV reservoir. According to P. R. Bowser, “This is important because it suggests that these infected fish may serve as a reservoir for the virus in the Great Lakes ecosystem. While we don’t fully understand the lack of recent mortality events, the potential presence or absence of concurrent stressors on the fish may be playing a role.” The kidney, spleen, brain, and digestive tract are sites where VHSV is most abundant and clinically infected fish shed virus in the feces, urine, and reproductive fluids. Although survivor fish generally do not shed virus, a pathogen source during absence of clinical disease can likely be attributed to an asymptomatic infected fish species (Jørgensen 1982). Wolf (1988) noted that more VHSV-resistant salmonids (brown trout) may be virus reservoirs but as shown with isolates from turbot and cod, these isolates are often avirulent. Jørgensen (1982) reported that VHSVinfected rainbow trout (45% of those tested) were found in a stream below the outfall of a hatchery in Denmark, but it could not be determined if infected fish were hatchery escapees or had become infected via contaminated water. Fish above the hatchery were not infected. Opinions differ as to whether birds are a virus source. Eskildsen and Jørgensen (1973) reported that VHSV was not present in the feces of 41 sea gulls but other reports claim that herons can be vectors for the virus.

Pathology VHSV produces hemorrhages, usually petechiae, throughout the musculature, visceral organs, and gills of affected fish (Yasutake

1975; Meier and Jørgensen 1980). These hemorrhages occur because the endothelial cells are major targets for VHSV replication (de Kinkelin et al. 1979). Hematopoietic tissue of the kidney is severely affected and is characterized by hyperplasia and necrosis. Melanomacrophage centers are destroyed and leukopenia is present along with congestion. The liver exhibits focal necrosis and degeneration with vacuolated pyknotic and karyolytic hepatic cells. Although small hemorrhages are present in the musculature, muscle cells are essentially normal. In natural VHSV infections, gill epithelium is a possible route of virus entry, where it may or may not replicate (Neukirch 1984). Virus does replicate near its portal of entry in blood capillary walls of the skin and underlying muscle and is then transported to the kidney and spleen where more active replication occurs. According to de Kinkelin et al. (1979), the time between waterborne exposure to VHSV and recovery of the replicated agent from internal organs is about 24 hours. Peripheral blood of VHSV infected rainbow trout is adversely affected and may be ˇ 2003). instrumental in fish death (Rehulka Acute-infected fish with clinical signs had significantly lower red blood cell counts, hematocrits, and hemoglobin levels. These fish also had less total protein, creatine, glucose, triacylglycerol, inorganic phosphate, total calcium, and sodium than noninfected fish. Fish with chronic VHS, showing minor clinical signs were in worse condition: reduced biochemical parameters and a higher hepatosomatic and visceral somatic index.

Significance Historically, VHSV has been the most serious viral disease to affect European trout culture, where it continues to be a major health concern. With its detection in Japan and Korea, along the northwest coast of North America and now its presence in the Great Lakes region of North America, new emphasis has been

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placed on its potential to affect salmon culture and other species of cultured and wild populations throughout the world, particularly the northern hemisphere. The fact that VHSV is established in nonsalmonid marine species and infects a wide variety of freshwater species emphasizes its importance in aquatic resources. It is now recognized that where VHSV occurs, there is always the possibility of a highly communicable disease outbreak among various fish species. The spread of the virus throughout the Great Lakes region and the frequent observation of VHS associated fish kills are especially troubling. The recent detection of VHSV in Lake Superior means all of the Great Lakes are VHSV endemic. Recreational fishing contributes significantly to the economy of the Great Lakes states and the potential is great that the virus could negatively impact the recreational fishery (P. R. Bowser, personal communication). He states that, “On a worldwide basis, VHSV is considered one of the most serious pathogens of fish because it kills so many fish, is not treatable, and infects a broad range of fish species.” Therefore, a maximum effort should be expended to manage and prevent the spread of this virus. Currently, however, VHSV is not a problem in salmon culture in the Pacific Northwest because the indigenous strain appears to be avirulent to salmonids. VHS is also important as an OIE reportable disease and occurs on the United States Title 50 list (American Fisheries Society 2006; OIE 2008). This virus has caused great economic impact because various regulatory bodies are focused to prevent its spread. Regulations on transport of live fish from endemic areas have had substantial regional impacts on the marketing of aquaculture products.

Erythrocytic inclusion body syndrome Erythrocytic inclusion body syndrome (EIBS), a viral infection causing anemia, was first rec-

ognized in 1982 in cultured coho and chinook salmon, in the northwestern United States (Holt and Rohovec 1984; Leek 1987). The disease is caused by the erythrocytic inclusion body syndrome virus (EIBSV).

Geographical range and species susceptibility EIBS was originally identified in cultured coho and chinook salmon along the Columbia River and its tributaries in Oregon and Washington (Holt and Rohovec 1984; Leek 1987; Rodger 2007). Since then it has been recognized in freshwater reared Atlantic salmon in Norway (Lunder et al. 1990), freshwaterand seawater-reared Atlantic salmon and rainbow trout in Iceland and Scotland (Rodger et al. 1991; Rodger and Richards 1998) and in Pacific salmon in Japan (Takahashi et al. 1992b) and Alaska (Meyers 2007). Okamoto et al. (1992) found that chum and masu salmon are as susceptible to EIBS virus as are coho salmon. In experimental exposures, rainbow and cutthroat trout become infected, but disease severity is mild compared to that in Pacific salmon (Piacentini et al. 1989).

Clinical signs Other than severe erythrocytic anemia and pale gills and internal organs, no behavioral or external clinical signs were described (Holt and Rohovec 1984). Takahashi et al. (1992a) found that EIBS-infected fish had hyperemia of the intestine, dilation of the spleen, hyperemia and hemorrhage of the atrium, mucus in the stomach, and yellowish liver. Graham et al. (2002) reported necrosis of the hematopoietic tissue and hyperplasia of the gills. Cells of these affected tissues contained virus particles in electron micrographs. Infected erythrocytes develop cytoplasmic inclusions that stain more lightly than the cell nucleus (Figure 8.7). Hematocrits of normal salmon are 40–45%, whereas those of EIBS-infected fish drop to the mid-30% range and can plummet to 15%

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(a)

(b)

Figure 8.7 Erythrocytic inclusion body syndrome (EIBS) of salmon. (a) Electron micrograph of EIBS virus within membrane-bound organelle (Arakawa et al. 1989; reprinted with permission of Springer-Verlag Publishing Co.). (b) Erythrocytes with multiple cytoplasmic inclusion bodies (arrow) typical of EIBS infected fish. (Photo courtesy of S. Leek.)

and to 1–4% late in an infection. Rodger (2007) found no clinical disease or hematological changes associated with presence or absence of viral inclusions in red blood cells of EIBSV-infected Atlantic salmon.

Diagnosis EIBS is diagnosed by staining erythrocytes with pinacyanol chloride or Leishman-Giemsa

to detect basophilic cytoplasmic, round to ovoid inclusion bodies that measure 1–8 µm in diameter and appear as either large single inclusions or as smaller multiple inclusions (Figure 8.7) (Leek 1987; Arakawa et al. 1989). These inclusions stain pale purple, lavender, or pink. Their presence in electron micrographs is confirmatory of EIBSV. The virus was not isolated in any of nine different piscine cell lines (Leek 1987; Piacentini et al. 1989).

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Virus characteristics The EIBS virus is distinctly different from the iridovirus of viral erythrocytic necrosis. Because of its size, possession of a lipid envelope, and its presence in organelles, the EIBS virus most resembles members of the family Togaviridae (Arakawa et al. 1989; Rodger 2007). Membrane-bound inclusions in the erythrocytes contain spherical viral particles with an icosahedral core that measure 75–100 nm in diameter (Figure 8.7). The viruses, which consists of a single strand RNA, are either associated with viroplasm-like material or are contained within membrane-bound organelles.

Epizootiology An EIBS infection is not necessarily lethal but mortality can occur during severe anemia. In fact Rodger (2007) reported that of the 4–28% of Atlantic salmon that were EIBS positive by electron microscopy they demonstrated neither hematological nor clinical disease. Piacentini et al. (1989) described five disease stages of EIBS when fish were held at 12◦ C; each of these stages is characterized by specific changes in hematocrit values and presence of erythrocytic cytoplasmic inclusions (Table 8.8). The entire cycle of infection from exposure to recovery required about 45 days, after which surviving fish appeared normal and were resistant to reinfection. Takahashi et al. (1992b) described similar stages of EIBS development in Japanese sea-cage cultured coho salmon at 10◦ C.

Although Takahashi et al. (1992b) attributed a mortality of about 23% directly to EIBS, affected fish were severely infected with opportunistic pathogens (i.e., external fungi) and as a result suffered much higher than normal mortality. The majority of deaths attributed to EIBS are thought to be opportunistic bacterial and fungal infections that complicate the disease process. Fish infected with EIBS generally survive if they do not succumb to secondary pathogens. However, Takahashi et al. (1992a) believe that EIBS is a major contributor to increased mortality of saltwater net-cage-reared coho salmon in Japan where the virus is responsible for great economic losses to aquaculturists. They also reported that water temperature affects the progression and duration of EIBS; the disease cycle begins more quickly at 15–18◦ C (5 days) and is more prolonged at 6–9◦ C (32 days). Atlantic salmon that suffer from EIBS in freshwater appear to be less susceptible to recrudescence when transferred to seawater (Jarp et al. 1996). Fish classified as EIBSpositive prior to seawater transfer suffered 0.5% cumulative mortality but fish groups that were EIBS-negative in freshwater had 4.9% mortality in seawater. Exposure to EIBS virus in freshwater appears to provide some protection after smoltification. Experimental transmission of EIBS is achieved by injecting naive fish with filtrates (0.22 µm) of homogenized kidney, spleen, or blood from afflicted fish (Piacentini et al. 1989). Typical EIBSV inclusions appear in erythrocytes in about 14 days. The disease was

Table 8.8 The progressive stages of epizootic erythrocytic inclusion body syndrome in coho salmon held at 12◦ C. Stage

Days Post Infection

Hematocrit (% Packed RBC )

Cytoplasmic Inclusions

Fish Condition

I II III IV V

10 10–13 18–25 30–40 45

45 35–40 25+ 35–40 45

None Low High Low None

Normal Normal RBC Lysis Recovery Normal

Source: Piacentini et al. (1989).

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also successfully transmitted to coho salmon by cohabitation with infected fish in 14–28 days. A wide range of fish sizes are susceptible to the EIBS virus. Okamoto et al. (1992) found that 1.2–220 g coho salmon are equally sensitive. Takahashi et al. (1992a) speculated that the EIBS virus could have been introduced into Japan via imported infected eggs.

Pathology Pathology of EIBS has not been fully described but the major histopathology change is lysis of infected erythrocytes. Approximately 35% of erythrocytes and hemoblasts of infected fish will have cytoplasmic inclusions (Takahashi 1992a). The number of erythrocytes in the anterior kidney is reduced and the parenchyma and hematopoietic areas show necrosis and congestion. Phagocytes, many containing eosinophilic vacuoles, infiltrate the kidney and epithelial cells of the renal tubules show hyaline droplet degeneration. Similar pathology occurs in the spleen with cellular vacuolation, while limited necrosis of hepatocytes occurs in the liver sinusoids. Graham et al. (2002) reported on an EIBS outbreak in freshwater-reared Atlantic salmon in which the affected fish demonstrated hyperplasia of the gill epithelium with fusion of lamellae and extensive necrosis of the hematopoietic centers of the spleen and kidney. They indicated that phagocytolytic syndrome described previously was likely caused by the EIBS virus. Watanabe et al. (2006) investigated viruses associated with heart and skeletal muscle inflammation, a disease that is characterized by cardiac muscle degeneration and inflammation, in Atlantic salmon smolts. They found a consistent association with EIBS virus.

Significance Alone, the EIBS virus may not be a highly significant pathogen, but it reduces the host’s nat-

ural resistance to other secondary infectious agents that cause EIBSV-affected fish to die. In some aquaculture situations, EIBS is thought to cause significant economic losses (Takahashi et al. 1992b).

Salmonid herpesviruses As many as 10 herpesviruses have been isolated from trout and salmon (Tanaka et al. 1992); however, the two United States isolates are considered the same [salmonid herpesvirus 1 (SalHV-1)], and three from Japan are the same [salmonid herpesvirus 2 (SalHV-2)]. Five other isolates from coho and masu salmon in Japan are unresolved. Since SalHV1 and SalHV-2 are distinctly different viruses and cause distinct clinical disease, they are discussed separately. A third salmonid herpesvirus, Epizootic epitheliotropic disease virus, causes disease in lake trout. It has been partially characterized but not cultured.

Salmonid herpesvirus 1 Salmonid herpesvirus 1 (SalHV-1) causes a mild viral disease of rainbow trout first described by Wolf and Taylor (1975), who named the virus Herpesvirus salmonis. Isolation of a very similar herpesvirus from steelhead trout was reported and designated salmonid herpesvirus 1 (Hedrick et al. 1986).

Geographical range and species susceptibility The original SalHV-1 isolate was made in the state of Washington (Wolf and Taylor 1975) and later the virus was isolated from fish in seven hatcheries and two lakes in California (Eaton et al. 1989). Although steelhead and rainbow trout are natural hosts of SalHV-1, chum salmon fry and chinook salmon are experimentally susceptible, while coho and Atlantic salmon, and brook and brown trout are resistant.

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Clinical signs Adult fish from which the original SalHV-1 isolate was obtained showed distinctly darkened pigmentation that was the only clinical sign noted. Approximately 2 weeks after experimental exposure, juvenile rainbow trout stopped eating, became lethargic, laid on the tank bottom, and responded to physical stimulation with erratic swimming (Wolf and Smith 1981). Some fish also became dark and most developed exophthalmia accompanied by hemorrhage. Gills were pale, abdomens were distended, hemorrhages developed on the fins of some fish, and mucoid fecal casts were evident. Internally, the peritoneal cavity had abundant ascites that occasionally was bloody and gelatinous. The intestines were flaccid, the liver hemorrhagic, mottled or friable, and the kidneys pale.

Diagnosis Salmonid herpesvirus 1 disease is diagnosed by isolation of virus in CHSE-214, rainbow trout fin (RTF-l) and RTG-2 cells; however, three nonsalmonid cell lines (BB, BF-2 or FHM) are refractive to the virus (Wolf and Taylor 1975). Virus replication occurs at 5–10◦ C. Cytopathic effect, consisting principally of syncytia, was depressed or variable at 15◦ C with complete inhibition at higher temperatures. Identification of SalHV-1 is obtained by serum neutralization.

Virus characteristics Salmonid herpesvirus 1 is a typical icosahedral herpesvirus that measuring 90–95 nm. The virion is enveloped and contains a DNA genome (Wolf et al. 1978; Hedrick et al. 1987). The herpesvirus isolate made from steelhead trout in California proved to be similar, but distinguishable from the initial isolate; therefore, two strains of SalHV-1 are recognized and no further distinction will be made between them. Hedrick et al. (1987) serologically compared North American isolates of SalHV-

1 to Japanese salmonid herpesvirus isolates (Oncorhynchus masou virus, OMV; Yamame tumor virus, YTV; and nerca virus Tawata Lake, NeVTA) and found that the United States isolates were distinctly different from those of Japan. On the basis of these differences, isolates from North America were named salmonid herpesvirus 1 and those from Japan, salmonid herpesvirus 2 (Hedrick and Sano 1989). Eaton et al. (1991a) also compared the five herpesviruses from salmonids by DNA homology and confirmed that SalHV-1 was distinctly different from SalHV-2. Davison (1998) using restriction endonuclease mapping, cosmid cloning, DNA hybridization, and targeted DNA sequencing experiments showed that the genome of SalHV-1 is 174.4 kbp in size, consisting of a 133.4 kbp long unique region linked to a 25.6 kbp short unique region that is flanked by 7.7 kbp inverted repeats. This genome structure is different from CyHV-3, ranid herpesvirus 1 and ranid herpesvirus 2 from frogs and CCV; all of these are composed of a single unique region flanked by terminal direct repeats. The genomic sequence of SalHV-1 is distinct but related to SalHV-2 and both are distantly related to CCV. The virus has been placed within the family Alloherpesviridae (ICTVdB 2008).

Epizootiology The first isolation of SalHV-1 was made from ovarian fluid of postspawning adult rainbow trout that experienced up to 50% mortality at a federal hatchery in Washington (Wolf and Taylor 1975). All brood stock infected with the initial SalHV-1 were destroyed and the hatchery thoroughly disinfected. The virus was not isolated again until more than 10 years later in California (Hedrick et al. 1987). During the California outbreaks in 1985–86, virus (SalHV-1) was isolated from 5.8% of ovarian fluid samples and 4.5% of pooled tissue samples collected from steelhead trout at one hatchery (Eaton et al. 1989). It has been

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theorized that vertical transmission of SalHV1 occurs because of its presence in ovarian fluids. Salmonid herpesvirus 1 epizootics occur from January through April when water temperatures are 10◦ C or less. Incubation period in experimental infections is 25 days or more. Wolf and Smith (1981) reported that mortality continued for about 50 days after onset of disease but Eaton et al. (1989) were unable to kill fish with their SalHV-1 isolates.

Pathology Wolf and Smith (1981) described histopathologic changes in fish that had been experimentally infected with SalHV-1. Heart tissue was edematous and necrotic with loss of striations in muscle fibers; kidneys, livers, and spleens were edematous, hyperplastic, congested, and necrotic with syncytia in the spleens. Eaton et al. (1989) described hypertrophy, edema, and hemorrhaged gill epithelium that separated from connective tissue. Also fused liver cells had enlarged nuclei containing Feulgen positive, eosinophilic Cowdry Type A inclusion bodies; skeletal muscle of some infected fish was hemorrhaged and necrotic, and syncytia was present in liver hepatocytes.

Significance Because SalHV-1 has low pathogenicity and is infrequently encountered, its importance to trout culture is believed to be minimal.

Salmonid herpesvirus 2 Salmonid herpesvirus 2 (SalHV-2) was isolated from adult masu salmon (also known as Seema, Japanese salmon, or cherry salmon) in Japan (Kimura et al. 1981a). The virus is also known as Oncorhynchus masou virus (OMV). Two other fish viruses isolated in Japan are similar to OMV, namely nerka virus Towada Lake, Akita Prefecture (NeVTA) and yamame

tumor virus (YTV) (Sano et al. 1983; Hedrick and Sano 1989). Salmonid herpesvirus 2 is an important pathogen in Salmonid culture in Japan, and it is an OIE notifiable pathogen (OIE 2008).

Geographical range and species susceptibility Salmonid herpesvirus 2 (OMV) is known only in Japan where it causes mortality in juvenile masu and coho salmon and tumors on adults (Kimura et al. 1981b). Experimentally, sockeye (kokanee) and chum salmon, and rainbow trout are also susceptible (Kimura et al. 1981c, 1983).

Clinical signs Clinically, SalHV-2 manifests itself in two different ways depending upon age of fish. In juveniles, OMV produces an acute disease that kills 30- to 150-day-old fish while neoplasia develops in some older fish. Infected juvenile fish become inappetent, exophthalmic, and petechiae appear on body surfaces especially under the jaw and belly (Figure 8.8). Internally, livers of infected salmon fry are mottled with white areas and in advanced cases, the liver is totally white. The spleen may be swollen and macroscopically the kidney appears normal (Kimura et al. 1981b, 1981c). Beginning about 4 months postvirus exposure and persisting for at least 1 year, neoplastic tumors develop on surviving kokanee, chum, coho, masu salmon, and rainbow trout. These tumors occur mainly around the mouth in the maxillary and mandibular (Figure 8.8) regions but may also be found on gill covers, body surface, and cornea, but rarely in the kidney (Kimura 1981b).

Diagnosis During active infections, SalHV-2 is isolated in cell cultures from clinically ill salmon less than 6-months-old, normal adult salmon or

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(a)

(b)

(c)

Figure 8.8 Salmonid herpesvirus—type 2 (Oncorhynchus masou virus) in chum salmon. (a) Experimentally infected young fish with petechial hemorrhage on underside (arrow) and exophthalmia. (b) OMV tumor on mandible of young fish(Yoshimizu et al. 1987; reprinted with permission of Springer-Verlag Publishing Co.). (c) Tumor on upper jaw of adult fish. (Photo by M. Yoshimizu; reprinted with permission of CRC Press.)

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fish with tumors. The virus will replicate in all cell lines of salmonid origin, including CHSE-214 and RTG-2, where yield is about 106 TCID50 /mL (Kimura et al. 1981a). Optimum cell culture incubation temperature is 10–15◦ C, but viral replication will occur at 5–16◦ C. Cytopathic effect develops in 5–7 days at 10–15◦ C and is characterized by cells becoming rounded, syncytium formation, and eventually lysis. Cowdry type A inclusion bodies are present in syncytia cells. Inoculated cultures that do not develop CPE in 10–14 days should be blind passed. Isolates of OMV are distinguished from other fish herpesviruses by serum neutralization tests (Hedrick et al. 1987). Gou et al. (1991) prepared a recombinant plasmid DNA probe and were able to detect SalHV-2 DNA in salmonid tissues 2 weeks before virus was isolated from asymptomatic fish. A monoclonal antibody preparation against a recent isolate was cross-reactive with OMV and SalHV-1 but not with IPNV or IHNV (Hayashi et al. 1993). Tumors may be confirmed by histology. The SalHV-2 detected in fish tissue and infected cell cultures by indirect FAT using antisera to the nucleocapsid (Kumagai et al. 1995a). This simple, rapid, and reliable procedure detected the virus in liver smears that were over 6 months old.

Virus characteristics Salmonid herpesvirus 2 is a typical herpesvirus. Without the envelope, the icosahedral nucleocapsid measures 100–115 nm in diameter and the enveloped particle measures 220–240 nm (Tanaka et al. 1987). The SalHV-2 agent is serologically and morphologically similar, and possibly identical to YTV and NeVTA agents (Sano et al. 1983). Using DNA, Eaton et al. (1991b) further demonstrated that OMV and YTV are similar but distinctly different from NeVTA; however, they still placed the three strains in SalHV-2. Sequencing of two cloned fragments of the SalHV-2 genome revealed that this herpesvirus

was related to CCV and subsequent analysis of the SalHV-1 genome demonstrated higher homology to each other and supports its inclusion in the Alloherpesviridae family (Bernard and Mercier 1993; Davison 1998).

Epizootiology Salmonid herpesvirus type 2 (OMV, YTV, and NeVTA) is widespread in Japan (Sano 1988); however, due to management practices, the disease has disappeared from anadromous runs in some regions (Tanaka et al. 1992). Salmonid herpesvirus 2 is probably transmitted vertically during spawning via contaminated ovarian fluid and horizontally via water. Studies of SalHV-2 by Kimura et al. (1981b) indicated that 35–60% of 3- to 5-month-old chum salmon died during a 60-day period following infection by immersion in water containing 100 TCID50 /mL. Kimura et al. (1981c) compared the susceptibility of 0- to 7-monthold chum, masu, kokanee, coho salmon, and rainbow trout at 10–15◦ C. Cumulative mortalities in chum salmon ranged from 3% for 0 age to as high as 98% for 3-month-old fish, but only 7% and 2% mortality occurred in 6- and 7-month-old chum salmon respectively. Similar cumulative mortalities were recorded for sockeye and masu salmon; however, l-monthold sockeye suffered 100% mortality. Coho salmon and rainbow trout were less susceptible to SalHV-2 (OMV strain) with the highest mortality reaching 39% and 29% respectively. Sano et al. (1983) reported a 65% mortality of 5-month-old yamame (masu) when injected IP with 103 TCID50 of SalHV-2 (YTV strain), but only 15% mortality of those injected with 102 TCID50 . Tanaka et al. (1984) noted an 87% mortality of masu salmon over a 4-month period when infected at l month of age and 63% mortality when infected at 3 months. The oncogenic nature of SalHV-2 (OMV) provides a study model for herpesvirus induced tumors. Sixty percent of masu that survived SalHV-2 (OMV) epizootics exhibited tumors, 45% of which occurred around the mouth

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Tumor bearing rate (%)

40

30

20

10

0

100

150 Days postinfection

200

Figure 8.9 Increasing incidence of SHV-2 (OMV) tumor induction in chum salmon. (From Yoshimizu et al. (1987).)

(Kimura 1981a, 1981b). Sano et al. (1983) provided the first information to authenticate the oncogenicity of a herpesvirus isolated directly from fish tumor and for which River’s postulates were fulfilled. Working with the SalHV-2 (YTV) agent, they found that tumors developed primarily on mandibles or premaxillae of masu and chum salmon 10–13 months after exposure. A comparison of tumor incidence in four different salmonid species exposed to SalHV-2 between 1 and 7 months of age was conducted by Yoshimizu et al. (1987) (Figure 8.9). Tumors took 120–360 days to develop but finally occurred in 100% of 3month-old challenged masu, 12% of rainbow trout, 35% of coho salmon, and 40–71% of chum salmon.

Pathology Histopathology of OMV disease varies with species and age of infected fish (Tanaka et al. 1984). In the viremia stage the kidneys of masu, chum, and coho salmon are usually the principal target organ as judged by severity changes. Epithelium of the mouth, jaw, operculum, and head in l-month-old masu

was necrotic, with focal necrosis of hepatic cells in the liver, spleen; necrosis of the pancreas developed later. In 3-monthold masu, necrosis of hematopoietic tissue is seen simultaneously with hyperplasia, and syncytia cells were present in the kidney. As infection progresses, the kidney returns to normal but the liver, initially marked only with focal necrosis, becomes more extensively necrotic. Also, hepatocytes display margination of chromatin. Cell degeneration occurs in the spleen, pancreas, cardiac muscle, and brain. Tumors of differentiated epithelium develop 4 months to 1 year postinfection in 12–100% of surviving chum, coho, and masu salmon and rainbow trout. Kumagai et al. (1995b) found that coho salmon survivors injected with SalHV-2 developed basal cell tumors on the gills 18 months after inoculation. Generally tumors develop earlier in chum and coho salmon than in rainbow trout and masu salmon (Yoshimizu et al. 1989). Most tumors form around the mouth and head but tumors are also present in decreasing frequency on/in caudal fin, gill cover, body surface, cornea, and kidney. Tumors include abundantly proliferative, well-differentiated squamous epithelial

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cells in which the papilloma is supported by a stroma of fine connective tissue, and several layers of squamous epithelial cells are present in a papillomatous array (Yoshimizu et al. 1988b).

Significance Salmonid herpesvirus 2 is a major problem among land-locked salmon populations and coastal cage reared coho salmon in northern Japan (Yoshimizu 1989). The virus was isolated from all but four locations sampled in northern Japan between 1976 and 1987. A significant number of 1- to 5-month-old salmon may be killed during a SalHV-2 outbreak and epizootic survivors tend to develop tumors and become virus reservoirs. Since SalHV-2 has been isolated from ovarian fluids of adult fish, transmission during spawning is most likely; however, treatment of eggs with iodophores significantly reduces this possibility.

Epizootic epitheliotropic disease Epizootic epitheliotropic disease (EED), so named because it is primarily a disease of the skin, is an acute viral disease of lake trout (Bradley et al. 1989; McAllister and Herman 1989). Acute mortality occurred in cultured juvenile lake trout populations in several state and federal hatcheries in the Great Lakes basin between 1985 and 1989; it was estimated that 15 million cultured lake trout died from an undetermined disease. Much speculation was made as to the cause of these mortalities that included epitheliocystis (Bradley et al. 1988) and Chlamydia (McAllister and Herman 1987) but no definitive etiology was established until transmission studies showed that the disease was infectious. It was demonstrated that a transmittable, filterable agent was involved and transmission electron micrographs showed putative herpesvirus particles in tissues from naturally and experi-

mentally infected fish (McAllister and Herman 1989). Similarly, Bradley et al. (1989) demonstrated that a filterable agent was involved and virus capsids could be recovered from affected fish by isopycnic centrifugation. It is now accepted that the etiological agent of EED was a virus named “epizootic epitheliotropic disease virus” (EEDV). The condition has also been called lake trout epidermal hyperplasia, lake trout hyperplasia, and salmonid herpesvirus (SalHV-3).

Geographical range and species susceptibility EED is known to occur only in the Great Lakes region of North America, particularly in the Lake Michigan and Lake Superior water sheds. In addition to infecting lake trout, the disease infects lake trout X brook trout hybrids. The virus has not been observed in wild or feral stocks, although they are considered the probable virus reservoir. Brown, brook, and rainbow trout as well as chinook and Atlantic salmon are experimentally refractive to EEDV.

Clinical signs Epizootics of EED were characterized by rapidly developing mortality and nonspecific clinical signs including lethargy, riding high in the water, sporadic hyper activity and corkscrew swimming, (Mcallister and Herman 1989; Bradley et al. 1989). Hemorrhages occur in the eyes and occasionally the mouth. Hyperplastic epithelial lesions are gray to white mucoid blotches on the jaws, inside the mouth, and on the body and fins. Secondary fungal infections (presumably Saprolegnia) develop on the eyes, fins, and body surface. The only visible internal lesion is a swollen spleen. Hematocrits of infected lake trout are slightly elevated, averaging about 45% compared to a normal 40%.

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Diagnosis

Epizootiology

EED is presumptively diagnosed in lake trout by recognition of clinical signs in fish that are experiencing acute mortality. Transmission electron micrographs of epithelial tissue reveal EEDV particles that can be recovered from skin tissue by ultracentrifugation and visualization in electron micrographs. Attempts to isolate the virus in CHSE-214, EPC, FHM, and RTG-2 cell cultures were largely unsuccessful; therefore, no in vitro virology techniques are currently available for proper identification. The disease has been diagnosed by virus extraction from skin, partial purification by isopyogenic centrifugation, and visualization of negative-stained virions in electron micrographs. No EEDV antisera are available for diagnostic and virus identification purposes. However Kurobe et al. (2009) developed a PCR procedure that detects and identifies EEDV DNA in clinically healthy juveniles and in the ovarian fluids of spawning adults. The PCR procedure can be used in diagnosing EED.

Epizootics of EEDV occur in spring or fall when water temperatures are 6–15◦ C, but tend to be more prevalent in spring when water temperatures are 8–10◦ C. The disease has been diagnosed only in juvenile cultured lake trout where mortality may approach 100% in fish up to 14 months of age. Survivors of EED shed the infectious agent because EEDV was experimentally transmitted to naive lake trout and lake trout X brook trout hybrids during cohabitation and via exposure to contaminated water containing filtered (0.22 µm) material from homogenates of diseased fish (Bradley et al. 1989; Mcallister and Herman 1989). The viral etiological agent of EED has been transmitted by filtrates from infected fish. McAllister (1993) experimentally transmitted EEDV from infected fish to na¨ıve lake trout via water exposure from skin scrapings/homogenates and/or contaminated equipment. Mortality in the na¨ıve fish began 7–14 days later. The recently developed PCR detection method by Kurobe et al. (2009) detected EEDV in ovarian fluid of spawning lake trout lends credence to the theory that the virus can be transmitted vertically from adult fish to progeny lake trout during spawning. EED in aquaculture is stress mediated and almost any stressful condition will precipitate its onset. The process of anesthetizing and clipping fins for identification purposes prior to stocking appears to be a major predisposing factor in precipitating onset of the disease and mortality of fingerling and yearling lake trout (Bradley et al. 1989). Since its discovery in the mid-1980s, EEDV has not been a great problem. Some biologist working closely with lake trout aquaculture believe that this reduced impact because of lower stress on the fish due to improved hatchery management practices that led to the reduction in disease outbreaks (Sue Marcquenski, Wisconsin Department of Natural Resources, personal communication). These practices include lower fish density in culture units, no recirculating water, better feed, and generally improved husbandry

Virus characteristics EEDV virus has been placed in the family Herpesviridae. Electron microscopy of harvested material from infected tissues by ultracentrifugation revealed hexagonal, unenveloped viral particles that measures 100–105 nm in diameter and enveloped particles that measures 220–235 nm (Bradley et al. 1989). McAllister and Herman (1989) found two types of ellipsoid to spherical viral particles, one of which measured 130–175 nm in diameter with a diffused electron-dense nucleoid with a diameter of 75–100 nm. The other particle type measured 150–200 nm in diameter with a diffuse nucleoid measuring 75–100 nm. According to Waltzek et al. (2009), EEDV shares >81% similarity for deduced amino acids sequence of regions of the DNA polymerase and terminase genes with SalHV-1 and SalHV2. These data support designation EEDV as SalHV-3.

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Trout and Salmon Viruses 191

practices. Future studies can be carried out to test this theory but these observations agree with evidence from other herpesvirus diseases of fish that are often stress related. Apparently few new outbreaks of EEDV have occurred since infected hatcheries were depopulated and sterilized in the early 1990s. Although, millions of virus-infected lake trout were stocked into the Great Lakes before the disease was known to be a virus, no cases of EEDV have been detected in these wild populations. However typical of herpesvirus, it could be present in wild fish as a latent stage.

Pathology Histopathology is the same in EEDV infected lake trout whether disease occurs naturally or is experimentally induced (Bradley et al. 1989). The epithelium contains hypertrophied cells and areas of hyperplasia with lymphocytic infiltration, cell degeneration, and necrosis. Affected cells in the epidermis contained swollen cytoplasm and enlarged nucleus. Macrophages laden with cellular debris are common in the kidneys and sinusoids of the liver. In advanced cases, gills develop inconsistent edema. Intranuclear inclusion bodies are sometimes seen in epidermal cells.

Significance A filterable, infectious agent has been implicated in at least some EED cases and strong evidence points to a herpesvirus etiology. The virus is capable of killing large numbers of juvenile lake trout under certain conditions and continues to be a threat to lake trout aquaculture.

etiology was unknown until Nelson et al. (1995) isolated a togavirus like agent from diseased Atlantic salmon in CHSE-214 cells where cytopathic effect developed in 28 days at 15◦ C. When filtrates from infected cells were injected into naive Atlantic salmon, clinical signs and pathology were consistent with those described for pancreas disease of Atlantic salmon. A similar disease known as sleeping disease is recognized in freshwater rainbow trout culture in Europe. Similar lesions and viral particles were associated with both disease and Boucher and Lauencin (1996) demonstrated cross protection between the two agents in experimentally infected rainbow trout. Sleeping disease virus (SDV) and salmon pancreas disease virus (SPDV) are considered strains of the salmon alphavirus (SAV). The diseases strains of SAV are discussed in a review by McLoughlin and Graham (2007).

Geographical range and species susceptibility Sleeping disease occurs in freshwater rainbow trout aquaculture. It is problematic in France and it was been diagnosed in England, Scotland, Germany, Italy, and Spain (McLoughlin and Graham 2007). Pancreas disease has been recognized in Atlantic salmon in salt water in Ireland, Scotland, France, Norway, Spain, and the United States (Munro et al. 1984; Graham et al. 2007). In addition to Atlantic salmon, the disease has been frequently found in rainbow trout in the marine environment in Norway (Taksdal et al. 2007). Recent data suggests that the Pancreas disease in Atlantic salmon and rainbow trout in Norway is caused by a different strain of virus than that of the other regions.

Salmon pancreas disease/sleeping disease in rainbow trout Clinical signs A pathological condition in the pancreas of cultured Atlantic salmon was recognized in Ireland, Scotland, France, Norway, Spain, and the United States (Munro et al. 1984). The

Fish with pancreas disease display a loss of appetite, lethargy, and abnormal swimming behavior. They may produce fecal casts, be

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in generally poor condition, respond poorly to physical stress, and often have dermal lesions due to inability to maintain their position in the water column and the associate abrasions from rubbing against the pen. Affected fish may not be able to swallow food pellets due to muscle degeneration of the esophagus. Gross lesions can include edema of scale pockets exophthamia and ascites. Hearts may be pale and occasionally a ruptured heart is seen upon necropsy (Taksdal et al. 2007). Rainbow trout with sleeping disease lay lethargic at the bottom of the raceway and display extensive necrosis of the skeletal red muscle and degenerative and inflammatory lesions of the exocrine pancreas.

Diagnosis All three strains of SAV have been cultured on CHSE 214 and RTG-2 cells at 10-15◦ C. The CPE consists of small plaques of vacuolated and pyknotic cells. However, the initial isolation culture may not produce obvious CPE and passages may be needed. The use of IFAT or immunostaining on inoculated cell cultures greatly enhances the ability to detect the virus (Todd et al. 2001). When this method was used on rainbow trout artificially infected with sleeping disease virus, it was isolated from the brain, kidney, and serum; highest virus titers were seen in the serum (Kerbart Boscher et al. 2006). They also found that many of the clinical signs of the disease were first evident after the virus was no longer detectable by cell culture. Detection of SAV specific antibodies provides a wider window of detection to evaluate exposure to the pathogen. A virus neutralization test that utilizes immunostaining was much more sensitive for detecting previous exposure than was cell culture (Graham et al. 2005). The use of RT-PCR has gained popularity for diagnosing the presence of SAV. Villoing et al. (2000b) used RT-PCR targeting a fragment of the E2 glycoprotein gene of

SAV 2 and demonstrated its utility in detecting the virus 70 days postinfection, long after the virus could no longer be detected by cell culture. They showed that the kidney and pyloric ceaca were more reliable for detecting the virus than were heart or kidney tissues. Furthermore they were able to detect SAV 1 using the same primers. SYBR Green-based (Graham et al. 2006) and Taqman-based realtime RTPCR assays have been developed for SAV (Hodneland and Endresen 2006). Three different assays were developed for detection of SAV 1, SAV 3, or all three strains.

Virus characteristics Nelson et al. (1995) first isolated and characterized SPDV. They found that the virus had the characteristics of a togavirus. It is a spherical, enveloped RNA virus with a diameter of 65.5 nm that can be inactivated by chloroform, pH 3.0 and 50◦ C and has a buoyant density in cesium chloride of 1.20 g/mL. Graham et al. (2007), evaluating virus stability in various environments, found that the virus was effectively inactivated by in organically rich medium at 60◦ C and at 4◦ C if the pH was adjusted to pH 4.0 or below or above pH 12. The virus survives a few days in fish serum at 4◦ C. For long-term storage, virus in fish serum or cell culture medium survived much better when held at −80◦ C than −20◦ C. Villoing et al. (2000a) sequenced the ∼12 kb RNA genome of the SDV and demonstrated that it was in the Alphavirus genus of the family Togaviridae. Subsequently Weston et al. (2002) sequenced the genome of SPDV and compared the pathogenicity and antigenicity between SPDV and SDV. They concluded that there were differences between the two, but they are closely related isolates of the same virus species designated Salmonid alphavirus (SAV). Hodneland et al. (2005) recently showed that pancreas disease virus from Atlantic salmon and rainbow trout in Norway is a distinct virus subtype and named it

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Norwegian salmonid alphavirus (NSAV). The salmonid aphaviruses are designated subtypes 1, 2, and 3 for SPDV, SDV, and NSAV respectively.

Epizootiology Pancreas disease occurs mainly in Atlantic salmon in their first or second year at sea. The disease is also seen in rainbow trout in the marine environment in Norway. Outbreaks are most prevalent July through September. Sleeping disease is highly transmissible and affects all ages of rainbow trout in freshwater aquaculture in Europe. Cumulative mortality may exceed 22% of the population.

Pathology Histopathological lesions in association with pancreas disease include heart and skeletal muscle myopathy and lesions in exocrine pancreatic tissue (McLoughlin et al. 2002).

are primarily confined to very young fish and usually result in low mortality. However, challenge studies show that the pathogen is highly virulent and has a broad host range. Because of its regional confinement, pathogenic potential and potential broad host range, this virus is listed as an OIE reportable pathogen. Because of this virus’s potentially broad host and the fact that it is restricted to a region where salmonids are not native, we discuss this disease in detail in Chapter 10, Miscellaneous Viral Diseases of Fish.

Other salmonid virus diseases Other virus diseases have been detected in trout and salmon, but reported incidences are low and consequently they have received little attention. However, some of these viruses have serious disease potential. In his review of fish viruses and viral diseases of salmonids, Sano (1995) listed a total of 24 conditions of salmonids in which a virus was the known or suspect etiological agent, many of which have been discussed.

Significance Aquareovirus diseases Pancreas disease causes substantial losses in pen reared Atlantic salmon in Western Norway and Ireland. This highly contagious virus disease produces a chronic progression in which cumulative losses can be high. Also, diseased fish have poor growth and the quality of the flesh is often reduced due to changes caused by the disease induced muscular degeneration.

Epizootic hematopoietic necrosis virus Epizootic hematopoietic necrosis (EHN) is a disease of rainbow trout and redfin (European) perch confined to Australia (Whittington et al. 1999). Outbreaks of EHN in rainbow trout

New aquareoviruses are being isolated more frequently and from a variety of aquatic animals. LaPatra and Hedrick (1995) reported that at least 25 new aquareovirus isolates were made in the state of Washington alone between 1990 and 1995. Many of these aquareoviruses are not pathogenic but were isolated from apparently healthy fish during routine virus assays. However, chum salmon reovirus (CSV) is pathogenic, results in mortality, and causes moderate to severe multifocal liver necrosis (LaPatra and Hedrick 1995). Landlocked salmon virus (LSV) is a reovirus like agent that was isolated in Japan and apparently causes little injury (Hsu et al. 1989). Several cell lines, including AS, BF-2, BB, CCO, and CHSE-214 cells, are sensitive to this virus.

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Cutthroat trout virus A small hexagonal RNA virus was isolated from ovarian fluid of adult cutthroat trout (Hedrick et al. 1991). Although there was no virus-induced mortality, the virus was experimentally infective to brown and brook trout. Following waterborne exposure, virus was reisolated from the kidney and spleen in CHSE-214 cells at 15◦ C for up to 5 weeks postinfection.

Salmonid retroviruses Retroviruses can cause a variety of chronic diseases. They are often host specific and may be difficult to culture in vitro. Most diseases of fish that are associated with retroviruses are neoplastic diseases. Two diseases have been associated with retroviruses in salmonids: plasmacytoid leukemia virus and salmon swim bladder sarcoma virus

Plasmacytoid leukemia virus Plasmacytoid leukemia (PL) is an important disease of farmed chinook salmon in British Columbia, Canada. The disease, also known as marine anemia, causes pale gills, exopthalmia, ascites, and enlarged spleens and kidneys. Cumulative mortality may approach 50% (Kent et al. 1990). The disease is diagnosed primarily by histology and tissue imprints. Fish with PL have large numbers of plasmablasts in the posterior kidney and at least one other tissue. Plasmablasts are large immature lymphocytes that have dark cytoplasmic staining with Giemsa. These cells are immunoglobulin positive and ultrastructuraly similar to plasma cells (Kent et al. 1990). Kent and Dawe (1990) demonstrated an infectious etiology of PL by experimentally transmitting the disease to chinook salmon, Atlantic salmon, and sockeye salmon using kidney homogenates from a chinook salmon with PL.

A retrovirus-designated salmon leukemia virus (SLV) was isolated from eye and kidney tumors of affected fish by Eaton and Kent (1992). Later cell lines derived from kidney and eye tissue from a salmon with plasmacytoid leukemia were persistently infected with the retrovirus (Eaton et al. 1993). This virus induced syncytia and vacuole formation in the infected cell lines, and produced 110 nm in diameter, enveloped virions by budding from the cytoplasm. These virions had a density of 1.16 g/ml in sucrose and were associated with a manganese-dependent reverse transcriptase. Purified virions, evaluated by polyacrylamide electrophoresis, produced seven virus-specific protein bands (Eaton and Kent 1992; Eaton et al. 1993). A microsporidian Nucleospora salmonis has also been implicated as an etiological agent of PL (Hedrick et al. 1990; Morrison et al. 1990). Anemia and proliferation of plasmacytes may be clinical signs of two separate diseases in chinook salmon. Hedrick et al. (1991) demonstrated transmission of the disease using cells from infected fish and prevention of disease using the anti-microsporidian drug fumagillin. Also, Wongtavatchai et al. (1995) were able to transmit the disease using the microsporidian produced in cell culture, whereas Kent and Dawe (1993) were able to transmit the disease using homogenates after filtration through 0.22 µm filters and were not able to suppress disease transmission using fumagillin. Also, Eaton et al. (1994) demonstrated a bimodal viremia in fish infected with tissue. The first being 3–4 weeks and the second at 7 weeks postinjection. Diagnosis of PL is problematic especially during early stages of the disease. It is done primarily by clinical signs and histological examination. Yet, there is considerable variation in interpretation of histological samples by diagnosticians (Stephen et al. 1995). Monoclonal antibodies have been established for SLV (Newbound et al. 1993). Using IFAT with these antibodies on tissue imprints was more

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specific than histology in identifying PL in experimentally infected fish (Newbound et al. 1995). However, when used on field samples, this test proved unreliable (Saksida et al. 1999). Factors that make diagnosis more difficult are the confounding effects of concurrent bacterial kidney disease and Nucleospora salmonis infection. Stephen et al. (1996) evaluated the epidemiology of PL in farmed pacific salmon in British Columbia. They found the PL is endemic to the region and the mean losses from PL outbreaks are approximately 6% and that PL was diagnosed most often in August and September, when water temperatures were at their highest. They also found an association of PL with concurrent presence of other infectious and inflammatory diseases.

affected tumor using RT-PCR with degenerate primers (Quackenbush et al. 2001). This virus was designated salmon swim bladder sarcoma virus (SSSV). Since then the entire SSSV genome has been sequenced from provirus isolated from affect tissue (Paul et al. 2006). They found that this virus was most closely related to an endogenous retrovirus in zebrafish and was phylogenetically between the Epsilonretrovirus (containing walleye dermal sarcoma virus and walleye epidermal hyperplasia viruses 1 and 2) and Gammaretrovirus genera. The factors leading to disease and the importance of SSSV to commercial and wild stocks of Atlantic salmon are not known.

Salmon swim bladder sarcoma virus

Intracellular pathogens most often associated with epitheliocystis of gills have been Chlamydia and Rickettsia. However Fridell et al. (2004) isolated a paramyxovirus from the gills of Atlantic salmon afflicted with epitheliocystis associated with mortality in two salmon farms in Norway. Atlantic salmon paramyxovirus (ASPV) was suggested as the name of the disease. Virus was isolated in rainbow trout gills (RTgill-Wl) but little is known about the diseases importance. Proliferative gill inflammation (PGI) has been recognized for over 40 years with numerous multifactorial etiologies in several fish species including salmonids. Kvellestad et al. (2005) produced evidence that the condition may have had a viral etiology in Atlantic salmon in Norway. They detected ASPV in Atlantic salmon gills by immunohostochemistry on sections of formalin fixed, paraffin imbedded tissue. Pathological changes included pale gills, inflammation, circulatory disturbances, cell death, and epithelial cell proliferation. The ASPV was seen in lamellar epithelial and endothelial cells of physiologically altered tissues. The authors concluded that ASPV is at least a contributing cause of PGI.

There are two reports of outbreaks of a neoplasic disease involving the swim bladder in Atlantic salmon. The first case was in 1975, involving a commercial marine net pen facility in Scotland where a detected prevalence of swim bladder tumors was 4.6% in second-year juveniles held at sea (McKnight 1978). Affected fish had large tumors classified as leiomyosarcomas or fibrosarcomas. These tumors often protruded from the swim bladder and occupied a large portion of the abdominal cavity. Tumors apparently arose from the junction between the inner smooth muscle and the areolar tissue zone of the swim bladder. Electron microscopy revealed retrovirus-like particles in the affected tissue (Duncan 1978). The second case occurred in 1997–1998 in a population of juvenile Atlantic salmon collected from Pleasant River in Maine and maintained at the North Attleboro National Fish Hatchery in Massachusetts. The population experienced chronic mortality and a cumulative loss of 35%. Affected fish had sarcoma of the swim bladder and a portion of a retrovirus pol gene was amplified from the

Atlantic salmon paramyxovirus (ASPV)

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Viral erythrocytic necrosis Viral erythrocytic necrosis is a relatively common disease of pacific salmon in the marine environment. It is caused by the erythocytic necrosis virus (ENV) and is characterized by moderate to severe anemia, poor growth and secondary infections. The disease is usually diagnosed by the presence of eosinophilic cytoplasmic inclusion bodies in giemsa-stained erythrocytes. This disease affects a variety of species of marine fish and is discussed in more detail in Chapter 10, Miscellaneous Viral Diseases of Fish.

Management of salmonid viruses Fish virus diseases are not amenable to treatment with drugs or chemotherapeutics but some may be controlled entirely through management: avoidance, prevention, and vaccination.

Avoidance Avoidance is the most effective and economical way of controlling some salmonid viruses. However, many salmonid viruses may be carried in asymptomatic fish throughout their life. These carrier fish provide a source of infection either through reproduction, physical contact, or in the water source. Specific pathogen free (SPF) certification is used to insure that eggs of a specified brood stock population are free of specific pathogens. This certification is based on a sample assay of broodfish from an egg production facility prior to using those fish as an egg source (OIE 2006; USFWS and AFSFHS 2007). These procedures have been used widely in trout and salmon aquaculture since the mid-1960s and have undoubtedly thwarted the spread of IPNV as well as other fish viral agents. Ideally, brood stock should be certified free of a target virus for three consecutive years with two assays each year. When

eggs are brought into a production facility, they should be hatched and reared through the susceptible fingerling stage in a closed, fishfree water supply. Because non-salmonid fishes can be virus carriers, fish that come in contact with cultured salmonids should also be virus free. The SPF technique is used to help prevent the transmission of IPNV, IHNV, VHSV, and other viruses is effective in providing brood stock from which eggs are taken are held in a closed water system with no contact with waters containing possible carrier fish. While this approach is a credible procedure, it is not a perfect system, because there are examples where the spread of virus has not been totally prevented through SPF egg certification. An example of successfully preventing transmission of salmonid herpesvirus type 2 by screening brood stock for virus and disinfecting eggs by immersion in iodophores at 50 mg/L for 20 minutes was reported by Kimura and Yoshimizu (1989). Egg disinfection can also reduce the impact of other salmonid viruses by eliminating pathogens that might be attached to egg surfaces. Eggs should be incubated in water from a fish-free source, or if eggs are incubated in open water supplies, water should be irradiated with ultraviolet light with 3.0 × 103 µw.sec/cm2 (Yoshimizu et al. 1986). Fingerlings should also be grown in UV treated water. The incidence of SalHV-2 has declined in Japan since initiation of egg disinfection with iodophores and UV irradiation of incubation and culture waters. Avoidance of ISA disease can be enhanced by locating Atlantic salmon culture sites no closer than 5 km to another site and by decontamination of virus positive sites (Jarp and Karlsen 1997). Implementation of good sanitation practices by fish farmers can also reduce risk of ISA. Currently, no management or control procedures, other than avoidance, are known to prevent dissemination of the EIBS virus or salmonid herpesvirus 1; however, Wolf (1988) recommended that sound hatchery sanitation practices be followed as a control for SalHV-1.

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Management and control of EEDV relies upon avoidance. Hatcheries in which EEDV had been reported were disinfected with chlorine and stringent regulations imposed on reintroduction of fish. Experimentally, treating water with high doses of UV irradiation prevents waterborne transmission of EEDV.

Prevention of virus diseases If IHNV-positive brood stock cannot be avoided (i.e., sea-run Pacific salmon) the management approach outlined for Alaskan salmon operations by Meyers et al. (1990) should be applied. According to K. Amos (personal communication), these same procedures have been used in the state of Washington to reduce potential for IHNV transmission during manual spawning of steelhead and all Pacific salmon. These brood stock/egg management procedures can reduce hatched fry mortality attributable to IHNV from the 90% range to less than 10% if: 1. IHNV-free water is used for egg incubation and rearing. If fish-free water is not available it is treated with ultraviolet light. 2. A strict disinfection policy is used for utensils, facilities, field clothing, personnel, and external surfaces of brood fish during egg and milt collection. 3. Eggs from each female are fertilized separately with milt from 1 or 2 males. Each spawn of eggs from a specific mating is water-hardened separately in 100 mg/L solution of iodophor for 60 minutes. 4. Eggs from 80 to 100 females are pooled into stacked incubators with upwelling water. 5. Eggs in each hatching unit are kept physically isolated from eggs in other hatching units. 6. Alevins are transported and released when yolk is depleted or after rearing fry for 3–6 weeks. For management of IHNV after clinical disease has already occurred, Amend (1970) rec-

ommended increasing water temperature to 18◦ C to help arrest IHNV epizootics. However, Hetrick et al. (1979) pointed out that increasing water temperature does not stop mortality of already infected fish and epizootic survivors may still become virus carriers. Reducing water temperature to 4 or 5◦ C during peak susceptibility periods may have a beneficial effect on reducing effects of IPNV virus (Frantsi and Savan 1971). To control VHSV, it is essential to avoid wide fluctuations in water temperature and to carefully manage other environmental stressors. Programs to manage VHSV in Europe advocate controlled movement of fish, destruction of infected populations when feasible, and disinfection of facilities before restocking with VHSV SPF fish or eggs (Daelman 1996). On the basis of epidemiology, McClure et al. (2005) made several management suggestions to reduce losses to ISA in salmon net pens. They advocated frequent sea lice treatments, use care in seawater adapting smolts to minimize stress, avoid stocking large smolts, and establish boat operations that minimize travel between production sites. Additionally, Hammel and Dohoo (2005) advocated the production of single year classes for ISA prevention on a facility and extended feeding of moist pellets. In areas where IPNV, IHNV, or VHSV are indigenous, the culture of more resistant fish strains or species should be considered. Using male and female rainbow trout from a resistant stock, Kaastrup et al. (1991) showed that inherited resistance to VHSV was carried by the male rather than the female. In view of this, a breeding program to enhance resistance would appear to be feasible and has been used on Danish rainbow trout farms. Also, Dorson et al. (1991) showed that triploid rainbow and brook trout hybrids were resistant to VHSV and IHNV, but remained susceptible to IPNV. Several antiviral agents have been used experimentally to retard virus development in fish and cell cultures (Fryer et al. 1976).

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Addition of 0.14 mg/L of free iodine to hatchery waters helped prevent rainbow trout from becoming infected by waterborne exposure to IHNV (Batts et al. 1991). Some antiviral agents have been noted to reduce SalHV-2 mortality. Kimura et al. (1983) showed that Acyclovir quinine reduced tumor development when fish were immersed in 25 µg/mL of the drug for 30 minutes each day for 2 months. Iododeoxyuridine gave even better results with an oral dosage of 15 µg/fish per day for 2 months; however, neither drug is approved for use in food fish and this approach is probably too expensive and labor intensive to be used on a large scale.

Vaccination Considerable research has been expended to develop a vaccine for IPNV, IHNV, or VHSV utilizing several different kinds of antigen preparations. Attenuated and killed virus and subunit preparations have been developed and tested experimentally. When fish were exposed to an attenuated IHNV and then challenged, a significant degree of protection was detected (Fryer et al. 1976). A formalin-killed preparation of IHNV was protective when administered by intraperitoneal injection or by hyperosmotic infiltration (Nishimura et al. 1985). Engelking and Leong (1989) protected kokanee salmon from five strains of IHNV by injection with a glycoprotein from a single strain of the virus. Gillmore et al. (1988) reported successful preliminary results from a genetically engineered IHNV subunit vaccine that was composed of viral glycoprotein produced through recombinant DNA and introduced in Escherichia coli. Oberg et al. (1991) expressed the ribonucleoprotein gene of IHNV in E. coli through plasmid induction but this protein alone does not induce immunity or antibody. However, if it is administered with the bacterially expressed IHNV glycoprotein enhanced, resistance to IHNV occurs.

In an attempt to show protective humoral antibody in rainbow trout to multiple IHNV vaccine exposures, Ristow et al. (1993) found that when results of ELISA and plaque neutralization tests (PNT) were correlated, antibody levels in 5X vaccinated fish were significantly higher at 20 days postvaccination but at 80 days there was no difference in antibody levels. These data indicate only a short term benefit to multiple vaccinations. Exposure of rainbow trout to an avirulent cutthroat trout virus (CTV) provided significant protection when they were exposed to virulent IHNV (Hedrick et al. 1995). Although the protective mechanism was not identified, it was thought to be associated with interferon-like activity in anterior kidney cells. Passive immunization also occurred in rainbow trout following injection of sera from survivors of an IHNV epizootic with neutralization titers of 640–1,280 being recorded (LaPatra et al. 1993). Relative protection ranged from 63 to 100% in these fish, clearly demonstrating that antibodies do provide fish immunological protection. Using monoclonal and polyclonal antibodies, LaPatra et al. (1994a) identified ten antigenic IHNV groups, 91% of which fell into three groups. When susceptible fish were exposed to seven of these antigenic groups they experienced mortality of 14–92%, but more importantly following vaccination with immune serum relative protection of infected fish ranged from 91 to 100%. Following passive immunotherapy, rainbow trout were protected against IHNV by injection even though their virus neutralization antibody titers were low (LaPatra et al. 1994b). These data indicate that vaccination against one strain of IHNV may protect against other strains as well. Emmenegger et al. (1997) synthesized three putative antigenic peptide determinants on the IHNV glycoprotein and injected them into rainbow trout followed by antibody analysis using ELISA and virus neutralization. These antibody responses were generally low and

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inconsistent thus leading to the conclusion that a better understanding of how a fish’s immune mechanism is needed in order to develop an effective IHN vaccine. Research to develop vaccines for prevention and control of IPNV and other viruses has led to availability of several products. In an effort to develop a viral subunit vaccine for IPNV, Manning and Leong (1990) expressed the cDNA of IPNV in E. coli. Rainbow trout were vaccinated by immersion in the bacterial lysate for 20 minutes and challenged 20 days later at which time IPNV protection was acquired. A commercially available subunit IPNV vaccine is now being used in Europe (see Table 4.5). Bootland et al. (1990) showed that immersion of 2-, 3-, and 6week-old brook trout in a formalin-inactivated IPNV preparation enhanced relative percent survival, but as fish became older protection decreased. Vaccination did not prevent IPNV infection in any age group of fry. The recombinant subunit vaccine has shown good application in Atlantic salmon smolts. A multivalent vaccine containing the subunit IPN antigen provided protection in laboratory challenge and in a natural IPN outbreak in marine net pens of Atlantic salmon (Ramstad et al 2007). Much progress in fish vaccinations have been made using naked DNA vaccination using plasmids that express the glycoprotein gene of fish rhabdoviruses and results are very encouraging (Lorenzen et al. 1998; Corbeil et al. 1999; Traxler et al. 1999,). Studies suggest that induction of proinflammatory cytokines may contribute to the observed protection (Acosta et al. 2005). Also, encouraging results have been reported using DNA vaccination against ISA with plasmids expressing the HE gene (Mikalsen et al. 2005) and against IPN using a plasmid expressing the polyprotein from segment A (Mikalsen et al. 2004). With the development of methods to generate recombinant rhabdoviruses (Biacchesi et al. 2000), new attenuated constructs have been

evaluated for use as live vaccines in salmon. Thoulouze et al. (2004) showed a strong protective response of rainbow trout to IHN after immersion exposing them to an NV gene knock out construct of IHNV. Romero et al (2008) demonstrated protective responses of rainbow trout to VHS and IHN in rainbow trout that were immersion exposed to a NV knockout IHNV or an IHNV that expressed the VHSV G protein. In spite of encouraging vaccination results with older fish, it may be difficult to successfully vaccinate salmonid fry against IHNV or IPNV for several reasons: (1) very young fish are most susceptible to these viruses, (2) it takes time to develop a protective immunity, and (3) temperatures at which salmonids hatch and grow are not very conducive to rapid development of a strong immune response. The possibility of immunizing trout and salmon against VHSV was proposed in the mid-1970s by de Kinkelin and Le Berre (1977), but to date it has not evolved into an effective practice. Experimental vaccination programs for VHSV in Europe that employ both killed and live preparations were reviewed by de Kinkelin (1988). Killed VHSV vaccines require injection but are effective in eliciting protection as are DNA vaccines (Lorenzen et al. 1998). Live, attenuated vaccines can be applied via immersion and have provided a moderate to high degree of protection in laboratory trials.

References Acosta, F., A. Petrie, et al. 2005. Kinetics of Mx expression in rainbow trout (Oncorhynchus mykiss) and Atlantic salmon (Salmo salar L.) parr in response to VHS-DNA vaccination. Fish and Shellfish Immunology 18:81–89. Ahne, W. 1982. Vergleichende untersuchungen uber die stabilitat von vier fischpathgenen viren (VHSV, PFR, SVCV, IPNV). Zeits Veternarski (B)29; 457–476. Ahne, W. 1983. Presence of infectious pancreatic necrosis virus in the seminal fluid of rainbow

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susceptibility of rainbow trout (Salmo gairdneri) and hybrids (S. gairdneri x Oncorhynchus kisutch) to experimental infection. Journal of the Fisheries Research Board of Canada 33:1205–1208. Ortega, S. C., R. M. de Oca, et al. 2002. Case report: viral infectious pancreatic necrosis in farmed rainbow trout from Mexico. Journal of Aquatic Animal Health 14:305–310. Park, M. A., S. G. Sohn, et al. 1993. Infectious haematopoietic necrosis virus from salmonids cultured in Korea. Journal of Fish Diseases 16:471–478. Paul, T. A., S. L. Quackenbush, et al. 2006. Identification and characterization of an exogenous retrovirus from atlantic salmon swim bladder sarcomas. Journal of Virology 80:2941–2948. Peters, F., and M. Neukirch. 1986. Transmission of some fish pathogenic viruses by the heron, Ardea cinerea. Journal of Fish Diseases 9:539–544. Piacentini, J. S., J. S. Rohovec, and J. L. Fryer. 1989. Epizootiology of erythrocytic inclusion body syndrome. Journal of Aquatic Animal Health 1:173–179. Pietsch, J. P., D. F. Amend, and C. M. Miller. 1977. Survival of infectious hematopoietic necrosis virus held under various environmental conditions. Journal of the Fisheries Research Board of Canada 34:1360–1364. Pilcher, K. S., and J. L. Fryer 1980. The viral diseases of fish: a review through 1978. Part I: diseases of proven etiology. Critical Reviews in Microbiology 7:289–364. Plumb, J. A. 1972. A virus-caused epizootic of rainbow trout (Salmo gairdneri) in Minnesota. Transactions of the American Fisheries Society 101:121–123. Quackenbush, S. L., J. Rovnak, et al. 2001. Genetic relationship of tumor-associated piscine retroviruses. Marine Biotechnology 3:S88–S99. Ramstad, A., A. B. Romstad, et al. 2007. Field validation of experimental challenge models for IPN vaccines. Journal of Fish Diseases 30:723–31. Rasmussen, C. J. 1965. A biological study of the Egtved disease (IHNV). Annals New York Academy of Science 126:427–460. Raynard, R. S., A. G. Murray, and A. Gregory. 2001. Infectious salmon anaemia virus in wild fish from Scotland. Diseases of Aquatic Organisms 46:93–100.

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Chapter 9

Sturgeon viruses

Intensive culture of white sturgeon is steadily increasing in United States (especially in northern California) and Europe. As a result of more intensive culture, infectious diseases that affect the species have emerged. Five viruses have been associated with mortality in cultured, and to some extent, wild white sturgeon populations; white sturgeon adenovirus (WSAV), white sturgeon iridovirus (WSIV), two white sturgeon herpesviruses (WSHV-l and WSHV-2), and fish nodavirus.

White sturgeon adenovirus WSAV was identified in diseased juvenile white sturgeon between 1984 and 1986 (Hedrick et al. 1985), but the disease has not evolved into a serious health problem.

Geographical range and species susceptibility WSAV was initially observed in 0.5 g sturgeon at a farm in northern California. 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.

Clinical signs Affected fish appear lethargic, anorexic, emaciated, have pale livers, and intestines are void of food. No epidermal lesions have been described.

Diagnosis The virus was first identified only from electron microscopy of infected fish tissue that has large numbers of electron dense hexagonal virions in the cell nuclei. Later a white sturgeon cell line derived from the spleen (WSS-2) was shown to be susceptible to the virus (Hedrick et al. 1991a; Benko et al. 2002).

Virus characteristics WSAV virions average 74 nm in diameter. Methyl green pyronine-stained intestinal tissue from infected fish showed nuclear inclusions that contained DNA. Hedrick et al. (1985) tentatively placed the virus in the family Adenoviridae based on DNA genome, virus morphology, and absence of an envelope. Attempts to isolate the virus in white sturgeon spleen (WSS-l) and white sturgeon heart (WSH-l) cell cultures were unsuccessful, but later the virus 219

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was produced on WSS-2 cells. The evaluation of genomic sequence from cell culture produced virus demonstrated that the sturgeon adenovirus is a unique genus of Adenoviridae (Benko et al. 2002; Kovacs et al 2003).

in northern California (Hedrick et al. 1990). Since then it has been diagnosed as an important cause of losses in white sturgeon on aquaculture facilities from California to British Columbia.

Epizootiology

Geographical range and species susceptibility

The initial WSAV epizootic was not explosive, but accumulative mortality approached 50% over 4 months. Transmission to noninfected sturgeon was only partially successful. Intraperitoneal injections of filtrates into naive sturgeon resulted in enlarged cell nuclei in the gut epithelium, but reaction was not as severe as that observed in naturally infected fish. However, experimentally infected sturgeons were larger than those naturally infected, which may have affected results.

WSIV disease has been detected in Aquaculture facilities in Oregon, Idaho, California, and British Columbia and the virus is considered endemic in the Sacramento-San Joaquin, Columbia, Snake, Kootenai, and Fraser river systems (LaPatra et al. 1994; Raverty et al. 2003). The virus was isolated from juvenile sturgeon that ranged in length from 7 to 46 cm and has only been identified in white sturgeons.

Pathology

Clinical signs and findings

Histologically, epithelial cells lining the straight intestine and spiral valve exhibited nuclear hypertrophy. Nuclei of some infected cells (up to one-third of the cells in some fish) were five times larger than those in noninfected cells. Infected cells continued to enlarge until they ruptured and cell content was released into the gut lumen.

Affected fish appear weak, experience weight loss, cease swimming, and drop to the tank bottom where they die. Gills are pale and display hyperplasia and necrosis of the pillar cells lining the lamellar vascular channels, and some petechia are present. Internally, there is no body fat, livers are pale, and the gut is void of food.

Significance

Diagnosis

When WSAV was first isolated and diagnosed, the disease caused significant mortality in that particular population, but whether or not it will adversely affect future sturgeon seed production is unknown.

WSIV has been diagnosed by electron microscopy and by isolation in cell culture. Hedrick et al. (1992) reported isolation of WSIV in white sturgeon spleen (WSS-2) cells where it induced cell enlargement and slow but progressive cell degeneration. Virus replication occurred at 10, 15, and 20◦ C (optimum temperature) but not at 5 or 25◦ C. Cell culture is slow, requiring over 14 days for CPE to develop. Therefore, presumptive diagnosis is primarily done by histology of cranial skin

White sturgeon iridovirus WSIV was first isolated in 1988 from cultured juvenile white sturgeon at several fish farms

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samples followed by PCR, EM, or cell culture. The PCR method is the most sensitive method of detecting the virus (Kwak et al. 2006). Both histopathology and PCR have been used in nonlethal diagnostics using tissues from fin biopsies (Drennan et al. 2007).

Virus characteristics Electron microscopy of infected degenerating gill tissue shows abundant numbers of icosahedral virions in the cell cytoplasm that measure about 262 nm. WSIV has been placed in the iridovirus-like group because of its DNA genome, large size, and apparent virion assembly in the cytoplasm.

Epizootiology WSIV is highly virulent to juveniles. In the original epizootic, one farm lost 95% of 200,000 juvenile sturgeon in 4 months (Hedrick et al. 1990). Initial clinical signs of disease began 10 days postexperimental exposure to the virus followed by 80% mortality in 50 days at 15◦ C (Hedrick et al. 1992). No deaths occurred in experimentally infected channel catfish, striped bass, or chinook salmon; however, lake sturgeon did suffer low mortality. In studies evaluating the influence of various factors on WSIV disease, Georgiadis et al. (2001) and Drennan et al. (2006) showed a strong association of disease with a predisposing stressor or high-density culture conditions. Georgiadis et al. (2001) suggested possible vertical transmission as the source of virus infection where as Drennan et al. (2006) showed that in the system they evaluated, the most likely source was contaminated river water.

Pathology Histologically, gills, skin, esophagus, and olfactory organ have numerous hypertrophic

and occasionally basophilic cells with swollen nuclei in the epithelium and epidermis.

Significance Because of its pathogenic potential, WSIV is an important pathogen on white sturgeon aquaculture facilities in endemic areas. The virus may appear in other areas where white sturgeon has been introduced.

White sturgeon herpesvirus Two herpesviruses have been isolated from white sturgeon: WSHV-l in 1989 (Hedrick et al. 1991c) and WSHV-2 in 1991 (Watson et al. 1995). These have been renamed Acipenserid herpesviruses 1 and 2, respectively.

Geographical range and species susceptibility WSHV-l was initially reported in California in sturgeon less than 10 cm in length (Hedrick et al. 1991c). WSHV-2 was initially found in subadult and adult sturgeons during routine sampling on commercial farms in California and was identified in diseased subadult sturgeons in 1993 (Watson et al. 1995). This virus was also isolated in wild sturgeons in Oregon, which were trapped in the Columbia River below Bonneville Dam (Engelking and Kaufman 1996). Since then several herpesvirus isolates have been made from other locations and host species, including Idaho, northeastern Canada (in shortnose sturgeon), and in Italy (Kelly et al. 2005). The initial sequence analyses of a portion of the gene encoding the DNA polymerase of these isolates suggested a third distinct sturgeon herpesvius (Kelly et al. 2005). A more thorough evaluation and resolution of amplification errors indicated that the Italian isolate was WSHV-1, the isolate from Idaho was WSHV-2, and the isolate from short

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nose sturgeon from northeastern Canada (designated shortnose sturgeon herpesvirus-SSHV) was closely related to WSHV-2 (Kurobe et al. 2008). Both studies indicated that the WSHV2 is more closely related to channel catfish virus than to WSHV-1 and both were distant from Cyprinid herpesvirus 3. In challenge studies WSHV-2 has been shown to cause disease in pallid sturgeon and shovelnose sturgeon (Kurobe et al. 2008).

CHSE-214, EPC, RTG-2, and FHM were resistant to WSHV-2 (Watson et al. 1995). Virus identical to that seen in cells of infected fish was observed in inoculated WSSK-l cells where it grew at 10, 15, and 20◦ C but not at 5 or 25◦ C. Antiserum to WSHV-l can be used to distinguish WSHV-l from WSHV-2 by neutralization.

Clinical signs

Mature particles were hexagonal, enveloped in cytoplasm vacuoles, and possessed all morphological characteristics of a herpesvirus. The WSHV-l virion nucleocapsids, 110 nm in diameter, were found within the nucleus and cytoplasm of infected cells (Hedrick et al. 1991b). Similarly, WSHV-2 nucleocapsids were 107 nm in diameter, and enveloped particles were 177 nm in diameter (Watson et al. 1995). Sequence analyses of portions of the DNA polymerase gene and the terminase gene confirm that both WSHV-1 and WSHV-2 are herpesviruses. They are more closely related to the other fish and amphibian herpesviruses than to herpesviruses of mollusks, reptiles, birds, or mammals (Kurobe et al. 2008). Thus they are likely members of the Alloherpesviridae family.

There are no specific external clinical signs associated with WSHV-1. Fish continue to feed until death. Internally the stomach and intestines are filled with fluid but other tissues appear normal. In experimental challenges and case reports, WSHV-2-infected fish display raised pale skin lesions on the head and pectoral regions. These lesions are often covered by thick mucus. The affected fish display lethargy and erratic swimming.

