SELECTED PATHOGENS OF ORNAMENTAL FISHES

By

PREEYANAN SRIWANAYOS

A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE

UNIVERSITY OF FLORIDA

2012

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© 2012 Preeyanan Sriwanayos

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To my parents for all their love and support throughout my life

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ACKNOWLEDGMENTS

I thank my advisor, Dr. Ruth Francis-Floyd, for her guidance and support during my 3 years of study. I also thank my co-advisor, Dr. Denise Petty, for her tutelage and help throughout my study. Additionally, I gratefully acknowledge the enthusiastic supervision of the other members of my supervisory committee throughout this work:

Drs. Thomas Waltzek, Andrew Kane, and Jeffrey Hill.

I am grateful to the Royal Thai Government for financial support throughout my study. I thank Segrest Farms, Gibsonton, Florida and Sun Pet Ltd., Atlanta, Georgia for their generous contributions.

I thank Drs. Jim Wellehan, Galaxia Cortes-Hinojosa, and Ms. Linda Archer of the

Aquatic Pathobiology Laboratory, University of Florida for their assistance and use of their laboratories. I am appreciative to Dr. Mark Stidworthy of International Zoo

Veterinary Group, UK for his pathological expertise. I am in debt to Ms. Heather

Maness, Dr. Natalie Steckler, and Dr. Claire Erlacher-Reid for their help with data collection. I thank Dr. Charles Cichra of the University of Florida, Department of

Fisheries and Aquatic Sciences and Mr. James Colee of the University of Florida,

Institute of Food and Agricultural Sciences (IFAS) Statistics Department for their statistical guidance. I also thank Ms. Karen Kelly for her mentoring and guidance in the area of electron microscopy.

I am appreciative of my best friends, Atthapol Charoenkietkrai, Pasicha

Chaikaew, Suwussa Bamrungsab, and Akeapot Srifa, for their encouragement and friendship provided throughout my study aboard time and especially during this project.

And finally to my parents, thank you for your support and love over the many years.

Both of them helped me became the woman I am today.

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TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 7

LIST OF FIGURES ...... 8

ABSTRACT ...... 10

CHAPTER

1 INTRODUCTION ...... 12

Ornamental Fish Trade ...... 12 Diseases of Ornamental Fishes ...... 13 Tetrahymena Infection in Imported Guppies ...... 16 Guppies ...... 16 Tetrahymena ...... 19 Megalocytivirus Infection in Fishes, with Emphasis on the Orbiculate Batfish (Platax orbicularis) ...... 23 Orbiculate Batfish ...... 23 Megalocytivirus ...... 24

2 PRELIMINARY ASSESSMENT OF PARASITE LOAD IN SELECTED IMPORTED GUPPIES (POECILIA RETICULATA) ...... 37

Materials and Methods...... 39 Imported Guppy Population ...... 39 Gross Examination ...... 40 Collection of Tissues for Evaluation of Microscopically Visible Parasites ...... 40 Parasite Identification Protocols ...... 41 Data Analyses ...... 41 Results ...... 42 Discussion ...... 43

3 PRELIMINARY STUDY OF TRANSMISSION OF TETRAHYMENA FROM INFECTED TO TETRAHYMENA-FREE GUPPIES (POECILIA RETICULATA) ..... 51

Materials and Methods...... 52 Experimental Animals ...... 52 Tetrahymena-free fish ...... 52 Tetrahymena-infected fish ...... 54 Experimental Infection ...... 55 Histological Examination ...... 57 Statistical Analysis ...... 57

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Results ...... 57 Experimental Infection ...... 57 Histological Examination ...... 59 Discussion ...... 59

4 MEGALOCYTIVIRUS INFECTION IN ORBICULATE BATFISH (PLATAX ORBICULARIS) ...... 72

Materials and Methods...... 75 Clinical History ...... 75 Histopathology ...... 76 Transmission Electron Microscopy ...... 76 DNA Extraction, Polymerase Chain Reaction Amplification, and Sequencing.. 77 BLASTN, Molecular Dataset, Sequence Alignment, and Phylogenetic Analysis ...... 77 Results ...... 78 Histopathology ...... 78 Transmission Electron Microscopy ...... 79 Sequencing, BLASTP, Molecular Dataset, and Phylogenetic Analysis ...... 79 Discussion ...... 80

LIST OF REFERENCES ...... 89

BIOGRAPHICAL SKETCH ...... 101

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LIST OF TABLES

Table page

1-1 Reports of viral diseases in ornamental fishes ...... 28

1-2 Comparison of known megalocytiviruses infecting fishes...... 29

2-1 Microscopically visible parasite were found on external surfaces and within the gastrointestinal tract of guppies imported to the U.S. from Singapore ...... 47

2-2 Determining the number of fish that need to be sampled in order to assess pathogen prevalence in a lot (or population) of fish ...... 48

3-1 Classification of parasite intensity per field of view (FOV) at predetermined magnifications ...... 63

3-2 Mortality rates, infection rates, and biopsy findings of 42 exposed Tetrahymena-free female guppies from each treatment during the 14-day experimental period...... 64

3-3 Mortality rates, infection rates, and biopsy findings of infected male guppies during the 14-day experimental period...... 65

4-1 Primers used to amplify fragments of the major capsid protein (MCP) gene sequence from the orbiculate batfish iridovirus (OBIV) ...... 82

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LIST OF FIGURES

Figure page

1-1 An example of a fancy guppy (Poecilia reticulata) ...... 30

1-2 A photomicrograph of Tetrahymena sp. on the skin mucus of a guppy ...... 31

1-3 A diagram shows the standard shapes of guppy caudal fin ...... 32

1-4 Differentiation of male and female guppies ...... 33

1-5 Orbiculate batfish (Platax orbicularis) ...... 34

1-6 Distribution of orbiculate batfish ...... 35

1-7 Orbiculate batfish iridovirus particles ...... 36

2-1 Camallanus spp. recovered from the gastrointestinal tract of a guppy imported from Singapore ...... 49

2-2 Gyrodactylus spp. observed on the fin of a guppy ...... 50

3-1 Holding tanks ...... 66

3-2 External lesions observed in guppies infected with Tetrahymena ...... 67

3-3 Mortality rate (%) of exposed Tetrahymena-free guppies during the 14-day experimental period ...... 68

3-4 Cumulative mortality (%) of guppies used in the experiment during the 14- day experimental period ...... 69

3-5 Tetrahymena infection rate (%) in exposed Tetrahymena-free guppies during the 14-day experimental period ...... 70

3-6 Guppies infected with Tetrahymena ...... 71

4-1 Kidney from infected orbiculate batfish with intracytoplasmic basophilic iridovirus inclusions in the interstitium and glomerul ...... 83

4-2 Heart from infected orbiculate batfish with abundant intracytoplasmic basophilic iridovirus inclusions ...... 84

4-3 Intracytoplasmic basophilic inclusions in infected orbiculate batfish ...... 85

4-4 Transmission electron photomicrographs of iridoviral infected cells in the heart of an orbiculate batfish ...... 86

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4-5 Phylogenetic analysis of the megalocytivirus MCP ...... 87

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Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science

SELECTED PATHOGENS OF ORNAMENTAL FISHES By

Preeyanan Sriwanayos

December 2012

Chair: Ruth Francis-Floyd Cochair: B. Denise Petty Major: Fisheries and Aquatic Sciences

Infectious diseases impact fish production worldwide, resulting in economic loss from mortality and decreased value from poor appearance. To improve understanding of selected pathogens, I attempted to assess parasitism in recently imported guppies

(Poecilia reticulata), develop a transmission model for an important parasite, and characterize an emerging viral disease.

Guppies are a mainstay of the ornamental fish trade and are produced in large numbers in the U.S. and Thailand. To assess parasitism in this representative species, a preliminary health assessment was conducted to evaluate parasitic infection on a

shipment of guppies recently imported to the U.S. from Singapore. Two parasite genera

(Camallanus and Gyrodactylus) were observed on guppies in this shipment.

Although Tetrahymena was not found in the preliminary health assessment, it is

an important parasitic disease of freshwater fishes, particularly members of the family

Poeciliidae, and can result in high mortality. A pilot study was conducted to assess

Tetrahymena transmission between guppies, by cohabitation, under conditions that

were similar to those present in Florida wholesale facilities. Although the parasite was

successfully transmitted by cohabitation of Tetrahymena-free fish with Tetrahymena-

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infected fish, the parasite transmission was limited and the process is still poorly understood.

Megalocytiviruses are important emerging pathogens of ornamental and food fishes and are poorly understood. I was able to characterize a megalocytivirus newly discovered in orbiculate batfish (Platax orbicularis). Based on histopathology, electron

microscopy, and phylogenetic findings, I confirmed that this virus is identical to a megalocytivirus previously reported in Banggai cardinalfish (Pterapogon kauderni).

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CHAPTER 1 INTRODUCTION

Ornamental Fish Trade

The ornamental fish industry is an important sector of the world’s aquaculture

industry. More than 1 billion ornamental fishes, including 4000 freshwater and 1400 marine species, are traded globally each year (Whittington and Chong 2007). According

to the Food and Agriculture Organization of the United Nations (FAO), annual

international export of ornamental fishes was approximately US$ 251 million in 2004

and it has been increasing at an average growth rate of 14 percent per year since 1985

(FAO 2012). Total value of the ornamental fish industry worldwide, which includes non-

exported products, aquarium-associated materials, retail sales, and labor costs, is

estimated to be as much as US$15 billion annually (Bartley 2004). The global value of

the wholesale ornamental fish trade is estimated at US$ 1 billion, and retail value about

US$ 3 billion (FAO 2012).

The European Union is the world’s largest ornamental fish importer. The United

States, however, is the single largest importer in the world with a reported value of

imported ornamental fishes of US $37.2 million in 2011 (USDA-ERS 2012).

Approximately 80% of the imported value into the U.S. aquarium market is from

freshwater ornamental fishes (Chapman et al. 1997). Most freshwater ornamental fishes

originate from Asia, including Singapore, Hong Kong, Malaysia, Thailand, the

Philippines, Sri Lanka, Taiwan, Indonesia, and India. These account for about two thirds

of the world’s total export value (Chapman et al. 1997; Adams et al. 2001). Greater than

90% of freshwater ornamental fish species are captive bred and reared in ponds, tanks,

or vats until they reach a marketable size in 4 to 6 months (FAO 2012). The top 10

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freshwater ornamental fish species imported to the U.S. in 1992 were guppy (Poecilia reticulata), neon tetra (Paracheirodon innesi), platy (Xiphophorus maculatus), Siamese

fighting fish (Betta splendens), goldfish (Carassius auratus), Chinese algae-eater

(Gyrinocheilus aymonieri), molly (Poecilia sphenops), cardinal tetra (Paracheirodon

axelrodi), glassfish (Parambassis lala), and tiger barb (Puntius tetrazona) (Chapman et

al. 1997; Adams et al. 2001).

Unlike freshwater ornamental fish species that are mostly farm-bred, marine

species are mainly wild-caught (FAO 2012). The Philippines and Indonesia were the

most important exporting countries supplying approximately 59% of the total number of

marine ornamental fishes exported worldwide (Wood 2001; Wabnitz et al. 2003). More

than 1,470 species of marine fishes are traded globally, with an estimate of 20–24

million individuals (Wabnitz et al. 2003). The top 10 marine ornamental fish species

imported to the U.S. between years 1997-2002 were green chromis (Chromis viridis),

Koran angelfish (Pomacanthus semicirculatus), yellow tang (Zebrasoma flavescens),

three-stripe damsel (Dascyllus aruanus), Ocellaris clownfish (Amphiprion ocellaris),

yellowtail damsel (Chrysiptera parasema), blue damsel (Chrysiptera cyanea), half-blue

damsel (Chrysiptera hemicyanea), domino damsel (Dascyllus trimaculatus), and

bluestreak cleaner wrasse (Labroides dimidiatus). These fishes accounted for 39% of all

marine fish species imported into the U.S. between years 1997 and 2002 (Wabnitz et al.

2003).

Diseases of Ornamental Fishes

Infectious agents including parasites, viruses, bacteria, and water molds are important health concerns for both freshwater and marine ornamental fishes. The imported ornamental fishes, especially those from Asia, have been reported to carry a

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wide range of pathogens (Mouton et al. 2001; Ariel 2005; Whittington and Chong 2007;

Musa et al. 2008). There is concern that the international ornamental fish trade has the potential to transfer unwanted or novel organisms and disease agents to importing countries (Font and Tate 1994; Evans and Lester 2001; Ariel 2005; Moravec and

Justine 2006).

Parasitic infections are very common in the ornamental fish industry worldwide.

