© 2017 natalie katherine stilwell

RANAVIRUSES IN AQUACULTURE: GENETIC DIVERSITY, IMPROVED MOLECULAR

TOOLS, AND EXPERIMENTAL ANALYSIS OF HUSBANDRY FACTORS INFLUENCING

MORBIDITY

By

NATALIE KATHERINE STILWELL

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL

OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT

OF THE REQUIREMENTS FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2017

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ACKNOWLEDGMENTS

My PhD studies were made possible by a University of Florida (UF) Alumni Fellowship,

and I am grateful to the United States Department of Agriculture National Institute of Food and

Agriculture for providing grant funding for the live sturgeon project. I would also like to thank

the UF Graduate Student Council and Veterinary Graduate Student Association for providing

travel grant funding, which made it possible to present and distribute my research at several

conferences.

I wish to extend my gratitude to my major professor, Dr. Thomas Waltzek, for his

constant dedication to the advancement of wildlife and aquatic animal virology, and for his

generosity towards his peers and students. It has been an honor working in the UF Wildlife and

Aquatic Animal Veterinary Disease Laboratory (WAVDL) as his graduate student. I would also

like to thank my advisory committee members, Drs. Lisa Farina, Salvatore Frasca Jr., Marco

Salemi, and James Wellehan, not only for providing insight in their respective fields but for their

helpful life and career advice. I am grateful to the faculty, staff, and students at UF WAVDL

who contributed time and effort to my PhD project over the past four years, including: Linda

Archer, Allison Cauvin, Abigail Clark, Sieara Claytor, Dr. Galaxia Cortes-Hinojosa, Dr. Jason

Ferrante, Jared Freitas, Jaime Haggard, Rachel Henriquez, Dr. Shohreh Hesami, Kamonchai

Imnoi, Jessica Jacob, Samantha Koda, Nelmarie Landrau-Giovanetti, Dr. Denise Petty, Dr. Maria

Robles, Elizabeth Scherbatskoy, Dr. Preeyanan Sriwanayos, Dr. Kuttichantran Subramaniam,

Patrick Thompson, Luke Trimmer-Smith, and Dr. Pedro Viadanna. Working in the UF WAVDL

lab among such enthusiastic and kind individuals has truly been a highlight of my career. I am

also grateful to the staff within the UF College of Veterinary Medicine (CVM) for their

assistance, including Kia Hendrix, Erin Sanetz, Debbie Couch, and Sally O’Connell.

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The live sturgeon work would not have been possible without the aid of several

individuals. I am grateful to Jeffrey Powell, Diane von Eschen, and personnel at the Gavin’s

Point Fish Hatchery for their generous donation of pallid sturgeon, and to Lacey Hopper and the

USFWS Bozeman Fish Health Center for performing health assessments. I would like to extend

thanks to Dr. Andrew Kane and Ross Brooks at the UF Aquatic Pathobiology Laboratory for

their guidance and help with aquatic life support systems, and to Dr. Brian Stacy for the use of

necropsy facilities. I am grateful to James Colee of the UF Department of Statistics for his

assistance with statistical analyses, and to Dr. Vsevolod Popov of the University of Texas

Medical Branch for processing electron microscopy samples. I would also like to thank the

members of the UF CVM diagnostic laboratories, particularly Melissa Brown, for their help

processing histological and microbiological samples. Several members of the international

ranavirus community generously contributed resources for the enhancement of my research.

These individuals include: Dr. Ellen Ariel, Dr. Joy Becker, Dr. Paul Hick, Dr. Riikka

Holopainen, Dr. James Jancovich, Dr. Diana Jaramillo, Dr. Somkiat Kanchanakhan, Dr. Debra

Miller, Dr. Niels Jorgen Olesen, Dr. Allan Pessier, Dr. Jaree Polchana, Dr. Anna Toffan, Dr.

Steven van Beurden, and Dr. Richard Whittington.

Finally, I want to extend my utmost gratitude to my family, particularly my parents Dr.

Munro and Katherine Steckler, my brother Alec Steckler, and my dear husband Dr. Justin

Stilwell, for their love and patience as I pursue my career dreams.

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TABLE OF CONTENTS

page

ACKNOWLEDGMENTS .............................................................................................................. 3

LIST OF FIGURES ........................................................................................................................ 8

ABSTRACT .................................................................................................................................. 10

CHAPTER

1 LITERATURE REVIEW ......................................................................................................... 12

Introduction to Ranaviruses ...................................................................................................... 12 Summary ................................................................................................................................... 24

2 REPEATED DETECTIONS OF RANAVIRUSES IN AQUACULTURE ............................. 27

Introduction ............................................................................................................................... 27

Materials and Methods .............................................................................................................. 29 Results ....................................................................................................................................... 35 Discussion ................................................................................................................................. 37

3 VALIDATION OF A TAQMAN REAL-TIME QUANTITATIVE PCR FOR THE

DETECTION OF RANAVIRUSES ......................................................................................... 78

Introduction ............................................................................................................................... 78 Materials and Methods .............................................................................................................. 80 Results ....................................................................................................................................... 84

Discussion ................................................................................................................................. 86

4 THE EFFECT OF WATER TEMPERATURE ON FROG VIRUS 3 DISEASE IN

HATCHERY-REARED PALLID STURGEON (SCAPHIRHYNCHUS ALBUS) ................... 99

Introduction ............................................................................................................................... 99

Materials and Methods ............................................................................................................ 101 Results ..................................................................................................................................... 108 Discussion ............................................................................................................................... 112

5 CONCLUDING STATEMENTS ........................................................................................... 129

LIST OF REFERENCES ............................................................................................................ 131

BIOGRAPHICAL SKETCH ...................................................................................................... 146

6

LIST OF TABLES

Table page

1-1 Characteristics for members of the family Iridoviridae and genus Ranavirus. .................26

2-1 Conserved iridoviral/ranaviral genes and their predicted functions if known...................43

2-2 Documented ranavirus detections in fish hosts. ................................................................45

2-3 Fully sequenced ranavirus genomes used for phylogenomic analyses within this

chapter…. ...........................................................................................................................46

2-4 In vitro growth chracteristics and ultrastructural analyses for the 12 aquaculture isolates

included in this chapter. .....................................................................................................47

2-5 Genome characteristics for the eight fish ranaviruses sequenced in this chapter ..............48

2-6 Predicted open reading frames for the fathead minnow ranavirus isolate (FHMRV).. .....49

2-7 Predicted open reading frames for the northern pike ranavirus isolate (NPRV) ...............52

2-8 Predicted open reading frames for the 2001 pallid sturgeon ranavirus isolate

(PSRV01)…. ......................................................................................................................55


(PSRV09)…. ......................................................................................................................58


(PSRV13)…. ......................................................................................................................61


(PSRV15)…. ......................................................................................................................64

2-12 Predicted open reading frames for the Russian sturgeon ranavirus isolate (RSRV) .........67

2-13 Predicted open reading frames for the white sturgeon ranavirus isolate (WSRV) ............70

3-1 Quantitative PCR assays developed for the detection of ranaviruses ................................89

3-2 Primers and probes designed against the ranavirus major capsid protein (MCP) gene for

development of the plasmid standard and for use in the diagnostic assay.........................90

3-3 Panel of ranaviruses used for primer and probe design and/or validation of the TaqMan

qPCR assay ........................................................................................................................91

3-4 Results for the TaqMan qPCR assay on fish tissue homogenates with EHNV infection

status determined by virus isolation and confirmed by PCR .............................................93

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3-5 Inter-assay variability (reproducibility) of the pan-ranavirus qPCR across twelve

experiments (plates) at the WAVDL .................................................................................94

4-1 Study 1 results summary..................................................................................................115

4-2 Study 2 results summary..................................................................................................116

4-3 Log mean (± SE) qPCR viral copy number for external and internal tissue homogenates

in the warmwater and coldwater treatments over the 28 d study (n=4 fish per treatment

per day) ............................................................................................................................117

4-4 RNAscope® ISH results from cold- and warmwater fish corresponding with the highest

average TaqMan qPCR viral load per sampling date ......................................................118

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LIST OF FIGURES

Figure page

2-1 Cytopathic effect (CPE) typical of the 12 ranavirus isolates characterized within this

chapter ................................................................................................................................73

2-2 Transmission electron microscopy photomicrographs illustrating typical ranavirus virion

morphogenesis ...................................................................................................................74

2-3 Maximum likelihood (ML) cladogram and phylogram depicting the relationships of 28

ranaviruses based on their aligned genomes ......................................................................75

2-4 Whole-genome alignments of 28 ranaviruses displaying 6 locally collinear blocks to

indicate genome arrangement ............................................................................................77

3-1 Aligned partial (97 bp) major capsid protein (MCP) sequences for 36 ranaviruses

illustrating the in silico specificity of the qPCR primers (RanaF1 and RanaR1) and

TaqMan probe (RanaP1) ....................................................................................................95

3-2 Quantification of a standard curve for Frog virus 3 using the TaqMan real-time

polymerase chain reaction (qPCR) assay ...........................................................................96

3-3 Aligned partial (94 bp) major capsid protein (MCP) sequences for 36 ranaviruses

illustrating in silico specificity of the SYBR green qPCR primers developed by Jaramillo

and colleagues ....................................................................................................................98

4-1 Experimental tank design for studies 1 and 2 ..................................................................119

4-2 Study 1 survival curve. ....................................................................................................120

4-3 Representative photos of gross pathology associated with FV3 disease at 23°C in studies

1 and 2 ..............................................................................................................................121

4-4 Comparison of log mean (±SE) qPCR copy number for external (Ext.) and internal (Int.)

tissue homogenates in the warmwater and coldwater treatments over the 28 d study. ...122

4-5 Prevalence of FV3 infection in study 2 following bath exposure at 17°C. .....................123

4-6 Prevalence of FV3 infection in study 2 following bath exposure at 23°C ......................124

4-7 Hematoxylin and eosin (H&E) and RNAscope® in situ hybridization (ISH) results from

the spleen of a warmwater exposed sturgeon sampled on day 7 of study 2 ....................125


the gill of a warmwater exposed sturgeon sampled on day 7 of study 2 .........................126


the posterior kidney of a warmwater exposed sturgeon sampled on day 7 of study 2. ...127

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the heart, including pericardial lymphomyeloid tissue, from a warmwater exposed

sturgeon sampled on day 7 of study 2 ..............................................................................128

10

Abstract of Dissertation Presented to the Graduate School

of the University of Florida in Partial Fulfillment of the

Requirements for the Degree of Doctor of Philosophy

RANAVIRUSES IN AQUACULTURE: GENETIC DIVERSITY, IMPROVED MOLECULAR

TOOLS, AND EXPERIMENTAL ANALYSIS OF HUSBANDRY FACTORS INFLUENCING

MORBIDITY

By

Natalie Katherine Stilwell

August 2017

Chair: Thomas B. Waltzek

Major: Veterinary Medical Sciences

Ranaviruses are globally emerging pathogens negatively impacting wild and cultured

fish, amphibians, and reptiles. Since their first discovery in the 1960s, ranaviruses have been

detected with increasing frequency in farmed freshwater and marine fish species, including

several species of federally endangered, US-farmed sturgeon (e.g., pallid Scaphirhynchus albus,

lake Acipenser fulvescens, and Russian A. gueldenstaedtii). This dissertation outlines several

research projects aimed at furthering our understanding of ranaviral infections in fish. Chapter 1

provides an overview of fish ranaviruses emphasizing their taxonomy, biology, pathology,

diagnostics, and factors influencing their impact on aquaculture and wild stocks. Viral

characterization and phylogenomic analyses were used to examine biologic and epidemiologic

trends in aquaculture (Chapter 2). The results support the fact that ranaviruses originated in fish

prior to spreading into other ecothermic vertebrates. Furthermore, some ranavirus species such as

Frog virus 3 exhibit low host specificity infecting amphibians, reptiles, and fish on nearly every

continent. The global emergence of ranaviruses underscores the role international trade has

played in the dissemination of these pathogens and the need for diagnostic tools capable of

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detecting them. Genomic sequences generated for fish ranaviruses (Chapter 2) were combined

with previously available ranavirus genomic sequences to design improved molecular assays for

the diagnosis and characterization of ranaviruses, including a pan-ranavirus quantitative real-

time TaqMan PCR (Chapter 3) and an in situ hybridization assay using RNAscope® technology

(Chapter 4). The qPCR assay, which was validated against an extensive pool of 36 ranavirus

isolates, detected the majority of isolates and may serve as a useful, single-step diagnostic tool

for both ranavirus surveillance and research purposes. Finally, bath FV3 challenges were

conducted at 17˚C and 23˚C to examine the effect of water temperature on ranaviral disease in

hatchery-reared young-of-year pallid sturgeon (Chapter 4). The clinical signs, gross and

microscopic pathology, viral load (qPCR), viral titer (virus isolation), and cumulative mortality

clearly revealed that elevated water temperature significantly increases disease in juvenile pallid

sturgeon. These data suggest that temperature manipulation may serve as an effective

management tool for sturgeon hatcheries afflicted with ranavirus epizootics.