Diagnosis WSHV-1 and WSHV-2 were isolated from epithelial tissue only in white sturgeon skin (WSSK-l) cells where syncytia were detected 3 days after inoculation (Hedrick et al. 1991c). Cytopathic effect of WSHV-2 consists of grape-like clusters at the foci of plaques with heteromorphic and fused cells that eventually float off the culture substrate. Total CPE occurred 5–7 days after inoculation at 15◦ C. White sturgeon spleen (WSS-2) and white sturgeon gonad (WSGO) cells are also susceptible to WSHV-1 and WSHV-2. Optimum virus production was seen in WSSK-1 cells for WSHV-1 and WSGO cells for WSHV-2. A fourth cell line, white sturgeon liver (WSLV), developed CPE when exposed to either virus but produced very little virus (Watson et al. 1995). The non-sturgeon cells lines, CCO,

Virus characteristics

Epizootiology The original sturgeon population from which WSHV-l was isolated suffered 97% mortality after being moved into a wet laboratory. However, the disease process was complicated by the presence of columnaris. Experimental virus transmission from cell cultures resulted in 35% mortality after sturgeon were exposed by immersion to 105.3 TCID50 /mL of water for 30 minutes at 15◦ C. Virus was recovered for 2 weeks postinfection but was not isolated at 4 weeks. WSHV-2 was originally isolated from internal organs of captive adult brood stock

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during routine screening and later from a diseased juvenile fish on commercial farms (Watson et al. 1995). In the fall of 1994, apparent WSHV-2 was isolated from approximately 2-year-old wild white sturgeon (Engelking and Kaufman 1996). The virus isolated from these fish was completely neutralized by WSHV-2 antisera, therefore, confirming a second isolation of that virus. WSHV has been implicated as an important cause of mortality in juvenile white sturgeon populations on aquaculture facilities (Georgiadis et al. 2000). WSHV-2 appears to be more pathogenic under experimental conditions than does WSHV-l and can be found in both skin and internal organs. It is more lytic in cell culture and replicates in several white sturgeon cell lines. Also, experimental challenges indicate that other species of sturgeons are susceptible to WSHV-2 (Kurobe et al. 2008).

Pathology Histopathology of WSHV-1-infected fish included focal or diffused dermatitis and epidermal lesions were characterized by intercellular edema. Hydropic degeneration and hypertrophy of malphigian cells with loss of intercellular junctions were common. Chromatin margination was associated with flocculent nonmembrane-bound intranuclear inclusion (Hedrick et al. 1991b). Sturgeons challenged with WSHV-2 demonstrated epithelial hyperplasia of the skin, mouth, pharynx, esophagus, and gills. These hyperplastic lesions led to epithelial erosion. Affected cells demonstrated enlarged nuclei with marginated chromatin (Watson et al. 1995).

where as WSHV-2 has been associated with disease in older fish. Also this virus appears to be more pathogenic in challenge experiments. The occurrence of WSHV-2 associated disease is much more frequent on commercial aquaculture facilities than WSHV-1 (Kelly et al. 2005).

Other sturgeon virus diseases Sturgeon are often cultured on facilities that contain other fish species and are sometimes cocultured with these species and are therefore evaluated as potential vectors for pathogens of their cohorts’ viruses. LaPatra et al. (1995) demonstrated that white sturgeon fry were susceptible to IHNV and that white sturgeons cultured with IHNV-infected rainbow trout had neutralizing antibodies to the virus. In another case, Athanassopoulou et al. (2004) demonstrated that white sturgeon on aquaculture facilities in Greece became infected by Noda virus in a farm that contained infected sea bass. The two species that were separate did not share water, so direct contact was not likely but sequence analysis demonstrated a close relationship between the viruses from the two sources. The fish were in freshwater systems and the authors speculate that the sea bass was the source of the virus and horizontal transmission may have occurred by mechanical means on contaminated equipment. The Nodavirus infections in the sturgeon caused vacuolating neuropathy similar to the disease in other fish, but it did not cause the retinopathy seen in the infected sea bass. Nodavirus is discussed in more detail in Chapter 10 (Miscellaneous Viral Diseases of Fish).

Management of sturgeon viruses Significance WSHV diseases are important to sturgeon production systems. WSHV-1 appears to be primarily a problem in fry and young fingerlings

Some sturgeon aquaculture facilities have adopted the 100 ppm iodine treatment of eggs and attempt to avoid using fish from known WSIV and WSHV-2 carrier populations as

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broodstock. Drennan et al. (2006) demonstrated that avoiding the use of river water in endemic areas greatly reduced the incidence of WSIV infection. All of the virus associated diseases in sturgeons have been shown to be correlated with density and environmental or nutritional stress. Also, because white sturgeon could be vectors for IHNV, their presence on a facility should be taken into account when limiting the transmission of this virus.

References Athanassopoulou, F., C. Billinis, and T. Prapas. 2004. Important disease conditions of newly cultured species in intensive freshwater farms in Greece: first incidence of nodavirus infection in Acipenser sp. Diseases of Aquatic Organisms 60: 247–252. Benko, M., P. Elo, et al. 2002. First molecular evidence for the existence of distinct fish and snake adenoviruses. Journal of Virology 76: 10056–10059. Drennan, J. D., S. E. LaPatra, et al. 2006. Transmission of white sturgeon iridovirus in Kootenai River white sturgeon Acipenser transmontanus. Diseases of Aquatic Organisms 70: 37– 45. Drennan, J. D., S. E. Lapatra, et al. 2007. Evaluation of lethal and non-lethal sampling methods for the detection of white sturgeon iridovirus infection in white sturgeon, Acipenser transmontanus (Richardson). Journal of Fish Diseases 30: 367–79. Engelking, H. M., and J. Kaufman. 1996. White sturgeon viruses isolated from Columbia River white sturgeon. American Fisheries Society/Fish Health Section News Letter 24 (3): 4–5. Georgiadis, M. P., R. P. Hedrick, et al. 2000. Risk factors for outbreaks of disease attributable to white sturgeon iridovirus and white sturgeon herpesvirus-2 at a commercial sturgeon farm. American Journal of Veterinary Research 61: 1232–1240. Georgiadis, M. P., R. P. Hedrick, et al. 2001. Factors influencing transmission, onset and severity of outbreaks due to white sturgeon iridovirus in a commercial hatchery. Aquaculture 194: 21–35.

Hedrick, R. P., J. M. Groff, et al. 1990. An iridovirus infection of the integument of the white sturgeon Acipenser transmontanus. Diseases of Aquatic Organisms 8: 39–44. Hedrick, R. P., T. S. McDowell, et al. 1991a. Two cell lines from white sturgeon. Transactions of the American Fisheries Society 120: 528–534. Hedrick, R. P., T. S. McDowell, et al. 1991b. Isolation of an epitheliotropic herpesvirus from white sturgeon Acipenser transmontanus. Diseases of Aquatic Organisms ll: 49–56. Hedrick, R. P., T. S. McDowell, et al. 1991c. Characteristics of two viruses isolated from white sturgeon Acipenser transmontanus. In: Proceedings Second International Symposium on Viruses of Lower Vertebrates., Corvallis, Oregon, Oregon State University, pp. 165–174. Hedrick, R. P., T. S. McDowell, et al. 1992. Isolation and some properties of an iridoviruslike agent from white sturgeon Acipenser transmontanus. Diseases of Aquatic Organisms 12: 75–81. Hedrick, R. P., J. Spears, et al. 1985. Adenoviruslike particles associated with a disease of cultured white sturgeon, Acipenser transmontanus. Canadian Journal of Fisheries and Aquatic Sciences 42: 1321–1325. Kelley, G. O., T. B. Waltzek, et al. 2005. Genetic relationships among herpes-like viruses isolated from sturgeon. Journal of Aquatic Animal Health 17: 297–303. Kovacs, G. M., S. E. LaPatra, et al. 2003. Phylogenetic analysis of the hexon and protease genes of a fish adenovirus isolated from white sturgeon (Acipenser transmontanus) supports the proposal for a new adenovirus genus. Virus Research 98: 27–34. Kurobe, T., G. O. Kelley, et al. 2008. Revised phylogenetic relationships among herpesviruses isolated from sturgeons. Journal of Aquatic Animal Health 20: 96–102. Kwak, K. T., I. A. Gardner, et al. 2006. Rapid detection of white sturgeon iridovirus (WSIV) using a polymerase chain reaction (PCR) assay. Aquaculture 254: 92–101. Lapatra, S. E., J. M. Groff, et al. 1994. Occurrence of white sturgeon iridovirus infections among cultured white sturgeon in the Pacific Northwest. Aquaculture 126: 201–210. LaPatra, S. E., G. R. Jones, et al. 1995. White sturgeon as a potential vector of infectious

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hematopoietic necrosis virus. Journal of Aquatic Animal Health 7: 225–230. Raverty, S., R. Hedrick, et al. 2003. Diagnosis of sturgeon iridovirus infection in farmed white sturgeon in British Columbia. Canadian Veterinary Journal 44: 327–328.

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.

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Chapter 10

Other viral diseases of fish

Some viral fish diseases are less species specific or group specific than those previously discussed, and/or the fish group they infect has not been represented. The viruses in this chapter that have a broad host range discussed first are epizootic hematopoietic necrosis, lymphocystis virus disease, viral erythrocytic necrosis (VEN) virus, Red Sea bream virus, the fish nodaviruses, other iridoviruses, aquabirnaviruses, and virus associated with epizootic ulcerative syndrome (EUS). Other important viruses that have a limited host range, but were not represented in earlier chapters, discussed here include pilchard herpesvirus (PHV), largemouth bass virus (LMBV), Pike fry rhabdovirus (PFR), other rhabdoviruses, a herpesvirus of turbot, and fish tumor-producing viruses. Virus diseases included in this chapter occur in a wide variety of fish, cause a diverse range of disease conditions, and represent several different virus types (Table 10.1).

Epizootic hematopoietic necrosis Epizootic hematopoietic necrosis (EHN) disease is caused by epizootic hematopoietic 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.

necrosis virus (EHNV), an iridovirus found in Australia (Langdon et al. 1986); it is thought to be the first fish virus found in Australia.

Geographical range and species susceptibility EHN virus was initially isolated from feral redfin perch (European perch) in New South Wales, Australia (Langdon et al. 1986) and a short time later from farmed rainbow trout (Langdon et al. 1988). The virus has also been found in Victoria and South Australia (Whittington et al. 1996). Macquarie perch, silver perch, mosquito fish, and mountain galaxias are also susceptible to EHNV by bath exposure, and Atlantic salmon are susceptible by intraperitoneal injection (Langdon 1989).

Clinical signs EHN in redfin perch and rainbow trout is characterized by a slow or rapid spiral to the surface where they become listless accompanied by ataxic swimming. The fish may even school and “headstand” above submerged objects or swim erratically. Redfin perch infected with 227

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Table 10.1 Viruses that cause important diseases in fish other than catfish, salmonids, carp and minnows, and eels and sturgeons. Virus Type

Virus

Host

Disease Type

Iridovirus Ranavirus Ranavirus

Epizootic hematopoietic necrosis virus Largemouth bass virus

Rainbow trout, redfin perch, and others Black Bass

Acute viremia

Ranavirus Lymphocystivirus

Grouper iridovirus Lymphocystis disease virus

Megalocytivirus

Red Sea bream iridovirus Erythocytic necrosis virus Pike fry rhabdovirus

Fish nodavirus, nervous necrosis virus Pilchard herpesvirus

Grouper Many species of marine and freshwater fish–higher teleosts Many higher teleosts in Asia Many marine and anadromous fish species Northern Pike, other freshwater fish in Europe European perch, largemouth bass, pike perch, grayling Many species of marine fish Pilchard

Esocid herpesvirus 1 (blue spot disease)

Northern pike and muskellunge

Walleye herpesvirus

Walleye

Walleye dermal sarcoma virus Walleye epidermal hyperplasia virus Esox lymphosarcoma virus

Walleye

Unclassified Rhabdovirus Vesiculovirus Vesiculovirus

Nodavirus betanodavirus Herpesvirus Unclassified Unclassified

Unclassified Retrovirus epsilon epsilon Unclassified

Perch rhabdovirus disease

Walleye Northern Pike and muskellunge

EHNV become dark and anorexic. Petechial hemorrhages appear in skin, at the base of fins, and on several internal organs (Langdon et al. 1986; 1988). Infected fish develop a reddening of the head and nostrils and have focal or irregular widespread muscular pallor, particularly in juvenile fish. Based on the studies by Reddacliff and Whittington (1996), rainbow trout infected with EHNV have dark skin pigmentation, are inappetent and sometimes ataxic. Gross lesions include abdominal distension, swelling of spleen and kidney, and occasional pale foci in the liver.

Acute viremia-hyperinflation of air bladder Acute viremia Chronic tumor-like skin lesions caused by enlarged cells Anemia – enlarged cells in viscera Anemia – chronic loss, secondary infection Acute hemorrhagic viremia Acute hemorrhagic viremia

High losses – neurological and vision disorders Acute losses with gill inflammation Flat, granular, bluish white skin lesions caused by enlarged cells diffuse epidermal hyperplasia Invasive tumors of the skin Discrete epidermal hyperplasia Tumors in the dermis may metastasize

Diagnosis EHN virus can be isolated from infected fish in RTG-2 or BF-2 cells with CPE developing in 24 to 36 hours at 25◦ C (Langdon and Humphrey 1987). The virus is also infective in seven other fish cell lines, but CPE severity varies. Cytopathic effect begins as focal rounding and retraction of individual cells, followed by progressive cell lysis or detachment to form cell-free plaques. Identification and detection methods were summarized by Whittington and Hyatt (1997).

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A direct antigen capture ELISA method for EHNV detection was developed by Steiner et al. (1991), utilizing rabbit anti-EHNV primary immunoglobulin and sheep-anti-rabbit secondary antibody. Serological methods for identifying EHNV include polyclonal antiserum for virus neutralization tests, immunoperoxidase staining of histological sections, and immunogold electron microscopy. A PCR procedure was also developed for EHNV detection (Gould et al. 1995). The ranaviruses are very closely related, and differential diagnosis of this genus can be accomplished by PCR that targets a portion of the major capsid protein (MCP) gene followed by sequencing (Mao et al. 1997) or by restriction fragment length polymorphism (Marsh et al. 2002).

Virus characteristics EHN virus has an icosahedral capsid that measures 150–170 nm in diameter; the virus assembles in the cytoplasm and obtains an envelope by budding from the cytoplasm membrane. The virus is classified in the genus Ranavirus of the family Iridoviridae and is similar to several other iridoviruses that have been isolated from fish and frogs (Hedrick et al. 1992; Hengstberger et al. 1993; Ahne et al. 1997). This virus is related to but distinct from the catfish iridovirus and the LMBV (Mao et al. 1997). Like other ranaviruses, EHNV has a large heavily methylated DNA genome that is linear and circularly permuted. Studies to determine the virus’ sensitivity/resistance to desiccation, disinfectants, high and low pH, and temperature indicate that EHNV can survive for extended periods of time in nature and on equipment (Langdon 1989).

Epizootiology EHN virus is the first known fish iridovirus to cause severe necrosis of internal organ tissue. It

also appears to be among the most virulent iridoviruses isolated from fish to date. Outbreaks of EHNV in wild redfin perch populations have resulted in high mortality, ranging from about 40 to several thousand dead juvenile fish per 100 m of shoreline per day (Langdon and Humphrey 1987). An estimated 95% decline in a wild redfin perch population was anecdotally described in one outbreak in New South Wales (Whittington et al. 1996). In another epizootic that lasted 2–3 weeks, mortality may have reached 100% in adult fish. The virus spreads easily on equipment (nets, boats, etc.) and can be transferred via live or frozen fish, or in intestines of piscivorous birds (Whittington et al. 1996). The virus resists drying and physical and chemical agents, and can survive in frozen fish carcasses for at least a year. The virus has been found in healthy adult redfin perch, which are thought to be virus reservoirs in the ecosystem. However, Whittington et al. (1994) found that redfin perch exposed to EHNV showed no sign of infection 2 and 4 months later. Initial experimental EHNV transmission was by injection of infected cell culture media into three adult redfin perch; two died at 4 days and virus was isolated from both; however, the third yielded no virus when killed 21 days later (Langdon 1989). Whittington and Reddacliff (1995) discovered that redfin perch were highly susceptible to EHNV by bath inoculation with virus doses as low as 0.08 TCID50 /mL of water. At 19–21◦ C, incubation was 11 days – longer at cooler temperatures, and no infection occurred at temperatures below 12◦ C. Rainbow trout were susceptible by intraperitoneal injection at 8–21◦ C and were persistently infected after 63 days but were not susceptible via experimental bath exposure. Rainbow trout less than 12.5 cm in length are susceptible with low mortality ranging from 1 to 8%. Recrudescence was thought to result from contact with infected wild redfin perch. Most EHNV outbreaks at rainbow trout hatcheries can be correlated with water

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quality and poor management practices; when these problems are corrected, disease often disappears. Most likely, the redfin perch and rainbow trout epizootics were caused by the same virus because of similar clinical signs, nature of the virus isolates, and in all probability, the trout contracted the disease from redfin perch residing in the watershed (Langdon 1989). Losses in EHN-affected rainbow trout populations are generally low. In these populations, the prevalence of the virus and seroprevalence are also low. However, laboratory findings indicate that it is highly virulent. Therefore, it is believed that the virus causes rapid death to infected individuals with low transmission rate to new individuals. The higher prevalence in affected redfin perch was used to suggest that this is the reservoir in positive aquaculture facilities. However, Whittington et al. (1999) evaluated infected farms and found EHNV in 4- to 6-week old fingerling trout as well as in larger grow-out fish. Also, on another farm, EHNV was detected in adult fish with nocardiosis. When the fingerling source of this farm was evaluated, EHNV-positive fish were found after evaluating routine mortalities. The virus may not be associated with clinically detectable disease; therefore, it is believed that the virus may be transferred with shipments of live fish. Because of the low prevalence in a carrier population and the high prevalence in the low number of dying fish, it was suggested that inspections focus entirely on routine mortalities.

nal hematopoietic necrosis, small multiple focal hepatic necrosis, focal acute splenic necrosis, mild gill hyperplasia with occasional focal necrosis, congestion, and edema in the wall of the swim bladder. Focal acute necrosis of myocardial cells with necrotic cells and debris in the cardiac lumen were also observed.

Significance The significance of EHNV is not clear, but when it becomes established in populations of redfin perch or rainbow trout, high losses can occur. The virus also has the capability to infect a wider range of fish hosts. This is an OIElisted pathogen that is currently restricted to Australia. The potential impact of this virus on a variety of hosts makes it especially important to prevent the spread of EHNV to nonendemic areas.

Lymphocystis Lymphocystis, a cellular hypertrophic disease primarily found on the body and fins, is the oldest and perhaps best known of all fish virus diseases. Although lymphocystis has been recognized since 1874, and theorized to be of a viral nature in 1920 (Weissenberg 1965), its viral etiology was not actually proven until the early 1960s (Wolf 1962). Its etiological agent is lymphocystis disease virus (LCDV), for which Rivers’s postulates were fulfilled in 1964 (Wolf et al. 1966).

Pathology In redfin perch, EHNV produces focal necrosis of the renal hematopoietic tissue, hepatic tissue, spleen, and gastrointestinal tract epithelium, and it is believed that death results from injuries to the kidney and spleen (Langdon 1989). Infected fish also develop encephalopathy in various degrees of severity. On the other hand, clinically diseased rainbow trout develop focal to extensive acute re-

Geographical range and species susceptibility LCDV is the most widely distributed of all fish viruses. It affects freshwater fish in most areas of the world including North and South America, Europe, Africa, Australia, and Asia (Nigrelli and Ruggieri 1965; Wolf 1988). Lymphocystis is also found in fish from nearly all

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marine waters including the Atlantic and Pacific Oceans, Gulf of Mexico, the Red Sea, Bering Sea, and the Mediterranean Sea to name a few (Anders 1989; Faisal 1989). Lymphocystis had not been reported in the Caribbean Sea until it was reported in Caitipa mojarras and the exotic Indian glassfish (Bunkley-Williams et al. 1996; Williams et al. 1996). Anders (1989) named 11 orders, 45 families, and 141 fish species that are susceptible to LCDV, and the list continues to grow. The most prominent orders affected are Perciformes and Pleuronectiformes, which contribute a large percentage of the susceptible species. Sunfishes (black bass and bluegills), perch, sea trout, drum, butterfly fish, angelfish, cichlids, and flatfishes are most frequently infected. Other susceptible fishes include cultured, wild, and traditional aquarium or ornamental species (Durham and Anderson 1978). Since the international ornamental fish trade is vast and essentially unregulated in terms of disease, spread of lymphocystis has likely been enhanced by this industry (Williams et al. (1996). Salmonids, cyprinids, and ictalurids, or members of other low phylogenetic orders do not appear to be susceptible.

Clinical signs LCDV infection produces greatly hypertrophied cells in connective tissue below the epidermis. The cells are large, grey or whitish in color, and may measure up to 2 mm in diameter. These cells, which are easily seen with the unaided eye, may occur singularly or grouped together in “grape-like” clusters and can appear tumor like (Figure 10.1). Lymphocystis lesions develop on any external surface including eyes and gills but are most prevalent on fins, head, and lateral body surfaces. Rarely, lymphocystis lesions are found in the spleen, liver, and mesenteries of juvenile marine fish (Lawler et al. 1974). Colorni and Diamant (1995) described a significant LCDV in-

fection in the spleen of red drum. There are no behavioral abnormalities associated with LCDV infections unless lesions in the oral cavity or on the head interfere with respiration or feeding.

Diagnosis LCDV infections can be identified by gross clinical signs coupled with the presence of hypertrophied cells with enlarged nuclei. These cells may contain Feulgen-positive inclusions in the cytoplasm that can be seen microscopically in stained sections of the lesions (Figure 10.2). Generally, virus isolation in tissue culture is not easily accomplished because of a long incubation period (Wolf et al. 1966). However, LCDV has been isolated in BF-2 cells and largemouth bass cells. Also Bowden et al. (1995) used red drum dorsal fin cells (RDDF-1) incubated at 25◦ C to isolate the virus. Initial CPE became apparent in RDDF1 cells 8–10 days postinoculation, and after 2 weeks, developed enlarged cells ranging from 40 to 90 µm that floated in the media. The cytopathic effect in infected cell cultures is similar to lymphocystis lesions on infected fish (Wolf and Carlson 1965). At 23–25◦ C, in vitro-infected cells increase in size and become basophilic. Six days postinfection, Feulgenpositive inclusions can be seen in the cytoplasm followed by formation of a hyaline capsule at day 10. The cells reach maturity in 3–4 weeks and are identical in appearance to those seen in infected fish, including an enlarged nucleus; however, the cells are smaller. Although cyprinids are generally not susceptible to lymphocystis, in China, Zhang et al. (2003) tested 13 established cell lines for LCDV isolation and successfully isolated the virus from flounder in a cell line developed from grass carp ovary (GCO). Obvious CPE developed in the cell monolayer at 1-day postinoculation. By 3 days, CPE involved about 75% of the cell sheet.

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232 Viral Diseases

(a)

(b)

(c)

Figure 10.1 Lymphocystis virus. (a) Large lymphocystis virus-induced lesions (arrows) on fins of a cultured two-month-old spotted bass; (b) Lymphocystis lesions on a 40-cm smallmouth bass from a wild population; (c) Lymphocystis lesion on an 8-cm croaker taken from a wild population and held in captivity where lesions developed (formalin preserved).

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(a)

(b)

Figure 10.2 Lymphocystis virus. (a) Histological section of a lymphocystis lesion showing the hypertrophy cell with an enlarged nucleus (bar = 100 µm) and (b) electron micrograph of lymphocystis virus (bar = 400 nm).

Virus characteristics Lymphocystis virus is in the genus Lymphocystivirus, family Iridoviridae. The virion is icosahedral with a diameter of 145–330 nm (Robin and Bertholimue 1981) (Figure 10.2). The genome is a linear double-stranded DNA that is terminally redundant, circularly permuted, and heavily methylated. Initial stages of LCDV replication occurs in the nucleus, but virions are assembled in the cytoplasm of infected cells. It is likely that there are numerous species or strains of LCDV. The first indications of species or strain diversity were based on fish species susceptibility; virus taken from one species was not always pathogenic to other species (Overstreet and Howse 1977). Peters and Schmidt (1995) found that lymphocystis

virus in plaice developed anomalies in mature lymphocystis cells and underwent disintegration within the cells inclusion body. The conclusion was drawn that the LCDV in plaice is not totally adapted to that species thus supporting the theory of more than one lymphocystis virus strain. Genomic sequence data also indicate that there are more than one species of LCDV. The entire genomes of two different isolates from flatfish demonstrate a high amount of sequence and genomic structural divergence. Tidona and Darai (1997) sequenced the type strain LCDV-1 isolated from flounder. They found that the genome was 102,653 bp and contained 195 putative genes. Zhang et al. (2004) sequenced an LCDV isolated from cultured Japanese flounder in China (LCDV-C)

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and found that this isolate was 186,250 bp in length and contained 240 potential genes. The most highly conserved gene was the MCP gene. Comparison of the deduced amino acid sequences of the encoded proteins demonstrated only 87.6% identity. Further analyses of MCP genes from Asian isolates indicate substantial sequence divergence between these strains, a rockfish isolate, and LCDV isolates from freshwater ornamental fish, leading to the identification of six genogroups (Kim and Lee 2007; Hossain et al. 2008). Sequencing a portion of the DNA polymerase gene from a largemouth bass isolate obtained in the United States demonstrated 77.7% deduced amino acid identity to LCDV-C and 69% deduced identity to LCDV-1 (Hanson et al. 2006).

Epizootiology Lymphocystis is a chronic, benign condition that seldom results in morbidity or death of fish, with cultured red drum being a possible exception (Colorni and Diamant 1995). The primary problem caused by the LCDV is that infected fish are unattractive and less marketable. However, studies in California indicated that lesions can become so large that the oral cavity is occluded and fish die of starvation (McCosker 1969). Less severely affected fish may experience weight loss, and infected walleye from Lake Michigan weighed less than noninfected fish (Petty and Magnuson 1974). The presence of large lesions on the body or fin may also render fish more vulnerable to gill nets. Transmission of lymphocystis virus is influenced by several factors: fish species, population density, injury, parasite load, environmental conditions, and water pollution levels. The virus can be transmitted by cohabitation or spraying infected material on scarified skin (Wolf et al. 1966). Lymphocystis was also experimentally transmitted in filtrates of tissue from cultured red drum to naive drum by

intramuscular injection, but attempts at oral transmission were unsuccessful (Bowden et al. 1995). In natural waters, lymphocystis virus is presumably transmitted to healthy fish when infected cells rupture, and virus is released into the water with infection being acquired through the skin. Ryder (1961) found that injuries sustained during spawning activities led to a higher incidence of lymphocystis. Handling, netting, tagging, etc., may also increase lymphocystis incidence because lesions tend to develop where scales are disturbed or fins damaged (Clifford and Applegate 1970). Some external parasites (copepods or isopods) that disrupt the protective mucous layer may increase transmission efficiency (Nigrelli 1950). Lawler et al. (1974) postulated that parasitic isopods (Lironeca ovalis) may have been instrumental in a lymphocystis virus infection of silver perch by causing skin irritation, thus allowing virus to invade the tissue. Although lymphocystis is usually selflimiting, Bowden et al. (1995) noted that “lymphocystis in intensive red drum culture systems is frequently a disease of high morbidity” and can result in considerable economic loss because of unsightly lesions. They also pointed out that lymphocystis predisposes red drum to more pathogenic diseases such as amyloodinosis and vibriosis. It has been suggested that an increased occurrence of lymphocystis is influenced by water pollution (Alpers et al. 1977). Anders and Yoshimizu (1994) proposed that pollutants adversely affect the immune and endocrine systems of fish, thus allowing activation of latent skin tumor-producing viruses. In Norway, Reierson and Fugelli (1984) described a higher lymphocystis incidence in flounder in polluted waters than in flounder in less polluted waters, but cautioned against drawing any definitive conclusions because of a complex disease–environmental relationship. Based on limited sampling, Bunkley-Williams et al. (1996) concluded that virus prevalence was higher in areas where lymphocystisinfected fish were cleaned and offal returned

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to the sea, enabling other fish to come into contact with the virus. Lymphocystis occurs in all sizes and ages of fish, but Yamamoto et al. (1985) did report a higher virus incidence in small walleye (250–600g) than in medium (600–1,200 g) or larger fish. They also noted that the rate of infection in females was three times greater than that of males. In a population dynamics study of flounder, Lorenzen et al. (1991) determined that a higher incidence of lymphocystis occurred in younger fish because they were held in high-density conditions and had not yet acquired disease immunity as had older fish. In Israel, lymphocystis lesions were seen on the body and fins of cultured red drum that had been shipped from Texas. Lesions first appeared on only a few fish when they weighed about 20 g, but 2 months later, disease had spread to several hundred fish in a population of 10,000. Lesions identical to those on the fins and body were subsequently seen in the spleen and heart, but none were found in the liver or kidney. The infected cells measured about 430 µm in diameter. Under experimental conditions, lymphocystis incubation time depends upon temperature. At 10–15◦ C, incubation may take up to 6 weeks for lesions to develop compared to 5–12 days at 20–25◦ C (Cook 1972). Incidence of lymphocystis infections may reach 100% in some crowded fish populations (Durham and Anderson 1978). By contrast, in wild marine populations, infection incidence may range from 4 to 57%, with the highest incidence usually occurring in spring and summer (Amin 1979). However, Reierson and Fugelli (1984) reported a disease increase in winter. Lymphocystis lesions usually do not cover large areas of the body or fins, but Overstreet (1988) reported that naturally infected croakers had lesions over 60–80% of their body surface, which was an unusually high level of infection. Experimental infections or those that develop under confined conditions tend to produce lesions that cover a larger portion of the body surface.

Pathology Lymphocystis virus infects fibroblast cells of the skin and occasionally the internal organs. Hypertrophied cells may be 20–1,000 times larger than normal 10–100 µm cells (Figure 10.2) (Anders 1989). These cells are generally round to oval and easily visible. In histological sections, cells contain an enlarged, centrally located nucleus and Feulgen-positive, basophilic, ribbon-shaped cytoplasmic inclusions containing viral DNA (Dunbar and Wolf 1966). Morphology of the inclusions can vary with fish species, but are considered sites of viral maturation (Wolf and Carlson 1965). Little inflammation is associated with lymphocystis, but infected cells are often clustered in a collagen hyaline matrix (Howse and Christmas 1970). Electron micrographs show that inclusion bodies are dense areas of hexagonal-shaped virus particles, which are transported to different sites by the blood stream. In most instances, lymphocystis lesions regress after a period of time, cells undergo lyses or slough, and infected areas heal leaving little evidence of infection.

Significance Lymphocystis virus-infected fish generally do not die as a result of infection, but they do have reduced economic value. Entire populations of juveniles or adult fish may be rejected by sport and commercial fishermen if heavily infected with lesions even though lesions are transient.

Viral erythrocytic necrosis Viral erythrocytic necrosis (VEN), a viral infection of erythrocytes, affects a wide variety of marine and anadromous fish species. VEN was known as piscine erythrocytic necrosis (PEN), until Walker and Sherburne (1977) suggested that the nature of the disease would be better

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reflected if emphasis were placed on blood instead of fish, thus “VEN.” Laird and Bullock (1969) proposed a viral etiology for VEN, and Walker (1971) later provided electron microscopic evidence to support that etiology. The etiological agent of VEN is erythrocytic necrosis virus (ENV). Wolf (1988) cited literature from the early 1900s that described the disease, but at that time, its cause was attributed to an intracellular parasite.

Geographical range and species susceptibility VEN was reported in Atlantic cod, Atlantic seasnail, and shorthorn sculpin in waters of the North Atlantic of Canada and the U.S. coast (Laird and Bullock 1969). Later, VEN was reported in the United Kingdom (Smail and Egglestone 1980), Portugal (Eiras 1984), Mediterranean Sea (Pinto et al. 1989), and the Pacific region of the United States (Rohovec and Amandi 1981). Essentially, VEN is confined to marine environments but has been reported in eels in Taiwan (Chen et al. 1985) and freshwater stocks of anadromous salmon

(Rohovec and Amandi 1981). Walker and Sherburne (1977) listed 12 species of fish from the Atlantic coast of North America that are affected by VEN, 5 of questionable susceptibility, and 25 that appear to be refractive. Affected species are primarily alewife, rainbow smelt, sea raven, and European sea bass (Pinto et al. 1989). Lamas et al. (1995) reported VEN in cultured turbot in the Mediterranean Sea along the Spanish coast. Other economically important susceptible fish species are chum, coho, chinook, and pink salmon, steelhead trout, Atlantic herring, and striped mullet (MacMillan et al. 1980; Rohovec and Amandi 1981; Eiras 1984).

Clinical signs The principal clinical sign of VEN is severe anemia characterized by pale gills, watery colorless blood, and discolored livers (Meyers et al. 1986). Stricken fish will be listless or swim in circles and have hematocrits as low as 2–10% (Evelyn and Traxler 1978). Pacific herring with VEN have hemorrhages in the epithelium (Figure 10.3). Walker and

Figure 10.3 Pacific herring with viral erythrocytic necrosis virus. Note the hemorrhages in the epithelium (arrows). (Photo courtesy of J. R. Winton.)

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(a)

(b)

Figure 10.4 Viral erythrocytic necrosis. (a) Erythrocytes with singular cytoplasmic inclusion bodies (arrows) and (b) electron micrograph of the hexagonal viruses in Atlantic cod erythrocytes (bar = 200 nm). (Photos courtesy of B. Nicholson.)

Sherburne (1977) reported that captured infected cod were somewhat emaciated and darker than noninfected fish.

Diagnosis VEN is confirmed by electron microscopy because virus has not been propagated in estab-

lished cell lines (Evelyn and Traxler 1978). However, Reno and Nicholson (1980) successfully replicated ENV in vitro in cod erythrocytes at 4◦ C. VEN infections can be detected by staining blood smears and observing the erythrocytic cytoplasm inclusion bodies. Nuclei of infected cells may be round, bilobed, or “U” shaped (Figure 10.4) (Walker and Sherburne 1977). Infected erythrocytes contained round

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Table 10.2 Summary of pathology of viral erythrocytic necrosis infection in some fish. Fish Host

Percent Erythrocytes Affected

Pathology

Atlantic cod Common blenny Alewife Atlantic herring Pacific herring Chum salmon

0.01–99, mature 3–60, mature 0.17, mature 90, mature and immature Mature and immature 80, mature and immature

Skin darkening, no mortality None None Hyperemic liver, hemolysed blood, two types of inclusions Erythroblastosis 0.3% mortality, prone to BKD, vibriosis and anemia

Source: Laird and Bullock (1969); Evelyn and Traxler (1978); Reno and Nicholson (1980); Piento et al. (1989).

to lobate eosinophilic (Giemsa stain) inclusion bodies in the cytoplasm (Figure 10.4). Depending upon infection stage and severity, 9% to 100% of erythrocytes will contain these inclusions. Pinto et al. (1989) found erythrocytic inclusions in 2.7–25% of VEN-infected European sea bass. The inclusion body is also Feulgen-positive, fluoresce bright green with acridine orange, and most likely contains the replicating pool of viral nucleic acid.

Virus characteristics Electron microscopy of ENV-infected erythrocytes will reveal hexagonal virions in the cytoplasm (Figure 10.4). Although serological methods for ENV identification are not available, Pinto et al. (1989) did purify viral material from infected erythrocytes of sea bass and developed an immunofluorescent reagent that allowed viral differentiation from similar viral fish infections. The ENV that has been described to date is an icosahedral virion with a DNA genome classified as a member of the family Iridoviridae. The size of ENV virions varies with afflicted fish species. Virus described in erythrocytes of affected Atlantic herring and salmonids was 140–190 nm (Evelyn and Traxler 1978), but a larger 300–350 nm virus was reported in infected Atlantic cod (Walker and Sherburne 1977). Without benefit of serological procedures and in vitro culture, it is impossible to determine if all of these viruses

are the same; however, with the exception of particle size, they are very similar.

Epizootiology The characteristics of a VEN outbreak vary depending on fish species and degree of infection (Table 10.2). Mean hematocrits decline to less than 10% from a normal of near 40% in chum salmon (MacMillan et al. 1980) and even lower in other species. It is proposed that newly infected circulating erythrocytes are destroyed in the hematopoietic centers of the spleen and anterior kidney, thus facilitating virus release. The host compensates for this anemia by erythroblastosis, but additional infection of erythroblasts with virus may prolong infection. Smail and Egglestone (1980) recorded a 32% incidence of VEN in young-of-the-year Atlantic cod compared to an almost zero incidence in 3- to 4-year-olds, although the older fish may have recovered from an earlier infection. Similarly, MacMillan and Mulcahy (1979) detected 43% incidence of VEN virus in young Pacific herring compared to 4% incidence in 4- to 8-year-old fish. Meyers et al. (1986) found a VEN prevalence of 56–l00% in 1 year olds and 17–80% in older Pacific herring in Alaska. Captive European sea bass had a 20–100% VEN infection compared to 6% in wild sea bass (Pinto et al. 1989). These collective data indicate that younger and confined fish are more susceptible to VEN. In their

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study of VEN in Pacific herring in Puget Sound, Washington, in 2005–2007, Hershberger et al. (2009) found 67% prevalence in 1-year-old fish in October, 2005, that waned to 0% infection in August 2006. A second epizootic occurred in the summer of 2007 in the 2006-year class. The infection route of ENV is most likely from fish to fish through the water. Experimentally ENV can be transmitted by injection with filtrates derived from infected fish tissue. Eaton (1990) transmitted ENV from Pacific herring to pink, chum, and sockeye salmon with chum salmon being the most susceptible. These studies show that ENV from one fish species will infect another species. Blood-sucking parasites such as the salmon louse (Lepeophtheirus sp.) and gill maggot (Salmincola sp.) may also serve as vectors (Smail 1982). Evelyn and Traxler (1978) found that chum and pink salmon that were injected with cell-free filtrates became clinically diseased in 12 days (chum) and 3 weeks (pink). Infection rate in chum salmon increased from 50% at 12 days to 100% at 48 days postinoculation. These researchers also reported that at 7 months postinoculation, four infected fish were ENV negative, indicating that recovery from VEN is possible. Rohovec and Amandi (1981) reported an ENV infection in 20 g coho salmon in freshwater. The fish were experiencing mortalities, but no clinical signs other than hematocrits of less than l0% were noted. Reno et al. (1985) reported that ENVinfected Atlantic cod and Atlantic herring exhibited lower hematocrits and erythrocyte counts, but a reduced hemoglobin concentration occurred in cod. There was no effect on plasma electrolyte or protein concentration in either species. Artificially, ENV-infected chum salmon developed lowered hematocrits and hemoglobin concentrations (Haney et al. 1992). The ENV-infected salmon also had higher white blood cell counts and more fragile red blood cells than the controls. After 4 weeks of infection, the fish began to recover. However, infected fish recovered more slowly

from stress and had greater osmoregulatory difficulties than did control fish. Meyers et al. (1986) also found evidence that ENV-infected Pacific herring suffered osmoregulatory stress that could precipitate death, particularly in young fish. Survival of ENV-infected fish varies. MacMillan et al. (1980) and Rohovec and Amandi (1981) assayed 35 stocks of sea-run salmonids and found 25% to be positive for erythrocytic inclusions. Percentage of ENVpositive fish within the respective populations ranged from 1.0 to 15.1%. When evaluating the effect of VEN on released salmon, mortality directly due to the viral disease was limited; however, environmental stressors caused higher mortality in infected fish than in uninfected fish. It appears that in most cases VEN is not directly responsible for deaths of infected fish, but it renders the fish more susceptible to environmental insults and secondary bacterial and/or fungal infections. Hershberger et al. (2009) found many persistent and recurrent outbreaks of VEN in Pacific herring in the Puget Sound in Washington and Prince Williams Sound in Alaska and concluded that the disease is probably common among Pacific herring throughout the eastern North Pacific Ocean. They also suggested that with this prevalence, VEN likely impacts Pacific herring at the population level, and its influence may be coverted and not easily detected.

Pathology Appy et al. (1976) described pathology that takes place in the erythrocytes of cod infected with VEN. The nucleus becomes more rounded and densely stained with the presence of magenta-colored cytoplasm inclusions that measure 1–4 µm in diameter (Figure 10.4). In some cells, inclusions appeared to possess a small central vacuole, which increased in size with subsequent degeneration. Infected cells appeared more rounded and irregular than

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uninfected cells. Some cells fractured and resulted in the formation of fragments containing remains of the inclusion. Cytoplasmic inclusions of infected cells are composed of a pool of icosahedral viral particles (Figure 10.4) with incomplete viruses at the edge of the pool.

Significance Prevalence of VEN in some fish populations can be high, but the direct impact on wild populations is unclear, and its actual importance to these stocks is unknown. However, it poses a threat to maricultured populations. While limited mortality can be directly attributed to VEN, the virus makes infected fish more susceptible to opportunistic bacterial diseases (i.e., Vibrio anguillarum) as well as more sensitive to low dissolved oxygen and handling stressors (Reno et al. 1985; Haney et al. 1992).

Viral nervous necrosis Viral nervous necrosis (VNN), caused by betanodaviruses, is also called viral encephalopathy and retinopathy (VER), or piscine nodavirus disease. Genomic sequence differences have been used to distinguish five main fish nodavirus groups based on the type virus; these are striped jack nervous necrosis virus (SJNNV), red spotted grouper nervous necrosis virus (RGNNV), barfin flounder nervous necrosis virus (BFNNV), tiger puffer nervous necrosis virus (TPNNV) (Nishizawa et al. 1997), and turbot nervous necrosis virus (TNNV) (Johansen et al. 2004). The pathology and disease processes appear to be similar in most of the VNN outbreaks reported.

worldwide (Nakai et al. 1995; Curtis et al. 2001). This disease has been diagnosed in Japan, Spain, Portugal, Tahiti, Greece, Martinique, France, Norway, India, Israel, Italy, Tunisia, the United States (California), and the Atlantic coast of Canada (Breuil et al. 1991; Muroga 1995; Le Breton et al. 1997; Dalla Valle et al. 2000; Castric et al. 2001; Curtis et al. 2001; Barker et al. 2002; Athanassopoulou et al. 2004; Thiery et al. 2004; Azad et al. 2005). Essentially, VNN virus is typically endemic in marine waters but occasionally occurs in freshwater fishes. A variety of marine fish species are affected by VNN. Muroga (1995) listed 14 fish species susceptible to VNN including barramundi, groupers, parrot fish, flounders and turbot, puffers, striped sea bass, and several other species of sea bass (Dalla Valle et al. 2000; Castric et al. 2001; Curtis et al. 2001; Azad et al. 2005). Ucko et al. (2004) isolated VNN from five species of fish in the Red Sea and Mediterranean Sea from clinical diseased fish as well as healthy fish. Larval striped jack are prominently affected by VNN, and Le Breton et al. (1997) reported VNN disease in production-size European sea bass. Arimoto et al. (1993) showed that larval yellowtail and gold-striped amberjack were refractive to SJNNV, and Muroga (1995) listed 14 other fish species, including barramundi, groupers, other jacks, parrot fish, flounders, and puffers that are affected by VNN. Although, most cases of VNN have been found in saltwater, Athanassopoulou et al. (2004) demonstrated that white sturgeon in freshwater aquaculture facilities in Greece became infected by VNN and developed clinical disease on a farm that contained infected sea bass. Later, Bigarre et al. (2009) identified VNN in freshwater-reared tilapia in Europe.

Species susceptibility and geographical range

Clinical signs

VNN has been found in a variety of wild marine and cultured fish species in numerous geographical regions and the virus may be found

Externally, fish infected with VNN are darker in color with an opaque cornea or bilateral exophthalmia, and they become hyperactive and

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erratic when disturbed. When larval striped jack are injected with virus, clinical signs indicate that the central nervous system is affected (Arimoto et al. 1994; Curtis et al. 2001). Infected fish become inappetent, emaciated, and larger fry develop enlarged swim bladders and vertebrae deformities. Larger VNNinfected European sea bass exhibited reduced feeding, lethargy, and abnormal swimming (Le Breton et al. 1997). Fish rotated on the long axis, swimming up and down; moribund fish came to the surface belly up with a curved body.

Diagnosis Piscine nodaviruses do not replicate in many fish cell lines, but VNN virus does replicate in striped snakehead cells (SSN-1) cells at 25◦ C (Frerichs et al. 1991; Iwamoto et al. 1999). Dannevig et al. (2000) also isolated VNN from Atlantic halibut in Norway using SNN-1 cells in which CPE, characterized by rounded granular cells with heavy cytoplasm vacuoles, appeared in 5 days. The monolayer either partially or completely disintegrated over 3- to 6-day postinoculation, but the monolayer regenerated in 10–14 days. The virus was identified by indirect immunofluorescence and RTPCR. Other definitive methods of diagnosing the virus in infected larvae include ELISA (Arimoto et al. 1992; Nishizawa et al. 1995), FAT utilizing both monoclonal and rabbit polyclonal antibody (Nguyen et al. 1997), and RTPCR (Nishizawa et al. 1994; Dalla Valle et al. 2000; Hick and Whittington 2010). A sensitive diagnostic molecular test described by Dalla Valle et al. (2000) was based on RT-PCR plus a nested PCR in combination. The two-step molecular method was 104 times more sensitive than isolation in cell culture, and it identified VNN in sperm and ovarian tissue. Hick and Whittington (2010) describe a quantitative RT-PCR method to detect VNN and found it even more sensitive compared to previously reported methods. Using ELISA, Breuil et al. (2001) provided a sensitive method for detect-

ing two nodavirus strains (Sb1 and Sb2) from sea bass along the Mediterranean and Atlantic French coast and recommended the method to be used by OIE for detection of these viruses.

Virus characteristics Fish nodavirus infectious particles are 25– 34 nm in diameter icosahedral unenveloped virions that replicate in cell cytoplasm (Mori et al. 1992; Azad et al. 2005). The virus genome consists of two single strands of positive-sense RNA. The largest, RNA1 (3.1 kb), encodes RNA-dependent RNA polymerase while RNA2 (1.6 kb) encodes the coat protein. A subgenomic mRNA generated from RNA1 during infection produces a third protein, which inhibits RNA silencingsuppression activity, enhancing the ability of the nodavirus to replicate (Iwamoto et al. 2005). Sequence comparisons among fish nodavirus isolates have allowed the identification of a variable region in RNA2 that can be used to differentiate the fish nodaviruses into four distinct types: SJNNV, RGNNV, BFNNV, and TPNNV (Nishizawa et al. 1997). A system for regenerating fish nodavirus by transfecting RNA produced from cDNA clones was used to compare two strains to identify a region of the genome that imparts host specificity. The region responsible was the variable portion of the coat protein (Ito et al. 2008).

Epizootiology In the last 10–15 years, incidences of nodavirus-associated diseases of fish have increased dramatically in marine environments around the world. These viruses occur in wild fish populations as well as in mariculture and ornamental populations. Striped jack nervous necrosis virus severely affects larval striped jack when they are 2–20 days old (Arimoto et al. 1994). Arimoto et al. (1993) also reported disease occurrence in 3.5–4.4 mm long striped jack larvae, which remained susceptible

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at 78 mm in length. The SJNNV creates a viremia that affects nearly all organs and tissues and is most likely transmitted horizontally. Sea bream appeared to be potential carriers of VNN, as they did not show clinical infection, but the virus was detected in their tissues by PCR and juvenile sea bass cohabiting with them developed VNN (Castric et al. 2001). Using PCR, Mushiake et al. (1994) demonstrated the potential for vertical transmission when they detected SJNNV in spawning striped jack. They found that by examining fish for virus just before spawning, SJNNVpositive fish could be identified and eliminated from spawning activities, thus preventing vertical transmission. However, Nishizawa et al. (1996) showed that this procedure is not 100% effective in detecting fish with very low levels of SJNNV. Nevertheless, PCR screening of adults and utilizing only SJNNV-negative fish for spawning resulted in reduced numbers of virus-infected larvae. Using FAT and PCR methods, Nguyen et al. (1997) detected nodavirus in the reproductive fluid of a 13-year-old adult striped jack during spawning but found no virus in fluids of 4-year-old fish that had not spawned. Virus was also detected in the gonad, intestine, stomach, kidney, and liver of a 13-year-old fish by FAT using rabbit anti-SJNNV serum, suggesting that virus originates in these organs of striped jack and is shed from intestines and gonads during spawning, resulting in contaminated eggs. This theory was supported by Dalla Valle et al. (2000), who detected VNN virus in sperm and ovarian tissue of European sea bass using the two-step molecular detection method (RT-PCR plus nested PCR). Mortality of SJNNV-infected striped jack is often near 100% in larvae less than 10 days old, but older fish are less severely affected (Arimoto et al. 1994). VNN may occur in striped jack at water temperatures from 20 to 26◦ C, but progresses most rapidly at 24◦ C. Larger European sea bass were affected by VNN, but Breuil et al. (1991) noted that mortalities ceased when fish reached 5 g in weight.

However, Le Breton et al. (1997) reported VNN in European sea bass that were 10–45 g and 350–580 g; both groups suffered mortalities of 11–60% during July and September when water temperatures were 24.5–26.5◦ C.

Pathology Naturally (acute and subacute stages of natural infections) and experimentally, SJNNVinfected striped jack were studied by histopathology for progression of VNN (Nguyen et al. 1996). General features of pathogenesis in these three groups were similar. Nerve cells in the spinal cord, especially above the swim bladder, were initially affected, and later the brain and retina became necrotic and vacuolated. Deaths occurred 1–2 days after lytic degeneration and vacuolation of nervous tissue cells. Results of FAT, histopathology, and electron microscopic observations indicate that SJNNV exhibits a tropism for nerve cells and initial multiplication occurs in the spinal cord from which virus spreads to the brain and finally the retina and presumably other tissues and organs. Le Breton et al. (1997) described necrosis of the cornea and frontal area of the head with blood vessels of the brain severely congested. Large vacuoles were present in the brain and retina.

Significance In Japan VNN is most severe in larval striped jack where it can be devastating. The presence of VNN in other parts of the world and its potential to affect a large number of fish species make the nodaviruses potentially dangerous pathogens to mariculture.

Red Sea bream iridoviral disease The red Sea bream iridoviral disease (RSIV), also known as megalocytivirus diseases, was

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first recognized in 1990, causing high mortality in cultured red Sea bream in Japan (Inouye et al. 1992). The disease is caused by a member of a newly recognized Megalocytivirus genus of Iridoviridae. There are several related viruses representing at least three distinct genogroups that cause similar diseases (Song et al. 2008). This group of pathogens include infectious spleen and kidney necrosis virus (ISKNV), turbot iridovirus, grouper sleepy disease iridovirus, orangespotted grouper iridovirus (OSGIV), rock bream iridovirus (RBIV), dwarf gourami iridovirus, African lampeye iridovirus, and the large yellow croaker iridovirus (Wang et al. 2007). Red Sea bream iridovirus is an OIE reportable pathogen (OIE 2009).

Species susceptibility and geographical range Since its detection in the early 1990s, RSIV disease has been found throughout East and Southeast Asia. Naturally occurring clinical RSIVD has been documented in over 30 species of marine fish, most of which are Perciformes or flatfish (Matsuoka et al. 1996; Kawakami and Nakajima 2002).

Clinical signs Diseased fish become lethargic, exhibit severe anemia, petechiae on the gills, and an enlarged spleen. Stained imprints of kidney and spleen reveal enlarged darkly staining cells.

et al. 1995). The M10 antibody will detect both RSIV and ISKNV. Molecular detection can be done using PCR (Kurita et al. 1998). These viruses can also be cultured on Grunt fin (GF) cells and CRF-1, a Red Sea bream fibroblast line, at 25◦ C. They produce CPE consisting of rounded enlarged cells usually within 4 days postinoculation (Nakajima and Sorimachi 1994; Imajoh et al. 2007). Other cell lines such as BF-2 cells may initially produce CPE with RSIV, but the virus may be lost after passage (Imajoh et al. 2007). Confirmation of the virus in cell culture requires IFAT or PCR.

Virus characteristics The RSIV and related megalocytiviruses have similar characteristics to other fish iridoviruses. The virions are large 200–240 nm icosahedral particles that assemble in the cytoplasm and contain a large (>110 kb), linear, heavily methylated double-stranded DNA genome that is terminally redundant and circularly permuted. The virions are sensitive to chloroform, ether, heat, and acid pH, and replication is inhibited by 5-iodo-2deoxyuridine (Nakajima and Sorimachi 1994). He et al. (2001) were the first to sequence the entire genome of a megalocytivirus. They sequenced ISKNV and demonstrated that it was substantially different from the ranaviruses and lymphocystis viruses. Since then, the genomes of OSGIV and RBIV have been sequenced and compared. Eaton et al. (2007) re-annotated the published sequences and demonstrated very close similarity and the presence of 110 orthologous genes.

Diagnosis Epizootiology This disease can be presumptively diagnosed based on clinical signs and the identification of enlarged dark staining cells in Giemsa-stained spleen imprints. The disease can be confirmed if RSIV-specific antibody (such as the monoclonal antibody M10) labels the enlarged cells using indirect immunofluorescence (Nakajima

Infection by megalocytiviruses related to RSIV is widespread in many species of wild and cultured fish in Asia. Most cases of the megalocytivirus diseases are in the marine environment, but this disease has been found in freshwater cultured ornamental fish. The disease is

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primarily a problem at warm temperatures (above 20◦ C), and therefore it is usually seen in the summer and fall. The data do not suggest vertical transmission. Research suggests that fish that have recovered from infection may become resistant to reinfection. In view of this, a vaccine, a formalinkilled preparation, for RSIV was developed by Nakajima et al. (1999). In a field trial, juvenile Red Sea bream were vaccinated by IP injection and, along with nonvaccinated control fish, were moved to marine net pens for natural exposure to the virus and observed for 12 weeks. Cumulative mortality caused by RSIV in the vaccinated fish was 19.2% compared to 68.5% in the control fish. Virus antigen was not detected in spleen of vaccinated but was present in the spleen of control fish. Also, the body weight of the vaccinated fish was significantly higher than the control fish. The results suggest that a vaccine against RSIVD can be an effective tool to combat the disease. Recently DNA vaccines using plasmids expressing the MCP gene and a transmembrane protein gene were evaluated and shown to provide protection. The formalin-inactivated RSIVD vaccine is commercially available for Red Sea bream, striped jack, and members of the genus Seriola (amberjack and yellowtail) in Japan (OIE 2009).

Pathology Fish with megalocytivirus infections display enlarged basophilic staining cells in the heart, gills, spleen, and hematopoietic tissue. These cells stain blue with Giemsa stain and are Feulgen positive. Necrosis is observed in the hematopoietic tissue.

Significance The megalocytivirus diseases cause substantial losses in marine aquaculture facilities in Asia. The potential for high losses and the broad

host range make it an important disease concern. The RSIV is an OIE reportable disease (OIE 2009).

Other fish viruses Numerous fish viruses cause important diseases of fish that are not among the most commonly cultured species for commercial production. For the most part, these viruses infect wild fish populations, but some affect fry and fingerling populations being raised for stocking natural waters. This group of pathogens includes cytolytic viruses, i.e., rhabdoviruses, iridoviruses, and tumor-causing viruses.

Pike fry rhabdovirus disease Pike fry rhabdovirus disease (PFRD) is an acute, highly infectious disease commonly called “head disease” in cultured fry and “red disease” in cultured fingerling northern pike (Bootsma 1971; de Kinkelin et al. 1973). The etiological agent, pike fry rhabdovirus (PFR), was characterized by de Kinkelin et al. (1973) and classified as a vesiculovirus that is similar to spring viremia of carp virus. It was noted that “head disease” had previously been reported in The Netherlands in 1959 and “red disease” in 1956; however, B´ek´esi et al. (1984) deemphasized the importance of this disease. PFR was isolated from pike in hatcheries in The Netherlands and Germany, serologically similar rhabdoviruses were isolated from grass carp (Ahne 1975a), tench, white bream (Ahne et al. 1982), European catfish (Jørgensen et al. 1989), brown trout (Adair and McLoughlin 1986), roach (Haenen and Davidse 1989), and gudgeon (an ornamental fish accidentally introduced into Europe with grass carp) (Ahne 1975b; Ahne and Thomsen 1986). These isolates have subsequently been identified as PFR. PFR has also been found in Hungary, France, and the United Kingdom but has not been reported in North America.

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Figure 10.5 (a) Pike infected with pike rhabdovirus showing “head disease” with hydrocephalus and (b) pike with “red disease” showing hemorrhaging in the lateral musculature. (Photos courtesy of R. Bootsma.)

Two forms of PFRD occur in northern pike, one of which is commonly known as “head disease,” affects swimming fry (de Kinkelin et al. 1973). Infected fry have a raised cranium (Figure 10.5) as a result of hydrocephalus, have exophthalmia, and exhibit poor growth. The second form occurs in larger fish, which develop severe hemorrhages along the lateral trunk musculature, thus the name “red disease” (Figure 10.5). Hemorrhagic areas are swollen and gills are pale, and infected fish exhibit bilateral exophthalmia, abdominal distension, and ascetic fluid accumulation in the body cavity. In both forms, fish lose schooling behavior, and individuals either swim lethargically at the surface or lie listlessly on the bottom. Although

fish species other than northern pike can become infected with PFR, clinical signs of these infections are poorly defined. PFR is detected by isolation of virus in FHM or EPC cells (de Kinkelin et al. 1973). Kidney, spleen, and intestines provide the best tissue sources for virus isolation. Infected cells incubated at 20◦ C develop focal areas of rounded cells in 2–3 days. Virus identification is obtained by serum neutralization, but when using homologous rabbit antiserum, PFR neutralization is dependent upon complement (Clerx et al. 1978). Also, other viruses (i.e., SVC), are partially neutralized by PFR-specific antiserum (Ahne 1986). This partial cross-reactivity can be removed by reacting PFR with the

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antiserum against other rhabdoviruses of fish. PFR is serologically distinct from VHSV and IHNV, but some cross-reactivity occurs with SVCV (see Table 6.l). Cross-reactivity with SVC, an OIE reportable disease, has led to the development of more definitive nucleic acid-based identification methods. Ahne et al. (1998) reported on the use of RNAse protection assays to distinguish the two viruses, and Stone et al. (2003) used degenerate RT-PCR and sequencing of the fragment of the glycoprotein gene to place the fish vesiculoviruses into four genogroups with genogroup I being SVCV and genogroups II–IV containing viruses identified as PFR. The causative agent of PFRD is a rhabdovirus in the genus Vesiculovirus that measures l25–155 nm in length and about 80 nm in diameter (Bootsma and van Vorstenbosch 1973). It is enveloped; the virus has a singlestrand, nonsegmented RNA genome (Ahne 1975b). The genome consists of 11,097 nucleotides and contains five genes, encoding the nucleoprotein, phosphoprotein, matrix protein, glycoprotein, and RNA-dependent RNA polymerase (Chen et al. 2009). PFR replicates well in FHM cells at 10–28◦ C with titers reaching greater than 109 PFU/mL (de Kinkelin et al. 1973). Epithelioma papillosum of carp, BB, and RTG-2 cells are also susceptible to PFR, but virus yield is lower than that in FHM cells. Infectivity persists in tissue culture medium with serum at 4◦ C for 5 months but decreases at higher temperatures. Seventy percent of infectivity is lost in 3 days in water at 14◦ C. Fry and small fingerling pike (up to 5 cm) are susceptible to PFR with up to l00% mortality (Bootsma 1971). Haenen and Davidse (1993) showed that a PFR isolate from asymptomatic roach was highly pathogenic to young roach (85% mortality) following waterborne exposure, mildly pathogenic to carp (45% mortality), and avirulent to rainbow trout, concluding that PFR could pose a health threat to cyprinid farms as well as cultured pike farms. The disease occurs in early spring when water temperatures are in the 10◦ C range.

The natural transmission mechanism for PFR is not known. Bootsma et al. (1975) were unable to isolate the virus from frozen kidney, liver, spleen, intestine, or gonads of brood pike even though progeny of these fish were infected. Bootsma and van Vorstenbosch (1973) also experimentally infected pike eggs with PFR, and within weeks of hatching, there was 100% fry mortality; however, definitive proof of vertical transmission is lacking. Horizontal transmission of PFR has been experimentally demonstrated following waterborne exposure and injection of juvenile northern pike (Ahne 1985). Fry (0.2g) bathed in virus and then fed to larger pike (60 g) transmitted PFR to recipient fish, but deaths or overt disease did not occur. PFRD can be confused with other viruses that affect pike. Ahne (1985) demonstrated susceptibility of 0.2-g pike fry to other viruses by exposing them to water bath containing IPNV, SVCV, or VHSV, resulting in 42–90% mortality. Grass carp reovirus caused 64% mortality under similar circumstances. Dorson et al. (1987) confirmed that experimentally 10day to 2-month-old pike fry are susceptible to IPNV, IHNV, VHSV, and perch rhabdovirus, all of which can cause hemorrhaging, exophthalmia, and up to 74% mortality. Therefore, when virus is isolated from pike fry, definite virus identification should be made. It was postulated by Ahne (1986) that pike could play the role of vector for other pathogenic viruses normally not found in this species. In “head disease,” there is an accumulation of cerebral fluid in the ventricle of the mesencephalon, causing swelling (Figure 10.5) (Bootsma 1971). Petechia can be found in the brain, spinal cord, and spleen; kidney tubules show degeneration with necrosis of epithelial cells. “Red disease” is characterized by petechia throughout the spinal cord, pancreas, and hematopoietic tissue of the kidney, but severe muscle hemorrhage is the most dramatic pathological lesion with erythrocytes and inflammatory cells congregating between bundles of necrotic muscle. There is extensive

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necrosis of excretory kidney tubules, and the cerebral region becomes edematous. The liver, heart, and gastrointestinal tract appear normal in both forms of disease. When PFRD occurs in juvenile pike, it can be devastating. In 1972, only 0.6% of 1.85 million cultured young pike in The Netherlands reached a length of 4–5 cm, and it was estimated that approximately 86% of the mortality was due to PFRD. PFRD has had a significant effect on northern pike fry culture in the past, but due to egg disinfection with iodine at spawning, incidence of disease has been reduced (P. de Kinkelin, Laboratoire de Ichtyopathologie, Thiverval-Grignon, France, personal communication). Research by Haenen and Davidse (1993) indicated that seed carp farms could also be at risk for the virus.

Perch rhabdoviruses Two independent studies indicate that in addition to PFR and spring viremia of carp virus, there is at least one other genogroup of the Vesiculovirus genus of rhabdoviruses in Europe, the perch rhabdovirus. Rowley et al. (2001) compared rhabdovirus isolates from Northern Ireland in 1998–1999 to other PFRlike vesiculovirus isolates from Ireland and Holland and to the type strains of SVCV and PFR. They found that all PFR-like viruses reacted with PFR-specific antiserum. However, nucleic acid sequencing of a portion of the glycoprotein gene revealed that the Northern Ireland isolates were closely related to the other PFR-like viruses but nevertheless distinct from PFR and SVCV. The isolates studied came from wild common bream during a disease outbreak accompanied by high mortality in May, 1998, healthy farmed rainbow trout and brown trout at the same time, and healthy wild bream and roach in the same stretch of river 11 months later. Experimental infection was established in mirror carp producing 12% mortality. Serological testing by ELISA and serum

neutralization indicated that the 1998 isolates were somewhat similar to PFR but distinct from spring viremia of carp virus. Betts et al. (2003) evaluated rhabdovirus isolates from perch, grayling, northern pike and pike perch, and largemouth bass in France between 1981 and 1995 and compared them to a northern pike isolate from Denmark, brown trout isolates from Finland, and to the type specimens of PFR and SVCV. Using serum neutralization tests, immunofluorescence, SDS-PAGE, and nucleotide sequencing, they found that these French and Danish isolates grouped with perch rhabdovirus and were distinct from lake trout rhabdovirus, PFR, and SVCV. They proposed that four distinct vesiculo-type viruses are endemic to Europe. Many of these isolates were obtained from diseased fish during high-mortality events, and several have been shown to cause disease in experimental challenges; however, the economic and environmental impact of these rhabdoviruses is not clear.

Aquabirnaviruses Aquabirnaviruses are becoming more prevalent in freshwater and marine waters. Aquabirnavirus, of which there are several serogroups, is a genus of the family Birnaviridae. These viruses are non-enveloped, icosahedral and have a two-segmented double-stranded RNA genome (Dobos et al. 1995). Infectious pancreatic necrosis virus is the type species, but “new” viruses are being detected in a wide variety of fish species. Isshike et al. (2001) determined the infectivity of seven strains of marine birnaviruses (MABV) in five fish species and found only three of the virus strains caused mortality in yellowtail (0–64%) and amberjack (0–28%). Some of the MABV strains replicated in the host fish but caused no overt disease or mortality. They concluded as follows: “The infectivity of MABV for different fish species is considered to change, which is an important factor in the development of

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the infection cycle of this virus among marine organisms.” An aquabirnavirus was isolated from wild flounder and mummichog in the Chesapeake Bay, Virginia (Ahne et al. 2003). In their RTPCR analysis of the isolate, it was shown to belong to serogroup A, which is the most common serogroup in the United States.

Epizootic ulcerative syndrome associated viruses Epizootic ulcerative syndrome (EUS) is a multifactoral disease of many species of fish in Australia, India, Thailand, The Philippines, and numerous other south and southeastern Asian countries (Lilly et al. 1998; Lio-Po et al. 2003). The number of affected species is possibly nearly 100 cultured and wild fish species, but the snakehead and walking catfish appear to be the most severely affected cultured fish. The disease, primarily one of freshwater but may also occur in estuaries, is characterized by shallow or deep ulcers, with red centers and white margins anywhere on the body. Histological lesions are characterized by granulomatous dermatitis and muscle inflammation. EUS is associated with multiple etiologies including bacterial (Aeromonas hydrophila), fungal (Aphanomyces sp.), and now two viruses have been isolated from EUS-affected fish. Lio-Po et al. (2000) isolated a rhabdovirus from snakehead in The Philippines (EUS-P) and John et al. (2001) described snakehead reovirus (SKRV) from EUS-infected snakehead in India (EUS-I). The Philippine rhabdovirus was isolated from gills, liver, spleen, kidney, and muscle lesions of snakehead in snakehead spleen cells (SHS) incubated at 15–25◦ C (Lio-Po et al. 2000). Channel catfish ovary (CCO) and BF-2 cells were also susceptible. Using serological procedures, the EUS rhabdovirus somewhat resembled ulcerative disease rhabdovirus and infectious hematopoietic necrosis virus, but the Philippine isolate was distinctly different. The

rhabdovirus in infected cells was about 55 × 175 nm in size. However the study did not conclusively establish the virus’ role in the epizootiology of EUS. Lio-Po (2003) transmitted EUS rhabdovirus horizontally from virus infected to naive snakehead by cohabitation in lake water where lesions appeared in 6–14 days on the naive fish. Naive fish exposed to lake water only developed lesions in 16–21 days. Rhabdovirus was isolated from fish in both treatments, suggesting transmission via water from infected fish. Fish in aquifer water only without contact with EUS-infected fish did not develop the disease. The snakehead reovirus isolated from EUS-I had a double-stranded RNA genome with icosahedral symmetry and double capsid. The virus, measuring approximately 71 nm in diameter, was isolated in SSN-1 and SSN-3 cells at 25◦ –30◦ C. However, this virus did not produce typical syncytia CPE associated with other fish aquareoviruses. Serological comparison also indicated the antigenic distinctness of the SKRV isolate from several American and European aquareovirus isolates. These observations demonstrate that EUS remains a condition associated with multiple etiologies, leaving a specific etiology unresolved. However, either the rhabdovirus or aquareovirus could be serious complicating factors in the diseases. Epizootic ulcerative disease syndrome develops primarily in summer and is often associated with environmental stress or poor water quality. The overall impact of EUS is unclear, but while mortality is generally chronic, the long-term effects on a fish population could be significant. Some degree of prevention/control may be obtained by liming ponds, improving water quality, and removing infected fish.

LMBV No debilitating viremia had been reported from largemouth bass or any other Centrarchidae until LMBV was isolated from adult largemouth bass from Santee Cooper Reservoir,

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South Carolina (Plumb et al. 1996). This virus is a member of the Ranavirus genus in the family Iridoviridae and was given the name SanteeCooper ranavirus (Mao et al. 1999). Grizzle et al. (2002) compared the Santee-Cooper LMBV isolate to a virus isolated from largemouth bass in Lake Weir, Florida, in 1991 and showed that the two viruses were identical by measuring several parameters. These authors proposed that the name largemouth bass virus be retained. LMBV is known only in the United States. The disease has been associated with largemouth bass kills or isolated from moribund or dead largemouth bass in as many as 18 southern, eastern, central, and northern states from Florida north to Vermont, Michigan, and Wisconsin and west to Texas and Oklahoma. While most LMBV disease outbreaks have occurred in largemouth bass, other centrarchids have been diagnosed with LMBV infection. Bluegills, bullfrog tadpoles, and metamorphosing bullfrogs were experimentally refractive (unpublished). Moribund largemouth bass infected with LMBV lose equilibrium and float at the sur-

(a)

(b)

face, apparently due to enlarged swim bladders, but otherwise appear normal with no significant or remarkable external lesions. The South Carolina virus isolate came from fish that were frozen in the moribund state and assayed for virus within 48 hours (Plumb et al. 1996). Gills of these fish remained red but soft, eyes were clear, and body surface and fins retained normal color with no macroscopic epidermal lesions. Internally, infected fish looked normal except for the gas gland located on the ventral surface of the air bladder, which appeared more red than normal; the swim bladder was sometimes greatly enlarged. Hanson et al. (2001) noted a brown to yellow residue in the swim bladders in LMBV-infected largemouth bass sampled from a population after LMBV-associated mortality had subsided. LMBV disease is diagnosed by isolation of the virus in FHM or BF-2 cells inoculated with filtrates of homogenized internal organs. Initial focal CPE, which may be visible within 24 hours, consists of a few pyknotic cells that form circular, cell-free areas with rounded cells at the margins (Figure 10.6). As CPE progresses, additional cells become

(c)

Figure 10.6 Largemouth bass virus (LMBV) in FHM cells. (a) Focal Cytopathic effect (CPE) 24 hours postinoculation and incubation at 30◦ C, (b) total CPE of FHM cells 36 hours postinoculation and incubation at 30◦ C, and (c) electron micrograph of LMBV in FHM cytoplasm (bar = 300 nm).