High prevalence of parasites, both external and internal, has been reported frequently in fishes imported from Asia (Evans and Lester 2001; Mouton et al. 2001; Kim et al. 2002;

Thilakaratne et al. 2003). Evans and Lester (2001) examined 5 freshwater ornamental fishes commonly imported to Australia, including guppy (P. reticulata), platy (X. maculatus), neon tetra (P. innesi), cardinal tetra (P. axelrodi), and sucking catfish

(Hypostomus plecostomus), for parasite infection and found relatively high prevalence of 10 parasite species in different shipments originating from Southeast Asia. Another study found a total of 12 species of parasites among 13 species of freshwater ornamental fishes originating from 23 export farms in Sri Lanka, and an overall prevalence of parasitism of 45.3% (Thilakaratne et al. 2003). Furthermore, Kim et al.

(2002) reported 7 species of parasites in 15 species of freshwater tropical ornamental fishes imported to Korea from Southeast Asia. Approximately 15 species of parasites were identified in imported ornamental fishes during the quarantine process in Australia between 1999-2004 (Whittington and Chong 2007).

In addition to parasitic infections, bacterial infections are among the most

common diseases of recently transported ornamental fishes. Gram-negative bacteria, including Aeromonas, Citrobacter, Edwardsiella, Flavobacterium, Pseudomonas, and

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Vibrio, are the most frequently isolated genera from ornamental fishes. Moreover, gram-

positive bacteria such as Mycobacterium and Streptococcus have also been reported in many important ornamental fishes (Lewbart 2001). Bacterial infections can be a primary cause of disease or may be opportunistic, causing disease and mortality in fish with predisposing conditions such as ectoparasites or trauma from transport and handling.

Several factors may predispose fish to bacterial infections. Stress is defined as

the physiological responses of the body to any demand made upon an organism that

helps in the maintenance of homeostasis (Barton 2002). Stressors impacting recently

transported ornamental fishes include poor water quality, shipping conditions, trauma,

and crowding. Many cases of bacterial infection have been reported in recently imported

ornamental fishes. For example, Whittington and Chong (2007) identified 9 genera of

bacteria in diseased ornamental fishes imported into Australia during 1999-2004. One

study reported that Mycobacterium sp. was isolated from goldfish and koi imported to

South Africa (Mouton et al. 2001). Gratzek et al. (1978) examined 77 bags of fishes

imported from Southeast Asia to the U.S. and found a total of 11 genera of bacteria in

fishes from 51 bags. Fishes in those bags were noted with bacteremia. Moreover, 14

additional genera of bacteria were isolated from the shipping water.

Viral infections are also of concern in ornamental fishes (Armstrong and

Ferguson 1989; Anderson et al. 1993; Bernoth and Crane 1995; Rodger et al. 1997;

Sudthongkong et al. 2002; Jeong et al. 2008), although historically there is less information available about viruses than for parasitic and bacterial diseases. Several viruses have been reported to cause diseases in ornamental fishes including megalocytivirus, lymphocystivirus, betanodavirus, spring viremia of carp virus (SVCV),

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koi herpesvirus (KHV), and cyprinid herpesvirus-1 (CyHV-1, or carp pox) (Table 1-1).

Many of these viruses have been associated with significant mortality (30% to 70% mortality rates) in important marine and freshwater fishes resulting in economic losses

(Schuh and Shirley 1990; Hedrick et al. 2000; Weber et al. 2009; Yanong and Waltzek

2010). Clinical disease may only occur within certain temperature ranges which are optimal for replication of specific viral agents (Petty and Fraser 2005).

Tetrahymena Infection in Imported Guppies

Guppies

Guppy Poecilia reticulata (Figure 1-1) is an euryhaline ornamental fish belonging to the class Actinopterygii, order Cyprinodontiformes, family Poeciliidae (livebearing fishes). It is one of the most popular aquarium pet fishes in the U.S. and many parts of the world because of its spectacular coloration, and ease of reproduction and maintenance (Mojetta 1993; Chapman et al. 1997).

Guppies are sold in 2 forms, wild-type and fancy. Wild-type guppies have a short colorful caudal fin with many pattern variations such as pintail, double sword, and bottom sword; nevertheless, they are smaller and less colorful than the selectively bred fancy guppy. The genetics of fancy guppies have been developed by selective breeding to produce numerous varieties since the mid-1960s (Dawes 1991). Today’s fancy guppies have large, elongated bodies and long caudal fins with many different color and pattern variations. Several countries around the world including Singapore, the United

States (mainly in Florida), Japan, Germany, the United Kingdom, Russia, and some

African countries now produce about 40 or more varieties of fancy guppy. Currently, there are approximately 5 common fancy guppy varieties that are most popular with aquarium hobbyists. These are Cobra, Tuxedo, Mosaic, Grass, and Swordtail (Thai

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Department of Fisheries 2002). In addition to the different varieties and color patterns of guppies, the caudal fins also have many shape variations which are shown in Figure 1-3

(Jacobs 1971).

Guppies are native to warm, freshwater streams of Trinidad and Tobago,

Venezuela, Guyana, Barbados, and Northern Brazil (Axelrod and Vorderwinkler 1988).

They have been introduced into many lakes and ponds throughout the world for control

of mosquitoes (Mills 1993; Houde 1997). Guppies are an important aquaculture product

in Asia, where several strains are bred and exported to pet stores worldwide.

Guppies thrive in water temperatures ranging from 24 to 28°C (75–82°F) (Jacobs

1971). They can tolerate salinity equal to seawater for up to 7 days (Schelkle et al.

2011), but favor a lower salinity level of 0.5 - 9 psu (Fernando and Phang 1985). They

are adaptable to a range of water chemistries, but hardness of 374 mg/L with pH of

about 7.8 seems to be ideal (RMGA 2010).

Guppies are omnivorous, feeding on insect larvae, small crustaceans, benthic

detritus, and algae. They do cannibalize their own young and the eggs and fry of other

fishes (Jacobs 1971; Houde 1997). They are also highly sexual dimorphic (Houde

1997). Male guppies are smaller, but they have a brilliantly colored body and large

colorful tail. The larger females are plainer and have a smaller tail with little decoration

(Mills 1993). Female guppies can grow to 5 cm but males only grow to 3 cm (Dawes

1991; Axelrod and Vorderwinkler 1988). Females are easily differentiated from males by

their larger body size, drab color, and the presence of a dark gravid spot at the

urogenital opening (Figure 1-4A). Males are more streamlined than females when

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viewed dorsally and they have a gonopodium which is a modified anal fin (Fernando and Phang 1985; Houde 1997) (Figure 1-4B).

Guppies are ovoviviparous which means that the egg develops into embryos inside the female’s body, resulting in the release of free-swimming fry (Dawes 1991;

Houde 1997). They have a polygamous mating system with internal fertilization, and they can reproduce year round. Male guppies perform courtship displays in combination with a change in color pattern. When the females respond to the males, the males then inseminate the female using the gonopodium (Houde 1997). Male guppies, and many other poeciliids, have a very short gonopodium, thus they need to make contact with a female in order to complete courtship and sperm transfer (Dawes 1991). Female guppies can store sperm from several males, and may continue to produce offspring many times from a single spawning event. A large female can produce as many as twenty or more fry in each brood (Houde 1997).

Mating preferences of female guppies are primarily based on male color patterns, courtship behavior, and other characteristics of the male including total length, tail length and degree of carotenoid coloration (Houde 1997). Female guppies are believed to choose the males that have conspicuous color patterns, high display rates, and long caudal fins. The display frequency of male guppies is affected by the behavior, reproductive status, and size of the females (Houde 1997). Parasitism may affect the fish’s ability to attract a mate, as color and behavior may be changed. The sigmoid display (moving in 2 directions, like the letter S) rate of parasitized males is likely to be lower than that of males with no parasites, which renders them less attractive to females

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(Kennedy et al. 1987). While healthy female guppies do not desire parasitized males, the healthy males also discriminate against parasitized females (Lopez 1999).

Tetrahymena

Tetrahymena (Figure 1-2) is a common parasite infecting many species of

ornamental and food fishes; however, guppies and other poeciliids are most sensitive to

the parasite. Tetrahymena can infect both vertebrates and invertebrates (Thompson

1958; Hoffman et al. 1975). It has been reported to infect a broad range of fish species,

including guppy (P. reticulata), neon tetra (P. innesi), pristella tetra (Pristella maxillaris),

cherry barb (Puntius titteya), zebrafish (Danio rerio), Atlantic salmon (Salmo salar), and

hybrid striped bass (Morone chrysops x Morone saxatilis) (Ferguson et al. 1987;

Ponpornpisit et al. 2000; Smith et al. 2003). Furthermore, they have been reported to

infect other animals including amphibians, chick embryos, and insects (Thompson 1958;

Hoffman et al. 1975).

Tetrahymena represents a genus of holotrichous ciliated protozoans belonging to

the class Oligohymenophorea, order Hymenostomatida, family Tetrahymenidae (Lom

1995; Basson and Van As 2006; Colorni 2008). It possess an ovate to pear-shaped

body (approximately 100 μm × 30 μm in size), with a pointed anterior end. A radially

symmetrical body is covered with 20-35 meridional kineties over the entire body

surface. A caudal cilium is presented in some species (Hoffman et al. 1975; Imai et al.

2000; Basson and Van As 2006; Colorni 2008). Tetrahymena has a small deep pocket

buccal cavity with an undulating membrane on the right side and 3 diagonal

membranellae on the left side of the cytostome (Sleigh 1973; Hoffman et al. 1975;

Hoffman 1999; Colorni 2008).

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Tetrahymena has been reported worldwide and it is generally ubiquitous in aquatic ecosystems (Basson and Van As 2006). It is a free-living, normally saprozoic ciliate that feeds on organic matter and bacteria in natural habitats or the bottom of an aquarium (Leibowitz et al. 2005; Lom 1995). However, several species are known to be

facultative histophagous parasites that are capable of invading host tissue and feeding

on cell debris (Thompson 1958; Lom 1995).

Tetrahymena is not an obligate pathogen; however, it can become highly

invasive to fish hosts (Leibowitz et al. 2005). It has been reported to spread systemically

via the blood vessels or musculature to internal organs (Colorni 2008; Noga 2010).

Tetrahymena has been found in many tissues of infected fish including scale pockets,

muscle fiber, intestine, liver, kidney, heart, eye socket, cranial cavity, and spinal cord

(Imai et al. 2000; Leibowitz and Zilberg 2009). Clinical signs of Tetrahymena infection

may include white areas on the skin and fins, epidermal sloughing, raised scales,

clamped fins, fin erosion, excessive mucus and loss of equilibrium (Hoffman et al. 1975;

Imai et al. 2000; Colorni 2008). In severe cases, an inflammatory reaction may be

observed in tissues that have been invaded by the parasite (Hoffman et al. 1975).

Although this parasite can be seen on living fish using a dissecting microscope, it is

more easily demonstrated by examining wet mounts of infected tissues with a light

microscope (Hoffman et al. 1975). Species identification requires silver impregnation

methods, such as Klein’s dry silver impregnation technique, Protargol method or

Chatton-Lwoff technique, allowing visualization of the ciliary rows (Hoffman et al. 1975;

Lom and Dykova 1992; Imai et al. 2000). Species can also be identified using the

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mitochondrial cytochrome-c oxidase subunit 1 (cox1) gene as a DNA barcode

(Chantangsi et al. 2007).

Currently, there are 4 pathogenic species of Tetrahymena that have been described infecting fish; T. corlissi, T. pyriformis, T faurei, and T. rostrata (Lom 1995).

The two most prevalent species of Tetrahymena reported as causative agents of “Tet”

or “Guppy Killer” disease in guppies and other tropical ornamental fishes are T. corlissi

and T. pyriformis (Kim et al. 2002; Lom 1995; Noga 2010). Tetrahymena corlissi has a size range of 55 μm × 30 μm, with 25-31 meridional kineties and one caudal cilium. It has a round to oval macronucleus, a single micronucleus, and a single posterior contractile vacuole. It produces reproductive cysts that can divide to produce 2–8 tomites. Tetrahymena pyriformis is about 50 to 80 μm in its long axis. It has 17–21

kineties and lacks the caudal cilium (Lom and Dykova 1992; Ponpornpisit et al, 2001;

Basson and Van As 2006; Colorni 2008).

Tetrahymena infection has been reported in guppies imported from Asia. Kim et

al. (2002) reported the first finding of T. corlissi in guppies imported to Korea and they

considered this parasite to be a cause of mass mortality. Imai et al. (2000) also

identified ciliates that infected 55% of guppies imported from Singapore as T. corlissi.

Other studies found 65% prevalence of T. corlissi infection of guppies imported from

Singapore (Evans and Lester 2001). Moreover, T. corlissi and T. pyriformis were

identified in guppies exported from Sri Lanka (Thilakaratne et al. 2003).