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CHAPTER 1

LITERATURE REVIEW

Introduction to Ranaviruses

The family Iridoviridae, along with six other virus families (Ascoviridae, Asfarviridae,

Marseilleviridae, Mimiviridae, Phycodnaviridae, and Poxviridae), are categorized as

Nucleocytoplasmic Large DNA Viruses (NCLDVs). NCLDVs are unique among DNA viruses

for their large genomes and ability to replicate either partially (Ascoviridae, Iridoviridae) or

completely (Marseilleviridae, Mimiviridae, Phycodnaviridae, Poxviridae) in the cytoplasm of

host cells (Yutin et al. 2009, Yutin and Koonin 2012). The family Iridoviridae has recently been

reorganized into the subfamily Alphairidovirinae that infect ectothermic vertebrates (genera:

Ranavirus, Lymphocystivirus, and Megalocytivirus) and subfamily Betairidovirinae that infect

invertebrates such as insects and crustaceans (genera: Iridovirus and Chloroiridovirus) (Chinchar

et al. 2017a). Lymphocystiviruses and megalocytiviruses infect bony fish, whereas members of

the genus ranavirus infect bony fish, amphibians, and reptiles.

Characteristics for members of the family Iridoviridae and genus Ranavirus are outlined

in Table 1-1. Iridoviruses possess nucleocapsids with icosahedral symmetry (120-350 nm in

diameter) and an electron-dense core of double-stranded DNA observed within the cytoplasm of

infected cells (Chinchar et al. 2017a). Ranavirus virions may be naked or enveloped (Braunwald

et al. 1979), with enveloped virions averaging 160-200 nm in diameter (Chinchar et al. 2017a).

The linear, double-stranded DNA ranavirus genome is circularly permuted and terminally

redundant (Goorha and Murti 1982). Ranavirus genomes range from 104-140 kbp with 73-139

open reading frames (ORFs) and 48-57% G+C content (Jancovich et al. 2015, Chinchar et al.

2017a). As NCLDVs, some ranaviruses possess both nuclear and cytoplasmic phases within the

13

life cycle, with viral transcription occurring in the nucleus of the host cell prior to viral capsid

formation and final virion assembly in the host cytoplasm.

Seven ranaviral species are recognized by the International Committee on Taxonomy of

Viruses (ICTV) including: Ambystoma tigrinum virus (ATV), Bohle iridovirus (BIV), Epizootic

hematopoietic necrosis virus (EHNV), European catfish virus (ECV), Frog virus 3 (FV3),

Santee-Cooper ranavirus (SCRV), and the recently added Singapore grouper iridovirus (SGIV)

(Chinchar et al. 2017a). Certain ranaviruses (e.g., FV3, BIV) possess among the lowest known

specificity for any double-stranded DNA virus, infecting three classes of ectothermic vertebrates:

Osteichthyes, Reptilia, and Amphibia. Due to their expanding host and geographic ranges, as

well as high associated morbidity and mortality rates, all amphibian ranavirus detections and

EHNV in fish hosts are notifiable to the World Organization for Animal Health (OIE 2016a,b).

The apparent rapid expansion of ranaviruses into novel fish, amphibian, and reptile hosts

around the world underscores the need for an improved understanding of the phylogenetic and

taxonomic relationships of these emerging pathogens (Duffus et al. 2015, Jancovich et al. 2015).

Increased awareness and surveillance for ranaviruses as well as the advent of high-throughput

sequencing technologies have resulted in the discovery and genetic characterization of a number

of fish ranaviruses yet to be classified by the ICTV (Ariel et al. 2016, Holopainen et al. 2016,

Subramaniam et al. 2016b). Ranavirus phylogenetics has historically relied upon the analysis of

a few conserved genes (e.g., major capsid protein, MCP) (Mao et al. 1997, Tidona et al. 1998,

Hyatt et al. 2002, Marsh et al. 2002, Holopainen et al. 2009). However, the MCP alone is

insufficient to resolve all branches within the growing ranavirus phylogenetic tree (Duffus and

Andrews 2013, Jancovich et al. 2015). Therefore, recent advances in sequencing efficiency (i.e.,

Next Generation Sequencing approaches) have facilitated phylogenetic analyses based upon 26

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genes conserved among all iridoviruses and an additional 27 conserved ranaviral genes (Eaton et

al. 2007, Jancovich et al. 2015, Ariel et al. 2016, Hick et al. 2016, Holopainen et al. 2016,

Subramaniam et al. 2016b, Claytor et al. 2017). Other criteria commonly used to classify

iridoviruses include: restriction endonuclease fragment length polymorphism (RFLP) profiles,

virus protein profiles, genomic organization, virion morphology, in vitro growth characteristics,

antigenic properties, geographic and/or host range (Jancovich et al. 2015).

Ranaviral Detections in Fish

Ranaviruses have been detected in a variety of teleost fishes with the majority of cases

occurring in aquaculture. Notable cases in farmed fishes include multiple species of federally

endangered and threatened sturgeon (family Acipenseridae). Hatchery-reared young-of-year

pallid sturgeon (Scaphyrhynchus albus) experienced high mortality and severe pathology at a

Missouri state hatchery during epizootics in 2001, 2009, 2013, and 2015 (Waltzek et al. 2014,

Chapter 2). Ranavirus infection has also caused high mortality in farmed white (Acipenser

transmontanus) and Russian sturgeon (Acipenser gueldenstaedtii) in California in 1998 and

Georgia in 2004, respectively (Waltzek et al. 2014).

Although FV3 infections in cultured and wild fish have been reported on several

occasions, the role of the virus in disease remains unclear in several of these detections. For

example, the first detection of FV3 in fish occurred in a single moribund three-spined stickleback

(Gasterosteus aculeatus) during a sympatric outbreak in a wild population of moribund red-

legged frogs (Rana aurora, Mao et al. 1999a). The FV3 strains infecting these two hosts were

genetically identical; however, the role of the ranavirus in disease was complicated as both hosts

were co-infected with other pathogens (e.g., Myxobolus sp. in the stickleback and multiple

species of bacteria in the frogs). In 2008 in a natural pond in Japan, a die-off of invasive North

American bullfrogs (Rana catesbeiana) was attributed to FV3, with cohabitating asymptomatic

15

cyprinids (Gnathopogon spp.) also testing positive by PCR (Une et al. 2009). In Thailand, an

FV3-like ranavirus associated with epizootics in ranaculture facilities was also attributed to an

outbreak involving farmed marbled sleeper goby (Oxyeleotris marmoratus) (Prasankok et al.

2005). Incidental FV3 isolations have also occurred in several asymptomatic US-farmed fishes

including fathead minnow (Pimephales promelas) and northern pike (Esox lucius) (Waltzek et al.

2014). Ranavirus detections in asymptomatic individuals suggest certain fish species may serve

as dead-end hosts or subclinical reservoirs and potentially transmit ranavirus to susceptible

populations. Furthermore, transmission of ranaviruses among different host species or even

among classes of ectothermic vertebrates has been demonstrated experimentally, raising

concerns about the impact of these emerging pathogens on aquaculture and aquatic ecosystems

(Whittington et al. 1994, 1999, Brenes et al. 2014).

In addition to FV3, several other ranavirus species have been detected in fish. Epizootic

hematopoietic necrosis virus (EHNV), an OIE-notifiable ranavirus detected only in Australia,

results in marked disease within wild redfin perch (Perca fluviatilis) populations and less severe

disease in aquacultured rainbow trout (Oncorhynchus mykiss) (Langdon et al. 1986, 1988,

Whitttington et al. 1994, 1996). Experimental challenges performed by Langdon (1989) and

Becker et al. (2013) revealed EHNV susceptibility in five additional fish species including:

Murray-Darling rainbowfish (Melanotaenia fluviatilis), dewfish (Tandanus tandanus), eastern

mosquitofish (Gambusia holbrooki), silver perch (Bidyanus bidyanus), and Macquarie perch

(Macquaria australasica). Bohle iridovirus (BIV), a ranavirus species initially isolated from

ornate burrowing frogs (Limnodynastes ornatus) in Australia (Speare and Smith 1992), was more

recently the suspected cause of neurologic disease in cultured Nile tilapia (Oreochromis

niloticus) (Ariel and Owens 1997). Barramundi fingerlings (Lates calcarifer), a host known to be

16

susceptible to BIV, were fed infected tilapia and subsequently developed neurologic symptoms

similar to those previously seen in BIV-infected barramundi. However, viral isolation from

moribund tilapia was unsuccessful and confirmation of BIV infection by molecular diagnostic

methods was not performed.

Multiple ranaviruses have been isolated in farmed and wild European fishes. Short-finned

eel ranavirus (SERV) was detected in short-finned eel (Anguilla australis) imported from New

Zealand into Italy (Bovo et al. 1999, Bang Jensen et al. 2009). Two nearly identical ranaviruses

known as European catfish virus (ECV) and European sheatfish virus (ESV) have impacted the

production of brown and black bullhead catfish (Ameiurus nebulosus and melas) and sheatfish

(Silurus glanis) (Ahne et al. 1989, 1991, Pozet et al. 1992). Other ranaviruses from fishes in

Europe include the pike-perch iridovirus (PPIV) isolated from overtly healthy Finnish pike-perch

fingerlings in 1995 (Sander lucioperca, Tapiovaara et al. 1998), cod iridovirus (CoIV) as the

cause of ulcerative disease in wild cod (Gadus morhua) in Denmark (Jensen et al. 1979, Ariel et

al. 2010), and Ranavirus maxima (Rmax) in farmed turbot fry (Scophthalmus maximus) in

Denmark (Ariel et al. 2010).

Two fish ranaviruses, the Singapore grouper iridovirus (SGIV) and Santee-Cooper

ranavirus (SCRV), represent the most divergent species and their inclusion in the genus has been

questioned (Hyatt et al. 2000, Whittington et al. 2010, Jancovich et al. 2015); however, they are

included here given their significance in aquaculture. One clade includes strains of SCRV such

as largemouth bass virus (LMBV), doctor fish virus (DFV), and guppy virus 6 (GV6) (Hedrick

and McDowell 1995, Mao et al. 1999b, Plumb et al. 1996). Historically, SCRV was believed to

primarily affect North American largemouth bass (hence the name LMBV) including an early

epizootic within the Santee-Cooper Reservoir in South Carolina, USA. However, natural and

17

experimental infections have now confirmed SCRV in >16 fish species including cultured koi

(Cyprinus carpio) in India (George et al. 2015) and the invasive northern snakehead (Channa

argus) within the Chesapeake Bay, USA (Groocock et al. 2008, Iwanowicz et al. 2013). Whether

LMBV is pathogenic to largemouth bass remains controversial, and it appears that several hosts

including other centrarchids (e.g., smallmouth bass Micropterus dolomieu, bluegill Lepomis

macrochirus, redear sunfish Lepomis microlophus) experience asymptomatic infections

(Getchell and Groocock 2017).

The other divergent ranavirus clade includes SGIV, which was recently accepted as the

seventh ranavirus species by the ICTV (Chinchar et al. 2017a), and grouper iridovirus (GIV).

These two nearly identical ranaviruses affect multiple cultured grouper and non-grouper species

in Asia (Duffus et al. 2015). The SGIV was originally nicknamed Sleepy Grouper Disease based

on the clinical signs associated with mass mortality events in brown-spotted grouper

(Epinephelus tauvina) in Singapore in 1994 and 1998 (Chua et la. 1994, Qin et al. 2003). The

GIV was characterized from mortalities observed in Taiwanese yellow grouper (Epinephelus

awoara) beginning in 1998 (Murali et al. 2002). The SGIV/GIV have recently been isolated in

other cultured fishes in Taiwan including largemouth bass (Micropterus salmoides) and

barramundi (Huang et al. 2011).

Ranavirus Pathology and Transmission

Although clinical presentation may vary among host species, ranaviruses generally

induce systemic disease with external and internal hemorrhage, necrosis, edema (particularly in

amphibians and chelonians), behavioral changes (e.g., lethargy, anorexia, erratic movement), and

significant morbidity/mortality (Miller et al. 2015, Rijks et al. 2016). Histopathological changes

associated with infection can be quite similar among species and have been described for both

natural and experimental infections. Ranaviral infections are typically systemic, with

18

hemorrhage, edema, and necrosis of affected internal and external tissues noted particularly in

epithelium, vascular endothelium, and hematopoietic tissues (Miller et al. 2015). Hemorrhage

and necrosis may be focal, multifocal, or diffuse (Whittington et al. 2010). Tissue necrosis is

indicated by pyknosis, karyorrhexis, karyolysis, absent nuclei, eosinophilic homogeneous

cytoplasm, loss of cell adherence to basement membranes, and cellular rupture with loss of

integrity (Zachary and McGavin 2012). Basophilic to amphophilic cytoplasmic inclusions are

characteristic of ranavirus infected cells and may range from absent or rare (Stöhr et al. 2013) to

widespread (Docherty et al. 2003, personal observation).

Virtually all tissues are targeted in systemic ranavirus infections including: skin, muscle,

heart, blood vessels, lymphoid tissue (e.g., thymus), spleen, liver, kidney, pancreas,

gastrointestinal tract (spanning from the oropharynx to the large intestine), neural tissue (e.g.,

neuroepithelium, brain and meninges), bone, adipose tissue, and the respiratory tract including

the trachea, lung, and gill (Docherty et al. 2003, Miller et al. 2015). Hyperplasia and necrosis

have been noted in the gill epithelium of fish with occasional fusion of the secondary lamellae

(Whittington et al. 2010). The hematopoietic tissues (e.g., kidney, spleen, liver, bone marrow)

are among the most commonly targeted tissues and are generally recommended for ranavirus

testing regardless of host class (Miller et al. 2015). Both hematopoietic and non-hematopoietic

areas of the liver, spleen, and kidney may exhibit degeneration/necrosis and basophilic to

amphophilic cytoplasmic ranaviral inclusions. In the kidney, glomeruli and renal tubular

epithelial cells may both be affected (Balseiro et al. 2010). In the liver, hepatocytes,

melanomacrophage aggregates, and sinusoids are typically affected.