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pyknotic, rounded, and detached until eventually the entire cell sheet is affected. Culture media harvested from infected FHM cells contained 108.9 TCID50 /mL. Grizzle et al. (2003) developed a PCR procedure to detect LMBV in tissue of overtly diseased fish as well as asymptomatic fish using primers designed to specifically amplify the MCP gene of LMBV. The PCR method was significantly more sensitive to virus detection in tissue of overtly diseased fish and asymptomatic fish than inoculated cell cultures. More recently quantitative PCR method developed by Getchell et al. (2007) has proven useful. The LMBV capsid is assembled in the cytoplasm and measures 132–145 nm in diameter (Figure 10.6). With the envelop, acquired after passing through the plasma membrane, the virion measures about 174 nm in diameter. Infected FHM cell cultures stained with acridine orange develop an apple green cytoplasm, indicative of a double-stranded nucleic acid. Using viral DNA, restriction fragment length polymorphism and sequence determination of the MCP and DNA methyltransferase assays showed that LMBV is a member of Ranavirus genus within the Iridoviridae family (Mao et al. 1999). Furthermore, they found that LMBV was nearly identical to the iridovirus from doctorfish and guppies isolated by Hedrick and McDowell (1995). Goldberg et al. (2003) used partial sequencing of the MCP gene and a non-coding region and found no differences in sequences between a highly virulent LMBV isolate and two less virulent isolates from different regions of the United States. However, amplified fragment length polymorphism analysis demonstrated genetic differences in the three isolates. Experimental studies show that LMBV can be transmitted to smaller, naive largemouth bass by injection or horizontally by cohabitation. However, no experimentally infected fish showed signs of disease other than an inflamed lesion measuring about 0.5 × 1.0 cm at the injection site. Virus was isolated from

the swim bladder, liver, spleen, trunk kidney, ovary, blood, normal muscle, and necrotic muscle from the injection site at 8 and 26 days postinjection. Viremia in largemouth bass that were injected with serially diluted media from virus-infected FHM cells was clearly established with internal organs and muscle, developing virus titers that ranged from 102.3 to 107.5 TCID50 /g. Oral transmission of LMBV was accomplished by injecting guppies with the virus and feeding them by gavage to largemouth bass (Woodland et al. 2002). During a 7-week sampling period, none of the bass developed clinical signs, but virus was isolated from five of seven fish fed the virus. In cohabitation studies, largemouth bass, both with and without direct contact with infected individuals, developed LMBV infections, indicating that waterborne transmission occurs (Grant et al. 2005). Beck et al. (2006) were able to produce clinical LMBV disease (air bladder hyperinflation) in a small percentage of largemouth bass in immersion exposure challenges using larger experimental fish. Whether or not LMBV is totally responsible for epizootics in wild fish is debatable. It is theorized by some researchers that environmentally induced stressors predispose fish to clinical disease and death. In view of this, the significance of LMBV is not clear. Experimental transmission studies indicate that the virus does not appear to be highly pathogenic. However, LMBV has been associated with many largemouth bass-specific fish kills especially when the virus first enters the population, suggesting that LMBV may spread rapidly and cause high losses under certain conditions. In several documented outbreaks of LMBVassociated mortality, losses were substantial among large fish, and infection persisted in the population long after the deaths subsided (Piaskoski 1997; Hanson et al. 2001). There have been no reports of LMBV-associated fish kills in subsequent years after an initial outbreak. Neal et al. (2009) evaluated the population impact of LMBV on a small lake and

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found no substantial change in the population makeup after the event.

logical effect of fish iridoviruses ranges from insignificant to serious.

Other iridoviruses

Other herpesviruses

During the past 20 years, in addition to the previously discussed iridovirus, a relatively large number of fish viruses have been isolated and classified as members of the family Iridoviridae (Williams 1996). Heretofore, most viruses in this group were from insects, other arthropods, and amphibians, but the number of iridovirus isolations from fish has increased (Mao et al. 1997). Some iridoviruses from widely different geographical areas and divergent taxonomic fish species are similar; EHNV from redfin perch (Australia), the virus from European catfish (Germany), and black bullhead (France) were similar in spite of diverse geographical origin (Hedrick et al. 1992). Using molecular characterization and sequence analysis, Mao et al. (1997) determined that nine isolates of iridovirus from fish, reptiles, and amphibians were more similar to frog virus-3 (genus Ranavirus) than to lymphocystis (genus Lymphocystisvirus). These authors suggested that the viruses shared features based on geographical origin rather than group of vertebrates from which they were isolated. The best known iridovirus-induced fish diseases are lymphocystis, EHNV, and VEN virus, which have been discussed. Whittington and Hyatt (1996) listed ten iridoviruses that cause systemic viremia in fish. Other iridoviruses have been isolated from sturgeon, carp, goldfish (Berry et al. 1983), guppy, doctorfish (Hedrick and McDowell 1995), turbot in Denmark (Block and Larsen 1993), and the previously discussed LMBV. An iridovirus was also isolated from cultured, imported, brown-spotted grouper in Singapore (Chua et al. 1994), where the disease, known as sleepy grouper disease, has been responsible for significant economic losses in net-cage populations from April to August. The patho-

Herpesviruses are very common pathogens of fish. Many are very host specific and require very specific cell types for culture, and so in many instances susceptible cell lines have not been developed. These herpesvirus diseases have only been diagnosed by electron microscopy or using molecular methods. Most herpesviruses establish long-term latency in their hosts, and several appear to be vertically transmitted. Often herpesvirus diseases cause epidermal lesions such as mild to substantial cellular proliferation or patches of hypertrophic cells.

Pilchard herpesvirus In 1995, a massive mortality attributed to a herpesvirus occurred in Australian pilchard (a sardine-like marine fish) in Australia and New Zealand (Jones et al. 1997; Whittington et al. 1997). At the time of detection in March of 1995, pilchard herpesvirus (PHV) disease was confined to the western, southern, and eastern coasts of Australia where it occurred along 5,000 km of the coastline and northeastern New Zealand where it occurred along 500 km of coastline, which constitutes the general geographical range of the Australian pilchard. Infection progressed from mild to subacute to acute as it moved through the pilchard schools. The epizootic began on the southern coast of Australia, spread to the east and west coast, and then to New Zealand in June and continued in both localities until September. In Australia, the disease spread at about 30 km/day and generally followed the pilchard migration pattern, which was contrary to prevailing currents. Loss estimates were about 10% in some pilchard schools. To date, the

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virus has only been reported in pilchards but has not been reported since the initial epizootics; however, it is considered endemic to the Australian–New Zealand region (Whittington et al. 2008). The incident was thought to have been the most extensive mortality recorded in any fish species in terms of numbers of fish affected and geographical area covered. It serves as an example of how a highly virulent pathogen can affect a dense, schooling population of aquatic animals. Primarily adult pilchards larger than 11 cm were affected. (Hyatt et al. 1997). As stated by Whittington et al. (1997), “Affected fish left the school, ceased swimming, turned onto their sides, resumed swimming sluggishly, and then died within a few minutes.” Once startled, a flashing behavior began within 15 seconds and death occurred in three to five minutes. Some fish died with mouth agape and flared opercula. Tissue around the eyes of fresh dead and moribund fish was congested; gills were dark red to dark brown as compared to a bright red gill in healthy fish. Subacute to acute inflammation of gill lesions was followed by epithelial hypertrophy and hyperplasia. Initially lesions were small and focal but became generalized over a 4-day period. Internally the spleen varied from pale brown to dark red or black. PHV disease was diagnosed by clinical signs and confirmed by histopathology and detection of virus in gill tissues by electron microscopy (Whittington et al. 1997). The virus could not be isolated in several permanent fish cell lines (CHSE-214, RTG-2, BF-2, FHM, or EPC) inoculated with filtrates from homogenized gill and internal organ tissue (Hyatt et al. 1997), but bacteria, parasites, and environmental disturbances were ruled out as possible causes of mortality. A molecular method employing PCR was developed by Crockford et al. (2005) to detect PHV, and it was determined that this virus was molecularly different from Ictalurid herpesvirus 1 (CCV) and Salmonid herpesvirus 2 (Oncorhynchus masou

virus 2). A replicating herpes-like virus was seen in electron micrographs of gill tissue cells from all affected fish examined but not unaffected fish. The virus particles are 92 nm in diameter with many virions containing an envelope. Viral replication and assembly occurs in the nucleus. Virus release was associated with lyses of gill epithelial cells. Pilchards comprise a large part of the gannet’s diet, as well as other marine animals, and since these seabirds were observed eating affected fish, it was speculated that they may have contributed to the spread of disease (Whittington et al. (1997). A small pilchard fishery exists in Western Australia, New South Wales, and New Zealand, but the economic impact of PHV disease on this enterprise was undetermined. A significant loss of pilchards could have an ecological impact because of their importance in the diet of many marine predators.

Herpesvirus of turbot A herpesvirus infection was described in turbot by Buchanan (1978) and again by Block and Larsen (1994) in Europe. In a survey of juvenile turbot, imported from France into Norway, they were examined by histopathology and transmission electron microscopy of head, gill, and epithelium tissue for evidence of a viral infection (Hellberg et al. 2002). Large rounded cells of varying size were found in gill and epithelium; the largest cells were about 80 µm in diameter. These cells occasionally contained vacuoles peripherally to the nucleus. Electron microscopy of these cells showed numerous enveloped icosahedral virus particles of 200–220 nm in the affected cells. The structure of the virus particles were identical to previously described for Herpesvirus scophthalmi virions. Little is known about herpesvirus of turbot. Its geographical range is not identified nor have any massive mortality as a result of the virus been described.

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Figure 10.7 Northern pike with blue spot disease lesions (arrows) on dorsal skin. (Photo courtesy of Tom Jones)

Blue spot disease of pike and muskellunge Blue spot disease of pike and muskellunge is an epidermal disease reported in northern United States, Canada, and Ireland (Graham et al. 2004). The disease is characterized by flat epidermal lesions that are bluish white in color and a granular texture mostly on the dorsal skin and fins (Figure 10.7). Blue spot is more prevalent on spawning age females (Yamamoto et al. 1984; Anonymous 2002). Lesions that are 3–10 mm in diameter and 0.25-mm thick contain hypertrophied cells that have enlarged nuclei and a granular cytoplasm. These cells lack the ribbon-like cytoplasm inclusions seen in lymphocystis virus disease, but the granular cytoplasm consists of a large number of darkly staining virusladen inclusions. Electron microscopy suggests that the causative agent is a herpesvirus designated esocid herpesvirus 1 (EHV-1) (Yamamoto et al. 1984). The virus has not been propagated in cell culture, and diagnosis is based on histopathology with electron microscopy for confirmation. Appearance of

lesions varies by season when water temperature is between 2 and 13◦ C in late winter–early spring (Margenau et al. 1995). Lesions disappear when water temperature rises above 14◦ C. The prevalence of EHV-1 in central Canada was 1–7% of northern pike populations, but prevalence in Wisconsin was as high as 34%. The importance of this disease is not known, but it has not been associated with fish kills.

Fish tumor viruses Fish tumors have received considerable attention because of a perceived relationship between the health of fish and the quality of the aquatic environment in which they live. However, fish tumors are not a recent occurrence but have been known and recognized for centuries (Anders and Yoshimizu 1994). Although fish skin tumors have a variety of etiologies, the cause of many is undetermined. According to Anders and Yoshimizu (1994), there are about 50 types of fish tumors or tumor-like

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Table 10.3 Comparison of virus-caused epidermal lesions on walleye and the prevalence of each type of disease on 25 fish examined from a single population in Canada.

Disease

Number Positive

Epidermal

10

Dermal sarcoma Diffuse epidermal hyperplasia Lymphocystis

7 4 12

Lesion Characteristic

Virus Family

Size

Isolated

Clear, raised, hyperplasia distinct margin Smooth, rounded Mucoid, indistinct flat, translucent White, red, large spherical cells, grape-like clusters

Retroviridae

120 nm

No

Retroviridae Herpesviridae

135 nm 100 nm

No Yes

Iridoviridae

260 nm

Yes

Source: Yamamoto et al. (1985a).

proliferations that range from benign epidermal papillomas to melanomas and carcinomas. About 50% of fish tumors affect the skin, and approximately half of these are in some way virus related. In some fish tumor diseases where virus is suspected, a viral agent is actually isolated in tissue culture, in others virus detection is only by electron micrographs of tumors, and still others merely remain suspected of having a viral etiology. Most tumor viruses are known from wild fishes, but several play important roles in aquaculture. Several of these diseases have already been discussed in greater detail: fish pox in common carp and Asagi carp (see Chapter 6), stomatopapilloma of eel (see Chapter 7), plasmacytoid leukemia, salmon swim bladder sarcoma, and salmonid herpesvirus type 2 (see Chapter 8).

Tumors in walleye Four different viruses of walleye that cause tumor-like lesions of the epidermis are discrete epidermal hyperplasia, diffuse epidermal hyperplasia (walleye herpesvirus), walleye dermal sarcoma (WDS), and lymphocystis (Table 10.3) (Yamamoto et al. 1985). Tumors in primarily wild walleye have received considerable attention, and a given population can be affected by more than one of these tumor-causing viruses. These diseases have not been associated with substantial losses,

so the primary impact is public concern over the presence of tumors on these important game fish. Discrete epidermal hyperplasia and WDS viruses are among the best characterized retrovirus-associated diseases in fish.

Diffuse epidermal hyperplasia In central Canada, a herpesvirus was isolated from diffuse epidermal hyperplasia lesions on the skin of spawning walleye (Kelly et al. 1980). After further characterization, the virus was named Herpesvirus vitreum or walleye herpesvirus (Kelly et al. 1983). The herpesvirus infection was named “diffuse epidermal hyperplasia” because of the flat, diffused appearance of the skin lesion (Yamamoto et al. 1985). Walleye herpesvirus causes epidermal lesions that resemble thick areas of slime (Figure 10.8). The growths are flat and translucent with soft, swollen underlying tissue (Yamamoto et al. 1985). These lesions measure up to several centimeters in diameter, spread laterally on the surface of the fish, and are not as pronounced as lymphocystis or dermal sarcoma. The transient lesions appear during spawning but have no apparent ill effects on the host. Diffuse epidermal hyperplasia (walleye herpesvirus) is diagnosed by lesion recognition, isolation of virus from lesions in walleye cell lines, and histological sectioning of diseased

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(a)

(b)

(c)

Figure 10.8 Skin virus diseases of walleyes. (a) Walleye dermal sarcoma (arrows), (b) diffuse epidermal hyperplasia (arrow), and (c) Lymphocystis disease. (Photos courtesy of P. R. Bowser.)

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tissue. Cells in diffuse epidermal hyperplasia lesions are disorganized with slightly enlarged nuclei that contain granular inclusions (Figure 10.9). Electron microscopy show typical herpesvirus particles within the nucleus and mature enveloped virions near the cell periphery. After walleye herpesvirus was initially isolated in cell cultures, it was characterized by Kelly et al. (1983). The virion is icosahedral, measuring 200 nm in diameter, presumably with an envelope, and has a DNA genome. It replicates in walleye ovary (WO) and walleye embryo (We-2) cells but not in CHSE-214, BB, FHM, or RTG-2 cells. Primary CPE consists of syncytia formation followed by cell lysis. Walleye herpesvirus replication occurs at 4–15◦ C in cell cultures; maximum replication occurs at 15◦ C, and no replication takes place at or above 20◦ C. At 15◦ C, the maximum titer in cell cultures is 105 PFU/0.1 mL after 10–13 days of incubation. In western Canada, diffuse epidermal hyperplasia is found in spawning walleye during early spring, but lesions disappear by late spring and early summer. Little is known about the effects of the virus or its associated lesions on infected fish.

Discrete epidermal hyperplasia Discrete epidermal hyperplasia (DEH) is synonymous with walleye epidermal hyperplasia (Wolf 1988). Walker (1969) was the first to suspect that the disease was caused by a virus, and 15 years later, Yamamoto et al. (1985) described C-type retrovirus particles in electron micrographs of the lesions. Discrete epidermal hyperplasia has been found in adult walleye in Saskatchewan and Manitoba, Canada, and Lake Oneida, New York, and throughout the Great Lakes region. Walleye is the only known naturally susceptible species. Discrete epidermal hyperplasia lesions are gently raised, clear, translucent, mucoid-like patches and are more discrete and less gran-

ular than lymphocystis or dermal sarcoma lesions (Yamamoto et al. 1985) (Figure 10.8). The more or less circular lesions appear singularly or in groups and can cover large areas of the body and/or fins. Lesions vary in diameter from a few millimeters to several centimeters and are 1–2 mm thick. Demarcation between normal epidermis and tumor is sharply defined as opposed to the more diffused, flatter nature of herpesvirus hyperplasia (diffuse epidermal hyperplasia). Discrete epidermal hyperplasia is confined to the epidermis. A presumptive field diagnosis is based on morphology of the lesions; however, confirmation should be made by histological procedures and/or electron microscopy. Histopathology indicates that DEH lesions contain predominately cuboidal cells with mucous cells at the surface (Figure 10.9). Lesions do not extend below the basal lamina (Yamamoto et al. 1985). Viral particles are randomly distributed in microvilli-like extensions of the cell. Virions are not found in the cytoplasm matrix, but occur at the cell periphery. The virus, which has not yet been isolated in tissue culture, is a C-type, irregularshaped retrovirus, measuring approximately 120 nm in diameter. Sequence analysis of a portion of the polymerase gene amplified from affected tissue indicates that there are two closely related retroviruses that cause this condition, and they have been designated walleye epidermal hyperplasia virus type 1 and type 2 (WEHV1 and WEHV2) (LaPierre et al. 1998). The genomes of these viruses have been sequenced; they are 13 kb in length, the genes encode, and they are arranged similar to WDS virus (LaPierre et al. 1999). Discrete epidermal hyperplasia appears to have little deleterious effect on infected walleye. Infection rates of 0–20% were noted in a survey of nine separate Canadian walleye populations (Yamamoto et al. 1985), but when the same populations were surveyed in spring

Figure 10.9 Histopathology of the four types of skin tumors on walleye. (a) Discrete epidermal hyperplasia (arrow), (b) walleye dermal sarcoma, (c) diffuse epidermal hyperplasia, and (d) lymphocystis. (Photos a, b, and c. courtesy of P. R. Bowser.)

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(spawning season) and autumn, epidermal hyperplasia was found only in spring. However, some reports indicate that the disease occurs throughout the year. Incidence of disease was reported to be about 5% in New York walleye populations. Getchell et al. (2004) studied the disease in 3- to 8-year-old walleye in Lake Oneida, New York. They found the incidence to be less than 20%, but the prevalence was less in 8-year-old fish, indicating that age of fish may affect its presence. In the Canadian survey by Yamamoto et al. (1985), some fish were infected with lymphocystis or dermal sarcoma in addition to discrete epidermal hyperplasia. Two types of epidermal hyperplasia were found on Canadian walleye; one was the discrete epidermal hyperplasia and the other was the superficial epidermal hyperplasia lesion containing retrovirus and a more diffuse growth (diffuse epidermal hyperplasia). Transmission of discrete hyperplasia may be by direct contact with infected fish, particularly on spawning grounds in the spring. Bowser et al. (1998) transmitted the disease to 97% of 1-year-old walleye by injection of cell-free filtrates prepared from lesions. Application of lesion filtrates to skin was also used to experimentally transmit the disease (Quackenbush et al. 2001).

Walleye dermal sarcoma According to Wolf (1988), Walleye dermal sarcoma (WDS), caused by walleye dermal sarcoma virus (WDSV), was first described by Walker (1947), who recognized that the lesions were different from classical lymphocystis. Dermal sarcoma lesions have been reported from adult walleye in Lake Oneida (New York), Lake Champlain (Vermont), and a number of other lakes and rivers in New York, the Great Lakes region, and in central Canada. These tumors have not been reported from cultured walleye. Experimental transmission studies using filtrates from WDSV tumors collected in the spring indicate that saugar and

yellow perch are also susceptible to WDSV (Holzschu et al. 1998; Bowser et al. 2001). WDS are spherical lesions that occur anywhere on the body and fins of adult walleye (Figure 10.8). The sarcomas are fine textured, variably firm, vascularized, and color varies from pink to white. Tumor size ranges from 1 to more than 10 mm with a mean diameter of approximately 5 mm (Yamamoto et al. 1976). Although field observations may indicate the presence of dermal sarcoma because of morphological differences from lymphocystis, the only definitive way to identify the virus is to section tumors and examine the tissue histology and detect the virus in electron micrographs. Martineau et al. (1991) homogenized 20 tumors, and after centrifugation, a C-type retrovirus was observed by electron microscopy in the sediment pellets. Further evidence that the virus is a member of Retroviridae was presented by Bowser and Wooster (1994) when they demonstrated the presence of a lipid envelope. Viral RNA from the WDSV was electrophoresed, and a resultant cDNA synthesized from viral RNA hybridized with viral DNA in walleye tumors. This suggested that WDS results from an infection caused by a unique exogenous retrovirus of which the large genome is predominantly unintegrated in the tumor cells. There is some disagreement about diameter of the retrovirus: Walker (1969) reported a diameter of l00 nm, Yamamoto et al. (1976) gave a slightly larger size of 135 nm, and Martineau et al. (1991) described a 90-nm retrovirus. These variances could be due to different methods of tissue fixation. The WDSV was the first fish retrovirus genome to be sequenced (Holzschu et al. 1995). The genome is large (12.7 Kb) and complex compared to most other characterized retroviruses, and it represents the type species of the epsilon retroviruses that includes WEHV1 and WEHV2. Recently a cell line, spring tumor explant cells (STECs), has been established, which is persistently infected with replicating WDSV

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(Rovnak et al. 2007). This in vitro virus production system promises to be helpful in intricate molecular studies, including the production of recombinant products. Martineau et al. (1990) transmitted the disease to healthy 4-month-old walleye by intramuscular injection with homogenized tumors from adult fish. Tumors developed in recipient fish 4 months later. In a subsequent study, transmission of walleye tumorproducing retrovirus was greater at 15◦ C than at 10 or 20◦ C (Bowser et al. 1990). Tumors developed in 8 weeks at 15◦ C. Bowser and Wooster (1991) showed that tumors regressed either totally, or partially, during an 18-week study period when water temperatures rose from 7 to 29◦ C, but total regression was observed only in females. Also Bowser et al. (1996) reported successful transmission of WDSV to 82% of young-of-the-year walleye by injecting them with infective material from regressing tumors obtained in the spring. Dermal sarcoma lesions developed in yellow perch 20 weeks postexposure by topical application of cell-free filtrates, made from lesion material, on the fish’s flank and held at 15◦ C (Bowser et al. 2001). In support of the theory that WDSV is transmitted in spring, tumor material taken during autumn failed to elicit tumors in injected fish (Bowser et al. 1996). The authors identified WDSV RNA in fish injected with spring infectious material, but virus RNA was absent in fish injected with extracts from fall tumors. Analysis of WDSV virus transcripts at developmental and regression stages demonstrated that production of the virus is tightly regulated with virus primarily being produced during the regression phase (Quackenbush et al. 1997). These data suggest that the transmission of WDSV occurs primarily during the spring and may be optimally efficient during direct contact between adults during spawning. In some walleye populations, frequency of neoplasia can be high (Table 10.3), and an individual fish may have more than one type of tumor. Yamamoto et al. (1985) examined 25

walleye and found that 12 had lymphocystis, 7 had dermal sarcoma, 4 had diffused epidermal hyperplasia (walleye herpesvirus), and 10 had discrete epidermal hyperplasia. Bowser et al. (1988) noted that, on a seasonal basis, up to 27% of adult walleye in North America are affected with dermal sarcoma. They also noted that the number of infected fish in Lake Oneida was higher in spring and fall than during the summer. Histopathology can be used to definitively identify different virus-induced walleye growths (Figure 10.9). Gross dermal sarcoma lesions may be confused with lymphocystis, but probably not with epidermal hyperplasia (retrovirus) or diffuse epidermal hyperplasia (walleye herpesvirus) (Table 10.3; Figure 10.9). However, lymphocystis and dermal sarcoma tumor cells may also occur within the same lesion. Dermal sarcoma lesions consist of irregularly shaped tumors containing normal size cells, which are often arranged in whorls (Figure 10.9) whereas lymphocystis cells are hypertrophied. These tumors were thought to be noninvasive, but Ernest-Koons et al. (1996) discovered that, following experimental exposure of 9-week-old walleye, some of the neoplasms did not remain strictly cutaneous, especially those collected 84 days postexposure and later. Lesions were locally invasive and replaced normal muscle tissue. In one case, a head tumor had invaded the skull causing a deformed brain. Using in situ hybridization and immunohistochemical procedures, Poulet et al. (1995) showed that dermal sarcomas were associated with elevated, transcriptionally active WDSV in neoplastic cells. The virus tropism extended beyond the mesenchymal, fibroblast-like neoplastic cells to include mononuclear inflammatory and epidermal cells. The appearance of dermal sarcoma-infected fish makes them unacceptable to fishermen and consumers. If walleye culture were to become a viable commercial food fish industry, these virus-induced tumors could attract more attention.

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Figure 10.10 Northern pike with massive Esox lymphosarcoma lesions on the body. (Photo courtesy of Tom Jones.)

Esox lymphosarcoma Esox lymphosarcoma, also known as esocid lymphosarcoma or Esox sarcoma, is an epidermal tumor disease of muskellunge, northern pike, and tiger muskellunge (hybrid Esox lucius × Esox masquinongy) that occurs primarily on the skin but in advanced cases may invade internal organs. Esox lymphosarcoma has been reported across Canada, as well as Sweden, Finland, Ireland, and northern United States including Minnesota and Wisconsin (Wingvist et al. 1968; Sonstegard 1976), Colorado (Bowser et al. 2002), and New York and Vermont (Anonymous 2004). Esocid lymphosarcoma is characterized by macroscopically visible nodular lesions in epidermal connective tissue (Figure 10.10) from which tumors may metastasize into the muscle and then in advanced cases into internal organs. Skin lesions measuring about 1–3 mm thick, 5–10 mm in diameter, and up to the “size of a small orange” develop anywhere on the body surface (Sonstegard 1976; Tom

Jones, Vermont Fish & Wildlife Department, personal communication). Epidermal lesions are smooth, convex, whitish gray, red or pink; lesions are translucent tissue consisting largely of randomly oriented normal-sized undifferentiated cells. Tumors are caused by a putative C-type retrovirus, known only by electron microscopy, measuring about 100–150 nm in diameter that buds from the cytoplasm of infected cells (Sonstegard 1976; Yamamoto et al. 1984). Esocid lymphosarcoma, described as a stem cell lymphoreticular neoplasm, appears to be temperature related occurring primarily in the winter to spring (Sonstegard 1976). Incidence of esocid lymphosarcoma in wild northern pike and muskellunge has been as high as 6–21% in some pike and muskellunge populations (Winqvist et al. 1968; Sonstegard 1976; McAllister 1979). Esocid lymphosarcoma can transfer from fish to fish by physical contact especially during spring spawning activities (Anonymous 2004). After identifying the disease in

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hatchery-reared yearling tiger muskellunge in Colorado, esocid lymphosarcoma was experimentally transmitted by exposing juvenile tiger muskellunge to the injection of filtered tumor homogenates, implantation of neoplastic tissue, cohabitation, and exposure to cellfree filtrates in water (Bowser et al. 2002). At 32-week postchallenge, grossly visible lesions, confirmed as esocid lymphosarcoma by histopathology, were observed on the challenged fish. It is believed that this is the first report of the disease in tiger muskellunge. The significance of Esox lymphosarcoma is not known to be great in terms of impact on susceptible fish populations other than from an aesthetic point of view (McAllister 1979). The disease is not known to be transmitted to humans, but it is recommended that infected fish should not be eaten. The disease has received attention because some fish tumors are morphologically similar to structures found in several types of human lymphoma and leukemia (Dawe et al. 1976).

Other tumor viruses of fish Most isolated viruses known to cause tumors are herpesviruses and have been discussed in previous chapters. Other fish tumors from which viruses have been isolated in cell culture but may not actually have caused the tumor are as follows: a picornavirus from European smelt (Ahne et al. 1990), rhabdovirus from European eel (Ahne et al. 1987), and birnavirus from North Sea dab (Olesen et al. 1988). In addition, several tumor-associated viruses of fish have not been isolated but are known only from electron microscopy or by experimental transmission by exposing naive fish to cell-free filtrates from the tumors. In Japan, several groups of larval Japanese flounder (10–30 days old) contracted a tumor disease that affect the fins and body surface, which were opaque due to the presence of proliferated epithelial cells (Iida et al. 1989). Mortality reached 80–90% in a few weeks at

water temperatures of about 20◦ C. Numerous hexagonal virus-like particles were seen in the nucleus and cytoplasm of the tumor cells in electron micrographs. The disease was successfully transmitted to naive fish via exposure to 0.45-µm filtrates, but isolation attempts in 33 fish cell lines were unsuccessful. Sensitivity of the agent to ether and pH 3, in conjunction with its physical structure, indicates that the virus was a herpesvirus. Other experimentally induced skin tumors of fish associated with viruses are as follows: squamous cell carcinoma of rudd (Hanjavanit and Mulcahy 1989) and a neurofibromatosislike disease in damselfish (Schmale and Hensley 1988). Anders and Yoshimizu (1994) stated as follows: “It is notable that successful experimental induction of benign tumors was mostly associated with herpesviruses, whereas, formation of malignant forms seems to be restricted to the involvement of retroviruses of the subfamily Oncovirinae.” Viruses detected only by electron microscopy of infected tissues have been associated with a large number of fish tumors. According to Anders and Yoshimizu (1994), these include members of the herpesvirus, adenovirus, and retrovirus groups, but some suspected virus particles are questionable. Most fish herpesvirus diseases have been demonstrated by isolation and experimental transmission, but several putative herpesviruses are known only by electron microscopy. Adenovirus-like particles have been associated with epidermal hyperplasia and papillomas in Baltic cod (Jensen and Bloch 1980) and North Sea dab (Block et al. 1986). According to Anders and Yoshimizu (1994), 17 different retrovirus-like particles have been found in fish, 10 of which are associated with skin tumors. Some of these have been isolated in cell culture (Frerichs et al. 1991), while others have not. In addition, reverse transcriptase activity in tissue extracts indicated the presence of retrovirus in skin tumors in northern pike (Papas et al. 1976) and stone roller (Sonstegard 1977).

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Factors that affect virus-induced tumors in fish A number of factors affect development of fish tumors (Anders and Yoshimizu 1994). The season of the year affects prevalence of tumors with highest incidence occurring during spawning season, smoltification, gonadal development, or metamorphosis. During these periods, usually in spring or early summer, endocrine systems and often immune systems are in states of flux, and most likely this instability plays a role in the activation of latent viruses to produce tumors. Interaction between skin, virus, and biological and environmental factors play a major role in epidermal tumor development. This is mediated by immune suppression and increased endocrine activity, which are perceived to be influenced by environmental quality. Environmental parameters that can depress defense mechanisms are fluctuations in water temperature and salinity, high population density, condition factors, high concentrations of certain pollutants, and carcinogens. With regard to fish tumor viruses, the above factors could favor virus stimulation and proliferation of epidermal and fibroblastic cells to form tumors (Anders and Yoshimizu 1994). These authors also noted that epidermal fish tumors are induced by herpesviruses, adenoviruses, papovaviruses, and retroviruses all of which are responsible for skin tumors in humans.

Management of other fish viruses Control and management of lymphocystis is difficult. When encountered in aquaculture, avoidance of known carrier populations is recommended. When ornamental fish are involved, removing affected fish before infected cells rupture prevents release of virus. Also, replacement of 50% of the water in aquaria every 2–3 days will dilute virus and possibly reduce infection incidence. Proper sanitation and disinfection of equipment and holding contain-

ers are essential to prevent transmission of the virus to noninfected fish. In the case of VEN, because fry and youngof-the-year fish seem to be more susceptible, they should be reared in water that is free of infected fish. Adverse environmental conditions, such as low dissolved oxygen and other stressors, should be avoided, especially during disease outbreaks. A management scheme to deal with EHNV by avoidance and possible disease-free inspections was proposed by Whittington and Hyatt (1997). They indicated that EHNV cannot be eradicated from farmed rainbow trout or in water supplies from reservoirs in Australia because of carrier wild redfin perch. However, in an effort to restrict EHNV to those areas of Australia where it is endemic, stringent guidelines for production farms have been established. Infected farms may sell only killed fish or fillets, and export of live fish from these zones is prohibited. Government-owned farms practice SPF certification for EHNV prior to distribution of fish or eggs. Additionally, in the event that new infection sites are identified that have fish-free closed water systems, eradication of infected fish is recommended. Equipment and culture units should be completely disinfected with 200 mg/L of sodium hypochlorite, dried, and organic matter removed (Langdon 1989). Facilities should be repopulated only with stocks certified to be EHNV free. Control of skip jack VNN may vary with geographical region, but absence of stress on susceptible fish is uniformly required (Nakai et al. 1995). The PCR method for detecting SJNNV-positive spawning striped jack combined with hatchery hygiene and husbandry procedures recommended by Nishizawa et al. (1994) are helpful in preventing and controlling the virus in a particular species. A similar approach to that used for striped jack may be applied to European sea bass operations in the Mediterranean region, but Le Breton et al. (1997) suggested that this procedure may not always be effective because of widespread distribution of the pathogen in the region. They

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also suggested that development of a vaccine may be required to combat the disease. PFRD has been effectively managed by preventing spread of the pathogen and by disinfecting eggs. In The Netherlands, management practices initiated by Bootsma et al. (1975) reduced PFR impact on northern pike. A solution of 25 mg/L of iodine in the form of Wescodyne inactivated 99.99% of PFR in 30 sec. Routinely bathing pike eggs in the iodine solution following fertilization interrupts the PFR infection cycle. No particular management practices apply to fish tumor viruses other than avoidance of known infected populations. Do not use fish from affected waters for brood stock under any circumstances and disinfect all equipment that come in contact with fish bearing tumors. Also, when fish with tumors are found in culture units, the fish should be removed and destroyed immediately. Vaccines have proven useful against several of these miscellaneous virus diseases. In addition to the vaccine mentioned earlier for Red Sea brim iridovirus (Nakajima et al. 1999), Husgard et al. (2001) evaluated the immune ´ response to an injectable recombinant capsid protein vaccine for skip jack nervous necrosis. In an experimental trial, significant protection was observed when vaccinates were challenged 10 weeks postvaccination. Also a patent was applied for in the United States for an inactivated vaccine against VNN (Dos Santos et al. 2009). The further development and implementation of vaccines are highly encouraging because of their positive results; vaccines, not only increase survival of fish but in many cases increase growth efficiency.

References Adair, B. M., and M. McLoughlin. 1986. Isolation of pike fry rhabdovirus from brown trout (Salmo trutta). Bulletin of the European Association of Fish Pathologists 6(3):85. Ahne, W. 1975a. A rhabdovirus isolated from grass carp (Ctenopharyngodon idella Val). Archives of Virology 48:181–185.

Ahne, W. 1975b. Biological properties of a virus isolated from grass carp (Ctenopharyngodon idella Val.). In: L. A. Page (ed.) Wildlife Diseases. New York, Plenum Press, pp. 135–140. Ahne, W. 1985. Viral infection cycles in pike (Esox lucius L). Journal of Applied Ichthyology 1:90–91. Ahne, W. 1986. Unterschiedliche biologische eigenschaften 4 cyprinidenpathogener rhabdovirusisolate. Journal of Veterinary Medicine 33:253–259. Ahne, W., K. Anders, et al. 1990. Isolation of picornavirus-like particles from the European smelt (Osmerus eperlanus). Journal of Fish Diseases 13:167–168. Ahne, W., S. Blake, et al. 2003. Characterization of aquabirnaviruses from flounder Pseudopleuronectes americanus and mummichog Fundulus heterclitus in the Chesapeake Bay, Virginia, USA. Diseases of Aquatic Organisms 56:201–206. Ahne, W., M. Bremont, et al. 1997. Iridoviruses associated with epizootic haematopoietic necrosis (EHN) in aquaculture. World Journal of Microbiology and Biotechnology 13:367–373. Ahne, W., G. Kurath, and J. R. Winton. 1998. A ribonuclease protection assay can distinguish SVCV from PFR. Bulletin of the European Association of Fish Pathologists 18:220–224. Ahne, W., H. Mahnel, and P. Steinhagen. 1982. Isolation of pike fry rhabdovirus from tench, Tinca tinca L., and white bream, Blicca bioerkna (L.). Journal of Fish Diseases 5:535–537. Ahne, W., I. Schwanz-Pfitzner, and I. Thomsen. 1987. Serological identification of 9 viral isolates from European eels (Anguilla anguilla) with stomatopapilloma by means of neutralization tests. Journal of Applied Ichthyology 3:30–32. Ahne, W., and I. Thomsen. 1986. Isolation of pike fry rhabdovirus from Pseudorasbora parva (Temminck & Schlegel). Journal of Fish Diseases 9:555–556. Alpers, C. E., B. B. McCain, et al. 1977. Lymphocystis disease in yellowfin sole (Limanda aspera) in the Bering Sea. Journal of the Fisheries Research Board of Canada 34:611–616. Amin, O. M. 1979. Lymphocystis disease in Wisconsin fishes. Journal of Fish Diseases 2:207–217. Anders, K. 1989. Lymphocystis disease of fishes. In: W. Ahne, and E. Kurstak (eds) Viruses of Lower Vertebrates. Berlin, Springer-Verlag, pp. 141–160.

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virus (SJNNV) in adult striped jack. Diseases of Aquatic Organisms 28:87–91. Nguyen, H. D., T. Nakai, and K. Muroga. 1996. Progression of striped jack nervous necrosis virus (SJNNV) infection in naturally and experimentally injected striped jack (Pseudocaranx dentex) larvae. Diseases of Aquatic Organisms 24:99–105. Nigrelli, R. F. 1950. Lymphocystis disease and ergasilid parasites in fishes. Journal of Parasitology 36:36. Nigrelli, R. F., and G. D. Ruggieri. 1965. Studies on virus diseases of fishes. Spontaneous and experimentally induced cellular hypertrophy (lymphocystis disease) in fishes of the New York Aquarium with a report of new cases and an annotated bibliography (1874–1965). Zoologica 50:83–96. Nishizawa, T., M. Furuhashi, et al. 1997. Genomic classification of fish nodaviruses by molecular phylogenetic analysis of the coat protein gene. Applied and Environmental Microbiology 63:1633–1636. Nishizawa, T., M. Kise, et al. 1995. Neutralizing monoclonal antibodies to striped jack nervous necrosis virus (SJNNV). Fish Pathology 30:111–114. Nishizawa, T., K. Mori, et al. 1994. Polymerase chain reaction (PCR) amplification of RNA of striped jack nervous necrosis virus (SJNNV). Diseases of Aquatic Organisms 18:103–107. Nishizawa, T., K. Muroga, and M. Arimoto. 1996. Failure of the polymerase chain reaction (PCR) method to detect striped jack nervous necrosis virus (SJNNV) in striped jack (Pseudocaranx dentex) selected as spawners. Journal of Aquatic Animal Health 8:332–334. OIE (2009). Red sea bream iridoviral disease. OIE Diagnostic Manual for Aquatic Animal Diseases, Sixth Edition. Paris, World Organization for Animal Health (Chapter 2.3.7.). Olesen, N. J., P. E. V. Jørgensen, et al. 1988. Isolation of an IPN-like virus belonging to the serogroup II of the aquatic birnaviruses from dab, Limanda limanda L. Journal of Fish Diseases 11:449–451. Overstreet, R. M. 1988. Aquatic pollution, Southeastern U.S. coasts: histopathologic indicators. Aquatic Toxicology 11:213. Overstreet, R. M., and H. D. Howse. 1977. Some parasites and diseases of estuarine fishes in pol-

luted habitats of Mississippi. Annals of the New York Academy of Science 298:427–462. Papas, T. S., J. E. Dahlberg, and R. A. Sonstegard. 1976. Type C virus in lymphosarcoma in northern pike (Esox lucius). Nature 261:506–508. Peters, N., and W. Schmidt. 1995. Formation and disintegration of virions in lymphocystis cells of plaice Pleuronectes platessa. Diseases of Aquatic Organisms 21:109–113. Petty. L. L., and J. J. Magnuson. 1974. Lymphocystis in age 0 bluegills (Lepomis macrochirus) relative to heated effluent in Lake Monona, Wisconsin. Journal of the Fisheries Research Board of Canada 31:189–193. Piaskoski, T. O. 1997. Characterization of the largemouth bass virus in cell culture and protein profile comparison to frog virus-3 and lymphocystis disease virus. M.S. Thesis, Auburn University, Auburn, Alabama. Pinto, R. M., P. Alvarez-Pellitero, et al. 1989. Occurrence of viral erythrocytic infection in the Mediterranean sea bass, Dicentrarchus labrax (L.). Journal of Fish Diseases 12:185–191. Plumb, J. A., J. M. Grizzle, et al. 1996. An iridovirus isolated from wild largemouth bass. Journal of Aquatic Animal Health 8:265–270. Poulet, F. M., V. M. Vogt, et al. 1995. In situ hybridization and immunohistochemical study of walleye dermal sarcoma virus (WDSV) nucleic acids and proteins in spontaneous sarcomas of adult walleyes (Stizostedion vitreum). Veterinary Pathology 32:162–172. Quackenbush, S. L., D. L. Holzschu, et al. 1997. Transcriptional analysis of walleye dermal sarcoma virus (WDSV). Virology 237:107– 112. Quackenbush, S. L., J. Rovnak, et al. 2001. Genetic relationship of tumor-associated piscine retroviruses. Marine Biotechnology 3:S88–S99. Reddacliff, L. A., and R. J. Whittington. 1996. Pathology of epizootic haematopoietic necrosis virus (EHNV) infection in rainbow trout (Oncorhynchus mykiss Walbaum) and redfin perch (Perca fluviatilis L). Journal of Comparative Pathology 115:103–115. Reiersen, L. O., and K. Fugelli. 1984. Annual variation in lymphocystis infection frequency in flounder, Platichthys flesus (L.). Journal of Fish Biology 24:187–191. Reno, P. W., and B. L. Nicholson. 1980. Viral erythrocytic necrosis (VEN in Atlantic cod (Gadus

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morhua): in vitro studies. Canadian Journal of Fisheries and Aquatic Sciences 37:2276–2281. Reno, P. W., D. V. Serreze, et al. 1985. Hematological and physiological effects of viral erythrocytic necrosis (VEN) in Atlantic cod and herring. Fish Pathology 20:353–360. Robin, J., and L. Bertholimue. 1981. Purification of lymphocystis disease virus (LDV) grown in tissue culture, evidences for the presence of two types of viral particles. Reviews in Canadian Biology 40:323–329. Rohovec, J. S., and A. Amandi. 1981. Incidence of viral erythrocytic necrosis among hatchery reared salmonids of Oregon. Fish Pathology 15:135–141. Rovnak, J., R. N. Casey, et al. 2007. Establishment of productively infected walleye dermal sarcoma explant cells. Journal of General Virology 88:2583–2589. Rowley, H., D. A. Graham, et al. 2001. Isolation and characterization of rhabdovirus from wild common bream Abramis brama, roach Rutilus rutilus, farmed brown trout Salmo trutta and rainbow trout Oncorhynchus mykiss in Northern Ireland. Diseases of Aquatic Organisms 48:7–15. Ryder, R. A. 1961. Lymphocystis as a mortality factor in a walleye population. The Progressive Fish-Culturist 23:183–186. Schmale, M. C., and G. T. Hensley. 1988. Transmissibility of a neurofibromatosis-like disease in bicolor damselfish. Cancer Research 48:3828–3833. Smail, D. A. 1982. Viral erythrocytic necrosis in fish: a review. Proceedings of the Royal Society of Edinburgh (B) 81:169–175. Smail, D. A., and S. I. Egglestone. 1980. Virus infections of marine fish erythrocytes: prevalence of piscine erythrocytic necrosis in cod Gadus morhua L. and blenny Blennius pholis L. in coastal and off shore waters of the United Kingdom. Journal of Fish Diseases 3:41–46. Song, J. Y., S. Kitamura, et al. 2008. Genetic variation and geographic distribution of megalocytiviruses. Journal of Microbiology 46:29–33. Sonstegard, R. A. 1976. Studies of the etiology and epizootiology of lymphosarcoma in Esox lucius L. and Esox masquinongy). Progress in Experimental Tumor Research 20:141–155. Sonstegard, R. A. 1977. Environmental carcinogenesis studies in fishes of the Great Lakes of North

America. Annals New York Academy of Science 298:261–269. Steiner, K. A., R. J. Whittington, et al. 1991. Purification of epizootic haematopoietic necrosis virus and its detection using ELISA. Journal of Virological Methods 33:199–209. Stone, D. M., W. Ahne, et al. 2003. Nucleotide sequence analysis of the glycoprotein gene of putative spring viraemia of carp virus and pike fry rhabdovirus isolates reveals four genogroups. Diseases of Aquatic Organisms 53:203–210. Thiery, R., J. Cozien, et al. 2004. Genomic classification of new betanodavirus isolates by phylogenetic analysis of the coat protein gene suggests a low host-fish species specificity. Journal of General Virology 85:3079–3087. Tidona, C. A., and G. Darai. 1997. The complete DNA sequence of lymphocystis disease virus. Virology 230:207–216. Ucko, M., A. Colorni, and A. Diamant. 2004. Nodavirus infection in Israeli mariculture. Journal of Fish Diseases 27:459–469. Walker, R. 1947. Lymphocystis disease and neoplasia in fish. Analytical Record 99:559–560. Walker, R. 1969. Virus associated with epidermal hyperplasia in fish. National Cancer Institute Monographs 31:195–207. Walker, R. 1971. PEN, a viral lesion of erythrocytes. American Zoologist 11:707. Walker, R., and S. W. Sherburne. 1977. Piscine erythrocytic necrosis virus in Atlantic cod, Gadus morhua, and other fish: ultrastructure and distribution. Journal of the Fisheries Research Board of Canada 34:1188–1195. Wang, Y. Q., L. Lu, et al. 2007. Molecular epidemiology and phylogenetic analysis of a marine fish infectious spleen and kidney necrosis virus-like (ISKNV-like) virus. Archives of Virology 152:763–773. Weissenberg, R. 1965. Fifty years of research on the lymphocystis virus disease of fishes (1914–1964). Annals New York Academy of Science 126:362–374. Whittington, R. J., M. Crockford, et al. 2008. Herpesvirus that caused epizootic mortality in 1995 and 1998 in pilchard, Sardinops sagax neopilchardus (Steindachner), in Australia is now endemic. Journal of Fish Diseases 31:97–105. Whittington, R. J., and A. D. Hyatt. 1996. Contingency planning for control of epizootic

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haematopoietic necrosis disease. Singapore Veterinary Journal 20:79–87. Whittington, R. J., and A. D. Hyatt. 1997. Diagnosis and prevention of epizootic haematopoietic necrosis virus (EHNV) infection. In: New Approaches to Viral Diseases of Aquatic Animals. Hahsei, Miem, Japan, National Research Institute of Aquaculture, pp. 80–93. Whittington, R. J., J. B. Jones, et al. 1997. Epizootic mortality in the pilchard Sardinops sagax neopilchardus in Australia and New Zealand in 1995. I. Pathology and epizootiology. Diseases of Aquatic Organisms 28:1–16. Whittington, R. J., C. Kearns, et al. 1996. Spread of epizootic haematopoietic necrosis virus (EHNV) in redfin perch (Perca fluviatilis) in southern Australia. Australian Veterinary Journal 73(3):112–114. Whittington, R. J., A. Philbey, et al. 1994. Epidemiology of epizootic haematopietic necrosis virus (EHNV) infection in farmed rainbow trout, Oncorhynchus mykiss (Walbaum): findings based on virus isolation, antigen capture ELISA and serology. Journal of Fish Diseases 17:205– 218. Whittington, R. J., and G. L. Reddacliff. 1995. Influence of environmental temperature on experimental infection of redfin perch (Perca fluviatilis) and rainbow trout (Oncorhynchus mykiss) with epizootic haematopoietic necrosis virus, an Australian iridovirus. Australian Veterinary Journal 72:421–424. Whittington, R. J., L. A. Reddacliff, et al. 1999. Further observations on the epidemiology and spread of epizootic haematopoietic necrosis virus (EHNV) in farmed rainbow trout Oncorhynchus mykiss in southeastern Australia and a recommended sampling strategy for surveillance. Diseases of Aquatic Organisms 35:125–130. Williams, E. H., Jr., J. M. Grizzle, and L. BunkleyWilliams. 1996. Lymphocystis in Indian glassfish Chanda ranga imported from Thailand to Puerto Rico. Journal of Aquatic Animal Health 8:173–175.

Williams, T. 1996. The iridoviruses. Advances in Virus Research 46:345–412. Winqvist, G., Ljungberg, and B. Hellstroem. 1968. Skin tumors of northern pike (Esox lucius L.). II. Viral particles in epidermal proliferations. Bulletin of the Office of International Epizooties 69:1023–1031. Wolf, K. 1962. Experimental propagation of lymphocystis disease of fishes. Virology 18:249–256. Wolf, K. 1988. Fish Viruses and Fish Viral Diseases. Ithaca, New York, Cornell University Press. Wolf, K., and C. P. Carlson. 1965. Multiplication of lymphocystis virus in the bluegill (Lepomis macrochirus). Annals of the New York Academy of Science 126:414–419. Wolf, K., M. Gravell, and R. G. Malsberger. 1966. Lymphocystis virus: isolation and propagation in centrarchid fish cell lines. Science 151:1004–1005. Woodland, J. E., C. J. Brunner, et al. 2002. Experimental oral transmission of largemouth bass virus. Journal of Fish Diseases. 25:669–672. Yamamoto, T., R. K. Kelly, and O. Nielsen. 1984. Epidermal hyperplasias of northern pike (Esox lucius) associated with herpesvirus and C-type particles. Archives of Virology 79:255–72. Yamamoto, T., R. K. Kelly, and O. Nielsen. 1985. Morphological differentiation of virus associated skin tumors of walleye (Stizostedion vitreum vitreum). Fish Pathology 20:361–372. Yamamoto, T., R. D. MacDonald, et al. 1976. Viruses associated with lymphocystis disease and dermal sarcoma of walleye (Stizostedion vitreum vitreum). Journal of the Fisheries Research Board of Canada 33:2408–2419. Zhang, O-Y., H-M. Ruan, et al. 2003. Infection and propagation of lymphocystis virus isolated from the cultured flounder Paralichthys olivaceus in grass carp cell lines. Diseases of Aquatic Organisms 57:27–34. Zhang, Q. Y., F. Xiao, et al. 2004. Complete genome sequence of lymphocystis disease virus isolated from China. Journal of Virology 78:6982–6994.

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Part III

Bacterial Diseases

Numerous bacterial species inhabit the aquatic environment, many of which are essential to the balance of nature with no direct consequence in fish disease. However, some 60 to 70 bacterial species are capable of causing disease in aquatic animals, several of these are also infectious to humans. The aquatic environment, especially aquacultural and eutrophic waters and rivers, provides a natural habitat for growth and proliferation of bacteria because of nutrient-rich organic substrates that enhance bacterial growth. Bacterial flora of water is influenced by nutrient availability, pH, temperature, and other factors that affect their growth pattern, virulence, and pathogenicity. To grow, bacteria need an organic substrate that provide nutrients; some survive as free living organisms or serve as fish pathogens (facultative), while others are fastidious and survive indefinitely only within a host (obligate pathogens). Also, the level of salinity in water or culture media may affect growth and survival of some bacteria. Generally, an optimum pH 6 to 8 is desirable for growth of most bacteria; many die at pH above 11 or below 5. A temperature of 20–42◦ C is necessary for optimum growth of most bacteria but some will grow above 50◦ C (thermophiles), while others grow at near 0◦ C (psychrophiles). Bacteria that are most pro-

lific at temperatures of 18–45◦ C (mesophils) include many bacteria that are pathogenic to fish. Three basic bacterial cell morphologies are spherical (coccus), rod (bacillus), and spiral shaped (spirillum). Bacteria have a particular staining characteristic (Gram’s) referred to as gram positive (blue) or gram negative (red or pink). Gram-positive organisms may also be acid fast, which relates to the presence or absence of mycotic acid in the cell wall. Most bacteria responsible for fish disease are gramnegative rods but some pathogens that are gram-positive rods or cocci and a few that are acid-fast rods also cause disease in aquatic animals. In terms of pathology, there are basically two types of disease producing bacteria infectious to fish: (1) Obligate pathogens and (2) nonobligate or facultative pathogens. Although there are few true obligate pathogenic bacteria that cause fish diseases, Renibacterium salmoninarum, the etiological agent of bacterial kidney disease, and Mycobacterium spp. are examples rarely found in the absence of a host. However, facultative bacteria survive indefinitely in natural waters, and when environmental conditions are conducive, infectious fish diseases may result. Aeromonas hydrophila, a primary bacterial species involved

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in the motile Aeromonas septicemia complex, is an example of facultative bacteria. Fish bacterial infections can occur as a bacteremia, which implies the presence of bacterial organisms in the blood stream without clinical infection, or as a septicemia, which indicates that bacteria and toxins are actually present in the circulatory system and usually precipitate disease and clinical signs. Inflammation, hemorrhage, and necrosis are clinical signs associated with septicemia. Pathogenic bacteria can cause disease-producing exotoxins, which is generally characteristic, but not exclusively, of gram-positive organisms.

Pathogenic gram-negative bacteria produce either exotoxins or endotoxins, which consist of proteolytic enzymes that kill host cells and cause necrosis or make blood vessels more porous causing hemorrhage. Bacterial diseases are categorized according to fish species they infect, staining reaction of the cell, or by bacterial family, genus, and species (taxonomic grouping). In this book, bacterial diseases are organized according to family of cultured fishes that they most severely affect; therefore, it should not be concluded that the organism or disease discussed is limited to a particular fish group.

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Chapter 11

Catfish bacterial diseases

Channel catfish may be the most extensively cultured fish in the United States. Three major bacterial infections that affect these fish in the culture environment are columnaris (Flavobacterium columnare), enteric septicemia of catfish (ESC) (Edwardsiella ictaluri), and motile Aeromonas septicemia (MAS) (Aeromonas hydrophila and related motile aeromonads). Occasional other bacterial diseases of catfish are Edwardsiellosis (Edwardsiella tarda), pseudomonas septicemia (Pseudomonas fluorescens), and other incidental bacterial infections.

Columnaris Columnaris, occasionally referred to as cotton wool or mouth fungus, is an acute to chronic infectious skin disease of many fish species, but especially in channel catfish. The disease was first described by Davis (1922), but it was not until 22 years later that the organism was isolated, characterized, and named Bacterium columnaris (Ordal and Rucker 1944). The causative agent of columnaris has gone 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.

through several reclassifications and name changes since its original specific epitaph. Cytophaga columnaris, Flexibacter columnaris, and again Cytophaga columnaris. However, Bernardet and Grimont (1989) presented DNA relatedness and phenotypic characterization to justify retaining the name Flexibacter columnaris. Bernardet et al. (1996) redescribed this group of bacteria and renamed the etiology of columnaris in freshwater fish Flavobacterium columnare, which is used throughout this text. The genus Flavobacterium, yellow pigmented bacteria, contains two additional fish pathogens that affect salmonids (see Chapter 14): Flavobacterium psychrophila [Flexibacter (cytophaga) psychrophilum] (Borg 1960), etiological agent of cold-water disease, and Flavobacterium branchiophilum, etiological agent of bacterial gill disease.

Geographical range and species susceptibility Columnaris exists worldwide in freshwater habitats with channel catfish and other ictalurids being severely affected (Meyer 1970). No wild or cultured freshwater fish, including ornamental fish in aquaria, are totally resistant to columnaris. Cultured eels (fresh and brackish water) are highly susceptible to 275

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F. columnare (Wakabayashi et al. 1970) as are salmonids, particularly hatchery-reared trout and migrating adult salmon; common carp in Europe (Bernardet 1989); and cultured centrarchids, golden shiners, fathead minnows, and gold fish in the United States and tilapia wherever cultured.

Clinical signs Clinical signs of columnaris are easily recognized and differ little between species; however, lesion location will vary from outbreak to outbreak. Disease severity, type, and location of lesions, and pathogen virulence may correspond to the particular strain of F. columnare responsible for infection. Clinical signs of columnaris are nearly pathognomonic for the disease, but diagnosis can be complicated by simultaneous viral, other bacterial pathogens, and/or parasitic infections. Columnaris generally begins as an external infection on fins, body surface, or gills. The fins become frayed (necrotic) with grayish to white margins; initial skin lesions appear as discrete bluish gray areas, which evolve into depigmented necrotic lesions, causing fish to lose their metallic sheen (Figure 13.1). Skin lesions have yellowish (mucoid material) or pale margins accompanied by mild inflammation. These same lesion types occur on eel, trout, cyprinids, centrarchids, tilapia, and other fish groups infected with columnaris. Gill lesions appear as white to brown necrotic areas. The color is dependent upon the presence of debris and/or secondary fungus in the lesion (Figure 11.1). Lesions can develop exclusively on the gills, which usually results in subacute disease and mortality. In some instances, columnaris becomes systemic with little or no pathological change occurring in the visceral organs. Whether bacteria isolated from internal organs of systemically infected fish was taxonomically F. columnare is not clear, but bacteria can be isolated from kidneys of more than 50%

of necropsied catfish infected with epidermal columnaris (Hawke and Thune 1992).

Diagnosis Columnaris is normally diagnosed by recognition of typical lesions on the body, fins, and gills of diseased fish and presence of long, slender rods in wet mounts made from suspect lesions (Figure 11.2). These nonflagellated bacteria display gliding motility and form “haystacks” or columns in wet mounts prepared from gill tissue, which is basically confirmatory of the disease. A moist medium with a low nutrient and agar content such as cytophaga media (Ordal’s) (Anacker and Ordal 1959) and Hsu–Shotts (Shotts 1991) is generally used for isolation of F. columnare. Ordal’s and Hsu–Shotts media can also be prepared as broth by eliminating the agar in which F. columnare forms a distinct yellow, mucoid pellicle at the meniscus. Hawke and Thune (1992) found that a modification of media described by Fijan (1969) worked best for isolation of F. columnare, especially in the presence of other bacteria. This medium, selective cytophaga agar (SCA), is prepared by supplementing Ordal’s media with 5 µg of neomycin/mL and 200 IU polymyxin B/mL. Song et al. (1988a), however, found that medium of Shieh (1980) was superior to any other. The organism grows poorly or not at all on conventional highnutrient media. Immunofluorescent test, using conjugated anti-F. columnare sera, can also be used to rapidly detect and identify the pathogen in culture (Panangala et al. 2006a). Welker et al. (2005) described a rapid PCR detection method of F. columnare from channel catfish. The system is based on the 16S-23S rDNA intergenic spacer region of the ribosomal RNA operon. The pathogen was detected by PCR in tank water and catfish tissue samples with a higher frequency and in less time than standard microbiological methods.

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(a)

(b)

Figure 11.1 Columnaris lesion. (a) Columnaris lesion on caudal peduncle (arrow) and frayed caudal fin, and (b) lesion on gill (arrow) of channel catfish.

Bacterial characteristics When a columnaris isolate is incubated at 25–30◦ C for 48 hours, growth of most isolates will appear as spreading, rhizoid, discrete colonies with yellow centers that adhere tightly to the media. In comparing 27 isolates of F. columnare from channel catfish in the southeastern United States, Davidson (1996) found colonies with subtly different morphologies, which were also described by Shamsudin and Plumb (1996). When grown on Hsu–Shotts media, these isolates produced three colony morphologies: (1) bright yellow,

dry, umbonate and spreading with irregular edges (most colonies were of this type); (2) bright yellow, moist, and spreading with uneven edges’ and (3) pale yellow, dry, and flat, with uneven edges and spreading was noted more than in other colony types. Eleven of 27 isolates also grew weakly in the presence of 1% NaCl. An additional 11 isolates studied by Shamsudin and Plumb (1996) from blue catfish, channel catfish, largemouth bass, and fathead minnows had no morphological or physiological differences relative to host. All isolates were confirmed as F. columnare based on colony and cellular morphology, as well

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(a)

(b)

Figure 11.2 Wet mounts of Flavobacterium columnare from channel catfish showing (a) long slender, flexing rods, and (b) the haystacking (arrows) typical of virulent F. columnare.

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as biochemical and physiological characterization, most of which were uniform and similar to the ATCC No. 49512 isolate. Isolation of F. columnare is enhanced by addition of polymyxin B and neomycin to the media because they inhibit growth of noncolumnaris organisms (Fijan 1969). Columnaris is then confirmed by slide agglutination using specific F. columnare antisera. According to Bernardet et al. (1996), the emended genus Flavobacterium is a Gramnegative rod that measure 2–10 µm long and about 0.5 µm in diameter, motile by gliding, produce yellow colonies on agar, is a chemoorganotroph and facultative anaerobe, and decomposes several polysaccharides but not cellulose. These organisms are widely distributed in soil and freshwater habitats. The G + C contents of Flavobacterium DNAs range from 32 to 37 mol%. Major distinguishing factors between F. columnare, F. branchiophilum, and F. psychrophila are acid production from glucose, H2 S production, catalase, optimum growth temperature, salinity tolerance, and the presence of chondroitinase (Table 11.1). Griffin (1992) devised a simplified method of identifying F. columnare using five characteristics that distinguish it from other yellow pigment producing aquatic bacteria: (1) ability to grow in the presence of neomycin sulfate and Polymyxin B, (2) typical thin, rhizoid, yellowish colonies, (3) ability to degrade gelatin, (4) bind congo red, and (5) production of chondroitin lyase. Using DNA homology, Pyle and Shotts (1981) suggested that strains of columnaris from salmonids and those from warmwater fish were indeed different, and that three distinct groups existed within the cold-water isolates. Song et al. (1988b) found three distinct groups among 26 F. columnare isolates from Canada, Chile, Japan, Korea, Republic of China, and the United States based on DNA homology, although there was diversity in colony morphology and some biochemical characteristics not necessarily related to DNA

homology. Twenty isolates in one homologous group included representatives from each country. Analysis by SDS-PAGE of outer membrane profiles (OMP) of 27 channel catfish F. columnare isolates indicated four distinct groups based on OMP molecular mass, which ranged between 40 and 60 kDa (Davidson 1996). If all isolates examined in these studies were in fact F. columnare, different strains of the pathogen apparently exist within the species. Furthermore, Zhang et al. (2006) clearly differentiated pathogenic F. columnare from an attenuated strain used as a vaccine using LPS protein profiles. To help clarify differentiation of F. columnare strains, 31 isolates of F. columnare from fish and 3 from American Type Culture Collection were analyzed by the gas chromatography microbial identification system (MIS; Microbial ID, Newark, Delaware) and compared whole-cell fatty acid profiles (Shoemaker et al. 2005). Some differences were noted between the two methods with minor differences within each procedure; however, if biochemical results are known, either method can be used to rapidly identify F. columnare and distinguish them from other yellowpigmented bacteria. The authors concluded that the fatty acid profiles of F. columnare are rapid and accurate, but there is insufficient variability between isolates to allow biotyping or tracking a single isolate. Three genomovars of F. columnare were identified by Triyanto and Wakabayashi (1999). Subsequently, it was shown that genomovars I and II possess different pathogenicity; genomovar I resulted in 0–45% mortality in channel catfish, while genomovar II resulted in 92–100% mortality (Shoemaker et al. 2008). In addition, Olivares-Fuster et al. (2007) found that genomovar II was more frequently found in channel and blue catfish, and that genomovar I was found more frequently in nonictalurid species. These data indicate that genomovar II is a more serious pathogen for channel catfish.

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Table 11.1 Biophysical and biochemical characteristics of Flavobacterium columnare and Flexibacter maritimus. Characteristic

Flavobacterium columnare

Flexibacter maritimus

Cell morphology Colony morphology Cell size (µm) Yellow pigmented colony Motility Flexirubin pigment Binds Congo red Resistant to neomycin sulfate, polymyxin B Chondroitin lyase O-nitrophenyl- β-D-galactopymanoride Growth on peptone Glucose source of carbon Acid from carbohydrates Degradation of Gelatin Casein Starch Tyrosine Urease H2 S Nitrate reduced Catalase Cytochrome oxidase Optimum growth at (◦ C) Growth tolerance (◦ C) Growth in 0% SW-HS∗ 33% SW-HS 66% SW-HS 100% SW-HS G + C content (mol%) Habitat

Long, Gram negative rods Flat, rhizoid adheres to agar 0.3–0.5 × 3–10 + Gliding + +

Long, Gram negative rods Flat, irregular 0.5 × 2–30 + Gliding − +

+ + − + − −

+ − − + − +

+ − − − ? + − + + 25–30 10–37 + − − − 32–37 Freshwater (saprophytic)

+ + − + + − + + + 30 15–34 + + + + 33–42 Marine (saprophytic)

Source: Wakabayashi et al. (1986); Chen et al. (1995); Bernardet et al. (1996). +, positive reaction or characteristic; −, negative. ∗ SW-HS is Hsu–Shotts made with percentage of sea water.

Epizootiology Columnaris disease may occasionally occur as a primary infection without significant predisposing stress to the host, but more commonly the disease develops as a secondary pathogen due to environmental stress or trauma (handling, netting, crowding, etc.) or possibly nutritional stress. In either case, the disease can become acute accompanied with rapidly developing mortality. Columnaris may occur as the lone pathogen or in combination with external and systemic

infection with other pathogens of the body, fins, or gills. Hawke and Thune (1992) found that in 53 cases involving F. columnare in channel catfish, 11% were solely external, 17% were solely internal, and 72% were a combination of the two. Columnaris often appears in association with one or more other pathogens and is secondary to a more primary but less lethal organism (i.e., external protozoan parasites) or bacteria. Of the 53 F. columnare infections studied, 46 involved more than one bacterial infection, most of which were E. ictaluri or Aeromonas

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spp. Marks et al. (1980) were unable to experimentally induce a F. columnare infection unless a Corynebacterium sp. was also present. Chowdhury and Wakabayashi (1989) reported that F. columnare invaded fish when several other bacteria were present but not when A. hydrophila or P. fluorescens were present. They also showed that F. columnare did not survive well in vitro when density of A. hydrophila was approximately 100 times higher than that of F. columnare. In view of these conflicting reports, the actual role of F. columnare in primary or secondary infections and its relationship with other pathogens is unclear. In cultured channel catfish populations where no other fish species are present and the water supply comes from a well, catfish would be considered the pathogen source. Fish living in stream or reservoir water can also serve as a source for the bacteria. It was proposed by Bullock et al. (1986) that coarse fish (suckers, carp, etc.) are reservoirs of F. columnare. Transmission of columnaris is generally from fish to fish via water, but numerous factors can affect its transmission and contraction. A PCR detection procedure confirmed that F. columnare can be transmitted horizontally through water without fish-to-fish contact (Welker et al. 2005). However, the most common precursors to columnaris as a secondary infection in channel catfish include handling, seining, transportation, temperature shock, water quality (low dissolved or supersaturation of gasses—oxygen, nitrogen, etc.), and presence of other infectious diseases (Hanson and Grizzle 1985). Channel catfish are susceptible to columnaris at temperatures from 15 to 30◦ C, and young fish are more severely affected than adult fish. Centrarchids are especially susceptible when held in abnormally cool water during summer. While feed deprivation has been suggested to help reduce effects of some bacterial infections of channel catfish, i.e., ESC, this practice is detrimental to fish infected with columnaris and in fact may even preclude infection of F. columnare (Shoemaker

et al. 2003). Feed deprivation of 7 days reduces resistance to infection, but moreover deprivation for 2 and 4 weeks severely affects the fish’s weight and other health-related parameters, thus increasing susceptibility to columnaris. Mortality due to F. columnare in feeddeprived fish was as high as 78% compared to 1.7% in fed control fish. F. columnare infections can be chronic and cause lingering, gradually accelerating mortality in channel catfish, but more often it appears suddenly and accelerates to subacute mortality in a matter of days. Cases have been reported where columnaris infected juvenile channel catfish suffered about 50% mortality in a 24hour period, and 90% mortality in tank-held fingerling channel catfish is not uncommon during optimum disease conditions. Mortality in pond populations is usually lower, but may reach 50–60%. Bait minnows (fathead minnows and golden shiners) held in bait shop tanks are highly susceptible to columnaris, particularly if improperly handled and are crowded. Hussain and Summerfelt (1991) experimentally infected a group of walleye with columnaris in which up to 70% of mechanically injured fish contracted the disease, but no infection occurred in uninjured bait minnows. While columnaris occurs in every month of the year, it tends to be seasonal, especially in temperate climates where it has two peak periods (Figure 11.3). Infections increase in late March through April and low incidence occurs during summer but increases again in autumn. This pattern is probably dependent upon optimum water temperatures for the pathogen, the fact that greater numbers of susceptible size and age fish are present in spring and to movement and transport of fingerlings in the fall. Bowser (1973) found that black bullheads in Clear Lake, Iowa, had widespread columnaris infections in May and June with no reported incidences after July. F. columnare infections in Taiwan were reported by Kuo et al. (1981) to be the highest in tilapia and eel populations during September and October, lower in March through June,

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Pathology

Percentage columnaris per month

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Figure 11.3 Monthly distribution of F. columnare infections (all fish species) in Alabama from 1991 through 1996 (N = 1,274). (Diagnostic data compiled from diagnostic records at the Southeastern Cooperative Fish Disease Laboratory, Auburn University, Alabama, and the Alabama Fish Farming Center, Greensboro, Alabama.)

and very low in January through February and June through August. The ability of F. columnare to survive in water has been studied by several researchers. Fijan (1968) showed that pathogen survival was reduced at pH 7 or less, in waters with hardness less than 50 mg/L CaCO3 , and/or in waters containing low organic matter. Chowdhury and Wakabayashi (1988) found that F. columnare survival decreased little over 7 days in chemically defined water containing 0.03% NaCl, 0.01% KCl, 0.002% CaCl2 .H2 O, and 0.004% MgCl2 .6H2 O, but survival was reduced in water with higher concentrations of these ions. When sterile mud was seeded with F. columnare, 62% of the cells survived after 77 hours at 10◦ C, but only 35% survived at 20◦ C (Becker and Fujihara 1978). These data imply that under normal conditions the organism does not survive well in the environment for extended periods without a fish host; however, this possibility has not been totally eliminated.

Initial infections of F. columnare usually result from mechanical or physiological injury or environmental stressors, but can develop independent of stressors. Abrasion of the skin hastens columnaris invasion and dissemination to organs and tissues, but skin injury is not essential for infection to occur (Bader et al. 2003). However, columnaris cells were detected in all tissues more quickly in injured fish than in fish in which the skin remained intact with no injury. Darwish and Mitchell (2009) produced an acute columnaris infection in channel catfish by abrading the skin prior to exposure to the pathogen; however, it was proposed that lesion severity is dependent on strain virulence, and that necrotic lesions likely result from the organism’s proteolytic enzyme activity. Gill lesions start at the margins of filaments, and necrosis progresses toward the gill arch (Figure 11.1). Necrotic gills become congested and epithelium separates from the lamellae. When skin lesions occur, the dermis and underlying musculature become necrotic. Capillaries become congested and deteriorate, resulting in hemorrhaging at margins of the ulceration. Once integument is compromised by the bacterium, a systemic infection may follow. Generally very little pathology is associated with systemic columnaris infections of channel catfish; however, Hawke and Thune (1992) did note swelling of the trunk kidney in some case studies.

Significance From 1987 to 1989, columnaris was the most frequently reported infectious disease in the catfish industry accounting for 58% of all bacterial cases (Thune 1991) and continues to be a major disease factor. In fact columnaris constituted an average of 44.3% of the disease cases submitted to the Aquatic Diagnostic Laboratory, Stoneville, MS, from 1997 through 2005

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(Anonymous 2005). The broad geographic range of disease and extensive species susceptibility add to its significance, which has often been overlooked by fish pathologist because of its frequent role as a secondary pathogen. In reality, each year columnaris is probably responsible for killing as many cultured fish, irrespective of species, as any other bacterial organism.

Enteric septicemia of catfish (ESC) The genus Edwardsiella includes two bacterial species, which cause major disease problems in fish: E. tarda (Ewing et al. 1965) and E. ictaluri (Hawke et al. 1981). As reported by Mitchell and Goodwin (1999), E. ictaluri may have been present in cultured channel catfish as early as the 1960s. E. tarda, the causative agent of Edwardsiella septicemia, is discussed more extensively in Chapter 13 on eel bacterial diseases. E. ictaluri causes enteric septicemia of catfish (ESC), also known as “hole-in-the-head” disease (Hawke 1979; Hawke et al. 1981). This disease and columnaris are the two most significant infectious diseases of cultured catfish. Being the most frequently reported infectious fish disease in the southeastern United States, ESC causes losses in the millions of dollars and continues to be a major problem.