Many factors may influence the severity of Tetrahymena infection in ornamental

fishes. Recent transport, rough handling, the presence of other diseases, as well as

environmental factors such as temperature, photoperiod, and water quality may all

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impact the severity of Tetrahymena infections. Leibowitz et al. (2005) reported that high

levels of ammonia, high organic loads, and low water temperatures contribute to the

severity of Tetrahymena infection in guppies and other fish species. They revealed that

the infection rate of fish kept at water temperature of 16–17°C (61–62.6°F), 4-6 mg/L

total ammonia nitrogen, pH 7.6, and 10% organic matter was significantly higher than it

was in fish exposed to temperature of 24–25°C (75–77°F), 0–2 mg/L total ammonia

nitrogen, pH 7.6, and 0–1% organic matter. Furthermore, they also suggested that

infection was significantly increased when fish were exposed to shipping conditions,

which included high stocking densities, and associated deterioration of water quality

(high carbon dioxide levels, lower pH, supersaturation with oxygen and accumulations

of ammonia).

In addition to water quality and shipping conditions, photoperiod may also affect

prevalence of infection. Leibowitz et al. (2005) found that mortalities due to

Tetrahymena infection were higher in fish kept under a 12 h light: 12 h dark photoperiod

than fish kept in a 0 h light: 24 h dark photoperiod. Susceptibility of fish to Tetrahymena

infection also increased with inappropriate husbandry practices. For example, high

mortality of guppies caused by Tetrahymena infection was reported in a facility that

operated 24 h, 4 days a week (S. Moore, Segrest Farms, personal communication).

This meant that the fish were exposed to a 24 h light: 0 h dark photoperiod.

Concurrently, fish were frequently exposed to nets and handling. This facility also had

multiple inputs of new fish from different sources weekly, multiple species were held in

the same system, and the recirculating systems were not all-in-all-out (S. Moore,

Segrest Farms, personal communication). Consequently, Tetrahymena may have

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become established in the recirculating system where the guppies were held and may have been transmitted to other fish by cohabitation, resulting in the mortality events reported at that facility.

Many therapeutic chemicals are available for treatment of external tetrahymenosis including formalin, sodium chloride (salt), potassium permanganate, malachite green, methylene blue, and copper sulphate (Ponpornpisit et al. 2001; Klinger

and Francis Floyd 2009; Noga 2010). Ponpornpisit et al. (2001) also reported successful control of experimental Tetrahymena infection in guppies using a

combination of 0.5% salt bath and food containing C-UPIII (Chinese herbal mix

containing mainly polysaccharides). The results from their study showed that guppies

treated with both chemoprophylaxis (salt) and immunoprophylaxis (C-UPIII) had

significantly lower infection rates than guppies treated with a single dose of an

antiparasitic chemical. However, Herbert and Graham (2008) reported ineffectiveness of

salt and formalin baths in treating Tetrahymena corlissi infection in golden perch

(Macquaria ambigua). Once the parasite penetrates tissues and becomes systemic, it is

untreatable and often fatal to the fish (Kim et al. 2002; Chettri et al. 2009; Leibowitz et

al. 2010; Noga 2010).

Megalocytivirus Infection in Fishes, with Emphasis on the Orbiculate Batfish (Platax orbicularis)

Orbiculate Batfish

Orbiculate batfish (Platax orbicularis) (Figure 1-5) are marine ornamental fish

belonging to the class Actinopterygii, order Perciformes, family Ephippidae. They have a

round shape and a strongly laterally compressed shape with dark dorsal and anal fins

that almost encircle the body. Adults have a yellowish silvery or dusky dark bar through

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the eyes and another bar behind the head with a few small, scattered black spots on the body. Juveniles are reddish brown, with irregular black dots and small white dots (black-

edged) on the body. Males can grow up to 50 cm in length. Orbiculate batfish are

omnivorous and may feed on algae, invertebrates and small fishes (Mills 1993; Capuli

and Ortanez 2011).

Orbiculate batfish are endemic to the tropical Indo-Pacific region, including the

Red Sea extending to East Africa and including the Tuamoto Islands. Their range

extends north to southern Japan and south to northern Australia and New Caledonia,

but they do not reach Hawaii or the eastern Pacific (Figure 1-6). Orbiculate batfish have

also been recorded off the coast of Florida and in the Western Central Atlantic Ocean.

(Mills 1993; Mojetta 1993; Capuli and Ortanez 2011). Orbiculate batfish inhabit shallow

coastal waters and seaweed reefs (depth range 5 - 30 m) and have a preferred water

temperature range of 22 to 28°C (71.7–82.4°F). Juveniles form small groups among

mangroves and other sheltered waters, whereas adults move out to open waters and

deeper areas (Mills 1993; Capuli and Ortanez 2011). Currently, these fish are usually

wild-caught for sale in the ornamental fish trade. Most of these collections occur in the

Indo-Pacific region.

Megalocytivirus

Fish viruses belonging to the genus Megalocytivirus are important causes of

systemic disease and mortality in various species of marine and freshwater fishes

(Weber et al. 2009; Yanong and Waltzek 2010; Zhang et al. 2011). Megalocytiviruses

have been associated with mass mortalities and serious economic losses in Asian

aquaculture industries, and have been reported in other parts of the world (Oh et al.

2006; Wang et al. 2009; Weber et al. 2009). Many species of important marine and

24

freshwater ornamental, food, and game finfish are known to be susceptible to megalocytiviruses. Infection has been reported in Banggai cardinalfish (Pterapogon kauderni) (Weber et al. 2009), barramundi (Lates calcarifer) (Wang et al. 2009), dwarf

gourami (Trichogaster lalius) (Anderson et al. 1993; Sudthongkong et al. 2002),

freshwater angelfish (Pterophyllum scalare) (Rodger et al. 1997) and other cichlids

(Armstrong and Ferguson 1989), grouper (Family Serranidae) (Yanong and Waltzek

2010), largemouth bass (Micropterus salmoides) (Yanong and Waltzek 2010), livebearing fishes (Family Poecilidae) (Jeong et al. 2008), Nile tilapia (Oreochromis niloticus) (McGrogan et al. 1998), red drum (Sciaenops ocellatus) (Weng et al. 2002), and threespine stickleback (Gasterosteus aculeatus) (Waltzek et al. 2012).

Currently, 6 megalocytiviruses have been completely sequenced at the full genome scale including infectious spleen and kidney necrosis virus (ISKNV) (He et al.

2001), red sea bream iridovirus (RSIV) (Kurita et al. 2002), large yellow croaker iridovirus (LYCIV) (Chen et al. 2003), rock bream iridovirus (RBIV) (Do et al. 2004),

orange-spotted grouper iridovirus (OSGIV) (Lu et al. 2005), and turbot reddish body

iridovirus (TRBIV) (Shi et al. 2010) (Table 1-2).

Megalocytiviruses are in the family Iridoviridae. These are large, double-stranded

DNA viruses (dsDNA) with an icosahedral capsid diameter of 120–200 nm (Figure 1-7).

The family Iridoviridae is subdivided into five genera, Iridovirus, Chloriridovirus,

Ranavirus, Megalocytivirus, and Lymphocystivirus. Members of this family infect a

range of poikilothermic hosts, including invertebrates, fish, amphibians, and reptiles

(William et al. 2005; Chinchar et al. 2009).

25

Megalocytiviruses have been reported to cause diseases in a wide variety of fish species at water temperatures of 7.5–32°C (46–89.6°F) (Yanong and Waltzek 2010;

Waltzek et al. 2012). Clinical signs of megalocytivirus infection are non-specific and may include lethargy, loss of appetite, darkening of body pigmentation, erratic swimming behavior, ulceration, hemorrhages on the skin, fins and gills, severe anemia, pale gills, and white feces. High mortality rates are common and may approach 100% of the population (Weber et al. 2009; Yanong and Waltzek 2010; Zhang et al. 2011; Waltzek et al. 2012). At necropsy, fish infected with megalocytivirus may have a markedly enlarged spleen, reno- and hepatomegaly, petechiae in the liver, and hemorrhagic fluid in the coelom. Histopathologically, cytomegaly and tissue necrosis may be visible in several internal organs, especially in the spleen, liver, and kidney (Chen et al. 2003; Weber et al. 2009; Yanong and Waltzek 2010; Waltzek et al. 2012). High numbers of cytomegalic cells characterized by strongly basophilic granular intracytoplasmic inclusions are often observed within various organs including spleen, kidney, intestine, eyes, gonads, and connective tissues. Cytomegalic cells may also be found in lower numbers in the liver, heart, brain, and gills (Gibson-Kueh et al. 2003; Weber et al. 2009; Yanong and Waltzek

2010; Zhang et al. 2011; Waltzek et al. 2012).

To confirm megalocytivirus infection in fish, special diagnostic tests are necessary. These may include histopathologic examination of whole fish or tissue samples (spleen, kidney, liver, heart, intestine, stomach, gonad, and brain) that have been fixed in 10% neutral buffered formalin, embedded in paraffin, sectioned at 3–4 μm, and stained with hematoxylin and eosin (H&E); virus isolation by cell culture using homogenized fresh or frozen tissue inoculated onto Grunt Fin (GF) cell line (derived

26

from fin tissues of Blue Striped Grunt, Haemulon sciurus); use of transmission electron microscopy to search for virus particles in cytomegalic cells from suspected tissue samples; and use of the molecular techniques, which include polymerase chain reaction

(PCR) assays to detect viral DNA (Weber et al. 2009; Yanong and Waltzek 2010; Zhang

et al. 2011; Waltzek et al. 2012).

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Table 1-1. Reports of viral diseases in ornamental fishes. Virus Susceptible fishes References Common Name Species or Family Nodaviridae Betanodavirus Guppy Poecilia reticulata Hegde et al. 2003 Convict surgeonfish Acanthurus triostegus Yanong 2010 Orbiculate batfish Platax orbicularis David et al. 2010

Alloherpesviridae Cyprinivirus Cyprinid herpesvirus 1 Common carp/Koi Cyprinus carpino Wolf 1988 (CyHV-1) or carp pox Cyprinid Goldfish Carassius auratus Jung and Miyazaki herpesvirus 2 (CyHV- 1995 2) or hematopoietic necrosis herpesvirus of goldfish Cyprinid herpesvirus 3 Common carp/Koi Cyprinus carpino Hedrick et al. 2000 (CyHV-3) or koi herpesvirus (KHV)

Iridoviridae Lymphocystivirus Glassfish Chanda ranga Wolf 1988; Paperna The marine angels Pomacanthidae et al. 2001 Cichlids Cichlidae Butterflyfishes Chaetodontidae Megalocytivirus Banggai cardinalfish Pterapogon kauderni Weber et al. 2009 Freshwater angelfish Pterophyllum scalare Rodger et al. 1997 Dwarf gourami Trichogaster lalius Paperna et al. 2001; Sudthongkong et al. 2002 Poeciliids Poeciliidae Paperna et al. 2001 Orange chromide Etroplus maculatus Armstrong and Ferguson 1989 African lampeye Aplocheilichthys Sudthongkong et al. killifish normani 2002

Rhabdoviridae Vesiculovirus Spring viremia of carp Common carp/Koi Cyprinus carpino Wolf 1988; virus Goldfish Carassius auratus Hoole et al. 2001

28

Table 1-2. Comparison of known megalocytiviruses infecting fishes. Virus Susceptible Disease Gross signs Histologic Lesions References Species Temp (°C) Infectious Mandarin fish 25–34°C Swollen and brownish kidney; enlarged Cytomegaly of infected cells in the He et al. spleen and (Siniperca spleen; pale, and distended liver with spleen, kidney, cranial connective 2000; He et kidney chuatsi) petechial hemorrhages; pale heart; pale tissue, and endocardium; diffuse al. 2002 necrosis virus gills; petechial hemorrhages in the necrosis in the hematopoietic (ISKNV) operculum, mandible, orbit around the eye, tissues of kidney and spleen. base of dorsal and ventral fins, caudal fin, and abdomen.

Red sea Red sea bream 25–27°C Severe anemia; petechiae of the gills; Enlarged cells in the spleen, heart, Wang et al. bream (Pagrus major) enlargement of the spleen, kidney, liver, kidney, intestine and gills; necrosis 2003 iridovirus and more than heart, and gills. and degeneration of splenic pulp (RSIV) 30 other and ellipsoid sheaths. species of cultured marine fish

Large yellow Large yellow 27–30°C Pale and ulcerative gills; red spotted liver; The widespread necrosis of Chen et al. croaker croaker swollen congested spleen and kidney. tissues and enlargement of cells, 2003 iridovirus (Pseudosciaena mainly in the spleen and kidney, (LYCIV) crocea) but also in the gills, liver, and intestinal epithelium.