Ranaviral disease in the skin may manifest as erosive to ulcerative lesions with the

underlying muscle layers displaying hemorrhage and necrosis of fibers. An ulcerative dermatitis

19

has been observed in rainbow trout (Oncorhynchus mykiss, Reddacliff and Whittington 1996)

infected with EHNV and pallid sturgeon (Scaphirhynchus albus) infected with FV3 (Waltzek et

al. 2014). Amphibians appear to have a strong cutaneous component to disease; specifically,

degeneration and ulceration of the epidermis has been documented in several species including

the common frog (Rana temporaria, Cunningham et al. 2007) and Chinese giant salamander

(Andrias davidianus, Geng et al. 2011). Affected amphibians may or may not display cutaneous

edema (Bollinger et al. 1999). Skin lesions in reptiles and amphibians are often chronic in nature

and affected animals may lack systemic disease (Cunningham et al. 2008). Ranaviral-induced

cutaneous disease has been recorded in squamate reptiles including: brown anole (Anolis sagrei),

green anole (Anolis carolinensis), Asian glass lizard (Dopasia gracilis), green iguana (Iguana

iguana), and central bearded dragon (Pogona vitticeps) (Stöhr et al. 2013).

Lesions of the gastrointestinal mucosa range from erosive to ulcerative (Bollinger et al.

1999). Interestingly, edema and necrosis of the swim bladder have been noted in EHNV

(Reddacliff and Whittington 1996) and LMBV (Zilberg et al. 2000) infections, with the latter

also causing a fibrinous peritonitis along the serosal surface of all coelomic organs. A neural

component to disease has been observed in a few amphibian cases, including neuroepithelial

necrosis (Docherty et al. 2003) and vestibular hemorrhage and necrosis associated with spinning

and erratic swimming in anurans (Mazzoni et al. 2015). Finally, FV3 DNA has been detected in

a granulomatous ocular lesion of a wild American bullfrog (Burton et al. 2008).

Several routes have been proposed for ranaviral transmission, including direct contact

with affected individuals, ingestion of infected tissues via predation or scavenging, and contact

with affected water or fomites (Langdon 1989, Reddacliff and Whittington 1996, Harp and

Petranka 2006, Brenes et al. 2014). Transmission routes are often indicated by tissue-specific

20

pathology. For example, Robert et al. (2011) observed high ranaviral DNA loads in the distal

intestine of experimentally infected African clawed frogs (Xenopus laevis) soon after infection,

suggesting adsorption of the virus through the cloaca. Only later was viral DNA detected in

several tissues throughout the body, suggesting a viremia. Severity of ranaviral infection is dose-

dependent (Brunner et al. 2005), and the route of exposure may affect the presentation and extent

of disease (Hoverman et al. 2010). Ranaviruses may persist in water and soil, even after

dessication or freezing. Largemouth bass virus (LMBV) DNA is detectable in water samples for

at least 7 days, although infectivity is mostly lost within 24 hours (Grizzle and Brunner 2003),

and FV3 is demonstrated experimentally to remain viable for more than a month in soil (Nazir et

al. 2012). Ranaviruses are also somewhat resistant to drying and freezing, which may help them

to overwinter (Bollinger et al. 1999, Johnson and Brunner 2014).

Diagnosis of Ranavirus

The OIE has outlined several recommended diagnostic methods for the isolation and

identification of ranaviruses (OIE 2016a,b). According to the OIE diagnostic manual for

ranaviral infection in amphibians (OIE 2016a), a case is suspect for ranaviral infection if the skin

and/or parenchymal tissues of an apparently healthy, moribund, or dead individual contain

histological evidence of necrosis with or without the presence of cytoplasmic basophilic

inclusion bodies. A case is confirmed when the suspect animal’s tissues or cell culture test

positive via: 1) immunoperoxidase test/stain, 2) antigen-capture ELISA, 3) PCR followed by

restriction enzyme analysis (REA) or sequencing, and/or 4) immunoelectron microscopy (tissue

only). For EHNV infections in fish (OIE 2016b), a suspect case is defined when one or more

animals demonstrates characteristic histopathology (e.g., liquefactive or coagulative necrosis)

with or without the presence of cytoplasmic inclusion bodies. Suspect EHNV cases are

confirmed via PCR (Hyatt et al. 2000) with sequencing or REA, plus one or more of the

21

following: 1) immunoperoxidase test/stain (Reddacliff and Whittington 1996); 2) antigen-capture

ELISA; 3) immunoelectron microscopy.

End-point PCR (conventional) and real-time quantitative PCR (qPCR) assays have been

developed for the detection of ranaviruses. While conventional PCR uses a set of primers

specifically designed to amplify a targeted nucleic acid region, qPCR additionally utilizes a

fluorescent reporter signal to quantify the amount of target nucleic acid in the original sample

(Green and Sambrook 2012). Two types of qPCR chemistries exist: DNA-binding dyes (e.g.,

SYBR Green) chemistries use fluorogenic dyes to bind to double-stranded DNA in a non-

sequence-specific manner, while probe-based (e.g., TaqMan) chemistries use one or more

fluorescently labeled oligonucleotides to hybridize to a specific internal sequence (Green and

Sambrook 2012). Most ranavirus-specific assays target the major capsid protein (MCP) gene

(Mao et al. 1997, Tidona et al. 1998, Hyatt et al. 2002, Marsh et al. 2002, Holopainen et al.

2009). The MCP gene is ideal for molecular diagnostic assays because it is highly conserved and

maintains stable base mutations that allow for the differentiation of species (Mao et al. 1997).

Two conventional PCR assays targeting the MCP are recommended by the OIE (2016a,b) for

diagnosis of ranaviruses. One assay allows for the differentiation of Australian ranaviral species

(EHNV and BIV) from American (FV3) and European (ECV) species by performing REA of

PCR amplicons (Marsh et al. 2002). The second conventional assay targets a 580 bp region of

the MCP which can be sequenced for ranaviral species identification (Hyatt et al. 2000). Real-

time qPCR assays have been designed to detect specific ranaviruses (Goldberg et al. 2003,

Getchell et al. 2007, Allender et al. 2013a) or a range of ranaviruses (Pallister et al. 2007,

Holopainen et al. 2011, Jaramillo et al. 2012) (Table 2-1). Of these assays, only one SYBR

22

Green qPCR assay has been rigorously validated against a number of ranaviral taxa (Jaramillo et

al. 2012).

In cases of viral identification, it is prudent and clinically helpful to distinguish infection

from disease. PCR methods used alone detect viral nucleic acid but cannot determine whether

infection is causing disease. Positive PCR results may indicate residual DNA from previous

infection, or even an asymptomatic carrier state. PCR can, however, be used in combination with

supporting diagnostic evidence to assess the extent of disease. For example, the presence of

ranavirus-suggestive histopathological changes (e.g., hemorrhage, necrosis) and/or identification

of viral nucleic acid or protein via in situ methods (e.g., in situ hybridization (ISH) and

immunohistochemistry (IHC)) are helpful indicators of productive infection and ranavirus-

associated pathology. In situ methods are particularly useful for identifying pathogens that

exhibit inconsistent and/or non-specific pathology. Ranaviruses often display non-specific

histopathological signs characterized by hemorrhage and necrosis within the epithelium, vascular

endothelium, and hematopoietic tissues. Furthermore, small basophilic to amphophilic

cytoplasmic inclusions present in ranavirus infected cells may be either rare in number or

difficult to distinguish from similar looking histological findings such as phagocytosed material

in leukocytes (Stöhr et al. 2013). Thus, ISH and IHC aid in giving histological context to

pathogens located within affected tissues.

Host, Viral, and Environmental Factors Influencing Ranaviral Disease

Currently, little is known about the host, viral, and environmental factors that influence

ranaviral disease outbreaks in aquaculture. Common environmental stressors that can increase

infectious disease episodes in production include: poor water quality, inappropriate temperature,

and malnutrition (Barton et al. 1991, Chua et al. 1994, LaPatra et al. 1996, Buentello et al. 2000,

Georgiadis et al. 2000, 2001, Drennan et al. 2005, Savin et al. 2011). Ranavirus outbreaks under

23

natural settings and experimental challenge studies have both revealed that increasing host

density facilitates transmission by increasing contact rates between animals as well as pathogen

concentrations within the water (Woodland et al. 2002, Brunner et al. 2007, Brenes et al. 2014,

Brunner et al. 2015).

Ranaviral experimental challenge studies and observations during natural outbreaks

suggest that host factors can influence ranaviral pathogenicity. Host age may contribute to

ranaviral infection, as mortality often varies between age classes of the same species (Cullen et

al. 1995, Haislip et al. 2011, Hoverman et al. 2011, Brenes et al. 2014). Typically, larval and

metamorphic amphibians are believed to be more susceptible to ranaviruses than adults. Multiple

hypotheses exist, including the assumption that increased metabolic rates in rapidly developing

juveniles may increase viral replication rates and overwhelm the host’s immune response (Robert

et al. 2005, Brunner et al. 2015). As seen in some fish species such as the redfin perch (Perca

fluviatilis), juveniles tend to congregate in shallow warm waters that are optimal for viral

replication (Whittington et al. 2010). Furthermore, experimental infections with EHNV in

rainbow trout showed highest mortality at temperatures beyond the host’s upper threshold (Ariel

et al. 2009). Similarly, increased water temperature is believed to increase mortality rates during

FV3 epizootics in North American hatchery-reared pallid sturgeon (Waltzek et al. 2014).

Finally, viral factors are believed to influence ranaviral outbreaks. Several genes related

to increased virulence and pathogenicity have been identified and studied in ranaviruses

including a viral equivalent of the eukaryotic translation initiation factor 2 (eIF-2alpha), which

contributes to synthesis of host proteins. Additional virulence genes present in some or all

ranaviruses that may aid in evading the host’s immune reponse include: RNAse-III like protein,

homologues of beta-hydroxysteroid dehydrogenase, tissue necrosis factor receptor, dUTPase,

24

cytosine methyltransferase, and US22 proteins (Grayfer et al. 2015, Claytor et al. 2017). Cell

culture studies have revealed ranavirus isolates typically have a permissive temperature range

(Miller et al. 2015). Outbreaks are often seasonal in nature (Brunner et al. 2015) and may reflect

temperature preferences of the virus in combination with temperature-dependent immune status

of the ectothermic host. Several temperature studies performed in laboratory settings have

demonstrated a significant effect of temperature on mortality (Grant et al. 2003, Allender et al.

2013b, Echaubard et al. 2014). Ultimately, a combination of viral, host, and environmental

factors likely contribute to the severity of ranaviral disease.

Summary

The emerging nature of ranaviruses, evidenced by their expanding host and geographic

ranges, has spurred a growing interest in the scientific community. As a result, a global group of

scientists, veterinarians, and citizens has formed the Global Ranavirus Consortium in recent

years to facilitate communication, collaboration, and ultimately a greater understanding of these

complex, lethal pathogens. At the Third International Symposium on Ranavirus in Gainesville,

Florida leading scientists expressed the need for ranavirus research to advance our

understanding/capabilities in the following areas: 1) phylogenetic and taxonomic classification,

2) development and validation of improved molecular diagnostic methods, and 3) mitigation

strategies to prevent high-risk populations from exposure to ranaviruses (Duffus et al. 2017,

www.ranavirus.org). The chapters of my dissertation were aimed at addressing these three focal

areas within aquaculture. First, viral characterization and phylogenomic analyses were used to

elucidate biologic and epidemiologic trends in aquaculture (Chapter 2). Comparative genomic

analyses facilitated the design of two new molecular assays, a pan-ranavirus quantitative real-

time TaqMan qPCR and an in situ hybridization assay using RNAscope® technology (Chapters

3 and 4). Finally, bath FV3 challenges were conducted at 17˚C and 23˚C to examine the effect of

http://www.ranavirus.org/

25

water temperature on ranaviral disease in hatchery-reared young-of-year pallid sturgeon (Chapter

4).

26

Table 1-1. Characteristics for members of the family Iridoviridae and genus Ranavirus.

Characteristic Family Iridoviridae Genus Ranavirus

Nucleocapsid symmetry icosahedral icosahedral

Virion diameter (nm) 120-350 160-200

Genome nucleic acid structure dsDNA dsDNA

Genome size (kbp) 104-303 104-140

Unique genome features Circularly permuted,

terminally redundant

Circularly permuted,

terminally redundant

G+C content (%) 28-57 48-57

Open reading frames (#) 73-211 73-139

27

CHAPTER 2

REPEATED DETECTIONS OF RANAVIRUSES IN AQUACULTURE

Introduction

Of the seven Ranavirus species recognized by the International Committee on the

Taxonomy of Viruses (ICTV), only one (e.g., Ambystoma tigrinum virus ATV) is known to

occur exclusively in amphibians, while four ranaviral species (e.g., European catfish virus ECV,

Epizootic hematopoietic necrosis virus EHNV, Santee-Cooper ranavirus SCRV, and Singapore

grouper iridovirus SGIV) are documented only in fish hosts. The final two ranaviral species

(e.g., Bohle iridovirus BIV and Frog virus 3 FV3) and certain unrecognized ranaviruses (e.g.,

common midwife toad ranavirus CMTV) possess the lowest known specificity among double-

stranded DNA viruses with documented infections in fish, amphibians, and reptiles.