Geographical range and species susceptibility E. ictaluri has been confirmed in the United States, Thailand (Kasornchandra et al. 1987), and Australia (Humphrey et al. 1986). In a discussion and literature review, Evans et al. (2008) reported that the pathogen has also been found in Vietnam, Taiwan, Singapore, Croatia, and Spain. In the United States, ESC has been isolated from channel catfish, primarily from Florida to Texas and north to Missouri and Kentucky. However, the bacterium

has been reported among cultured channel catfish in other states including Arizona, California, Idaho, Indiana, Kansas, New Mexico, and Virginia. As channel catfish propagation expands around the world, the dissemination of E. ictaluri into new geographical areas will likely occur. E. ictaluri infects a narrower host range than many other warmwater fish diseaseproducing bacteria, but host susceptibility is expanding. Cultured channel catfish are much more susceptible to this bacterium than other ictalurids; however, white catfish, blue catfish, and brown bullhead have on occasion been naturally infected with E. ictaluri. Although there have been natural E. ictaluri infections reported in blue catfish, Wolters et al. (1996) showed that under experimental conditions, juvenile blue catfish had the highest rate of survival (90%), channel catfish the lowest (62%) and channel X blue hybrids had intermediate survival (74%) following pathogen exposure. Natural infections have been reported in walking catfish in Thailand (Kasornchandra et al. 1987) and in two aquarium species: Bengal danio (Waltman et al. 1985) and green knife fish in the United States (Kent and Lyons 1982). Experimental infections were established in chinook salmon and rainbow trout, but because of temperature constraints, the pathogen is unlikely to produce serious problems in salmonid species (Baxa et al. 1990). European catfish (sheatfish) are only slightly susceptible (Plumb and Hilge 1987), while several commonly cultured warmwater species were shown to be refractive (Plumb and Sanchez 1983). However, Evans et al. (2008) lists 23 fish species from which E. ictaluri has been isolated or are experimentally susceptible. These species include freshwater and marine fishes and several aquarium species. While considered a pathogen of fish, E. ictaluri was isolated from intestines of fish-eating great blue heron, snowy egret, and double-crested cormorants inhabiting catfish culture operations in the United States (Taylor 1992). However, Waterstrat et al (1999) were unable to culture

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E. ictaluri from great blue herons that were fed E. ictaluri-infected fish.

Clinical signs Enteric septicemia is a chronic, to subacute disease with almost pathognomonic clinical signs in channel catfish (Hawke 1979). Diseased fish hang listlessly at the surface with a “headup–tail-down” posture, sometimes spinning in circles followed by morbidity and death. Affected fish have pale gills, exophthalmia, and occasionally enlarged abdomens. Small depigmented lesions of 1–3 mm in diameter appear on the flanks and backs of infected fish. Lesions then progress into similar-sized inflamed cutaneous ulcers (Figure 11.4). In chronically ill fish, an open lesion may develop along the central skull line between the eyes at the insertion of the two frontal bones, thus the name “holein-the-head” disease. Petechia or inflammation in the skin under the jaw and on the operculum and belly may be so extensive that skin becomes bright red (paintbrush hemorrhage). Hemorrhage may also occur at the base of fins. Internally, the body cavity may contain a cloudy or bloody fluid, and on rare occasions, a clear yellow fluid, the kidney and spleen are hypertrophied, and the spleen is dark red; adipose tissue, peritoneum, and intestine are inflamed; and the liver is either pale or mottled with congestion (Figure 11.4).

Diagnosis Generally, E. ictaluri is isolated from visceral organs or brain of clinically infected fish on BHI agar or TSA, where punctate colonies form in 36–48 hours when incubated at 28–30◦ C. Careful observation of primary isolation on BHI agar culture plates is essential because of the possibility that other more rapidly growing bacteria, such as Aeromonas spp., will overgrow the E. ictaluri. Shotts and Waltman (1990) developed an Edward-

siella isolation media (EIM), which is selective for the bacteria and distinguishes it from other fish bacterial pathogens. On EIM, E. ictaluri produces clear greenish colonies, while the medium inhibits growth of Gram-positive bacteria. Other Gram-negative fish pathogens will grow on the media, but are separated by colony morphology and color (Figure 11.5). E. tarda colonies are small with black centers, A. hydrophila colonies are larger than E. ictaluri and are brownish, and Pseudomonas fluorescence colonies are blackish and punctate. A completely defined medium consisting of 46 individual components was developed for E. ictaluri by Collins and Thune (1996). By reducing the essential components from 46 to 8, they also described a “minimum essential medium” on which E. ictaluri grew. Confirmation of E. ictaluri requires identification by conventional biochemical characterization, serological or PCR procedures. Serological FAT and ELISA methods developed by Rogers (1981) utilized rabbit-anti E. ictaluri sera to identify E. ictaluri in either in vitro or in vivo. No cross-reactivity was detected with E. tarda, Salmonella sp., or A. hydrophila. A lack of serological cross-reactivity of E. ictaluri with E. tarda or A. hydrophila was also demonstrated by Klesius et al. (1991) using ELISA and monoclonal antibody. The specificity of E. ictaluri antibody was further shown by Chen and Light (1994), who found no correlation between agglutinating E. ictaluri antibody and A. hydrophila antibody. Ainsworth et al. (1986) used monoclonal antibodies in an indirect FAT in E. ictaluri diagnosis. When compared to bacterial culture of brain, liver, spleen, and anterior and posterior kidney, 71% of the fish were positive by culture and 62% were positive by FAT using the monoclonal antibody, but by combining the two methods, 90.3% of the fish were E. ictaluri positive. Other studies support the timesaving advantage of using immunoassay to obtain results in 2 hours vs. 48 hours required for culture and conventional bacteriological methods; however, pathogen isolation is still

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Catfish Bacterial Diseases 285

(a)

(b)

(c)

Figure 11.4 Channel catfish infected with Edwardsiella ictaluri. (a) Lower fish is not infected and the upper fish shows the early stages of disease development with depigmented areas(arrowhead). The middle fish has petechial hemorrhages (arrowheads) in the skin. (b) Channel catfish exhibiting open lesions in the cranial region (large arrowheads), inflamed nares (small arrows) typical of chronic infection and exophthalmia. (c) Viscera of E. ictaluri-infected channel catfish with mottled liver (large arrow) and enlarged spleen (small arrow) and bloody ascites pooled in body cavity (arrowhead).

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Figure 11.5 Colonies of four bacterial pathogens isolated from channel catfish growing on Edwardsiella isolation media (EIM). All cultures are 48 hours old incubated at 30◦ C. (a) Aeromonas hydrophila colonies are large and brownish; (b) Edwardsiella tarda colonies are medium size, greenish, and have black centers; (c) E. ictaluri colonies are medium size and pale green with darker green centers; (d) Pseudomonas fluorescence are small and black with a pale halo. (Photographs courtesy of D. Earlix.)

the most reliable method of demonstrating its presence in a fish population. Immunofluorescent tests were used simultaneously to detect E. ictaluri and F. columnare in tissue of diseased fish and the cultured bacteria taken from infected fish (Panangala et al. 2006a). Bilodeau et al. (2003) described a specific real-time PCR method that detected E. ictaluri in blood and other tissue; however, this method is limited by its ability to also detect dead bacteria. A multiplex-PCR procedure described by Panangala et al. (2007) simultaneously detected E. ictaluri, F. columnaris, and A. hydrophila. Baxa-Antonio et al. (1992) suggested that detection of humoral antibody in channel catfish could be used to identify populations that have been exposed to E. ictaluri. Klesius (1993) further refined the ELISA system by developing FAST-ELISA to detect antibodies against E. ictaluri “exoantigen” in 30 minutes. Monoclonal antibody in the FAST-ELISA could be valuable in detecting covert carrier fish or predicting overt infection. Tyler and Klesius (1994) showed that recognition of E. ictaluri exoantigen by ELISA is highly specific.

However, an absence of antibody may not always indicate whether a fish has been exposed to the pathogen as all individuals do not uniformly produce antibody following exposure (Vinitnantharat and Plumb 1993). Earlix et al. (1996) digested homogenized kidney tissue from infected channel catfish in Triton X-100 and filtered it with a 0.45-µm nitrocellulose membrane to trap the bacteria. Following culture of the membrane for 24 hours at 30◦ C on agar, an ELISA application with E. ictaluri-specific monoclonal antibody detected 80% E. ictaluri prevalence in asymptomatic channel catfish. This was compared to 24% prevalence by conventional bacteriological isolation. Implementing these techniques in diagnostic laboratories could be useful in determining E. ictaluri carrier populations.

Bacterial characteristics When biochemical characteristics of E. ictaluri (Table 11.2) were established, there

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Catfish Bacterial Diseases 287 Table 11.2 Biochemical and biophysical characteristics of Edwardsiella ictaluri, E. tarda, and E. hoshinae∗ . Characteristic

E. ictaluri

E. tarda

Cell morphology Motility 25◦ C 37◦ C Growth at 40◦ C NaCl tolerance 1.5% 4.0% Cytochrome oxidase Indole Methyl red Citrate (Christensen’s) H2 S on triple sugar iron Peptone iron agar Lysine decarboxylase Ornithine decarboxylase Malonate utilization Gas from glucose Acid production from D-mannose, maltose D-mannitol, sucrose Trehalose L-arabinose Jordans tartrate Nitrate reduced to nitrite Tetrathionate reductase On E. ictaluri media (EIM) Mol% G + C of DNA

Short gramnegative, single or paired rods + + − + − + + + − + − − − + − + − + − + − + + + + + − − + +

+ + + + + − ± + + − + + + + ±

+ − − − − + ? Green translucent 56–57

+ + ± ± − + + ? 53

+ − − − + + + Black centers 55–58

E. hoshinae

Source: Ewing et al. (1965); Grimont et al. (1980); Hawke et al. (1981); Waltman et al. (1986). +, positive for 90–100% of strains; −, negative for 90–100% of strains; ±, mixed reaction. ∗ All strains of E. ictaluri, E. tarda and E. hoshinae tested are negative for Voges-Proskauer, Simmons citrate, urea, phenylalanine deaminase, arginine dihydrolase, gelatin hydrolysis, growth on KCN; acid production from glycerol, salacin, adonitol, D-arabitol, celobiose, dulcitol, erythritol, lactose, L-rhamanose and D-xylose; acid from mucate, esculin hydrolysis, acetate utilization, deoxyribonuclease, lipase, Bgalactosidase (ONPG), pectate hydrolysis, pigment production, tyrosine clearing and oxidase test.

was very little diversity among the many isolates (Hawke et al. 1981; Waltman et al. 1986; Plumb and Vinitnantharat 1989). The organism is a short, Gram-negative rod (0.8 × 1–3 µm) that tends to elongate in actively growing cultures. It is cytochrome oxidase negative, weakly motile at 25–28◦ C, and lacks motility at 30◦ C or above. E. ictaluri grows poorly or not at all at 37◦ C. At 20–30◦ C, it ferments and oxidizes glucose while producing gas. It will not tolerate a NaCl level higher than 1.5% in the media. E. ictaluri can easily be separated from E. tarda by its indole negative reaction and lack of H2 S production on TSI. Although some reports claim that the Minitek and API20 identification systems were not suitable for E. ictaluri, Klesius et al. (2003) accurately ap-

plied API-20E, FAME, and Biolog systems to E. ictaluri isolates. The lipopolysaccharide (LPS) of E. ictaluri was characterized by Weete et al. (1988); the lipid A molecule was similar to other enteric bacterial species; however, E. ictaluri LPS differed from other species in some key characteristic. Furthermore, Arias et al. (2003) showed that LPS analysis patterns between a mutant E. ictaluri and its parent strain were nearly identical and concluded that the only way of fingerprinting E. ictaluri strains was by phenotypic profiles and fatty acid composition. Accumulated evidence indicates that generally there is only one serological strain of E. ictaluri and no cross-reaction occurs with other aquatic bacteria. Antibodies to E. ictaluri in

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naturally infected channel catfish were not removed by adsorption with nine other species of bacteria commonly found in fish intestines and fish ponds (Chen and Light 1994). Also channel catfish immunized with these nine bacterial species failed to develop E. ictaluriagglutinating antibodies. Studies by Panangala et al. (2005; 2006b) indicate close phenotypic similarities and analysis of 16S–23S intergenic spacer regions of the rRNA operons between E. ictaluri isolates, but there are some distinct differences in these characteristics between E. ictaluri and its close relative E. tarda. Lobb and Rhoades (1987) and Newton et al. (1988) described one to three plasmids in E. ictaluri isolates and indicated that all E. ictaluri isolates, regardless of origin, possess plasmids. The DNA of these plasmids is specific enough that Speyerer and Boyle (1987) suggested using them as detection probes for E. ictaluri in infected fish.

Epizootiology ESC is endemic throughout the U.S. channel catfish industry. During the early years of the known presence of ESC, relatively few cases were diagnosed, but a higher incidence of the disease occurred as awareness of it escalated. In 1981, 47 outbreaks were diagnosed in southeastern United States and 1420 diseases cases were attributed to E. ictaluri in 1985 (A. J. Mitchell, Fish Farming Experiment Station, Stuttgart, Arkansas, personal communication). In 1988, 1605 cases of E. ictaluri were documented constituting 30.4% of all reported fish disease cases in this region. In a survey by Wagner et al. (2002), they found that 78.1% of all channel catfish culture operations experienced columnaris and/or ESC or a combination of the two diseases. Average loss in these outbreaks ranged from 90 to 900 kg per episode. When first described, E. ictaluri was thought to be an obligate pathogen that could not survive for an extended period outside the

host (Hawke 1979). It has now been determined that it can survive in pond bottom mud for over 90 days at 25◦ C and in water for up to 15 days at 25◦ C or less (Plumb and Quinlan 1986). Survival of E. ictaluri in the environment may be influenced by microbial competition because survival is shortened in water or mud that contains other microbes (Earlix 1995). Questions concerning how long E. ictaluri remain in epizootic survivors have not been fully answered; however, Mgolomba and Plumb (1992) found significant numbers of bacteria in blood and all organs of fish 81 days after exposure. Klesius (1992) isolated E. ictaluri from channel catfish in a population 280 days after initial pathogen detection. Nearly all E. ictaluri epizootics have occurred in channel catfish culture environments, but Chen et al. (1995) found serological evidence that catfish living in natural waters had been exposed to the bacterium. E. ictaluri is readily phagocytized by macrophages in the blood, but indications are that bacteria are not destroyed in these cells (Figure 11.6) (Miyazaki and Plumb 1985); therefore, phagocytes may contribute to the bacteria’s longevity in carrier fish. Transmission from adult channel catfish to offspring at spawning also seems possible, but as yet is unproven. The primary mode for E. ictaluri transmission is through water via carrier channel catfish, which sequester the bacteria in their intestines and shed it into the water. Transmission of E. ictaluri from fish that died of ESC to nondiseased contact fish occurred by cannibalism or from bacteria shed by dead fish (Klesius 1994). The practicality of daily removal of dead fish from a culture unit was shown by Earlix (1995), who found that during an ESC epizootic, significantly higher concentrations of E. ictaluri were present in pond water in areas containing large numbers of dead fish than in areas with no carcasses. Because dead fish shed large numbers of bacteria, their removal can reduce bacterial concentrations to which noninfected

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(a)

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Figure 11.6 (a) Smear from lesion in the skull of channel catfish showing intracellular E. ictaluri (arrow) in macrophages (H & E stain) (photo by T. Miyazaki). (b) Electron micrograph of olfactory organ from channel catfish illustrating E. ictaluri (arrow) invading the tissue (×15,000). (Photo by E. Morrison.)

fish are exposed. It also is possible that cormorants, herons, etc., can be vectors of E. ictaluri (Taylor 1992). It was demonstrated by ELISA that viable E. ictaluri can survive in the intestines of these birds, thus suggest-

ing that fish-eating birds can be a pathogen source. Although ESC has been diagnosed during every month of the year and in a wide range of water temperatures, it is considered a

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Percentage ESC per month

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Figure 11.7 Seasonal occurrence of E. ictaluri (N = 1,252) in Alabama from 1991 through 1996 showing greatest incidence of disease in May–June and September–October when water temperatures are 20–28◦ C. (Diagnostic data compiled from diagnostic records at the Southeastern Cooperative Fish Disease Laboratory, Auburn University, Alabama, and the Alabama Fish Farming Center, Greensboro, Alabama.)

seasonal disease occurring primarily when temperatures range from 18 to 28◦ C in late spring to early summer and again in autumn (Figure 11.7). Mortality in experimentally E. ictaluri-infected channel catfish fingerlings was highest at 25◦ C, slightly lower at 23 and 28◦ C, and no deaths occurred at 17, 21, or 32◦ C (Francis-Floyd et al. 1987). BaxaAntonio et al. (1992) found that channel catfish experimentally infected with E. ictaluri by immersion experienced 0% mortality at 15◦ C, 46.6% at 20◦ C, 97.8% at 25◦ C, 25% at 30◦ C, and 4% at 35◦ C. However, an increase of ESC outbreaks during July and August has been noted in case records, which could indicate an expanding temperature tolerance for the pathogen. Mortality rate in E. ictaluri-infected catfish populations in ponds varies from less than 10% to over 50%. The pathogen infects fingerling as well as production-size fish and occurs in ponds, raceways, recirculating systems, and cages. Even though ESC can develop inde-

pendent of extrinsic influences, adverse environmental circumstances can intensify disease severity. The precise relationship between environmental quality and E. ictaluri outbreaks has not been fully elucidated, but during experimental infections, Wise et al. (1993a) established a correlation between confinementinduced stress and increased susceptibility of channel catfish to the pathogen. They also concluded that stress induced by handling and hauling likely increases mortality due to ESC. The effect of stress on ESC was shown by Ciembar et al. (1995), who determined that stressed channel catfish suffered higher cumulative infections (47%) 3 weeks after immersion exposure to E. ictaluri than did unstressed fish (16%). Injection of the corticosteroid Kenolog increased susceptibility of channel catfish to an initial as well as a second exposure to E. ictaluri. It also increased carrier rate of survivors from one out of eight (12.5%) in controls to eight out of eight (100%) in Kenolog-injected fish (Antonio and Hedrick 1994). From these studies, there is little doubt that increased stressful conditions play a role in susceptibility and severity of E. ictaluri. Blood cortisol concentration is one measure of stress on animals. Sink and Strange (2004) found by stressing fish for 30–60 minutes in a plastic net and then exposing the fish to E. ictaluri that blood cortisol was significantly elevated and the stressed fish were significantly more susceptible to the pathogen. Also, the longer the period of stress (30–60 minutes), the greater the susceptibility of fish to E. ictaluri. Increasing salinity of water up to 5000 mg/L (0.5%) for short periods of time or during transport has been historically used as a fish prophylactic. Antidotal evidence indicates that catfish farms with elevated salinity in artesian well water supplies have less ESC problems as well as other diseases. In view of this, it was shown that channel catfish were less susceptible to E. ictaluri in water up to 4000 mg/L salinity (G. Whitis, Aquaculture

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Extension Specialist, Alabama Fish Farming Center, Greensboro, Alabama). Certain dietary nutrients may affect ESC severity. It was shown by Paripatananont and Lovell (1995) that increased dietary zinc (15–30 mg Zn/day) enhanced channel catfish resistance to E. ictaluri, and organic zinc methionine was of greater value than inorganic zinc sulfate. Diets containing corn materials with fumoninsin (product of the fungus Fusarium moniliforme) reduced channel catfish growth and resistance to E. ictaluri infections (Lumlertdacha and Lovell 1995). Wise et al. (1993b) noted that addition of 60 mg or more of vitamin E into diets of channel catfish increased agglutinating antibody titers and enhanced ability of macrophages to phagocytize virulent bacteria. The effects of elevated dietary arginine on resistance of channel catfish to E. ictaluri were studied by Buentello and Gatlin (2001) and found that fish fed 2% arginine for two weeks prior to exposure to the pathogen positively influenced reduced mortality of fish and maximized survival. However, it is clear from the environmental and nutritional studies that effects of these factors on susceptibility of channel catfish to ESC are poorly understood. There is no indication that E. ictaluri poses a health threat to aquatic animals other than fish. Temperature limitation under which E. ictaluri grows precludes the bacterium from being a pathogen for warm-blooded animals.

Pathology The most severely damaged organs in E. ictaluri-infected catfish are the trunk kidney and spleen. Both organs develop necrosis, and the liver becomes edematous. Interlamellar gill tissue proliferates, and skin epidermis is destroyed. Mild focal infiltration occurs in the musculature underlying the epidermis, where bacteria apparently colonize the capillaries causing skin depigmentation and necrosis (Miyazaki and Plumb 1985; Shotts et al.

1986). Ulcerative head lesions are necrotic and hemorrhaged, whereas systemic infections are associated with necrosis of hepatocytes and pancreatic cells. Intact, apparently dividing E. ictaluri cells are seen within macrophages (Figure 11.6). This phenomenon was also seen in electron micrographs of macrophages in tissues of the olfactory sac (Morrison and Plumb 1992). After experimental exposure to 5 × 108 CFU/mL of E. ictaluri via immersion, 93% of channel catfish developed acute ESC and 7% developed chronic disease (Newton et al. 1989). Acute infection was characterized by hemorrhage, ulceration, and enteritis, olfactory sacculitis at 2 days postexposure followed by hepatitis, and dermatitis. Chronic ESC, most commonly observed at 3–4 weeks postexposure, was characterized by cranial swelling, ulceration, granulomatous inflammation, and meningeoencephalitis of the olfactory bulbs, tracts, and lobes of the brain. Fish can be infected with E. ictaluri by injection, waterborne exposure, or ingestion. Indications are that in waterborne exposure, the olfactory sac of channel catfish is a primary invasion site for E. ictaluri, and phagocytes play a role in establishing a septicemia. Waterborne bacteria invades the olfactory organ via the nasal opening, migrates into the olfactory nerve, enters the brain (Shotts et al. 1986), and spreads from the meninges to the skull and skin, which creates a “hole-in-the-head” condition. Morrison and Plumb (1992) showed that E. ictaluri can attach to the olfactory epithelium and migrate into the submucosa (Figure 11.6). Injury included loss of sensory cilia and microvilli from the olfactory mucosal surface within 1 hour of exposure. Degeneration of olfactory receptors and supporting cells was evident 24 hours postinfection. Electron microscopy confirmed E. ictaluri on the mucosal surface and within the epithelium. Host leukocytes migrated through the olfactory epithelium into the interlamellar lumen and phagocytized the bacteria that did not appear to be destroyed.

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The gills are also a primary site of E. ictaluri invasion during immersion (Nusbaum and Morrison 1996). Using radiolabeled E. ictaluri, they demonstrated that the organism colonizes the gill epithelium in large numbers in 2–72 hours. Following increasing bacterial numbers on the gill, they rapidly migrate to the liver and less rapidly to the trunk kidney, gut, and brain. Nusbaum and Morrison (2002) showed by fluorescent antibody that E. ictaluri appeared on gills within 5 minutes of immersion and could be isolated from the circulatory system within 15 minutes. When infected fish were placed in clean water, the bacteria were cleared in 15 minutes from the blood but reappeared at 12-h postinfection. In this experiment, A. hydrophila appeared in the blood of 100% of fish 7 hours postchallenge with E. ictaluri, but prior to E. ictaluri challenge, only 20% carried A. hydrophila. These studies suggest that a latent infection of A. hydrophila coupled with a superinfection of E. ictaluri may enhance the appearance of clinical ESC. Ingested E. ictaluri entering the bloodstream through the intestine results in a septicemia (Newton et al. 1989). Catfish orally exposed to E. ictaluri developed enteritis, hepatitis, interstitial nephritis, and myositis within 2 weeks of infection. Baldwin and Newton (1993) showed that at 0.25 hours postintestinal exposure via oral gavage, E. ictaluri was found in the kidneys, thus indicating a rapid transmucosal passage. They also suggested that E. ictaluri may have invasion and survival potential in the host that is similar to other invasive Enterobacteriaceae. Little is known about the pathogenic mechanism of E. ictaluri. However, regarding the attachment mechanism, Wolfe et al. (1998) showed that bacterial lectins were instrumental in the attachment mechanism of E. ictaluri by utilizing specific sugar residues, particularly d-mannose, N-acetylneuraminic acid and l-fucose in the nasal mucosa. Stanley et al. (1994) demonstrated by electron microscopy a fibril network of connecting cells that were

similar to attachment fibrils seen by Morrison and Plumb (1994) when E. ictaluri attached to the olfactory mucosa. When comparing virulent to avirulent isolates, virulent cells had a greater amount of capsular material and surface proteins and increased chondroitin degradation. Williams et al. (2003) compared cellular and extracellular products of virulent and attenuated strains of E. ictaluri and showed that hemolysin activity was significantly greater in virulent strains. However, scanning electron microscopy found no conclusive difference in surface structure expression between pathogenic and attenuated strains of E. ictaluri. Intracellular replication of E. ictaluri in channel catfish macrophages was reported by Booth et al. (2006). They showed that E. ictaluri enters, survives, and replicates in head kidney-derived macrophages from channel catfish. Microscopic examination of infected macrophages revealed numerous intracellular bacteria that were contained in clear vacuoles. Invasion by the bacteria and replication were related to opsonization of the cells by normal serum, but it did not relate to the increase in the number of bacteria once it entered the macrophage.

Significance ESC is one of the most economically important infectious diseases of cultured channel catfish in the United States, where it is the causative agent of a large percentage of diagnosed diseases. Enteric septicemia constituted an average of 35.8% of the disease cases submitted to the Aquatic Diagnostic Laboratory, Stoneville, MS, from 1997 through 2005 (Anonymous 2005). E. ictaluri infections cost the aquaculture industry millions of dollars annually in killed fish and expenditures for prevention and chemotherapy. The effect of ESC on reduced growth and higher feed conversion ratios is only speculative, but this may be significant. Because channel catfish depend on olfactory

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functions while feeding, any injury to these sensitive tissues most likely affects the feeding process and would help explain why affected fish do not feed once disease has reached a certain level of severity.

Motile Aeromonas septicemia (MAS) MAS is associated with infections caused by motile members of the genus Aeromonas. The disease has a number of synonyms including hemorrhagic septicemia, infectious dropsy, infectious abdominal dropsy, red pest, red disease, red sore, rubella, and others. Currently known as MAS, the disease became part of modern fish health in the 1930s when it was named infectious dropsy of carp in Europe. In North America, the syndrome was known as hemorrhagic septicemia until the mid-1970s when its name was changed to “motile Aeromonas septicemia” at which time it became apparent that more than one motile member of the Aeromonas genus could cause the same disease syndrome. A. hydrophila (punctata, liquefaciens), A. sobria, A. veronii, and A. caviae are the principal motile Aeromonas species that affect fish. Carnahan et al. (1991) listed seven motile species of the genus, but found only the four mentioned to be regularly associated with diseased fish.

Geographical range and species susceptibility Motile Aeromonas septicemia is a ubiquitous disease that affects fishes in warm-, cool-, and cold freshwater around the world, and occasionally those in brackish water, especially in Europe, North and South America, and Asia. Motile Aeromonas septicemia is generally, but not exclusively, associated with warmwater fish. Channel catfish, other ictalurids, silurids, clariads, carp and other cyprinids, eels, centrarchids, and true basses (striped bass) are susceptible. Trout and salmon are also susceptible but are usually affected only when wa-

ter temperatures reach their upper tolerance (stressful) limits (Nieto et al. 1985). As far as is known, no fish species is totally immune or resistant to bacteria that cause MAS. Motile aeromonads infect other aquatic animals as well, causing “red-leg” disease in frogs and fatal disease in reptiles (Shotts et al. 1972). They can also produce fatal septicemia and localized infections in humans (Goncalves et al. 1992).

Clinical signs Clinical signs of MAS are varied, diverse, and generally are categorized as either behavioral or external signs, but internal lesions do occur. Motile Aeromonas septicemia-infected fish lose their appetite, become lethargic, and swim lazily at the surface. When diseased fish first appear at the surface, they usually dive when disturbed but eventually lose their equilibrium, return to the surface, and move into shallow water. External signs of MAS are varied with none being specific, but slight differences in disease manifestation can be noted between fish types. In catfish and other fishes without scales, fins are frayed, hemorrhaged or hyperemic, and congested. Epidermal lesions begin as irregularly shaped depigmented areas that eventually develop into necrotic patches of skin, which slough, leaving open, ulcerated lesions with exposed muscle (Figure 11.8). Lesion margins are whitish or hemorrhaged. External lesions can occur at any location: caudal peduncle, dorsally, ventrally, laterally, or on top of the head. On scaled fish, fins with whitish margins become frayed; skin lesions begin as hemorrhages at the base of scales and have a red (inflamed) central area surrounded by whitish (necrotic) tissue from which scales have been lost (Figure 11.8). In scaleless and scaled fish, the saprophytic fungus Saprolegnia spp. will often attack necrotic tissue, giving the lesion a fuzzy, brownish appearance. Motile Aeromonas septicemia-infected fish may have exophthalmia with hemorrhages or

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(a)

(b)

(c)

Figure 11.8 Motile Aeromonas septicemia (Aeromonas hydrophila)-infected fish. (a) Channel catfish with necrotic lesions (arrows) with exposed musculature, (b) early stage of A. hydrophila-induced lesion with hemorrhage on margin of necrotic tissue (arrows), and (c) largemouth bass with ulcerative skin lesions.

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opaqueness of the eye, enlarged abdomens with ascites, pale gills indicative of anemia, edematous musculature, and, in scaled fish, scales that show lepidorthosis before being lost. Internally, organs are friable, have a generalized hyperemia, swollen kidney and spleen, and the liver is often mottled with hemorrhage interspersed with pale areas. The body cavity may contain a clear fluid (ascites), but more often the fluid is bloody and cloudy. The intestine is flaccid, hyperemic, contains yellowish mucus, and is void of food.

Diagnosis Diagnosis of MAS is not based solely on clinical signs because other bacterial organisms (i.e., Pseudomonas sp.) and protozoan parasites (i.e., Epistylis sp. and Ichthyobodo sp.) often produce similar clinical signs and identical external lesions. A definitive MAS diagnosis can only be made by a complete necropsy of diseased fish in conjunction with isolation and identification of the causative organism. Primary isolation of motile aeromonads (A. hydrophila, A. sobria, or A. caviae) is made on either BHI or TSA. The cultures incubated at 25–30◦ C for 24–48 hours develop entire, slightly convex colonies, which are mucoid and white to yellowish in color. A. hydrophila can be isolated and identified on Rimler-Shotts (RS) selective media where an orange-yellow colony will form when incubated at 35◦ C (Shotts and Rimler 1973). Nielsen et al. (2001) used a PCR procedure to identify A. hydrophila in aquaculture in China, where it was the most accurate method of A. hydrophila identification.

Bacterial characteristics Motile Aeromonas spp. are Gram-negative, short, motile rods, which are cytochrome oxidase positive and ferment glucose; these characteristics constitute presumptive identity for motile Aeromonas spp. These bacteria mea-

sure 0.8–0.9 × 1.5 µm, are polar flagellated, produce no soluble pigments, and are resistant to vibriostat (0/129) (2,4-diamino-6,7disopropyl pteridine phosphate) and Novobiocin (Table 11.3). The genus Aeromonas is in the family Vibrionaceae; however, Colwell et al. (1986) proposed that the family Aeromonodaceae, which includes Aeromonas spp., be recognized. A. hydrophila, A. sobria, and A. caviae are distinguished by biochemical tests that include gas from glucose, esculin hydrolysis, and acid from arabinose (Table 11.3). Serological identification of motile aeromonads is not a common tool because of antigenic diversity and genetic complexity within the species (Janda 1991). This is particularly true of the ubiquitous A. hydrophila, which has as many as twelve 0-antigens and nine H-antigens. Historically, monovalent antiserum to a specific strain of A. hydrophila agglutinates only a small percentage of heterologous A. hydrophila. However, a virulent strain of A. hydrophila from epizootic ulcerative syndrome (EUS) in Asia was used to produce a monoclonal antibody (designated F26P5C8) that identified virulent strains of this bacterium from other epizootics as serotype I (Cartwright et al. 1994). This monoclonal antibody also identified a large number of A. hydrophila isolates from Australia and Japan, but their virulence was unknown.

Epizootiology Motile Aeromonas septicemia has been one of the most frequently diagnosed bacterial fish diseases since 1972 and was the most severe disease problem encountered by catfish farmers until the mid-1980s (A. J. Mitchell, Fish Farming Experiment Station, Stuttgart, Arkansas, personal communication). The disease accounted for as many as 60% of total bacterial diagnostic cases reported in some years during that period. However, at the Aquatic Diagnostic Laboratory, Stoneville, MS, MAS constituted less than 1% of the

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Table 11.3 Biochemical and biophysical characteristics of Aeromonas hydrophila, A. caviae, and A. Sobria. Character/substrate

A. hydrophila

A. caviae

A. sobria

Morphology Cytochrome oxidase Catalase Growth on nutrient agar (37◦ C) Ornithine decarboxylase Indole production Fermentation Glucose, sucrose, manitol Dulcitol, rhamnose, xylose Raffinose, inositol, adonitol NO3 reduction to NO2 Growth on peptone without NaCl 0/129 resistant Hydrolysis of starch, gelatin ONPG, RNA, and DNA hydrolysis Gas from glucose∗ Salacin B-D xylosidase Arbutine G-glucosidase Acid from arabinose∗ Voges-Proskauer Elastase Esculin Growth on KCN L-histidine utilization L-arginine utilization H2 S from cystine

Short, gram negative, single or paired rods + + + + + + − − + +

+ + + − +

+ − − + + + +

+ − − + + + +

+ − − + + + +

+ + + + + + + + + + + + +

− + + + + + − − + + + + −

+ − − −(14%+) −(29%+) − + − − − − − +

Source: Shotts and Rimler (1973); Popoff (1984) +, positive reaction; −, negative reaction for substrate. ∗ Characteristics most often used to separate the three species if they are isolated from fish.

total cases between 1997 and 2005 (Anonymous 2005). Recently, MAS has caused severe losses in catfish production facilities in West Alabama. In 2009, there were 99 cases of MAS in West Alabama with losses exceeding 3 million pounds. All of the disease outbreaks in West Alabama appear to be caused by the same strain. Producers have obtained an INAD to use Florfenicol to treat these recent outbreaks and work toward a label for MAS in catfish (Personal communication William Hemstreet, Alabama Fish Farming Center). Motile Aeromonas septicemia is generally a seasonal disease, but has been diagnosed every month of the year across the southern United States where it peaks in the spring (Figure 11.9). As one moves northward, in-

fections tend to occur in late spring or early summer. This disease pattern can be attributed to lower resistance of fish following winter, and spring environmentally induced stressful periods, hormonal imbalance during spawning, presence of large numbers of susceptible young fish, and the fact that water temperatures in spring are more conducive to bacterial infections. As summer progresses, fish tend to develop an immunity and/or natural resistance to Aeromonas spp., thus MAS outbreaks decrease only to rise again in fall when water temperatures are more favorable, and juvenile fish are being handled and transported. The 2009 MAS outbreak in West Alabama did not follow this bimodal trend; the highest losses occurred in late summer and the loses were

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Percentage MAS per month

20

10

0 J

F

M A

M

J J A Month

S

O

N

D

Figure 11.9 Monthly distribution of motile Aeromonas septicemia infections in fish in Alabama from 1991 through 1996 (N = 919). (Diagnostic data compiled from diagnostic records at the Southeastern Cooperative Fish Disease Laboratory, Auburn University, Alabama, and the Alabama Fish Farming Center, Greensboro, Alabama.)

not associated with a particular stressor, indicating that the A. hydrophila was a primary pathogen (William Hemstreet personal communication). Fish mortality associated with motile Aeromonas infection is usually chronic with obscure peaks of deaths per day; however, cumulative mortality can be significant. High mortality may result when a highly virulent bacterial strain is involved, but normally total losses will be below 50%. Infections of A. hydrophila can occur in very young to adult fish. Epizootics of young fish may be subacute, whereas die-offs in older fish are more chronic. Kage et al. (1992) found that 90–100% of 7day-old Japanese catfish died as a result of an A. hydrophila infection. Infections may be external, internal (systemic), or more often both. In external infections, skin lesions from which bacteria can be isolated are the only obvious clinical sign of disease, and bacteria cannot be isolated from internal organs. It can be argued that this phase of the syndrome should not be considered

motile Aeromonas septicemia because there is no septicemia. Systemic infections are characterized by a septicemia, and the causative organism can easily be isolated from any internal organ as well as skin lesions. Infections of A. hydrophila in fish are often associated with a predisposing stress such as temperature shock, low oxygen (See Figure 1.5), high ammonia and other adverse water quality problems, trauma from improper handling or hauling, and presence of other disease organisms (Plumb et al. 1976; Grizzle and Kiryu 1993). Peters et al. (1988) found that even social stress of juvenile rainbow trout enhanced ventilation, elevated glucose and leukocyte volume, and increased susceptibility to A. hydrophila. Motile Aeromonas infections in most animals, including humans, usually occur as a secondary infection when a debilitating condition already exists; however, it can be a primary infection, and if A. hydrophila becomes established, it is often the cause of morbidity and death. A. hydrophila infections in wild fish are often associated with rivers, eutrophic lakes, and ponds that receive large amounts of organic enrichment (Shotts et al. 1972), conditions similarly found in catfish culture ponds that receive large daily feed quantities. The relationship between water quality-induced stress and an A. hydrophila infection in an enriched channel catfish culture pond was shown by Plumb et al. (1976) and later duplicated in the laboratory (Walters and Plumb 1980). An abrupt decrease in the ponds dissolved oxygen concentration resulted in a drop in pH and increased concentrations of ammonia and carbon dioxide, followed in 6 days by an A. hydrophila infection (See Figure 1.3). Susceptibility of some fish species to A. hydrophila is also linked to water temperature. Groberg et al. (1978) reported that when exposed to A. hydrophila, coho, chinook salmon, and steelhead trout suffered 64–100% mortality at 18◦ C compared to 0% mortality at 9.4◦ C. Rainbow trout demonstrated an increased susceptibility to A. hydrophila when

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water temperature was raised from 5.5 to 8◦ C to more than 11◦ C (Nieto et al. 1985). Algae in unclean holding tanks can also contribute to higher A. hydrophila concentrations and incidence of infection (Levanon et al. 1986). Although rare, A. hydrophila may cause high mortality among cultured fish without presence of severe external (stressful) influences as was the case in West Alabama in 2009. This inconsistency may result from the presence of A. hydrophila strains that possess specific virulent or pathogenic characteristics. There is evidence that the motile Aeromonas complex involves secondary and opportunistic pathogens, but A. hydrophila’s ability to cause disease and death of fish should not be overlooked because occasionally highly virulent strains emerge. Regardless of whether or not the organism serves as a primary or secondary invader of stressed fish, it is often the final insult that leads to death. In Asia, A. hydrophila has been closely associated with EUS, a disease that has plagued both wild and cultured fish populations from Indonesia to India since the early 1980s (Boonyaratpalin 1989). The definitive etiology of EUS is uncertain because a virus (snakehead rhabdovirus), fungus (Aphanomyces invaderis), and A. hydrophila have all been associated with the lesions. In Southeast Asia, Aphanomyces was found in most fish lesions associated with EUS (Roberts et al. 1993; Willoughby et al. 1995). The massive tissue damage incurred was blamed on the fungus, but researchers were skeptical as to whether or not this organism was the primary pathogen. While clinical signs of EUS and MAS are almost identical, investigators have suggested that A. hydrophila is secondary to the primary cause, but may have been what actually killed the fish. A. hydrophila, which adapts readily to its environment, is found in most natural freshwater ponds, streams, reservoirs, and bottom mud where it exists as a facultative organism utilizing any available organic material as a nutrient source (Hazen et al. 1978). Often dif-

ferent bodies of water, or watersheds, harbor a unique strain of A. hydrophila because of its serological diversity and the widespread facultative nature of the bacterium. If fish resistant to a particular strain of the bacterium are moved to a different body of water, the possibility exists that they will be exposed to a different A. hydrophila strain for which they have no immunity and may become infected as a result of handling and transport. As noted earlier, A. hydrophila primarily affects fish, but other aquatic animals and humans can also be infected. In humans, these infections can become quite severe, causing enteritis, meningitis, and localized infections on extremities (Ketover et al. 1973). Puncture wounds inflicted by catfish spines may result in serious A. hydrophila infections (Hargraves and Lucey 1990). Several human deaths due to A. hydrophila have been reported, but often the patient was debilitated by another ailment. Goncalves et al. (1992) attributed a case of pneumonia in a healthy male to an A. hydrophila septicemia when he became ill 3 days after swimming in the ocean, suggesting that ocean water was the source. The fact that A. hydrophila is a freshwater organism casts some doubt as to the source. King et al. (1992) reported an Aeromonas spp. isolation rate in humans of 10.6 cases per one million people in California. Although 2% of the patients died, all had serious underlying medical conditions in addition to the Aeromonas infection. They concluded that Aeromonas spp. infections in humans is not an important public health problem and is largely unpreventable; however, Aeromonas infection in humans is a reportable disease in California. Because of an increased incidence of A. hydrophila infections in humans, handling infected fish with caution is advised. Pathogenic capabilities of A. hydrophila, A. sobria, and A. caviae are still unclear. Gray et al. (1990) found that in 61 isolates of motile Aeromonas spp. taken from pig and cow feces and other environmental sources (none were from fish), 96.4% of A. hydrophila, 36.4%

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of A. sobria, and only 11.6% of A. caviae were cytotoxic. The conclusion was that A. hydrophila was highly pathogenic, A. sobria was only moderately pathogenic, and A. caviae was not pathogenic. This may not hold true for isolates from fish because of cultures taken from MAS diseased channel catfish in Mississippi: A. sobria was more frequently found than A. hydrophila (M. Johnson, Mississippi State University, Stoneville, Mississippi, personal communication). In view of this, it is possible that A. sobria may be the primary cause of MAS infections in cultured channel catfish but often misidentified as A. hydrophila. However, recent MAS epizootics in West Alabama were caused by A. hydrophila. In some regions of the world and non-catfish species, A. sobria may be a more serious problem; as Wahil et al. (2005) points out, this pathogen caused 1% mortality per day in cultured perch in Switzerland. The clinical signs of A. sobria were similar to those of A. hydrophila.

Pathology Pathology of MAS is not distinctly different from most other septicemic bacterial infections in fish (Miyazaki and Jo 1985; Ventura and Grizzle 1988). Lesions are typical of those caused by bacterially produced protease and hemolysins. Epidermal infections are characterized by necrotic lesions that have spongy centers and hemorrhagic margins. The epidermis adjacent to lesions is edematous, and the dermis becomes hemorrhagic with macrophage and lymphocyte accumulation accompanied by severe inflammation. In scaled fish, scale pockets become edematous, causing lepidorthosis. In systemic infections, most internal organs are edematous with diffused necrosis of the liver, kidney, and spleen. Inflammation is not apparent in diseased internal organs, but hemorrhage and/or erythema does occur. Virulence of A. hydrophila may be influenced by specific biochemical characteris-

tics or by production of extracellular products that independently produce pathological effects when injected into fish (Santos et al. 1987). It was reported by Wakabayashi et al. (1980) that virulence of A. hydrophila is related to proteolytic casein and elastin hydrolysis. By correlating extracellular enzymatic activity of 127 strains of A. hydrophila, they showed that elastase positive strains produced lesions and mortality when injected into channel catfish. It was suggested that other extracellular substances such as hemolysins and proteases may also be involved with A. hydrophila’s pathogenic mechanism (Chabot and Thune 1991). Additional factors that may influence pathogenesis of the organism are its ability to adhere to tissue surfaces, resistance to phagocytosis, production of surface proteins, siderophores, and LPS; presence of pili, S-layer or outer membrane proteins; and the bacteriostatic activity of host serum. Also, water isolates of the bacterium may be avirulent or less pathogenic to fish than are isolates from diseased fish (de Figueiredo and Plumb 1977). The significance of these potentially pathogenic factors is as yet unclear; therefore, in the final analysis, the only accurate measure of the organism’s virulence and pathogenicity is whether or not it kills fish.

Significance There is disagreement among fish pathologists as to the significance of MAS. The frequency of its appearance cannot be disputed, but the fact that MAS is usually a secondary disease reduces its importance for many fish pathologists. However, when MAS, especially A. hydrophila and A. sobria, are present in an aquaculture environment where a high potential for stress exists, the disease cannot be ignored because, in most instances, it is not a primary pathogen, but it is often what kills the fish. Also, the fact that the motile Aeromonas group occurs worldwide and affects such a large variety of fish species in a variety of freshwater

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environments makes the disease syndrome an important factor in both cultured and wild fish populations.

Other bacterial diseases of catfish Several other bacterial organisms occasionally cause infectious diseases in channel catfish. The most significant of these diseases are Edwardsiellosis (E. tarda) and infections caused by P. fluorescence or Bacillus mycoides. Other Gram-positive bacteria that have rarely been detected in diseased channel catfish are Carnobacterium spp., Staphylococcus sp. (Chapter 17), and Streptococcus sp., but these are considered insignificant in terms of impact on channel catfish and are not discussed.

Pseudomonas septicemia P. fluorescence causes a disease, Pseudomonas septicemia, that is nearly identical to MAS and at one time was considered as one cause of “hemorrhagic septicemia.” This organism is a facultative bacterium in many water sources, where it occasionally causes infections in channel catfish during and/or following environmental stress. Clinical signs, morbidity, and mortality patterns are similar to those of MAS, and when P. fluorescence is present, it usually causes septicemia. P. fluorescence is isolated from skin, muscle lesions, and internal organs on either BHI or TSA agars. The organism is a Gram-negative bacillus, slightly motile and cytochrome oxidase positive. Colonies of P. fluorescens on EIM are small and black (Figure 11.5). It is distinguished from motile aeromonads and other fish pathogenic bacteria because it is oxidative only in glucose motility deep, it does not produce gas, and it produces a diffusible fluorescent pigment on hard agar.

and is discussed in more detail in Chapter 13 (Meyer and Bullock 1973). This organism is seen in about 1–2% of catfish disease cases, but when present, it can be a major problem. Generally, prevalence of EPDC in an infected population is less than 5%, but when fish are crowded in holding tanks, incidence can increase to 50% or higher. An E. tarda infection in channel catfish is characterized by lethargic swimming and the presence of small, 3–5 mm cutaneous lesions located dorsolaterally on the body. Within the flank muscles or caudal peduncle, these small lesions progress into larger abscesses and develop obvious convex, swollen areas. The skin loses its pigmentation, and incised lesions emit a foul-smelling odor. As infection progresses, posterior body mobility is lost. Internally, there is a general hyperemia characteristic of septicemia, and the body cavity has a putrid odor. The kidney, in particular, is enlarged, and the liver mottled or abscessed. E. tarda infections are usually prevalent in channel catfish of approximately 0.4 kg or larger; however, subadult channel catfish are not resistant. In channel catfish experimentally infected with E. tarda, pathogenesis and histopathology developed rapidly, following skin injury and waterborne exposure, but the process was relatively short lived (Darwish et al. 2000). The organism can be isolated from most internal organs in 2–3 days postexposure and disappears in about 8 days. Histopathology consists of necrosis and granulomatous inflammation of the muscle, liver, kidney, and spleen and peaks at 6 days postinfection with tissue healing being nearly complete at 8 days. Recently, E. tarda has been observed causing lesions usually associated with E. ictaluri including septicemia and the typical “hole in the head”, ulcerative lesion of the frontal portion of the head (L. Hanson unpublished observation).

Edwardsiellosis Bacillus mycoides E. tarda produces a disease in channel catfish known as Edwardsiellosis or “emphysematous putrefactive disease of catfish” (EPDC)

B. mycoides was reported to be a potential pathogen of channel catfish by Goodwin et al.

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(1994). This organism was isolated from diseased channel catfish during an epizootic in May when water temperatures were 25–27◦ C. No assessment of morbidity or mortality was given. Diseased fish had pale areas and/or open ulcers on their backs and focal necrosis of epaxial muscles. A Gram-positive bacillus was the only pathogen isolated from the fish. The isolation was facilitated by the use of tissue explants rather than an inoculating loop. B. mycoides produces a distinct whorllike growth on Muller-Hinton agar with no ¨ distinct colonies. The organism appeared as long chains of bacteria in diseased tissue and on culture plates. A similar pathological condition was produced by intramuscular injection of the organism into naive channel catfish. Intraperitoneal injection or skin abrasion failed to facilitate infection.

Management of catfish bacterial diseases Controlling bacterial diseases of channel catfish is a combination of wise management, judicious use of available chemotherapeutics, and vaccination when feasible.

Management Management of any bacterial infection of channel catfish, whether columnaris, enteric septicemia, motile Aeromonas septicemia, or other pathogens, entails proper handling, maintaining a quality environment, and reducing environmental stressors. Ideally, fish should never be handled in a weakened condition or when environmentally stressed. Water temperature control appears to be an important environmental management tool for channel catfish, particularly in tanks, raceways, and aquaria. When running water is used, flows must be sufficient to flush away metabolic waste while optimum oxygen concentrations and other water quality parameters are maintained.

Channel catfish reared in ponds that utilize artesian well water that contains up to 4,000 mg/L of NaCl seldom contract ESC (G. Whitis, Alabama Fish Farming Center, Greensboro, Alabama, personal communication). Plumb and Shoemaker (1995) exposed juvenile channel catfish, with a 10% carrier rate of E. ictaluri at 15◦ C, to water with concentrations of NaCl of near 0–3,000 mg/L at 25◦ C. Fish held in 0 and 100 mg/L NaCl suffered 100 and 96% mortality, respectively. Fish held in concentrations of 1,000, 2,000, and 3,000 mg/L NaCl had 33, 43, and 17% mortality, respectively. Based on these experimental results and personal observations, some channel catfish farmers have added low-grade salt to ponds as an ESC preventive measure. Although field trial results have not been as dramatic as experimental results, the effect of increasing salt concentrations to over 1,000 mg/L has proven beneficial for ESC control and may also be effective in preventing columnaris infections. Disadvantages of adding large amounts of salt to ponds are cost, its corrosion effect on equipment, dilution of salt if ponds are flushed, and possible environmental repercussions. There is a tendency for the aquaculturist to “do something” when ESC strikes; however, changing the feeding regime from everyday to once every few days, or complete cessation of feeding during epizootics, may be as effective in limiting mortality as applying antibiotics. Cessation of feeding or feeding medicated feed with Romet-30 every third day resulted in higher survival of E. ictaluri-infected channel catfish than in fish that received daily feeding with a normal ration (D. Wise, Delta Branch Research Station, Stoneville, Mississippi, personal communication). This procedure is practiced by many catfish farmers, and generally with satisfying results. When evaluating ESC susceptibility of production-size channel catfish, as previously noted, Kim and Lovell (1995) demonstrated that fish held over winter without receiving feed were more resistant to E. ictaluri in spring than fish that had received normal or partial feeding during winter months. Although withholding feed may

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be used in ESC, this practice may not be applied to other bacterial diseases. Shoemaker et al. (2003) demonstrated that reducing feed to fish with columnaris was detrimental to the fish, leading to increased susceptibility. Culturing blue catfish or channel × blue hybrids rather than pure channel catfish in farms where E. ictaluri is endemic is a feasible management approach (Wolters and Johnson 1994; Wolters et al. 1996). E. ictaluri is generally less pathogenic to blue catfish and channel × blue hybrids to pure channel catfish strains; however, all catfish species appear to be equally susceptible to columnaris and MAS.

Chemotherapy Bacterial infections in channel catfish vary in response to chemotherapy, but there are now four drugs or chemicals that are FDA approved for treating bacterial diseases in channel catfish (Tables 5.1, 5.3,  R and 5.4): Terramycin (oxytetracycline),  R Romet-30 (sulfadimethoxine-ormetoprim),   R R Aquaflor (Florfenicol), and Perox-Aid (35% hydrogen peroxide). There are a couple of compounds approved by the FDA for non-disease application on channel catfish that have some therapeutic value.  R Terramycin (Pfizer Inc.) incorporated into feed at 2.2 g of active oxytetracycline per kilogram of feed (approximately 1 g/lb) is fed at 2% of body weight that gives an application rate of 50–75 mg/kg of body weight per day and is fed for 10 days. A 21-day withdrawal period is required for Terramycin.  R Romet-30 (Hoffman-La Roche Inc.) is also fed at 50 mg/kg of fish per day (Plumb et al. 1987), but for only 5 days, followed by a 3day withdrawal period. These treatments are effective against ESC, and MAS infections providing treatment is initiated early. Johnson and Smith (1994) found that pellet size and formulation of Romet-30 medicated feed affected the efficacy of feeding the

drug. They determined that small pellets formulated at 5 kg of drug per ton of feed resulted in a significantly (p < 0.05) higher survival rate with the same medicated formulation that was noted in fish fed large pellets. The smaller pellets allowed feeding at 3% of body weight and resulted in fish consuming the medicated pellets. These fish gained more weight than did infected nonmedicated controls or fish fed Romet-medicated feed formulated to be fed at 1% of body weight with 15 kg Romet per ton of feed. Indiscriminate use of any antibiotics should be avoided to reduce potential for antibiotic resistance. Fish disease diagnostic laboratories have reported that over 45% of A. hydrophila isolates are resistant to Terramycin. This acquired resistance is due, in part, to many years of exposure to improper drug application. Also, plasmid-mediated R-factors are involved in antibiotic resistance of A. hydrophila (Shotts et al. 1976). It has been noted that E. ictaluri has developed a resistance to Terramycin and/or Romet-30 with as many as 10–15% of the isolates studied being resistant to one or both drugs. Waltman et al. (1989) found that E. ictaluri resistance to Romet-30 was mediated by an R plasmid, and Starliper et al. (1993) showed that the plasmid could be transferred to nonresistant isolates. Cooper et al. (1993) suggested that bacteria present in agricultural runoff are potential sources for the plasmid responsible for E. ictaluri being resistant to Romet-30. Feed containing Romet-30 enhances selection for E. ictaluri that have received the R-plasmid. While this resistance can be plasmid or genetically induced, the problem is likely exacerbated by improper use of antibiotics such as applying medicated feed when it is not necessary, improper application rates, and/or continuing treatment longer than prescribed. Aquaflor (Florfenicol) is approved for feeding all life stages of channel catfish with enteric septicemia. The drug is incorporated into manufactured feed to provide 10 mg/kg of body weight per day for 10 days with a withdrawal

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time of 12 days. The efficacy of Florfenicol at a feeding rate of 10–40 mg/kg of body weight per day for 10 days will control mortality from ESC (Gaunt et al. 2003). They also showed the drug was efficacious and palatable. It should be noted that Romet and Terramycin and Florfenicol are approved by the FDA for treating ESC. Florfenicol is also approved for columnaris and Terramycin is also approved for Pseudomonas infection. These drugs can be used to treat an unapproved disease is under extra-label use provisions with a veterinarian. Perox-Aid is approved for use as a bath on fingerling and adult channel catfish with columnaris infection. The normal dose is 50–75 mg/L (ppm) of water for 60 minutes. The compound can be used as a static treatment or as a continuous flow through application as long as the effective concentration is maintained for the duration. Perox-Aid is also effective for treating channel catfish eggs to prevent and control fungus (Saprolegnia sp.). Sensitivity of fin fish to Perox-Aid increases as water temperature rises; therefore, at higher temperatures, sensitivity of a small number of fish should be determined before the entire population is treated. Potassium permanganate (KMnO4 ) is EPA approved for treating aquaculture water for organic enrichment reduction, but is commonly used for prevention and treatment for columnaris. The chemical is applied at 2–4 mg/L in ponds indefinitely and up to 10 mg/L in tanks for up to 1 hour depending upon organic water load (Phelps et al. 1977). Tucker (1984) showed how potassium permanganate demand can be measured colorimetrically, and Lau and Plumb (1981) reported that 2 mg/L above the potassium permanganate demand was necessary to control columnaris. The most effective way to treat a pond of channel catfish infected with columnaris is to apply potassium permanganate in combination with feeding Terramycin medicated feed. Prophylactic bath treatments of 1–3% salt (NaCl) or 4–10 mg/L of potassium permanganate for 1 hour

will reduce incidence of posthandling infections. Copper sulfate (CuSO4 ) has EPA approval for treating food fish ponds for vegetation control but is not approved by FDA for treating diseased fish. Griffin and Mitchell (2007) tested the use of CuSO4 to pretreat channel catfish before exposure to E. ictaluri and found that fish treated with the chemical suffered 14.1% mortality due to E. ictaluri compared to 35.5% for fish not treated with CuSO4 prior to infection. The reason for this differential was unknown. It is essential to know the alkalinity of the water to be treated because CuSO4 is toxic in soft, low alkalinity water. Generally the FDA will not interfere with the use of CuSO4 or potassium permanganate on diseased food fish as long as the EPA guidelines are followed. Thomas-Juno and Goodwin (2004) compared several therapeutic regimes for treating columnaris in channel catfish. Oxytetracline or combination of oxytetracycline + sulphadimethoxine/ormetoprim in feed prior to bacterial infection resulted in 0% mortality. Water bath treatment with Diquat, Potassium permanganate, chloramine-T, hydrogen peroxide, and CuSO4 were also compared. Untreated fish suffered 100% mortality. Of the water applied drugs, Diquat was the most effective treatment with 0% mortality, while potassium permanganate-treated fish had 69% mortality, and chloramine-T-treated fish had 75% mortality. CuSO4 and hydrogen peroxide were not effective, indicating that these drugs have very little potential for preventing columnaris in fish. Diquat, an EPA-approved herbicide for aquaculture, has been reported to have some therapeutic value for columnaris but not FDA approved for this application. Darwish and Mitchell (2009) treated experimentally columnaris-infected channel catfish with 5.0, 10.0, and 15.0 mg/L of Diquat for 6-, 22-hour postinfection. At 21 days of posttreatment, nontreated control fish suffered 95% mortality compared to 68, 59, and 49% mortality,

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respectively, for 5.0, 10.0, and 15.0 mg/L of Diquat, indicating that there is some therapeutic efficacy for Diquat against columnaris in channel catfish; however, the herbicide is not FDA approved for disease control in fish.

Vaccination During the past 20 years, great strides have been made in developing and evaluating vaccines for cultured channel catfish, which have resulted in several now being licensed for bacterial diseases of these fish. Research has shown that E. ictaluri has several components that are immunogenic in channel catfish. Much research preceded actual development of vaccines against E. ictaluri and F. columnare, both of which are becoming popular disease management tools in channel catfish culture (Table 4.5). When injected, E. ictaluri is strongly immunogenic, but injection is impractical for immunizing large numbers of juvenile channel catfish. Vaccination studies of E. ictaluri have included use of whole-cell bacterins and cell extracts, such as LPS and purified “immunodominant” antigens as well as live attenuated preparations. The LPS is immunologically protective when injected into channel catfish and the LPS antibody is detectable in serum (Saeed and Plumb 1986; 1987). Several studies showed that fractionating E. ictaluri and using a purified immunodominant antigen can provide protection against ESC (Plumb and Klesius 1988; Vinitnantharat et al. 1993). Protein with a molecular mass of 36 kDa is a primary immunodominant antigen in the cell wall of E. ictaluri and is retained by the bacteria when subcultured up to 30 passages (Vinitnantharat et al. 1993). The bacterium is antigenic when juvenile fish are immersed in a solution containing E. ictaluri at a concentration of 1:10 of the vaccine in water with exposure for 1–2 minutes (Vinitnantharat and Plumb 1992). A killed, coated E. ictaluri vaccine applied orally was effective for increasing

survival of challenged fish (Plumb et al. 1994). Fish feed containing 1% vaccine in the feed had 77% survival compared to 23% survival of unvaccinated controls. It was also shown by Ainsworth et al. (1995) that oral application of particulate E. ictaluri antigen elicited a humoral response and resulted in antibodies in the mucus, gut washings, and gut content, but titers were not as high as in the serum. Antibody titers against E. ictaluri are easily quantified, but antibody is yet to be proven as a measure of protection. However, channel catfish that survived a natural epizootic of E. ictaluri, and with serum antibody titers greater than 1:512, were protected against subsequent experimental exposure to the pathogen (Vinitnantharat and Plumb 1993). By contrast, Klesius and Sealey (1995) showed that E. ictalurispecific antibodies were not protective in fish, but titers were less than 1:512. Klesius and Shoemaker (1997) reported that a live, attenuated E. ictaluri elicits strong immunity, and that vaccination by immersion in one E. ictaluri strain cross-protected fish against multiple pathogenic strains of E. ictaluri. Shoemaker et al. (1999) further showed conclusively that the modified, live E. ictaluri vaccine was protective to 7-day- to 31-dayold posthatch channel catfish fry when immersed in the vaccine. The research of these individuals led to the developing of a licensed  R commercial vaccine (Aquavac-ESC , Intervet Inc., USA) that is currently used successfully in the catfish industry as an ESC prevention management tool. The vaccine contains an avirulent strain of E. ictaluri, designated RE-33. Catfish should be at least 7 days posthatch when vaccinated. For maximum efficacy, fish are immersed for a minimum of 2 minutes at water temperature between 21 (70◦ F) and 29◦ C (85◦ F). It is interesting to note that research on the development of the immune system of channel catfish demonstrates that the lymphoid tissues are not organized until 21 days posthatch, and fry that were immersion exposed to wild-type E. ictaluri did not develop antibody responses or protection

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(Petrie-Hanson and Ainsworth 1999, 2001). This suggests that persistence or another mechanism allows protection to develop when fry are vaccinated with the attenuated bacterium. Vaccination of channel catfish against F. columnare showed some promise as early as 1972 (Schachte and Mora 1973) when highantibody titers were achieved in channel catfish injected with the vaccine. Moore et al. (1990) vaccinated fingerling channel catfish by immersion in a formalin killed bacterin of F. columnare that resulted in better survival and less need for chemotherapy treatment in vaccinated fish than in controls. In another vaccination study, Maas and Bootsma (1982) found that common carp will absorb F. columnare by immersion and indicated that this could be a means of vaccinating against columnaris. However, Shoemaker et al. (2007) demonstrated the efficacy of a modified, live F. columnare vaccine by immersing eyed channel catfish eggs in the vaccine followed by a booster immersion vaccination of 34-day-old fry. This research led to licensing of a live  R attenuated vaccine, Aquavac-Col (Intervet Inc., USA), to protect channel catfish against columnaris. Indications for use is to immunize at least 7-day posthatch fry, with maximum benefit by immersion for at least 2 minutes per day for 14 days prior to anticipated exposure to F. columnare. It was also shown that this modified F. columnare vaccine could be made into a bivalent product when combined with E. ictaluri Equavac-ESC vaccine. Vaccinated fingerlings were challenged 109, 116, and 137 days later with both pathogens and results suggest that the administration of a live bivalent vaccine at the eyed egg stage is immunogenic, safe, and elicits significant protection upon a single pathogen of either pathogen challenge. The vaccination application can be integrated into routine culture of channel catfish. Research to develop a vaccine to protect against A. hydrophila dates back to the middle 1950s, but the greatest obstacle is the fact that the organism is ubiquitous in the aquatic

environment. Since the early attempts little effort has gone into the development of motile Aeromonas vaccines for channel catfish, but studies have explored its application for other fish species. Researchers used heat or formalinkilled bacterins and cell extracts such as LPS with some success. Carp immunized by immersion in LPS from A. hydrophila were protected by the cell-mediated system (Baba et al. 1988). Rainbow trout fry were protected against challenge following injection of heat killed A. hydrophila (Khalifa and Post 1976) and Japanese eel were protected from the bacterium by injection with attenuated, heat or formalin-killed bacteria (Song et al. 1976). Neither of the latter studies, however, used heterologous antigen for challenge. Tiecco et al. (1988) vaccinated eel with formalin and ampicillin-inactivated A. hydrophila by injection and immersion. Both methods provided some protection, but the formalin-inactivated bacterin was not as effective as was the ampicillin preparation. While vaccination of channel catfish against bacterial infections will protect fish under normal conditions against the target pathogen, this method of disease prevention should not be perceived as absolute because poor management or stressful environmental conditions can negate protection provided by vaccines. However, if a disease for which fish have been vaccinated does develop, mortalities will be lower in vaccinated populations; therefore, a manager will need to analyze the benefit:cost ratio to determine if vaccination is feasible. Generally, benefits of vaccinated channel catfish are that they grow faster, show a lower feed conversion rate, and have significantly better survival than do unvaccinated fish.

References Ainsworth, A. J., G. Capley, et al. 1986. Use of monoclonal antibodies in the indirect fluorescent antibody technique (IFA) for the diagnosis of Edwardsiella ictaluri. Journal of Fish Diseases 9:439–444.

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Ainsworth, A. J., C. D. Rice, and L. Xue. 1995. Immune responses of channel catfish, Ictalurus punctatus (Rafinesque). After oral or intraperitoneal vaccination with particulate or soluble Edwardsiella ictaluri antigen. Journal of Fish Diseases 18:397–409. Anacker, R. L., and E. J. Ordal. 1959. Studies on the myxobacterium Chondrococcus columnaris, I. Serological typing. Journal of Bacteriology 78:25–32. Anonymous. 2005. Annual Case Summary Report. Aquatic Diagnostic Laboratory, Mississippi State University College of Veterinary Medicine, Stoneville, Mississippi, 12pp. Antonio, D. B., and R. P. Hedrick. 1994. Effects of the corticosteroid Kenalog on the carrier state of juvenile channel catfish exposed to Edwardsiella ictaluri. Journal of Aquatic Animal Health 6:44–52. Arias, C. R., C. S. Shoemaker, et al. 2003. A comparative study of Edwardsiella ictaluri parent (EILO) and E. ictaluri rifampicin-mutant (RE-33) isolates using lipopolysaccharides, outer membrane proteins, fatty acids, Biolog, API 20E and genomic analysis. Journal of Fish Diseases 26:415–421. Baba, T., J. Immura, et al. 1988. Immune protection in carp, Cyprinus carpio L., after immunization with Aeromonas hydrophila crude lipopolysaccharide. Journal of Fish Diseases 11:237– 244. Bader, J. A, K. E. Nusbaum, and Shoemaker. 2003. Comparative challenge of Flavobacterium columnare using abraded and unabraded channel catfish, Ictalurus punctatus (Rafinesque) Journal of Fish Diseases 26:461–467. Baldwin, T. J., and J. C. Newton. 1993. Early events in the pathogenesis of enteric septicemia of channel catfish caused by Edwardsiella ictaluri: light and electron microscopic and bacteriologic findings. Journal of Aquatic Animal Health 5:189–198. Baxa-Antonio, D., J. M. Groff, and R. P. Hedrick. 1992. Effect of water temperature on experimental Edwardsiella ictaluri infections in immersion exposed channel catfish. Journal of Aquatic Animal Health 4:148–151. Baxa, D. V., V. J. Groff, et al. 1990. Susceptibility of nonictalurid fishes to experimental infection with Edwardsiella ictaluri. Diseases of Aquatic Organisms 8:113–117.

Becker, C. D., and M. P. Fujihara. 1978. The bacterial pathogen Flexibacter columnaris and its epizootiology among Columbia River fish. American Fisheries Society Monograph No. 2. Bernardet, J. F. 1989. Flexibacter columnaris: first description in France and comparison with bacterial strains from other origins. Diseases of Aquatic Organisms 6:37–44. Bernardet, J-F., and P. A. D. Grimont. 1989 Deoxyribonucleicacid relatedness and phenotypic characterization of Flexibacter columnaris sp. nov., nom. rev., Flexibacter psychrophilus sp. nov., nom. rev., and Flexibacter maritimus Wakabayashi, Hikida, and Masumura 1986. International Journal of Systematic Bacteriology 39:346–354. Bernardet, J. F., P. Segers, et al. 1996. Cutting the Gordian knot: emended classification and description of the genus Flavobacterium, emended description of the family Flavobacteriaceae, and proposal of Flavobacterium hydatis nom. nov. (Basonym, Cytophaga aquatilis Strohl and Tait 1978). International Journal of Systematic Bacteriology 46:128–148. Bilodeau, A. L., G. C. Waldbieser, et al. 2003. A real-time polymerase chain reaction assay of the bacterium Edwardsiella ictaluri in channel catfish. Journal of Aquatic Animal Health 15:80–86. Boonyaratpalin, S. 1989. Bacterial pathogens involved in the epizootic ulcerative syndrome of fish in Southeast Asia. Journal of Aquatic Animal Health 1:272–276. Booth, N. J., A. Elkamel, and R. L. Thune. 2006. Intracellular replication of Edwardsiella ictaluri in channel catfish macrophages. Journal of Aquatic Animal Health 18:101–108. Borg, A. F. 1960. Studies on myxobacteria associated with diseases in salmonid fishes. Journal of Wildlife Diseases 8:1–85. Bowser, P. R. 1973. Seasonal prevalence of Chondrococcus columnaris infection in black bullheads from Clear Lake, Iowa. Journal of Wildlife Diseases 9:115–119. Buentello, J. A., and D. M. Gatlin. 2001. Effects of elevated dietary arginine of resistance of channel catfish exposure to Edwardsiella ictaluri. Journal of Aquatic Animal Health 13:194–201. Bullock, G. L., T. C. Hsu, and E. B. Shotts. 1986. Columnaris disease of fishes, United States

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resistant Escherichia coli. Journal of Aquatic Animal Health 5:9–15. Darwish, A. M., and A. J. Mitchell. 2009. Evaluation of diquat against an acute experimental infection of Flavobacterium columnare in channel catfish, Ictalurus punctatus (Rafinesque). Journal of Fish Diseases 32:401–408. Darwish, A., J. A. Plumb, and J. C. Newton. 2000. Histopathology and pathogenesis of experimental infection with Edwardsiella tarda in channel catfish. Journal of Aquatic Animal Health 12:255–266. Davidson, M. L. 1996. Characterization of the outer membrane proteins of Flavobacterium columnare and examination of their antigenicity to channel catfish (Ictalurus punctatus). Masters Thesis, Auburn University, Auburn, Alabama, USA. Davis, H. S. 1922. A new bacterial disease of freshwater fishes. U. S. Bureau of Fisheries Bulletin 38:261–280. de Figueiredo, J., and J. A. Plumb. 1977. Virulence of different isolates of Aeromonas hydrophila in channel catfish. Aquaculture 11:349–354. Earlix, D. J. 1995. Host, pathogen, and environmental interactions of enteric septicemia of catfish. Ph.D. Dissertation, Auburn, Alabama, USA, Auburn University, 102pp. Earlix, D., J. A. Plumb, and W. A. Rogers. 1996. Isolation of Edwardsiella ictaluri from channel catfish by tissue homogenization, filtration and enzyme linked immunosorbent assay. Diseases of Aquatic Organisms 27:19–24. Ewing, W. H., A. C. McWhorter, et al. 1965. Edwardsiella, a new genus of Enterobacteriaceae based on a new species, Edwardsiella tarda. International Bulletin of Bacterial Nomenclature and Taxonomy 15:33–38. Evans, J. J., P. H. Klesius, et al. 2008. Edwardsiella septicemias. Wallingford, UK, CAB International. http://www.cabicompendium.org/ac Fijan, N. N. 1968. The survival of Chondrococcus columnaris in waters of different quality. Stockholm, In Symposium II de la Commission de l’ Office Internatinal Des Epizooties Des Maladies Des Poissons. Fijan, N. N. 1969. Antibiotic additives for the isolation of Chondrococcus columnaris from fish. Applied Microbiology 17:333–334. Francis-Floyd, R., M. H. Beleau, et al. 1987. Effect of water temperature on the clinicaloutcome

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of infection with Edwardsiella ictaluri in channel catfish. Journal of the American Veterinary Medical Association 191:1413–416. Goncalves, J. R., G. Braum, et al. 1992. Aeromonas hydrophila fulminant pneumonea in a fit young man. Thorax 47:482–483. Gaunt, P., E. Endris, et al. 2003. Preliminary assessment of the tolerance and efficacy of florfenicol against Edwardsiella ictaluri administered in feed to channel catfish. Journal of Aquatic Animal Health 15:239–247. Goodwin, A. E., J. S. Roy, et al. 1994. Bacillus mycoides: A bacterial pathogen of channel catfish. Diseases of Aquatic Organisms 18:173– 178. Gray, S. J., D. J. Stickler, and T. N. Bryant. 1990. The incidence of virulence factors in mesophilic Aeromonas species isolated from farm animals and their environment. Epidemiology and Infections 105:277–294. Griffin, B. R. 1992. A simple procedure for identification of Cytophaga columnaris. Journal of Aquatic Animal Health 4:63–66. Grimont, P. A., D. F. Grimont, et al. 1980. Edwardsiella hoshinae, a new species of Enterobacteriaceae. Current Microbiology 4:347–351. Griffin, B. R., and A. J. Mitchell. 2007. Susceptibility of channel catfish, Ictalurus punctatus (Rafinesque), to Edwardsiella ictaluri following copper sulfate exposure. Journal of Fish Diseases 30:581–585. Grizzle, J. M., and Y. Kiryu. 1993. Histopathology of gill, liver, and pancreas, and serum enzyme levels of channel catfish infected with Aeromonas hydrophila complex. Journal of Aquatic Animal Health 5:36–50. Groberg, W. J., Jr., R. H. McCoy, et al. 1978. Relation of water temperature to infections of coho salmon (Oncorhynchus kisutch), chinook salmon (O. tshawytscha), and steelhead trout (Salmo gairdneri) with Aeromonas salmonicida and A. hydrophila. Journal of Fisheries Research Board Canada 35:1–7. Hanson, L. A., and J. M. Grizzle. 1985. Nitriteinduced predisposition of channel catfish to bacterial diseases. The Progressive Fish-Culturist 47:98–101. Hargraves, J. E., and D. R. Lucey. 1990. Edwardsiella tarda soft tissue infection associated with catfish puncture wound. Journal of Infectious Diseases 162:1416.