Rock bream Rock bream, 23–27°C Petechial hemorrhage of the fins, tail, and Enlarged cells in the kidney, Jung and iridovirus Striped gills; congestion of the liver; enlargement spleen, heart, liver, and gills; Oh 2000 (RBIV) beakperch of the spleen and kidney. necrosis in the hematopoietic (Oplegnathus tissues of kidney; necrosis and fasciatus) hemorrhage around the splenic pulp.

Orange Orange spotted 25–30°C Severe anemia; petechiae on the gills; Enlargement of cells and necrosis Lu et al. spotted grouper enlarged spleen. of renal and splenic hematopoietic 2005 grouper (Epinephelus tissues. iridovirus coioides) (OSGIV)

Turbot reddish Turbot 25–30°C Pale gills with local hemorrhages; petechial Enlarged cells in the spleen, Shi et al. body iridovirus (Scophthalmus hemorrhages in fins and fin bases, muscle, kidney, and gill. 2004 (TRBIV) maximus) and skin.

29

Figure 1-1. An example of a fancy guppy (Poecilia reticulata). Photo courtesy of Thai Department of Fisheries.

30

Figure 1-2. A photomicrograph of Tetrahymena sp. on the skin mucus of a guppy. Klein’s dry silver impregnation. (Bar = 20 μm). Photo courtesy of Preeyanan Sriwanayos.

31

Figure 1-3. A diagram shows the standard shapes of guppy caudal fin. A) fan tail; B) triangle or delta tail; C) veil tail; D) flag tail; E) double sword; F) top sword; G) bottom sword; H) lyre tail; I) spade tail; J) pointed tail or spear tail; K) round tail; and L) pintail. Photo courtesy of Steve Challis.

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A

B

Figure 1-4. Differentiation of male and female guppies. Guppies are highly sexual dimorphic. Female guppies have larger body size, plain color, and a dark gravid spot at the urogenital opening on both sides (A). Male guppies can be easily differentiated from females by their gonopodium (a modified anal fin) and colorful body and tail variation (B). Photos courtesy of wereallwet.com; Akvaristika slike.

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Figure 1-5. Orbiculate batfish (Platax orbicularis). Photo courtesy of Joe DE VROE.

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Figure 1-6. Distribution of orbiculate batfish. Photo courtesy of Kaschner et al., 2010.

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Figure 1-7. Orbiculate batfish iridovirus particles. (Bar = 200 nm). Photo courtesy of Preeyanan Sriwanayos.

36

CHAPTER 2 PRELIMINARY ASSESSMENT OF PARASITE LOAD IN SELECTED IMPORTED GUPPIES (POECILIA RETICULATA)

Parasitic infections are among the most important health concerns in the ornamental fish industry worldwide (Molnár 1987; Paperna 1991; Moravec et al. 1999;

Piazza et al. 2006). Parasitism may cause morbidity and mortality in ornamental fishes, and may also affect the growth and behavior of fishes. Parasitic disease can impact marketability (appearance, size, health status) and cause a reduction in value, resulting in economic losses that exceed loss related to mortality alone (Schloz 1999).

Many species of parasites have been reported in ornamental fishes imported to the U.S. and other countries (Gratzek et al. 1978; Moravec et al. 1999; Evans and

Lester 2001; Mouton et al. 2001; Kim et al. 2002; Thilakaratne et al. 2003; Piazza et al.

2006). Parasites, including protistans, monogeneans, nematodes, digeneans, cestodes, and crustaceans, are commonly detected on ornamental fishes after arrival at import facilities. More than 20 species of parasites have been found in recently imported ornamental fishes during investigations of disease prevalence (Moravec et al. 1999; Kim et al. 2002; Thilakaratne et al. 2003; Whittington and Chong 2007).

Guppy (Poecilia reticulata) is a livebearing ornamental fish in the order

Cyprinodontiformes, family Poeciliidae, and is one of the most popular imported ornamental fish because of its coloration and tail varieties. However, guppies have been reported to have a high loss rate at many ornamental fish import facilities due to parasitic infections (Lawhavinit et al. 2002; Kim et al. 2002; Thilakaratne et al. 2003).

Several studies suggest that guppies and other poeciliids cultured in Asia may be a source of new parasite introductions to importing countries and there is concern that

37

these organisms may become established among native fish species (Font and Tate

1994; Rigby et al. 1997; Evans and Lester 2001; Moravec and Justine 2006).

The most common parasites found in recently imported guppies are a ciliated

protozoan (Tetrahymena), a monogenean (Gyrodactylus), and a nematode

(Camallanus). Kim et al. (2002) first reported finding Tetrahymena corlissi and

Camallanus cotti from guppies imported from Southeast Asia into Korea. These two

parasites were associated with mass mortalities of guppies on many tropical fish import

facilities in Korea. Thilakaratne et al. (2003) also reported a high prevalence of parasitic

infections in guppies housed at export facilities in Sri Lanka prior to shipping. These

included Dactylogyrus spp., Gyrodactylus spp., Tetrahymena spp., and Trichodina spp.

Moreover, guppies imported from Singapore into Australia were found to be infected with C. cotti and T. corlissi when they were released from quarantine (Evans and Lester

2001).

Approximately 600 million ornamental fishes, including guppies, are imported into the United States from Southeast Asia each year (Gratzek et al. 1978; Chapman et al.

1997). These fish may cause problems for the U.S. ornamental fish import industry since they can die from parasitic disease during transportation or after their arrival, resulting in economic losses as reported in many countries (Lawhavinit et al. 2002; Kim et al. 2002; Thilakaratne et al. 2003). However, few studies have been done to investigate parasites of ornamental fishes arriving in the U.S. (Gratzek et al. 1978).

Another risk is the potential for accidental introduction of non-native parasites to native fish populations. Understanding the occurrence of parasites in ornamental fishes

38

imported to the U.S. is necessary to evaluate the risks they may pose to the U.S.

ornamental and food fish industries, as well as to native fauna.

Based on the results of diagnostic screening of a small sample of newly imported

guppies for an ornamental fish wholesale facility, I conducted this preliminary health

assessment to 1) determine the prevalence of parasites on 1 shipment of guppies

imported into the U.S. from Asia; 2) identify the parasites present on these imported

guppies; and 3) assess the intensity of parasitic infection on these imported guppies. I

hypothesized that 1) guppies imported to the U.S. from Asia would not have heavy parasite burdens, 2) guppies imported to the U.S. from Asia would not have diverse parasite populations, and 3) guppies imported to the U.S. from Asia would not to be infected with Tetrahymena.

Materials and Methods

Imported Guppy Population

In the preliminary health assessment described here, a parasite investigation was conducted on a shipment of imported guppies. The fish had been packed in polystyrene shipping boxes and transported via airplane to Atlanta, GA from Singapore

on September 1st, 2010. The fish were then transported by truck for an estimated 6

hours to a research laboratory, Fisheries and Aquatic Sciences, University of Florida,

Gainesville. The total shipment was 6 bags of fish packed at 100 fish per bag. Fish in

this shipment had been separated by gender and variety.

A total of 30 imported guppies were examined during the investigation. Ten fish

were randomly removed from each of 3 pre-designated shipping bags (3 groups of 10

fish each). Prior to transport from Atlanta, these fish were rebagged into 3 smaller plastic bags with clean water.

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Gross Examination

Prior to removal from shipping bags, fish were evaluated for appearance including characteristics of body and fin color, body condition, behavior, body shape, and finnage. Moreover, fish were examined carefully by gross observation to screen for the presence of macroscopically visible parasites. Fish were euthanized, and then weighed, and measured for total length (TL), and tissues were collected as described below.

Collection of Tissues for Evaluation of Microscopically Visible Parasites

To evaluate parasite load at the microscopic level, tissue samples of skin mucus, caudal fin, and gill were examined with a light microscope at 40x, 100x, and 400x magnifications. Tissue samples of caudal fin and skin mucus were collected following standard techniques (Noga 2010); however, all 4 gill arches on both the left and right sides were removed for inspection.

In addition to external examinations described above, the entire gastrointestinal

(GI) tract from esophagus to the anus was dissected free and placed on a slide with drops of water. A coverslip was used to gently compress the sample and the GI tract

was examined for intestinal parasites using a light microscope as described above.

The total number of parasites seen with the light microscope was averaged so

that the number of parasites per field of view could be used to assess the intensity of

infection. Parasite load was defined as light, moderate, or heavy based on criteria

modified from DiMaggio et al. (2008). For ciliated protozoans, a light infection was

defined as < 10 organisms per 400x field of view; moderate was 10-50 organisms per

400x field of view; and heavy was > 50 organisms per 400x field of view. For nematodes

and monogeneans, a light infection was defined as < 10 organisms per 40x field of view;

40

moderate was 10-25 organisms per 40x field of view; and heavy was > 25 organisms per 40x field of view.

Parasite Identification Protocols

If trichodinids, Chilodonella, or Tetrahymena were detected, slides were air-dried and stained using Klein’s dry silver impregnation technique, in which slides were stained with an aqueous solution of 2% silver nitrate (AgNO3) for 10 minutes followed by 20

minutes exposure to ultraviolet (UV) light (Lom and Dykova 1992; Buchmann 2007). If other ciliates or flagellates were detected, new smears were prepared, air-dried, and stained with Diff-Quik® (Medion-Diagnostics Ag, Duedingen, Switzerland). For

identification of monogeneans, parasites were placed on a slide and mounted using a

glycerin jelly technique (Woodland 2006). For nematodes, parasites were preserved in a

hot mixture of 70% ethanol and glycerin or mounted by a glycerin jelly technique.

Digeneans were fixed in hot Bouin’s fixative overnight and then transferred to 70% alcohol. Myxosporeans, leeches, and copepods were preserved in 10% neutral buffered formalin (Hoffman 1999). All parasites were identified to genus based on morphological characters and standard taxonomic keys as described by Hoffman (1999).

Data Analyses

The percentage of fish that were positive for each type of parasite was

determined by dividing the number of infected fish by the total number of fish examined

(Bush et al. 1997). Preliminary assessment of the prevalence of parasites was

attempted by defining a population as the shipment group. Prevalence is the number of

infected animals as a percentage of the total number of animals in the sample

population at a given time (OIE 2009).

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Results

At arrival, guppies (average weight of 0.49 g and average total length of 40 mm) were robust and approximately 93.3% were active and had bright coloration. There were no grossly visible parasites evident on external examination of any fish, but 10% of these fish had mild frayed fins. At necropsy, mild to severe telangiectasia at the tips of the gills were found in 26.7% of fish.

Of the 30 guppies examined, 2 genera of parasites including a nematode and a monogenean, were observed in 26.7% of fish.

A total of 49 nematodes (Figure 2-1) were recovered from the gastrointestinal tract of 8 guppies (26.7%). The infection was graded as “light” except for 1 fish that had a “moderate” infection level (Table 2-1). All nematodes were identified as the genus

Camallanus by their red color and laterally compressed buccal capsule with a basal ring and 2 lateral valves, with longitudinal ridges (Rigby et al. 1997; Hoffman 1999).

Camallanus is medium-sized and slim worm in the class Secernentea, order

Camallanida, family Camallanidae (Hoffman 1999). They often appear reddish in color because of feeding on the host’s blood.

Specimens of Gyrodactylus (Figure 2-2) were collected from the skin mucus and the caudal fin of 2 guppies (6.67%), with light infection levels (Table 2-1). Parasites were identified by their pairs of anchors with supporting dorsal and ventral transverse bars, the presence of a pair anchors on the embryo in the adult worm, and the absence of eye spots (Hoffman 1999). Gyrodactylus belongs to the class Monogenea, order

Monopisthocotylea, family Gyrodactylidae (Hoffman 1999). It has a fusiform body with 2

anterior cephalic lobes, each bearing spike sensillium. The body length of living,

42

unflattened, moderately contracted specimens is approximately 700-800 μm depending on species (Harris 1986).

Discussion

Despite previous reports of a high prevalence of parasite infection of guppies

(Evans and Lester 2001; Mouton et al. 2001; Kim et al. 2002; Thilakaratne et al. 2003),

only 2 genera of parasites (Camallanus and Gyrodactylus) were observed on guppies

arriving in the U.S. from Singapore. A wider range of parasites in guppies newly

imported from Singapore was expected than was found in this study. Although the

intensity of infection of the monogenean parasite, Gyrodactylus, found on infected fish

in this preliminary health assessment was relatively low, the results for Camallanus

were different. One fish was observed with moderate Camallanus infection.

The most prevalent parasite observed in fish in this preliminary health

assessment was Camallanus which is a nematode parasite commonly found in the

intestine of guppies (Rigby et al. 1997; Kim et al. 2002). The preliminary finding of

Camallanus infections of imported guppies are consistent with previous reports

indicating that Camallanus is an important concern in guppies imported from Asia

(Mouton et al. 2001; Kim et al. 2002; Thilakaratne et al. 2003). Camallanus has been

reported in many freshwater species of fishes in East, South and Southeast Asia

(Moravec and Nagasawa 1989; Kim et al. 2002; Moravec et al. 2003; Thilakaratne et al.