Most studies exploring the taxonomic and phylogenetic diversity of ranaviruses have

relied upon sequences from a small number of genes including the major capsid protein (MCP)

gene (Mao et al. 1997, Tidona et al. 1998, Hyatt et al. 2002, Marsh et al. 2002, Holopainen et al.

2009). The MCP gene is invariably 1392 nucleotides in all ranaviruses and contains several

stable nucleotide mutations historically used to separate ranaviruses into distinct clades

(Holopainen et al. 2009, Jancovich et al. 2015). Despite the popularity of the MCP gene for

ranavirus phylogenetics, analysis of the MCP alone does not effectively resolve the relationships

among very closely related ranaviruses (e.g., FV3 strains) (Duffus and Andrews 2013, Jancovich

et al. 2015). More recently, researchers have generated robust phylogenetic trees based on the

concatenation of varying numbers of 26 core iridoviral and 27 core ranaviral genes (Eaton et al.

2007, Jancovich et al. 2015, Ariel et al. 2016, Hick et al. 2016, Holopainen et al. 2016,

Subramaniam et al. 2016b). The core genes represent proteins with known functions as well as

predicted proteins with unknown functions (Table 2-1); however, phylogenetic signal has not

28

been determined for all core genes. A recent study utilized full genome alignments in

phylogenetic analyses of ranaviruses (Claytor et al. 2017).

Several additional criteria (e.g., genomic organization, virion morphology, in vitro

growth characteristics) have been used to characterize iridoviruses (Jancovich et al. 2012).

Ranaviruses are easily propagated in several fish cell lines (e.g., bluegill fry (BF-2), chinook

salmon embryo (CHSE-214), epithelioma papulosum cyprini (EPC), and fathead minnow

(FHM)) and can even replicate in some mammalian cell lines at 30°C (Gravell and Granoff 1970,

Ariel et al. 2009). The majority of ranaviruses readily grow at room temperature (22-25°C)

causing cytopathic effect (CPE) within 24-72 hr. Several related ranaviruses isolated from

marine fishes including cod iridovirus (CoIV), Ranavirus maxima (Rmax), and short-finned eel

ranavirus (SERV) can be cultivated at colder temperatures of 12-15°C (Ariel et al. 2010, Bang

Jensen et al. 2009). Ranavirus CPE is first characterized by changes in individual cells (e.g.,

rounding, becoming refractile, and lysing) leading to focal plaques that coalesce until the entire

monolayer is destroyed (Miller et al. 2015).

Negative stain electron microscopy (EM) and transmission EM have been used to assist

with the diagnosis of ranavirus infections by characterizing virion structure and morphogenesis

(OIE 2016a,b). Ultrastructural characteristics of ranaviruses include hexagonal nucleocapsids

with a vertex-vertex diameter measurement ranging from 120-200 nm. A ranaviral assembly site

and paracrystalline array are typically located adjacently within the host cytoplasm, and

individual nucleocapsids may be observed obtaining the viral envelope in completion of their

maturation as they bud out from the host cell membrane (Chinchar et al. 2017b). Tissues to be

examined by TEM for ranaviruses are typically fixed in EM grade buffered fixatives containing

paraformaldehyde and glutaraldehyde (e.g., Karnovsky’s fixative). Negative stain EM can

29

provide ultrastructural details of the virion exterior by staining concentrated ranavirus

preparations with 1% phosphotungstic acid prior to examination (Waltzek et al. 2014).

Six of seven ranavirus species (e.g., BIV, ECV, EHNV, FV3, SGIV, ESV/ECV, SCRV,

SGIV) are documented in fish hosts (Duffus et al. 2015). Although few ranaviruses have been

detected within North American aquaculture facilities, FV3 has resulted in high mortality

epizootics in multiple farmed North American sturgeon species (Chapter 1, Waltzek et al. 2014),

underscoring the need for further study of fish ranaviruses. In this study, we utilized a suite of

techniques and analyses (e.g., in vitro growth characteristics, ultrastructural features, genome

annotation, and phylogenetics) to characterize 12 ranaviruses isolated from fishes.

Materials and Methods

Aquaculture Case Histories

Ranaviruses characterized here include six sturgeon isolates from epizootics that occurred

at three North American hatcheries. FV3 epizootics in young-of-year pallid sturgeon

(Scaphyrhynchus albus) have been reported at the Blind Pony State Fish Hatchery (BPSFH) in

Sweet Springs, Missouri resulting in cumulative mortalities up to 90-100% (Waltzek et al. 2014,

Chapter 4). Bath challenges recreated the lethal disease in fulfilment of Koch’s postulates

(Waltzek et al. 2014). Ranaviruses have also been isolated during high mortality epizootics at

other United States hatcheries involving white (Acipenser transmontanus) and Russian

(Acipenser gueldenstaedtii) sturgeon in California in 1998 and Georgia in 2004, respectively

(Waltzek et al. 2014). Experimental challenges involving intracoelomic injection of the Russian

sturgeon isolate into juvenile Russian and lake (Acipenser fulvescens) sturgeon reproduced the

disease resulting in 33% and 100% cumulative mortality, respectively (Robert Bakal personal

communication).

30

Two additional ranaviruses were isolated during US health certifications of apparently

healthy farmed fathead minnows (Pimephales promelas) in Arkansas in 2005 and northern pike

(Esox lucius) in Ohio in 2008 (Waltzek et al. 2014). In both cases, pooled internal tissue

homogenates were inoculated onto the epithelioma papulosum cyprini (EPC) cell line resulting

in cytopathic effect. The isolates were preliminarily identified as ranaviruses based on

transmission electron microscopy and PCR and Sanger sequencing performed at the National

Veterinary Services Laboratories in Ames, IA (Janet Warg personal communication).

Four previously characterized ranavirus isolates from European fishes were included for

comparison to the US fish isolates. The short-finned eel ranavirus (SERV) was isolated during a

health certification of apparently healthy short-finned eels (Anguilla australis) imported into

Italy from New Zealand (Bovo et al. 1999, Bang Jensen et al. 2009). The pike-perch iridovirus

(PPIV) was isolated from healthy-appearing cultured pike-perch (Sander lucioperca) fingerlings

in Finland in 1995 (Tapiovaara et al. 1998). The cod iridovirus (CoIV) was isolated from wild

Atlantic cod (Gadus morhua) in Danish coastal waters displaying an ulcerative skin disease

(Jensen et al. 1979, Ariel et al. 2010). The Ranavirus maxima (Rmax) was isolated during an

export certification of clinically healthy turbot (Scophthalmus maximus) fry from a Danish

aquaculture facility in 1999 (Ariel et al. 2010).

Isolation of Fish Ranaviruses

In the summer of 2015, the UF Wildlife and Aquatic Veterinary Disease Laboratory

(WAVDL) in Gainesville, FL received moribund pallid sturgeon fry from an ongoing epizootic

at the BPSFH. Four samples, each containing pooled internal tissues (liver, kidney, spleen) from

five fry, were diluted 1:25 in Minimal Essential Medium (MEM) with 2% Fetal Bovine Serum

(FBS) (MEM-2). Tissues were then homogenized using a stomacher (Seward Stomacher® 80

31

Biomaster) on high speed for 120 s. The homogenate was then centrifuged at 3000 x g (10 min at

4°C) to pellet cellular debris. An equal volume of the clarified tissue homogenate was added to a

MEM-2 antibiotic solution resulting in a final concentration of 500 IU penicillin ml−1

, 500 µg

streptomycin ml−1

, and 12.5 µg fungizone ml−1

, and 14 mM HEPES (4-(2-hydroxyethyl)-1-

piperazine ethane sulfonic acid) buffer. After incubating overnight at 4°C, the samples were

again clarified at 3000 x g (10 min at 4°C) and the tissue homogenates were inoculated onto 25

cm2 flasks containing confluent monolayers of EPC cells. After a 1 hr viral adsorption period,

the inoculum was pipetted from each flask and replaced with MEM containing 5% FBS (MEM-

5) and 1% HEPES buffer. Flasks were incubated at room temperature (24°C) and observed daily

for cytopathic effect.

Propagation of Fish Ranaviral Isolates

Eleven of the fish ranaviral isolates archived at the WAVDL have been previously

described (Tables 2-2 and 2-3). Isolates were thawed and inoculated onto duplicate 25 cm2

flasks

containing confluent monolayers of either EPC or bluegill fry (BF-2) cells. After a 1 hr

adsorption period, supernatant was pipetted from each flask and replaced with MEM-5

containing 1% HEPES buffer. Flasks were incubated at room temperature (24°C) and observed

and photographed every 24 hours until cytopathic effect (CPE) was complete. The contents of

one 25 cm2 flask were fixed for TEM and the other was expanded to four 175 cm

2 flasks

permitting virus purification prior to DNA library construction and genomic sequencing as

described below.

Transmission Electron Microscopy

Fixation for TEM was accomplished by adding an equal volume (5 ml) of modified

Karnovsky’s fixative containing 2% glutaraldehyde and 2% paraformaldehyde (Electron

Microscopy Sciences) to the 25 cm2 flask. After a 15 min fixation period, cells were scraped

32

from the flask with a cell scraper and the resulting cell suspension centrifuged at 3000 x g (10

min at 4°C). The resulting cell pellet was resuspended in 0.1 M neutral buffered cacodylate

buffer (Electron Microscopy Sciences) and stored at 4°C until TEM was performed. Fixed

samples were shipped to the Center for Biodefense and Emerging Infectious Diseases, Institute

for Human Infections and Immunity at The University of Texas Medical Branch (Galveston, TX)

for TEM. Photomicrographs were examined to quantify virion size and development. Mean

nucleocapsid (vertex to vertex) diameter and complete enveloped virion diameter measurements

were calculated from measurements of 20 nucleocapsids and 5 enveloped virions per isolate

using ImageJ2 (Schindelin et al. 2015). Image scale bars were used to calibrate the measurement

tool prior to analyzing each photomicrograph.

Cell Culture, Virus Purification, and DNA Extraction

The eight US fish ranaviruses with genomes not yet determined were propagated in EPC

cells grown to confluence in four 175 cm2 flasks at 24°C in MEM containing 10% FBS, 50 IU

penicillin ml−1

, 50 μg streptomycin ml−1

, and 2 mM L-glutamine. To reduce the risk of

contamination, isolates were grown at separate time points. Following a 1 hr virus adsorption

period, 50 ml of MEM-2 with 1% HEPES buffer was added and flasks were incubated at room

temperature (24°C) until cytopathic effect was complete. Cells were then harvested using a cell

scraper.

Viral purification was performed using a protocol slightly modified from Majji et al.

(2006) as described by Claytor et al. (2017). Scraped suspended cells underwent three freeze-

thaw events to release cell-associated virus and were then centrifuged at 5,509 x g (4°C for 20

min) using a Beckman JA-14 fixed angle rotor to remove cellular debris. Ultracentrifugation at

100,000 x g (4°C for 60 min) in a Beckman Type 50.2 Ti rotor was then performed to pellet the

33

virus, which was resuspended in 3 ml resuspension buffer (RSB; 10 mM Tris-HCl, pH 7.6, 10

mM KCl, 1.5 mM MgCl2). The 3 ml sample underwent a DNAse (200 μg ml–1

, Sigma) treatment

in the presence of 10 mM MgCl2 (60 min at 37°C) to remove non-target cell-associated DNA.

The reaction was stopped after 1 hr by adding EDTA to a final concentration of 50 mM, and the

virus sample was layered over a 20% (w/w) sucrose cushion and ultracentrifuged at 150,000 x g

(90 min at 4°C) in a Beckman Type 50.2 Ti rotor. Pelleted, purified virus was resuspended in

RSB and DNA extraction was performed using the Qiagen DNeasy extraction kit according to

manufacturer’s instructions. Finally, a Qubit® dsDNA BR Assay Kit was used to determine

concentration of the extracted DNA.

Library Preparation, Next Generation Sequencing, Assembly, Genome Annotation,

BLASTp Analyses

DNA libraries for the eight ranaviral samples were prepared using the TruSeq Dual Index

HT DNA PCR-free Library Preparation Kit (Illumina) as previously described (Claytor et al.

2017). Sequencing was performed using v3 chemistry on an Illumina MiSeq sequencer.

Following sequencing, Velvet (Zerbino and Birney 2008) and SPAdes (Bankevich et al. 2012)

software were used to perform de novo genome assembly. Velvet was first used to generate

contigs using k-mer values ranging from 19 to 191 and a search step size of 2. Output trusted

contigs were entered into SPAdes for assembly using k-mers 21, 33, 55, 77, 99, and 127 with

automatic cutoff setting for coverage. Reads were mapped against the final genome consensus

using Bowtie 2 within Unipro UGENE 1.26 (Langmead and Salzberg 2012) to determine quality

of the assembly and the resulting data were visualized in Tablet 1.14.10.20 (Milne et al. 2010).

Genome annotation was performed using Genome Annotation Transfer Utility (Tcherepanov et

al. 2006) using FV3 (Genbank Reference Sequence NC_005946) as the reference genome.