Hawke, J. P. 1979. A bacterium associated with disease of pond cultured channel catfish. Journal of the Fisheries Research Board of Canada 36:1508–1512. Hawke, J. P., A. C. McWhorter, et al. 1981. Edwardsiella ictaluri sp. nov., the causative agent of enteric septicemia of catfish. International Journal of Systematic Bacteriology 31:396–400. Hawke, J. P., and R. L. Thune. 1992. Systemic isolation and antimicrobial susceptibility of Cytophaga columnaris from commercially reared channel catfish. Journal of Aquatic Animal Health 4:109–113. Hazen, T. C., C. B. Fliermans, et al. 1978. Prevalence and distribution of Aeromonas hydrophila in the United States. Applied Environmental Microbiology 36:731–738. Humphrey, J. D., C. Lancaster, et al. 1986. Exotic bacterial pathogens Edwardsiella tarda and Edwardsiella ictaluri from imported ornamental fish Betta splendens and Puntius conchonius respectively; isolation and quarantine significance. Australian Veterinary Journal 63:363–369. Hussain, M., and R. C. Summerfelt. 1991. The role of mechanical injury in an experimental transmission of Flexibacter columnaris to fingerling walleye. Journal of the Iowa Academy of Science 98:93–98. Janda, J. M. 1991. Recent advances in the study of the taxonomy, pathogenicity, and infectious syndromes associated with the genus Aeromonas. Clinical Microbiology Review 4:397–410. Johnson, M. R., and K. L. Smith. 1994. Effect of pellet size and drug concentration on the efficacy of Romet-medicated feed for controlling Edwardsiella ictaluri infections in channel catfish fingerlings. Journal of Aquatic Animal Health 6:53–58. Kage, T., R. Takahashi, et al. 1992. Aeromonas hydrophila, a causative agent of mass mortality in cultured Japanese catfish larvae (Silurus asotus). Fish Pathology 27:57–62. Kasornchandra, J., W. A. Rogers, and J. A. Plumb. 1987. Edwardsiella ictaluri from walking catfish, Clarias batrachus L., in Thailand. Journal of Fish Disease 10:137–138. Kent, M. L., and J. M. Lyons. 1982. Edwardsiella ictaluri in the green knife fish, Eigemannia virescens. Fish Health News 2:ii. Ketover, B. P., L. S. Young, and D. Armstrong. 1973. Septicemia due to Aeromonas hydrophila;

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clinical and immunological aspects. Journal of Infectious Diseases 127:284–290. Khalifa, K. A., and G. Post. 1976. Immune response of advanced rainbow trout fry to Aeromonas liquefations. The Progressive FishCulturist 38:66–68. Kim, M. K., and R. T. Lovell. 1995. Effect of overwinter feeding regimen on body weight, body composition and resistance to Edwardsiella ictaluri in channel catfish, Ictalurus punctatus. Aquaculture 134:237–246. King, G. E., S. B. Werner, and W. Rizer. 1992. Epidemiology of aeromonas infections in California. Clinical Infectious Diseases 15:449–452. Klesius, P. 1992. Carrier state of Edwardsiella ictaluri in channel catfish, Ictalurus punctatus. Journal of Aquatic Animal Health 4:227– 230. Klesius, P. H. 1993. Rapid enzyme-linked immunosorbent tests for detecting antibodies to Edwardsiella ictaluri in channel catfish, Ictalurus punctatus, using exoantigen. Veterinary Immunology and Immunopathology 36:359– 368. Klesius, P. 1994. Transmission of Edwardsiella ictaluri from infected, dead to noninfected channel catfish. Journal of Aquatic Animal Health 6:180–182. Klesius, P., K. Johnson, et al. 1991. Development and evaluation of an enzyme-linked immunosorbent assay for catfish serum antibody to Edwardsiella ictaluri. Journal of Aquatic Animal Health 3:94–99. Klesius, P., and W. M. Sealey. 1995. Characteristics of serum antibody in enteric septicemia of catfish. Journal of Aquatic Animal Health 7:205–210. Klesius, P. H., and C. A. Shoemaker. 1997. Heterologous isolates challenge of channel catfish, Ictalurus punctatus, immune to Edwardsiella ictaluri. Aquaculture 157:147–155. Klesius, P., J. Lovy, et al. 2003. Isolation of Edwardsiella ictaluri from tadpole madtom in southwestern New Jersey River. Journal of Aquatic Animal Health 15:295–301. Kuo, S-C., H-Y. Chung, and G.-H. 1981. Studies on artificial infection of the gliding bacteria in cultured fishes. Fish Pathology 15:309– 314. Lau, K. J., and J. A. Plumb. 1981. Effects of organic load on potassium permanganate as a treatment

for Flexibacter columnaris. Transactions of the American Fisheries Society 110:86–89. Levanon, N., B. Motro, et al. 1986. The dynamics of Aeromonas hydrophila in the water of tanks used to nurse elvers of the European eel Anguilla anguilla. Bamidgeh 38:55–63. Lobb, C. J., and M. Rhoades. 1987. Rapid plasmid analysis for identification of Edwardsiella ictaluri from infected channel catfish Ictalurus punctatus. Applied Environmental Microbiology 53:1267–1272. Lumlertdacha, S., and R. T. Lovell. 1995. Fumonisin-contaminated dietary corn-reduced survival and antibody production by channel catfish challenged with Edwardsiella ictaluri. Journal of Aquatic Animal Health 7:1–8. Maas, M. G., and R. Bootsma. 1982. Uptake of bacterial antigens in the spleen of carp (Cyprinus carpio L). Developments in Comparative Immunology Supl. 2:47. Marks, J. E., D. H. Lewis, and C. S. Trevino. 1980. Mixed infection in columnaris disease of fish. Journal of the American Veterinary Medical Association 177:811–814. Meyer, F. P. 1970. Seasonal fluctuations in the incidence of diseases on fish farms. In A Symposium on Diseases of Fishes and Shellfishes edited by S. F. Snieszko, 21–29. American Fisheries Society Special Publication No. 5. Meyer, F. P., and G. L. Bullock. 1973. Edwardsiella tarda, a new pathogen of channel catfish (Ictalurus punctatus). Applied Microbiology 25:155–156. Mgolomba, T. N., and J. A. Plumb. 1992. Longevity of Edwardsiella ictaluri in the organs of experimentally infected channel catfish, Ictalurus punctatus. Aquaculture 101:1–6. Mitchell, A. J., and A. E. Goodwin. 1999. Evidence that enteric septicemia of catfish (ESC) was present in Arkansas by the late 1960’s: New insights into the epidemiology of ESC. Journal of Aquatic Animal Health 11:157–178. Miyazaki, T., and J. A. Plumb. 1985. Histopathology of Edwardsiella ictaluri in channel catfish, Ictalurus punctatus (Rafinesque). Journal of Fish Diseases 8:389–392. Miyazaki, T., and Y. Jo. 1985. A histopathological study of motile aeromonad disease in ayu. Fish Pathology 20:55–59. Moore, A. A., M. E. Eimers, et al. 1990. Attempts to control Flexibacter columnaris

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epizootics in pond-reared channel catfish by vaccination. Journal of Aquatic Animal Health 2:109–111. Morrison, E., and J. A. Plumb. 1992. The chemosensory system of channel catfish, Ictalurus punctatus, following immersion exposure to Edwardsiella ictaluri. American Chemosensory Society, Orlando, Florida, April 8–10, 1992 (Abstract). Morrison, E. E., and J. A. Plumb. 1994. Olfactory organ of channel catfish as a site of experimental Edwardsiella ictaluri infection. Journal of Aquatic Animal Health 6:101–109. Newton, J. C., R. C. Bird, et al. 1988. Isolation, characterization, and molecular cloning of cryptic plasmids isolated from Edwardsiella ictaluri. American Journal of Veterinary Research 49:1856–1860. Newton J. C., L. G. Wolfe, et al. 1989. Pathology of experimental enteric septicemia in channel catfish Ictalurus punctatus Rafinesque following immersion exposure to Edwardsiella ictaluri. Journal of Fish Diseases 12:335–348. Nielsen, M. E., L. Høi, et al. 2001. Is Aeromonas hydrophila the dominant motile Aeromonas species that causes disease outbreaks in aquaculture production in the Zheijiang province of China? Diseases of Aquatic Organisms 46:23–29. Nieto, T. P., M. J. R. Corcobado, et al. 1985. Relation of water temperature to infection of Salmo gairdneri with motile Aeromonas. Fish Pathology 20:99–105. Nusbaum, K. E., and E. E. Morrison. 1996. Entry of 35 S-labeled Edwardsiella ictaluri into channel catfish. Journal of Aquatic Animal Health 8:146–149. Nusbaum, K. E., and E. E. Morrison. 2002. Edwardsiella ictaluri bacteraemia elicits shedding of Aeromonas hydrophila complex in latently infected channel catfish, Ictalurus punctatus (Rafinesque). Journal of Fish Diseases 25:343–350. Olivares-Fuster, O., J. L. Baker, et al. 2007. Host specific association between Flavobacterium columnare genomvars and fish species. Systematic and Applied Microbiology 30:624–633. Ordal, E. J., and R. R. Rucker. 1944. Pathogenic myxobacteria. Society of Experimental Biology and Medicine Proceedings 56:15–18. Panangala, V. S., R. A. Shelby, et al. 2006a. Immunofluorescent test for simultaneous detec-

tion of Edwardsiella ictaluri and Flavobacterium columnare. Diseases of Aquatic Organisms 68:197–207. Panangala, V. S., C. A. Shoemaker, et al. 2006b. Intra- and interspecific phenotypic characteristics of fish-pathogenic Edwardsiella ictaluri and E. tarda. Aquaculture Research 37:49–60. Panangala, V. S., V. L. Van Santen, et al. 2005. Analysis of 16S-23S intergenic spacer regions of the rRNA operons in Edwardsiella ictaluri and Edwardsiella tarda isolates form fish. Journal of Applied Microbiology 99:657–669. Panangala, V. S., C. A. Shoemaker, et al. 2007. Multiplex-PCR for simultaneous detection of 3 bacterial fish pathogens, Flavobacterium columnare, Edwardsiella ictaluri, and Aeromonas hydrophila. Diseases of Aquatic Organisms 74:199–208. Paripatananont, T., and R. T. Lovell. 1995. Responses of channel catfish fed organic and inorganic sources of zinc to Edwardsiella ictaluri challenge. Journal of Aquatic Animal Health 7:147–154. Peters, G., M. Faisal, et al. 1988. Stress caused by social interaction and its effect on susceptibility to Aeromonas hydrophila infection in rainbow trout Salmo gairdneri. Diseases of Aquatic Organisms 4:83–89. Petrie-Hanson, L., and A. J. Ainsworth. 1999. Humoral immune responses of channel catfish (Ictalurus punctatus) fry and fingerlings exposed to Edwardsiella ictaluri. Fish and Shellfish Immunology 9:579–589. Petrie-Hanson, L., and A. J. Ainsworth. 2001. Ontogeny of channel catfish lymphoid organs. Veterinary Immunology and Immunopathology 81:113–27. Phelps, R. P., J. A. Plumb, and H. C. Harris. 1977. Control of external bacterial infections of bluegills with potassium permanganate. The Progressive Fish-Culturist 39:142–143. Plumb, J. A., J. M. Grizzle, and J. DeFiqueiredo. 1976. Necrosis and bacterial infection in channel catfish (Ictalurus punctatus) following hypoxia. Journal of Wildlife Diseases 12:247–253. Plumb, J. A., and V. Hilge. 1987. Susceptibility of European catfish (Silurus glanis) to Edwardsiella ictaluri. Journal of Applied Ichthyology 3:45–48. Plumb, J. A., and P. Klesius. 1988. An assessment of the antigenic homogeneity of Edwardsiella

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ictaluri using monoclonal antibody. Journal of Fish Diseases 11:499–510. Plumb, J. A., G. Maestrone, and E. Quinlan. 1987. Use of a potentiated sulfonamide to control Edwardsiella ictaluri infection in channel catfish (Ictalurus punctatus). Aquaculture 62:187–194. Plumb, J. A., and E. E. Quinlan. 1986. Survival of Edwardsiella ictaluri in pond water and bottom mud. The Progressive Fish-Culturist 48:212–214. Plumb, J. A., and D. J. Sanchez. 1983. Susceptibility of 5 species of fish to Edwardsiella ictaluri. Journal of Fish Diseases 6:261–266. Plumb, J. A., and C. Shoemaker. 1995. Effects of temperature and salt concentration on latent Edwardsiella ictaluri infections in channel catfish. Diseases of Aquatic Organisms 21:171– 175. Plumb, J. A., and S. Vinitnantharat. 1989. Biochemical biophysical, and serological homogeneity of Edwardsiella ictaluri. Journal of Aquatic Animal Health 1:51–56. Plumb, J. A., S. Vinitnantharat, and W. D. Paterson. 1994. Optimum concentration of Edwardsiella ictaluri vaccine in feed for oral vaccination of channel catfish. Journal of Aquatic Animal Health 6:118–121. Pyle, S. W., and E. B. Shotts, Jr. 1981. DNA homology studies of selected flexibacteria associated with fish diseases. Canadian Journal of Fisheries and Aquatic Sciences 38:146–151. Roberts, R. J., L. G. Willoughby, et al. 1993. Mycotic aspects of epizootic ulcerative syndrome (EUS) of Asian fishes. Journal of Fish Diseases 16:169–183. Rogers, W. A. 1981. Serological detection of two species of Edwardsiella infecting catfish. Developments in Biological Standards 49:169–172. Saeed, M. O., and J. A. Plumb. 1986. Immune response of channel catfish to lipopolysaccharide and whole cell Edwardsiella ictaluri vaccines. Diseases of Aquatic Organisms 2:21–25. Saeed, M. O., and J. A. Plumb. 1987. Serological detection of Edwardsiella ictaluri Hawke lipopolysaccharide antibody in serum of channel catfish, Ictalurus punctatus Rafinesque. Journal of Fish Diseases 10:205–209. Santos, Y., A. E. Toranzo, et al. 1987. Relationships among virulence for fish, enterotoxigenicity, and phenotypic characteristics of motile Aeromonas. Aquaculture 67:29–39.

Schachte, J. H., Jr., and E. C. Mora. 1973. Production of agglutinating antibodies in the channel catfish (Ictalurus punctatus) against Chondrococcus columnaris. Journal of the Fisheries Research Board of Canada 30:116–118. Shamsudin, M. N., and J. A. Plumb. 1996. Morphological, biochemical, and physiological characterization of Flexibacter columnaris isolates from four species of fish. Journal of Aquatic Animal Health 8:335–339. Shieh, H. S. 1980. Studies on the nutrition of a fish pathogen, Flexibacter columnaris. Microbiological Letters 13:129–133. Shoemaker, C. A., P. H. Klesius, and J. L. Bricker. 1999. Efficacy of a modified live Edwardsiella ictaluri vaccine in channel catfish as young as seven days post hatch. Aquaculture 176:189– 193. Shoemaker, C. A., C. R. Arias, et al. 2005. Technique for identifying Flavobacterium columnare using whole cell fatty acid profiles. Journal of Aquatic Animal Health 17:267–274. Shoemaker, C. A., P. H. Klesius, and J. J. Evans. 2007. Immunization of eyed channel catfish, Ictalurus punctatus, eggs with monovalent Flavobacterium columnare vaccine and bivalent F. columnare and Edwardsiella ictaluri vaccine. Vaccine 25:1126–1131. Shoemaker, C. A., P. H. Klesius, et al. 2003. Feed deprivation of channel catfish, Ictalurus punctatus (Rafinesque), influences organosomatic indices, chemical composition and susceptibility to Flavobacterium columnare. Journal of Fish Diseases 26:553–561. Shoemaker C. A., O. Olivares-Fuster, et al. 2008. Flavobacterium columnare genomovar influences mortality in channel catfish (Ictalurus punctatus). Veterinary Microbiology 127:353–359. Shotts, E. B. 1991. Selective isolation methods for fish pathogens. Journal of Applied Bacteriology, Symposium Supplement 70:75S–80s. Shotts, E. B., V. S. Blazer, and W. D. Waltman. 1986. Pathogenesis of experimental Edwardsiella ictaluri infections in channel catfish (Ictalurus punctatus). Canadian Journal of Fisheries and Aquatic Sciences 43:36–42. Shotts, E. B., J. L. Gaines, et al. 1972. Aeromonasinduced deaths among fish and reptiles in an eutrophic inland lake Journal American Veterinary Medicine Association 161:603–607.

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Shotts, E. B., and R. Rimler. 1973. Medium for the isolation of Aeromonas hydrophila. Applied Microbiology 26:550–553. Shotts, E. B., V. L. Vanderwork, et al. 1976. Occurrence of R factors associated with Aeromonas hydrophila isolated from aquarium fish and waters. Journal of Fisheries Research Board of Canada 33:736–740. Shotts, E. B., and W. D. Waltman. 1990. An isolation medium for Edwardsiella ictaluri. Journal Wildlife Diseases 26:214–218. Sink, T. D., and R. J. Strange. 2004. Linking stress to increased susceptibility of channel catfish to enteric septicemia using cortisol. Journal of Aquatic Animal Health 16:93–98. Song, Y-L., S-N. Chen, and G. Kou. 1976. Agglutinating antibodies production and protection in eel (Anguilla japonica) inoculated with Aeromonas hydrophila (A. liquefaciens) antigens. Journal of Fisheries Society of Taiwan 4:25–29. Song, Y-L., J. L. Fryer, and J. S. Rohovec. 1988a. Comparison of six media for the cultivation of Flexibacter columnaris. Fish Pathology 23:91–94. Song, Y-L., J. L. Fryer, J. S. Rohovec. 1988b. Comparison of gliding bacteria isolated from fish in North America and other areas of the Pacific rim. Fish Pathology 23:197–202. Speyerer, P. D., and J. A. Boyle. 1987. The plasmid profile of Edwardsiella ictaluri. Journal of Fish Diseases 10:461–469. Stanley, L. A., J. S. Hudson, et al. 1994. Extracellular products associated with virulent and avirulent strains of Edwardsiella ictaluri from channel catfish. Journal of Aquatic Animal Health 7:141–146. Starliper, C. E., R. K. Cooper, et al. 1993. Plasmidmediated Romet resistance of Edwardsiella ictaluri. Journal of Aquatic Animal Health 5:1–8. Taylor, P. 1992. Fish-eating birds as potential vectors for Edwardsiella ictaluri. Journal of Aquatic Animal Health 4:240–243. Thomas-Juno, S., and A. E. Goodwin. 2004. Acute columnaris infection in channel catfish, Ictalurus punctatus (Rafinesque): efficacy of practical treatments for warmwater aquaculture ponds. Journal of Fish Diseases 27:23–28. Thune, R. L. 1991. Major infectious and parasitic diseases of channel catfish. Veterinary and Human Toxicology 33 (Suppl. 1): 14–18.

Tiecco, G., C. Sebastio, et al. 1988. Vaccination trials against “red plaque” in eels. Diseases of Aquatic Organisms 4:105–107. Triyanto, A, and H. Wakabayashi. 1999. Genotypic diversity of strains of Flavobacterium columnare from diseased fish. Fish Pathology 34:65–71. Tyler, J. W., and P. H. Klesius. 1994. Decreased resistance to Edwardsiella ictaluri infection in channel catfish intraperitoneally administered an oil adjuvant. Journal of Aquatic Animal Health 6:275–278. Tucker, C. S. 1984. Potassium permanganate demand of pond waters. The Progressive FishCulturist 46:24–28. Ventura, M. T., and J. M. Grizzle. 1988. Lesions associated with natural and experimental infections of Aeromonas hydrophila in channel catfish, Ictalurus Punctatus (Rafinesque). Journal of Fish Diseases 11:397–407. Vinitnantharat, S., and J. A. Plumb. 1992. Kinetics of the immune response of channel catfish to Edwardsiella ictaluri. Journal of Aquatic Animal Health 4:207–214. Vinitnantharat, S., and J. A. Plumb. 1993. Protection of channel catfish following exposure to Edwardsiella ictaluri and effects of feeding antigen on antibody titer. Diseases of Aquatic Organisms 15:31–34. Vinitnantharat, S., J. A. Plumb, and A. E. Brown. 1993. Isolation and purification of an outer membrane protein of Edwardsiella ictaluri and its antigenicity to channel catfish (Ictalurus punctatus). Fish & Shellfish Immunology 3:401–409. Wagner, B. A., D. J. Wise, et al. 2002. The epidemiology of bacterial diseases in food-size channel catfish. Journal of Aquatic Animal Health 14:263–272. Wahil, T., S. E. Burr, et al. 2005. Aeromonas sobria, a causative agent of disease in farmed perch, Perca fluviatillis L. Journal of Fish Diseases 28:141–150. Wakabayashi, J., K. Ganai, et al. 1980. Pathogenic activities of Aeromonas biovar hydrophila (Chester) Popoff and Vernon. 1976 to fishes. Fish Pathology 15:319–325. Wakabayashi, H., K. Kira, and S. Egusa. 1970. Studies on columnaris of eels – I. Characteristics and pathogenicity of Chondrococcus columnaris isolated from pond-cultured eels. Bulletin of the Japanese Society of Scientific Fisheries 36:147–155.

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Walters, G. R., and J. A. Plumb. 1980. Environmental stress and bacterial infection in channel catfish, Ictalurus punctatus Rafinesque. Journal Fish Biology 17:177–185. Waltman, W. D., E. B. Shotts, and V. S. Blazer. 1985. Recovery of Edwardsiella ictaluri from Danio (Danio devario). Aquaculture 46:63–66. Waltman, W. D., E. B. Shotts, and T. C. Hsu. 1986. Biochemical characteristics of Edwardsiella ictaluri. Applied and Environmental Microbiology 51:101–104. Waltman, W. D., E. B. Shotts, and R. E. Wooley. 1989. Development and transfer of plasmidmediated antimicrobial resistance in Edwardsiella ictaluri. Canadian Journal of Fisheries and Aquatic Sciences 46:1114–1117. Waterstrat, P. R., B. Dorr, et al. 1999. Recovery and viability of Edwardsiella ictaluri from great blue herons Ardea herodias fed E. ictaluriinfected channel catfish Ictalurus punctatus fingerlings. Journal of the World Aquaculture Society 30:115–122. Weete, J. D., W. T. Blevins, et al. 1988. Chemical characterization of lipopolysaccharide from Edwardsiella ictaluri, a fish pathogen. Canadian Journal of Microbiology 34:1224–1229. Welker, T. L., C. A. Shoemaker, et al. 2005. Transmission and detection of Flavobacterium columnare in channel catfish Ictalurus punctatus. Diseases of Aquatic Organisms 63:129–138. Williams, M. L., P. Azadi, and M. L. Lawrence. 2003. Comparison of cellular extracellular products expressed by virulent and attenuated strains of Edwardsiella ictaluri. Journal of Aquatic Animal Health 15:264–273.

Willoughby, L. G., R. J. Roberts, and S. Chinabut. 1995. Aphanomyces invaderis sp. Nov., the fungal pathogen of freshwater tropical fish affected by epizootic ulcerative syndrome. Journal of Fish Diseases 18:273–275. Wise, D. J., T. E. Schwedler, and D. L. Otis. 1993a. Effects of stress on susceptibility of naive channel catfish in immersion challenge with Edwardsiella ictaluri. Journal of Aquatic Animal Health 5:92–97. Wise, D. J., J. R. Tomasso, et al. 1993b. Effects of vitamin E on the immune response of channel catfish to Edwardsiella ictaluri. Journal of Aquatic Animal Health 5:183–188. Wolfe, K., J. A. Plumb, and E. E. Morrison. 1998. Lectin binding characteristics of the olfactory mucosa of channel catfish: potential factors in attachment of Edwardsiella ictaluri. Journal of Aquatic Animal Health 10:348– 360. Wolters, W. R., and M. R. Johnson. 1994. Enteric septicemia resistance in blue catfish and three channel catfish strains. Journal of Aquatic Animal Health 6:329–334. Wolters, W. R., D. J. Wise, and P. H. Klesius. 1996. Survival and antibody response of channel catfish, blue catfish, and channel catfish female x blue catfish male hybrids after exposure to Edwardsiella ictaluri. Journal of Aquatic Animal Health 8:249–254. Zhang, Y., C. R. Arias, et al. 2006. Comparison of lipopolysaccharide and protein profiles between Flavobacterium columnare strains from different genomovars. Journal of Fish Diseases 29:657–663.

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Chapter 12

Carp and minnow bacterial diseases

Carp and goldfish culture (family Cyprinidae) is probably the oldest known type of aquaculture. Common carp have been cultured in Europe since the middle ages and the Chinese have grown carp and goldfish in captivity for 2,500 years. Currently golden shiners and fathead minnows are cultured extensively in the United States for bait fish, while goldfish and koi carp are popularly cultured for aquaria and ornamental ponds. Two bacterial diseases most often associated with cyprinids are “carp erythrodermatitis” (CE) (Fijan 1972) and “ulcer disease of goldfish” (UDG) (Elliott and Shotts 1980). Ironically, both disease syndromes are caused by the same organism, “atypical” Aeromonas salmonicida, also known as Aeromonas salmonicida subspecies achromogens and “atypical nonmotile Aeromonas.” The following discussion includes CE, UDG, and other bacterial pathogens that cause disease in cyprinids that include motile Aeromonas septicemia (MAS), Pseudomonas spp., and columnaris (See Chapter 11).

Atypical nonmotile aeromonas infections Atypical Aeromonas salmonicida was recognized as a fish pathogen by Smith (1963) in Great Britain. Paterson et al. (1980) later showed that the organism was identical to Haemophilus piscium, the etiological agent of “ulcer disease” of trout. Since the early 1970s, atypical A. salmonicida has been a major disease organism in cultured cyprinids where it produces a subacute to chronic infection. It has also been associated with similar diseases in increasing numbers of other fish species. CE was originally considered a stage of infectious dropsy of carp (IDC); however, CE is now recognized as a distinct disease (Fijan 1972). The precise etiology of CE was not determined until Bootsma et al. (1977) reported a nonmotile “atypical” A. salmonicida (subspecies achromogens) from carp at five fish farms in Yugoslavia and Germany, at which time Koch’s postulates were fulfilled. Goldfish ulcer disease followed a similar scenario until Shotts et al. (1980) solved the etiological problem.

Geographical range and species susceptibility 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.

Atypical A. salmonicida infections (CE and UDG) have been documented in most 315

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European countries, Iceland, Great Britain, the United States, Canada, Japan, Israel, several Asian countries, and Australia (Fijan 1972; Fijan and Petrinec 1973; McCarthy 1975; Bootsma et al. 1977; Paterson et al. 1980; Gudmandsotter 1988). Cultured carp with ´ clinical signs typical of CE have also been reported in China. CE per se has not been reported in North America because the syndrome is referred to as ulcerative disease of goldfish (Elliott and Shotts 1980; Shotts et al. 1980). Common carp, including scaled and mirror varieties, koi carp, and goldfish are the principal species affected by atypical A. salmonicida, but similar types of lesions have been described in a variety of other freshwater and marine fish species. Other notable susceptible freshwater fish are northern pike (Wiklund 1990), eels, roach, silver bream, common bream, perch, and common minnow. Several species of salmonids, including Atlantic, chum, and coho salmon and rainbow, brook, and brown trout (Evelyn 1971; Wichardt 1983) are also susceptible. Marine fish that have been either naturally or experimentally infected with atypical A. salmonicida include European flounder (Wiklund and Byland 1991), dab, plaice (Wiklund and Dalsgaard 1995), turbot (Pedersen et al. 1994), and Atlantic tomcod (Williams et al. 1997). Most of these infections have occurred, in either wild, captured, or cultured fishes in the North Sea-Baltic Sea area of Scandanavia or the western North Atlantic Ocean.

Clinical signs Clinical signs of atypical A. salmonicida infections are similar in most fish (Fijan 1976; Elliott and Shotts 1980). Moribund CEinfected fish rest near the surface, at the bottom, or close to pond banks, or concentrate near fresh water inflow; a behavioral pattern that has not been described for UDG. Infected fish become darkly pigmented with slightly ex-

tended abdomens and slight to extreme exophthalmia. Deep ulcers in the skin and muscle tissue are the most obvious clinical signs of CE and UDG (Figure 12.1). These lesions begin as small to large hemorrhages in the skin and scale pockets and progress to extensive necrosis in the superficial epithelium and into the musculature. Hemorrhagic inflammation also occurs on the fins. In other fish species, lesions associated with atypical A. salmonicida are generally very similar to those described for CE and UDG (McCarthy 1975; Hubert and Williams 1980). However, Iida et al. (1984) found that the most common sign of disease associated with atypical A. salmonicida in eels was a swollen head and presence of ulcerative lesions on the jaws and cheeks. Affected goldfish have pronounced anemia, exhibit body edema along with lepidorthosis, and proteinacious material may collect on lesions giving the appearance of a fungal infection (Figure 12.1). Internal lesions are often lacking but when present include hemorrhagic appearance, pale liver, and inflamed intestine. Atlantic salmon (30–40 g) infected with atypical A. salmonicida exhibited clinical signs that closely resembled those of furunculosis of salmonids (Paterson et al. 1980). Clinical signs displayed by diseased turbot in saltwater tanks in Denmark were lethargy and necrotic lesions, which usually started as erosion on the tips of skin nodules (Pedersen et al. 1994). These nodules contained white centers surrounded by a narrow hemorrhagic zone. Some erosion progressed into ulcers of 0.5–3 cm in diameter, which were distributed over the body. Similar signs were described in atypical A. salmonicida infected European flounder, dab, and plaice (Wiklund and Dalsgaard 1995).

Diagnosis The first step in diagnosing atypical A. salmonicida infections is recognizing clinical signs followed by necropsy and bacterial

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(a)

(b)

(c)

Figure 12.1 Diseases caused by atypical Aeromonas salmonicida. (a) Ulcerative lesion (arrow) on the skin of common carp with carp erythrodermatitis (Photo courtesy of N. Fijan). (b) Ulcerative lesion (arrow) on the skin of goldfish with ulcer disease of goldfish. (c) Ulcerative lesion (arrow) on lateral surface of Atlantic tomcod. (Photo courtesy of S. Bastien-Daigle and P. J. Williams and with permission of the American Fisheries Society.)

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isolation. The pathogen can be isolated from necrotic muscle or hemorrhagic skin lesions, but seldom from the visceral organs of carp or goldfish. Contrastingly, the pathogen is often isolated from visceral organs of other infected fish species. For example, Cornick et al. (1984) isolated atypical A. salmonicida from kidneys of 22% of Atlantic cod with skin lesions, and Hubert and Williams (1980) working with roach isolated the causative organism from 95% of skin ulcers and 67% and 24% of blood and kidney samples, respectively. Wiklund and Dahlsgaard (1995) isolated the organism from skin ulcers of 73% of affected fish and were unable to isolate it from internal organs of European flounder, dab, or plaice. Bootsma et al. (1977) proposed that isolation media be supplemented with tryptone and/or serum, but Paterson et al. (1980) found that blood agar produced satisfactory isolation results. More recently, Tinman and Maurice (2009) improved isolation efficiency of the pathogen by including 0.01% Coomassie Brilliant BlueR-250 to TSA. Atypical A. salmonicida is fastidious, especially upon primary isolation, but growth capability improves upon repeated subcultivation. Upon isolation, slow growing punctate, nonpigmented, cream-colored colonies appear after 24–72 hours when incubated at 20◦ C. Muscle lesions are often contaminated with other, less fastidious bacteria (A. hydrophila or Listonella (Vibrio) anguillarum) that grow on media more rapidly than does atypical A. salmonicida.

Bacterial characteristics Atypical A. salmonicida is a short Gramnegative, nonmotile bacillus, but in contrast to typical A. salmonicida salmonicida, produces no diffused brown pigment below 25◦ C (Shotts et al. 1980). The pathogen may be either cytochrome oxidase positive, as is the case in isolates from carp and goldfish, or cytochrome oxidase-negative as in isolates from coho salmon (Chapman et al. 1991), European

flounder, dab, plaice, and turbot (Pedersen et al. 1994). Atypical A. salmonicida cells are about 0.5 µm wide and 0.7–1.4 µm long. Optimum growth temperature is 27◦ C with no growth taking place at 37◦ C. As suggested by Pedersen et al. (1994) and Wiklund and Dalsgaard (1995), it appears that atypical A. salmonicida isolated in the North Sea-Baltic Sea region from European flounder, turbot, etc. forms a distinct group from isolates of carp, goldfish, and other species (Shotts et al. 1980). Major differences in the isolates include fastidiousness, cytochrome oxidase reaction, degradation of casein, esculin hydrolysis, and susceptibility to ampicillin and cephalothin. All atypical A. salmonicida isolates are less biochemically reactive than other bacterial isolates from fish (Table 12.1). Restriction endonuclease analysis of atypical A. salmonicida isolates in Australia indicates there are distinctly different strains of the organism (Whittington et al. 1995). They showed that an isolate from silver perch was the same as that from Australian goldfish on the same farm and similar to a goldfish isolate in the United States, but differed from an isolate of goldfish origin in Singapore. However, an isolate from farmed greenback flounder in Australia was a completely different strain and was probably brought to the farm by wild-caught marine fish. Pulsed-field gel electrophoresis (PFGE) analysis with XbaI restriction enzyme was used to study the genetic heterogeneity of 88 atypical A. salmonicida isolates from 17 fish species in Finland, Iceland, Norway, Sweden, and Denmark (Hanninen and Hirvela-Koski ¨ ¨ 1999). Analysis of Norwegian and Icelandic strains had the same ribotype and their PFGE patterns suggested a genetic relatedness. Moreover, atypical A. salmonicida isolates from the same Finish fish farm over time had closely related PFGE patterns suggesting genetic stability. It was also suggested by these authors that PFGE is a distinguishing method to study genetic heterogeneity of atypical A. salmonicida and is applicable to epidemiological studies.

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Table 12.1 Biochemical and biophysical characteristics of nonmotile aeromonads (typical Aeromonas salmonicida salmonicida, atypical A. salmonicida (subsp.achromogens) and atypical A. salmonicida from North Sea and Baltic Sea).

Characteristic/Reaction∗

Cell morphology Agglutinates antitypical A. salmonicida serum Motility Cytochrome oxidase Oxidation/fermentation glucose Gas from glucose Growth in or at 0% NaCl 3% NaCl 4% NaCl 5◦ C 30◦ C 37◦ C Degredation of Casein Esculin Gelatin Starch Voges Prauskaur ß-galactosidase Indole Susceptibility to Ampicilin (33 µg) Cephalothin (66 µg) Acid from Glycerol L-arabinose Ribose Galactose Fructose D-mannose Mannitol N-acetylglucosamine Arbutine Salicine Cellobiose Maltose Saccharose Trehalose Glycogen

Typical A. salmonicida

+ − + +/+ +

Atypical A. salmonicida

short, Gram-negative rods + − + +/+ −

North, Baltic Sea A. salmonicida

+ − + +/+ −

+ + + + + −

+ + + + + −

+ + +(V)† + + −

+ + + + − − −

+ − − + + − +

+ + + + + +(V) −

S S

R R

S S

+ + + + + − + + + + − + − − +

+ − + + + + + − − − − − + + +

− − −(V) −(V) + V − −(V) − V −(V) −(V) V +(V) −(V)

Source: Kimura (1969), McCarthy (1975), Munroe and H˚astings (1993), Wicklund and Dalsgaard (1995). +, positive; −, negative; V, most strains variable; R, resistant; S, susceptible; (V), variable.

Epizootiology The epizootiology of atypical A. salmonicida is similar in most fish species, especially in

terms of seasonal and temperature relationship, age susceptibility, and mortality patterns but differ to some degree in location of the causative organism in infected fish. CE

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generally occurs in spring following stocking of production ponds, with less severe outbreaks being reported in autumn. Negligible losses from CE occur in summer. Atypical A. salmonicida infections in spring are associated with optimum water temperatures (15–20◦ C) and to spawning activities, which can lead to superficial skin injury. During periods when water temperatures fluctuate between 6 and 20◦ C, carp losses due to CE can be greater than 50% (Fijan and Petrinec 1973). Tinman and Maurice (2009) reported cases of CE in common carp in Israel increased by 96% between 1997 and 1998 and increased again in 1999. The disease emerged across a broad temperature range of 11–23◦ C with a mean of 16◦ C. The disease was first observed in association with decrease in ambient temperature during the fall and persisted throughout the colder winter days. As water temperature warmed over 23◦ C the ulcers rapidly healed. Disease caused by atypical A. salmonica in other fish species also tends to be more prevalent in spring and fall, with little evidence of its presence in summer (McCarthy 1975; Cornick et al. 1984). Atypical A. salmonicida can be experimentally transmitted by cohabitation of healthy and diseased fish or inoculation of infected skin material into dermis or scale pockets of naive fish. However, the most consistent experimental transmission method is to rub infected material onto superficially scarified skin (Fijan 1976; Bootsma et al. 1977; Wiklund 1995a). Young carp are highly susceptible to CE, but severity of disease changes from an acute infection in young fish to a subacute and/or chronic infection as fish become older. A 15% survival occurred in 3 and 5 month old carp, but when these fish reached 10 months of age survival was 60% (Wiegertjes et al. 1993). It was suggested that resistance may be an inherited trait because mortality of offspring from one female was 100% when challenged while a companion population from a second female had 25–50% mortality. The source of atypical A. salmonicida is presumably latent carrier fish, although under cer-

tain conditions, the pathogen may survive for a period of weeks in the environment (Wiklund 1995b). In sterilized brackish water, the pathogen will survive for <14 days, but if sediment is added to the water, it survives for up to 63 days. Atypical A. salmonicida was recoverable for a shorter time in small containers with sterile fresh water and survival was better at 4◦ C than 15◦ C. Conclusions drawn from these data are that atypical A. salmonicida may survive in bottom sediment of brackish water environments for a long time; therefore, this substrate could serve as a pathogen reservoir. Atlantic tomcod captured in the Atlantic Ocean and held in tanks for experimental purposes invariably developed atypical A. salmonicida infections, displaying epithelial and muscular lesions in about 10 days following confinement. Mortality occurred at 25 days when held at 11◦ C; incubation time was doubled at 6◦ C (Williams et al. 1997). Ulcerative disease of goldfish is a chronic condition which primarily affects cultured fish that are 1 year of age or older, including brood stock. The disease usually occurs in early spring following stocking of spawning ponds with adult fish and when water temperatures range between 18 and 25◦ C. Losses of larger subadult and adult goldfish during this time may reach 45–90%. Epizootic survivors appear to have no long lasting disease effects with the exception of reduced egg yield. Saleable size goldfish (5–10 cm) become more susceptible as fish density, parasite load, and environmental stressors increase. Other potential pathogenic bacteria such as F. columnare, A. hydrophila, and Listonella (Vibrio) anguillarum (in marine fish) are often found in skin lesions in association with atypical A. salmonicida in goldfish. In Denmark, an epizootic of atypical A. salmonicida in turbot revealed that the onset and progression of disease paralleled an increase in water temperature from about 5–15◦ C, during which time mortality reached 15–20% (Pedersen et al. 1994). After initial outbreak and drug treatment, a secondary L. anguillarum infection occurred.

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Atypical A. salmonicida isolated from European flounder showed low virulence in rainbow trout, while infectious experiments in roach and bleak were inconclusive (Wiklund 1995a). However, when European flounder were exposed to the bacteria by ingestion or contaminated water following a skin abrasion, skin lesions resulted. While it was concluded that the atypical A. salmonicida from European flounder was not a threat to rainbow trout, it was noted by Morrison et al. (1984) that Atlantic cod could become carriers of the bacterium and serve as reservoirs. Also, in experimental studies by Carson and Handlinger (1988), atypical A. salmonicida isolated from goldfish was pathogenic to Atlantic salmon, brown, rainbow, and brook trout, while isolates from marine fish in Scandanavia were not pathogenic to rainbow trout.

Pathology Infections of atypical A. salmonicida are generally localized in the skin where small or large hemorrhagic, inflamed lesions occur in the center of which necrosis develops on the outer layer of the skin (Fijan 1976). Even though lesions may disappear, infection can remain and result in hydropsy, ascites, exophthalmia, and anemia. It was noted by Bootsma et al. (1977) that bacteria responsible for CE could only be found in dermal and subdermal lesions, but nevertheless a generalized edema did occur. As previously discussed, atypical A. salmonicida infections in other fish species (cod and Atlantic salmon) may be systemic, but still the skin will be most severely affected. Histopathology of atypical A. salmonicida in Atlantic cod showed that the host developed a defined reaction including encystment of the bacteria (Morrison et al. 1984). The spleen and kidney showed degenerative changes, leukocyte accumulation, and cyst formation. It was theorized by Pol et al. (1980) that inflammation, tissue necrosis, and possibly generalized edema present in CE-infected

fish could be caused by bacterial exotoxins. Maurice et al. (1999) showed that virulence of atypical A. salmonicida in goldfish ulcer disease was associated with production of a paracrystaline outer membrane A-layer protein.

Significance Infections of atypical A. salmonicida achromogens assumed a more important role in aquaculture and wild fish populations because of expanded known species susceptibility after 1980. That it occurs in a variety of wild and cultured fish in fresh and saltwaters, particularly tomcod, flounder, and halibut enhances its importance. Under certain conditions, CE can be a very serious disease in European carp and goldfish culture ponds. Reduced egg production and loss of yearling and adult fish have resulted from atypical A. salmonicida infections, a problem exacerbated by its poor response to conventional chemotherapeutics.

Other bacterial diseases of cyprinids Motile aeromonads (See Chapter 11) can cause disease problems in cultured carp (common, koi, grass, silver and bighead), golden shiners, fathead minnows, and goldfish, especially following handling, stocking, or transport. Generally, A. hydrophila, the principle bacterial organism in IDC syndrome in Europe, is the etiological agent. The bacterium has also been identified as the etiology in a variety of infections and mortalities found in wild cyprinid populations. Clinical signs of A. hydrophila in cyprinids are necrotic (frayed) fins, hemorrhaged scale pockets, scale loss, and development of necrotic skin lesions. Infections become systemic and produce edema, anemia, and hyperemia of internal organs. Carp with IDC are swollen with fluid in the muscles, scales exhibit lepidorthosis, and bloody fluid is present in the body cavity which produces dropsy. Cyprinid mortalities caused by A. hydrophila are usually chronic and seldom

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subacute. Pseudomonas spp. and P. fluorescens produce clinical signs similar to those observed in MAS in cyprinids and can be distinguished from each other by bacterial isolation and identification. Cyprinids, especially shiners, fathead minnows, and the Asian carps, are extremely susceptible to columnaris infections. This organism can be responsible for chronic losses in ponds following stocking and other periods of stress. Shiners and fathead minnows crowded into holding tanks for extended periods of time are highly susceptible to columnaris and mortalities may approach 100% if untreated. Altinok and Grizzle (2001) found that goldfish were more sensitive to columnaris in freshwater than in water containing ≥1.0 ppt NaCl. Water quality has a definitive effect on susceptibility of common carp to columnaris (Decostere et al. 1999). They studied the ability of different strains of F. columnare to attach to gills and determined that highly virulent strains attached more readily than low virulent strains and that the bivalent ion rich water and presence of nitrite, organic matter, and high water temperature enhanced the pathogens attachment ability. Cyprinids infected with columnaris display gray frayed fins, open ulcerative skin lesions where scales have sloughed, grayish color as a result of injury to the mucous layer and epithelium, and hemorrhages at the base of fins and around the operculum and mouth; lesions around the mouth may be yellowish due to the presence of large numbers of bacteria. Also, lesions may contain a variety of other waterborne bacteria, including A. hydrophila and P. fluorescens, as well as protozoan parasites.

Management of cyprinid bacterial diseases To avoid atypical A. salmonicida infections in carp, goldfish, and other susceptible fish species handling, high fish densities in ponds and holding tanks, or skin injuries should be kept to a minimum particularly in the cooler

months when fish are more susceptible. Brood goldfish, stocked into ponds in early spring just prior to spawning, often contract atypical A. salmonicida infections due to stress. When brood fish are stocked into spawning ponds in late fall or early winter, spring prespawning stress can be eliminated and UDG generally avoided; therefore, fall brood fish stocking is common on goldfish farms where atypical A. salmonicida is endemic. Fijan (1976) recommended that general prophylactic measures be taken on farms to reduce occurrence and/or severity of CE and UDG. Antibiotic baths, where permitted, can be applied to carp and goldfish after handling. Medicated feed containing oxytetracycline can be fed at 50 mg/kg of fish daily for 10 days to diseased fish; however, success of this procedure to control atypical A. salmonicida has been limited. A high rate of drug resistance for atypical A. salmonicida was reported by Barnes et al. (1994), who found that essentially all isolates from Scotland were resistant to Amoxycillin (>500 mg/L MIC); this resistance was chromosomal rather than plasmid mediated. In the Canadian Maritime Providences, an intraperitoneal injection of enrofloxacin was used to prevent atypical A. salmonicida infections in captured Atlantic tomcod (Williams et al. 1997). An injection of 5 mg/kg of the drug at time of capture, followed by additional injections at 10 and 45 days postcapture successfully prevented formation of skin and muscle lesions and mortality due to the pathogen. Several antigen preparations from atypical A. salmonicida were tested for their immunogenic potential to protect carp against CE (Evenberg et al. 1988). An injected, formalinkilled, whole cell bacterin provided consistent but moderate protection. However, a concentrated, deactivated culture supernatant afforded better protection against subsequent lethal challenge with the pathogen. A bath challenge system was used by Daly et al. (1994) to demonstrate the efficacy of vaccination with live A. salmonicida. Exposure to high

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concentrations of bacteria resulted in carp developing infection and typical clinical signs, but carp exposed to a sublethal dose of the bacteria developed only skin lesions, which healed after 1 week. When challenged by subcutaneous injection, 100% of naive carp died, fish exposed to high bacterial concentrations suffered 40–60% mortality, but all fish exposed to the sublethal dose of live bacteria recovered. Protection of these fish lasted for 5 months. These data and those of previous studies indicate that extracellular products of atypical A. salmonicida are involved in CE and that vaccination of carp with sublethal concentrations of live, attenuated bacteria against the disease is feasible. Using the knowledge that virulence of atypical A. salmonicida associated with production of a paracrystaline outer membrane A-layer protein, Maurice et al. (1999) cloned the Alayer protein into a pET-3d plasmid in order to express and produce a recombinant product for protein in E. coli (BL21[DE3]). The A-layer protein compared favorably by biochemical, immunological, and molecular characteristics with A-layer protein isolated from atypical A. salmonicida; the generated protein was indistinguishable from the wild type when examined by SDS-PAGE and gel filtration chromatography. This was the first large scale production of biologically active recombinant type A protein from atypical A. salmonicida and could possibly be used as a basis for a vaccine against the disease. Infections of MAS, Pseudomonas sp., and columnaris are treated the same in all fish species by using good management practices and reduce environmental stress, maintaining good water quality and/or prevention of skin injury from handling. Application of medicated feed with Terramycin may be used for active infections. Bath treatments with potassium permanganate at 2–4 ppm indefinitely or short term bath of up to 1 hour in 1–3 ppt NaCl are generally effective proactive disease prevention approaches for columnaris in goldfish, golden shiners, and fathead minnows.

Holding tanks of bait fish are often given prolonged treatments of commercially prepared concoctions that contain a variety of chemicals for columnaris. When used according to instructions these compounds usually have some beneficial effect.

References Altinok. I., and J. M. Grizzle. 2001. Effects of low salinities on Flavobacterium columnare infection of euryhaline and freshwater stenohaline fish. Journal of Fish Diseases 24:363–367. Barnes, A. C., T. S. Hastings, and S. G. B. Amyes. ˚ 1994. Amoxycillin resistance of Scottish isolates of Aeromonas salmonicida. Journal of Fish Diseases 17:357–363. Bootsma, R., N. Fijan, and J. Blommaert. 1977. Isolation and preliminary identification of the causative agent of carp erythrodermatitis. Veterinary Arhives 47:291–302. Carson, J., and J. Handlinger. 1988. Virulence of the aetiological agent of goldfish ulcer disease in Atlantic salmon, Salmo salar L. Journal of Fish Diseases 11:471–479. Chapman, P. F., R. C. Cipriano, and J. D. Teska. 1991. Isolation and phenotypic characterization of an oxidase-negative Aeromonas salmonicida causing furunculosis in coho salmon (Oncorhynchus kisutch). Journal of Wildlife Diseases 27:61–67. Cornick, J. W., C. M. Morrison, et al. 1984. Atypical Aeromonas salmonicida infection in Atlantic cod, Gadus morhua L. Journal Fish Diseases 7:495–499. Daly, J. G., G. F. Wiegertjes, and W. B. Van Muiswinkel. 1994. Protection against carp erythrodermatitis following bath or subcutaneous exposure to sublethal numbers of virulent Aeromonas salmonicida susp. nova. Journal of Fish Diseases 17:67–75. Decostere, A., and F. Haesebrouck. 1999. Influence of water quality and temperature on adhesion and high and low virulence Flavobacterium columnare strains to isolated gill arches. Journal of Fish Diseases 22:1–11. Elliott, D. G., and E. B. Shotts, Jr. 1980. Aetiology of an ulcerative disease in goldfish Carassius auratus (L.): microbiological examination of

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diseased fish from seven locations. Journal of Fish Diseases 3:133–143. Evelyn, T. P. T. 1971. An aberrant strain of the bacterial fish pathogen Aeromonas salmonicida isolated from a marine host, the Sable fish (Anoplopoma finbria), and from two species of cultured Pacific salmon. Journal of the Fisheries Research Board of Canada 28:1629–1634. Evenberg, D., P. DeGraaff, et al. 1988. Vaccineinduced protective immunity against Aeromonas salmonicida tested in experimental carp erythrodermatitis. Journal of Fish Diseases 11:337– 350. Fijan, N. 1972. Infectious dropsy in carp—a disease complex. Symposium of the Zoological Society of London 30:39–51. Fijan, N. 1976. Diseases of cyprinids in Europe. Fish Pathology 10:129–134 Fijan, N., and Z. Petrinec. 1973. Mortality in a pond caused by carp erythrodermatitis. Rivista Italiana Piscicoltura E Ittiopathologia 8:45– 49. Gudmandsottir, B. K. 1988. Infections of atypical ´ strains of the bacterium Aeromonas salmonicida (Abstract). Icelandic Agriculture Science 12:61–62. Hanninen, M.-L., and V. Hirvela-Koski. 1999. Ge¨ ¨ netic diversity of atypical Aeromonas salmonicida studies by pulsed-field gel electrophoresis. Epidemiology and Infection 123:299–307. Hubert, R. M., and W. P. Williams. 1980. Ulcer disease of roach, Rutilus rutilus (L). Bamidgeh 32:46–52. Iida, T., K. Nakakoshi, and H. Wakabayashi. 1984. Isolation of atypical Aeromonas salmonicida from diseased eel, Anguilla japonica. Fish Pathology 19:109–112. Kimura, T. 1969. A new subspecies of Aeromonas salmonicida as an etiological agent of furunculosis on “Sakuramasu” (Oncorhynchus masou) and pink salmon (O. gorbuscha) rearing for maturity, Part 1. On the morphological and physiological properties. Fish Pathology 3:34–44. Maurice, S., K. Hadge, et al. 1999. A-protein from ¨ achromogenic atypical Aeromonas salmonicida: molecular cloning, expression, purification, and characterization. Protein Expression and Purification 16:396–404. McCarthy, D. H. 1975. Fish furunculosis caused by Aeromonas salmonicida var. achromoqenes. Journal of Wildlife Diseases 11:489–493.

Morrison, C. M., J. W. Cornick, et al. 1984. Histopathology of atypical Aeromonas salmonicida infection in Atlantic cod, Gadus morhua L. Journal of Fish Diseases 7:477–494. Munroe, A. L. S., and T. S. Hastings. 1993. Fu˚ runculosis. In: V. Inglis, R. J. Roberts, and N. R. Bromage (eds) Bacterial Diseases of Fish. Oxford, Blackwell Press, pp. 122–142. Paterson, W. D., D. Douey, and D. Desautels. 1980. Isolation and identification of an atypical Aeromonas salmonicida strain causing epizootic losses among Atlantic salmon (Salmo salar) reared in a Nova Scotian Hatchery. Canadian Journal of Fisheries and Aquatic Sciences 37:2236–2241. Pedersen, K., H. Kofod, et al. 1994. Isolation of oxidase-negative Aeromonas salmonicida from diseased turbot Scophthalmus maximus. Diseases of Aquatic Organisms 18:149–154. Pol, J. M. A., R. Bootsma, and J. M. BergBlommaert. 1980. Pathogenesis of carp erythrodermatitis (CE): role of bacterial endo- and exotoxin. In: W. Ahne (ed.) Fish Diseases Third COPRAQ-Session. Berlin, Springer-Verlag, pp. 120–125. Shotts, E. B., Jr., F. D. Talkington, et al. 1980. Aetiology of an ulcerative disease in goldfish, (Carassius auratus L.): characterization of the causative agent. Journal of Fish Diseases 3:181– 186. Smith, I. W. 1963. The classification of “Bacterium salmonicida.” Journal of General Microbiology 33:263–274. Tinman, S., and S. Maurice. 2009. Carp erythrodermatitis a new and increasingly prevalent disease in Israel. http://www.rockymountainkoi.com/ new-page-5.htm. Accessed 08/17/09. Whittington, R. J., S. P. Djordjevic, et al. 1995. Restriction endonuclease analysis from goldfish Carassius auratus, silver perch Bidyanus bidanus, and greenback flounder Rhombosolea tapirina in Australia. Diseases of Aquatic Organisms 22:185–191. Wichardt, U.-P. 1983. Atypical Aeromonas salmonicida-infection in sea-trout (Salmo trutta, L.). I. Epizootiological studies, clinical signs and bacteriology. Lasforskningsinstitutet Meddelande (Salmon Research Institute Report) No. 6:1–l0. Wiegertjes, G. F., J. G. Daly, and W. B. Muiswinkel. 1993. Disease resistance of carp, Cyprinus

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carpio L.: identification of individual genetic differences by bath challenge with atypical Aeromonas salmonicida. Journal of Fish Diseases 16:569–576. Wiklund, T. 1990. Atypical Aeromonas salmonicida isolated from ulcers of pike, Esox lucius L. Journal of Fish Diseases 13:541–544. Wiklund, T. 1995a. Virulence of “atypical” Aeromonas salmonicida isolated from ulcerated flounder Platichthys flesus. Diseases of Aquatic Organisms 21:145–150. Wiklund, T. 1995b. Survival of “atypical” Aeromonas salmonicida in water and sediment microcosms of different salinities and temperatures. Diseases of Aquatic Organisms 21:137–143.

Wiklund, T., and G. Bylund. 1991. A cytochrome oxidase negative bacterium (presumptively and atypical Aeromonas salmonicida) isolated from ulcerated flounders (Platichthys flesus (L.)) in the northern Baltic Sea. Bulletin of the European Association of Fish Pathologists 11:74–76. Wiklund, T., and I. Dalsgaard. 1995. Atypical Aeromonas salmonicida associated with ulcerated flatfish species in the Baltic Sea and the North Sea. Journal of Aquatic Animal Health 7:218–224. Williams, P. J., S. C. Courtenay, and C. Vardy. 1997. Use of enrofloxacin to control atypical Aeromonas salmonicida in the Atlantic tomcod (Microgadus tomcod Walbaum). Journal of Aquatic Animal Health 9:216–222.

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Chapter 13

Eel bacterial diseases

Generally eels contract many of the same bacterial diseases as do other warmwater fishes; however, two infectious diseases, Edwardsiellosis (Edwardsiella tarda), also known as hepatonephritis, and red spot disease (Pseudomonas anguilliseptica), seem to have a greater affinity for cultured eel than for other fish species. Other bacterial fish pathogens Flavobacterium columnare, motile Aeromonas septicemia (See Chapter 11), atypical Aeromonas salmonicida (Chapter 12), and Vibrio (Listonella) anguillarum (See Chapter 14) also occur in eel.

Edwardsiellosis Edwardsiellosis is a subacute to chronic disease of cultured eels, channel catfish, and numerous other fish species (Hoshina 1962; Meyer and Bullock 1973; CAB International 2006). The disease is also known as hepatonephritis in eel, and emphysematous putrefactive disease in channel catfish, or fish gangrene because of the foul odor emitted from gas-filled pockets in necrotic muscle. The etiology of Edwardsiel-

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.

losis is Edwardsiella tarda, which was earlier known as Paracolobacterum anguillimortifera (Wakabayashi and Egusa 1973).

Geographical range and species susceptibility Edwardsiella tarda is a cosmopolitan bacteria having been found in natural and freshwater culture facilities as well as marine environments around the world. It has been reported from 25 countries in North and Central America, the Caribbean, Europe, Asia, Australia, Africa, and the Middle East (Austin and Austin 1987; CAB International 2006). Although the most prominent fish species infected by E. tarda are eels (American, European, and Japanese) and channel catfish, the organism has been isolated from many diseased freshwater and marine fish species, including carp, largemouth bass, striped bass, red seabream, flounder, tilapia, yellowtail (Hoshina 1962; Meyer and Bullock 1973; Austin and Austin 1987; Francis-Floyd et al. 1993; Benli and Yildiz 2004). CAB International (2006) lists over 47 fish species that are either naturally or experimentally susceptible to E. tarda. Edwardsiella tarda has also been isolated from intestinal content of apparently healthy fish during routine bacteriological 327

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surveys (Wyatt et al. 1979; van Damme and Vandepitte 1980). Generally considered to be a warm water inhabitant, E. tarda occasionally has been implicated in salmonid infections. It was isolated from migrating adult chinook salmon in the Rogue River in Oregon when water temperatures were above normal (Amandi et al. 1982). In Nova Scotia, Canada, it was isolated from a small number of migrating adult Atlantic salmon (Martin 1983) and from brook trout in Quebec (Uhland et al. 2000). Edwardsiella tarda occasionally causes disease in ornamental fish in aquaria (Humphrey et al. 1986). Edwardsiella tarda infections are not limited to fish; the pathogen often exist as part of normal intestinal microflora of many fisheating birds and mammals, reptiles, amphibians, and marine mammals, as well as cattle, and swine (White et al. 1969; Coles et al. 1978). Humans have also been known to become infected with E. tarda, usually as an opportunistic pathogen (Hargreaves and Lucey 1990).

Clinical signs and findings Clinical signs of E. tarda infections differ slightly between locality and fish species. Acutely infected eels exhibit lethargic swimming, tend to float at the surface, fins become congested and hyperemic, petechial hemorrhages develop on the underside, the anal region is swollen and hyperemic, and gas filled pockets may develop between dermis and muscle (Figure 13.1). Ulcerative, necrotic lesions may develop in the skin and muscle. Internally, the liver, kidney, and spleen are whitish and may develop abscesses. Infected channel catfish develop small to large abscesses in the lateral musculature, which emit a foul odor when incised. A variety of nonspecific clinical signs occur in other fish species that include exophthalmia, opaqueness of the eye, swollen and necrotic skin and muscle lesions, rectal protrusion, and abscesses in internal organs.

Diagnosis Edwardsiella tarda is easily isolated from internal organs and muscle lesions of clinically infected fish and identified by conventional bacteriological or serological methods. When working with eels, it is necessary to isolate bacteria from diseased fish for diagnostic purposes because clinical signs of E. tarda and other bacterial septicemias cannot be differentiated (Nichibuchi et al. 1980). Edwardsiella tarda isolation is easily made on BHI agar, TSA, or any other general purpose medium (Meyer and Bullock 1973; Amandi et al. 1982). Incubation at 26–30◦ C for 24–48 hours yields small, circular, convex, transparent colonies approximately 0.5 mm in diameter. The incidence of E. tarda isolation from the brain of chinook salmon was improved from 2 to 19% by inoculating thioglycolate media and then transferring the inoculum to BHI agar rather than inoculating directly from fish onto BHI agar (Amandi et al. 1982). Edwardsiella tarda form small, green colonies with black centers on EIM (Shotts and Teska 1989). Methods for rapid E. tarda diagnosis and identification, in addition to conventional methods of isolation and biochemical and biophysical characteristics, include serologicalfluorescent antibody technique (FAT) and enzyme-linked immunosorbent assay (ELISA) (Swain and Nayak 2003). Serological techniques of bacterial agglutination with specific antisera can be used for positive identification. Rabbit polyclonal antibodies against outer membrane proteins (OMPs) were used for detection of E. tarda by ELISA (Kumar et al. 2007). OMPs from 16 isolates of E. tarda recovered from snakeheads in India had one major common protein and nine other variable proteins. Rabbit polyclonal antibodies acted strongly against the OMPs and detected one common 44 kd protein that showed no crossreaction with five common aquatic bacteria including Escherichia coli, Pseudomonas fluorescens, and Aeromonas hydrophila. The study indicated that rabbit polyclonal antibodies

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(a)

(b)

Figure 13.1 (a) Japanese eel infected with Edwardsiella tarda. Note the pale, mottled liver with an abscess (large arrow) and slight petechiae on the throat (small arrow). (Photo by E. B. Shotts.) (b) Japanese eel infected with Pseudomonas anguilliseptica showing petechia (arrow). (Photos by T. Miyazaki.)

against the OMPs could be useful in detecting E. tarda in an ELISA system.

Bacterial characteristics Edwardsiella tarda, a member of the family Enterobacteriaceae, is a small, straight rod about 1 µm in diameter and 2–3 µm in length with peritrichous flagella (Farmer and McWhorter 1984). It is Gram-negative, facultatively anaerobic. Key presumptive characteristics of E. tarda are its motility, indole production in tryptone broth, H2 S produc-

tion on triple sugar iron (TSI) agar slants, tolerantes up to 4% salt, produces gas during glucose fermentation, and grows at 40◦ C (Table 11.2 in Chapter 11 Catfish bacterial diseases). It is catalase-positive, cytochrome oxidase negative, ferments glucose, reduces nitrate to nitrite, and is lactose-negative. Positive identification of E. tarda can be made by specific serum agglutination or fluorescent antibody tests because no serological cross reactivity occurs between it and Edwardsiella ictaluri or other enteric bacteria. Edwardsiella tarda is phenotypically a homogeneous

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organism. Waltman et al. (1986), as well as others, found only slight variations in biochemical and biophysical characteristics of 116 E. tarda isolates from the United States and Taiwan. Using the Minitek Numerical Identification System, Taylor et al. (1995) correctly identified E. tarda 100% of the time compared to 83% positive identification with the API 20E system. However, Panangala et al. (2005) showed that analysis of 16S-23S inergenic spacer regions of the rRNA operons of E. tarda and E. ictaluri have a close genetic relationship and that E. tarda is more genetically diverse than E. ictaluri. Maiti et al. (2009) used biochemical analysis, plasmid profiling SDS-PAGE, and enterobacterial repetitive intergenic consensus (ERIC)-PCR of E. tarda isolated from pond sediments to reveal five genogroups with similarity ranging from 50 to 100%, but no correlations were found between the tests. Protein patterns of E. tarda isolates analyzed by SDS-PAGE indicated that the isolates could be divided into four heterogenous groups. Maiti et al. (2008) also found that BOX-PCR (a method for DNA fingerprint analysis) and PCR showed some genotypes to be habitat specific. Farmer and McWhorter (1984) identified a wild type E. tarda and a second Biogroup 1, both of which are easily distinguished from E. ictaluri and Edwardsiella hoshinae (a pathogen of reptiles and amphibians) (Grimont et al. 1980). Park et al. (1983) identified four E. tarda serotypes (A, B, C, and D) among 445 isolates collected from fish and other environmental sources. Seventy-two percent of fish isolates were serotype A, indicating that it may be the predominant disease-causing type in fish. Also, 28 strains of E. tarda isolated from diseased flounder were identical to serotype A (Rashid et al. 1994a). Analysis of 16S-23S intergenic spacer regions (ISR) of rRNA operons in E. tarda showed a high degree of size and sequence between E. ictaluri and E. tarda and within each species (Panangala et al. 2005). These results

confirm a close genetic relationship between the two species of bacteria and that E. ictaluri is more homogeneous than E. tarda. Panangala et al. (2006) further showed that E. tarda isolates are less homogeneous than E. ictaluri with regard to antiserum protein profiles and fatty acid profiles, but a close relatedness of the two bacterial species was demonstrated.

Epizootiology According to Egusa (1976), E. tarda infections of Japanese eels were more prevalent in summer when water temperatures were highest. This trend was also reported by Kuo et al. (1977), who found that the optimum water temperature for Edwardsiellosis in eel populations was 30◦ C. Edwardsiellosis of catfish occurs most often in summer when water temperatures exceed 30◦ C. However, the disease is not universally restricted to warm temperatures; Liu and Tsai (1982) found that eel infections in Taiwan were most common during January to April when temperatures fluctuated. In the United States, Amandi et al. (1982) isolated E. tarda from 19% of dead or dying Rogue River chinook salmon in Oregon when water temperatures rose from 17 to 20◦ C. The various temperatures at which E. tarda can cause disease may reflect an opportunistic capability that could be more closely associated with increased species susceptibility during temperature related stress than to pathogenicity of the organism. Under similar conditions at 20◦ C, Japanese flounder and eel reacted very differently to E. tarda endotoxins. A heat labile endotoxin from virulent E. tarda was 15 times more toxic to Japanese flounder than to Japanese eel. The main source of E. tarda is presumably intestinal content of carrier aquatic animals, primarily catfish and eels; however, amphibians and reptiles must also be considered bacterial sources. Edwardsiella tarda is probably released from infected animals into the water or is naturally present in the mud and/or water. It has been reported that cell counts of

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E. tarda in eel pond water are higher when clinical disease is present (Hidaka et al. 1983). Edwardsiella tarda cell counts at 22–26◦ C were four times higher when clinical disease was present, which agrees with observations made in E. ictaluri-infected channel catfish culture pond water (Earlix 1995). Fish are experimentally infected with E. tarda by IP or IM injection, stomach gavage, or immersion in water containing the pathogen but infection by immersion is not guaranteed. Huang and Liu (1986) killed 100% of IPinjected eels when E. tarda was mixed with Aeromonas hydrophila but waterborne exposure to the same mixture failed to induce disease and death without sublethal concentrations of nitrogenous compounds. Darwish et al. (2000) had little success in infecting channel catfish by immersion in an E. tarda bath unless the skin was first superficially abraded. Mekuchi et al. (1995) successfully infected Japanese flounder with E. tarda by IM and IP injection, immersion, and orally. Before introducing E. tarda into the intestinal lumen via silicon tube, Miyazaki et al. (1992) injured the intestine with hydrogen peroxide. Infected fish developed pathological lesions in the kidney and liver that were nearly identical to those in natural infections and resulted in death 5–23 days postinfection. Edwardsiella tarda is a hardy, facultative bacterium that can survive for extended periods in water at temperatures of 15–45◦ C (optimum 30–37◦ C), pH 4.0–10.0 (optimum pH 7.5–8.0), and in salt solutions of 0–4% (optimum 0.5–1.0% NaCl) (Chowdhury and Wakabayashi 1990). Isolation of E. tarda for over 76 days from 20◦ C pond water indicates that water can serve as a pathogen source. The organism also survives for days in net fowling material (Wyatt et al. 1979). Because E. tarda is a common inhabitant of catfish pond waters, its presence poses a constant threat of disease. In E. tarda-positive catfish ponds, the organism was isolated from 75% of pond water samples, 64% of mud samples, and from l00% of apparently healthy

frog, turtle, and crayfish sampled (Wyatt et al. 1979). Also, E. tarda was isolated from the flesh of as many as 88% of dressed domestic catfish but from the flesh of only 30% of imported dressed fish. In an ecological study, Rashid et al. (1994b) isolated E. tarda in 86% of water samples from one salt water flounder mariculture pond and in 22% from a second pond. In the first pond, E. tarda was present in 44% of sediment samples and 14% of asymptomatic fish but was found in 0% of sediment and only 2% of flounder from a second pond. Ironically, no clinical disease, morbidity, or mortality occurred in either pond. An environmental stressor may not be essential for E. tarda to initiate infections in fish but fluctuating and high water temperatures, high water organic content and generally poor water quality, and crowded conditions can contribute to the onset and severity of disease (Meyer and Bullock 1973). When exposed to environmental stressors 25–50% of juvenile channel catfish experimentally infected with A. hydrophila also developed an E. tarda infection (Walters and Plumb 1980). Only 4–13% of nonstressed, A. hydrophila-uninfected fish developed E. tarda infections. These data indicate that environmental stress, in conjunction with other bacterial infections, can predispose channel catfish to disease from naturally present E. tarda. Also, sublethal concentrations of copper (100–250 µg/L) in the water reduced resistance of Japanese eels to E. tarda infections (Mushiake et al. 1984). Uhland et al. (2000) postulated that stress-associated immunosupression due to an increase in summer water temperature, and the lack of precipitation were primary causes of Edwardsiellosis in brook trout in Quebec. Transmission of E. tarda in catfish, eels, and most other fish species appears to be enhanced at water temperatures from 20 to 30◦ C. Japanese flounder were most susceptible at 20–25◦ C by IM injection in which an LD50 of 7.1 × 101 CFU was established (Mekuchi et al. 1995). This compared to an LD50 of 1.7 × 102 CFU for IP injection and 3.6 × 106 CFU/mL and 1.3 × 106

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CFU/fish for immersion and oral exposure, respectively, at the same temperature. Suprapto et al. (1995) also showed that the LD50 was 101.1 CFU/g of flounder and 106 CFU/g of eel. On several occasions, one of the author observed intensively cultured Nile tilapia, which were environmentally stressed or had a moderate to heavy Trichodina infection (protozoan ectoparasite), become clinically infected with E. tarda and/or Streptococcus spp. (unpublished). While stressed, neither bacterial infection responded to chemotherapy but as soon as the stressor was relieved and parasites eliminated, bacterial infections disappeared without antibiotic medication. In experimental infectivity studies, 100 g eels were immersed in E. tarda and 60% mortality occurred (Liu and Tsai 1982). Kodoma et al. (1987) reported that natural E. tarda infections can cause an 80% loss in eels. A cumulative mortality of 83% in 4 days in conjunction with a spontaneous infection in Nile tilapia was reported by Benli and Yildis (2004). Edwardsiella tarda can be a health threat to humans where it usually manifests itself as gastroenteritis and diarrhea. However, extraintestinal infections can produce typhoid like illness, meningitis, peritonitis with sepsis, cellulitis, and hepatic abscess (Clarridge et al. 1980). The organism has been associated with wound infections precipitated by fish hooks or catfish spine punctures (Hargreaves and Lucey 1990). It has also been implicated in diarrhea associated with an intestinal infection of Entamoeba histolytica, other tropical diarrheas, and from consumption of contaminated freshwater fish (Gilman et al. 1971). According to Janda and Abbott (1993), in English-language literature concerning E. tarda, there have been at least 250 reported cases of E. tarda infections in humans. The ability of E. tarda to infect warm blooded animals and humans in particular, was shown by Janda et al. (1991) who demonstrated its ability to invade HEp-2 cell monolayer cell cul-

tures, produce cell-associated hemolysin and siderophores, and express mannose-resistant hemagglutination against guinea pig erythrocytes. Janda and Abbott (1993) also showed that strains of E. tarda produced 30–40% higher levels of cell associated hemolytic activity (hemolysins) than did E. ictaluri which affects only fish. This increased hemolytic activity could contribute to E. tarda pathogenicity to humans. Infections in humans are usually successfully treated with antibiotics but deaths have been reported.