2003). In the past few decades it has been reported worldwide in many species of

fishes in Australia, Brazil, Europe, Hawaii, and North America. Several reports have

speculated that it may have been introduced with the introduction of aquarium fishes

from Asia (Hoffman and Schubert 1984; Moravec and Sey 1988; Font and Tate 1994;

43

Moravec 1994, Rigby et al. 1997; Levsen 2001; Menezes et al. 2006; Morevec and

Justine 2006).

Gyrodactylus, a monogenean parasite and a common ectoparasite of guppies,

was the second most prevalent parasite observed in this study. Gyrodactylus

bullatarudis, G. costaricensis, G. gambusiae, G. katharineri, G. rasini, and G. turnbulli,

have been reported in ornamental fishes in many countries around the world including

Australia, England, Korea, Sri Lanka, the United State, Trinidad, and Venezuela

(Rogers and Wellborn 1965; Harris 1986; Harris and Lyles 1992; Cone 1995; Dove and

Ernst 1998; Harris and Cable 2000; Kim et al. 2002; Thilakaratne et al. 2003; Harris et

al. 2004; Buchmann and Bresciani 2006). Two species of Gyrodactylus, G. bullatarudis

and G. turnbulli, are common parasites of guppies and other poeciliids (Harris 1986;

Buchmann and Bresciani 2006). G. bullatarudis has been reported in wild poeciliids in

central and North America and G. turnbulli has been reported to infect a range of fishes

and amphibians in North America (Rogers and Wellborn 1965; Harris 1986). The origin

of G. turnbulli infection on guppies is unclear, since guppies ranged from the north-

eastern coast of South America, extending north into Trinidad, and the Lesser Antilles

(Jacobs 1971). G. bullatarudis and G. turnbulli have been identified in guppies in the

U.S. since 1956 (Rogers and Wellborn 1965; Hoffman 1999). It is possible that these 2

species of Gyrodactylus may have already existed in wild population of fishes in the

U.S. However, identifying gyrodactylids to species was beyond the scope of my

investigation.

The low diversity of parasites observed in this study could be attributed to several

factors. First, although the shipment of guppies in this assessment was evaluated at a

44

level that did allow an assessment of prevalence at the 10% level, the small numbers of fish examined were not sufficient to determine lower prevalence of other parasites that

may have been present in these populations. Amos (1985) suggested that the minimum

sample size for assessment of fish pathogens in a population of interest must be based

on statistical methodology which provides 95% confidence that infected fish will be

observed. Estimated sample sizes in accordance with assumed prevalence of infection

for the population of interest are shown in Table 2-2. In order to estimate a presumed

parasite prevalence of 5% in this shipment which had 600 fish, I would have had to

evaluate at least 55 individual fish per shipment (Table 2-2).

Second, these fish may have come from parasite-free sources or they may have

been treated for ectoparasites before exporting. Many guppy farms in Singapore hold

fish in quarantine tanks before shipping for 1-3 days in an effort to prevent export of fish

carrying potential pathogens (Fernando and Phang 1985). It is possible that fish arriving

at these pre-shipment holding facilities may have been treated for common ectoparasite

infections. Treating the fish prior to export would minimize infection and reduce the

prevalence of parasites on transported fish. Some export farms in Sri Lanka are

reported to routinely use antiparasitic compounds such as formalin, malachite green,

methylene blue, and acriflavine at their facilities (Thilakaratne et al. 2003). It is

interesting to note the guppies still had monogeneans despite being placed into clean

water prior to truck transport to Florida. This suggests water change alone may not be

enough to inhibit or remove the parasite from infected fish. Further, treatments for

ectoparasites would not be expected to impact nematode infections in the

45

gastrointestinal tract, which may explain the relatively high prevalence of Camallanus- positive fish.

It is important to note that this study was conducted as a preliminary health assessment of guppies arriving into the U.S. from 1 Asian country. Although the intent was to improve understanding of parasite loads on imported fish, the scope of the project was very small and consequently, interpretation of data is limited. Results from this study should not be considered representative of the parasite infection in the whole population of guppies imported from Asia into the U.S.

Parasitic infections are of concern in ornamental fish industries worldwide. Future

investigation of parasitic disease of imported ornamental fishes should be performed

using more appropriate sampling procedures including examination of a sufficient

number of fish to detect parasites even at the level of 2% prevalence. Numbers of fish

examined for the population of interest must be high enough to provide 95% confidence

that infected fish will be included in the fish sampled, based on presumed prevalence of

infection in that population.

Despite the important findings from this health assessment, I was not able to

pursue a full-scale investigation due to the intensive labor and time required to properly

perform the investigation. This preliminary study involved 6 people and 9 hours to be

performed in a timely manner.

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Table 2-1. Microscopically visible parasite were found on external surfaces and within the gastrointestinal tract of guppies imported to the U.S. from Singapore (n = 30). The location of parasites and percent of examined fish that were found to be infected is shown. (40x magnification) Parasites Site of Infection Infection levels* % of Infected Fish Monogenean Gyrodactylus Caudal fin, Skin L 7 Nematode Camallanus GI tract L to M 27 *L = Light (< 10 organisms per field of view), M = Moderate (10-25 organisms per field of view).

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Table 2-2. Determining the number of fish that need to be sampled in order to assess pathogen prevalence in a lot (or population) of fish (Amos 1985). Lot size At 2% prevalence At 5% prevalence At 10% prevalence size of sample size of sample size of sample 50 50 35 20 100 75 45 23 250 110 50 25 500 130 55 26 1000 140 55 27 1500 140 55 27 2000 145 60 27 4000 145 60 27 10,000 145 60 27 100,000 or more 150 60 30

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Figure 2-1. Camallanus spp. recovered from the gastrointestinal tract of a guppy imported from Singapore. Photo courtesy of Preeyanan Sriwanayos.

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Figure 2-2. Gyrodactylus spp. observed on the fin of a guppy. (Bar = 100 μm). Photo courtesy of Preeyanan Sriwanayos.

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CHAPTER 3 PRELIMINARY STUDY OF TRANSMISSION OF TETRAHYMENA FROM INFECTED TO TETRAHYMENA-FREE GUPPIES (POECILIA RETICULATA)

Tetrahymena are important parasites of freshwater ornamental and food fish

species (Hoffman et al. 1975, Ferguson et al. 1987; Ponpornpisit et al. 2000; Smith et

al. 2003; Basson and Van As 2006). Several cases of Tetrahymena infection have been reported in imported guppies and the parasite is considered to be a cause of mass mortality of guppies in ornamental fish farms worldwide (Imai et al. 2000; Evans and

Lester 2001; Kim et al. 2002). It has been also reported to cause disease and mortality in guppies during transportation or after arrival at wholesale facilities (Evans and Lester

2001; Lawhavinit et al. 2002). Tetrahymena was associated with a 9-22% mortality rate

of newly imported guppies at an ornamental fish wholesale facility in the U.S. (S. Moore,

Segrest Farms, personal communication). However, in a preliminary health assessment

that examined 30 guppies newly imported into the U.S. from Singapore, Tetrahymena

was not observed on any fish (Chapter 2).

Despite previous reports of occurrence, there is little known about the infection

process of Tetrahymena in guppies. Leibowitz and Zilberg (2009) reported successful

infection of guppies with Tetrahymena corlissi by cohabitation; however, very high

stocking densities were used. Their cohabitation model used guppy fry (0.06 ± 0.01 g)

stocked in Petri dishes with 40 mL of water at an average stocking density of 250 fish/L

(15 g of fish/L). This stocking density is much higher than the typical stocking densities

encountered in wholesale facilities in Florida. A stocking density of 1.2–6.25 fish/L (1.5–

3.75 g of fish/L, average weight of 0.30–0.60 g) is more typical of wholesale facilities in

Florida (R. P. E. Yanong, University of Florida, personal communication).

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In addition to high stocking densities, Leibowitz and Zilberg (2009) maintained experimental fish at water temperatures of 24 ± 1°C (73.4–77°F), an acceptable temperature for guppies, but in sub-optimal water quality parameters (4–6 mg/L total ammonia nitrogen and 1–2 mg/L nitrite). These conditions may have increased the susceptibility of the fish to Tetrahymena infection or enhanced the ability of the parasite to replicate, thus increasing the inoculum presented to the fry (Leibowitz et al. 2005;

Leibowitz and Zilberg 2009).

The present study was conducted to assess Tetrahymena transmission between guppies, by cohabitation, under conditions that were similar to those present in Florida wholesale facilities. Experimental objectives were to determine whether 3 Tetrahymena- infected guppies were able to transmit the parasite by cohabitation to Tetrahymena-free guppies within 14 days; to determine whether 10 Tetrahymena-infected guppies were able to transmit the parasite by cohabitation to Tetrahymena-free guppies within 14 days; and to determine if the Tetrahymena-free guppies developed more severe disease when exposed to the heavier inoculum of 10 Tetrahymena-infected fish during the 2-week observation period.

Materials and Methods

Experimental Animals

Tetrahymena-free fish

A group of 150 Tetrahymena-free female guppies were obtained from an import facility in Atlanta, Georgia on September 14, 2011 and shipped to the Fish Disease

Research Laboratory, Fisheries and Aquatic Sciences, University of Florida, Gainesville.

Five of these fish were collected, euthanized, weighed, and measured for total length

(TL). They were then examined for the presence of external parasites using standard

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diagnostic techniques (mucus scraping, fin and gill biopsies) (Noga 2010). Samples of skin mucus, caudal fin, and gill tissue were examined using a light microscope at 40x,

100x, and 400x magnifications to ascertain that fish were free of Tetrahymena and other external parasites.

Remaining Tetrahymena-free female fish (n = 145) were housed in two 80-L tanks containing 44 L of well water in a flow-through system supplied with supplemental aeration (Figure 3-1) for 10 weeks.

Water temperature and pH were measured daily using HI 98128 pHep®5

pH/Temperature Tester (Hanna Instrument, Inc., Smithfield, Rhode Island). An LDO

(Luminescent Dissolved Oxygen) digital meter (HQ20, Hach Company, Loveland,

Colorado) was used to measure dissolved oxygen (DO) once a week. Tanks were

maintained at a water temperature of 24±1°C (73.4–77°F), pH 8.0-8.5, and dissolved

oxygen 8.3 mg/L. All fish were fed commercial flake food (tropical flake, containing

crude protein minimum 45.0%, crude fat minimum 7.0%, crude fiber maximum 3.0%,

moisture maximum 10.0%, Florida Tropical Fish Farms Association Co-op Store,

Tampa, FL) at 3% of their body weight once daily. Fish were exposed to a 12 h light: 12

h dark photoperiod.

Any fish that died during the 10-week holding period was examined as described

above for the presence of ectoparasites. During this time, 9.3% of female fish died for

unknown reasons. Tetrahymena and other parasites were never observed on these fish.

One hundred twenty-six fish (average weight of 0.65 g and the average total length of

39 mm) were selected for the experiment at the end of the 10-week holding period.

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These fish were not examined for the presence of parasites immediately prior to the experiment to prevent handling stress.

Tetrahymena-infected fish

Two hundred and fifty male guppies from a population of imported guppies known to be infected with Tetrahymena were obtained from a wholesale facility. Staff at the facility had examined skin mucus, fin and gill biopsies with a light microscope to confirm the infection.

Upon receipt of these fish at the Fish Disease Research Laboratory, Gainesville,

Florida on August 31, 2011, ten fish were removed and sedated with 10 mg/L buffered tricaine methanesulfonate (MS-222, Western Chemical, Inc., Ferndale, Washington),

weighed and measured for total length (average weight of 0.51 g and average total

length of 37 mm). Following examination, these fish were returned to the general

population.

On day 4 post-arrival, these guppies were found to be infected with columnaris

and 51.2% of them died within 5 days of arrival. Surviving fish were treated with a short-

term bath of 25 mg/L tetracycline for 1 hour followed by a 100% water change for 3

consecutive days. Although the columnaris infection resolved, a 19.7% mortality rate

attributed to the Tetrahymena infection continued during the first 2 weeks of the holding

period. Light Chilodonella infection was also found on 45% of fish that died.

Consequently, fish were also treated with 125 mg/L formalin for 30 min to control the

Chilodonella infection. Remaining fish (n = 98) were kept in an 80-L tank containing 44 L

of well water in flow-through system. The tank was maintained at a water temperature of

25±1°C (75.2–78.8°F) and pH 8.3–8.5.