Annotated gene functions were predicted by performing BLASTP searches against the National

34

Center for Biotechnology Information (NCBI) GenBank non-redundant protein sequence

database. Criteria utilized for designating open reading frames (ORFs) included: 1) >120 nt in

length, 2) must not overlap with another ORF by more than 25%, 3) in the case of overlapping

ORFs, only the larger ORF was kept (O’Dea et al. 2016).

Multiple Genome-wide Alignments and Phylogenomic Analyses

The genomes of 28 ranaviruses genomes including the eight US fish ranavirus genomes

sequenced as part of this study (Table 2-3) were reordered to ensure the 5’ end of each genome

began with the arbitrarily chosen FV3 ORF1R as previously described (Claytor et al. 2017). The

reordered genomes were then aligned in MAUVE version 2.01 (Darling et al. 2004) to visualize

genomic inversions and obtain the six locally collinear blocks (LCB1-6) alignments that were

then concatenated using Geneious 10.0.2 (www.geneious.com, Kearse et al. 2012).

Maximum likelihood (ML) and Bayesian methods were each utilized for tree

construction. A ML phylogram was generated from the concatenated LCB nucleotide alignment

in IQ-TREE 1.5 software with 1000 bootstrap replicates to determine robustness at each node

(Trifinopoulos et al. 2016) using the best fit general time reversible model, gamma distributed

with invariant sites (GTR+I+G4). Clades with bootstrap values of >70% were considered well

supported. The same evolutionary model was implemented in MrBayes version 3.2 (Ronquist et

al. 2011) for the Bayesian analysis. Markov-chain Monte Carlo (MCMC) analysis was run with

default priors for topology, branch lengths, the four stationary frequencies of the nucleotides, the

six different nucleotide substitution rates, the proportion of invariable sites, and the shape

parameter of the gamma distribution of rate variation. Analyses were run with 3 hot chains and 1

cold chain with the default heating parameter (temperature = 0.2). Bayesian analyses were run

for a length of 1,100,000 generations, with a stopping rule implemented when average standard

deviation of split frequencies fell below 0.001%. Chains were sampled at a diagnostic frequency

http://www.geneious.com/

35

of every 1,000 generations and the first 25% of MCMC samples were discarded as burn-in. The

maximum clade credibility tree was chosen based on the highest product of clade probabilities.

Clades were considered well supported if they possessed posterior probabilities of >0.90.

Topologies of ML and Bayesian trees were examined for congruency, and bootstrap and

posterior probability support values were reported at each node. Phylogenetic trees were

visualized and edited in FigTree v1.4.2 (http://tree.bio.ed.ac.uk/software/figtree/).

Results

Isolation of Fish Ranaviruses

All four pooled internal tissue samples from the 2015 BPSFH epizootic in juvenile pallid

sturgeon resulted in CPE on EPC cells within 48 hours of inoculation. CPE was characterized by

the development of focal plaques that coalesced until the entire cell monolayer was destroyed

(Figure 2-1, Table 2-4).

Propagation of Fish Ranaviral Isolates

Nine of the eleven remaining ranaviral isolates (excluding Rmax and CoIV) were

successfully cultivated in EPC cells, while Rmax and CoIV isolates were successfully cultured

on BF-2 cells. CPE occurred in all isolates within 72 hr of incubation at 24°C and was similar to

the 2015 pallid sturgeon samples described above (Figure 2-1, Table 2-4).

Transmission Electron Microscopy

All 12 fish ranaviruses displayed characteristic hexagonal nucleocapsids with electron-

dense cores and vertex- vertex diameter measurements ranging from 124-159 nm as expected for

ranaviruses (Table 2-4) (Chinchar et al. 2017a). Photomicrographs of several isolates showed

virions arranged in characteristic viral assembly sites with associated paracrystalline arrays

within the cytoplasm of infected cells. Virions were observed acquiring an outer envelope as

http://tree.bio.ed.ac.uk/software/figtree/

36

they budded through the cell membrane in all isolates except WSRV for which only a few

photomicrographs were available (Figure 2-2, Table 2-4).

Next Generation Sequencing, Assembly, Genome Annotation, and Phylogenetic Analyses

The eight assembled genomes ranged from 105,472-106,777 bp, encoded between 96-98

ORFs, and displayed a G+C content ranging from 55.42-57.74%, all within established

parameters for members of the genus Ranavirus (Chinchar et al. 2017a, Tables 1-1 and 2-5).

Average mapped reads coverage ranged 6,240-11,047 reads/nt. All 8 genomes possessed the 26

core iridoviral and 27 core ranaviral genes outlined by Eaton et al. (2007) (Tables 2-6 through 2-

13).

The multiple genome alignment revealed distinct groups of ranaviruses based on genome

structure (Figure 2-4). The 28 ranavirus genomes included in the analysis were arranged into six

locally collinear blocks (LCB1-6). Using SERV to represent the ancestral genomic arrangement,

a series of genome inversions were revealed. While other basal ranaviruses (e.g., ESV, EHNV,

and ATV) share SERV’s ancestral genome arrangement, the Rmax/CoIV group alone display a

small inversion of the LCB5, creating a unique genome arrangement not observed in any other

examined taxa. The CMTV members, including the newly characterized WSRV, all display a

medium-sized inversion of the LCB3. Finally, the most derived clades (e.g., TFV, BIV, and

FV3) all display a large inversion of the LCB2 through LCB5 region that likely occurred

following their split from most recent common ancestor shared with the CMTV clade) Chen et

al. 2013, Mavian et al. 2012a, Claytor et al. 2017).

Phylogenomic analyses indicated that the eight fish ranavirus genomes fully

characterized here and the four genomes previously sequenced each grouped within specific

clades within the Ranavirus genus (Figure 2-3). The European fish isolates SERV, Rmax and

CoIV all lie at the base of the tree, with Rmax and CoIV forming their own clade and SERV

37

joining the EHNV-like group (Claytor et al. 2017). PPIV and WSRV grouped within the CMTV-

like clade, while the FHMRV, NPRV, and the five other sturgeon isolates grouped within the

FV3 clade. Within the FV3 clade, the FHMRV, NPRV, and RSRV isolates grouped together as

the sister group to the clade formed by FV3, SSME, and PSRV01, -09, -13, and -15.

Genome annotations revealed several notable differences among the eight US fish

ranavirus genomes, including genes involved with viral replication and pathogenesis such as The

US22 family protein present in members of several double-stranded DNA viral families (Chen et

al. 2013, Claytor et al. 2017) and the the ranaviral homologue of the eukaryotic translation

initiation factor 2 (vIF-2alpha). All eight fish ranavirus genomes studied possessed one or two

US22 genes at three possible locations within their genomes as previously described (Claytor et

al. 2017). Six of fish ranaviruses (FHMRV, NPRV, PSRV01, -09, -11, -15) possessed two US22

copies as previously described for most strains of FV3 (e.g., orthologous to FV3 ORFs 5R and

98R; Reference Sequence NC_005946). The Russian sturgeon ranavirus (RSRV) lacks ORFs 5R

and 6R; however, it encodes the ADRV ORF50 ortholog (GenBank Reference Sequence

KF033124) and the FV3 ORF98R ortholog similar to a recently described recombinant bullfrog

ranavirus (Claytor et al. 2017). Similar to other members of CMTV clade, WSRV possesses the

FV3 ORF5R ortholog and the ADRV ORF50 ortholog. The viral homologue for the eukaryotic

translation initiation factor 2 (evIF-2alpha), was truncated in the FHMRV, NPRV, and PSRV01

genomes.

Discussion

Ranaviruses are a global threat to farmed and wild fishes, with the majority of cases

occurring in captive facilities (Whittington et al. 2010, Table 2-2). Herein, we characterized 11

ranaviruses isolated from aquaculture facilities and a single ranavirus from a wild case. The fish

ranaviral isolates exhibited similar in vitro characteristics, virion architecture, and

38

morphogenesis typical for ranaviruses (Table 2-4) (Chinchar et al. 2017a). The eight sequenced

genomes sizes, number of ORFs, and %G+C content were all within established parameters for

members of the genus Ranavirus (Chinchar et al. 2017a, Table 2-5). The eight sequenced

genomes possessed the 26 core iridoviral genes and additional 27 for ranaviruses outlined by

Eaton et al. (2007). Finally, the gene order and inversion patterns of each of the analyzed fish

ranaviral genomes were congruent with the previously defined ranaviral genomic orientations

(Figure 2-4, Claytor et al. 2017).

The work presented herein is the first to incorporate full genome comparisons of fish

ranaviruses in an effort to resolve common issues associated with partial genome analyses

including poorly resolved or supported trees (Figure 2-3). Claytor and colleagues (2017) reported

that full genomic analyses generated ranaviral trees congruent to analyses based on a smaller

number of concatenated genes and even overcome regions of poor phylogenetic signal that arise

from recombination. The maximum likelihood and Bayesian trees presented here suggest

ranaviruses likely originated in fish, as the most basal Ranavirus members have been

documented in fish hosts (Jancovich et al. 2010, 2015). These include short-finned eel ranavirus

(SERV), Epizootic hematopoietic necrosis virus (EHNV), European catfish virus (ECV),

European sheatfish virus (ESV), cod iridovirus (CoIV), and Ranavirus maxima (Rmax) isolated

from turbot (Figure 2-3). The highly divergent Santee-Cooper ranavirus (SCRV) and Singapore

grouper iridovirus (SGIV) ranaviruses were excluded from the full genome analyses as there is

almost no collinearity of their genomes to other ranaviruses, supporting their position as the most

basal members of the genus (Jancovich et al. 2015). Ambystoma tigrinum virus (ATV), which

branches off the tree immediately after the aforementioned basal fish ranaviruses, appears to be

the first evolutionary jump into amphibians (Jancovich et al. 2010). The more derived ranavirus

39

lineages, the Frog virus 3-like ranaviruses (e.g., BIV, TFV, STIV, RGV, SSME, NPRV,

FHMRV, PSRV01, -09, -11, -15) and the CMTV-like ranaviruses (e.g., PPIV, ADRV, CH8/96,

RCV-Z, WSRV), exhibit very low host specificity infecting three different classes of vertebrates

(Mao et al. 1999a, Brenes et al. 2014, Waltzek et al. 2014).

Our phylogenomic analyses revealed seven new fish strains of FV3 including five

isolated from epizootics at sturgeon hatcheries (Figure 2-3). Interestingly, the four pallid

sturgeon isolates from the Blind Pony State Fish Hatchery (BPSFH) in Sweet Springs, MO

formed a monophyletic group indicating that the same FV3 strain is responsible for repeated

epizootics negatively impacting the recovery efforts of this endangered species. The isolate from

the Russian sturgeon epizootic grouped together with the fathead minnow ranavirus and northern

pike ranavirus that were both isolated from visibly healthy fish as part of health certifications.

These findings support fish as potential asymptomatic carriers of FV3 (Une et al. 2009, Brenes et

al. 2014).

Similar to FV3, the CMTV-like ranaviruses include closely related strains isolated from

diverse hosts from around the world. For example, Andrias davidianus ranavirus (ADRV) has

caused epidemics in highly endangered giant salamanders in Chinese rearing facilities since 2010

while the related common midwife toad virus (CMTV) is an important pathogen of wild

European amphibians (Chen et al. 2013, Price et al. 2014). Also in Europe (Switzerland), a

reptile CMTV member known as CH8/96 caused 100% mortality in a captive population of

Hermann’s tortoises (Testudo hermanni) in 1996 (Stöhr et al. 2015). Sequencing of the pike-

perch iridovirus (PPIV) genome, isolated in Finland from apparently healthy pike-perch (Sander

lucioperca) fingerlings in 1995, revealed PPIV is the earliest known member of the

CMTV/ADRV-like ranavirus clade and the sole isolate from a fish (Holopainen et al. 2016). Our

40

phylogenomic analysis revealed the white sturgeon ranavirus (WSRV), isolated in 1998 on a

Californian sturgeon farm, represents the second fish member of the CMTV clade (Figure 2-3).

Interestingly, the closest relatives of the WSRV are two geographically distant isolates:

ADRV from China and the Rana catesbeiana virus (RCV-Z) isolated from a US aquaculture

facility during a bullfrog epizootic in 1998 (Claytor et al. 2017, Figure 2-3b). The discovery of

closely related ranaviruses impacting diverse hosts on multiple continents over a short time

period of time suggests international trade is driving the spread of these emerging pathogens.

Unregulated movement of animals, particularly the American bullfrog (Lithobates catesbeianus)

for the human food, bait, and pet trades, has been proposed as an important route in

disseminating ranaviruses into new geographic regions with naïve populations (Schloegel et al.

2009, Altherr et al. 2011, Duffus et al. 2015, Claytor et al. 2017). Although not as well studied as

the international trade in bullfrogs, this work leaves open the possibility that the international

live sturgeon trade may also have contributed to the spread of FV3 and CMTV strains.

The US22 family proteins, present in members of several double-stranded DNA viral

families, are believed to counter host antiviral responses (Chen et al. 2013). Several FV3 and

CMTV members of the Ranavirus genus are documented to possess one or two US22 genes

likely gained from their amphibian hosts or even during recombination events with other

ranaviruses (Chen et al. 2013, Claytor et al. 2017). The eight fish ranavirus genomes examined

here possessed two copies of the US22 genes as previously described (Claytor et al. 2017).

US22 genes in strains of FV3 and CMTV have been argued to influence virulence and host range

evolution in amphibians; however, their role in fish hosts remains unknown. Further analysis will

be required to examine if RSRV, which possesses genes of high identity to both CMTV/ADRV

41

and FV3, may represent another recombinant virus similar to the recently described bullfrog

isolate RCV-Z2 (Claytor et al. 2017).