Pathology In Japanese eels, in which most pathology studies have occurred, E. tarda causes either a nephritic or hepatic histopathology (Miyazaki and Egusa 1976a, b; Miyazaki and Kaige 1985). In the more common nephritic form, the kidney is enlarged and contains varioussize abscesses that initially occur in the sinusoids of the hematopoietic tissue. This tissue becomes swollen and foci of infection develops into abscesses that progress into cavities filled with dark red, odiferous, purulent matter. These large abscesses that contain neutrophils, some of which phagocytize bacteria, are walled off by fibrin. Cell liquefaction follows and small abscesses can be found in other organs and the lateral musculature that adjoins the kidney. In the hepatitis form of disease, livers are usually enlarged and contain various-size abscesses, some of which leak fluid into the body cavity. Enlarged abscesses involve hepatocytes and blood vessels that form pus-like emboli and pyemia with extensive bacterial multiplication (Egusa 1976). Pathogenesis of experimental E. tarda infection in channel catfish following skin abrasion and water exposure to the pathogen results in lesions in a hyperemic spleen in 20 hours (Darwish et al. 2000). The infection spreads to the liver and other organs by day 2 but by 4–5 days PI, the whole body is infected. These data

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suggest that E. tarda is not a rapidly evolving disease following its access to an injured body surface. Histopathology, due to E. tarda, in turbot includes severe inflammation with a large number of abscesses primarily in the kidney and spleen where macrophages are the main inflammatory cells. Miwa and Mano (2000) found that Japanese flounder experimentally infected with E. tarda developed enlarged livers in the early stages of infection. However, they concluded that pathology was not the result of any observable histopathology changes and that E. tarda causes hypertrophy of the cells as well as preventing decrease in liver cell number. At least some E. tarda isolates produce toxic extracellular products (ECP). Ullah and Arai (1983) found evidence that E. tarda does not form endotoxins as do other Gram-negative bacteria but produces two exotoxins that may contribute to pathogenicity. However, factors that regulate pathogenicity of E. tarda are unclear. Suprapto et al. (1995) detected a heat-labile ECP in a serogroup A strain of E. tarda that was lethal to Japanese eel and Japanese flounder. The optimum incubation temperature for ECP production was 25–30◦ C, which coincides with most reports of optimum temperature for fish susceptibility. The ECP production peaked at 72–120 hour postmedia inoculation. An intracellular component (ICC), which occurred from 24 to 120 hours postmedia inoculation, was also associated with lyses of E. tarda cells. Han et al. (2006) stated that there is no difference between virulent and nonvirulent E. tarda isolates with regard to hemolytic and cytotoxic activities, but mortality titers ranged between 102.5 and 105.5 cfu/ml in olive flounder. Also, the results demonstrated that the ability of E. tarda to resist compliment activity and phagocytosis is conferred by its superoxide dismutase and catalase activities; two enzymes that are essential in E. tarda pathogenesis. Results of these studies suggest that toxin produced by E. tarda plays an important role in its virulence.

Significance Significance of E. tarda depends largely on fish species affected and presence of environmental stressors. However, with an increasing number of reported susceptible fish species and other animals including humans, E. tarda is an organism that can not be ignored. In Taiwan and Japan, it is one of the most serious bacterial diseases to affect cultured eels. However, in the United Sates, unless channel catfish are crowded, the disease is of minor consequence. A subclinical infection in market-size fish will affect fish quality at slaughter because of necrotic lesions in the muscle and can cause equipment contamination problems at such times (Meyer and Bullock 1973). When this occurs, fish processing lines are shut down, cleaned, and disinfected. In other fish species, including intensively cultured tilapia, E. tarda may be responsible for some losses but usually they are chronic rather than acute. Caution must always be taken when handling E. tarda-infected fish because of potential opportunistic infectivity to man.

Red spot disease Red spot disease is a mild to severe bacterial infection that affects mainly cultured eels, but recently the species susceptibility has expanded. The disease, known as “Sekiten-byo” in Japan, is caused by Pseudomonas anguilliseptica (Wakabayashi and Egusa 1972).

Geographical range and species susceptibility Red spot disease was initially identified in cultured Japanese eels (Wakabayashi and Egusa 1972). While most prevalent in Japan, the disease and its etiological agent have been identified in Taiwan, Malaysia, Scotland, Finland, the Netherlands, and France (Stewart

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et al. 1983; Nash et al. 1987;et al. Wicklund and Bylund 1990; Haenen and Davidse 2001). Red spot disease is present in marine as well as freshwater fish. Although Japanese eel appear to be the most susceptible fish species, European eels are also highly susceptible to the pathogen. Additionally the disease has occurred in, or isolated from, seabream, turbot, giant seaperch, and estuarine grouper in Malaysia (Nash et al. 1987); Atlantic salmon, brown and rainbow trout, and whitefish (Wicklund and Bylund 1990); wild Baltic herring along the southwest coast of Finland (Bylund 1983; Lonnstr om ¨ ¨ et al. 1994); maricultured gilthead seabream, seabass, and turbot in France (Berthe et al. 1995); sea bream in Spain (Blanco et al. 2002); and striped jack in Japan (Kusuda et al. 1995). Disease has been produced by experimental infections in ayu (Nakai et al. 1985a), but an isolate from Baltic herring was avirulent to rainbow trout (Lonnstr om ¨ ¨ et al. 1994).

eels (Miyazaki and Egusa 1977; Ellis et al. 1983). Kusuda et al. (1995) reported hemorrhages in the mouth, nose, brain, and on the operculum of infected striped jack.

Diagnosis Red spot disease of eels is conventionally diagnosed by isolation of P. anguilliseptica from internal organs of diseased fish. The slow growing organism is cultured on general purpose media (BHI or TSA) where it forms 1 mm diameter colonies in 3–4 days at 25◦ C. Isolation is enhanced by the addition of NaCl. The bacterium is similarly isolated from other fish species and is identified by conventional microbiological techniques. However, using a PCRbased system, Blanco et al. (2002) identified P. anguilliseptica from diseased and asymptomatic sea bream in Spain. Identification by this procedure was completed in 8 hours as opposed to 10 days required for conventional microbiological methods.

Clinical signs Japanese eels infected with P. anguilliseptica develop petechial hemorrhage in the subepidermal layer of the jaws, underside of the head, and along the ventral body surface (Figure 13.1). Internally, petechia can occur on the peritoneum, the liver is swollen, and the spleen and kidney become atrophied (Jo et al. 1975). European eels have similar pathology, but less severe than in Japanese eels (Ellis et al. 1983). Clinical signs in fish species other than eels vary. In Baltic herring, hemorrhages were present in eyes, fins, and on the heads; the cornea was ulcerated and bloodstained abdominal ascites were seen in several fish (Lonnstr om ¨ ¨ et al. 1994). Berthe et al. (1995) described petechial hemorrhages in the skin and livers of farmed gilthead seabream and moribund fish exhibited abdominal distension causing a “belly-up” syndrome. Skin lesions were also observed on these fish, but lesions and histopathology were less severe than in

Bacterial characteristics Pseudomonas anguilliseptica measures 0.8 µm in diameter by 5–10 µm in length. Growth takes place at 5–30◦ C, but not at 37◦ C. For presumptive identification, P. anguilliseptica is motile (more so at 15 than at 25◦ C), gramnegative, cytochrome oxidase-positive and negative for soluble pigments, H2 S production, and indole production (Palleroni 1984) (Table 13.1). The organism is not reactive in glucose oxidation-fermentation tests; however, it is positive for catalase, degradation of gelatin, and grows on media with 0–4% NaCl. The bacterium has low metabolic reactivity for many sources of carbon (Stewart et al. 1983). Thirteen isolates of P. anguilliseptica from France were identical phenotypically to the type strain NCIMB 1949T (Berthe et al. 1995) and to isolates from Finland (Lonnstr om ¨ ¨

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Table 13.1 Biophysical and biochemical characteristics of Pseudomonas fluorescens and P. anguilliseptica. Characteristic∗

P. fluorescens

P. anguilliseptica

Cell size (µm) Gram stain Motility Fluorescent pigment on agar Optimum temperature (◦ C) Growth at 4◦ C 30◦ C 40◦ C Glucose motility deep Growth in 4% NaCl Catalase Cytochrome oxidase Hydrolysis of Arginine Gelatin Starch Indole production Nitrate reduction ß-galactosidase Acid production from Arabinose Arginine D-alanine Fructose Galactose Glucose Glycerol Inositol Lactose Maltose Mannitol Manose Raffinose Salicin Sucrose Trehalose Xylose

0.8 × 2.3–2.8 − + (weak) + 25–30 + + − Oxidative − + + + + + − + −

0.4 × 2–5 − + (15◦ C best) − 25 + + − ? + + + − + − − − −

+ + + + + + + + − + + + ? − + + +

− − ? − − − − − − − − − − − − ? −

Source: Wakabayashi and Egusa (1972); Palleroni (1984). +, positive; −, negative; ?, unknown.

et al. 1994). Confirmation of the French isolates was achieved by agglutination with specific antisera of the type strain. There are two antigenic groups of P. anguilliseptica (Nakai et al. 1981): Type I antigen is heat labile (121◦ C for 30 minutes) while Type II is heat stable. The heat labile type from Japan possesses an R antigen and is pathogenic only to eels. On the basis of precipitin and agglutination test, P. anguilliseptica isolates

from Taiwan are identical to the Japanese R antigen type (Nakai et al. 1985b). Lopez-Romalde et al. (2003) determined ´ the phenotypic and genetic characterization of P. anguilliseptica from Spain, which showed close similarity to that of isolates from Japan. Genetic analysis by means of randomly amplified polymorphic DNA demonstrated existence of variability within P. anguilliseptica. Two major genetic groups were established,

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which were related to the host origin of the isolates.

Epizootiology Epizootiological characteristics of red spot were summarized by Muroga et al. (1973) and Muroga (1978) as follows: (1) prevailed in brackish-water ponds, (2) prevailed when water temperature was below 20◦ C and disappeared when above 27◦ C, and (3) prevailed among Japanese eels. In Japan, red spot disease of eels occurs primarily in April and May, becomes less frequent during summer, and reoccurs to a mild degree in October. Experimental infection of Japanese eels confirmed a relationship between water temperature and P. anguilliseptica epizootics (Muroga et al. 1975). Nearly all infected Japanese eels held below 20◦ C died, 71% of European eels died at 21◦ C, while few fish of either species held above 27◦ C succumbed to the disease. In France, disease in gilthead seabream occurred below 16◦ C with highest mortality occurring in the 9–16◦ C range. Striped jack in Japan were found to be infected with P. anguilliseptica at 14–16◦ C (Kusuda et al. 1995). European eel eelvers imported into the Netherlands, and held at 23–25◦ C, suffered 3% to 20% mortality within 2–3 weeks of the initial outbreak (Haenen and Davidse 2001). Because P. anquilliseptica tolerates up to 4% NaCl, infections may be as severe in brackish and salt water as in fresh water ponds. In Malaysia, the bacterium occurred in giant seaperch and estuarine grouper in off shore cages; mortality was 20–60% during the cooler season of November–December and February–March; this episode also coincided with poor water quality (Nash et al. 1987), leading to speculation that the bacterial infection was secondary to stressful environmental conditions and poor nutrition. Mortality among 300 g striped jack infected with P. anguilliseptica was 0.1–1% daily with a cumulative mortality of 30% from February through April (Kusuda et al. 1995).

Mushiake et al. (1984) noted that copper concentrations of 25–100 µg of Cu/L in water reduced the Japanese eel’s resistance to P. anguilliseptica by adversely affecting phagocytosis and other cell-mediated defense mechanisms. Stewart et al. (1983) demonstrated the affinity of the organism for young eels when 96% of eelvers and only 3.9% of adults died following experimental infection. After describing P. anguilliseptica in eye lesions of Baltic herring, Lonnstr om et al. ¨ ¨ (1994) suggested that P. anguilliseptica had been present in that region for a long time; Bylund (1983) had noted similar lesions and Gram-negative bacteria in smears from ulcerated fish eyes. The primary pathogenicity of P. anguilliseptica was questioned by Lonnstr om ¨ ¨ et al. (1994) because the organism was isolated from eyes or kidneys of only about 43% of Baltic herring assayed and from no livers or spleens. It was also noted that P. anguilliseptica had been isolated during disease outbreaks in rainbow trout farms along the Finish coast and that Baltic herring may have been the disease reservoir. Even though the bacterium showed low pathogenicity to trout in the laboratory, stress factors present in aquaculture could result in significant disease.

Pathology In histopathology studies of red spot disease in Japanese eels, Miyazaki and Egusa (1977) showed that lesions appeared initially in the dermis, subcutaneous adipose tissue, interstitial muscle, vascular walls, bulbous arteriosus, and heart. Bacteria multiplied in the vascular walls, producing inflammation with serous exudation and cellular infiltration. Many small hemorrhages occurred in dermal connective tissue, and there was congestive edema of visceral organs and fatty degeneration of liver hepatic cells, serous exudation and cellular proliferation of the spleen, glomerulitis, activation of reticuloendothelial cells lining the sinusoids, and atrophy of hematopoietic tissue of the kidney.

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Several days postinjection with P. anguilliseptica, eels became moribund, demonstrating clinical signs similar to naturally infected fish and mortality followed at 6–10 days postinfection (Wakabayashi and Egusa 1972). No exotoxin production by P. anguilliseptica has been demonstrated, but it is speculated that pathogenicity results from presence of these substances. Nakai et al. (1985b) traced the progression of a Japanese eel red spot infection after fish were injected IM with P. anguilliseptica and held at 12◦ C, 20◦ C, and 28◦ C. Viable bacterial cells in the blood decreased for 1–12 hours postinjection, followed by a stationary period when numbers of cells remained constant or increased slightly. Bacterial cell counts then increased to 108 –1010 CFU/g of tissue or per milliliter of blood, thus establishing a persistent septicemia until death. Bacterial growth from injection to maximum concentration took approximately 12 days at 12◦ C and 5 days at 20◦ C. When inoculated fish were held at 28◦ C, bacterial cells disappeared from internal organs within 2 days. Experimental exposure of 40 g seabream by IP and subcutaneous injection and skin swabs after abrasion resulted in 100% mortality and pathogen isolation was made from all carcasses (Berthe et al. 1995). All control fish survived.

Significance Red spot is one of the most serious diseases of cultured eels in Japan and with its appearance in Europe and in other locals it has now become a global concern. The ability of P. anguilliseptica to infect numerous fish species and its presence in wild populations that may serve as pathogen reservoirs adds to its importance. The potential for economic loss to eel producers caused by P. anguilliseptica due to its expanding geographical range and wider species susceptibility was emphasized by Berthe et al. (1995).

Other bacterial diseases of eel A variety of other bacterial diseases affect eels. Columnaris (See Chapter 11) infects most all freshwater fish including cultured eels in freshwater ponds. Fins of infected fish turn white, become necrotic and frayed, and affected body areas lose pigmentation, becoming white when the outer layer of skin is destroyed. Mortality due to columnaris is usually chronic but under stressful conditions can be acute. Motile Aeromonas septicemia (Aeromonas hydrophila) infections in eels are not unusual in freshwater (Chapter 11). Necrotic epidermal lesions develop on the head, opercula, or in the lateral musculature in freshwater. This chronic to subacute disease results in significant losses and is usually associated with some type of environmental stress, handling, or transport. Infections of atypical Aeromonas salmonicida have been infrequently reported in cultured eels (See Chapter 12). The disease follows a similar pattern to that which occurs in carp and goldfish in which necrotic ulcers are generally confined to the skin, and losses are more or less chronic. Vibriosis has been recognized disease in eels in brackish waters for many years (See Chapter 14). In fact, the specific epithet of Vibrio anguillarum (pseudonym Listonella anguillarum [MacDonnell and Colwell 1985]) was derived from a disease described in eels in Italy during the nineteenth century. Infection produces shallow ulcerative lesions at any body location, while eels are in salt or brackish water. Vibrio vulnificus has been reported to cause major disease problems in intensively cultured eels in Japan and the Netherlands (Austin and Austin 1987; Dalsgaard et al. 1999). The major isolate of V. vulnificus is serovar E (biotype 2) in Denmark is usually associated with brackish water culture units (Fouz et al. 2001). The isolate from a Danish eel farm was serologically identical to V. vulnificus (ATCC 33149) in Japan. Unpublished information suggests that

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V. vulnificus has provoked the closure of several farms in Europe and Canada (Fouz et al. 2001). The pathogen is also capable of opportunistically infecting humans.

Management of eel bacterial diseases Control of bacterial diseases of cultured eels revolves around good aquaculture management, use of chemotherapy, and vaccination when indicated.

Management Maintaining a high quality environment by keeping organic enrichment low and preventing shock and extreme temperature fluctuations are proactive approaches to management of eel diseases. Highly enriched pond waters appear to enhance occurrence of Edwardsiellosis especially in eels. Because fish, turtles, snakes, etc. may carry E. tarda in their intestines, it is desirable when practical to prevent fish contact with these potential reservoirs. Because red spot disease has a tendency to occur in brackish water ponds at temperatures between 15◦ C and 20◦ C, Muroga (1978) proposed the following management practices for controlling the disease in cultured Japanese eel: (1) maintain water temperature above 27◦ C, (2) culture eels in freshwater ponds, and (3) culture less susceptible European eels where disease is endemic.

Chemotherapeutics Application of medicated feed is the most commonly used control for bacterial eel infections. In Taiwan the drug of choice for treating E. tarda eel infections is nalidixic acid because a large percentage of isolates are resistant to  R most other antibiotics including Terramycin

(Liu and Wang 1986). In the United States, Terramycin (at a rate of 50–100 mg/kg of fish per day for 10 days) is the drug of choice  R for treating E. tarda, but Romet-30 is also widely used and is fed at 50 mg/kg of fish per day for 5 days; neither of these drugs is FDA approved for treating E. tarda infections and requires a cross-label application procedure. For fish not skinned during dressing, a withdrawal period for Terramycin is 21 days and 42 for Romet-30. A 3 day withdrawal period is required for Romet in catfish. Norfloxacin was shown to be highly effective against E. tarda in vitro (MIC of 0.016–0.031 µg/mL) and when incorporated into feed of naturally E. tarda-infected Japanese flounder for 3 days at 100 mg/kg of body weight (Heo and Kim 1996). Antibiotic sensitivity/resistance of E. tarda has been studied by several researchers and most concur that it is a significant problem. Fifty-four strains of E. tarda from cultured eels in Taiwan (Chen et al. 1984) were highly sensitive to gentamicin and sulfa-drugs; Waltman and Shotts (1986) also found that a high percentage of E. tarda isolates from Taiwan were resistant to numerous antimicrobials. Liu and Wang (1986) reported that nearly 92% of E. tarda isolates taken from water in eel culture ponds demonstrated some degree of antibiotic resistance. Antibiotic resistance of E. tarda in the United States has not yet become a significant problem (Plumb et al. 1995). Medicated feed is also used with some success to control P. anguilliseptica infections in eels and other fish. Wicklund and Bylund (1990) found that oxytetracycline was not effective when used in P. anguilliseptica infected salmon but Romet-30 was effective. Red spot disease of Japanese eels was controlled by 9 consecutive days of baths in oxolinic acid or nalidixic acid (2–10 mg/L) with 0 mortalities compared to 100% mortality in nontreated controls (Jo 1978). Also, when fish were fed oxolinic acid at 5–20 mg/kg of fish for 3 consecutive days 100% survived. No antibiotic resistance was noted in V. vulnificus-infecting

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eel in Denmark; however, treatment with antibiotics had little effect on the disease as it reoccurred (Dalsgaard et al. 1999). Antibiotic therapy is sometimes indicated, but managers should emphasize proactive reduction of environmental stressors and application of effective vaccines when available.

Vaccination Salati (1988) reviewed techniques and procedures for vaccinating eels against E. tarda and found that the two basic types of vaccines used were whole cell bacterins and bacterial extracts. Song and Kou (1981) reported that a single immersion of 6 g eelvers was effective against E. tarda but two or three vaccine exposures proved more effective in eliciting immunity that lasted for 10 weeks. Following eel immunization with E. tarda LPS, culture filtrates, and formalin-killed cells, Salati et al. (1983) concluded that LPS was the immunogenic component. It was later shown that polysaccharide, without the lipid component, was more antigenic than other preparations (Salati and Kusuda 1985). Eels immunized by injection with formalin killed E. tarda cells and LPS showed only slight to moderate protection from injection challenge with virulent E. tarda 21 days postvaccination (Gutierrez and Miyazaki 1994). Studies in Japanese flounder have shown marginal success when E. tarda extracellular and intracellular products were used in injection, immersion, and orally administered vaccines (Rashid et al. 1994a). While serum agglutinating antibodies increased following injection and oral exposure, no clear protection was acquired. With regard to E. tarda vaccine, Salati (1988) concluded that bacterins that are simple and cheap to produce on a large scale do provide protection to anguillettes, and while protection is not complete, it may be feasible for use on farms where Edwardsiellosis is severe. All of these preparations were immunogenic, especially by in-

jection, but a commercial vaccine is not available. Eel vaccination experiments with P. anguilliseptica have shown that formalin-killed bacterins are effective immunogens when applied by injection (Nakai et al. 1982). Injected vaccines stimulated high agglutinating antibodies that were highly protective but vaccination by injection is probably not practical. Immersion apparently had no immunological effect. Fouz et al. (2001) vaccinated 9.5 million glass eel on farms in Spain and Denmark against V. vulnificus (serovar E, (biotype 2) by immersion and boosted 12–14 and 24–28 days later. This protocol resulted in a relative percentage survival of 62–86% in vaccinates. The authors concluded that vaccination by immersion seems to be the best preventive method against V. vulnificus infections.

References Amandi, A., S. F. Hiu, et al. 1982. Isolation and characterization of Edwardsiella tarda from fall chinook salmon (Oncorhynchus tshawytscha). Applied Environmental Microbiology 43:1380– 1384. Austin, B., and D. A. Austin. 1987. Bacterial Fish Pathogens: Disease in Farmed and Wild Fish. Chichester, England, Ellis Horwood. Benli, A. C. K. and H. Y. Yildiz. 2004. Blood parameters in Nile tilapia (Oreochromis niloticus L.) spontaneously infected with Edwardsiella tarda. Aquaculture Research 35:1388–1390. Berthe, F. C. J., C. Miche., and J. F. Bernardet. 1995. Identification of Pseudomonas anguilliseptica isolated from several fish species in France. Diseases of Aquatic Organisms 21:145–150. Blanco, M. M., A. Gibello, et al. 2002. PCR detection and PFGE DNA macrorestriction analysis of clinical isolates of Pseudomonas anguilliseptica from winter disease outbreaks in sea bream Sparus aurata. Diseases of Aquatic Organisms 50:19–27. Bylund, G. 1983. The health of Baltic herring and the symptoms of eye disease in fish in Pori coastal waters. Meri 12:203–212.

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CAB International. 2006. Datasheet on Edwardsiella septicemia. (original text by Plumb, J., and Evans, J.; revision by Evans, J.). In: Aquaculture Compendium. Online at www.cabi compendium.org/as and on CD-ROM. Wallingford, UK. Chen, S. C., M. C. Tung, and S. T. Huang. 1984. Sensitivity in vitro of various chemotherapeutic agents to Edwardsiella tarda of pond-cultured eel. COA Fisheries Series No. 10, Fish Disease Research (VI) 2:135–141. Chowdhury, M. B. R., and H. Wakabayashi. 1990. Survival of four major bacterial fish pathogens in different types of experimental water. Bangladesh Journal of Microbiology 7:47–54. Clarridge, J. E., D. M. Musher, et al. 1980. Extraintestinal human infection caused by Edwardsiella tarda. Journal of Clinical Microbiology 11:511–514. Coles, B. M., R. K. Stroud, and S. Sheggeby. 1978. Isolation of Edwardsiella from 3 Oregon sea mammals. Journal of Wildlife Diseases 14:339–341. Dalsgaard, I., L. Hoi, et al. 1999. Indole-positive Vibrio vulnificus isolated from disease oiutbreaks on a Danish eel farm. Diseases of Aquatic Organisms 35:187–194. Darwish, A., J. A. Plumb, and J. C. Newton. 2000. Histopathology and pathogenesis of experimental infection with Edwardsiella trda in channel catfish. Journal of Aquatic Animal Health 12:255–266. Earlix, D. 1995. Host, pathogen, and environmental interactions of enteric septicemia of catfish. Doctoral Dissertation, Auburn University, Alabama. Egusa, S. 1976. Some bacterial diseases of freshwater fishes in Japan. Fish Pathology 10:103–114. Ellis, A. E., G. Dear, and D. J. Steward. 1983. Histopathology of Sekiten-byo caused by Pseudomonas anguilliseptica in the European eel, Anguilla anguilla L., in Scotland. Journal of Fish Diseases 6:77–79. Farmer, J. J., and A. L. McWhorter. 1984. Genus X Edwardsiella Ewing and McWhorter 1965, 37AL. In: Bergey’s Manual of Systematic Bacteriology, Vol. I. Baltimore, Williams and Wilkins, pp. 486–491. Fouz, B., M. D. Esteve-Gassent, et al. 2001. Field testing of a vaccine against eel diseases caused

by Vibrio vulnificus. Diseases of Aquatic Organisms 45:183–189. Francis-Floyd, R., P. Reed, et al. 1993. An epizootic of Edwardsiella tarda in largemouth bass Micropterus salmoides). Journal of Wildlife Diseases 29:334–336. Gilman, R. H., M. Madasamy, et al. 1971. Edwardsiella tarda in jungle diarrhea and a possible association with Entamoeba histolytica. Southeast Asian Journal of Tropical Medicine and Public Health 2:186–189. Grimont, P. A. D., F. Grimont, et al. 1980. Edwardsiella hoshinae, a new species of Enterobacteriaceae. Current Microbiology 4:347–351. Gutierrez, M. A., and T. Miyazaki. 1994. Responses of Japanese eels to oral challenge with Edwardsiella tarda after vaccination with formalin-killed cells or polysaccharide of the bacterium. Diseases of Aquatic Organisms 6:110–117. Han, H. J., D. H. Lee, et al. 2006. Pathogenicity of Edwardsiella tarda to olive flounder, Paralichthys olivaceus (Temmick and Schlegel). Journal of Fish Diseases 29:601–609. Haenen, O. L. M., and A. Davidse. 2001. First isolation and pathogenicity studies with Pseudomonas anguilliseptica from diseased European eel Anguilla anguilla (L.) in The Netherlands. Aquaculture 196:27–36. Hargreaves, J. E., and D. P. Lucey 1990. Edwardsiella tarda soft tissue infection associated with catfish puncture wound. Journal of Infectious Diseases 162:1416–1417. Heo, G. J., and J. H. Kim. 1996. Efficacy of norfloxacin for the control of Edwardsiella tarda infection in the flounder Paralichthys olivaceus. Journal of Aquatic Animal Health 8:255– 259. Hidaka, T., Y. Yanohara, and T. Shibata. 1983. On the causative bacterial of head ulcer disease in cultured eels Anguilla japonica. Memorandum of the Faculty of Fisheries Kagoshima University 32:147–166. Hoshina, T. 1962. On a new bacterium, Paracolobactrum anguillimortiferum sp. Bulletin of the Japanese Society of Scientific Fisheries 28:162–164. Huang, S. T., and C. I. Liu. 1986. Experimental studies on the pathogenicity of Edwardsiella tarda and Aeromonas hydrophila in eel, Anguilla japonica. COA Fisheries Series, No. 8. Fish Disease Research ( VIII): 40–55.

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Humphrey, J. D., C. Lancaster, et al. 1986. Exotic bacterial pathogens Edwardsiella tarda and Edwardsiella ictaluri from imported ornamenta fish Betta splendens and Puntius conchonius respectively; isolation and quarantine significance. Australian Veterinary Journal 63:93–69. Janda, J. M., and S. L. Abbott. 1993. Expression of an iron-regulated hemolysin by Edwardsiella tarda. FEMS Microbiology Letters 111:275–280. Janda, J., S. L. Abbott, et al. 1991. Pathogenic properties of Edwardsiella species. Journal of Clinical Microbiology 29:1997–2001. Jo, Y. 1978. Therapeutic experiments on red spot disease. Fish Pathology 13:41–42. Jo, Y., R. Muroga, and K. Onishi. 1975. Studies on red spot disease of pond cultured eels. III. A case of the disease in the European eels (Anguilla anguilla) cultured in Tokushima Prefecture. Fish Pathology 9:115–118. Kodoma, H., T. Murai, et al. 1987. Bacterial infection which produces high mortality in cultured Japanese flounder Paralichthys olivaceus in Hokkaido Japan. Japanese Journal of Veterinary Research 35:227–234. Kuo, S.-C., H.-Y. Chung, and G-H Kou. 1977. Edwardsiella anguillimortifera isolated from edwardsiellosis of cultured eel (Anguilla japonica). JCRR Fisheries Series No. 29, Fish Disease Research ( I): 1–6. Kumar, G., G. Rathore, et al. 2007. Isolation and characterization of outer membrand proteins of Edwardsiella tarda and its application in immunoassays. Aquaculture 272:98–104. Kusuda, R., N. Dohata, et al. 1995. Pseudomonas anguilliseptica infection of striped jack. Fish Pathology 30:121–122. Liu, C. I., and S. S. Tsai. 1982. Edwardsiellosis in pond-cultured eel in Taiwan. CAPD Fisheries Series No. 8, Report on Fish Disease Research IV:92–95. Liu, C. I., and J. H. Wang. 1986. Drug resistance of fish- pathogenic bacteria. II. Resistance of Edwardsiella tarda in aquaculture environment. COA Fisheries Series No. 8, Fish Disease Research 8:56–67. Lonnstr om, L., T. Wicklund, and G. Bylund. ¨ ¨ 1994. Pseudomonas anguilliseptica isolated from Baltic herring Clupea harengus membras with eye lesions. Diseases of Aquatic Organisms 18:143–147.

Lopez-Romalde, S., B. Magarinos, et al. 2003. Phenotypic and genetic characterization of Pseudomonas anguilliseptica strains isolated from fish. Journal of Aquatic Animal Health 15:39–47. MacDonnell, M. T., and R. R. Colwell. 1985. Phylogeny of the Vibrionaceae, and recommendation for two new genera, Listonella dnd Shewanella. Systematic Applied Microbiology 6:171–182. Maiti, N. K., A. Mandal, et al. 2008. Comparative analysis of genome of Edwardsiella tarda by BOX-PCR-ribotyping. Aquaculture 280:60– 63. Maiti, N. K., A. Mandal, et al. 2009. Phenotypic and genetic characterization of Edwardsiella tarda isolated from pond sediments. Comparative Immunology, Microbiology and Infectious Disease 32:1–8. Martin, J. D. 1983. Atlantic salmon and alewife passage through a pool and weir fishway of the Mayaguadaric River, New Brunswick, Canada during 1983. Canadian Annual Report on Fisheries and Aquatic Science (Abstract). Mekuchi, T., T. Kiyokawa, et al. 1995. Infection experiments with Edwardsiella tarda in the Japanese flounder. Fish Pathology 30:247– 250. Meyer, F. P., and G. L. Bullock. 1973. Edwardsiella tarda, a new pathogen of channel catfish (Ictalurus punctatus). Applied Microbiology 25:155–156. Miwa, S., and N. Mano. 2000. Infection with Edwardsiella tarda causes hypertrophy of liver cells in the Japanese flounder Paralichthys olivaceus. Diseases of Aquatic Organisms 42:277–231. Miyazaki, T., and S. Egusa. 1976a. Histopathological studies of edwardsiellosis of Japanese eel (Anguilla japonica). I. Suppurative interstitial nephritis form. Fish Pathology 11:33–43. Miyazaki, T., and S. Egusa. 1976b. Histopathological studies of edwardsiellosis of Japanese eel (Anguilla japonica). II. Suppurative hepatitis form. Fish Pathology 11:67–75. Miyazaki, T., and S. Egusa. 1977. Histopathological studies of red spot disease of the Japanese eel (Anguilla japonica). I. Natural infection. Fish Pathology 12:39–47. Miyazaki, T. M. A. Gutierrez, and S. Tanaka. 1992. Experimental infection of Edwardsiellosis in the Japanese eel. Fish Pathology 27:39–47.

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Miyazaki, T., and N. Kaige. 1985. Comparative histopathology of edwardsiellosis in fishes. Fish Pathology 20: 219–227. Muroga, R. 1978. Red spot disease of eels. Fish Pathology 13:35–39. Muroga, K., Y. Jo, and T. Sawada. 1975. Studies on red spot disease of pond-cultured eels. II. Pathogenicity of the causative bacterium, Pseudomonas anguilliseptica. Fish Pathology 9:107–114. Muroga, K., Y. Jo, and M. Yano. 1973. Studies on red spot disease of pond-cultured eels. I. The occurrence of the disease in eel culture ponds in Tokushima Prefecture in 1972. Fish Pathology 8:1–9. Mushiake, K., K. Muroga, and T. Nakai. 1984. Increased susceptibility of Japanese eel Anguilla japonica to Edwardsiella tarda and Pseudomonas anguilliseptica following exposure to copper. Bulletin of the Japanese Society of Scientific Fisheries 50:1797–1801. Nakai, T., H. Hanada, and K. Muroga. 1985a. First records of Pseudomonas anguilliseptica infection in cultured ayu Plecoglossus altivelis. Fish Pathology 20:481–484. Nakai, T., K. Muroga, et al. 1982. Studies on red spot disease of pond-cultured eels. IX. A field vaccination trial. Aquaculture 3:131– 135. Nakai, T., K. Muroga, et al. 1985b. A serological study on Pseudomonas anguilliseptica isolated from diseased eels in Taiwan. Fish Pathology 19:259–261. Nakai, T., K. Muroga, and H. Wakabayashi. 1981. Serological properties of Pseudomonas anguilliseptica in agglutination. Bulletin of the Japanese Society of Scientific Fisheries 47:699–703. Nash, G., I. G. Anderson, et al. 1987. Bacteriosis associated with epizootic in the giant sea perch, Lates calcarifer, and the estuarine grouper, Epinephelus tauvine, cage cultured in Malaysia. Aquaculture 67:105–111. Nichibuchi, M., K. Muroga, and Y. Jo. 1980. Pathogenic Vibrio isolated from eels, Diagnostic tests for the disease due to present bacterium. Fish Pathology 14:125–131. Palleroni, N. J. 1984. Family I. Pseudomonadaceae Winslow, Broadhurst, Buchanan, Krumwiede, Rogers and Smith 1917. In: N. R. Krieg and J. G. Holt (eds) Berqey’s Manual of Systematic Bacte-

riology, Vol I. Baltimore, Williams & Wilkins, pp. 141–199. Panangala, V. S., V. L. van Santon, et al. 2005. Analysis of 16S–23S intergenic spacer regions of the rRNA operons in Edwardsiella ictaluri and Edwardsiella tarda isolated from fish. Journal of Applied Microbiology 99:657–669. Panangala, V. S. C. A. Shoemaker, et al. 2006. Intra- and interspecific phenotypic characteristics of fish-pathogenic Edwardsiella ictaluri and E. tarda. Aquaculture Research 37:49–60. Park, S., H. Wakabayashi, and Y. Watanabe. 1983. Serotype and virulence of Edwardsiella tarda isolated from eel and their environment. Fish Pathology 18:85–89. Plumb, J. A., C. C. Sheifinger, et al. 1995. Susceptibility of six bacterial pathogens of channel catfish to six antibiotics. Journal of Aquatic Animal Health 7:211–217. Rashid, M. M., K. Honda, et al. 1994b. An ecological study on Edwardsiella tarda in flounder farms. Fish Pathology 29:221–227. Rashid, M. M., T. Mekuchi, et al. 1994a. A serological study on Edwardsiella tarda strains isolated from flounder (Paralichthys olivaceus). Fish Pathology 29:277. Salati, F. 1988. Vaccination against Edwardsiella tarda. In: A. E. Ellis (ed.) Fish Vaccination. London, Academic Press, pp. 135–151. Salati, F., K. Kawai, and R. Kusuda. 1983. Immunoresponse of eel against Edwardsiella tarda antigens. Fish Pathology 18:135–141. Salati, F., and R. Kusuda. 1985. Chemical composition of the lipopolysaccharide from Edwardsiella tarda. Fish Pathology 20:187–191. Shotts, E. B., and J. D. Teska. 1989. Bacterial pathogens of aquatic vertebrates. In: B. Austin and D. A. Austin (eds) Methods for Microbiological Examination of Fish and Shellfish. Chichester, Ellis Horwood, pp. 164–186. Song, Y. L., and G. H. Kou. 1981. The immunoresponses of eel (Anguilla japonica) against Edwardsiella anguillimortifera as studied by the immersion method. Fish Pathology 15:249–255. Stewart, D. J., R. Woldemariam, et al. 1983. An outbreak of ‘Sekiten-byo’ among cultured European eels, Anguilla anguilla L., in Scotland. Journal of Fish Diseases 6:75–76. Suprapto, H., T. Nakai, and K. Muroga. 1995. Toxicity of extracellular products and intracellular components of Edwardsiella tarda in the

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Japanese eel and flounder. Journal of Aquatic Animal Health 7:292–297. Swain, P. and S. K. Nayak. 2003. Comparative sensitivity of different serological tests for seromonitoring and surveillance of Edwardsiella tarda infection of Indian major carps. Fish and Shellfish Immunology 15:333–340. Taylor, P. W., J. E. Crawford, and E. B. Shotts, Jr. 1995. Comparison of two biochemical tests systems with conventional methods for identification of bacteria pathogenic to warmwater fish. Journal of Aquatic Animal Health 7:312–317. Uhland, F., C. P. Helie, and R. Higgins. 2000, Infections of Edwardsiella tarda among brook trout in Quebec. Journal of Aquatic Animal Health 12:74–77. Ullah, A., and T. Arai. 1983. Pathological activities of the naturally occurring strains of Edwardsiella tarda. Fish Pathology 18:65–70. van Damme, L. R., and J. Vandepitte. 1980. Frequent isolation of Edwardsiella tarda and Plesiomonas shigelloides from healthy Zairese freshwater fish: A possible source of sporadic diarrhea in the tropics. Applied Environmental Microbiology 39:475–479. Wakabayashi, H., and S. Egusa. 1972. Characteristics of a Pseudomonas sp. from an epizootic of pond-cultured eels (Anguilla japonica). Bulletin of the Japanese Society of Scientific Fisheries 38:577–587.

Wakabayashi, H., and S. Egusa. 1973. Edwardsiella tarda (Paracolobactrum anguillimortifera) associated with pond-cultured eel disease. Bulletin of the Japanese Society of Scientific Fisheries 39:931–936. Walters, G., and J. A. Plumb. 1980. Environmental stress and bacterial infection in channel catfish, Ictalurus punctatus Rafinesque. Journal of Fish Biology 17:177–185. Waltman, W. D., and E. B. Shotts. 1986. Antimicrobial susceptibility of Edwardsiella tarda from the United States and Taiwan. Veterinary Microbiology 12:277–282. Waltman, W. D., E. B. Shotts, and T. C. Hsu. 1986. Biochemical and enzymatic characterization of Edwardsiella tarda from the United States and Taiwan. Fish Pathology 21:1–8. White, F. H., F. C. Neal, et al. 1969. Isolation of Edwardsiella tarda from an ostrich and an Australian skink. Journal of the American Veterinary Medical Association 155:1057– 1058. Wicklund, T., and G. Bylund 1990. Pseudomonas anguilliseptica as a pathogen of salmonid fish in Finland. Diseases of Aquatic Organisms 8:13–19. Wyatt, L. E., R. Nickelson II, and C. Vanderzant. 1979. Edwardsiella tarda in freshwater catfish and their environment. Applied Environmental Microbiology 38:710–714.

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Chapter 14

Salmonid bacterial diseases

A certain mystique is associated with salmonids that no other family of fish enjoys, and this in part explains the attention they receive from environmentalist, aquaculturist, fish pathologist, and conservation agencies. Moreover, the high economic value of both wild and cultured salmonids demands that considerable attention be directed toward their health management. Bacterial infections that affect salmonids occur worldwide and have significant impact on cultured and wild populations. These diseases include furunculosis (Aeromonas salmonicida), vibriosis (Vibrio anguillarum and related species), cold-water vibriosis (Vibrio salmonicida), enteric redmouth (Yersinia ruckeri), bacterial gill disease (Flavobacterium branchiophilum), bacterial cold-water disease (F. psychrophilum), bacterial kidney disease (Renibacterium salmoninarum), and several other less-specific or wellknown diseases. Most of the salmonid bacterial pathogens are identified by conventional bacteriological procedures based on morphological and biochemical characteristics. However, serological procedures are widely used and molecular

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.

techniques for salmonid pathogens described in Cunningham (2002) are becoming more wide spread.

Furunculosis Furunculosis is one of the oldest and best known bacterial fish diseases having been first reported late in the nineteenth century. It is generally considered a disease of salmonids but is increasingly associated with other cool and occasionally warm-water fishes in both fresh and saltwater. Its etiological agent is A. salmonicida subsp. salmonicida, which is also known as “typical A. salmonicida” (Shotts and Bullock 1975). “Typical” A. salmonicida subsp. salmonicida, the cause of furunculosis in salmonids, should not be confused with “atypical” A. salmonicida, the cause of “ulcer disease” in cyprinids and other non-salmonid fishes. There are at least three subspecies in the atypical group: A. salmonicida subsp. achromogens, A. salmonicida subsp. masoucida, and A. salmonicida subsp. smithia that are discussed in Chapter 12 (Carp and Minnow Bacterial Diseases). For practical reasons, reference to furunculosis or A. salmonicida in this chapter indicates typical A. salmonicida subsp. salmonicida. 345

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Geographical range and species susceptibility Furunculosis is found in most regions of the world where salmonids occur, including Europe, Japan, Korea, North America, South Africa, and the United Kingdom (McCarthy 1975; Trust et al. 1980; Wichardt et al. 1989). Brook trout are generally considered the most susceptible trout species; however, there are resistant strains within this species. But all salmonids, including brown, rainbow, and lake trout, and anadromous Atlantic and Pacific salmon are susceptible to the disease. Wichardt et al. (1989) indicated that in Sweden, acute typical A. salmonicida infections also occur in Arctic char. Nomura et al. (1992) noted that in Japan, rainbow trout are not severely affected by A. salmonicida but cultured amago and masu salmon are susceptible, and as culture of the latter species increases A. salmonicida’s impact on them is likely to also increase. However, in experimental A. salmonicida infections of different pink salmon populations, no difference in susceptibility was evident (Beacham and Evelyn 1992a). A. salmonicida subsp. salmonicida may occur in species of cyprinids, pike, perch, bullheads, and experimentally in some nonsalmonid marine fish species such as sac-larvae turbot and halibut (Fryer and Rohovec 1984; Ostland et al. 1987; Bergh et al. 1997). Nonsalmonids become infected usually when they have been in close proximity to A. salmonicida infected salmonids. Natural and experimental A. salmonicida transmission to a species of wrasse was reported by Bricknell et al. (1996), but these fish were not as susceptible as Atlantic salmon. A serious outbreak of A. salmonicida occurred in wild smallmouth bass and redbreasted sunfish in three western Virginia rivers (Cipriano and McDaniel 2008).

Clinical signs Furunculosis is classified into four categories based upon severity: acute, subacute, chronic, or latent. Acute infection with sudden mass deaths without notable or marked external clinical signs, more commonly in juvenile salmonids, is usually associated with a septicemia. Older fish with A. salmonicida septicemia are listless and drift down stream or lie on the bottom of ponds, tanks, or net cages where they respire weakly before death. Few external clinical signs accompany acute furunculosis, but fish exhibit darkened skin pigmentation, lethargic swimming, and inappetence. Discrete petechia may occur at the base of fins. Internally, the spleen may be enlarged and septicemia develops with high mortalities occurring within 2 to 3 days. Clinical signs in older fish with subacute infection develop more gradually but are associated with high morbidity accompanied by a consistent rise in deaths. Various clinical signs are apparent, including dark pigmentation, “furuncles” or boil like lesions on the body surface, and hemorrhage at the base of frayed fins (Figure 14.1). In coho salmon, subdermal hemorrhaging resembling a bruise (hematoma) with a purplish color occurs but the skin is not ruptured. Internally, the peritoneal cavity may contain a bloody, cloudy fluid and the intestine is flaccid, inflamed, and filled with a bloody discharge that exudes from a red, prolapsed anus. The kidney is generally edematous and hemorrhagic, the spleen is usually enlarged and dark red, and the liver is pale. A general hyperemia is evident throughout the viscera. Chronic furunculosis results in low grade but consistent mortality over an extended period of time. Clinical findings described for subacute infections are also dramatically apparent in chronically infected moribund fish but prevalence throughout the population is

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Figure 14.1 Brown trout with furuncle (arrow), caused by Aeromonas salmonicida. (Photo courtesy of the U. S. Fish and Wildlife Service.)

lower. An inflamed intestine is often the only internal pathological manifestation. In the latent form of furunculosis, A. salmonicida lies dormant in a fish population with fish showing no clinical signs or internal pathology; however, they are still considered pathogen carriers. Nomura et al. (1992) indicated that salmon without frank infection had A. salmonicida in kidney and coelomic fluid, but Cipriano et al. (1992) successfully isolated the organism from skin mucus in a higher percentage of fish than from internal organs. They isolated A. salmonicida from skin mucus taken by nonlethal sampling; mucus from 56% of infected lake trout was positive compared to isolation from only 6% of kidneys. A similar ratio was seen in mucus versus kidney of Atlantic salmon.

Diagnosis External and internal visual observations are helpful in presumptively diagnosing A. salmonicida, but isolation of the bacterium from muscle lesions, kidney, spleen, or liver of diseased fish is essential for positive identification. A. salmonicida grows on TSA and

BHI agar or on most other general bacteriological media when incubated at less than 25◦ C; preferably at 20◦ C. Typical A. salmonicida grown on media with tyrosine produces a brown, water-soluble pigment that diffuses throughout the media in 2–4 days when incubated below 25◦ C. The pathogen grows poorly at 30◦ C and not at all at 37◦ C. Upon initial isolation, most colonies of A. salmonicida are hard and friable with some being smooth, soft, and glistening. Bacteria in hard colonies (rough) will autoagglutinate in broth and are virulent, while bacteria in smooth colonies do not autoagglutinate and are avirulent (Munro and Hastings 1993). Autoagglutination and virulence are associated with the presence of an additional cell-surface protein called the “A layer” (Udey and Fryer 1978). Application of the PCR method of identifying pathogens is becoming more widespread and applicable. Byers et al. (2002) used A. salmonicida subsp. salmonicida in infected salmonids and demonstrated that these PCR assays could replace biochemical tests to confirm identity of the pathogen. These PCR tests were also capable of direct detection of A. salmonicida in mucus, gill, and kidney samples from overtly infected fish. However,

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conventional bacterial culture was more reliable for detection of the pathogen in covertly infected salmonids than was direct PCR testing of tissues. It is not always possible to inoculate bacterial culture media on site of an epizootic or to perform nonlethal assays in which transport of samples from assayed fish to a laboratory is necessary. Fish can be iced down for short periods of time; however, Cipriano and Bullock (2001) evaluated commercial transport systems, Ames, Stuart’s, and Cary-Blair media, for transport of mucus for detection of A. salmonicida in an Atlantic salmon population. They determined that transport media was a suitable transport of mucus with A. salmonicida for as long as 48 hours at 18–20◦ C without significant increase or decrease of bacterial cell numbers.

Bacterial characteristics In addition to the presumptive characteristics noted above, typical A. salmonicida are small (0.9 × 1.3 µm), Gram-negative, nonmotile rods that ferment and oxidize glucose and are catalase and cytochrome oxidase positive (Shotts and Bullock 1975). Definitive identification is achieved by additional biochemical reactions (Table 12.1). A comparison of the biochemical reaction between typical A. salmonicida (salmonicida) and atypical A. salmonicida (achromogens) using API 50E and conventional tube media shows distinct differences in several key reactions (Hirvela¨ Koski et al. 1994). Typical A. salmonicida subsp. salmonicida is morphologically and biochemically homogeneous with some exceptions, the most notable being a variation in pigment production, Voges-Proskauer test results, and its ability to ferment selected sugars (Paterson et al. 1980; Popoff 1984). Also some isolates of typical A. salmonicida lose their ability to produce brown pigment after numerous transfers on media in the laboratory. Some motile A. hy-

drophila isolates acquire the ability to produce similar pigment after numerous subcultures but color does not develop as rapidly or intensely as A. salmonicida. Identification of A. salmonicida can be made more rapidly with serological rather than cultural procedures using serum agglutination, fluorescent antibody, ELISA, or polymerase chain reaction (PCR) techniques on either infected tissue or cultured bacterium. A specific DNA probe was developed with PCR by Mooney et al. (1995), a method that detected A. salmonicida DNA in 87% of wild Atlantic salmon in three Irish river systems. McCarthy and Whitehead (2008) described an immuno-India ink serological technique that permits laboratory diagnosis of furunculosis in less than 15 minutes from receipt of the specimen. There appears to be only one serological strain of A. salmonicida subsp. salmonicida. McCarthy and Rawle (1975) showed a strong serological homogeneity of typical A. salmonicida strains, but these strains will not cross-react to other aeromonads. Monoclonal antibody against A. salmonicida salmonicida lipopolysaccharides further demonstrates the homogeneity of this antigenic property (Rockey et al. 1988). Aoki and Holland (1985) found that at least three major proteins of the outer membranes of different A. salmonicida isolates were similar.

Epizootiology Since early descriptions of furunculosis, A. salmonicida has been considered an obligate pathogen passed from fish to fish or generation to generation through carrier fish (Popoff 1984). It was believed to have only a short life span outside its host; however, Michel and Dubois-Darnaudpeys (1980) demonstrated that the organism will survive and retain its pathogenicity for 6–9 months in pond bottom mud. Allen-Austin et al. (1984) reported that the bacterium will remain viable

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in freshwater at undetectable levels for over 1 month, but Rose et al. (1990) showed that A. salmonicida survived for less than 10 days in seawater. Nomura et al. (1992) maintained viable A. salmonicida for 60 days in sterilized freshwater but for only 4 days in nonsterile water, concluding that the organism could survive in water long enough to infect other fish. A. salmonicida survives for a short time in the marine environment; according to colony forming units on culture media, the bacteria disappeared in about 10 days after seeding into a sterilized environment with 25 ppt salinity and 16–22◦ C (Effendi and Austin 1994). However, by a variety of direct counts and other detection methods, they noted that the organism did not become less numerous. Also, in nonsterile environments and substrates, the organism did not survive much past 10 days, but on the same sterile microcosms, there was very little loss of cell numbers at 20 days, leaving the question of whether or not these remaining cells were viable and pathogenic, unanswered. In view of the discrepancies in these studies, and in spite of data supporting its extended survival outside of the host, A. salmonicida is still considered to be an obligate pathogen. Mortality rates of salmonids infected with typical A. salmonicida can vary from none in the latent or carrier state to very high in the acute state. Most commonly in subacute or chronic infections, the mortality will be moderate to high. Furunculosis is transmitted from fish to fish by contact either through the skin or by ingestion. Skin lesions, caused by injury or parasites, will enhance bacterial invasion. Cipriano (1982) demonstrated contact exposure of furunculosis in brook trout. Carrier fish, which show no overt signs of disease and from which bacteria cannot be easily isolated, can still be implicated in disease transmission. Infected nonsalmonid fish can also serve as pathogen reservoirs; Ostland et al. (1987) reported transmission of A. salmonicida from infected common shiners to coho salmon and brook trout.

Furunculosis has occasionally been isolated from non-hatchery fish in natural waters, but in most instances the affected fish were salmonids. In 2004, diseased smallmouth bass and redbreasted sunfish were found in the North Fork, South Fork, and main stream Shenandoah River in central Virginia. The river system lost about 80% of its adult smallmouth bass population; many moribund fish exhibited skin lesions indicative of bacterial infection. In the spring and fall of 2007, a study was conducted to establish the cause of the mortality in which moribund fish had clinical signs of infectious disease (Cipriano and McDaniel 2008). A. salmonicida subsp. salmonicida and other facultative bacteria were isolated from 8 of 10 necropsied fish from the North Fork and from 9 of 10 gills of fish from the South Fork. A. salmonicida was also isolated from smallmouth bass in the James, Cowpasture, and Jackson Rivers in west central Virginia. The study provided evidence that A. salmonicida was the putative etiological agent of lesions associated with smallmouth bass mortalities in the Shenandoah River but whether the pathogen was responsible for the extensive loss in 2004 is not clear; however, it clearly shows the pathogens potential to inflict disease in wild non-salmonid fish populations. Source of the A. salmonicida is unknown at this time. Contaminated water and/or equipment also serve as a source of disease transmission. In Sweden, fish at several trout and salmon farms became infected after A. salmonicida-infected fish were released into upstream water supplies (Wichardt et al. 1989). The role of aerosol transmission of A. salmonicida was investigated by Wooster and Bowser (1996), who found that the organism could travel at least 104 cm from its source through the air and then be recovered from water. This air borne potential makes management of A. salmonicida in closely associated culture units more critical. Experimentally, Hjeltnes et al. (1995) transmitted A. salmonicida to cultured cod,

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halibut, and wrasse from Atlantic salmon; however, this occurred rarely and did not result in serious disease in these species. Experiments by Bricknell et al. (1996) indicated that the pathogen could naturally infect wrasse but was unlikely to cause a primary infection; however, they served as pathogen reservoirs. According to P´erez et al. (1996), 30 g turbot in seawater were more susceptible to A. salmonicida than were 25 g rainbow trout in freshwater. Intragastric inoculation failed to result in infection in either species. Rainbow trout can carry A. salmonicida after initial infection for up to 2 years without re-exposure (Bullock and Stuckey 1975). It was also noted that when rainbow trout were stressed by water temperature of 18◦ C and injected with an immunosuppressant (Kenolog 40—triamcinolone acetonide), latently infected fish shed A. salmonicida. Following immunosuppression with Kenolog 40, 33% of the trout exhibited overt disease compared to 4% of fish stressed only by high water temperatures. There was also 73% mortality in chemically immunosuppressed fish compared to 33% mortality in those stressed solely by elevated water temperature. In natural trout, infections mortality can reach 5–6% per week and a total mortality of 85% has been reported in untreated populations. Nomura et al. (1992) isolated A. salmonicida from the kidney and coelomic fluid of chum, pink, and masu salmon, which showed no frank signs of disease after entering freshwater. Furthermore, the ability of typical A. salmonicida to occur and survive in wild fish populations was shown by Mooney et al. (1995), who used PCR to detect an 87% carrier rate in wild Atlantic salmon in Irish rivers. Although pathogen levels were extremely low in each fish, they speculated that the bacterium is wide spread among wild salmon populations in Irish rivers. When Cipriano et al. (1996a) examined an asymptomatic rainbow trout population in a recirculating water system that utilized a fluidized sand biofilter, A. salmonicida was iso-

lated from only 1% of spleens and no kidneys. However, the bacteria were isolated from 15% of gill and 19% of mucus samples. The pathogen was not isolated during subsequent examinations of the fluidized biofilters and tank water, thus indicating that the pathogen did not become established in the recirculation system. In a study by Nomura et al. (2002), evaluating the causative agent of furunculosis in chum salmon caught in coastal rivers in Japan found that the pathogen was present on the gill surface of 50% of the fish when caught and decreased on fish in holding ponds. The pathogen was not isolated from kidneys or intestine. They theorized that the bacteria were spread during fish migration within the river and during transport of fish from the capture site to holding ponds and suggested that furunculosis could be controlled in wild-caught salmon by stocking at low density in maturing ponds that should have been disinfected, and fish should be treated to reduce A. salmonicida on the gills. Risk factors involved in post-smolt Atlantic salmon contracting furunculosis after moving into sea cages were related to location, feeding, rearing, and presence of other disease organisms (Jarp et al. 1995). When a total of 116,480 fish died at 124 sea sites, 54% of the deaths resulted from furunculosis, 10.5% from vibriosis, and 39.5% from IPNV. According to McCarthy (1980), when an infected trout population overcomes clinical furunculosis, either as a result of natural defenses or chemotherapy, at least some survivors become carriers. In spite of extensive research in this area, the carrier state of A. salmonicida is still controversial but generally infected asymptomatic fish are considered the primary reservoir for typical A. salmonicida. For example, the bacterium was not isolated from any tissue of asymptomatic Atlantic salmon by Hiney et al. (1994). Upon stressing these same fish, A. salmonicida was isolated from the mucus, fins, and gills of about 50% of the fish and from about 10% of blood samples. Frequency of stress-induced infections was about

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65%. Before fish were stressed, an ELISA determined if A. salmonicida was present. The pathogen was found in 50% of intestinal samples but not in any kidney or mucus samples. Application of ELISA on poststressed fish indicated a positive reaction in about 25% of kidney samples and when results of kidney, intestinal content, and mucus were combined, 45% of the fish were positive. It was suggested that intestines may be the primary location of A. salmonicida in carrier fish and stress induces bacterial colonization of mucus on fins and gills. In comparative carrier experiments, P´erez et al. (1996) showed that rainbow trout that survived an A. salmonicida infection became pathogen carriers but infected turbot did not. In A. salmonicida carrier populations, clinical furunculosis can appear without apparent environmental insult to fish, but usually its recrudescence is stress mediated. If fish are stressed as a result of handling, elevated water temperatures, low oxygen, or other adverse environmental factors, a latent infection can become chronic, subacute, or acute. Kingsbury (1961) correlated outbreaks of furunculosis with depressed oxygen levels. When oxygen concentrations dropped to less than 5 mg/L during the night, losses due to A. salmonicida increased. Nomura et al. (1992) correlated A. salmonicida infection in chum salmon to fish density; infectivity was 12.4% in fish held at 14.7 fish/m2 compared to disease free fish stocked at 4.9 fish/m2 . They also found that A. salmonicida prevalence was significantly higher in fish held in water with low dissolved oxygen (6–7 mg/L) than in fish held in water with 10 mg/L O2 . Survival in high density-low oxygen water was approximately 40% less than in low density-high oxygen conditions. Water temperature also plays an important role in furunculosis outbreaks. In trout, the optimum water temperature for disease development is 15–20◦ C. When infected fish are held in water at 20◦ C, A. salmonicida in the blood increases dramatically within 48 hours, but at 10◦ C, 168 hours are re-

quired for bacteria to reach the same concentrations. In addition to temperature and oxygen concentration affecting appearance of furunculosis, fish density also affects the disease. In a transmission dynamics study, Ogut et al. (2005) stocked Chinook salmon at various densities and challenged each tank with a single A. salmonicida-infected fish. Furunculosisrelated mortality decreased as fish density decreased. The natural mortality rate followed a similar pattern but results indicated that transmission coefficient deaths due to A. salmonicida were dependent on fish density. Clearly the data presented above taken as a whole indicates that environmental conditions and associated stressors influence the occurrence and severity of furunculosis. A. salmonicida certainly occurs when diseased fish come in contact with naive fish but transmission of A. salmonicida during spawning has been thought to be important because the organism is readily isolated from ovaries, eggs, and ovarian fluid (McCarthy 1980). Nomura et al. (1992) showed that A. salmonicida on chum salmon eggs was naturally reduced by a log of about 106 colony forming units in 48 hours after manual stripping. They also showed that the organism could not be isolated from eggs after 5 days of incubation; therefore, they also concluded that vertical transmission of A. salmonicida was unlikely. Conversely, Bullock and Stucky (1975) were unable to establish spawning-related transmission in four attempts and concluded that vertical transmission does not occur. Furunculosis is most prevalent during late spring and summer when subacute epizootics often occur; however, the disease can occur at any time. Piper et al. (1982) noted that twice as many furunculosis cases occurred in July than in any other month. Some populations of brook and rainbow trout have been selected for their resistance to furunculosis and the use of these strains have helped reduce the impact of disease in certain areas (Ehlinger 1977). Naturally inherited

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defense mechanisms of trout account for part of their resistance to furunculosis, but serum bactericidal activity is also important (Hollebecq et al. 1995).

Pathology Death of trout infected with A. salmonicida is attributed to a massive septicemia and toxic extra cellular products (ECP) (toxins and enzymes) produced by the organism that interfere with the hosts blood supply, resulting in massive tissue necrosis. Skin lesions are characterized by loss of scales, necrosis of epithelium and muscle, capillary dilation, and peripheral hemorrhage of the lesion (McCarthy and Roberts 1980). Klontz et al. (1966) experimentally infected rainbow trout with A. salmonicida and found that the most consistent pathological aberration was an enlarged spleen 16 hours after injection that persisted throughout the infection. Sinusoids in the spleen became congested and engorged with erythrocytes. Bacteria could be isolated at 8 hours postinjection but not during the ensuing 48 hours at which time a septicemia developed. Extensive inflammation and muscle necrosis developed at the injection site. Using radiolabeled A. salmonicida, Svendsen et al. (1999) evaluated the disposition of the pathogen in Atlantic salmon following exposure by immersion. Three groups of fish were studied: (1) artificially wounded and (2) fish with reduced epidermal mucus layer caused by swabbing the skin and (3) uninjured control fish. Radiolabeled A. salmonicida was measured in mucus, skin, gills, wounded areas, blood, and five internal organs. The highest level of radioactivity was measured in the wound areas and the gills with a significant correlation between levels of radioactivity in gills, blood, mucus, and skin at 2 hours postchallenge. The bacterium was found 2 hours after challenge in blood of swabbed fish and injured fish but not in any tissues of control fish. These data indicate that A.

salmonicida infection is facilitated by wounds and removal of mucus from the skin. Histopathological changes were similar to those observed in naturally infected trout, especially in hematopoietic organs. Hematopoietic tissue of the kidney was most severely affected by increased lymphoid hemoblasts, macrophages, neutrophils, and lymphocytes until finally kidney hematopoiesis ceased. Renal tissues were largely spared from injury. ECP produced by A. salmonicida include several proteases, hemolysins, and a leukocidin (Munro et al. 1980; Cipriano et al. 1981; Titball and Munn 1981). A. salmonicida ECPs are capable of producing a syndrome similar to chronic furunculosis characterized by muscle necrosis and edema at the injection site. It appears that pathological manifestation of A. salmonicida can be attributed to a combination of cellular and extra cellular components. Pathogenicity of A. salmonicida was initially correlated with colony morphology on solid bacteriological media. Organisms from “smooth” colonies were considered pathogenic and those from “rough” colonies were not; however, this theory has been reversed and pathogenicity clarified. Udey and Fryer (1978) found that A. salmonicida virulence was associated with presence of an Alayer (additional-layer) and these cells actually formed the rough colonies but the A-layer was absent in bacteria in smooth nonvirulent strains that autoagglutinated. Other investigators concur that the A-layer (A-protein monomer) correlates with virulence (Evenberg and Lugtenberg 1982; Trust et al. 1983). The surface protein seems to protect A. salmonicida from its host’s natural defense mechanisms. The A-layer is composed of a 50 kD protein, the A-protein, which provides the pathogen with a protective barrier against the fish host defense mechanism. Lipopolysaccharide (LPS), another major cell envelope antigen, contains an O-polysaccharide (Oantigen), which is exposed at the cell surface. The O-antigen and A-protein appear to assist

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A. salmonicida to resist the host’s normal bactericidal mechanisms.

Significance The severity, host susceptibility, and wide geographic range make furunculosis a significant disease in salmonid aquaculture. In some geographical areas where culture of trout and other salmonids is intensive, furunculosis may be the “most” serious infectious disease to occur, particularly on farms in the United Kingdom and Scandinavia (Munro and Hastings 1993). Elevated stocking rates in culture units, leading to a general degradation of environmental conditions, also contributes to disease severity; therefore, good, sound fish health management is essential in coping with the disease. A. salmonicida’s significance is enhanced by the fact that it is capable of causing infections and some mortality in nonsalmonid fishes in both cultured and wild populations, thus producing possible carrier fish and pathogen reservoirs.

Vibriosis Since the mid-eighteenth century, it has been speculated that marine fish can be infected by members of the bacterial genus Vibrio (Colwell and Grimes 1984); Listonella is a synonym of Vibrio. It was noted by Bullock (2004a) that infections of Vibrio in fish were recorded as early as the 1500s. Up to 13 Vibrio species have been reported to cause fish infections and include Vibrio alginolyticus, V. anguillarum, V. carchariae, V. cholerae, V. damsella (Photobacterim damsella), V. harveyi, V. ordalii, V. parahaemolyticus, V. mimicus, V. vulnificus, and V. salmonicida (Colwell and Grimes 1984; Egidius et al. 1986; Saeed 1995). A new species, V. ichthyoenteri, was proposed by Ishimaru et al. (1996) and V. splendidus and V. pelagiius were iso-

lated from cod (Santos et al. 1997). However V. anguillarum, V. ordalii, and V. salmonicida (cold-water vibriosis) are best noted for their pathogenicity to fish. Others are sporadically associated with fish infections and will not be discussed at length; therefore, the following discussion includes diseases caused by V. anguillarum and V. ordalii, which are collectively referred to as “vibriosis.” Both organisms produce hemorrhagic septicemias, but the diseases they cause differ slightly. Cold-water vibriosis (Hitra disease), caused by V. salmonicida is discussed separately. Vibrio spp. infections have a variety of common names: vibriosis; red pest, saltwater furunculosis, boil disease, and ulcer disease.

Geographical range and species susceptibility Halophilic Vibrio spp. bacteria are a normal microbial flora in seawater and inhabit the gut of many marine organisms, including fish. These bacteria have caused infection in many different species of fish in brackish waters and seawater. Although vibriosis is considered a disease primarily of saltwater fish, it does occasionally occur in freshwater fish (Muroga 1975; Tajima et al. 1985) especially if the cultured fish have been given feeds containing raw marine fish products. V. anguillarum is found worldwide but is particularly significant along the Gulf of Mexico, Pacific, and Atlantic coasts of North America, the North Sea, Atlantic, and Mediterranean coasts of Europe and North Africa, the Middle East, and throughout Asia. V. ordalii occurs along the Northwest Pacific coast of North America and in Japan (Schiewe 1981; Muroga 1992). Other Vibrio spp. are not associated with any particular geographical region. Anderson and Conroy (1970) listed approximately 50 fish species, mostly marine, from which V. anguillarum has been isolated and numerous species have been added to the list.

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Colwell and Grimes (1984) listed 12 families and 42 species of marine and freshwater fishes affected by Vibrio spp.; V. anguillarum was the most frequently isolated and salmon the fish group most often affected. V. anguillarum occurs in wild salmon populations but most significant infections occur in culture populations, especially coho, chinook, and sockeye salmon. Essentially all salmonids are susceptible, particularly anadromous species cultured in saltwater cages or smolts released into saltwater after being reared in freshwater. Often the most severely affected nonsalmonid cultured species are maricultured Atlantic cod, striped bass, European eel, Japanese eel, milkfish grown in brackish water ponds, and ayu grown in fresh water ponds. Other commonly affected families include flounder, sole, turbot, mullets, seabass, and groupers. As mariculture of these fisheries expands worldwide outbreaks of V. anguillarum will most likely become more significant. It is unlikely that any group of marine or brackish water fish is resistant to V. anguillarum especially in mariculture. Ornamental marine fish in home and commercial aquaria are also vulnerable to the bacterium. V. ordalii is specific for anadromous salmon and freshwater trout (Schiewe 1981). V. carchariae is seldom involved in diseased fish but Soffientino et al. (1999) identified this species in diseased summer flounder under intensive culture in Rhode Island, USA.

Clinical signs Clinical signs of vibriosis are similar to those of motile Aeromonas septicemia as they both cause the same type of hemorrhagic syndrome (Figure 14.2). Most Vibrio infected salmonids, as well as other infected species, reduce their feeding and swim lethargically along the edges of ponds, raceways, or cages accompanied by intermittent erratic or spinning patterns (Novotny and Harrell 1975). Infected fish also display pale gills (anemic), skin discoloration,

and have hemorrhagic, ulcerative necrotic skin lesions that may expose underlying muscle. Lesions develop on the head, within the mouth, around the vent, or at the base of fins. The vent is often red, swollen, and exudes a bloody discharge. Abdominal distension and exophthalmia will occur in many fish. Hemorrhage in the eye is a common clinical sign in coho salmon and sometimes may be the only gross pathological sign. Internal gross pathology may or may not be striking; however, a generalized hyperemia can occur, body cavities often contain bloody ascitic fluid, intestines are inflamed and flaccid, and petechia are present in the liver, kidney, adipose tissue, and on the peritoneum. Evelyn (1971a) reported that death in Pacific salmon often occurred before large necrotic muscle lesions develop or massive changes occur within internal organs. However, Novotny and Harrell (1975) noted that the spleen can sometimes be 2–3 times larger than normal. Summer flounder with V. carchariae infection had reddening around the anus, distended abdomen filled with opaque fluid, enteritis, and necrosis of the posterior intestine. The bacterium was isolated from ascites fluid and kidney of moribund specimens (Soffientino et al. 1999).