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Following formalin treatment, 20 fish were selected for examination and assessment of parasite burden. Each of these fish was sedated with 10 mg/L MS-222 and placed in a sterile Petri dish containing tank water and sedative, and then examined for the presence of Tetrahymena using a stereo dissecting microscope. If parasites were present, the level of infection was graded into 1 of 3 assessments of infection

(light, moderate, or heavy) applying criteria modified from DiMaggio et al. (2008) (Table

3-1). Sedated fish were then placed into a recovery tank containing aerated tank water until they revived.

Of fish examined, the percentage of lightly to moderately infected fish was less than 30%. In an effort to exacerbate the infection, water temperature was lowered to

20– 22°C (68–71.6°F). Total ammonia nitrogen was allowed to increase to 0.2–0.3 mg/L at pH of 8.3–8.5; this resulted in unionized ammonia of 0.02-0.05 mg/L. The stocking density was increased to 7.5 fish/L (Leibowitz et al. 2005; Leibowitz and Zilberg 2009) by decreasing the water volume. Fish were then examined for the presence of parasite every other day until the percentage of lightly to moderately infected fish increased to desired inoculum range of 30%. Eighty-one lightly to moderately Tetrahymena-infected fish were used as an inoculum for the experiment.

Experimental Infection

The experimental design was a fully randomized design, and consisted of 4 treatment groups; low inoculum, high inoculum, negative control, and positive control.

Twelve 20-L tanks were individually numbered, and treatments were randomly assigned to these tanks (3 replicates per treatment). Excluding the positive control tanks, 14

Tetrahymena-free female guppies were placed into each tank. Fourteen Tetrahymena- infected male guppies were placed into each of the positive control tanks. In the low

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inoculum treatment, each tank received 3 Tetrahymena-infected male fish (1

Tetrahymena-infected fish: 4.67 Tetrahymena-free fish). Each tank of the high inoculum treatment received 10 Tetrahymena-infected male fish (1 Tetrahymena-infected fish: 1.4

Tetrahymena-free fish). The negative control tanks were stocked with 14 Tetrahymena-

free female guppies.

To maintain a uniform stocking density of 6.25 fish/L in all treatment groups, the

water volumes were adjusted to 2.60 L for low inoculum tanks, 3.63 L for high inoculum

tanks, and 2.12 L for negative control and positive control tanks. Fish were exposed to a

12 h light: 12 h dark photoperiod.

Tanks were maintained on a flow-through system with 2 turnovers per hour at water temperatures of 24±1°C (73.4–77°F), pH 8.4–8.8, and continuous aeration.

Dissolved oxygen, water temperature, and pH were recorded daily. A water sample was

collected from all tanks from each treatment 3 times a week and analyzed using a Hach

Freshwater Fish Farmers Kit (FF-1A, Hach Company, Loveland, Colorado) for total

- ammonia nitrogen (NH3-N) and nitrite (NO2 ). The flow-rate was measured in each tank daily using a graduated cylinder.

Fish were fed approximately 3% of their body weight daily. Uneaten food and feces were siphoned from the bottom of the tanks daily.

During the experiment, all fish were observed at least twice daily for clinical signs of Tetrahymena infection. Early clinical signs included white areas of the skin and frayed fins (Figure 3-2). These observations were recorded daily. Dead fish were removed daily and were examined for the presence of Tetrahymena. At the end of experiment, all

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surviving fish were euthanized, examined for the presence of Tetrahymena, and graded for infection level.

Histological Examination

Prior to beginning the experiment, 3 Tetrahymena-infected male fish were

selected for histological examination. After euthanasia, the coelomic cavity of these fish

was opened and the opercula were removed. Whole fish were then fixed in 10% neutral

buffered formalin and embedded in paraffin, sectioned at 3 μm, mounted onto glass

slides, and stained with hematoxylin and eosin (H&E). These representative processed

samples were examined under a light microscope for the presence of Tetrahymena.

Statistical Analysis

All statistical analyses were carried out with SAS 9.2 (SAS Institute Inc., Cary,

North Carolina). Infection rate, mortality rate, and mean intensity (mean number of parasites per fish) were compared by Kruskal-Wallis test. Differences were considered

significant at P < 0.05.

Results

Experimental Infection

Tetrahymena was transmitted to Tetrahymena-free females by cohabitation, though only in the low inoculum treatment. No Tetrahymena were observed in fish from the negative control treatment.

Only 3 female guppies died during the experiment, these mortalities occurred on days 11-13. Two of these fish were in the low inoculum treatment and light levels of

Tetrahymena were found in both of these fish. Tetrahymena was not found in the one female that died in the high inoculum treatment. No mortalities occurred in

Tetrahymena-free female fish from the negative control treatment (Table 3-2). The

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mortality rate was not significantly different among the treatment groups (P = 0.26)

(Figure 3-3).

Tetrahymena infection in the male fish that were used as the inoculum was variable. Light to heavy Tetrahymena infection (average of <10 to >50 ciliates per high power fields) was seen on these fish, and the infection rate ranged from 7% to 78%

(Table 3-3). Mortality of infected male fish continued throughout the duration of the experiment. A total of 40.4% infected male fish in the positive control treatment, 36.7% in the low inoculum treatment, and 66.7% in the high inoculum treatment died during the experiment. Tetrahymena-infected males in all treatments died at a higher rate than the females.

During the examination of dead fish, parasites other than Tetrahymena were found. While the experiment was in progress, light to heavy Gyrodactylus infections were observed on all 3 females that died during the experiment. Further, it was also found on 85% of female fish that were examined at the end of the experiment, with light

to heavy levels of infection (average of <10 to >25 monogeneans per high power fields)

present in every tank (Table 3-2). Further, light Chilodonella infection was found in 1

female fish from the high inoculum treatment group.

When male fish were examined, Tetrahymena was found in each tank; however,

infection level was variable (Table 3-3). Gyrodactylus was found in 64.5% of male fish

from all tanks in the high and low inoculum treatment groups, as well as from 2 fish in

one of the positive control tanks. Chilodonella was also found in low numbers on 23% of

male fish used in the high inoculum treatment group, as well as in 1 tank used in the

positive control treatment group (Table 3-3).

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Gross examination of the 3 female guppies that died during the experiment revealed light Tetrahymena infection (average of <10 ciliates per high power fields) on the skin of the 2 fish from the low inoculum treatment. The mean intensity of

Tetrahymena in exposed Tetrahymena-free fish was not significantly different among the treatment groups and locations (skin: P = 0.37; fin: P = 0.10; gill: P = 1.00). Mild fin erosion and white patchy areas were observed on the skin of these fish.

Histological Examination

High numbers of Tetrahymena had been found on all 3 infected male fish that were examined histologically. Histological examination revealed high numbers of

Tetrahymena under the scale pockets, on gill filaments, and between muscle fibers

(Figure 3-6A, B, D). The organism was also found internally in these fish. Tetrahymena was observed in the pericardium, gonads, adipose tissues, mesentery and peritoneum

(Figure 3-6C). Low numbers of the parasite were also seen around the eye sockets of 2 fish. An inflammatory response was not observed in any tissues invaded by

Tetrahymena.

Discussion

In this study, Tetrahymena was successfully transmitted from Tetrahymena- infected male guppies to Tetrahymena-free female guppies in the low inoculum treatment group within 11 days of exposure. This finding is similar to a report of successful transmission of Tetrahymena corlissi in guppy fry (Leibowitz and Zilberg

2009). They demonstrated successful transmission of the parasite within 5 days; however, fish in their study were held at a much higher stocking rate than that used in this study. The lack of Tetrahymena in the negative control treatment group further supports the conclusion that the parasite was transmitted by cohabitation in this study.

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Parasite transmission was limited, and did not occur in the high inoculum treatment. The concurrent Gyrodactylus infection may have affected parasite transmission. As the number of Gyrodactylus increased, there may have been competition between the 2 parasites, potentially disadvantaging Tetrahymena.

Buchmann et al (1999) reported that rainbow trout infected with Gyrodactylus derjavini had lower infection rates and mortality than uninfected fish when exposed to

Ichthyophthirius multifiliis, a ciliate protistan. They suggested that rainbow trout previously infected with the ectoparasitic monogenean may have developed a host response that was protective against the external ciliate parasite.

In my experiment, Gyrodactylus was not found on any fish during the holding

period, but was present in both males and females after the experiment began. This

parasite was likely present in low numbers but undetected until fish were examined

more closely while the experiment was in progress. This suggests that the screening

method using the dissecting microscope was inadequate to detect the low level of

infection that had to have been present. The fish should have been subjected to a more

thorough examination immediately prior to the start of the experiment.

Numerous Tetrahymena were observed histologically in both external and

internal organs of the 3 male fish that were examined prior to beginning the experiment.

The parasite was not seen in the gastrointestinal tract of these fish, which is consistent

with reports by Imai et al. 2000 and Leibowitz and Zilberg 2009, which suggested that

the oral route is not a possible site of internal infection. Further, Imai et al. (2000)

suggested that the organism penetrates from scales to internal organs via muscle

fibers. This is consistent with the lesions observed in experimental fish which clearly

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demonstrated the ability of this organism to invade the skin, skeletal muscle, and internal organs of a suitable host.

The main problem with the experimental design used in this study was the low

number of infected fish and replicates. The low level of statistical power in this study

was caused by the small sample size. This contributed to my inability to detect statistical

differences in the infection and mortality rates, mean intensity, and the presence-

absence data among the treatment groups.

Another problem found in this study was the difficulty in achieving a 30%

Tetrahymena infection rate in the inoculum fish. This resulted in an inconsistent

inoculum and I may have inadvertently used more heavily infected males in the low

inoculum treatment group (Table 3-3).

During the holding period, Chilodonella had been detected in male fish. In an

effort to eradicate this parasite from experimental fish, they were treated with formalin

(125 mg/L for 30 min), followed by water changes (Francis-Floyd 1996; Klinger and

Francis-Floyd 2009; Noga 2010). During this time salinity was maintained at 5 psu.

These treatments may have also killed some Tetrahymena, making it difficult to

maintain the desired 30% infection rate in inoculum fish. Herbert and Graham (2008) did

report that salt and formalin baths were ineffective when treating Tetrahymena corlissi

infection in golden perch (Macquaria ambigua). This would explain the survival of some

Tetrahymena in inoculum fish, but it is unclear whether the parasite was completely

unaffected by these chemicals.

Removal of dead fish from experimental tanks may have also decreased the

number of parasites in the environment, further compromising parasite transmission.

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Leibowitz and Zilberg (2009) demonstrated that guppies exposed to 1000 ciliates/mL had significantly higher infection rates than those exposed to 100 ciliates/mL.

The use of flow-through water systems and the daily cleaning routine, while

appropriate for the fish, did not provide the environmental factors thought to be optimal

for invasion. Leibowitz et al. (2005) demonstrated the infection rate of Tetrahymena in

guppies was significantly higher when exposed to poor environmental conditions that

included high ammonia nitrogen and high organic load. Fish in my experiment were not

exposed to these conditions.

The use of different genders for infected fish and Tetrahymena-free fish in this

study may also have affected Tetrahymena transmission. Kennedy et al. (1987)

suggested that parasitized male guppies are unattractive to females as their display

rates decreased. Moreover, parasite-free female guppies usually avoid parasitized

males and parasite-free males also discriminate against parasitized females (Lopez

1999). Since only male guppies were infected at the beginning of the experiment, it is

conceivable that Tetrahymena-free female fish tried to avoid them, but I did not directly

observe this. My experiment was not designed to evaluate behavior, so subtle

behavioral responses would not have been obvious.

Despite these difficulties, Tetrahymena was successfully transmitted from

infected guppies to Tetrahymena-free guppies; however, the process is still poorly

understood. In future work, experimental design should eliminate the confounding

factors described above. In conclusion, Tetrahymena can cause significant disease and

high mortality in a diverse species of freshwater fishes, and methods for transmission

and effective therapy deserve further study.

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Table 3-1. Classification of parasite intensity per field of view (FOV) at predetermined magnifications (DiMaggio et al. 2008). Parasite Magnification Light Moderate Heavy (Per FOV) (Per FOV) (Per FOV) Monogeneans 40x 1-10 > 10-25 > 25 Ciliates 57x 1-10 > 10-50 > 50 400x 1-10 > 10-25 > 25

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Table 3-2. Mortality rates, infection rates, and biopsy findings of 42 exposed Tetrahymena-free female guppies from each treatment during the 14-day experimental period. Treatments Tank No. Biopsy findings* Infection Mortality rate (%) (%) Negative control (n = 14) 1 L Gyrodactylus 100 0 2 L-M Gyrodactylus 100 0 3 L-H Gyrodactylus 100 0 High inoculum (n = 14) 1 L-H Gyrodactylus 100 7 2 L-H Gyrodactylus 100 0 3 L Gyrodactylus 93 0 L Chilodonella 7 Low inoculum (n = 14) 1 L Tetrahymena 7 7 L Gyrodactylus 71 2 L-H Gyrodactylus 86 0 3 L Tetrahymena 7 7 L Gyrodactylus 93 *L = Light, M = Moderate, H = Heavy.