Another gene involved in viral replication and pathogenesis is the viral homologue of the

eukaryotic translation initiation factor 2 (eIF-2alpha), which contributes to host protein synthesis

(Grayfer et al. 2012). As a core ranaviral gene, the vIF-2alpha acts as a pseudosubstrate for

protein kinase R (PKR), a kinase that arrests host protein synthesis when a host cell becomes

infected. If PKR binds to the vIF-2alpha pseudosubstrate instead of the host’s eIF-2alpha, the

host cell’s transcription machinery can be utilized by the virus to facilitate viral replication

(Grayfer et al. 2015). Of the eight examined genomes, truncated vIF-2alpha genes were found in

the FHMRV, NPRV, and PSRV01 genomes. Truncation of the vIF-2alpha gene may reduce its

function and allow PKR to effectively bind to the host’s eIF-2alpha, arresting protein synthesis

in infected cells. Two of the three genomes with truncated vIF-2alpha genes (FHMRV and

NPRV) were isolated from asymptomatic hosts; however, PSRV01 still caused significant

morbidity and mortality in juvenile pallid sturgeon despite possessing a truncated vIF-2alpha

gene as has been observed in other lethal FV3-like ranaviruses such as the soft-shelled turtle

iridovirus (Huang et al. 2009). Thus, the presence of a full-length vIF-2alpha gene cannot alone

explain virulence among these ranaviruses. Despite being each others’ closest relatives, the four

pallid sturgeon ranaviruses (PSRV01, -09, -13, and -15) possessed subtle differences in their

genomes.

Young-of-year sturgeon including several endangered or threatened species (e.g., lake,

pallid, Russian, white) appear to be highly susceptible to FV3- and CMTV-like ranaviruses.

Multiple reasons for increased susceptibility in sturgeon have been proposed, including low

genetic diversity and immune function amplified by aquaculture-related stressors (e.g.,

42

temperature stress, high densities, poor water quality) (LaPatra et al. 1996, Georgiadis et al.

2000, 2001, Drennan et al. 2005, Savin et al. 2011). Sturgeon may contribute to amplification

and spread of ranaviruses in cultured and wild populations. The pathogenicity of certain FV3 and

CMTV members isolated from asymptomatic non-sturgeon fish hosts (e.g., FHMRV, NPRV, and

PPIV) remains unknown. Certain fish species may indeed be less susceptible to ranavirus disease

and still serve as carriers or reservoirs, resulting in viral dissemination to more susceptible

species of fish, amphibians, and reptiles.

In conclusion, we characterized 12 ranaviruses isolated from a range of aquacultured and

wild freshwater and marine fishes. Genomic sequencing of eight fish ranaviruses suggests fish

are susceptible to strains of both FV and CMTV with some species such as juvenile sturgeon

being highly susceptible. Future studies considering host, environmental, and viral genetic

differences among strains are needed to explain the observed difference in pathogenicity between

the sturgeon ranavirus isolates and less pathogenic strains (Chapter 4). Experimental challenges

comparing the sturgeon ranaviruses across a range of hosts, such as those performed in

amphibians (Hoverman et al. 2010) and across multiple host classes (Brenes et al. 2014) are

warranted. Future efforts are also needed to determine how transmission of lethal ranaviruses

occurs at sturgeon hatcheries (e.g., use of contaminated surface waters, infected broodstock, etc.)

and whether less susceptible fish species serve as important ranaviral reservoirs. Finally, the risk

infected fish and water from aquaculture facilities may pose to naïve wild populations needs to

be determined.

43

Table 2-1. Conserved iridoviral/ranaviral genes and their predicted functions if known. Table is

modified from Eaton and colleagues (2007). Gene numbers correspond to the

Epizootic hematopoietic necrosis virus annotation (RefSeq# NC_028461).

Gene Predicted function

1L Myristylated membrane protein

4Ra,b

Unknown function

7R DNA-dependent RNA polymerase II largest subunit

8L NTPase/helicase

9Ra Unknown function

10L DNA repair enzyme RAD2

11R Unknown function

13L Immediate early protein ICP-46

14L Major capsid protein

15La Unknown function

16L Thiol oxidoreductase

18L Thymidine kinase

19L Proliferating cell nuclear antigen (PCNA)

22L Immediate early protein ICP-18

23L Transcription elongation factor S-II

24R Ribonuclease III


26Ra,c

Unknown function





38R Ribonucleoside reductase alpha subunit

41La Caspase recruitment domain protein

42La dUTPase

43R DNA-dependent RNA polymerase II second largest subunit

44L DNA polymerase


48L Phosphotransferase

53L Myristylated membrane protein






62R Tyrosine kinase


68Ra Neurofilament triplet H1-like protein


44

Table 2-1. Continued

Gene Predicted function





85L D5 family NTPase/ATPase


89L Serine/threonine protein kinase



92L ATPase



100R Replicating factor a Core ranaviral gene

b absent in GIV

c absent in ATV

45

Table 2-2. Documented ranavirus detections in fish hosts. Host common names refer to natural hosts only. Refer to Chapter 1 for

details of experimentally infected hosts.

Ranavirus name

Abbrevia-

tion Natural host common name

Wild/

cultured

Morbidity/

mortality Earliest citation

Bohle iridovirus BIV Nile tilapia Cultured Yes Ariel and Owens 1997

Cod iridovirus a

CoIV Atlantic cod Wild No Jensen et al. 1979

Doctor fish virusb DFV Doctor fish Wild Yes Hedrick and McDowell 1995

Epizootic hematopoietic necrosis

virus

EHNV Redfin perch, rainbow trout Both Yes Langdon et al. 1986, 1988

European catfish virus ECV Brown bullhead, black bullhead Both Yes Pozet et al. 1992

European sheatfish virus ESV Sheatfish Cultured Yes Ahne et al. 1989, 1991

Fathead minnow ranavirus a

FHMRV Fathead minnow Cultured No Waltzek et al. 2014

Frog virus 3 FV3 Three-spine stickleback Wild Yes Mao et al. 1999a

Grouper iridovirus GIV Yellow grouper Cultured Yes Murali et al. 2002

Guppy virus 6b GV6 Guppy Cultured Yes Hedrick and McDowell 1995

Largemouth bass virusb LMBV Largemouth bass, koi, northern

snakehead, et al.

Both Yes Plumb et al. 1996, Mao et al. 1999b

Northern pike ranavirus a

NPRV Northern pike Cultured No Waltzek et al. 2014

Pallid sturgeon ranavirus a

PSRV Pallid sturgeon Cultured Yes Waltzek et al. 2014

Russian sturgeon ranavirus a

RSRV Russian sturgeon Cultured Yes Waltzek et al. 2014

Pike-perch iridovirus a

PPIV Pike-perch Cultured No Tapiovaara et al. 1998

Ranavirus maxima a

Rmax Turbot Cultured No Ariel et al. 2010

Short-finned eel ranavirus a

SERV Short-finned eel Cultured No Bovo et al. 1999

Singapore grouper iridovirus SGIV Brown-spotted grouper Cultured Yes Chua et al. 1994

Tiger frog virus TFV Marbled sleeper goby Cultured Yes Prasankok et al. 2005

White sturgeon ranavirus a WSRV White sturgeon Cultured Yes Waltzek et al. 2014

a Sample characterized in this chapter

b DFV, GV6, and LMBV are all members of the species Santee-Cooper ranavirus

46

Table 2-3. Fully sequenced ranavirus genomes used for phylogenomic analyses within this chapter.

Ranavirus isolate Abbreviation Accession no. Reference

Ambystoma tigrinum virus ATV AY150217 Jancovich et al. 2003

Andrias davidianus ranavirus ADRV KC865735 Geng et a. 2011

Bohle iridovirus BIV KX185156 Hick et al. 2016

Cod iridovirus CoIV KX574342 Ariel et al. 2016

Common midwife toad virus-E CMTV_E JQ231222 Mavian et al. 2012a

Common midwife toad virus-NL CMTV_NL KP056312 Van Beurden et al. 2014

Epizootic hematopoietic necrosis virus EHNV FJ433873 Jancovich et al. 2010

European sheatfish virus ESV JQ724856 Mavian et al. 2012b

Fathead minnow ranavirus FHMRV Unpublished WAVDLa

Frog virus 3 FV3 AY548484 Tan et al. 2004

German gecko ranavirus GGRV KP266742 Stöhr et al. 2015

Northern pike ranavirus NPRV Unpublished WAVDLa

Pike-perch iridovirus PPIV KX574341 Holopainen et al. 2016

Pallid sturgeon ranavirus 2001 PSRV01 Unpublished WAVDLa




Rana catesbeiana virus-Z RCV-Z Unpublished WAVDL

Rana grylio virus RGV JQ654586 Lei et al. 2012

Ranavirus maximus Rmax KX574343 Ariel et al. 2016

Russian sturgeon ranavirus RSRV Unpublished WAVDLa

Short-finned eel ranavirus SERV KX353311 Subramaniam et al. 2016b

Soft-shelled turtle iridovirus STIV EU627010 Huang et al. 2009

Spotted salamander Maine virus SSME KJ175144 Morrison et al. 2014

Testudo hermanni ranavirus CH8/96 KP266741 Stöhr et al. 2015

Tiger frog virus TFV AF389451 He et al. 2002

Tortoise ranavirus ToRV KP266743 Stöhr et al. 2015

White sturgeon ranavirus WSRV Unpublished WAVDLa

a Ranavirus genomically sequenced at the WAVDL for the purposes of this chapter

47

Table 2-4. In vitro growth chracteristics and ultrastructural analyses for the 12 aquaculture isolates included in this chapter. Mean

measurements (±standard deviation SD) are based on n=20 for nucleocapsids and n=5 for complete enveloped virions due

to limited image availability for enveloped virions. NO = not observed due to the small number of available

photomicrographs.

Isolate

abbreviation Host common name (latin name)

Cell

line

Time to

complete

CPE (hr)

Nucleocapsid

vertex-vertex

diameter

(SD)(nm)

Enveloped virion

diameter (SD)(nm)

CoIV Atlantic cod (Gadus morhua) BF-2 48 150.8 (5.7) 188.3 (4.3)

FHMRV Fathead minnow (Pimephales promelas) EPC 24 142.9 (5.7) 172.4 (11.0)

NPRV Northern pike (Esox lucius) EPC 48 130.6 (7.8) 179.8 (18.4)

PPIV Pike-perch (Sander lucioperca) EPC 24 144.3 (7.0) 208.9 (11.4)

PSRV01 Pallid sturgeon (Scaphirhynchus albus) EPC 24 140.3 (6.0) 222.9 (8.7)




Rmax Turbot (Scophthalmus maximus) BF-2 48 141.4 (8.6) 181.7 (2.4)

RSRV Russian sturgeon (Acipenser gueldenstaedtii) EPC 24 153.6 (7.0) 189.6 (9.2)

SERV Short-finned eel (Anguilla australis) EPC 48 123.6 (5.4) 162.9 (9.2)

WSRV White sturgeon (Acipenser transmontanus) EPC 48 158.5 (4.5) NO

48

Table 2-5. Genome characteristics for the eight fish ranaviruses sequenced in this chapter. See Table 2-3 for taxa abbreviations.

Isolate Ranavirus clade Size (nt)

Mapped reads

(%) ORFs (#) G+C (%)

Mean coverage

(reads/nt)

FHMRV FV3 105,472 82.54 98 55.42 10,365.2

NPRV FV3 105,545 90.49 98 56.13 10,231.6

PSRV01 FV3 105,770 94.07 98 55.62 9,022.2

PSRV09 FV3 105,488 92.24 97 57.74 6,240.6

PSRV13 FV3 106,506 95.73 98 55.69 8,808.9

PSRV15 FV3 106,248 96.85 97 56.05 9,608.2

RSRV FV3 106,777 87.35 97 55.46 11,047.6

WSRV CMTV 105,934 63.21 96 55.50 9,835.8

49

Table 2-6. Predicted open reading frames for the fathead minnow ranavirus isolate (FHMRV). RUK13 = Ranavirus United Kingdom

13. See Table 2-3 for other taxa abbreviations.

ORF

Position (nt

range)

Product

size (aa) Predicted function and conserved domain or signature

Best BLAST hit

Ranavirus ORF % ID Accession no.