Diagnosis Vibriosis is diagnosed by isolating the pathogen on general laboratory media (BHI or TSA, etc.) but isolation can be enhanced by addition of 0.5–3.5% NaCl to the media. Santos et al. (1996) found that of 45 V. anguillarum or related isolates from around the world, none would grow in media without salt. Media inoculated with V. anguillarum incubated at 25–30◦ C for 24–48 hours form round, raised, entire, shiny, cream-colored colonies measuring about 1–2 mm in diameter (Colwell and Grimes 1984; Austin and Austin 1987). Isolates from fish with systemic infection can

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(a)

(c)

(b)

(d)

Figure 14.2 Clinical vibriosis. (a) Pacific salmon with epidermal lesion on the back and caudal peduncle (arrows). (Photo courtesy of J. Rohovec.) (b) Red grouper with hemorrhagic lesion (arrow) on the skin. (c) Red drum with diffused hemorrhage (arrow). (Photo by J. Hawke.) (d) Milkfish with hemorrhage (arrow) in the scale pockets and epidermis.

be obtained from any internal organ, especially those with copious blood supplies. Isolation of V. ordalii is greatly enhanced by using seawater agar with up to 3% NaCl. Incubation at 15–25◦ C for 7 days will produce colonies that are not dissimilar from those of V. anguillarum (Schiewe et al. 1981). Although V. ordalii and V. anguillarum share numerous biophysical and biochemical characteristics, as well as some serological cross-reactivity (Chart and Trust 1988), there are distinct taxonomical differences between these two pathogens and V. salmonicida.

Bacterial characteristics V. anguillarum is a slightly curved, Gramnegative rod that measures about 0.5 × 1.5 µm. The organism is motile, cytochrome ox-

idase positive, ferments carbohydrates without gas production, and is sensitive to novobiocin and vibriostat 0/129 (2-4-diamino-6, 7disopropyl pteridine phosphate) (Table 14.1). Biochemically, V. anguillarum is a heterogeneous organism with at least five different described biotypes (Schiewe et al. 1977; Austin and Austin 1987). V. anguillarum is further fractionated by serological diversity. Initially, three salmonid serotypes were identified in the northwestern United States and Europe. Serological groupings number as many as six serotypes; therefore, serology can be used to separate and identify V. anguillarum. Using slide agglutination, Santos et al. (1996) serologically characterized 45 V. anguillarum and related isolates taken from fish, shellfish, and the environments of Denmark, Spain, Chile, Norway, and the United States. Some phenotypic and

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Table 14.1 Biochemical and biophysical characteristicsa of Vibrio (Listonella) anguillarum, V. ordalii, and V. salmonicida. Test/Characteristic

V. anguillarum

V. ordalii

V. salmonicida

Growth on 0% NaCl 3% NaCl Growth at 37◦ C Voges–Proskauer Arginine Christiensens citrate Starch hydrolysis Gelatin hydrolysis ONPG Lipase Indole from tryptone Glycogen Nitrate reduction Acid from Celobiose Fructose Galactose Glycerol Glucose Sorbitol Trehalose Maltose Sucrose Gentiobiose Gluconate D-Mannose Sensitive to 0/129

+a + + + + + + + + + + + + + + + + + + + + + − + + +

− + − − − − − + − − − − (+) − − − − + − − + + − − − +

− + − − − − − − − − − − − (+) − − (+) + − + − − − + + +

Source: Pacha and Kien (1969); Colwell and Grimes (1984); Holm et al. (1985); Dalsgaard et al. (1988). a +, positive; −, negative; (+), weakly positive.

serologic relatedness was found among the strains but 22 (49%) were typed among seven O-antigen serogroups. Isolates from turbot and cod were serologically different from the others; however, Knappskog et al. (1993) found that all 14 isolates from cod in Norway were V. anguillarum serotype 02. There are a number of serologic groupings of V. anguillarum; as many as 23 serovars (designated 01 to 023) have been identified. Most of the serovars pathogenic to fish are serovar 01, 02, or 03. Gonzalez et al. (2004) evaluated three serological methods, AQUARAPID-Va, AQUAEIA, and dot-blot assays, for V. anguillarum identification. In 21 strains tested, AQUARAPID-Va and AQUAEIA identified all 01 and 02 strains, while the dot-blot identified all 01, 02, and 03 strains. Because of simplic-

ity, effectiveness, and speed, AQUARAPIDVa was determined to be the most suitable serological method for detection of V. anguillarum in field and small-scale laboratory studies. Serogroup 03 constitutes V. ordalii. Clearly V. anguillarum is a diverse species that may help explain its wide geographical distribution and multiple fish species susceptibility.

Epizootiology V. anguillarum, and other Vibrio spp., are quite varied in their habitat requirements: living in ocean or brackish water, bottom mud, fish, and other marine organisms. Isolates from fish possess low to high virulence. It is not known if different strains are

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obligate pathogens, while others are facultative, but Larsen (1983) showed that environmental strains of V. anguillarum were less pathogenic than reference strains from fish. Vibriosis in saltwater is equivalent to motile Aeromonas septicemia in freshwater with the brackish water environment as a transitional zone where either may occur. V. anguillarum causes infection in wild fish but has its greatest impact on cultured fish, often during periods of environmentally induced stress. The disease is more prevalent during summer when water temperatures are high and dissolved oxygen is lowest. High population density or poor hygiene will contribute to epizootics; however, when fish are exposed to only a low number of highly virulent isolates, disease can occur without presence of exogenous stressors. V. ordalii is not as widespread as V. anguillarum and has been isolated only from fish and not from water or bottom sediments, but this pathogen causes severe losses among cagecultured salmon in coastal waters of North America’s Pacific Northwest. This organism also preferentially infects the skeletal muscle, heart, gills, and intestinal tract of salmonids and does not produce as extensive septicemia as V. anguillarum. V. anguillarum and V. ordalii are both capable of infecting and killing large numbers of cultured fishes. Thorburn (1987) noted that mortalities on various Swedish rainbow trout farms ranged from 0 to 17% and were correlated to transport of fish, farm size, and number of years the farm had been in production. Less severe effects occurred at older and larger farms, suggesting that experience in management may have a positive influence on disease outcome. Up to 100% mortality was reported in Atlantic salmon (Sawyer et al. 1979) after experimental exposure to V. anguillarum. Experimental waterborne challenges in Atlantic cod produced mortality of 4–36% using 13 different isolates from cultured cod fry (Knappskog et al. 1993). Some less common vibrios may show differential fish species pathogenicity; V. splendidus and

V. pelagiius, isolated from cod in Spain and Denmark respectively, were more pathogenic to rainbow trout than to cod (Santos et al. 1997). Increased water temperature has an effect on mortality of V. anguillarum-infected salmonids. For example, coho salmon suffered 58–60% mortality when held at 18–20◦ C, 40% at 15◦ C, 28% at 12◦ C, and only 4% at 6◦ C (Groberg et al. 1983). Salmon smolts held in freshwater have little, if any, exposure to V. anguillarum; therefore, when they reach saltwater, they are highly vulnerable to infection. In a vaccination experiment, Harrell (1978) found that over 90% of unvaccinated 0-age sockeye salmon were killed by V. anguillarum during their first 50 days in saltwater cages and mortalities of 60–90% are common among young salmon when stocked into saltwater. In Norway, mortality of V. anguillarum-infected Atlantic salmon transferred from freshwater to saltwater was influenced by time of transfer; fish moved late in the growing season suffered higher mortality than those that were moved early (Groberg et al. 1983). In most instances, V. anguillarum is transmitted through the water column and by contact, but Grisez et al. (1996) conclusively transmitted the disease via the oral route in turbot. Artemia nauplii were incubated in a suspension of V. anguillarum and then fed to turbot, which showed clinical signs of disease 24 hours after feeding. All fish that had been fed contaminated nauplii died within 4 days and the pathogen was detected in all exposed fish by immunohistochemistry. It was concluded that V. anguillarum was first transported through the intestinal epithelium by endocytosis, released into the lamina propria, and transported by the blood to internal organs, resulting in septicemia and death. Young fish were also infected with V. anguillarum at a very young age via feed infected rotifers to first feeding turbot (Planas et al. 2005). This method of infection provides reproducible results in challenge studies.

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Although salmonids have been the subject of the majority of vibriosis research as noted other fishes are susceptible to vibriosis. For example, wild and cultured flounder, eels, and milkfish are susceptible to the pathogen and can develop acute or chronic infections (Watkins et al. 1981). In flounder, acute vibriosis occurs year-round with onset of clinical signs appearing 12–48 hours postinoculation and death within 2–4 days. Clinical signs in acute vibriosis of these fish are less evident than in the chronic form. Chronic vibriosis appears 1–4 days postexposure with duration of 2–6 weeks during which death or recovery of some fish may occur. In Japan, juvenile ayu are susceptible to V. anguillarum, where the problem is more severe in sea-run fish in which the incidences can be as high as 17% compared to less than 1% after stocking into freshwater (Muroga et al. 1984). Also, diseased ayu from saltwater environments have higher numbers of bacteria in their tissues. A similar pattern is true for cultured eels in Japan where V. anguillarum is a greater problem in brackish water ponds than in freshwater ponds. These observations indicate that V. anguillarum is introduced into fresh water via infected fish that are captured in salt water and transferred to fresh water. Mizuki et al. (2006), studying the distribution of V. (Listonella) anguillarum in Japanese flounder, found that of 439 isolates the source was primarily rearing water and live diets (rotifers). They strongly suggested that the pathogen is a transient bacterium of the intestinal microflora or Japanese flounder and permanently indigenous to Japanese hatcheries.

Pathology Experimental oral V. anguillarum infections in turbot showed that 50% had bacterial cells in the spleen and 80% of these cells remained attached to the intestinal wall following washing (Olsson et al. 1996). These authors proposed that the intestine is the portal of entry for V.

anguillarum and that bacterium penetrates the intestinal mucus that overlays the epithelial cells and then enters the blood stream. Pathogenesis of V. anguillarum is facilitated by an ECP that is heat stable at 100◦ C for 15 minutes (Umbreit and Tripp 1975). Live and heat-killed bacteria and heat-treated cell-free culture fluid killed 53–80% of injected goldfish. Conversely, no evidence was found by Harbell et al. (1979) that an endotoxin, culture supernatant, or cell lysate caused pathology or death when injected into coho salmon. It was proposed that V. ordalii invades the rectum and posterior gastro-intestinal tract or possibly the integument (Ransom et al. 1984). Its pathology differs from that of V. anguillarum in that V. ordalii is not dispersed evenly in various organs and tissues but produces microcolonies in skeletal and heart muscle, gill tissue, and throughout the gastrointestinal tract. Also a lower number of V. ordalii cells are present in the blood during an infection than in V. anguillarum infections, possibly because the bacteremia develops later in the disease cycle. While not as well known as V. anguillarum or V. ordalii, V. harvei can be pathogenic to salmon. Zhang and Austin (2003) found that rainbow trout and Atlantic salmon injected with 106 cells of V. harvei per fish suffered up to 100% mortality. Of the 19 isolates studied, 14 were pathogenic; however, the ECP’s of only 5 isolates were harmful to fish.

Significance Vibriosis occurs worldwide and affects a wide variety of fish species, making it an important fish disease in marine and brackish water fishes. Vibriosis is considered the most common bacterial disease of marine species in general and is probably the most significant disease of cultured marine fishes, particularly salmonids in the United States (Novotny and Harrell 1975). Muroga (1975) also stated that vibriosis is the most important infectious

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disease of cultured ayu in Japan. Although most reports of vibriosis involve cultured fish, wild marine fish and ornamental fish are also susceptible.

Cold-water vibriosis Cold-water vibriosis (CV), also known as “Hitra disease” and hemorrhagic syndrome, is a disease primarily of Norwegian Atlantic salmon farming (Hjeltnes and Roberts 1993); however, its presence in other geographical regions is expanding. The disease was first described by Egidius et al. (1981) on the island of Hitra and since that time, it has been reported annually throughout Norway and elsewhere. The causative agent of CV was described by Holm et al. (1985) and Egidius et al. (1986), who proposed a new species, V. salmonicida. However, the etiology was disputed by Fjølstad and Heyeroaas (1985) and Poppe et al. (1985), who hypothesized that Hitra disease is a multifactorial disease that could include a nutritional deficiency (vitamin E), as well as environmental stress. Generally, V. salmonicida is accepted as the etiological agent of Hitra disease (cold-water vibriosis) (Bruno et al. 1986; Hjeltnes et al. 1987a), but environmental conditions could affect severity of the disease.

Geographical range and species susceptibility Cold-water vibriosis has been reported in cultured Atlantic salmon along the coast of Norway, in the Shetland Islands of northern Scotland, and in the Faroe Islands (Hjeltnes et al. 1987b; Dalsgaard et al. 1988). Coldwater vibriosis had not been reported from other parts of Europe but was detected by H. Mitchel (Connors Aquaculture, Inc. Eastport, Maine, personal communication) among Atlantic salmon in Nova Scotia, Canada, and the

northeastern United States in 1989 and again in 1993. The disease is found primarily in saltwater or brackish water net cages. Atlantic salmon are most susceptible to CV; rainbow trout can be infected but are far less susceptible (Egidius et al. 1984). V. salmonicida was also isolated from highly stressed wild juvenile cod, but it was thought that this incidence was unusual and cod should not be considered a likely host (Jørgensen et al. 1989).

Clinical sign Cold-water diseased fish may show no evidence of infection or develop distinct clinical signs. The initial clinical sign of cold-water vibriosis is the tendency for infected fish to swim lethargically on their sides near the surface of cages, show imbalance, and are inappetent (Holm et al. 1985; Bruno et al. 1986). Infected fish, which generally appear well nourished, may or may not exhibit extensive skin hemorrhage or become darker depending upon stage of disease. When hemorrhages do appear, they are located at the base of fins and on the abdominal region (Figure 14.3). Gills are generally pale, but the opercular cavity may be hemorrhaged. The anus may be reddish and prolapsed. Internally, a bloody fluid accumulates in the peritoneal cavity with hemorrhages on tissue surfaces ranging from petechia to ecchymosis (Figure 14.3). The spleen is grayish; the enlarged liver is usually yellowish, red, brown, or anemic; hemorrhages occur on the air bladder, which may also be filled with clear or bloody fluid. Hemorrhages also occur in fatty tissue around the pyloric cecae, throughout the visceral cavity, and on the peritoneal wall. The gut, particularly the posterior region, is hemorrhagic and the lumen contains a watery, yellowish or bloody fluid. Occasionally hemorrhages are found in muscle. Stained blood smears show bacteria in the plasma (Figure 14.4).

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(a)

(b)

(c)

(d)

Figure 14.3 Atlantic salmon infected with Vibrio salmonicida. (a) Hemorrhage (arrow) on the skin (Photo by H. Mitchell.) (b) Petechia (large arrow) on pale liver and peritoneum (small arrow). (c) Granular appearing spleen. (d) Blood smear of Atlantic salmon infected with cold water vibriosis and a high number of V. salmonicida in the blood. (Photos b, c, and d courtesy of B. Hjeltness.)

Diagnosis While V. salmonicida is not easily isolated on culture media, it can be recovered from internal organs and blood of infected fish. Initially, V. salmonicida was isolated on nutrient agar with 5% human blood and 1.5–2% NaCl (Egidius et al. 1984), but either TSA or BHI are suitable medias for isolation if they contain salt. The bacterium prefers 0.5–4% NaCl, grows optimally at 15–17◦ C, and will grow at 1–22◦ C but not at 26◦ C. At 15◦ C, slight growth is evident in 24 hours and becomes more pronounced at 72 hours. Colonies are

smooth, grayish, opaque, slightly raised, with entire margins, and range in size from punctate to 1 or 2 mm in diameter. Espelid et al. (1988) utilized a monoclonal antibody (MAB) against the surface antigen of V. salmonicida to identify the organism by ELISA. All isolates possessed a common outer membrane antigen designated as VS-Pl, which is a protein–lipopolysaccharide complex. Immunohistochemistry, based on the avidin–biotin complex, was used by Evensen et al. (1991) to identify V. salmonicida in formalin-fixed and histological-prepared tissues from experimentally infected Atlantic

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salmon. This method was also used to identify the bacterium in tissue from the original Hitra disease episode in 1977 in Norway in preserved heart, liver, spleen, kidney, and gut tissue by incubation with monoclonal antibodies specific for V. salmonicida. These data provide strong evidence for V. salmonicida as the cause of the first outbreaks of the disease and is consistent with other reports identifying it as the etiological agent of cold-water vibriosis.

Bacterial characteristics Vibrio salmonicida is Gram-negative, cytochrome oxidase positive, motile, and sensitive to vibriostat 0/129 (Table 14.1). The bacterium is a slightly curved rod, measures 0.5 × 2–3 µm, and may be pleomorphic upon initial isolation (Holm et al. 1985; Egidius et al. 1986). It is not hemolytic to human erythrocytes. According to Holm et al. (1985), V. salmonicida is a uniformly distinct species that differs serologically from V. anguillarum and V. ordalii. Isolates of V. salmonicida from Atlantic salmon and Atlantic cod were identical according to DNA analysis (Jørgensen et al. 1989) but Schroder et al. (1992) found serological diversity among V. salmonicida isolates from Atlantic salmon and Atlantic cod. Wiik et al. (1989) identified four plasmid profiles among 32 isolates, which would indicate possible species diversity. In spite of possible serological and plasmid differences, all strains of V. salmonicida are biochemically similar (Table 14.1).

Epizootiology Although there is some disagreement about the etiology of Hitra disease, it is synonymous with cold-water vibriosis. Fjølstad and Heyeroaas (1985) found pathology of Hitra disease to be similar to vitamin E deficiency in higher vertebrates and because a pathogenic organism could not consistently be isolated, they proposed a nutritional etiology. Observa-

tions by Poppe et al. (1985) cannot be ignored, but most scientists working with V. salmonicida concur that cold-water vibriosis and Hitra disease are synonymous. Hjeltnes et al. (1987a) and Hjeltnes and Julshamn (1992) clearly correlated Hitra disease and cold-water vibriosis to V. salmonicida; however, they recognize that environmental conditions and nutrition may also play an important role in the disease. Hitra disease appeared in 1977 when it caused significant losses on fish farms on the Norwegian Island of Hitra. After that it was devastating to salmon farms along western and northern Norway (Egidius et al. 1981). This episode was followed by outbreaks in other regions (Bruno et al. 1986, Dalsgaard et al. 1988). On the basis of plasmid profiles, it was suggested that V. salmonicida was transmitted from cod to Atlantic salmon farms in northern Norway and then back to cod (Sørum et al. 1990). Cold-water vibriosis is transmitted to noninfected Atlantic salmon and to a lesser degree to rainbow trout through water; the pathogen is excreted from infected fish in excrement and is believed to survive in mud and can be carried long distances in lipid or air bubbles (Anonymous 2004). Hjeltnes et al. (1987a) fulfilled Koch’s postulates for V. salmonicida via transmission through water from infected fish. Superficial injury enhances infectivity but primarily the pathogen enters fish through the gills. Atlantic salmon infected via gills show behavioral and clinical signs after 1 week and 2 weeks postexposure. Mortality in V. salmonicida infected yearlings to market-size Atlantic salmon can be high. In natural infections, acute mortality rates of 5% per day have been noted. Injured fish exposed to waterborne bacteria suffered from 80 to near l00% mortality in about 20 days. Fish challenged more naturally by cohabitation suffered 24 and 46% mortality in 24 and 35 days, respectively. Experimental waterborne exposure of the more susceptible Atlantic salmon resulted in 90% mortality over a 45-day period compared to 40%

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ilar to naturally infected fish (Bruno et al. 1986). Significant pathology occurred in the pyloric cecae and the mid- and hindgut and necrotic tissue sloughs into the lumen. Blood vessels are dilated and congested; hemorrhage is present in the lamina propria; and the entire mucosal epithelium is necrotic in some cases. Focal necrosis occurs in hematopoietic tissue of the kidney where glomerulus cell nuclei are often swollen and occasionally necrotic. The ellipsoid system of the spleen is engorged with macrophage like cells, focal necrosis develops in the reticuloendothelial cells, and gills exhibit epithelial necrosis and sloughing; however, the liver and pancreas appear normal. Totland et al. (1988) found that in V. salmonicida infections, the blood was heavily laden with bacteria. They also detected bacteria in the cell membrane on the luminal side of endothelial cells of capillaries. Intracellular bacteria were later seen within endothelial cells. Histopathology substantiates the causative relationship of V. salmonicida in cold-water vibriosis, but pathogenesis of the disease is not fully understood because knowledge of factors responsible for pathogenicity is limited.

mortality in similarly infected cod (Schroder et al. 1992). Rainbow trout, infected by injection with an undetermined number of bacteria, developed initial signs of disease 9 days postinfection and suffered 80–100% mortality after 14 days when held at 9◦ C (Egidius et al. 1981). These results may be a little misleading because of the high number of bacteria injected; however, it illustrates the diseases potential for severity at this temperature. Generally, cod are not susceptible to V. salmonicida but in an unusual highly stressed situation, naturally infected juvenile cod on 15 farms suffered 10–90% (50% average) mortality from the pathogen (Jørgensen et al. 1989). These fish were stocked into net cages in close proximity to V. salmonicida-infected Atlantic salmon. The majority of V. salmonicida infections occur in autumn to late winter or early spring when water temperatures ranged between 4 and 9◦ C (Egidius et al. 1981); however, H. Mitchell (personal communication) noted 0.5% mortality per day in some cages in Maine when water temperatures were 1◦ C. Between 1977 and 1981, the maximum number of cold-water vibriosis cases diagnosed in one year at the National Veterinary Institute, Oslo, Norway, constituted only a small percentage of the total case load. In 1982, the number of cold-water vibriosis cases increased to over 70% of total cases and during ensuing years the disease caused increasing losses. More cases have been reported during the warmer months (B. Hjeltnes, Institute of Marine Research, Bergen, Norway, personal communication), a pattern that has also been noted in North American epizootics. Disease severity seems to increase during May and June when water temperatures rise to 5–6◦ C, but when it appears during winter, it is most difficult to control.

During the 1980s, 80% of lost revenue from fish diseases on Norwegian fish farms was attributed to Hitra disease (Poppe et al. 1985). However, since 1989, mortalities due to the disease have decreased (B. Hjeltnes, personal communication). The fact that the disease is no longer confined to Norway and not only affects Atlantic salmon, but to a lesser degree rainbow trout and cod, makes it a disease of concern where they are cultured.

Pathology

Enteric redmouth

Histopathology of cold-water vibriosis in experimentally infected Atlantic salmon is sim-

Enteric redmouth (ERM) is a systemic bacterial disease principally of trout caused by

Significance

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Yersinia ruckeri, a member of the family Enterobacteriaceae. It is a chronic to subacute infection that primarily affects farm reared trout. The disease is also known as Hagerman redmouth disease, redmouth, salmonid blood spot, and in Norway it is called “yersiniosis.”

Geographical range and species susceptibility Enteric redmouth, initially diagnosed in the early 1950s in cultured rainbow trout in Idaho, now occurs in most states where trout are grown. Bacterial isolates obtained from rainbow and brook trout in West Virginia in 1952 and from rainbow trout in Australia in 1963 appeared to be identical to Y. ruckeri (Bullock et al. 1978a); therefore, it is probable that detection of Y. ruckeri in various areas during the past may not be totally due to dissemination from Idaho but likely the organism is endemic to at least some of these areas. Essentially, enteric redmouth occurs worldwide where salmonids are cultured (Tobback et al., 2007). The disease has been confirmed in many countries including Australia, Bulgaria, Canada, Denmark, Finland, France, Germany, Great Britain (including Scotland), Italy, Norway, South Africa, and Switzerland (Austin and Austin 1987; Davies and Frerichs 1989; Stevenson et al. 1993). However, to date, it has not been reported in Japan’s extensive trout and salmon farms. Rainbow trout are most severely affected by ERM; however, all salmonids are susceptible to some degree and chinook salmon is the most severely affected anadromous salmon species in the Pacific Northwest (K. Amos, personal communication). Significant mortalities of Atlantic salmon have been reported on Norwegian farms and eight salmonid and seven nonsalmonid species were identified by Stevenson et al. (1993) in which Y. ruckeri caused disease. Nonsalmonid species include the emer-

ald shiner, fathead minnow, three species of whitefish, sturgeon, and turbot, but for the most part, mortalities among these species have been insignificant. One report describes Yersinia ruckeri being isolated from diseased, cultured channel catfish in Arkansas (Danley et al. 1999). The bacterium has also been isolated from numerous apparently healthy fish, invertebrates (crawfish), mammals (muskrat), sea gulls, a human, sewage, and river water (Busch 1983; Michel et al. 1986a; Stevenson et al. 1993). It is quite possible that Y. ruckeri is more common in the environment than was once thought.

Clinical signs Enteric redmouth is categorized into subacute, chronic, and latent phases, and clinical signs vary within each phase. In the subacute phase, fish often exhibit a red mouth, head, and jaw as a result of subcutaneous hemorrhaging (Figure 4.4) (Ross et al. 1966; Busch 1983). Affected fish are dark, have inflammation and/or hemorrhaging at the base of fins and around the vent, and exhibit unilateral or bilateral exophthalmia with orbital hemorrhage; gills are hemorrhaged near filaments tips. Fish are inappetent and sluggish with a tendency to accumulate downstream in raceways or along pond edges. However, Frerichs et al. (1985) described Y. ruckeri infections in Atlantic salmon in which there was no reddening of the mouth and opercula; therefore, Y. ruckeri should not be ruled out if the classic “redmouth” is absent. Fuhrmann et al. (1984) reported that Y. ruckeri often produces gross internal pathology that resembles “furunculosis”. Petechia may be present in the muscles, visceral fat, intestines, liver surface, pancreas, pyloric cecae, and swim bladder. The lower intestine is flaccid, hemorrhaged, inflamed, and contains a thick, opaque, purulent material. The kidney and spleen may be enlarged.

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(a)

(b)

Figure 14.4 Rainbow trout infected with Yersinia ruckeri. (a) Petechiae on the abdominal wall (small arrow), ecchymosis in posterior abdominal musculature (large arrow), hemorrhage in mandible and exophthalmia. (b) This fish has pale gills, mottled liver and petechiae in the pyloric cecae area and in visceral adipose tissue (arrows). (Photos courtesy of S. LaPatra, Clear Springs Food.)

As disease progresses into the chronic phase, exophthalmia increases (bilaterally or unilaterally), often causing the eye to rupture. Fins become frayed and eroded, gills are pale, the abdomen is distended, and there is an accumulation of bloody, ascitic fluid in the body cavity (Busch and Lingg 1975). Fish become dark, lethargic, and emaciated before death. As clinical signs disappear, mortality abates and survivors move into a latent phase in which a few darkly pigmented fish may be present.

Diagnosis Enteric redmouth is diagnosed by isolation of Y. ruckeri from internal organs on general purpose bacteriological media (BHI or TSA) (Ross et al. 1966). Rodgers (1992) described a selective Y. ruckeri media (ROD), which differentiates the organism by its yellow color. Waltman and Shotts (1984) also described a selective media (Shotts-Waltman, SW) for Y. ruckeri on which it forms a green colony surrounded by a zone of hydrolysis. Hastings

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and Bruno (1985) evaluated the SW selective media and concluded that it alone was not reliable in isolating and identifying Y. ruckeri because of insufficient specificity; however, the selective media can be helpful in identification of Y. ruckeri from carrier fish, particularly if inocula are taken from intestine or kidney. Amplification of Y. ruckeri DNA in PCR was used by Argenton et al. (1996) to diagnose enteric redmouth. The PCR assays detected Y. ruckeri in three cell preparations of seeded samples: serotype O1 and O2 and an unknown. The bacterium was also detected by PCR in infected trout kidney tissue after brief digestion with proteinase K. Because of speed, sensitivity, and specificity, it was concluded that PCR was preferable to conventional bacteriological diagnostic tests. Altinok et al. (2001) evaluated a nonlethal method of detecting Y. ruckeri in blood of experimentally infected rainbow trout. Following a 1 hour immersion in water with 4.5 × 106 CFU Y. ruckeri, PCR was positive for the pathogen and continued to be positive for 5 days; the fish were negative from 9 to 30

days postexposure during which time no signs of clinical disease were noted. The method applied to 42 naturally infected rainbow trout, from which Y. ruckeri had been isolated, was also positive for the pathogen in all fish. The PCR detected the pathogen from four fish showing signs of ERM and five additional fish from which no bacteria were isolated. Three of five fish collected from a stream below the ERM positive hatchery also yielded positive results in response to the PCR technique.

Bacterial characteristics Yersinia ruckeri grows aerobically and anaerobically at 9–37◦ C with an optimum temperature range of 22–25◦ C; colonies form in 24–48 hours. Isolates incubated at 37◦ C are generally avirulent to trout. The white to cream colored, translucent colonies are 1–2 mm in diameter, smooth, slightly convex, raised, and round with entire edges. Once the organism is isolated, it can be identified by conventional biochemical characteristics (Table 14.2)

Table 14.2 Biochemical and biophysical characteristics of Yersinia ruckeri. Test/Characteristic

% Positive

Test/Characteristic

% Positive

Motility Fermentation Oxidase Catalase ß-galactosidase Orinthine decarbosylase Lysine Arginine dihydrolase Tryptophan deaminase Urease H2 S production Indole Citrate utilization Methyl red Voges-Proskauer Gelatin hydrolysis Casein hydrolysis Tween 20, 80 hydrolysis Growth on MacConkey agar

82 100 0 100 100 100 100 0 0 0 0 0 99 91 93 77 74 82 99

Nitrite reduction Gas from glucose Acid from Amygdalin Arabinose Fructose Galactose Glucose Inositol Lactose Maltose Mannitol Mannose Melibiose Sorbitol Sucrose Trehalose

99 8 0 0 100 99 100 0 0 99 99 100 0 20 0 100

Source: Davies and Frerichs (1989).

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(Ewing et al. 1978; Stevenson and Daly 1982). It is a motile (peritrichous flagellation), Gramnegative, cytochrome oxidase negative rod that measures 1.0 × 1.0–3 µm. The most important biochemical reactions of Y. ruckeri, which are fairly homogeneous, are positive for fermentative metabolism, production of catalase and β-galactosidase, lysine and ornithine decarboxylation, methyl red test, nitrite reduction, and degradation of gelatin. It will grow on media containing up to 3% NaCl and utilizes citrate; two characteristics that separate it from Edwardsiella ictaluri. Yersinia ruckeri’s inability to produce H2 S, indole, oxidase and phenylanine deaminase, and its negative reaction for Voges-Proskauer are significant; however, some strains may vary in methyl red, Voges-Proskauer, lysine decarboxylase, arginine dihydrolase, and lactose fermentation tests (Ross et al. 1966; Busch 1983). Fermentation of sorbitol has received some attention in discriminating between pathogenic strains (Serotype I) and nonpathogenic strains (Serotype II). Serotype I does not ferment sorbitol as does Serotype II, consequently Cipriano and Pyle (1985) developed a sorbitol-based media used to distinguish them. However, Valtonen et al. (1992) questioned the accuracy of the sorbitol reaction test as a pathogenic indicator in Norwegian isolates. Austin and Austin (1987) stated that Y. ruckeri could be confused with Hafnia alvei if the API 20 system is used for identification; however, Grandis et al. (1988) showed that H. alvei is L-arabinose and L-rhamnose positive, while Y. ruckeri is uniformly negative for these reactions. Arias et al. (2007) described an isolate of atypical Y. ruckeri from brown trout in South Carolina, which was classified as biotype 2. The isolates were biochemically and genetically distinct from reference cultures including the type strain but otherwise were identical to the species based on API 20E, fatty acid methyl ester profiles, and 16S rRNA sequence analysis. Because the isolates were nonmotile and unable to hy-

drolyze Tween 20/80, they were classified as Y. ruckeri. Yersinia ruckeri can be positively identified by IFAT (Davies and Frerichs 1989), but one must be aware of potential cross-reactivity with other enteric bacteria such as H. alvei. Serological cross-reaction does not occur with other pathogens of fish (A. salmonicida, A. hydrophila, V. anguillarum or R. salmoninarum) that may be encountered in salmonids (Hansen and Lingg 1976). Based on agglutination, Y. ruckeri was originally separated into two serotypes; Serotype I, the Hagerman strain, which is pathogenic and most commonly encountered in clinical diagnosis of diseased fish (Ross et al. 1966; Busch 1983), and Serotype II, the Oregon strain, which is normally not highly pathogenic (Bullock et al. 1978b). However, Cipriano et al. (1986) found that Serotype II was pathogenic to chinook salmon. Currently, six whole cell serotypes of Y. ruckeri are recognized (Table 14.3). Stevenson and Daly (1982) and Stevenson and Airdrie (1984) divided the species into five serological groups (I and I’ through IV), concluding that the Australian isolate (Bullock et al. 1977) should be included in Serotype I (I’) and two additional Serotypes (V and VI) should be established. Daly et al. (1986) detected five strains of Y. ruckeri in Canada, most of which were in Serotype I and II.

Table 14.3 Serological groupings of Yersinia ruckeri Serological Group (Serovar)

I I II III IV V VI

Designation

Hagerman Salmonid Blood spot Oregon A Oregon B Australian (Excluded) Colorado Ontario

Source: Stevenson et al. (1993).

% of Isolates

59 6 15 8 6 0 3 3

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Sousa et al. (2001) established the molecular characterization of Portuguese strains of Y. ruckeri from cultured fish and by lipopolysaccharide (LPS) and outer membrane protein (OMP) analysis, plasmid profiles, and ribotyping to determine the heterogeneity among the isolates. Only two LPS profiles were observed among the isolates that corresponded to serotypes O1 and O3 of Y. ruckeri. The OMP analysis showed more heterogeneity with seven different patterns and a greater diversity in ribotyping was noted. Although OMP analysis and ribotyping can be useful for identifying strains of V. ordalii on the basis for farm and/or season of isolation, the authors considered ribotyping to be the best approach for epidemiological studies because it is easier to perform and offers slightly better discriminative power.

Epizootiology Although Y. ruckeri is generally considered an obligate pathogen, ERM epizootics are influenced by the fish’s environment and outbreaks are often allied with stress (Busch 1983), a fact that cannot be over emphasized. Hunter et al. (1980) noted that asymptomatic Y. ruckeri carrier steelhead trout did not transmit disease to uninfected fish unless stressed. However, when stressed at 25◦ C the pathogen was transmitted from carrier to uninfected fish, but no deaths were reported in the recipients. Disease outbreaks can be precipitated by handling apparently healthy trout and/or by presence of increased ammonia and metabolic waste in the water, which decreases oxygen levels (Bullock and Snieszko 1979). In a study to evaluate the effect of oxygen concentrations on Y. ruckeri-exposed rainbow trout, Caldwell and Hinshaw (1995) showed that a supersaturation of oxygen (150% of saturation) resulted in significantly higher mortality (17.9%) when fish were challenged with the pathogen as opposed to a mortality of 12.6% at hypoxic (70% of saturation) and 10% at normoxic (100% of

saturation) levels of dissolved oxygen. In view of these data, the 100% saturation is preferable to maintain health. Water salinity affects the rate of ERM in rainbow trout (Altinok and Grizzle 2001). Juvenile rainbow trout and brown trout exposed to Y. ruckeri were held in freshwater and salinities of up to 9.0
. Rainbow trout in freshwater suffered 96.5% mortality, whereas fish in the highest salinity suffered 75.0% mortality; however, there was no significance in the mortality between the treatments. These compared to 2.3% mortality in brown trout in all salinities. The pathogen was isolated from dead fish of both species. Infected trout and salmon can become Y. ruckeri reservoirs. Busch and Lingg (1975) reported that of rainbow trout tested 45 days after surviving an ERM epizootic 25% had Y. ruckeri established in their lower intestines, thus making them asymptomatic carriers. When held at 14.5◦ C, these fish developed a recurrent ERM infection. Yersinia ruckeri carrier trout shed the organism in higher numbers on 36–40 day cycles and bacterial shedding preceded clinical signs and mortality by 3–5 days. Also cyclic bacterial shedding and ensuing infections were influenced by water temperature and other water quality parameters; however, the full implication of this phenomenon is still not completely understood. Yersinia ruckeri was found in water and bottom sediments throughout the year at two trout hatcheries. High-risk periods for ERM outbreaks at these hatcheries were related to stressful conditions and peak bacterial counts in the water (Romalde et al. 1994). These researchers reported that the pathogen was isolated from water of an effluent stream 15 km below the outfall of contaminated hatcheries. Incubation time between exposure to Y. ruckeri and clinical ERM will vary inversely with water temperature, general health of fish, and presence of additional stressors (Ross et al. 1966; Busch 1983). In fish at 15◦ C, 5–7 days usually pass between exposure and first deaths, but if fish have a history of prior exposure,

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mortality may occur in 3–5 days. Onset of mortality may also depend upon fish size and level of exposure. Enteric redmouth outbreaks usually occur only after fish have been exposed to large numbers of the pathogen and deaths can continue for 30–60 days after initial onset of disease. Enteric redmouth occurs primarily between April and September when fish range in size from 5 to 200 g (Rodgers 1992). Larger fish can also be affected, but incidence of infection coincides with rise and fall of water temperature, poor water quality, over-crowding, and handling. Exposure to sublethal concentrations of copper (7 and 10 µg/L for 96 hour) significantly increases susceptibility of steelhead trout to ERM (Knittel 1981). Low numbers of Y. ruckeri could be cultured on selective media from intestines of carrier rainbow trout 6–8 weeks before infection appeared in the kidneys (Rodgers 1992). On a farm where fish were severely stressed by frequent handling and poor water exchange, a significantly higher number of bacteria were found in the feces and kidneys of fish. Horizontal transmission occurs primarily through water from fish shedding bacteria to uninfected fish. Transmission of Y. ruckeri during spawning has not yet been clearly shown; however, the organism can be isolated from broodstock at this time. The effect of Y. ruckeri on trout depends upon age and size of fish, their relative susceptibility, environmental conditions, and stress level. Naturally infected hatchery salmonids generally suffer less than 10% mortality during the subacute phase of disease and recurrent infections in survivor populations result in low, chronic mortality over an extended period. In laboratory infectivity studies, mortality depends upon method of challenge; IP injection produces more consistent results, while waterborne exposures are inconsistent producing mortality from near 0 to 100% (Busch and Lingg 1975). In Europe, Y. ruckeri’s presence in imported bait fish strongly suggests that they

were the original disease source on that continent (Michel et al. 1986a). Sources other than fish may contribute to the spread of Y. ruckeri. Willumsen (1989) found the pathogen in the intestinal contents of sea gulls during an ERM outbreak and Stevenson and Daly (1982) reported the pathogen in muskrat intestine. Yersinia ruckeri was also isolated from crayfish in a spring water supply. When the crayfish were eliminated, no more Y. ruckeri infections occurred, indicating the reality of a nonfish source (K. Amos, personal communication). Yersinia ruckeri may also survive in water and mud for an extended period. McDaniel (1972) reported it could adapt to a normal aquatic saprophytic state and live for 2 months in mud.

Pathology Histopathology of ERM is characterized by bacterial colonization of capillaries of heavily vascularized tissues. Submucosal hemorrhage of the mouth, jaws, and under the head is most striking but is not always present. Gills develop telangiectasis and petechial hemorrhage, and congestion; edema occurs in the muscle, liver, kidney, spleen, and heart (Busch 1983). Inflammation occurs in the reticuloendothelial tissue and necrotic foci develop in the liver with an accumulation of mononuclear cells. Macrophages phagocytize bacterial cells throughout the vascular system and hematopoietic tissue of the kidney. The spleen is necrotic, resulting in a loss of lymphoid tissue. The digestive tract is characteristically hemorrhaged, edematous, and necrotic with sloughing of the mucosa into the lumen. Because of septicemia and hemorrhaging, hematocrits, hemoglobin content, and blood protein are reduced to about half of normal (Quentel and Aldrin 1986).

Significance Enteric redmouth is a significant disease in cold-water aquaculture, particularly in

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rainbow trout. Its ability to cause high mortality in production-size trout emphasizes its impact on aquaculture. In the past ERM has accounted for up to 35% of overall losses in the trout industry at an annual cost of over $2.5 million in the Hagerman Valley (Busch 1978). In trout, detrimental effects of chronic Y. ruckeri infections on reduced feeding, higher feed conversions, and retarded growth rates is unknown but could be economically significant. Also the fact that its geographic range has increased, either by dissemination or because of better surveillance and detection methods, has elevated the disease to one of international concern. However, the common practice of vaccinating trout for ERM has significantly reduced the disease to a minor problem in many areas.

Bacterial gill disease Bacterial gill disease (BGD) was named by Davis (1926) when he observed large numbers of long filamentous bacteria on clubbed gills of brook and rainbow trout. The term has since been used to describe infections caused by several different species of bacteria that affect fish gills, but the principle etiology of classical BGD in salmonids is Flavobacterium branchiophilum (branchiophila) (Wakabayashi et al. 1989; Bernardet et al. 1996). Also, Strohl and Tait (1978) described Cytophaga aquatile as a causative organism of BGD in trout and salmon, and Bernardet et al. (1996) determined that this species of bacteria should be Flavobacterium aquatile. Apparently both organisms, F. branchiophilum and F. aquatile, can actually cause the same clinical disease but F. branchiophilum infections appear to be more prevalent. In addition, an “atypical” bacterial gill disease (ABGD), caused by a bacterium identified as F. branchiophilum, was described in Canada but the disease did not produce clinical signs typical of BGD (Ostland et al. 1999). Other bacteria can occasionally be involved with BGD,

especially in nonsalmonids but are generally not the primary pathogen. BGD is distinctly different from gill infections of F. columnare (columnaris) (Chapter 11). While recognizing that pathological gill changes are caused by many factors, including nutrition, toxicants, fungi, and bacteria, only those gill infections of juvenile salmonids caused by filamentous flavobacteria (F. branchiophilum) and the ABGD organism will be discussed.

Geographical range and species susceptibility Since first being described in northeastern United States (Davis 1926), BGD has been reported throughout North America (Wakabayashi et al. 1980; Ferguson et al. 1991), Japan (Kimura et al. 1978), Hungary, the Netherlands, and it likely occurs in most countries where trout are cultured. Essentially this pathogen is found worldwide. F. aquatile, described in Canada, is similar biochemically to isolates taken from fish gills in numerous geographical locations, suggesting that it also has a wide geographical range (Strohl and Tait 1978). Juveniles of many fish species are susceptible to BGD but cultured salmonids, principally rainbow and brook trout, are most severely affected. BGD has also been reported in carp, goldfish, catfish, eels, fathead minnows, and other fish species (Ostland et al. 1989). Whether or not the same bacterial species is responsible for BGD in all fish species is unclear. The ABGD described by Ostland (1999) was found in multiple episodes of five salmonid species, including Atlantic salmon, in Canada.

Clinical signs Fish affected by BGD experience inappetence, orient themselves up stream or move toward water inlets, and ride high in the water. They

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(a)

(b)

(c)

Figure 14.5 Bacterial gill disease of salmonids. (a) Juvenile rainbow trout showing flared gill cover with some debris protruding from under one cover (arrows). (Photo courtesy of J. Ferguson.) (b) Clear lamellar trough (arrow) of healthy gill. (c) Mucous in the lamellar trough with long filamentous Flavobacterium branchiophilum (arrow).

become lethargic, sometimes spiraling, and moribund individuals sink to the bottom (Bullock 2004b). Affected fish have dark pigmentation, gills become very pale, gill covers are flared, or fish show opercula flashing; gills become swollen and congested with mucus and debris, causing the gills to protrude from beneath the operculum (Figure 14.5). Fish with ABGD did not present typical BGD clinical signs in that they did not show marked respiratory distress with flashing opercula and did not orient toward water inflow. Mortality rates of BGD rises rapidly, while that of ABGD was not precipitous.

Diagnosis Clinical BGD is presumptively diagnosed by microscopic examination of gill tissue wet mounts or by Gram staining smears. Large numbers of Gram-negative filamentous bacteria are observed on the surface of gill intralamellar spaces where they form clumps of long slender rods. Gill filaments begin swelling at the distal end where hyperplasia of the epithelium produces a “clubbed gill” condition

and becomes so acute that filaments fuse. Huh and Wakabayashi (1989) compared light microscopy of wet mount material to IFAT for accuracy in detecting BGD; IFAT was more accurate and resulted in identification of greater numbers of infected trout. Unless clinical disease is in progress, neither method detects F. branchiophilum on fish gills or in water. The organism is usually confined to the gill epithelium, but some bacteria may be found within and beneath necrotic lamellar epithelial cells (Ostland et al. 1994). Gills with ABGD are stubby with less-clubbed lamellae. Flavobacterium branchiophilum and F. aquatile are difficult to culture but may be cultured aerobically on cytophaga agar con◦ taining 1.5% agar incubated at 10–18 C. Both pathogens form yellow pigmented colonies, thus leading to the designation of “yellow pigmented bacteria” (YPB). Suspect F. branchiophilum colonies contain long filamentous bacteria and have refractive circular “cysts” (Ostland et al. 1994). Incubation at 12–18◦ C does not affect quantitative isolation whether incubation is 6 or 12 days. These bacteria are not isolated from internal organs. Colonies of cultured F. branchiophilum

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are light yellow, round, translucent, mildly convex, smooth, and about 0.5–1 mm in diameter after being incubated at 18◦ C for 5 days (Wakabayashi et al. 1989). Following bath exposure of rainbow trout to F. branchiophilum, an ELISA system detects the bacterium in crude gill extracts (MacPhee et al. 1995a). Using F. branchiophilum polyclonal antiserum, the biotin–avidin system detected the pathogen in whole bacterial cell populations, in gill preparations seeded with bacteria or in extracts of infected gills. Although the authors propose that the ELISA method is adaptable to field-collected gill samples, they recognized that further antigenic specificity testing is required. Spear et al. (1995) developed a MAB to F. branchiophilum and Flavobacterium columnare and used the MAB as the basis for indirect fluorescent antibody (IFA) to identify archived cultures from eastern Canada. The MABs, in conjunction with IFA, identified 76.2% of tested cultures as F. branchiophilum and 18.7% as F. columnare.

Bacterial characteristics Morphologically, F. branchiophilum is a Gram-negative rod, which measures approximately 0.5 × 5–15 µm (average length is 8 µm), is nonmotile, does not glide or spread on agar, and is cytochrome oxidase positive. Cells are heavily fimbrianated (Ostland et al. 1994). Biochemical and biophysical characteristics are similar among all F. branchiophilum isolates (Table 14.4); however, there is variability in carbohydrate utilization. The organism does not grow on TSA agar unless the medium is diluted approximately 20-fold, grows slowly at 10, 18, and 25◦ C on Cytophaga media containing up to 0.1% NaCl, but some strains grow at 5 and 30◦ C but not at 37◦ C. The ABGD agent is a Gram-negative, motile rod that produces a fluorescent pigment similar to Pseudomonas fluorescence; however, based on clinical pre-

sentation and histopathology, the bacteria associated with the disease was determined by the investigators to be F. branchiophilum (Ostland et al. 1999). The ABGD agent was compatible with F. branchiophilum as determined by polyclonal antibody. Antigenically, isolates from brook and rainbow trout, and Atlantic salmon in Ontario, Canada, are similar to those isolated from salmonids in Japan and Oregon (reference strains ATCC 3505 and ATCC 3506, respectively) (Ostland et al. 1994). All strains have similar mol% G + C of DNA of 30–32. Flavobacterium aquatile is isolated on 2% tryptone agar plates (Strohl and Tait 1978). Within 7 days, spreading colonies with pigmentation that varies from yellow to reddishorange appeared (Table 14.4).

Epizootiology The epizootiology of BGD is confusing because it has been associated with several different bacteria; Flexibacter sp., Cytophaga sp. (both of which are now flavobacteria), and Flavobacterium spp. Bullock (1972) implicated Flavobacterium sp. (Cytophaga sp.) in BGD but did not identify the species and could not create infections at will without severe environmental stress. Strohl and Tait (1978) described F. (Cytophaga) aquatile isolated from diseased gills of salmon and trout in Michigan, and Wakabayashi et al. (1980) showed that organisms causing BGD in Japan, Oregon, and Hungary were all members of the genus Flavobacterium. Bacterial strains from these regions possess some identical antigens as well as slight differences; however, strains from the USA and Hungary are most similar (Huh and Wakabayashi 1989). In general, experimental BGD transmission is unpredictable. F. branchiophilum has been successfully transmitted in the laboratory, but often only in conjunction with environmental stress. When rainbow trout are infected with

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Table 14.4 Characteristics of Flavobacterium branchiophilum and Flavobacterium aquatile, the two filamentous bacteria associated with bacterial gill disease of trout. Test/Characteristic

F. branchiophilum

F. aquatile

Yellow-red colony Growth at 37◦ C Motility Growth on nutrient agar Hydrolysis of Gelatin Casein Chitin Esculin Starch Indole from tryptone Nitrate reduction Bind Congo red Catalase production H2 S production ONPG Growth in 0% NaCl 0.5% NaCl 1.0% NaCl 2.0% NaCl Acid from Glucose Fructose Lactose Sucrose Maltose Trehalose Cellobiose Arabinose Xylose Raffinose Mannitol

+ − − − + + − − + − − ? ? ? ? + − − − + + − + + + (+) − − (+) −

+ − Gliding + + + + + + + + − + (+) + + + + + + + − − − (+) + + + + +

Source: Strohl and Tait (1978); Wakabayashi et al. (1989). +, positive; −, negative; (+), weakly positive.

F. branchiophilum by bath challenge, infection develops in 24 hours and mortality in 48 hours (Ferguson et al. 1991). Lumsden et al. (1994) successfully transmitted BGD by immersion, which resulted in a 23–41% survival rate in infected fish compared to 49–70% in unexposed controls. Stressful conditions were not used in either of these studies. F. aquatile has not been experimentally transmitted under any condition. Factors associated with BGD outbreaks in Ontario, Canada, indicated that if a diagnosis is made early in the epizootics that 84–100% survival could be achieved with the lowest survival in brook trout (Good et al. 2008). Survival within hatcheries var-

ied between years and that the most serious problems occurred in spring (March-May). The results emphasized the importance of the species being reared and time of year. In an earlier study, Bebak et al. (1997) evaluated the presence of BGD on numerous commercial hatcheries. They found that (1) regardless of location of the outbreak, there was significant association between mortality from BGD and previous experience with outbreaks; (2) correlation with commercial trout hatcheries with >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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(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 <10◦ C only (Ostland et al. 1999. The reservoir of F. branchiophilum is not fully known. It is theorized that subclinically

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infected fish are a source of infection when they shed bacteria; however, trout reared entirely in well water have become infected. Heo et al. (1990) detected F. branchiophilum in the water immediately before and during an epizootic, but not after disease had abated. Ostland et al. (1990) enumerated total bacteria and percentage of YPB on the gills of clinically healthy and BGD-infected rainbow trout. Healthy gills had 4.9 × 105 bacterial CFU/g of tissue, 15% of which were YPB. Severely BGD-infected fish had 3.9 × 106 CFU/g, of which 35% were filamentous YPB. Overall, these data suggest a strong association between the presence of filamentous yellow pigmented bacteria and BGD severity.

Pathology BGD usually results from gill injuries caused by environmental factors but once the bacteria become established in the gills it plays a major role in mortalities. BGD generally begins with hyperplasia of the gill epithelium caused by mild but chronic toxins or water irritants. F. branchiophilum has large numbers of fimbriae-like surface structures that bridge the spaces between the bacterium and gill tissue and appear to be involved in attachment to the gill surface (Spear et al. 1991a). Bacterial colonization and lamellae fusion will begin on the distal end of filaments infected with BGD, while in “nutritional gill disease” (due to insufficient dietary pantothentic acid), lamellar and filament fusion begins at the proximal end of the filament and progress distally. Speare et al. (1991a, b) confirmed and expanded on earlier bacterial colonization observations (Kudo and Kimura 1983). They noted that several gill changes at an ultrastructure level occurred immediately preceding colonization. The cause of these changes was not determined but environmental conditions were not considered to be a factor. Microridges of superficial filament epithelium showed cytoplasm blistering and a slight irregularity of filament tips, all of

which suggest mild hyperplasia. “Explosive” morbidity and acute mortality coincide with extensive bacterial proliferation on the lamellar surfaces, epithelial hydropic degeneration, necrosis, and edema. Lamellar fusion and epithelial hyperplasia were later detected in subacute (2–5 day) or chronic (7–14 day) infections (Spear et al. 1991a). F. branchiophilum does not appear to become systemic, even when injected IP, as it cannot be isolated at a later time from internal organs. In the case of ABGD, gill epithelium appeared swollen without evidence of single cell pigment (Ostland et al. 1999). In several cases, lamellar syncytia occurred while the lamellae were shortened and stubby with swollen epithelial cells. It has been suggested that F. branchiophilum physically occludes the gill surfaces and inhibits respiratory exchange (Wakabayashi and Iwado 1985). However, Speare et al. (1991b) challenged that suggestion because fish have large areas of underutilized gill surfaces and massive numbers of bacteria do not substantially cover the tissue–water interface. Why BGD-infected fish behave in a manner that suggests a lack of sufficient oxygen has not been adequately explained. However, oxygen consumption data by Wakabayashi and Iwado (1985) supports the theory that increased opercula movement increase water flow across the gills for better oxygenation of the blood. Trout infected with BGD develop a significant decrease in serum Na+, Cl− and an osmolarity imbalance, which results in hemoconcentration (Byrne et al. 1991). Fish with BGD exhibit increased respiration (tachybranchia) but are not hypoxic; therefore, the tachybranchia may not be a product of impaired oxygen exchange. It was suggested that fluctuation in blood components, other than those which are acid-base related, is responsible for death in BGD infected fish. These circulatory disturbances result from a loss of blood electrolytes, which triggers a fluid shift from extracellular to intracellular, thus leading to death. Also, Wakabayashi and Iwado (1985) concluded that a breakdown in gas exchange

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at the gills causes failure to provide sufficient oxygen to remove excess muscle lactate.

Significance BGD is one of the most common diseases of juvenile salmonids. Because BGD is often a subacute to acute condition with high mortality, it is considered a potentially serious problem in trout and salmon culture. The Fish Pathology Laboratory at Ontario Veterinary College, University of Guelph, Ontario, Canada, reported that BGD accounted for approximately 21% of all samples submitted for diagnosis from fish farms (Spear and Ferguson 1989). In 1995, Spear et al. (1995) reported that BGD accounts for 18% of the total diagnostic case submissions to the Atlantic Veterinary College.

Bacterial cold-water disease Bacterial cold-water disease (BCWD), originally referred to as “peduncle disease” because of the principal lesion location, causes low to moderate mortality among fingerling to production size cultured salmonids and occasionally other species. A second condition, known as rainbow trout fry syndrome (RTFS) in salmonid fry, is caused by the same pathogen. The etiology of both of these syndromes is F. psychrophilum. Bernardet et al. (1996) proposed reclassification of the Flavobacterium—Flexibacteria—Cytophaga complex and suggested the pathogen be named Flexibacter psychrophila and has also been named Cytophaga psychrophila but these three specific epitaphs are synonyms; in this text we will use F. psychrophilum.

Geographical range and species susceptibility While bacterial cold-water disease occurs throughout most trout and salmon growing regions of North America, it most seriously

affects salmon culture in the northwestern United States and western Canada. The disease also occurs in Japan (Wakabayashi et al. 1991), the United Kingdom (Austin 1992), Europe (Wiklund et al. 1994), and Australia (Schmidtke and Carson 1995). Ekman et al. (1999) found the pathogen in Atlantic salmon returning from the Baltic Sea to spawn. All salmonid species are suspected to be susceptible to F. psychrophilum; however, rainbow, brook, lake, and cutthroat trout and coho, sockeye, chum, and Atlantic salmon are known to be susceptible (Holt 1993). F. psychrophilum occasionally cause disease in nonsalmonids having been detected in European eels, carp, tench, and crucian carp in Germany (Lehmann et al. 1991). The bacterium was also reported in wild ayu in Japan (Iida and Mizokami 1996).

Clinical signs Clinical signs of BCWD vary with size and age of fish. The caudal peduncle appears whitish and the skin becomes necrotic, detached, and sloughs, exposing underlying muscle (Figure 14.6). In fish up to 1 year of age, the caudal peduncle lesion is the most typical clinical sign of BCWD. Muscle necrosis continues to the point that the vertebral column becomes exposed and the tail is nearly separated from the body. Lesions may also appear on the cranium, dorsally or laterally on the body, on the isthmus, mandible, or the vent. Fin and operculum hemorrhages and pale gills can occasionally be seen in infected fish with darkly pigmented, moribund fish observed late in epizootics. Exophthalmia, gill hemorrhages, and anemia may be evident. Internally, petechia may be present on the liver, pyloric cecae, adipose tissue, heart, swim bladder, and occasionally on the peritoneal lining. Liver and kidney are often pale or brown and coelomic fluid may be red-tinged. Some BCWD survivors may develop vertebrae deformities and/or nervous disorders that

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Figure 14.6 Rainbow trout with typical lesion on the caudal peduncle (arrow) associated with bacterial cold-water disease (Flavobacterium psychrophilum).

cause them to swim in a spiral (Kent et al. 1989). In RTFS-infected sac fry, the yolk membrane becomes eroded (Wood 1974), while older fish exhibit lethargy with labored respiration and sometimes spiral swimming, loss of equilibrium and spinal curvature (Kent et al. 1989, Bebak et al. 2007). Ulcerative lesions may occur on the cranium and vertical surfaces. The fish may be anorexic with exophthalmia, pale red gills, and pale kidneys, intestine, and liver.

Diagnosis Bacterial cold-water disease is diagnosed by recognition of characteristic clinical signs, detection of elongated vegetative cells in stained smears of lesion and isolation of F. psychrophilum on appropriate media. Cytophaga and TYG agar can be used for isolation, but a modified cytophaga media, either broth or agar, is the usual culture substrate (Bullock et al. 1974a). Cipriano et al. (1996) failed to obtain F. psychrophilum growth on tryptic soy, MacConkey, or triple sugar iron agar. Generally, the organism can be isolated from skin mucus, muscle lesions, and internal organs. On agar F. psychrophilum, colonies are pale yellow, flat, and spreading with a thin

irregular edge or smooth, entire, and yellow. Cultures may be a mixture of the two colony types (Holt 1993). Development of yellow colonies on cytophaga agar requires 14 days at 8◦ C (Iida and Mizokami 1996). The pathogen is strictly aerobic with an optimum growth temperature of about 15◦ C but will grow at temperatures from 4 to 25◦ C. Poor growth occurs at the temperature extremes and not at all at 30◦ C (Bernadet and Kerault 1989; Holt et al. 1989). F. psychrophilum does not tolerate NaCl concentrations above 1% in media. F. psychrophilum can be detected in fixed wax-embedded tissues of heart and spleen from rainbow trout using a PCR procedure (Crumish et al. 2007).

Bacterial characteristics On stained smears from lesions the bacterium is a slender, Gram-negative, bacillus, with rounded ends, that measures 0.3 × 2–2.5 µm. In broth culture, cells may be several times this length with some branching (Holt 1993). F. psychrophilum is motile by gliding and does not produce fruiting bodies. Biochemical properties do not vary greatly among different strains of F. psychrophilum (Table 14.5) (Holt 1993). Generally, the bacterium does not use carbohydrates but

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Table 14.5 Biophysical and biochemical characteristics of 28 isolates of Flavobacterium psychrophilum. Test/Characteristic

Growth in broth 1% tryptone 1% Casamino acid 0.1% sodium lauryl sulfate Nitrate reduction Production of Ammonia Hydrogen sulfide Catalase Cytochrome oxidase Acetylmethyl carbinol Indole

Reaction

+ − − − + − + − − −

Test/Characteristic

Degradation of Cellulose Starch Casein Gelatin Albumin Elastin Collagen Chitin Tyrosine Carbohydrate oxidation Glucose Lactose Sucrose

Reaction

− − + + + + + − − − −

Source: Holt (1993). +, positive; −, negative.

metabolizes casein, gelatin, albumin, and collagen. It is cytochrome oxidase positive by most accounts, weakly catalase positive, does not produce H2 S, and contains flexirubin pigments, which are of diagnostic value. Some strains degrade elastin. It is rare for F. psychrophilum to be found in the Southern Hemisphere but Schmidtke and Carson (1995) did isolate it from Atlantic salmon in Australia. Characteristics of the Australian isolate agreed with those of isolates from other parts of the world. Most strains of F. psychrophilum are generally biophysically and biochemically similar and serologically homogeneous; isolates from New Hampshire, Michigan, and Alaska were antigenically similar to Oregon strains (Holt 1993). Comparing F. psychrophilum isolates from three Pacific Northwest (United States) salmon hatcheries, Cipriano et al. (1996c) confirmed the lack of biochemical diversity and there was almost no serological diversity among the isolates. However, in spite of biochemical and serological homogeneity, diversity in ribotyping among the four isolates was noted. Madetoja et al. (2001) compared eight isolates of F. psychrophilum from Finland to isolates from Estonia, Sweden, and the

type strain NCIMB 1947 using phenotypic and genotypic characters. Of the multiple isolates, they found generally biochemical homogeneity with only minor differences. However seven different antigenic isolates suggested a new F. psychrophilum serotype. Serum agglutination positively identifies F. psychrophilum (Holt 1993), but Lorenzen and Karas (1992) used a rapid immunofluorescent diagnostic method on spleen imprints of diseased rainbow trout fry. A slight crossreactivity with F. columnare may occur, but this can be avoided by adsorption with the “columnaris” organism. Bertoleni et al. (1994) examined cell-free ECP by SDS-PAGE and proteases of 32, 86, 114, and 152 kDa and found that isolates taken from ayu in Japan were similar to those taken from coho salmon in the United States. Lindstrom et al. (2009) successfully used PCR, a quantitative enzyme-linked immunosorbent assay and filtration-based fluorescent antibody test as tools to detect F. psychrophilum in kidney and ovarian fluid to screen broodstock for the pathogen. The FAT test detected the bacterium in 100% of kidney tissue from 50 asymptomatic and 74% of ovaries from coho salmon and 42% from ovaries of rainbow trout.

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Epizootiology Bacterial cold-water disease normally occurs in spring when water temperatures range from 4 to 10◦ C, but Lehmann et al. (1991) noted that nonsalmonid infections occurred at water temperatures of 8–12◦ C. Experimental F. psychrophilum infections were induced by Holt et al. (1989) in coho and chinook salmon and rainbow trout at temperatures of 3–15◦ C; as water temperatures rose past 15◦ C, disease severity decreased. Rainbow trout fry syndrome affect newly hatched rainbow swim-up fry and advanced fry less than a couple months of age but may also be associated with low water temperature. Bebak et al. (2007) discussed a case history of RTFS in 13-week-old rainbow trout in which the mortality was about 100/day out of 35,000 individuals. It was thought that the infection may have been introduced into the tank population by contaminated watertesting equipment and spread from tank to tank via equipment used daily in tending the fish. Other sources of the pathogen, water supply, or vertical transmission were not totally ruled out. The disease was arrested by instituting prudent hygienic management practices such as removing dead fish twice a day, disinfecting utensils between tanks and disinfecting hands. Infections of F. psychrophilum are generally more severe in young fish but are significant in older fish as well (Leek 1987). Mortality in yolk-sac fry generally ranges from 30 to 50% and as fish begin feeding, it is usually in the 20% range. In Australia, mortality among Atlantic salmon reached as high as 80% (Schmidtke and Carson 1995). Chronic infections may develop in yearling coho and other Pacific salmon during winter months when typical peduncle lesions are seen and low-grade mortality occurs. Infected fish can be anemic; a condition that is often compounded by presence of an erythrocytic inclusion body syndrome virus (Leek 1987). Experimentally, F. psychrophilum virulence varies with bacterial strain. Mortality in coho

salmon infected with different strains varied from 0 to 100% (Holt 1993). When Bertolini et al. (1994) compared the pathogenicity of 29 F. psychrophilum isolates and classified them into four protease enzymes group, they noted 2 groups were generally more virulent than the other groups. Two separate physical manifestations may be noted following an active BCWD infection. Some surviving fish may have spinal deformities; a problem first reported by Wood (1974) when he described infected fish that had compressed vertebrae, lordosis, and scoliosis. Other fish may display nervous disorders such as spinning, swelling in the posterior skull, and dark pigmentation on either side. F. psychrophilum can readily be isolated from brain tissue but less often from visceral organs (Meyers 1989). F. psychrophilum is a rather common microbe of water and soils; however in general, carrier fish are considered the principal reservoir for the pathogen. Although the natural reservoir for F. psychrophilum is not fully understood, it is accepted that adult coho and chinook salmon, and in all probability rainbow trout and other salmonids, serve as carriers (Bullock and Snieszko 1970). This does not rule out contaminated water, soil, or utensils. According to Cipriano et al. (1996c), F. psychrophilum was isolated from kidney and mucus of asymptomatic chinook and coho salmon even though no significant mortality was occurring. Bacterial concentrations were as high as 3.0 × 106 CFU/g of tissue. They found that by combining results from both kidney and mucus, F. psychrophilum could be isolated from 66% to 88% of tested fish. Horizontal transmission is highly likely, although experimental transmission from fish to fish cannot be achieved unless the mucous layer and epithelium are injured. It is generally concluded that members of the Flavobacterium group are a normal part of trout and salmon skin microflora, and if conditions are favorable, it can become pathogenic. F. psychrophilum has been isolated from the spleen, kidney, ovarian fluid, and milt of

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mature fish, therefore suggesting that transmission may occur on eggs during spawning (Holt 1972). The organism was detected in 20–76% of sexually mature female chinook and coho salmon in hatchery populations and in 0–66% of males at the same facility (Holt 1993). In their study of Atlantic salmon in the Baltic Sea returning to spawn, Ekman et al. (1999) found that salmon carried the pathogen in the sexual products of both male and female, indicating that transmission from brood fish to offspring should be considered an important route of infection. Vertical transmission of F. psychrophilum was also studied by other researchers by its detection in ovarian fluid of adult fish. Taylor (2004) used a nested PCR that targeted the 16S rRNA gene to detect the pathogen in ovarian fluid and the yolk material within the egg of coho, chum, and chinook salmon and steelhead trout. The pathogen was detected in ovarian fluids and unfertilized eggs from all species studied, thus supporting the strong possibility of vertical transmission. F. psychrophilum was detected within eggs of Atlantic salmon from seven rivers in New England by Cipriano (2005) who concluded that the pathogen was established in the ova of salmon in the different New England watersheds. In determining the occurrence of F. psychrophilum in hatcheries in Denmark, Madsen et al. (2005) found the pathogen in ovarian fluid and milt of rainbow trout, indicating that the brood fish were a reservoir for the pathogen. However, they suggested that actual transmission studies be carried out to prove vertical transmission. Lindstrom et al. (2009) showed strong evidence of the potential for vertical transmission of F. psychrophilum via reproductive products. To illustrate that F. psychrophilum can be transmitted via eggs, Ekman et al. (2003) performed nanoinjection of the pathogen into ova. Many injected eggs died before hatching but of those that hatched the pathogen was not only recovered from sick fish but clinical RTFS developed, thus demonstrating that the disease can be transmitted via reproductive products.

Whether or not anadromous salmon carry the organism through the marine maturation phase is not clear. Upon returning to freshwater, these fish have a low incidence of F. psychrophilum infection; however, the longer they remain in freshwater, the higher the infection incidence, suggesting that adults become either infected or reinfected upon returning to freshwater.

Pathology Bacterial cold-water disease is a septicemia; however, the most obvious pathology is a necrotic lesion on the caudal peduncle with sloughing skin. Using PCR Crumish et al. (2007) indicated that fry suffering from F. psychrophilum infection showed evidence of myocarditis. Although bacterial cells can be found throughout most organs and tissues of infected fish, death is generally attributed to severe heart lesions (Wood and Yasutake 1956). There is usually very little inflammation associated with BCWD, but some researchers have noted mild mononuclear infiltration of macrophages in diseased tissues. It has been suggested that F. psychrophilum ECP cause characteristic epithelial lesions in fish injected with either live bacteria or cell free ECP. Internal organ lesions were also found in fish injected with either live bacteria or crude cellfree substances. Skin and skin mucus abrasion dramatically enhance invasion of F. psychrophilum into fish during bath or cohabitation challenges; however, the shedding rate of F. psychrophilum by infected fish is associated with water temperature and the mortality of infected fish (Madetoja et al. 2001). Furthermore, high numbers of bacteria in the water from dead fish is greater than numbers of cells shed by live infected fish, which leads to enhanced transmission. The results emphasize the importance of removing dead and moribund fish from rearing tanks in order to diminish the transmission rate. Orally infected fish did not result in significant infections.

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Early pathogenicity studies of F. psychrophilum indicated that exotoxins did not affect the disease process, but Pacha (1968) noted that proteases may be involved. Later, it was found that steelhead trout infected with F. psychrophilum ECP developed microscopic lesions similar to those seen when fish were infected with live, virulent organisms. An immunohistochemical method was used by Evensen and Lorenzen (1996) to show that F. psychrophilum was localized in the monocyte–macrophage system, skin lesions, retina, and choroid gland of the eye. Dermal changes in superficial or deep ulcers extended to the subcutaneous tissue or the musculature included inflammatory cell infiltrates in which polymorphonuclear cells were contained in their cytoplasm. Inflammation was also noted in the retina. The virulence of F. psychrophilum may be related to their ability to attach to gill tissue. Nematollahi et al. (2003) found that highly virulent strains attached more readily than low virulent strains. Moreover the ability to adhere to gills was influenced by the amount of organic material and nitrite in the water, but temperature had no adhesion-enhancing effect.

Significance Bacterial cold-water disease and rainbow trout fry syndrome are significant salmonid culture problems in juvenile populations in the Pacific Northwest region of North America where it is not only the most frequently reported fish pathogen but also causes highest mortality (Holt 1993). However, the pathogen and associated diseases occurs in most regions of the world where salmonids are cultured.

narum (Sanders and Fryer 1980). The causative agent was once classified as a Corynebacterium. BKD was initially described in Atlantic salmon in Scotland in 1930 and called Dee disease (Smith 1964). It was reported for the first time in the United States in 1935 (Belding and Merill 1935). BKD has also been called corynebacterial kidney disease and salmonid kidney disease.

Geographical range and species susceptibility BKD exists in many freshwater and marine environments where trout and salmon occur with the possible exception of Australia, New Zealand, and the former Soviet Union (Fryer and Sanders 1981; Evelyn and ProsperiPorta 1992). There are also no reports of BKD in China. Severe BKD has been reported in Canada, Chile, Great Britain, France, Germany, Iceland, Italy, Japan, Spain, and the United States. Most outbreaks in the United States occur in the northwest, upper midwest, and northeast (Mangin 1991). All salmonids are susceptible to R. salmoninarum, but brook trout and chinook salmon are considered most severely affected (Fryer and Sanders 1981; Evelyn 1993). Natural BKD outbreaks occur only in cultured salmonids, but it is not unusual to find the disease organism in wild salmonids where clinical infections are rare. Experimental infections of BKD have been established in sablefish (Bell et al. 1990), Pacific herring (Traxler and Bell 1988), shiner perch (Evelyn 1993), common shiner, and fathead minnows (Hicks et al. 1986).

Clinical signs Bacterial kidney disease Bacterial kidney disease (BKD) is a chronic, rarely subacute, bacterial infection of salmonids caused by Renibacterium salmoni-

External clinical signs of BKD vary somewhat among fish species but are most obvious in terminal stages of disease (Fryer and Sanders 1981). The most common early signs of infection are dark pigmentation,

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Salmonid Bacterial Diseases 381

(a)

(b)

Figure 14.7 Bacterial kidney disease of salmonids. (a) Rainbow trout with epidermal, ulcerative lesions (arrow) typical of chronic Renibacterium salmoninarum infection. (b) Granulomatous lesions (large arrow) in the swollen, corrugated kidney of brook trout. Note the petechia on the adipose tissue and intestine (small arrow). (Photos courtesy of G. Camenisch.)

exophthalmia, hemorrhages at the base of fins, abdominal distension, and lethargic swimming. Occasionally, superficial blisters and ulcerative abscesses appear on the body surface, particularly in rainbow trout (Figure 14.7). A bloody acetic fluid may occur in the coelomic cavity and petechia may appear in muscle under the peritoneal lining. As BKD progresses, small grayish-white, granulomatous lesions

develop in the kidney and to a lesser extent in the liver and/or spleen; the granulomas contain leukocytes, cellular debris, macrophages with phagocytosed bacteria, and cell-free bacteria. As disease progresses, the number and size of granulomas increase in all internal organs and the kidney becomes swollen with a “corrugated” surface (Figure 14.7). Although systemic BKD infections are normal, infection

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may occasionally occur only in the eye or on the skin (Hoffman et al. 1984).