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Table 3-3. Mortality rates, infection rates, and biopsy findings of infected male guppies during the 14-day experimental period. Treatments Tank No. Biopsy findings Infection Mortality rate (%) rate (%) Positive control (n = 14) 1 L-H Tetrahymena 79 71 L Chilodonella 21 2 L-H Tetrahymena 36 36 L Gyrodactylus 21 3 H Tetrahymena 7 14 High inoculum (n = 10) 1 L Tetrahymena 20 30 L-H Gyrodactylus 90 L Chilodonella 10 2 L Tetrahymena 30 50 L Gyrodactylus 50 L Chilodonella 10 3 L-M Gyrodactylus 80 30 L Chilodonella 50 Low inoculum (n = 3) 1 L Tetrahymena 33 100 L Gyrodactylus 67 2 L Gyrodactylus 67 0 3 L Tetrahymena 67 100 L Gyrodactylus 33 *L = Light, M = Moderate, H = Heavy.

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Figure 3-1. Holding tanks. Fish were housed in 80-L tanks containing 44 L of well water in flow-through system with 2 turnovers per hour and supplemental aeration during the holding period. Photo courtesy of Traiwut Sriwanayos.

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A

B

Figure 3-2. External lesions observed in guppies infected with Tetrahymena. Lesions included A) scale loss and development of deep ulcer on the caudal peduncle (arrow), B) severe fin erosion was also are a common finding. Photos courtesy of Preeyanan Sriwanayos.

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(Infected males)

Figure 3-3. Mortality rate (%) of exposed Tetrahymena-free guppies during the 14-day experimental period. No significant difference was found in mortality rate among the treatment groups. Data correspond to the mean ± SE. P < 0.05.

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Figure 3-4. Cumulative mortality (%) of guppies used in the experiment during the 14- day experimental period.

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Figure 3-5. Tetrahymena infection rate (%) in exposed Tetrahymena-free guppies during the 14-day experimental period. No significant difference was found in infection rate among the treatment groups. Data correspond to the mean ± SE. P < 0.05.

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A B

C D

Figure 3-6. Guppies infected with Tetrahymena. A) Tetrahymena under the scale pockets (arrows). Bar = 50 µm, B) Tetrahymena in the gill filaments. Bar = 20 µm, C) Tetrahymena in the gonad. Bar = 20 µm, D) Tetrahymena penetrate between muscle fibers. Bar = 20 µm. Photos courtesy of Preeyanan Sriwanayos.

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CHAPTER 4 MEGALOCYTIVIRUS INFECTION IN ORBICULATE BATFISH (PLATAX ORBICULARIS)

Viruses have been identified from finfish and are important problems for aquaculture industries (Hetrick and Hedrick 1993; Bernoth and Crane 1995; Rodger et

al. 1997, He et al. 2000, Paperna et al. 2001, Weng et al. 2002, Wang et al. 2003, Shi et

al. 2004, Wang et al. 2011). Members of the genus Megalocytivirus are iridoviruses

pathogenic to a wide variety of freshwater and marine fishes, and they have negatively

impacted ornamental and food fish aquaculture worldwide (Armstrong and Ferguson

1989; Anderson et al. 1993; Rodger et al. 1997; Sudthongkong et al. 2002; Jeong et al.

2008; Chinchar et al. 2009; Weber et al. 2009; Yanong and Waltzek 2010; Zhang et al.

2011; Waltzek et al. 2012). Megalocytiviruses induce lethal systemic diseases at water

temperatures ranging from 7.5–32°C (45.5–89.6°F) (Chen et al. 2003; Wang et al. 2011;

Zhang et al. 2011; Yanong and Waltzek 2010; Waltzek et al. 2012). Epizootics may

result in 100% mortality under intensive aquaculture conditions (Anderson et al. 1993;

He et al. 2000; Paperna et al. 2001; Rodger et al. 1997; Sudthongkong et al. 2002).

These viruses can spread horizontally from fish to fish by cohabitation, exposure to

water or equipment carrying the virus, and by ingestion of infected fish or feed items (He

et al. 2002; Go and Whittington 2006; Yanong and Waltzek 2010).

Megalocytivirus-infected fish may exhibit non-specific clinical signs including

lethargy, anorexia, hyperpigmentation, exophthalmos, skin lesions, unusual swimming

behavior, severe anemia, and white feces (Chen et al. 2003; Weber et al. 2009; Wang

et al. 2011; Waltzek et al. 2012). On post-mortem examination, fish infected with megalocytiviruses may exhibit hemorrhagic lesions, renomegaly, splenomegaly, hepatomegaly, and coelomic distension resulting from hemorrhagic fluid accumulation

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(Chen et al. 2003; Weber et al. 2009; Yanong and Waltzek 2010; Wang et al. 2011;

Zhang et al. 2011; Waltzek et al. 2012). Histopathologic examination typically reveals

cytomegalic mesenchymal cells characterized by strongly basophilic or amphophilic

granular intracytoplasmic inclusions observed in multiple organs including the spleen,

kidney, liver, heart, brain, gills, intestine, eyes, and gonads (Chen et al. 2003; Gibson-

Kueh et al. 2003; Weber et al. 2009; Yanong and Waltzek 2010; Zhang et al. 2011;

Waltzek et al. 2012). Transmission electron microscopy invariably reveals numerous

icosahedral virus particles with a capsid diameter of between 130-160 nm in the

cytoplasm of infected cells (Chen et al. 2003; Weber et al. 2009; Yanong and Waltzek

2010; Wang et al. 2011; Zhang et al. 2011; Waltzek et al. 2012).

Phylogenetic analyses support 4 separate species within the genus

Megalocytivirus (Kurita and Nakajima 2012; Waltzek et al. 2012). Megalocytiviruses

related to Red Sea bream iridovirus (RSIV) cluster into 1 of 2 genotypes and have been

associated with mass mortality epizootics in more than 30 maricultured species in

Japan, Korea, China, and Southeast Asia. Infectious spleen and kidney necrosis virus

(ISKNV), originally isolated from mandarin fish (Siniperca chuatsi) raised for food in

China, has also resulted in epizootics in more than 10 species of freshwater ornamental

fishes (He et al. 2000; Paperna et al. 2001; Sudthongkong et al. 2002; Yanong and

Waltzek 2010). A second ISKNV genotype was recently recognized following disease

episodes in ornamental and food fish species (Kurita and Nakajima 2012), which

included Banggai cardinalfish iridovirus (BCIV) isolated from a marine ornamental

species, Banggai cardinalfish (Pterapogon kauderni) (Weber et al. 2009) and marbled

sleepy goby iridovirus (MSGIV) isolated from a freshwater species, marbled sleeper

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goby (Oxyeleotris marmorata), cultured for food in China, (Wang et al. 2011), The third

species, turbot reddish body iridovirus (TRBIV), has primarily been associated with

disease in cultured Asian flatfish species, turbot (Scophthalmus maximus) (Shi et al.

2004). A fourth megalocytivirus species, threespine stickleback iridovirus (TSIV), was

recently described from an epizootic that occurred in a Canadian collection of

threespine stickleback (Gasterosteus aculeatus) (Waltzek et al. 2012).

Orbiculate batfish (Platax orbicularis) are tropical marine ornamental species

belonging to the order Perciformes, family Ephippidae. These species can be found

throughout the Indo-western Pacific region (Mills 1993; Mojetta 1993; Capuli and

Ortanez 2011). Orbiculate batfish occur in a variety of habitats; mangroves in shallow

coastal waters, other sheltered waters, deep seaweed reefs, and open waters. They

prefer water temperatures ranging from 22–28°C (71.7–82.4°F) (Mills 1993; Capuli and

Ortanez 2011). Jayasankar (1998) reported that orbiculate batfish were becoming more

popular in the marine ornamental fish trade.

In the present study, I report a megalocytivirus infection in P. orbicularis that was

associated with mortality in recently imported wild-caught fish acquired by a public

aquarium in Belgium. The objective of this study was to compare clinical aspects of the

disease as well as microscopic and ultrastructural features of the virus to previously

described megalocytiviruses including the virus associated with the only other marine

ornamental epizootic reported in Banggai cardinalfish, order Perciformes, family

Apogonidae.

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Materials and Methods

Clinical History

A mortality event occurred in March 2010 among a group of 10 orbiculate batfish quarantined at a public aquarium in Belgium. The wild-caught juvenile orbiculate batfish

(average of 12 cm in total length) from Southeast Asia were purchased by an ornamental fish import facility in the United Kingdom and then were sent to a quarantine facility in the UK, where these fish were acclimatized and quarantined prior to delivery to the aquarium in Belgium in February of 2010. At the aquarium the batfish were kept in a quarantine tank equipped with an external power filter. Water quality parameters at the aquarium were maintained at a temperature of 24°C; pH 7.8; nitrite <0.3 mg/L; nitrate 0-

12.5 mg/L; salinity 32 psu; 96% oxygen saturation; and undetectable total ammonia nitrogen. The orbiculate batfish were fed mysid (Mysidopsis) and brine shrimp (Artemia)

5-6 times per day.

At arrival, several fish had moderately frayed fins and large fleshy white nodules

on the fins. Two days after arrival, one fish became anorexic, displayed an increased respiration rate, and was noticeably darker than other batfish in the same tank. A wet mount examination of skin mucus of this batfish was negative for parasites. This fish

was dipped in freshwater once a day for 2 days. All fish in the same tank were treated

with a formalin bath at 100 mg/L for 6 hours to control a suspected parasite infection.

Fish were also treated prophylactically with oxytetracycline (Aquatet®, PHARMAQ

Limited, Fordingbridge, Hampshire) bath at 100 mg/L daily for 7 days to prevent a

secondary bacterial infection. Mortality continued during the treatment and a total of 8

fish died within the 4-week quarantine period. Necropsy findings were unremarkable;

however, the fleshy white nodules were noted on the dorsal fins of all fish that died and

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were also present on the remaining 2 fish in the tank. Following necropsy, whole fish

with opened coelomic cavity were fixed in 10% neutral buffered formalin and sent to the

International Zoo Veterinary Group (IZVG) Pathology Laboratory, UK for histological

processing and examination.

Histopathology

For 2 separate formalin-fixed fish that had been necropsied, 5 transverse

sections and 1 sagittal section of the head were cut and dehydrated through a series of

water and alcohol mixtures to full alcohol. Tissues were then cleared through a clearant

and embedded into paraffin blocks (2 blocks per individual). The 4 paraffin blocks were

sectioned at 3 μm, mounted onto glass slides, and stained with hematoxylin and eosin

(H&E). In addition to examination by IZVG Pathology Laboratory, these blocks were

also submitted to the UF Aquatic Pathobiology Laboratory, Gainesville, Florida, U.S.

Transmission Electron Microscopy

A paraffin block was selected by evaluating an H&E-stained slide for features

consistent with megalocytivirus pathology. Heart tissue was selected from this block for

transmission electron microscopy (TEM) evaluation and sent to the Electron Microscopy

Laboratory (EML), Department of Medical Pathology and Laboratory Medicine, School

of Medicine, University of California at Davis. An area with characteristic

megalocytivirus pathology in the heart was removed from the paraffin block and placed

in 100% xylene; after clearing overnight, the tissue was rehydrated and processed using

a standard protocol (Johannessen 1977) as previously described (Hayat 1989).

Ultrathin sections (45–60 nm) were stained with uranyl acetate and lead citrate and

viewed in a transmission electron microscope at the EML. I also viewed these samples

with an electron microscope at the University of Florida Electron Microscopy Laboratory.

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DNA Extraction, Polymerase Chain Reaction Amplification, and Sequencing

I extracted DNA from 50-μm sections of formalin fixed, paraffin embedded

tissues using a commercial extraction kit (DNeasy® Blood and Tissue Kit, Qiagen Inc.,

Valencia, California) following the manufacturer’s instructions for paraffin-embedded

tissues. I performed polymerase chain reaction amplification of the viral full length major

capsid protein (MCP) gene sequence using 14 primer pairs designed from the complete

MCP gene sequence of the Banggai cardinalfish iridovirus (BCIV which is also known

as the Pterapogon kauderni iridovirus or PkIV; GenBank accession # AB669096.1)

(Table 4-1).