1 1-771 257 Replicating factor FV3 1R 99 YP_031579

2 1378-2340 321 Myristylated membrane protein FV3 2L 100 YP_031580

3 2378-3217 280 RGV 3L 100 AFG73045

4 3248-4462 405 FV3 3R 100 YP_031581

5 4503-4685 61 FV3 4R 98 YP_031582

6 5118-5732 205 US22 family protein FV3 5R 94 YP_031583

7 5735-5962 76 STIV 8R 99 ACF42227

8 6723-7151 143 STIV 9L 96 ACF42228

9 7230-11111 1294 DNA-dependent RNA polymerase II largest subunit RGV 9R 100 AFG73051

10 11480-14326 949 NTPase/helicase FV3 9L 100 YP_031587

11 14342-14755 138 FV3 10R 100 YP_031588

12 15105-15317 71 FV3 11R 100 YP_031589

13 15383-16276 298 FV3 12L 100 YP_031590

14 16817-17023 69 FV3 13R 100 YP_031591

15 17038-17397 120 FV3 14R 100 YP_031592

16 17493-18461 323 ATPase FV3 15R 100 YP_031593

17 18741-19568 276 FV3 16R 100 YP_031594

18 19809-21317 503 FV3 17L 100 YP_031595

19 21354-21590 79 FV3 18L 100 YP_031596

20 21642-24257 872 Serine/threonine protein kinase RUK13 19R 99 AIX94586

21 24305-24751 149 PPIV 22 100 ANZ57004

22 24974-25633 220 SSME 56L 100 AHM26100

23 25763-28684 974 D5 family NTPase/ATPase SSME 22R 100 AHM26101

24 29062-30210 383 RUK13 23R 99 AIX94588

25 30620-31690 357 SSME 24R 100 AHM26103

26 31884-32669 262 GGRV 21R 100 AJR29179

27 32736-32966 77 eIF-2alpha-like protein FV3 26R 100 YP_031604

28 33498-36410 971 Tyrosine kinase FV3 27R 100 YP_031605

29 36459-36947 163 FV3 28R 100 YP_031606

30 37126-37422 99 FV3 29L 100 YP_031607

31 37624-37776 51 FV3 30R 100 YP_031608

32 37838-38257 140 FV3 31R 100 YP_031609

33 38307-38780 158 Neurofilament triplet H1-like protein FV3 32R 100 YP_031610

50


ORF

Position (nt

range)

Product


Best BLAST hit


34 40280-40471 64 CGSIV 33R 100 AGK44962

35 40615-40935 107 SSME 34R 99 AHM26113

36 41027-41488 154 SSME 35L 100 AHM26114

37 41686-42108 141 FV3 36L 100 YP_031614

38 42503-43138 212 NIF/NLI interacting factor FV3 37R 100 YP_031615

39 43279-44976 566 Ribonucleoside reductase alpha subunit FV3 38R 100 YP_031616

40 45082-45432 117 FV3 39R 100 YP_031617

41 45521-46069 183 SSME 40R 99 AHM26119

42 46451-49948 1166 SSME 41R 100 AHM26120

43 50444-50701 86 FV3 42L 100 YP_031620

44 50700-51314 205 ATV 80R 95 ALN36670

45 51627-52037 137 FV3 45L 99 YP_031623

46 52091-52492 134 Neurofilament triplet H1-like protein BIV 46L 98 ANK57973

47 52617-53033 139 ToRV 47L 100 AJR29281

48 53036-53287 84 FV3 48L 100 YP_031626

49 53396-55051 552 SSME 50L 100 AHM26127

50 55131-56816 562 FV3 51R 100 YP_031629

51 57073-58140 356 3-beta-hydroxysteroid dehydrogenase RGV 52L 100 ABI36881

52 58478-60046 523 Myristylated membrane protein FV3 53R 100 YP_031631

53 60341-60571 77 Nuclear calmodulin-binding protein FV3 54L 100 YP_031632

54 60609-61904 432 Helicase-like protein FV3 55L 100 YP_031633

55 60753-61892 380 40 kDa protein FV3 55R 100 YP_031634

56 61992-62429 146 FV3 56R 100 YP_031635

57 62543-64039 499 Phosphotransferase FV3 57R 100 YP_031636

58 64364-65077 238 FV3 58R 100 YP_031637

59 65330-66388 353 FV3 59L 99 YP_031638

60 66550-69591 1014 DNA polymerase SSME 60R 100 AHM26138

61 69600-69782 61 FV3 61L 98 YP_031640

62 70219-73884 1222 DNA-dependent RNA polymerase II second largest subunit FV3 62L 100 YP_031641

63 74263-74757 165 dUTPase FV3 63R 100 YP_031642

64 74867-75154 96 Caspase recruitment domain protein FV3 64R 100 YP_031643

65 75547-75711 55 FV3 65L 100 YP_031644

66 75708-76064 119 FV3 66L 98 YP_031645

67 76313-77476 388 Ribonucleoside reductase beta subunit FV3 67L 100 YP_031646

68 77760-78047 96 FV3 68R 99 YP_031647

69 78183-78449 89 FV3 69R 100 YP_031648

51


ORF

Position (nt

range)

Product


Best BLAST hit


70 78467-78841 125 FV3 70R 99 YP_031649

71 78881-79114 78 FV3 71R 100 YP_031650

72 79171-79887 239 FV3 72L 100 YP_031651


74 81482-82594 371 FV3 74L 100 YP_031653

75 82626-82880 85 LITAF/PIG7 possible membrane associated motif in LPS-induced

tumor necrosis factor alpha factor

FV3 75L 100 YP_031654

76 82943-83164 74 FV3 76R 100 YP_031655

77 83161-83508 116 FV3 77L 100 YP_031656

78 84123-84761 213 FV3 78L 100 YP_031657

79 84897-86615 573 ATPase-dependent protease RUK13 79R 100 AIX94632

80 87238-88353 372 Ribonuclease III FV3 80L 100 YP_031659

81 88409-88687 93 Transcription elongation factor S-II FV3 81R 100 YP_031660

82 88816-89289 158 Immediate early protein ICP-18 FV3 82R 100 YP_031661

83 89739-90383 215 Cytosine DNA methyltransferase FV3 83R 100 YP_031662

84 90755-91492 246 Proliferating cell nuclear antigen (PCNA) PPIV 84R 100 ANZ56939

85 91567-92154 196 Thymidine kinase FV3 85R 100 YP_031664

86 92544-92729 62 FV3 44R 100 YP_031665

87 93082-94899 606 FV3 87L 100 YP_031666

88 94932-95384 151 Thiol oxidoreductase GGRV 65R 99 AJR29223

89 95452-96594 381 SSME 89R 100 AHM26165

90 96686-98077 464 Major capsid protein SSME 90R 100 AHM26166

91 98201-99388 396 Immediate early protein ICP-46 SSME 91R 100 AHM26167

92 99691-99936 82 SSME 92R 100 AHM26168

93 100118-100285 56 FV3 93L 100 YP_031672

94 100395-100862 156 FV3 94L 100 YP_031673

95 100955-102046 364 DNA repair enzyme RAD2 STIV 100R 100 ACF42318

96 102848-103519 224 FV3 96R 100 YP_031675

97 103602-104015 138 Myeloid cell leukemia protein STIV 103R 100 ACF42321

98 104579-105244 222 US22 family protein STIV 104R 100 ACF42322

52

Table 2-7. Predicted open reading frames for the northern pike ranavirus isolate (NPRV). RUK13 = Ranavirus United Kingdom 13.

See Table 2-3 for other taxa abbreviations.

ORF

Position (nt

range)

Product


Best BLAST hit



2 1378-2340 321 Myristylated membrane protein FV3 2L 100 YP_031580

3 2378-3217 280 RGV 3L 100 AFG73045

4 3248-4462 405 FV3 3R 100 YP_031581

5 4503-4685 61 FV3 4R 100 YP_031582


7 5735-5962 76 STIV 8R 100 ACF42227

8 6723-7151 143 STIV 9L 96 ACF42228



11 14343-14756 138 FV3 10R 99 YP_031588

12 15106-15318 71 FV3 11R 100 YP_031589

13 15384-16277 298 PPIV 12L 100 ANZ57123

14 16818-17024 69 FV3 13R 100 YP_031591

15 17038-17397 120 FV3 14R 100 YP_031592

16 17493-18461 323 ATPase FV3 15R 100 YP_031593

17 18741-19568 276 FV3 16R 100 YP_031594

18 19809-21317 503 FV3 17L 100 YP_031595

19 21355-21591 79 FV3 18L 100 YP_031596

20 21643-24225 861 Serine/threonine protein kinase SSME 19R 99 AHM26098

21 24273-24719 149 PPIV 22 100 ANZ57004

22 24942-25601 220 SSME 56L 100 AHM26100


24 29030-30178 383 FV3 23R 100 YP_031601

25 30561-31658 366 RUK13 24R 100 AIX94589

26 31852-32643 264 ToRV 63L 100 AJR29294

27 32710-32940 77 EIF-2alpha-like protein FV3 26R 100 YP_031604


29 36432-36920 163 FV3 28R 100 YP_031606

30 37099-37395 99 FV3 29L 100 YP_031607

31 37597-37749 51 FV3 30R 100 YP_031608

32 37811-38230 140 FV3 31R 100 YP_031609

53


ORF

Position (nt

range)

Product


Best BLAST hit


33 38280-40058 593 Neurofilament triplet H1-like protein FV3 32R 100 YP_031610

34 40141-40332 64 CGSIV 33R 100 AGK44962

35 40476-40796 107 SSME 34R 100 AHM26113

36 40888-41349 154 SSME 35L 100 AHM26114

37 41321-41962 214 RUK13 36L 98 AIX94599



40 44937-45287 117 FV3 39R 100 YP_031617

41 45376-45924 183 SSME 40R 99 AHM26119

42 46306-49803 1166 SSME 41R 100 AHM26120

43 50299-50556 86 FV3 42L 100 YP_031620

44 50555-51199 215 ATV 80R 95 ALN36670

45 51512-51922 137 FV3 45L 100 YP_031623

46 51976-52482 169 Neurofilament triplet H1-like protein SSME 46L 94 AHM26124

47 52607-53023 139 ToRV 47L 100 AJR29281

48 53026-53277 84 FV3 48L 100 YP_031626

49 53386-54942 519 SSME 50L 100 AHM26127

50 55022-56707 562 FV3 51R 100 YP_031629




54 60452-61747 432 Helicase-like protein STIV 57L 99 ACF42275


56 61835-62272 146 FV3 56R 99 YP_031635


58 64197-64910 238 FV3 58R 100 YP_031637

59 65461-66519 353 FV3 59L 100 YP_031638


61 69731-69913 61 FV3 61L 100 YP_031640


63 74408-74902 165 dUTPase FV3 63R 100 YP_031642


65 75722-75886 55 FV3 65L 100 YP_031644

54


ORF

Position (nt

range)

Product


Best BLAST hit


66 75883-76434 184 FV3 66L 100 YP_031645


68 77935-78222 96 FV3 68R 100 YP_031647

69 78358-78624 89 FV3 69R 100 YP_031648

70 78642-79016 125 FV3 70R 100 YP_031649

71 79056-79289 78 FV3 71R 100 YP_031650

72 79346-80062 239 FV3 72L 100 YP_031651


74 81615-82727 371 FV3 74L 100 YP_031653



FV3 75L 100 YP_031654

76 83076-83297 74 FV3 76R 100 YP_031655

77 83294-83641 116 FV3 77L 100 YP_031656

78 84241-84879 213 FV3 78L 100 YP_031657








86 92662-92847 62 FV3 86L 100 YP_031665

87 93200-95053 618 FV3 87L 100 YP_031666

88 95086-95538 151 Thiol oxidoreductase GGRV 65L 99 AJR29223

89 95606-96667 354 FV3 89R 99 YP_031668



92 99765-100010 82 SSME 92R 100 AHM26168

93 100192-100359 56 FV3 93L 100 YP_031672

94 100469-100936 156 FV3 94L 100 YP_031673


96 102921-103592 224 FV3 96R 100 YP_031675



55

Table 2-8. Predicted open reading frames for the 2001 pallid sturgeon ranavirus isolate (PSRV01). RUK13 = Ranavirus United

Kingdom 13. See Table 2-3 for other taxa abbreviations.

ORF

Position (nt

range)

Product


Best BLAST hit



2 1378-2349 324 Myristylated membrane protein SSME 2L 100 AHM26081

3 2387-3226 280 RGV 3L 100 AFG73045

4 3156-4472 439 FV3 3R 100 YP_031581

5 4513-4695 61 FV3 4R 100 YP_031582


7 5745-5942 66 STIV 8R 100 ACF42227

8 6733-7161 143 STIV 9L 96 ACF42228



11 14353-14766 138 FV3 10R 100 YP_031588

12 15116-15328 71 FV3 11R 100 YP_031589

13 15394-16287 298 FV3 12L 100 YP_031590

14 16828-17034 69 FV3 13R 100 YP_031591

15 17049-17408 120 FV3 14R 100 YP_031592

16 17504-18472 323 ATPase FV3 15R 100 YP_031593

17 18752-19579 276 FV3 16R 100 YP_031594

18 19820-21328 503 FV3 17L 100 YP_031595

19 21366-21602 79 FV3 18L 100 YP_031596


21 24320-24766 149 PPIV 92 99 ANZ57004

22 24989-25648 220 SSME 56L 100 AHM26100


24 29077-30225 383 SSME 23R 100 AHM26102

25 30635-31705 357 SSME 24R 100 AHM26103

26 31899-32687 263 ToRV 63L 100 AJR29294

27 32755-32985 77 eIF-2alpha-like protein FV3 26R 100 YP_031604


29 36477-36965 163 FV3 28R 99 YP_031606

30 37144-37440 99 FV3 29L 100 YP_031607

31 37642-37794 51 FV3 30R 100 YP_031608

32 37856-38275 140 FV3 31R 100 YP_031609

33 38325-40436 704 Neurofilament triplet H1-like protein CGSIV 82L 100 AHA80927

34 40519-40710 64 CGSIV 33R 100 AGK44962

35 40854-41174 107 SSME 34R 99 AHM26113

56


ORF

Position (nt

range)