Diagnosis Infections of R. salmoninarum are diagnosed by noting clinical signs and detection of Grampositive encapsulated (halo) diplobacillus bacteria in internal organs. Fluorescent antibody technique or ELISA are more sensitive than the Gram stain and may be used on smears from infected kidney tissue to detect and/or confirm identity of the organism (Bullock et al. 1980). Unless R. salmoninarum is present in abundance, Gram stained smears are less precise than FAT or ELISA because pigment granules (melanin) can be confused with the pathogen. To enhance confidence in Gramstained smears, they are compared to methylene blue and unstained smears. By comparing these slides, abundance of pigment granules that are brown and refractile, the presence of R. salmoninarum in preparations can be determined. Nevertheless Gram staining to detect R. salmoninarum often results in a subjective analysis. Isolation is the only method for detecting viable R. salmoninarum, but because of its fastidious nature, routine culture is difficult. The pathogen grows on general bacterial culture media, but Evelyn (1977) developed an improved medium, “kidney disease medium” (KDM-2), using peptone yeast extract agar supplemented with cysteine and serum. Growth of R. salmoninarum is improved on KDM-2 by introducing “cross feeding”, whereby, a nonfastidious feeder (nurse organism) is placed next to the BKD organism (Evelyn et al. 1989). Another innovation that further improves R. salmoninarum growth on KDM-2 is addition to the medium of a small amount of KDM-2 broth in which R. salmoninarum had grown (supplemental, SKDM) (Evelyn et al. 1990). At 15◦ C, the organism may require up to 20 days for appearance of nonpigmented, creamy, shiny, smooth, raised, entire colonies

measuring approximately 2 mm in diameter. Because the organism grows slowly, precautions must be taken to insure that culture media remains moist during incubation. Detecting R. salmoninarum in subclinical carrier fish is challenging, but with improved FAT and more sensitive ELISA procedures, detection is more common (Pascho and Mulcahy 1987). Elliott and Barila (1987) developed a membrane filtration-fluorescent antibody technique (MF-FAT) that detects less than 102 R. salmoninarum cells/mL of coelomic fluid. Lee (1989) detected less than 102 cells/g of enzyme digested kidney using FAT and membrane filtrates. Treatment of kidney smears with the organic solvents xylene, acetone, or methanol for 30–60 minutes prior to FAT staining improved fluorescent intensity but did not alter detection of R. salmoninarum (Cvitanich 1994). Combining MF-FAT with a culture on SKDM can be of value in detecting R. salmoninarum carriers, particularly when carrier rate is low and no single test is conclusive (Teska et al. 1995). As part of a screening program to detect BKD, Griffiths et al. (1996) used kidney tissue and ovarian fluid from Atlantic salmon and compared pathogen detection by S-KDM agar culture, FAT, ELISA, and Western blot. Cultures from kidney on S-KDM agar identified the highest percentage of samples positive (34.5%) for R. salmoninarum with lower sensitivity recorded by indirect IFAT (12%), ELISA (4%), and Western blot (1%). Culturing ovarian fluid on S-KDM agar was best in which 19.5% was positive compared to IFAT (7.5%) and ELISA (0%). In many instances, R. salmoninarum was detected in either kidney tissue or ovarian fluid but not both. These authors recommended using a combination of methods to increase reliability for detecting R. salmoninarum. By culturing ovarian fluid in S-KDM broth and then applying Western blot assay, positive samples were increased by 32%. Five methods for detecting subclinical R. salmoninarum in coho salmon were compared

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by Cipriano et al. (1985). They found that Gram stain and direct fluorescent antibody were the least sensitive and counter immunoelectrophoresis the most sensitive. Sakai et al. (1987) modified an ELISA technique to a peroxidase–antiperoxidase procedure for detecting R. salmoninarum in coho salmon and found the modified method approximately ten times more sensitive than conventional FAT. The procedure also gave excellent results in a BKD survey. Eight different methods for detecting R. salmoninarum in supernatants from rainbow trout kidneys were compared by Sakai et al. (1989). Gram stain and immunodiffusion were the least sensitive, FAT was very sensitive, but ELISA (direct and indirect) was most sensitive by detecting the lowest concentration of bacterial cells when used in conjunction with a dot blot assay. The latter method detected 102 –103 cells per gram of kidney tissue. In comparing a double sandwich ELISA procedure with culture on SKDM-2 to detect R. salmoninarum, Gudmundsdottir et al. (1993) sampled 1239 individual fish from 12 locations in Norway. Seven of the 12 groups tested negative by both methods, five groups tested positive by ELISA, and four were positive by culture. Within the 12 groups, a significantly higher number of fish were positive by ELISA than by culture. From these studies, as a whole it appears that no one single R. salmoninarum detection method in carrier fish can be relied upon but a combination of culture, serological, or dot-blot methods should be used for most dependable results. This was further borne out by Jansson et al. (1996), who attempted to improve BKD detection, sensitivity, and reliability by comparing isolation on SKDM with commercial monoclonal and polyclonal preparations. By culture, 45% of fish were positive and by combining culture with polyclonal ELISA 50% were positive. They determined that bacterial culture, polyclonal ELISA, and a commercially available monoclonal based ELISA system were equally accurate in detecting R. salmoninarum. A quantitative polymerase chain reaction (QPCR) was compared to ELISA by Powel

et al. (2005) to detect varying levels of R. salmoninarum in tissues and body fluids of Chinook salmon injected with varying levels of the pathogen. Fish receiving high concentrations of the bacterium showed good positive correlation between QPCR and ELISA; however, at lower levels of infection, there was little correlation. An ELISA was used to detect soluble R. salmoninarum 57 kDa protein (p57 or “F” antigen) in tissues and body fluids. Griffiths et al. (1991) showed that in overtly infected Atlantic salmon, detection of soluble p57 by Western blot was significantly more sensitive and reliable than detection of whole cells by direct FAT in tissue homogenates. Whether or not the Western blot is applicable in detection of R. salmoninarum carrier fish with low level infections is not clear; however, about one third of epizootic survivors were seropositive for the p57 using this method. Indications are that ELISA is the most sensitive method for detecting soluble antigens in subclinical R. salmoninarum infections. However, p57 can cross-react with Corynebacterium aquaticum and Carnobacterium piscicola, two other Gram-positive bacteria (Bandin et al. 1993; Toranzo et al. 1993). Wood et al. (1995) showed that a 60 kDa protein may actually be responsible for this cross-reactivity and by using MAB with p57 they eliminated cross-reactivity. Using rocket electrophoresis and cross adsorption analysis, Getchell et al. (1985) showed a common p57 antigen in seven isolates. The soluble, heat stable antigen is released into culture media and is considered a primary mediator of the bacterium’s virulence (Bruno 1990).

Bacterial characteristics Renibacterium salmoninarum is a Grampositive bacillus (usually in pairs) that measures 0.3–0.5 × 0.6–1.0 µm (Young and Chapman 1978). It does not produce spores nor is it

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384 Bacterial Diseases Table 14.6 Biophysical and biochemical characteristics∗ of Renibacterium salmoninarum. Characteristic

Reaction

Characteristic

Cell morphology Gram stain Production of Acid phosphatase Alkaline phosphatase Caprylate esterase Catalase ∂-glucosidase ß-glucosidase Leucine arylamidase ∂-mannosidase Oxidase Trypsinase Degredation of Casein Tributyrin Tween 40 Tween 60 Tween 80

Diplobacillus +

Acid from sugars Growth at pH 7.8 0.25% bile salts 0.0001% crystal violet 0.001% methylene blue 0.00001% Nile blue 1% sodium chloride Utilization of 4MU-butyrate 4MU-heptanoate 4MU-laurate 4MU-nonanoate 4MU-oleate 4MU-palmitate 4MU-propionate

+ + + + + − + + − + + + + + −

Reaction

+ − + − + + (poor) + + + + + − +

Source: Goodfellow et al. (1985); Austin and Austin (1987). +, positive; −, negative. ∗Negative for production of butyrate esterase, chymotryupsinase, cystine arylamidase, ∂-fucosidase, ∂-galactosidase, ß-galactosidase, ßglucosaminidase, ß-glucuronidase, myristate, esterase, and valine arylamidase. Negative for degredation of adenine, esculin, arbutin, chitin, chondroitin, DNA, elastin, gelatin, guanine, hyaluronic acid, hypoxanthine, lecithin, RNA, starch, tyrosine, and xanathine.

acid-fast or motile. Its optimum growth temperature is 15◦ C; growth slowed when temperature is reduced to 5◦ C or elevated to 22◦ C and no growth occurs at 37◦ C (Smith 1964). Renibacterium salmoninarum is biochemically homogeneous (Table 14.6). The bacterium is proteolytic in litmus milk, produces catalase, and does not liquefy gelatin. It produces acid and alkaline phosphatase, caprylate esterase, glucosidase, leucine arylamidase, a-mannosidase, and trypsinase. The bacterium degrades tributyrin, Tween 40 and 60, but not Tween 80. The organism grows at pH 7.8 (optimum) on 0.0001% crystal violet, 0.00001% Nile blue, and poorly in l% sodium chloride (Goodfellow et al. 1985). Isolates of R. salmoninarum from various parts of the world are serologically homogeneous when using polyclonal antisera (Bullock et al. 1974b). However, when Arakawa et al. (1987) used monoclonal antibodies in an ELISA system, they identified nine different R.

salmoninarum isolates from the United States, Norway and England, of which there were three serological groups. One monoclonal antibody to a Norwegian isolate reacted with all other isolates. Genomic DNA was extracted from Corynebacterium aquaticum, C. piscicola, and a Gram negative Pseudomonas maltophila, all of which had reacted with polyclonal antisera for R. salmoninarum in an indirect FAT (Brown et al. 1995). The genomic DNA of these organisms was negative by PCR designed to amplify a segment of the gene encoding p57 of R. salmoninarum. Although R. salmoninarum antibodies reacted with antigens of these three bacteria, the results further suggest that cross-reactivity is not associated with p57. To compliment the high degree of specificity of R. salmoninarum, Starliper (1996) showed that there is little genetic diversity in the North American isolates.

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Epizootiology BKD is primarily a disease of hatchery fish, but it also occurs in wild salmonids, including anadromous populations but is uncommon in salmon in seawater. Renibacterium salmoninarum can cause overt disease or can be present only in the carrier state. In proceedings of the National Workshop on Bacterial Kidney Disease, Mangin (1991) noted that between 1986 and 1991, the pathogen was found at 63 locations in the United States where it caused no disease and at 97 locations where overt disease occurred. Nine states reported asymptomatic presence of the pathogen, and 16 states reported its presence in either the carrier state or as overt disease. These data suggest that factors other than pathogen presence are related to BKD outbreaks and species dependency. The susceptibility to R. salmoninarum, and possibly other pathogens, may change over time. Purcell et al. (2008) suggested that susceptibility to R. salmoninarum changed in Chinook salmon in Lake Michigan since their introduction in the late 1960s. Susceptibility of the Lake Michigan population was less than that of the original source population (Green River, Washington), possibly in response to a pathogen-driven selection. The presence of very low numbers of R. salmoninarum in some brood rainbow trout without clinical disease may be explained by the possibility that unless the host becomes stressed, the pathogen remains dormant. Starliper and Teska (1995) found in a study of brood brook trout and their progeny that overall prevalence of R. salmoninarum decreased over a 3-year period; carrier rate ranged from 11.6 to 64.4% depending upon type of detection method used and year of study, but adults and progeny remained asymptomatic for the entire period. However, fish infected by injection with a high dose of the pathogen shed bacteria into water 12 days postinjection with the range of bacterial concentration in the water from undetectable to 1,850 cfu/mL of water based on bacteriological

culture (McKibben and Pascho 1999). Results suggest that Chinook salmon injected with R. salmoninarum can be used as a source of infection in cohabitation challenge studies. Water quality may influence cumulative mortality and disease severity of BKD. A higher BKD incidence was noted at hatcheries with soft water than those using water with high total hardness; however, this relationship has not proved to be absolute (Warren 1963). BKD has an adverse effect on salmon smolt survivability as they move from freshwater to saltwater. Over a 150-day period, coho salmon smolts experienced over 17% cumulative mortality in seawater compared to only 4% in a group of siblings held in freshwater (Fryer and Sanders 1981). Paterson et al. (1981b) confirmed that heavy losses of Atlantic salmon smolts caused by BKD occurred as they became acclimated to seawater. Sanders et al. (1992) found that 20% of hatchery released, and wild salmonids migrating down the Columbia River were infected with R. salmoninarum. Prevalence in fish held in freshwater was 9% compared to 46% prevalence in those held in saltwater. Implications are that BKD becomes more severe when salmon reach the marine environment but after an initial epizootic losses decrease. In a survey of six chinook salmon hatcheries in the Columbia River and Snake River basins, Maule et al. (1996) found that prevalence of R. salmoninarum infections on the Snake River decreased from >90% to <65% from the beginning to the end of a 5 year study; this decrease was attributed to initiation of health management practices on the hatcheries. It was noted that the increase in infection rate as fish migrated down the Snake River was not seen in the Columbia River. These variations may have resulted from differences in river conditions and distance from hatcheries to dams. Using ELISA, Meyers et al. (1999) detected a higher concentration of R. salmoninarum soluble antigen in adult chinook salmon that matured in freshwater than in the same

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cohort fish spawned after maturation in seawater. This leads to a practical strategy for maturing Chinook salmon broodstock in seawater to reduce prevalence of R. salmoninarum positive parent fish. Fish culturists who do not have the option of maturing fish in seawater should note the importance of optimizing the freshwater maturation environment to minimize the prevalence of R. salmoninarum antigen in adult Chinook salmon. BKD is a reportable disease in the United Kingdom and the European Union. In Scotland, new outbreaks of the disease declined from 1990 through 2002 for Atlantic salmon with some year to year variations (Bruno 2004). There was only one recorded case of BKD in freshwater reared Atlantic salmon. BKD is uncommon in rainbow trout cultured in Scottish seawater; it was identified only in 1993 and between 1998 and 2000. BKD is affected by water temperature. Most epizootics in chinook and sockeye salmon occur during fall (12–18◦ C) and winter (8–11◦ C) and generally not in summer; Smith (1964) observed that mortality accelerated only when temperatures rose above 8◦ C. Mortalities in cool water tend to be slow and chronic compared to a more subacute mortality pattern in warmer water. Water temperature also influences time between exposure and death. At 11◦ C, deaths occur at 30–35 days post exposure to the pathogen compared to 60–90 days at 7–10◦ C. Sanders et al. (1978) presented conflicting data concerning the relationship of temperature and mortality in BKD. At 6.7–12◦ C, 78–l00% mortality occurred, but as temperatures rose to 20◦ C, mortality declined to a low of 8–14%, indicating the probability that factors other than temperature play a role in BKD-related mortality. Lovely et al. (1994) established R. salmoninarum carrier populations of Atlantic salmon at 7–8◦ C. Bacteria were isolated for up to 29 weeks from these fish, after which time, only 4% of fish were carriers as determined by immunoblot of the soluble R. salmoninarum antigen.

In addition to differential species susceptibility, various strains within species respond differently to BKD. Beacham and Evelyn (1992b) compared R. salmoninarum susceptibility in three chinook salmon strains and found substantial differences in mortality rates. Susceptibility in the most affected strain was seven times higher than in the least affected. Breeding and genetic manipulation of fish stocks to increase BKD resistance is a management area that should be further exploited. Several studies indicate that feed ingredients affect R. salmoninarum susceptibility. Nutritional studies involving Atlantic salmon indicate that insufficient amounts of dietary vitamin A, iron, zinc, iodine, and other minerals increased R. salmoninarum susceptibility (Paterson 1981b). Woodall and LaRoche (1964) noted that salmon receiving insufficient amounts of iodine were more susceptible to BKD, and when 4.5 mg of iodine per kilogram of feed was added, prevalence of the pathogen was reduced from 95 to 3%. Addition of fluorine to feed at the same feeding rate also reduced prevalence of disease from 38 to 4% (Lall et al. 1985). According to Evelyn et al. (1973), R. salmoninarum was first reported in anadromous salmonids in the United States in the early 1950s. There are indications that the disease is becoming more widespread, but this increase may be due to more sensitive detection methods than to an actual increase in geographical range. There has been a high BKD incidence reported in adult Pacific and Atlantic salmon upon returning to spawn (Evelyn 1993). When salmon migrate from freshwater to seawater, infections continue to develop with some fish succumbing to disease while at sea and those that survive return to spawn and become a pathogen reservoir for the next generation (Banner et al. 1986). BKD also exists in wild or feral trout and salmon populations confined to freshwater. Mitchum et al. (1979) demonstrated BKD in wild populations of brook, brown, and

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rainbow trout in freshwater streams and when uninfected, hatchery-reared trout were stocked into these waters, R. salmoninarum was transmitted to newly stocked fish. The newly infected rainbow trout died in 9 months or less postexposure. In Lake Michigan, coho salmon have been so affected by BKD that a detrimental impact on the salmon fishery resulted (Hnath 1991). Souter et al. (1987) found R. salmoninarum infected Arctic char and lake trout in the Northwest Territory of Canada, an area where no hatchery fish had been stocked. They postulated that this constituted a “natural” presence of the organism. R. salmoninarum does not survive well outside the host; however, Austin and Rayment (1985) did isolate it for 28 days from spiked filter-sterilized river water at 15◦ C, and it is possible that the bacterium can survive undetected for longer periods. Balfry et al. (1996) showed that the organism survived for up to 1 week in seawater. Marking anadromous salmonids with binary coded wire tags (CWTs) for future identification is used extensively. Elliott and Pascho (2001) evaluated the correlation of CWTs and horizontal transmission of BKD. Four months after inserting CWTs in the snout of smolts, 41% and 24% of tagged BKD positive fish in two successive years developed lesions that were confined to the head, particularly tissue around the nares. No nontagged cohorts developed lesions during the 4-month period. Results suggested that CWT marking could be instrumental in BKD invasion. Therefore, to prevent tagging exacerbating BKD during tagging, it was suggested that strict sanitary procedures be implemented during tagging of fish in R. salmoninarum positive populations. Although Koch’s postulates for BKD were fulfilled in 1956 (Ordal and Earp 1956), horizontal transmission was achieved with varying degrees of success. Wood and Wallis (1955) demonstrated R. salmoninarum transmission by feeding infected raw fish viscera to young salmon. It can also be transmitted by cohabita-

tion in either freshwater or seawater. Bell et al. (1984) successfully transmitted R. salmoninarum from deliberately infected fish in cohabitation with naive fish. It was shown by Balfry et al. (1996) that feces containing R. salmoninarum significantly contributes to horizontal transmission via oral ingestion; fish naturally exposed to pathogen laden feces had 70% infectivity compared to 98% in fish orally intubated with contaminated feces. Murray et al. (1992) successfully transmitted BKD to uninfected fish in two ways; chinook salmon were experimentally immersed in a solution containing 104 –106 cells/mL for 15–30 minutes, and the pathogen was transmitted from injected fish to naive fish by cohabitation. Both procedures required 5–6 months for clinical disease to appear. Skin injury to the host enhances infection of R. salmoninarum (Hoffman et al. 1984). Vertical transmission of R. salmoninarum is important because the organism is uniquely adapted to this process. It is small enough to enter the micropyle of the egg, which facilitates transmission; therefore, eggs may become infected during oogenesis or while they are surrounded by bacterial contaminated coelomic fluid prior to, and during, the act of spawning (Bruno and Munro 1986; Lee and Evelyn 1989). Renibacterium salmoninarum becomes a chronic infection, as is the case in trout and Atlantic salmon; it remains through multiple spawning periods or throughout the fish’s life span, thus enhancing vertical transmission. Allison (1958) first reported BKD in offspring from infected fish. However, Amos (1977) proved that the pathogen was transmitted from adults to offspring during spawning and that injection of adults with erythromycin prior to spawning prevented the pathogen from being transmitted to progeny. Bullock et al. (1978b) further demonstrated vertical transmission of BKD and Evelyn et al. (1984) also reported R. salmoninarum within trout and salmon eggs, which further supports the case for vertical transmission.

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Pathology Austin and Austin (1987) hypothesized that R. salmoninarum is a normal resident of the kidney and possibly the gastrointestinal tract of salmonids where it remains dormant in low numbers. In a stressed host, the pathogen will multiply in the kidney or break through the gut epithelial barrier, becoming systemic. This theory is based on the normal nonaggressive activity of the organism. It has also been postulated that an unidentified substance may occur in the kidneys of rainbow trout, which inhibits pathogen growth, thus suppressing its ability to cause disease (Daly and Stevenson 1988). In fact, Hardie et al. (1996) showed that activated rainbow trout macrophages in vitro released H2 O2 , which aided in the inhibition of R. salmoninarum’s growth, suggesting that this event may be important in protective cellmediated immunity. Renibacterium salmoninarum infections are usually systemic and characterized by diffuse granulomatous inflammation primarily in the kidney (Figure 14.7) but they can also occur in the liver, heart, spleen, and muscle. The granulomas are often large, particularly those in the kidney, with zones of caseous necrosis surrounded by epitheloid cells accompanied by lymphoid cell infiltration. These granulomas may or may not be encapsulated. Postorbital eye lesions are also characterized by granulomas with chronic inflammation and infiltration by macrophages and leukocytes (Hoffman et al. 1984). Chronologically, R. salmoninarum develops a septicemia within 4–11 days of experimental infection (Young and Chapman 1978). At about 18 days, bacteria colonize the outer surface of organs and by day 25 penetrates deeper into tissues where it forms granulomas. These findings are congruous with long incubation periods between R. salmoninarum exposure and appearance of clinical disease and chronic mortality. At 12◦ C, Bruno (1986) found that experimentally infected rainbow trout and At-

lantic salmon exhibited initial pathological signs 10 days postinfection and suffered 75 and 80% mortality respectively by day 35. R. salmoninarum was present in kidney collecting ducts and may have affected their function, causing sufficient damage to disrupt normal organ filtration. Death, therefore, was attributed to obliteration of normal kidney and liver structure due to large granulomatous lesions. Death was also partly attributed to heart failure resulting from invasion of the myocardium by phagocytic cells containing R. salmoninarum and release of hydrocytic and oxidizing enzymes from disrupted macrophages. In rainbow trout, Sami et al. (1992) described extended pathogenesis in which glomerulonephritis was associated with chronic BKD. Fifteen months postinfection, the following pathology was noted: fibrosis, adhesions in the Bowman’s capsule, shrinkage or swelling of capillaries, and degeneration of proximal tubules. In experimentally infected coho salmon, three different infection levels were determined by light microscopy and characterized according to bacterial location in injured tissue (Flano ˜ et al. 1996). Changes included destruction of hematopoietic tissue and plasmacytopoietic foci in renal and splenic tissue. Also, epithelioid and barrier cells appeared, which were thought to possibly be a local defense response to the pathogen. Wiens and Kaattari (1991), using MAB showed that the p57 could agglutinate salmonid leukocytes in vitro but whether or not it serves the same function in vivo is not known. However, it was suggested by Daly and Stevenson (1990) that p57 aids the ability of R. salmoninarum to attach to host cells, thus allowing intracellular invasion. The relationship of a 57 kDa protein (“p57”) R. salmoninarum soluble antigen concentration and bacterial load in Pacific salmon was explored by Hamel and Anderson (2002), who determined that the soluble antigen eventually breaks down or otherwise is removed from free circulation. Bacterial load and soluble antigen

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concentration in tissue are strong indicators of fish health and in ovarian fluid are predictors of the survival success of offspring. Results indicate either an increasing antigen removal rate of p57 or decrease of per-bacterium antigen secretion.

Significance Due to BKD’s wide occurrence in anadromous salmonid hatchery stock as well as wild salmon and trout populations, it is considered one of the most serious and controversial diseases of these fish. Though it seldom causes acute mortality, cumulative losses are significant and R. salmoninarum’s unique ability to be transmitted vertically within the egg, its intriguing epizootiology, and absence of effective therapy emphasizes its importance and economic impact on salmonid aquaculture.

Piscirickettsiosis Rickettsia infections have been observed in fish since 1939, but few instances of substantial diseases were reported prior to the mid-1990s (Almendras and Fuentealba 1997). However, since then the disease has become more frequent with expanded geographical range. During 1988, seawater cage cultured coho salmon in Chile experienced a significant epizootic that was eventually determined to be caused by Piscirickettsia salmonis; the disease is called piscirickettsiosis or salmonid rickettsial septicemia (SRS) (Fryer et al. 1990; Cvitanich et al. 1991). In some instances where the etiology was clearly a Piscirickettsia but not necessarily P. salmonis, the pathogen is called rickettsia like organism (RLO). In their review of Piscirickettsia infections, Fryer and Hedrick (2003) discussed transmission, hosts, possible reservoirs, procedures for detection and identification, and current approaches for prevention and treatment.

Geographical range and species susceptibility Piscirickettsia salmonis causes disease primarily in salmonids farmed in seawater cages. The P. salmonis epizootics in Chile involved coho salmon, apparently the most susceptible species, but infections have also occurred in chinook and Atlantic salmon and rainbow trout (Garc´es et al. 1991). Piscirickettsia infections have also been reported in Ireland (Rodger and Drinan 1993), United States (Chen et al. 2000; Maule et al. 2003), Europe (Bravo 1994a), British Columbia, Canada (Brocklebank et al. 1993), and the Mediterranean Sea region (McCarthy et al. 2005); however, it has not been determined if all rickettsia-like organisms are the same. Although disease caused by P. salmonis primarily affect marine fish it has been found in freshwater coho salmon and rainbow trout (Bravo 1994b, Gaggero et al. 1995). Non-salmonid species in which RLO’s have been found are European seabass in the Mediterranean Sea (McCarthy et al. 2005), white seabass in California (Chen et al. 2000,), and tilapia in Hawaii (Maule et al. 2003). Although primarily a disease of salmonids it clearly may also affect non-salmonids (Arkush et al. 2005).

Clinical signs Piscirickettsia salmonis-infected fish are anorexic, anemic, exophthalmic, have pale gills and darkened coloration, and swim lethargically at the surface (Lannan and Fryer 1993). Skin lesions include hemorrhages on the operculum and around the anus, petechia on the abdomen and shallow hemorrhagic ulcers on the body ranging in size from 0.5 to 2 cm in diameter. Firm white nodules (1 cm in diameter) may also be present in the skin. The most characteristic internal lesions are off-white to yellow subcapsular nodules that measure up to 2 cm in diameter throughout the liver (Cvitanich et al. 1991). The kidney

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becomes swollen, spleen is enlarged, and the liver occasionally becomes yellow and mottled. Rodger and Drinan (1993) found RLO infected Atlantic salmon in Ireland. The affected fish had hemorrhages in the skeletal muscle. Other obvious pathology includes splenomegaly, congestion of the pyloric caeca, and the spleen and pale liver. Infected coho salmon and rainbow trout in freshwater had no skin ulcerations but the abdominal cavity was distended with ascites (Gaggero et al. 1995). Petechiae and ecchymosis occur on serosal surfaces of the pyloric caeca, swim bladder, and hind gut (Brocklebank et al. 1993). In the epizootic of tilapia with piscirickettsiosis in Hawaii, there were some clinical signs similar to the disease in salmonids, but there were also some differences (Maule et al. 2003). The fish swam erratically had trouble maintaining depth, and had scattered cutaneous hemorrhage and exophthalmia. Many dead fish had clinical signs. The primary clinical sign of the RLO in European seabass was severe encephalitis.

Diagnosis Diagnosis of P. salmonis involves recognition of gross lesions and use of histochemical stains (H & E, Gram, Giemsa, acridine orange, Gim´enez, Machivello, and PAS) to detect the pathogen in tissue sections or smears (Almendras and Fuentealba 1997). The pathogen can be isolated in time-consuming tissue culture for confirmation. The most susceptible cell lines are CHSE-214, CSE-119, CHH-1, and RTG-2 at 15–18◦ C with up to 14 days required for CPE to develop (Fryer et al. 1990). Cytopathic effect includes cell rounding and development of one or more vacuoles in the cytoplasm. Recently Mauel et al. (2008) reported on a method for culturing P. salmonis on enriched blood agar medium. An indirect FAT was developed for P. salmonis detection in blood films, tissue section, and smears from infected fish (Lannan

et al. 1991). Presumably, this method could also be used to positively identify the pathogen in inoculated cell cultures. P. salmonis can also be detected and identified by immunohistochemistry with peroxidase–anti peroxidase using rabbit anti-P. salmonis polyclonal antibody, ELISA, or PCR. Aguayo et al. (2002) utilized an ELISA, in combination with different monoclonal antibodies against P. salmonis for its detection in coho salmon tissue samples. The results detected seven different isolates and did not cross-react with several other fish pathogens; the test was highly sensitive with a correlation of 98% with the immunofluorescence assay.

Pathogen characteristics Piscirickettsia salmonis is an obligate intracellular bacterium (Fryer et al. 1992). The P. salmonis-type strain (LF-89), on deposit with the American Type Culture Collection as ATCC VR 1361, is a weakly Gram-negative, pleomorphic, usually paired, coccoid cell that varies in diameter from 0.5 to 1.5 µm. It is nonmotile, unencapsulated and PAS, acid fast, and Giem´enez-negative but stains well with H & E, Giemsa, and methylene blue (Cvitanich et al. 1991). Lannan and Fryer (1994) determined that extracellular survival of P. salmonis was a maximum of 3 weeks at 5◦ C and 1 week at 20◦ C when suspended in cell-free Eagles minimum essential medium with 10% fetal bovine serum. Viability of P. salmonis in freshwater was destroyed almost immediately, which may help explain why the disease is seldom seen in freshwater fish. Because RLO are being found in nonsalmonids, there is interest in whether these are similar to isolates from salmonids. Chen et al. (2000) concluded that the bacterium from white seabass in California expressed some antigenic differences from P. salmonis of salmonids, but it was pathogenic to juvenile coho salmon that suffered 80% mortality in 10 days. Arkush et al. (2005)

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compared P. salmonis from white seabass in California to isolates from Chile, Norway, and British Columbia, Canada, using rDNA sequencing and internal transcribed spacers. They determined that isolates from different locations were essentially the same. However, using immunohistochemistry and DNA sequencing, McCarthy et al. (2005) determined that the P. salmonis from European seabass were related to P. salmonis from salmonids but was possibly a different strain.

Epizootiology Piscirickettsiosis occurred from May to November in Chile and cumulative losses at some farms reached 90% with up to 40% mortality per month (Cvitanich et al. 1991). Disease first appeared about 6–12 weeks after fish were transferred from freshwater to saltwater cages. Experimental P. salmonis transmission has been successful via intraperitoneal injection but cohabitation transmission experiments yielded mixed results. Garc´es et al. (1991) failed to observe transmission by cohabitation, but experimental transmission studies by Cvitanich et al. (1991) were successful. It was suggested that natural transmission is through oral transmission. However, Smith et al. (2004) concluded that of three routs of experimental infection (skin, gill, or intestinal) that the primary point of P. salmonis entry through the skin was more effective than intestinal intubations or gill exposure. Vertical transmission of P. salmonis was thought to have occurred in coho salmon when 98.3% of fingerlings from P. salmonis positive broodstock were positive, but only 27% of fingerlings from negative brood fish were positive (Almendras and Fuentealba 1997). The piscirickettsiosis-like disease in tilapia was successfully transmitted horizontally by cohabitation (Maule et al. 2003). Coho salmon and Atlantic salmon were tested for P. salmonis susceptibility by Garc´es et al. (1991). Although typical clinical dis-

ease developed only in the coho salmon, both species suffered nearly 100% mortality at the highest pathogen concentration. The organism was recovered in pure culture from infected fish of both species. An experimental challenge of 16–17 g coho salmon and rainbow trout was carried out in freshwater by Smith et al. (1996). Subsequent studies suggested that the optimum route of entry of P. salmonis in coho salmon in Chile was through the skin and gills, and the oral route may not be the normal method by which the pathogen infests salmonids (Smith et al. 1999). However, it appears that salmonids can be infected by P. salmonis from multiple routes. After intraperitoneal injection, clinical disease occurred in both species in the third week following exposure. At the highest pathogen concentration, cumulative mortality was 60% in the coho salmon and 28% in the rainbow trout. House et al. (2006) showed that the P. salmonis from white seabass was fully virulent for coho salmon; therefore, concluding that these non-salmonid fish can potentially serve as a reservoir of the pathogen for salmonids. The virulence of three isolates P. salmonis from Chile, Norway and British Columbia were studied by House et al. (1999). The virulence of the isolates was significantly different from each other, but the isolate from Chile was most virulent followed by the isolate from British Columbia and the Norwegian isolate. Piscirickettsiosis is generally considered a disease of salmon in saltwater but Gaggero et al. (1995) indicated that coho salmon and rainbow trout developed the disease in freshwater when 60 to 90 days old. They noted that mortality usually occurred in fish 6–12 weeks after introduction to seawater cages. Also P. salmonis and BKD may occur in the same fish in seawater cultured salmonids in Chile (Smith et al. 1996). It is theorized that Piscirickettsiosis is exacerbated by poor environmental conditions. In view of this, Almendras and Fuentealba (1997) proposed that severity of the diseases

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can be alleviated by decreasing stress during grading and handling, decreasing biomass and stocking density, and removal and proper disposal of dead and moribund-infected fish. It was reported by Mauel et al. (2003) that the piscirickettsiosis-like disease in tilapia could be induced by cold-stress in populations where the pathogen is endemic. Also proper disposal of blood and offal from harvested and dressed fish will eliminate an infection source.

Pathology Hematopoietic tissue becomes necrotic with inflammation and P. salmonis cells are seen in cytoplasmic vacuoles in histological sections or in imprints and smears from the kidney, brain, skeletal muscle, skin, heart, intestine, gills, and ovary (Cvitanich et al. 1991; Lannan and Fryer 1993). Histopathological lesions show necrosis of intravascular thrombi (Branson and Diaz-Munoz 1991). Necrosis was most obvious in spleens, livers, and kidneys of infected Atlantic salmon in Ireland (Rodger and Drinan 1993). The liver may be mottled and occasionally with distinct nodules (Arkush et al. 2005). Tilapia with piscirickettsiosis-like disease had epithelial hyperplasia with severe multifocal consolidation of secondary gill lamellae. There were also multiple granulomas in gills, spleen, kidney, and testes (Maule et al. 2003). Actual pathogenesis of P. salmonis has not been established.

Significance When piscirickettsiosis appeared in Chile in the early 1990s, it killed approximately 1.5 million coho salmon and it has continued to have a major economic impact on that country’s salmon industry. Its presence in geographical areas where trout and salmonids are cultured is of great concern. The detection of RLO’s in other geographical areas and pathogenicity to non-salmonid fish species adds to the concern of aquaculturists.

Other bacterial diseases of salmonids Several bacterial diseases that affect salmonids are more important in other fish groups; therefore, they are discussed in greater depth in other chapters: motile Aeromonas septicemia (A. hydrophila), and columnaris (F. columnare) (Chapter 11), ulcer disease (atypical A. salmonicida) (Chapter 12), Edwardsiellosis (E. tarda) (Chapter 13), and mycobacteriosis (Mycobacterium marinum) (Chapter 15). A disease of trout and Pacific salmon known as pseudokidney disease was recognized in the mid-1970s (Hiu et al. 1984). The causative agent of this disease, Lactobacillus piscicola, was reclassified as Carnobacterium piscicola (Wallbanks et al. 1990). The organism has been reported in North America and Europe (Hiu et al. 1984; Foott 1994). Primary susceptible species are coho and chinook salmon, and rainbow and cutthroat trout, but other marine and freshwater fish may also be susceptible under certain conditions. Fish older than 1 year are most often infected but Michel et al. (1986b) showed that very young salmonids and carp may also be affected. External clinical signs are abdominal distension, erythema at the base of fins, and subdermal blood blisters. Internally the liver, spleen, and kidney may be enlarged and the peritoneal cavity often contains ascitic fluid. Hemorrhages may occur in the testes, intestine, and musculature. Isolation of C. piscicola is by streaking kidney or lesion material on TSA or BHIA and with aerobic incubation at 15◦ C to 24◦ C for 24–72 hours (Foott 1994). Colonies are pinpoint, opaque, entire, circular, and nonpigmented. The bacterium is a nonmotile, nonsporeforming, nonacid fast, Gram-positive rod or coccobacillus measuring 1.1–1.4 × 0.5–0.6 µm. As cultures age, the bacteria become Gram variable. The organism is negative for oxidase, catalase, urease, H2 S, nitrate reduction, and lactose fermentation and positive for arginine dihydrolase and lactic acid production from

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glucose, maltose, mannitol, and sucrose (Hiu et al. 1984). Disease caused by C. piscicola is usually associated with environmental or spawning induced stress. The role of C. piscicola in fish diseases appears to be genuine, although experimental infections are not easily established. An infection of E. tarda (Chapter 13), normally an infection of warm-water fish, was reported in brook trout from two different fish farms in Quebec, Canada (Uhland et al. 2000). Affected fish had nonspecific lesions indicative of a bacterial septicemia that included hemorrhages on the gills and viscera and exophthalmia. The disease was associated with immunosuppression due to increased summer water temperature and lack of precipitation. A common disease of many species of fishes that also infects salmonids is columnaris caused by Flexibacter columnare. While it is more common in warm-water fishes, this pathogen also causes significant problems in salmonids. Juvenile trout/salmon (all species) are more susceptible than older fish, particularly when held in water at temperatures approaching upper levels of tolerance. These infections can be chronic or acute but several reports have revealed over 90% mortality in some populations of juvenile salmonids. High mortality is usually associated with injured and/or if held at 18–20◦ C; however, very low mortality occurs in uninjured fish held under identical conditions. Adult migrating salmon also become more susceptible to columnaris the farther they move upstream toward spawning beds. F. columnare infections in trout and salmon are enhanced by low levels of dissolved oxygen and elevated ammonia especially when combined with elevated water temperature.

Management of salmonid bacterial diseases The philosophical approach to health maintenance and control of salmonid bacterial

diseases does not differ from that of nonsalmonids but more management, chemotherapy, and vaccination options are available. To best control these diseases, a comprehensive program should include good management procedures, proper use of drugs, and vaccination when indicated.

Management Some salmonid bacterial infections caused by obligate pathogens can produce clinical disease without adverse environmental influences, but many epizootics are stress related. In view of this fact, it is important to maintain a high standard of water quality and other associated environmental conditions because overall management practices are critical in controlling, preventing, and reducing effects of disease. To successfully manage most bacterial trout and salmon diseases: (1) maintain water temperature below 17◦ C, (2) employ moderate stocking densities, (3) aerate when needed to provide near 100% oxygen saturation, (4) maintain adequate water flow to remove metabolites, uneaten feed, and solid waste, and (5) incubate eggs, and raise fry and fingerlings in water from a fish-free source whenever possible. Management of facilities where A. salmonicida is enzootic should include stocking fish strains that are more resistant to furunculosis. Destruction of A. salmonicida-infected fish, facility sanitation, and restrict egg sources from certified uninfected broodstock have been successful in maintaining a disease rate of 2% on “health-controlled” farms in Sweden (Wichardt et al. 1989). High sanitation standards should be a top priority in prevention and/or control of bacterial disease. Newly arrived eggs should be disinfected to kill any surface-associated bacteria. In the United States, the most reliable disinfectants for this purpose are buffered PVP iodophores and hydrogen peroxide. Application of sodium hypochlorite at 200 mg/L for up to 24 hours

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effectively killed R. salmonicida cells and probably other pathogens in water (Hirvela-Koski ¨ 2004). Because opinions concerning the etiological agent of cold-water vibriosis differ and because it is a multifactorial disease, its severity is to a large extent dependent on fish culture practices and environmental conditions (Hjeltnes et al. 1987b). H. Mitchell (personal communication) suggested that to minimize effects of cold-water vibriosis, early disease detection is essential, and once confirmed, all handling of fish should be avoided. Moribund and dead fish should be promptly removed and buried, incinerated, or composted. Once surviving fish are removed from cages, nets should be cleaned and sanitized by drying in sunlight and all clothing and/or equipment exposed to diseased fish should be disinfected. Also effluence from slaughter areas should be disinfected to prevent large numbers of bacteria from being reintroduced into culture areas. Enteric redmouth disease control dictates that ponds and raceways are kept clean, culture unit loading limits not be exceeded, the highest quality environmental conditions including water temperature and adequate water flow be maintained, nets and other utensils be sanitized, and facilities be disinfected following fish removal. Also, removal of all possible Y. ruckeri carrier fish or invertebrates from the water supply is essential. Reducing environmental stress, maintaining a high-quality water supply with adequate water flow and using proper sanitation are the best methods of preventing bacterial gill disease (Snieszko 1981). A water supply should originate from a fish-free source, have a highoxygen concentration and low levels of accumulated metabolites such as ammonia, be free of abrasive suspended solids, and be sufficient to accommodate fish density. Feed with excessive “fines” (feed dust) should not be used for BGD susceptible fish and uneaten feed should not be allowed to accumulate in tanks. Increasing water temperatures above the optimum range helps to prevent bacterial cold-

water disease but K. Amos (personal communication) also suggests that it is beneficial to use a substrate when incubating eggs, keep loading densities down for yolk sac fry incubation, minimize yolk-sac activity, and maintain minimum flow rates. Also, coho salmon fry infections were less severe when fish were held in shallow troughs, rather than deep troughs. From these observations, it can be concluded that any procedures that reduce egg, skin, and fin abrasions of juvenile fish are helpful in reducing incidence and severity of F. psychrophilum. Management procedures outlined by Bebak et al. (2007) to reduce the effects of F. psychrophilum will improve survival of affected fish. Water should be free of industrial contaminants such as PCB’s. Ekman et al. (2004) nanoinjected rainbow trout eggs with PCBs, which is present in Puget Sound waters, and then nanoinjected the eggs with F. psychrophilum. A lower resistance to the pathogen occurred in eggs injected with PCB. Other environmental contaminants can also affect susceptibility to other bacterial diseases. Arkush et al. (2001) exposed Chinook salmon to aromatic and chlorinated compounds representative of contaminant in urban estuaries. Based on cumulative percent mortality, salmon exposed to these pollutants were significantly more susceptible to V. anguillarum. Because of its complex epizootiological nature, BKD is one of the most difficult infectious fish diseases to manage. Control requires a variety of integrated management procedures that include reducing the possibility of vertical and horizontal transmission, chemoprophylaxis, chemotherapy when applicable, and environmental and population manipulation (Elliott et al. 1989). Because of improved detection sensitivity of R. salmoninarum in broodstock “BKD free certification” is more feasible. In hatcheryreared salmonids, it is important to reduce potential vertical transmission during artificial spawning activities by culling and the segregation of brood fish and eggs. Gametes from

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brood fish that are most likely to transmit R. salmoninarum to offspring should be destroyed. Each brood fish should be tested for R. salmoninarum at spawning and their gametes held separately under refrigeration until BKD status of each adult is known. Gametes from BKD-positive fish can then be destroyed while those of BKD-negative fish may be fertilized. Pascho et al. (1991) suggested that using ELISA and FAT to test adults for R. salmoninarum in conjunction with segregation of eggs significantly reduces disease prevalence in chinook. Also BKD testing can be done when eggs are taken to get an accurate measure of carrier status in brood fish. The National Workshop on Bacterial Kidney Disease addressed BKD management and recommended the following for implementation by the U.S. Fish and Wildlife Service (Mangin 1991): 1. There is no reason to destroy stocks of fish that are infected with R. salmoninarum. 2. Approved R. salmoninarum detection methods in fish include ELISA, FAT, and FAT with membrane filtration. 3. If R. salmoninarum is present at any brood stock facility, stocks should be managed to reduce infection severity and BKD suspect fish should be stocked only in areas where the pathogen is already known to exist. 4. Eggs should not be taken from clinically diseased fish unless a specific program (i.e., endangered species) requires use of such eggs. 5. Asymptomatically infected fish should not be stocked in areas that are free of R. salmoninarum. 6. In the United States, no matter the locale, all BKD problems should be treated the same. According to Almendras and Fuentealba (1997), effects of piscirickettsiosis can be reduced by decreasing stress during grading and handling, decreasing fish biomass and stocking density, and removal and proper disposal of dead and moribund-infected fish. Also proper disposal of blood and offal from harvested and dressed fish will eliminate an infection source.

Establishing sanitary management on salmonid farms was shown to be effective in reducing severe effects of diseases (Bebak et al. 2007). When F. psychrophilum was thought to have been introduced on water testing equipment of a fish farm, initiating sanitary practices; removal of dead fish twice a day, separation of equipment used in each tank, cleaning with chlorine and rinsing, not using same equipment in other tanks, and cleaning pipes and tanks prevented the disease from reaching the pathogens devastating potential. Another management tool recently considered for salmonid bacterial diseases is the implementation of probiotics in the diet (Irianto and Austin 2002). This involves incorporation of certain bacteria or algae into the diet that reduces the fish’s susceptibility to pathogenic organisms. Although the positive process involved is not known, they suggest that the increased resistance may be due to competition for nutrients and/or space, alter microbial metabolism, or stimulation of host immunity. The bacteria Bacillus sp. (JB1) and A. sobria (GC2) recovered from digestive tract of rainbow trout and carp, respectively, were cultured and incorporated into the feed of juvenile rainbow trout (Brunt et al. 2007). Fish fed the probiotics experienced reduced susceptibility to A. salmonicida, Lactococcus garvieae, Streptococcus iniae, V. anguillarum, V. ordalii, and Y. ruckeri. Control fish receiving no probiotic bacteria in the diet suffered 80–100% mortality when challenged with the pathogens compared to 0–13% mortality for those fish that received the probiotics.

Chemotherapy Chemotherapy of bacterial diseases in salmonids begins with treating incubating eggs where bacteria and fungi (Saprolegnia sp.) can be devastating. Eyed eggs can be immersed in 25–200 mg of iodine/L (Betadine) for at least 10 minutes using well-oxygenated chlorinefree water, rinsed immediately, and put into

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flowing water incubators. Perox-Aid (35% hydrogen peroxide) is FDA approved for prophylactic treatment on all freshwater finfish eggs for fungus and bacterial infections. PeroxAid is applied at a continuous flow rate of 500–1,000 mg/L (ppm) for 15 minutes on alternate days for three applications. In the United States, chemotherapy for furunculosis, vibriosis, enteric redmouth, coldwater vibriosis, and other bacterial salmonid diseases is limited to feeding Terramycin, Romet-30, and Florfenicol, which is now available; however, several other drugs are being used under INAD exemption (Table 4.2). For furunculosis, vibriosis and ERM Terramycin is fed at a rate of 50–75 mg/kg of fish per day for 10 days followed by a 21 day withdrawal period before harvest and marketing. Romet30 is fed at 50 mg/kg of fish per day for 5 days with a 42 day withdrawal period. Florfenicol is approved in Canada and by the FDA for furunculosis fed at a rate of 10 mg/kg of fish per day for 10 days followed by a 12 day withdrawal. Before chemotherapy is applied, it is recommended that antibiotic sensitivity for a particular isolate of any bacterial pathogen to determine antibiotic resistance. Antibiotics should never be used indiscriminately, for long periods of time, or at less than recommended concentrations because these practices lead to increased drug resistance. Additional drugs to treat salmonid bacterial infections are approved in other countries but can only be used experimentally in the United States. Oxolinic acid is used extensively in Europe at 10 mg/kg body weight per day. Enrofloxacin was shown by Bowser et al. (1994) to significantly (p < 0.05) increase survival of experimentally A. salmonicida-infected rainbow trout when fed at rates of 1.25–2.5 mg/kg of fish daily for 10 days. During reduced feeding activity in winter, enrofloxacin fed at 10 mg/kg per day to lake trout and 5 mg/kg per day to Atlantic salmon for 20 days significantly reduced mortality due to furunculosis (Hsu et al. 1994). A second fluoroquinolone, difloxacin, was evaluated for controlling A.

salmonicida in Atlantic salmon by Elston et al. (1995). Survival was significantly increased by feeding difloxacin at doses of 2.5 to 5.0 mg/kg of fish daily for 5 days. No increased benefit was achieved by increasing medication for 10 days. Amoxycillin has been seriously considered for fish. Barnes et al. (1994) found that all atypical A. salmonicida tested in Scotland were sensitive to minimum inhibitory concentrations of 0.3–1.5 mg of amoxycillin per liter of water. Cipriano et al. (1992) showed that A. salmonicida resides in the mucus of carrier trout. The monitoring of an Atlantic salmon population for A. salmonicida via mucus analysis and subsequent oral treatment with oxytetracycline (77 mg/kg of fish for 10 days) and topical treatment with Chloramine-T (INAD exemption) (15 mg/L) in a 60 minute bath was described by Cipriano et al. (1996b). These treatments reduced the presence of A. salmonicida in the mucus by 90–100%, allowing affected fish to be stocked in accordance with established fish health regulations for salmonids in New England. Cipriano et al. (1996c) also used topical disinfection of Chloramine-T to control A. salmonicida on salmonids. Terramycin is generally used to treat coldwater vibriosis. However, Poppe et al. (1986) used Terramycin, Tribrissen, or furazolidone (the latter two are not approved in the United States) in feed at 75–100 mg/kg of body weight per day for 10 days and found that furazolidone was most effective against CV. However, Bruno et al. (1986) held cumulative mortality to 3% with Terramycin. Mortality due to V. salmonicida in Atlantic salmon can be reduced in 3–4 days by feeding oxolinic acid at 75–90 mg/kg of body weight per day for 10 days. Because a variety of antibiotics have been used to control CV, multiple antibiotic resistance of V. salmonicida to oxytetracycline, sulphonamides, and oxolinic acid have emerged (Hjeltnes et al. 1987b). BGD (F. branchiophilum) therapy is effective when applied expediently as infected fish tend to recover rapidly after gill bacteria are

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reduced. Perox-Aid is approved for treating this disease at a concentration of 100 mg/L (ppm) hydrogen peroxide as continuous flow or static treatment for 30- or 50–100 mg/L for 60 minutes once per day on alternate days for three applications. However Rach et al. (2000) showed that BGD in brown and rainbow trout and chinook salmon can be successfully treated with 230 mg/L for 60 minutes every other day. Other treatments that have been used for BGD are disinfectants containing quaternary ammonia, such as benzalkonium chlorides (Hyamine 1622, Hyamine 3500, or Roccal), that can be added to water as a prolonged treatment. A 1–2 mg/L application of active benzalkonium chloride for 1 hour on 3 consecutive days is recommended; however, caution must be taken in soft water in which the compound can be toxic to some trout. The herbicide Diquat has also been used successfully in treating BGD at 2–4 mg/L of active ingredient. Potassium permanganate can also be used at 2 mg/L for 1 hour, but the safety margin is very narrow. Chloramine-T (INAD exemption) applied at 8–10 mg/L for 1 hour for 3 days is the most effective treatment for BGD; however, 15–20 mg/L are required in stubborn cases in cold water. Byrne et al. (1991) suggested that salt concentrations in water should be increased during BGD outbreaks or during chemotherapy treatment to compensate for loss of fish electrolytes. Other than hydrogen peroxide, none of the aforementioned compounds are FDA approved for therapeutic use against BGD if fish are destined for United States markets but some are used under INAD exemption. Serious outbreaks of BCWD are generally difficult to treat because fish with systemic infection feed poorly, thus neutralizing any advantage of medicated feed (Holt 1993). Bullock and Snieszko (1970) and Schachte (1983) recommended bath treatments with watersoluble oxytetracycline at 10 to 50 mg/L or quaternary ammonium compounds at 2 mg/L while infections are confined to the skin. After disease becomes systemic oxytetracycline

in the diet at 50–75 mg/L of fish per day for 10 days is effective if fish are feeding. In Great Britain Amoxycillin is incorporated at 50 µg/kg of feed and fed for 10 days. Oxytetracycline is injected into adult fish at 50 mg/kg prior to spawning to prevent vertical transmission. These drugs are not FDA approved. Chemoprophylaxis is used to reduce vertical transmission of R. salmoninarum but none are FDA approved for food fish. Adult female salmon are injected once or twice with 10–20 mg of erythromycin phosphate per kg of body weight at 3–4 week intervals prior to spawning with a 10 day window between the last injections and spawning (Amos 1977). Erythromycin has also been widely used as an oral treatment at 4.5 g/45 kg (about 100 mg/kg) of body weight per day for 10–21 days for clinical BKD (Elliott et al. 1989). Moffitt (1992) found that erythromycin fed at 200 mg/kg of body weight per day for 21 days reduced mortality more than when lower dosages were fed for shorter periods. Subsequently, Peters and Moffitt (1996) showed that an optimum erythromycin dosage for BKD infected coho salmon was 100 mg/kg daily for 28 days. They noted that a higher dose was unpalatable, particularly late in the treatment regime or when water temperature was >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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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.

References Aguayo, J., A. Miquel, et al. 2002. Detection of Piscirickettsia salmonis in fish tissues by an enzymelinked immunosorbent assay using specific monoclonal antibodies. Diseases of Aquatic Organisms 49:33–38. Alcorn, S., A. L. Murray, et al. 2005. A cohabitation challenge to compare the efficacies of vaccines for bacterial kidney disease (BKD) in chinook salmon Oncorhynchus tshawytascha. Diseases of Aquatic Organisms 63:151–160. Allen-Austin, D., B. Austin, and R. R. Colwell. 1984. Survival of Aeromonas salmonicida in river water. FEMS Microbiological Leters 21:143–146. Allison, L. N. 1958. Multiple sulfa therapy of kidney disease among brook trout. The Progressive Fish Culturist 20:66–68. Almendras, F. E., and I. C. Fuentealba. 1997. Salmonid rickettsial septicemia caused by Piscirickettsia salmonis: a review. Diseases of Aquatic Organisms 29:137–144. Altinok, I. and J. M. Grizzle. 2001. Effects of salinity on Yersinia ruckeri infection of rainbow trout and brown trout. Journal of Aquatic Animal Health 13:334–339. Altinok, I, J. M. Grizzle, and L. Zhanjiang. 2001. Detection of Yersinia ruckeri in rainbow trout blood by use of the polymerase chain reaction. Diseases of Aquatic Organisms 44:29–34. Amend, D. F., and K. A. Johnson. 1981. Current status and future needs of Vibrio anguillarum bacterins. Developments in Biological Standardization 49:403–417.

Amos, K. 1977. The control of bacterial kidney disease in spring chinook salmon. M. S. Thesis, University of Idaho, Moscow. Anderson, J. I., and D. A. Conroy. 1970. Vibrio disease in marine fishes. In: S. F. Snieszko (ed.) Symposium on Diseases of Fishes and Shellfishes. 266–272. Special publication No. 5. Bethesda, MD: American Fisheries Society. Anonymous. 1994. Guide to drug, vaccine, and pesticide use in aquaculture. Prepared by the Federal Joint Subcommittee on Aquaculture, U.S. Department of Aqriculture. Anonymous. 2004. Diseases caused by Vibrio species. Aquafarma. http://www.holar.is/ aquafarmer/node144.html. Aoki, T., and F. I. Holland. 1985. The outer membrane proteins of the fish pathogens Aeromonas hydrophila, Aeromonas almonicida and Edwardsiella tarda. FEMS Microbiology Letters 27:299–305. Arakawa, C. K., J. E. Sanders, and J. L. Fryer. 1987. Production of monoclonal antibodies against Renibacterium salmoninarum. Journal of Fish Diseases 10:249–253. Argenton, F., S. DeMas, et al. 1996. Use of random DNA amplification to generate specific molecular probes for hybridization tests and PCRbased diagnosis of Yersinia ruckeri. Diseases of Aquatic Organisms 24:121–127. Arias, R., O. Olivares-Fuster, et al. 2007. First report of Yersinia ruckeri biotype 2 in the USA. Journal of Aquatic Animal Health 19:35–40. Arkush, K. D., A. M. McBride, et al. 2005. Genetic characterization and experimental pathogenesis of Piscirickettsia salmonis isolated from white seabass Astractoscion nobilis. Diseases of Aquatic Organisms 63:239–249. Austin, B. 1985. Evaluation of antimicrobial compounds for the control of bacterial kidney disease in rainbow trout, Salmo qairdneri Richardson. Journal of Fish Diseases 8:209–220. Austin, B. 1992. The recovery of Cytophaga psychrophila from two cases of rainbow trout (Oncorhynchus mykiss Waulbaum) fry syndrome in the UK. Bulletin of the European Association of Fish Pathologist 12:207–208. Austin, B., and D. A. Austin. 1987. Bacterial Fish Pathogens: Diseases in Farmed and Wild Fish. Chichester, Ellis Horwood, 364pp. Austin, B., and J. Rayment. 1985. Epizootiology of Renibacterium salmoninarum, the causal agent

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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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Striped Bass Bacterial Diseases

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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Contorer, P., and R. Jones. 1979. Minocycline therapy of aquarium granuloma. Case reports and literature review. Cutis 23(6):864–868. Decostere, A., K. Hermans, and F. Haesebrouck. 2004. Piscine mycobacteriosis: a literature review covering the agent and the disease it causes in fish and humans. Veterinary Microbiology 99:159–166. Diamant, A., A. Banet, et al. 2000. Mycobacteriosis in wild rabbitfish Siganus rivulatus associated with cage farming in the Gulf of Eliat, Red Sea. Diseases of Aquatic Organisms 39:211–219. dos Santos, N. M. S., A. do Vale, et al. 2002. Mycobacterial infections in farmed turbot Scophthalmus maximus. Diseases of Aquatic Organisms 52:87–91. Elkamel, A. A. 2002. Pathogenic mechanism of Photbacterium damselae subspecies piscidida in hybrid striped bass. Ph.D. Dissertation, Louisiana State University, Baton Rouge, Louisiana. Elkamel, A. A., and R. L. Thune. 2003. Invasion and replication of Photobacterium damselae subsp. piscicida in fish cell lines. Journal of Aquatic Animal Health 15:167–174. Elkamel, A. A., J. P. Hawke, et al. 2003. Photobacterium damsela subsp. piscicida is capable of replicating in hybrid striped bass macrophages. Journal of Aquatic Animal Health 15:175– 183. Egusa, S. 1983. Disease problems in Japanese yellowtail Seriola guinguiradiata culture: a review. In: J. E. Stewart (ed.) Diseases of Commercially Important Marine Fish and Shellfish. Copenhagen, l’Exploration de la Mer, pp. 10–18. Frerichs, G. N. 1993. Mycobacteriosis: Nocardiosis. In: V. Inglis, R. J. Roberts, and N. R. Bromage (eds) Bacterial Diseases of Fish. Oxford, Blackwell Scientific Press, pp. 219–233. Frerichs, G. N., and R. J. Roberts. 1989. The bacteriology of teleosts. In: R. J. Roberts (ed.) Fish Pathology, Second edition. London, BailliereTindall, pp. 289–319. Fryer, J. L., and J. E. Sanders. 1981. Bacterial kidney disease in salmonid fish. Annual Reviews in Microbiology 35:273–298. Fukuda, Y., and R. Kusuda. 1981. Efficacy of vaccination for pseudotuberculosis in cultured yellowtail by various of administration. Bulletin of the Japanese Society of Scientific Fisheries 47:147–150.

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Nedoluha, P. C., and D. Westhoff. 1997. Microbiology of striped bass grown in three aquaculture systems. Food Microbiology 14:255–264. Nigrelli, R. F., and H. Vogel. 1963. Spontaneous tuberculosis in fishes and in other cold-blooded vertebrates with special reference to Mycobacterium fortuitum Cruz from fish and human lesions. Zoologica 48:130–143. Nitzan, S., B. Swartsburd, and E. D. Heller. 2003. The effect of growth medium salinity of Photobacterium damselae subsp. piscicida on the immune response of hybrid bass (Morone saxatilis X M. chrysops. Fish and Shellfish Immunology 16:107–116. Noya, M., B. Magarinos, et al. 1995. Sequential ˜ pathology of experimental pasteurellosis in gilthead seabream Sparus aurata. A light-and electron microscopic study. Diseases of Aquatic Organisms 21:177–186. Ostland, V. E., J. A. Stannard, et al. 2006. Aquatic Francisella-like bacterium associated with mortality of intensively cultured hybrid striped bass Morone chrysops × M. saxatilis. Diseases of Aquatic Organisms 72:135–145. Overton, A. S., M. J. Margraf, et al. 2003. The prevalence of mycobacterial infections in striped bass in Chesapeake Bay. Fisheries Management and Ecology 10(5):301–308. Pedersen, K. H., F. Skall, et al. 2009. Photobacterium damselae subsp. damselae, an emerging pathogen in Danish rainbow trout, Oncorhynchus mykiss (Walmaum), mariculture. Journal of Fish Diseases 32:465–472. Plumb, J. A. 1997. Infectious Diseases of Striped Bass. In: R. M. Harrell (ed.) Striped Bass and Other Morone Culture, edited by Developments in Aquaculture and Fisheries Science. Amsterdam, Elsevier, 30:271–313. Post, G. 1987. Textbook of Fish Diseases. Neptune, New Jersey, T. F. H. Publication. Rajan, P. R., J. H. Lin, et al. 2003. Simple and rapid detection of Photobacterium damselae ssp. piscicida by PCR technique and plating method. Journal of Applied Microbiology 95:1375–1380. Ranger, B.S., E. Mahrous, et al. 2006. Globally distributed mycobacterial fish pathogens produce a novel plasmid-encoded toxic macrolide, Mycolactone F. Infection and Immunity 74:6037–6045. Rhodes, M. W., K. H. Kator, et al. 2003. Mycobacterium shottsii sp., a slow growing species

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Van Duijn, C. 1981. Tuberculosis in fishes. Journal of Small Animal Practices 22:391–411. Wang, G-L., S-P. Yuan, and S. Jin. 2005. Nocardiosis in large yellow croaker, Larimichthys crocea (Richardson). Journal of Fish Diseases 28:339–345. Wayne, L. G., and S. Kubica. 1986. Genus Mycobacterium. In: P. H. Sneath, N. S. Mair, M. E. Sharpe, and J. G. Holt (eds) Bergey’s Manual of Systematic Bacteriology. Baltimore, Williams and Wilkins, 2:1436–1457. Wheeler, A. P., and B. S. Graham. 1989. Saturday conference: Atypical mycobacterial infections. Southern Medical Journal 82:1250. Wolf, J. C., and S. A. Smith. 1999. Comparative severity of experimentally induced mycobacteriosis in striped bass Morone saxatilis and hybrid tilapia Oreochromis spp. Diseases of Aquatic Organisms 38:191–200. Wolke, R. E. 1975. Pathology of bacterial and fungal diseases affecting fish. In: R. Ribelin and G. Migaki (eds) The Pathology of Fishes. Madison, Wisconsin, University of Wisconsin Press, pp. 33–116. Xu, D., and W. A. Rogers. 1993. Oxytetracycline residue in hybrid striped bass muscle. Journal of the World Aquaculture Society 24:466–472. Zappulli, V., T. Patarnello, et al. 2005. Direct identification of Photobacterium damselae subspecies piscicida by PCR RFLP analysis. Diseases of Aquatic Organisms 65:53–61. Zorrilla, I, M. C. Balemona, et al. 1999. Isolation and characterization of the causative agent of pasteurellosis, Photobacterium damsela ssp. piscicida, from sole, Solea senegalensis. Journal of Fish Diseases 22:167–172.

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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.

References Abutbul, S., A. Golan-Goldhirsh, et al. 2004. Use of Rosmarinus officinalis as a treatment against Streptococcus iniae in tilapia (Oreochromis sp.). Aquaculture 238:97–105. Amin, N. E., I. S. Abdallah, et al. 1988. Columnaris infection among cultured Nile tilapia Oreochromis niloticus. Antonie van Leeuwenhoek 54:509–520.

Anonymous. 1996. Invasive infection with Streptococcus iniae in Ontario, 1995–1996. Morbidity and Mortality Weekly Report, Center for Disease Control, Atlanta, Georgia. 45, No. 30, 650pp. Anonymous. 2006. Streptococcus in tilapia. Intervet, Shefield, S35 9ZX, England. http://www. thefishsite.com. Badran, A. F. 1993. An outbreak of edwardsiellosis among Nile tilapia (Oreochromis niloticus) reared in ponds supplied with domestic wastewater. Zagazig Veterinary Journal 21:771–777. Barnes, A. C. Fiona, M. Young, et al. 2003. Streptococcus iniae: serological differences, presence of capsule and resistance to immune serum killing. Diseases of Aquatic Organisms 53:241–247. Bowser, P. R., G. A. Wooster, et al. 1998. Streptococcus iniae infection of tilapia Oreochromis niloticus in a recirculation production facility. Journal of the World Aquaculture Society 29:335–339. Chang, P. H., and J. A. Plumb. 1996. Effects of salinity on Streptococcus infection of Nile tilapia, Oreochromis niloticus. Journal of Applied Aquaculture 6:39–45. Chen, C.-Y, C.-B. Chao, and P. R. Bowser. 2007. Comparative histopathology of Streptococcus iniae and Streptococcus agalactiaeinfected tilapia. Bulletin of the European Association of Fish Pathologists 27(1):2–8. Chen, S. C., m.-C. Tung, et al. 1994. Systematic granulomas caused by a rickettsia-like organism in Nile tilapia, Oreochromis niloticus (L.), from southern Taiwan. Journal of Fish Diseases 17:591–599. Chen, S.-C., P.-C. Wang, et al. 2000. A pisciricketsia-like organism in grouper, Epinephelus melanoshoma, in Taiwan. Journal of Fish Diseases 23:415–418. Colorni, A., A. Diamant, et al. 2002. Streptococcus iniae infections in Red Sea cage-cultured and wild fishes. Diseases of Aquatic Organisms 49:165–170. Darwish, A. M., and M. S. Hobbs. 2005. Laboratory efficacy of amoxicillin for the control of Streptococcus iniae infection in blue tilapia. Journal of Aquatic Animal Health 17:197–202. Darwish, A. M., S. D. Rawles, and B. R. Griffin. 2002. Laboratory efficacy of oxytetracycline for the control of Streptococcus iniae infection in blue tilapia. Journal of Aquatic Animal Health 14:184–190.

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Dodson, S. V., J. J. Maurer, and E. B. Shotts. 1999. Biochemical and molecular typing of Streptococcus iniae isolated from fish and human cases. Journal of Fish Diseases 22:331–336. Duremdez, R. C., and G. D. Lio-Po. 1985. Studies on the causative organism of Sarotheradon niloticus (Linnaeus) fry mortalities-2: Isolation, identification and characterization of Pseudomonas fluorescence. Fish Pathology 20:115–123. Eldar, A., Y. Bejerano, and H. Bercovier. 1994. Streptococcus shiloi and Streptococcus dificile: two new streptococcal species causing a meningoencephalitis in fish. Current Microbiology 28:139–143. Eldar, A., P. Frelier, et al. 1995. Streptococcus shiloi, the name for an agent causing septicemic infection in fish is a junior synonym of Streptococcus iniae. International Journal of Systematic Bacteriology 45:840–842. Evans, J. J., P. H. Klesius, et al. 2002. Characterization of β-haemolytic Group B Streptococcus agalactiae in cultured seabream, Sparus auratus L., and wild mullet, Liza klunzingeri (Day), in Kuwait. Journal of fish Diseases 25:505–513. Evans, J. J., P. H. Klesius, et al. 2004a. Streptococcus agalactiae vaccination and infection stress in Nile tilapia, Oreochromis niloticus. Journal of Applied Aquaculture 16(3/4):105–115. Evans, J. J., P. H. Klesius, and C. A. Shoemaker. 2004b. Efficacy of Streptococcus agalactiae (group B) vaccine in tilapia (Oreochromis niloticus) by intraperitoneal and bath immersion administration. Vaccine 22:3769–3773. Evans, J. J., P.H. Klesius, and C.A. Shoemaker. 2009. First isolation and characterization of Lactococcus garvieae from Brazilian Nile tilapia, Oreochromis niloticus (L.), and pintado, Pseudoplathystoma corruscans (Spix & Agassiz). Journal of Fish Diseases 32:943–951. Evans, J. J., C. A. Shoemaker, et al. 2001. Distribution of Streptococcus iniae in hybrid striped bass (Morone chrysops X Morone saxatilis) following nare inoculation. Aquaculture 194:233–243. Faisal, M., and W. Popp. 1987. Plesiomonas shigelloides: a pathogen for the Nile tilapia, Oreochromis niloticus. Journal of the Egyptian Veterinary Medical Association 47:63–70. Ferguson, H. W., J. A. Morales, and V. E. Ostland. 1994. Streptococcosis in aquarium fish. Diseases of Aquatic Organisms 19:1–6.

Fukuda, Y., A. Okamura, et al. 2002. Granulomatosis of cultured three-line grunt Parapristipoma trilineatum caused by and intracellular bacterium. Fish Pathology 37:119–124. Gaunt, P. 2004. Efficacy of Florfenicol for control of mortality caused by Streptococcus iniae in tilapia. Proceedings of the World Aquaculture Society, Honolulu, HI, 216pp. Hawke, J. P. 2000. Bacterial Disease Agents. In: R.R. Stickney (ed.) Encyclopedia of Aquaculture. New York, USA, John Wiley and Sons, pp. 69–97. Hernandez, E., J. Fiqueroa, and C. Irequi. 2009. ¨ Streptococcosis on a red tilapia, Oreochromis sp., farm: a case study. Journal of Fish Diseases 32:247–252. Hoshina, T., T. Sano, and Y. Morimoto. 1958. A streptococcus pathogenic to fish. Journal of Tokyo University Fisheries 44:57–68. Hsieh, C. Y., M. C. Tung, et al. 2006. Enzootics of visceral granulomas associated with Francisellalike organism infection in tilapia (Oreochromis spp.). Aquaculture 254:129–138. Hubert, R. M. 1989. Bacterial diseases in warmwater aquaculture. In: M. Shilo and S. Sarig (eds) Fish Culture in Warm Water Systems: Problems and Trends. Boca Raton, Fl: CRC Press Inc., pp. 194–197. Khalil, R. H., and S. M. Aly. 2008. Photobacteriosis in some wild and cultured freshwater fishes in Egypt. Eighth International Symposium on Tilapia in Aquaculture. Cairo, Egypt, pp. 1211–1228. Kimura, H., and R. Kusuda. 1979. Studies on the pathogenesis of streptococcal infection in culture yellowtails Seriola spp.: effect of the cell free culture on experimental streptococcal infection. Journal of Fish Diseases 2:501–510. Kitao, T. 1982. The method for detection of Streptococcus sp. causative bacteria of streptococcal disease of cultured yellowtail (Seriola guingueradiata): especially their cultural, biochemical and serological properties. Fish Pathology 17:17–26. Kitao, T. 1993. Streptococcal infections. In: V. Inglis, R. J. Roberts, and N. R. Bromage (eds) Bacterial Diseases of Fish. London, Blackwell Press, pp. 196–210. Kitao, T., T. Aoki, and K. Iwata. 1979. Epidemiological study on streptococcosis of cultured

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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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470 Bacterial Diseases

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.)

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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.)

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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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482 Appendices

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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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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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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Index 489

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