The 20-μl PCR reaction mixtures consisted of 0.1 µl of Platinum Taq DNA

Polymerase (Invitrogen, Carlsbad, California), 2.0 µl of 10X PCR Buffer, 0.8 µl of 50 mM

MgCl2, 0.4 µl of 10mM dNTPs, 1.0 µl of 20 µM of forward and reverse primers, 11.7 µl

of molecular grade water, and 3 µl of DNA template. The PCR conditions used for all

reactions included an initial denaturation of 5 min at 95˚C, followed by 50 cycles of

denaturation at 95˚C for 1 min, annealing at 50˚C for 1 min, and extension at 72˚C for 1

min, followed by a final elongation step at 72˚C for 10 min. After electrophoresis, I cut

bands of interest from the gel and extracted their DNA using the QIAquick gel extraction

kit (Qiagen, Valencia, California). Purified DNA fragments were then submitted to the

University of Florida Interdisciplinary Center for Biotechnology Research (ICBR) for

sequencing on ABI 3130 DNA sequencers (Applied Biosystems Inc., Foster City,

California).

BLASTN, Molecular Dataset, Sequence Alignment, and Phylogenetic Analysis

Following sequence assembly and removal of primer sequences, I conducted general BLASTN searches (www.ncbi. nlm.nih.gov/blast/Blast.cgi) of the full length viral

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MCP gene sequence (Altschul et al. 1997) and combined viral taxa identified by these analyses with taxa from recent comprehensive phylogenetic analyses (Kurita and

Nakajima 2012; Waltzek et al. 2012) to build the final dataset. I performed sequence

alignments using Mafft 5.8 (Katoh et al. 2005) followed by minor manual adjustments in

ClustalW (Thompson et al. 1994). The E-INS-I alignment strategy was used with the following parameters: scoring matrix (BLOSUM62), gap open penalty (1.53), and offset value (0). The final sequence was trimmed to the first conserved nucleic acid at the 3’ end of the MCP sequence due to incomplete data available for certain important taxa.

The aligned dataset was imported into jModelTest version 0.1.1 ((Guindon and Gascuel

2003; Posada 2008) and the Akaike information criterion (AIC) was used to select a

best-fit model of evolution for phylogenetic analysis. Maximum likelihood analyses were

conducted using MEGA version 5 (Tamura et al. 2011) with 1000 bootstrap replicates

selected for determining node support.

Results

Histopathology

Histopathologic examination revealed multiple areas of necrosis with pyknotic

cellular debris and an associated infiltration of lymphocytes in the renal hematopoietic

tissue, liver, and spleen. Lysis of the epithelial cells of renal tubules was also noted

(Figure 4-1, 4-2). Numerous cytomegalic cells containing granular basophilic

intracytoplasmic inclusions were commonly observed in the renal interstitium and

glomeruli of both specimens (Figure 4-1). Similar cytomegaly with associated inclusions

were also observed within the heart, spleen, liver, gill lamellae, the lamina propria and

submucosa of the esophagus, stomach, and intestine (Figure 4-3). The presence of

cytomegalic cells with intracytoplasmic inclusions and necrosis in this study were similar

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to the histological lesions reported in Banggai cardinalfish infected with megalocytivirus

(Weber et al 2009).

Transmission Electron Microscopy

TEM revealed numerous icosahedral virus particles arranged in paracrystalline arrays consistent with an iridovirus within the cytoplasm of enlarged cardiac myocytes

(Figure 4-4A, B, C, D). The observed viral assembly sites appeared to disrupt the myocyte myofibrils (Figure 4B). Virus particles were naked with an electron-dense nucleic acid core surrounded by a translucent zone and an outer nucleocapsid layer

(Figure 4-4C, D). The average diameter of virus particles, measured from apex to apex was 158 nm (n = 50, SD = 7.5 nm), and from side to side was 128 nm (n = 50, SD = 5.6 nm). Virus particles identified in the cytoplasm of cytomegalic cells were consistent with

BCIV in size, icosahedral shape, and intracytoplasmic location of virions (Weber et al

2009).

Sequencing, BLASTP, Molecular Dataset, and Phylogenetic Analysis

The sequenced PCR amplicons generated in this study ranged in size from 121 -

214 bp (Table 4-1) and when assembled resulted in 1,470 contiguous bp of the viral genome that contained the full length major capsid protein sequence (1,362 bp). The

BLASTN search of the viral MCP sequence from the infected batfish revealed highest sequence identity with ISKNV genotype 2 megalocytiviruses including the Banggai cardinalfish iridovirus (BCIV; GenBank accession # AB669096; 100%) and the marbled sleepy goby iridovirus (MSGIV; GenBank accession # HM067835; 99%). The final aligned MCP dataset after truncation of the 3’ end of the sequence contained 1,357 nucleic acid characters for 91 taxa. jModelTest identified the TrN+G model to be the most suitable model for phylogenetic analyses.

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The results of the phylogenetic analysis of the MCP gene in this study are consistent with recent phylogenetic analyses (Kurita and Nakajima 2012; Waltzek et al.

2012) supporting the recognition of 4 species within the genus Megalocytivirus (Figure

4-5). The Maximum Likelihood analysis demonstrated with a high level of confidence of

the reliability that the virus from orbiculate batfish is a megalocytivirus (hereafter

referred to as the orbiculate batfish iridovirus, OBIV) and the sister group to the Banggai

cardinalfish (BCIV) which together form the sister group to marbled sleepy goby

iridovirus (MSGIV). These three viruses collectively form a megalocytivirus clade known

as ISKNV genotype 2 (Figure 4-5). I conclude, based on the evidence, that OBIV is identical to BCIV.

Discussion

This study adds to the growing literature on the emerging threat of megalocytiviruses to the international ornamental fish trade (Armstrong and Ferguson

1989; Anderson et al. 1993; Rodger et al. 1997; Sudthongkong et al. 2002; Weber et al.

2009; Kim et al. 2010; Yanong and Waltzek 2010). Here I present the first case of a megalocytivirus outbreak in orbiculate batfish based on histologic, ultrastructural, and genetic evidence. Infected cytomegalic cells displayed basophilic granular intracytoplasmic inclusions in various tissues similar to previous reports including the report of megalocytivirus infection in Banggai cardinalfish (Gibson-Kueh et al. 2003;

Weber et al. 2009; Yanong and Waltzek 2010; Zhang et al. 2011; Waltzek et al. 2012).

The size, icosahedral shape, and intracytoplasmic location of virions is also consistent with reports of megalocytivirus infections in tropical freshwater and marine ornamental fishes, including Banggai cardinalfish (Paperna et al. 2001; Sudthongkong et al. 2002;

Gibson-Kueh et al. 2003; Weber et al. 2009).

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The OBIV full length major capsid protein sequence was found to be identical to the only other megalocytivirus reported from a marine ornamental fish species, the

Banggai cardinalfish iridovirus (Weber et al. 2009). Not surprisingly, the phylogenetic analysis revealed these viruses cluster together as members of the ISKNV genotype 2 clade (Figure 4-5). Interestingly, concurrent with the orbiculate batfish mortality event, recently imported wild Banggai cardinalfish also died at the same public aquarium; they were found to be positive for BCIV by histopathology and PCR (M. Stidworthy and T. B.

Waltzek, unpublished data). Taken together with the genetic evidence, these data

suggest that OBIV and BCIV are the same virus. This agent may be capable of infecting

other species, similar to what has been reported for other megalocytivirus species

(Kurita and Nakajima 2012).

It cannot be determined whether the imported wild orbiculate batfish and Banggai

cardinalfish spread the virus to each other following importation into the United Kingdom

or whether these species had already acquired the virus in the wild or at the export

facilities. Although the Banggai cardinalfish is restricted to a small region in Indonesia

known as the Banggai archipelago, the orbiculate batfish occurs sympatrically and both

species are sometimes cultured on the same Indonesian ornamental fish farms for sale

in international ornamental fish markets (T. B. Waltzek, University of Florida, personal

communication). Future surveillance efforts are needed to determine where marine

ornamental fishes become infected with megalocytiviruses as well as the overall impact

on international trade (e.g. frequency of epizootics and number of susceptible species).

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Table 4-1. Primers used to amplify fragments of the major capsid protein (MCP) gene sequence from the orbiculate batfish iridovirus (OBIV). Primers Orientation Sequences (5’-3’) Amplicon size (base pairs)* MCPF1 Forward CTGTTGGTCTTGCTGAGTGC 158 BeginR Reverse CCGTACAAGTGGGTCTCCAT Angelnew1F Forward GTTCATCGACATCTCCGCGT 177 Angelmcp1R Reverse GCCACCGTGACACTAAACTC Angelmcp2fb Forward GGGGTGGCGACTACCTCATT 194 Angelmcp2R Reverse CCAGGTCGTTAAATGACACCG Angelmcp3F Forward CAGCTACATTCGCTGGTGCGAC 150 Angelmcp3R Reverse GCATGCCAATCATCTTGT SmallgapF Forward CTGGAACGCCTGCATGAT 124 SmallgapR Reverse ATAGTCTGGCCGTTGGTGAT MegalofixedF Forward ACAAGATGATTGGCATGCG 166 MegalofixedR Reverse TTGAAGTGGATGCGCACCT Biggap1Fa Forward GGCGTTGCCTACTGTGTCTC 163 Big1Ra Reverse CAGGGTGACGGTTGATATGG Biggap1Fb Forward GGACCTGCTCATCAGCCAGAG 165 Angelnew1R Reverse CTACGACTAGACTGGGCCA Angelnew2F Forward CTGACAAGCGAGGAGCGTG 214 Angelnew2R Reverse GGGGACTGGCCGCGGTGTAG Angelnew3F Forward TCACCCACCGCAACGTGC 188 Angelnew3R Reverse GGGCGCAAAGTAGTAGG Biggap2Fa Forward GCTCCACCAGATGGGAGTAG 153 Biggap2Ra Reverse GACAGGCGGCCGTAGTTG Biggap2Fb Forward GGACATGGGCAATATCAACC 132 Biggap2Rb Reverse GTGTAGCCGGAGCCGTTG Angelnew5F Forward GGACAATGCAAAGACCA 145 ISKNVMCPF Reverse TTACAGGATAGGGAAGCCTGC EndF Forward GGTCAAGTTTGAAAACCCGA 121 MCPR4 Reverse CATAGCTACCAGACACACGG *Amplicon size includes primers.

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Figure 4-1. Kidney from infected orbiculate batfish with intracytoplasmic basophilic iridovirus inclusions in the interstitium and glomeruli (arrows). H&E stain. (Bar = 20 μm). Photo courtesy of Preeyanan Sriwanayos.

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Figure 4-2. Heart from infected orbiculate batfish with abundant intracytoplasmic basophilic iridovirus inclusions (arrows). H&E stain. (Bar = 50 μm). Photo courtesy of Mark Stidworthy.

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A

B

C

Figure 4-3. Intracytoplasmic basophilic inclusions in infected orbiculate batfish (arrows). H&E stain. A. gill; B. esophagus; C. intestine (Bar = 20 μm). Photos courtesy of Preeyanan Sriwanayos.

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A B

C D

Figure 4-4. Transmission electron photomicrographs of iridoviral infected cells in the heart of an orbiculate batfish. A) cluster of virus particles in the cytoplasm of an infected myocyte (circle), Bar = 10 µm; B) higher magnification inset from (A) revealing numerous virus particles disrupting myocyte myofibrials (arrows) , Bar = 1 µm; C) Virions arranged in a paracrystaline array, Bar = 500 nm; D) higher magnification revealing the naked icosahedral virus shape and the electron-dense nucleic acid core surrounded by a pale zone and an outer nucleocapsid layer of moderate electron-density.( Bar = 200 nm). Photos courtesy of Preeyanan Sriwanayos.

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Figure 4-5. Phylogenetic analysis of the megalocytivirus MCP. Phylogram depicting the relationship of the orbiculate batfish iridovirus (OBIV) to representatives from each of the 4 Megalocytivirus species (RSIV, ISKNV, TRBIV, TSIV) in the family Iridoviridae, based on the sequence of the major capsid protein (1357 nucleic acid characters including gaps). Numbers above or below each node represent bootstrap support of the Maximum Likelihood analysis. Branch lengths are based on the number of inferred substitutions, as indicated by the scale. The red arrow shows the position of OBIV. Accession numbers, host, and location of isolation are attached to each operational taxonomic unit. Photo courtesy of Preeyanan Sriwanayos.

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BIOGRAPHICAL SKETCH

Preeyanan Sriwanayos was born in Chiang Rai Province, Northern region of

Thailand. After she graduated high school in 2002, Preeyanan attended the Faculty of

Veterinary Medicine, Kasetsart University, Bangkok, Thailand and received her Doctor of Veterinary Medicine in March 2008. After graduation, she received a scholarship from the Royal Thai government to pursue her master’s degree in aquatic animal health management. Preeyanan began a Master of Science program that focused on fish health management at the University of Florida, School of Forest Resources and

Conservation, Program in Fisheries and Aquatic Sciences in August 2009.

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