Product


Best BLAST hit


36 41266-41727 154 SSME 35L 100 AHM26114

37 41925-42347 141 FV3 36L 100 YP_031614



40 45323-45673 117 FV3 39R 100 YP_031617

41 45762-46310 183 SSME 40R 100 AHM26119

42 46692-50189 1166 SSME 41R 100 AHM26120

43 50685-50942 86 FV3 42L 99 YP_031620

44 50941-51582 214 ATV 80R 95 ALN36670

45 51895-52305 137 FV3 45L 100 YP_031623


47 52903-53319 139 ToRV 47L 100 AJR29281

48 53322-53573 84 FV3 48L 100 YP_031626

49 53682-55262 527 SSME 49/50L 100 AHM26127

50 55342-57027 562 FV3 51R 100 YP_031629





55 60868-61557 230 40 kDa protein RGV 57R 100 AFG73099

56 62107-62544 146 FV3 56R 100 YP_031635


58 64479-65192 238 FV3 58R 100 YP_031637

59 65743-66801 353 FV3 59L 100 YP_031638


61 70013-70195 61 FV3 61L 100 YP_031640


63 74678-75172 165 dUTPase FV3 63R 100 YP_031642


65 75962-76126 55 FV3 65L 94 YP_031644

66 76123-76674 184 FV3 66L 99 YP_031645


68 78175-78462 96 FV3 68R 99 YP_031647

69 78598-78864 89 FV3 69R 100 YP_031648

70 78882-79250 123 EHNV 35L 98 YP_009182034

71 79290-79523 78 FV3 71R 100 YP_031650

57


ORF

Position (nt

range)

Product


Best BLAST hit


72 79580-80296 239 FV3 72L 100 YP_031651


74 81851-82963 371 FV3 74L 100 YP_031653

75 82995-83249 85 LITAF/PIG7 possible membrane associated motif in LPS-

induced tumor necrosis factor alpha factor

FV3 75L 100 YP_031654

76 83312-83533 74 FV3 76R 100 YP_031655

77 83530-83877 116 FV3 77L 100 YP_031656

78 84462-85100 213 FV3 78L 100 YP_031657





83 90078-90722 215 Cytosine DNA methyltransferase SSME 83R 100 AHM26159



86 92896-93081 62 SSME 86L 100 AHM26162

87 93434-95269 612 FV3 87L 100 YP_031666


89 95822-96904 361 SSME 89R 100 AHM26165


91 98512-99699 396 Immediate early protein ICP-46 RUK13 91R 100 AIX94643

92 100056-100301 82 SSME 92R 99 AHM26168

93 100483-100650 56 FV3 93L 100 YP_031672

94 100760-101227 156 FV3 94L 100 YP_031673


96 103214-103885 224 FV3 96R 100 YP_031675



58

Table 2-9. Predicted open reading frames for the 2009 pallid sturgeon ranavirus isolate (PSRV09). NA = indicates ORF number was

not assigned. RUK13 = Ranavirus United Kingdom 13. See Table 2-3 for other taxa abbreviations.

ORF

Position (nt

range)

Product


Best BLAST hit



2 1378-2349 324 Myristylated membrane protein RGV NA 99 ABR08658

3 2387-3226 280 RGV 3L 100 AFG73045

4 3156-4472 439 FV3 3R 100 YP_031581

5 4512-4694 61 FV3 4R 100 YP_031582


7 5744-5971 76 STIV 8R 100 ACF42227

8 6732-7160 143 STIV 9L 96 ACF42228


10 11490-14336 949 NTPase/helicase FV3 10R 100 YP_031587

11 14352-14765 138 FV3 11R 99 YP_031588

12 15115-15327 71 FV3 12L 100 YP_031589

13 15393-16286 298 FV3 13R 100 YP_031590

14 16827-17033 69 FV3 14R 100 YP_031591

15 17048-17407 120 FV3 13R 100 YP_031592

16 17503-18471 323 ATPase FV3 14R 100 YP_031593

17 18751-19578 276 FV3 15R 100 YP_031594

18 19819-21327 503 FV3 17L 100 YP_031595

19 21365-21601 79 FV3 18L 100 YP_031596


21 24367-24813 149 PPIV 22 99 ANZ57004

22 25036-25695 220 SSME 56L 100 AHM26100

23 25825-28746 974 D5 family NTPase/ATPase RUK13 22R 100 AIX94587

24 29124-30272 383 SSME 23R 100 AHM26102

25 30682-31752 357 SSME 24R 100 AHM26103

26 31946-32731 262 ToRV 63L 100 AJR29294

27 33055-33756 234 eIF-2alpha-like protein ADRV 84L 100 AGV20615

28 34312-37224 971 Tyrosine kinase CGSIV 83L 99 AHA42351

29 37273-37761 163 FV3 28R 99 YP_031606

30 37940-38236 99 FV3 29L 100 YP_031607

31 38438-38590 51 FV3 30R 100 YP_031608

32 38652-39071 140 FV3 31R 100 YP_031609

33 39121-39441 107 Neurofilament triplet H1-like protein STIV 35R 100 ACF42253

59


ORF

Position (nt

range)

Product


Best BLAST hit


34 40768-40959 64 CGSIV 33R 100 AGK44962

35 41103-41423 107 SSME 34R 100 AHM26113

36 41515-41976 154 SSME 35L 100 AHM26114

37 42174-42605 144 RUK13 36L 100 AIX94599



40 45579-45929 117 FV3 39R 99 YP_031617

41 46018-46566 183 SSME 40R 99 AHM26119

42 46948-50445 1166 SSME 41R 100 AHM26120

43 50941-51198 86 FV3 42L 99 YP_031620

44 51197-51859 221 ATV 80R 95 ALN36670

45 52171-52581 137 FV3 45L 100 YP_031623

46 52635-53198 188 Neurofilament triplet H1-like protein SSME 46L 100 AHM26124

47 53323-53739 139 ToRV 47L 100 AJR29281

48 53742-53993 84 FV3 48L 100 YP_031626

49 54102-55571 490 SSME 49/50L 100 AHM26127

50 55651-57336 562 FV3 51R 100 YP_031629






56 62416-62853 146 FV3 56R 100 YP_031635


58 64788-65501 238 FV3 58R 100 YP_031637

59 66052-67110 353 FV3 59L 100 YP_031638


61 70322-70504 61 FV3 61L 100 YP_031640

62 70949-74614 1222 DNA-dependent RNA polymerase II second largest subunit RUK13 62L 100 AIX94618

63 74993-75487 165 dUTPase FV3 63R 100 YP_031642


65 76158-76247 30 PPIV 42 88 ANZ56958


67 77748-78035 96 FV3 68R 100 YP_031647

68 78171-78437 89 FV3 69R 100 YP_031648

60


ORF

Position (nt

range)

Product


Best BLAST hit


69 78455-78829 125 FV3 70R 99 YP_031649

70 78869-79102 78 FV3 71R 100 YP_031650

71 79159-79875 239 FV3 72L 100 YP_031651

72 80408-81382 325 NTPase/helicase SSME 73L 100 AHM26149

73 81558-82670 371 FV3 74L 100 YP_031653



FV3 75L 100 YP_031654

75 83019-83240 74 FV3 76R 100 YP_031655

76 83237-83584 116 FV3 77L 100 YP_031656

77 84160-84798 213 FV3 78L 100 YP_031657








85 92594-92779 62 SSME 86L 100 AHM26162

86 93132-95030 633 FV3 87L 100 YP_031666


88 95583-96644 354 SSME 89R 100 AHM26165



91 99774-100019 82 SSME 92R 100 AHM26168

92 100201-100368 56 FV3 93L 100 YP_031672

93 100478-100945 156 FV3 93L 100 YP_031673


95 102932-103603 224 FV3 96R 100 YP_031675



61



ORF

Position (nt

range)

Product


Best BLAST hit



2 1378-2352 325 Myristylated membrane protein GGRV 2L 99 AJR29160

3 2390-3229 280 RGV 3L 100 AFG73045

4 3159-4475 439 FV3 3R 100 YP_031581

5 4515-4697 61 FV3 4R 100 YP_031582


7 5747-5974 76 STIV 8R 100 ACF42227

8 6735-7163 143 STIV 9L 96 ACF42228



11 14355-14768 138 FV3 10R 100 YP_031588

12 15118-15330 71 FV3 11R 100 YP_031589

13 15396-16289 298 FV3 12L 100 YP_031590

14 16830-17036 69 FV3 13R 100 YP_031591

15 17051-17410 120 FV3 14R 100 YP_031592

16 17506-18474 323 ATPase FV3 15R 100 YP_031593

17 18754-19581 276 FV3 16R 100 YP_031594

18 19822-21330 503 FV3 17L 100 YP_031595

19 21368-21604 79 FV3 18L 100 YP_031596


21 24316-24762 149 PPIV 22 99 ANZ57004

22 24985-25644 220 SSME 56L 100 AHM26100


24 29073-30221 383 SSME 23R 100 AHM26102

25 30631-31701 357 SSME 24R 100 AHM26103

26 31895-32680 262 ToRV 63L 100 AJR29294



29 37222-37710 163 FV3 28R 99 YP_031606

30 37889-38185 99 FV3 29L 100 YP_031607

31 38387-38539 51 FV3 30R 100 YP_031608

32 38601-39020 140 FV3 31R 100 YP_031609

33 39070-41202 711 Neurofilament triplet H1-like protein CMTV_NL 76L 100 AIW68569

34 41285-41476 64 CGSIV 33R 100 AGK44962

62


ORF

Position (nt

range)

Product


Best BLAST hit


35 41620-41940 107 SSME 34R 100 AHM26113

36 42032-42493 154 SSME 35L 99 AHM26114

37 42691-43122 144 RUK13 36L 100 AIX94599

38 43517-44152 212 NIF/NLI interacting factor SSME 37R 100 AHM26116


40 46096-46446 117 FV3 39R 100 YP_031617

41 46535-47083 183 SSME 40R 99 AHM26119

42 47465-50962 1166 SSME 41R 100 AHM26120

43 51458-51715 86 FV3 42L 100 YP_031620

44 51714-52316 201 ATV 80R 95 ALN36670

45 52628-53038 137 FV3 45L 100 YP_031623


47 53618-54034 139 ToRV 47L 100 AJR29281

48 54037-54288 84 FV3 48L 100 YP_031626

49 54397-55929 511 SSME 49L 100 AHM26127

50 56009-57694 562 FV3 51R 100 YP_031629






56 62774-63211 146 FV3 56R 100 YP_031635


58 65146-65859 238 FV3 58R 100 YP_031637

59 66410-67468 353 FV3 59L 100 YP_031638


61 70680-70862 61 FV3 61L 100 YP_031640


63 75343-75837 165 dUTPase FV3 63R 100 YP_031642


65 76657-76821 55 FV3 65L 96 YP_031644

66 76818-77369 184 FV3 66L 99 YP_031645

63


ORF

Position (nt

range)

Product


Best BLAST hit



68 78870-79157 96 FV3 68R 100 YP_031647

69 79293-79559 89 FV3 69R 100 YP_031648

70 79577-79951 125 FV3 70R 99 YP_031649

71 79991-80224 78 FV3 71R 100 YP_031650

72 80281-80997 239 FV3 72L 100 YP_031651


74 82597-83709 371 FV3 74L 100 YP_031653



FV3 75L 100 YP_031654

76 84058-84279 74 FV3 76R 100 YP_031655

77 84276-84623 116 FV3 77L 100 YP_031656

78 85199-85837 213 FV3 78L 100 YP_031657








86 93633-93818 62 SSME 86L 100 AHM26162

87 94171-96042 624 FV3 87L 100 YP_031666


89 96595-97677 361 SSME 89R 100 AHM26165



92 100791-101036 82 SSME 92R 100 AHM26168

93 101218-101385 56 FV3 93L 100 YP_031672

94 101495-101962 156 FV3 94L 100 YP_031673


96 103950-104621 224 FV3 96R 100 YP_031675



64



ORF

Position (nt

range)

Product


Best BLAST hit



2 1378-2352 325 Myristylated membrane protein GGRV 2L 99 AJR29160

3 2390-3229 280 RGV 3L 100 AFG73045

4 3159-4475 439 FV3 3R 100 YP_031581

5 4515-4697 61 FV3 4R 100 YP_031582


7 5747-5974 76 STIV 8R 100 ACF42227

8 6735-7163 143 STIV 9L 96 ACF42228



11 14355-14768 138 FV3 10R 100 YP_031588

12 15118-15330 71 FV3 11R 100 YP_031589

13 15396-16289 298 FV3 12L 100 YP_031590

14 17052-17411 120 FV3 14R 100 YP_031592

15 17507-18475 323 ATPase FV3 15R 100 YP_031593

16 18755-19582 276 FV3 16R 100 YP_031594

17 19823-21331 503 FV3 17L 100 YP_031595

18 21369-21605 79 FV3 18L 100 YP_031596

19 21657-24278 874 SSME 19R 99 AHM26098

20 24326-24772 149 Serine/threonine protein kinase PPIV 22 99 ANZ57004

21 24995-25654 220 SSME 56L 100 AHM26100

22 25784-28705 974 SSME 22R 100 AHM26101


24 30641-31711 357 SSME 24R 100 AHM26103

25 31905-32690 262 ToRV 63L 100 AJR29294



28 37232-37720 163 FV3 28R 99 YP_031606

29 37899-38195 99 FV3 29L 100 YP_031607

30 38397-38549 51 FV3 30R 100 YP_031608

31 38611-39030 140 FV3 31R 100 YP_031609

32 39080-40822 581 Neurofilament triplet H1-like protein SSME 32R 100 AHM26111

33 40905-41096 64 CGSIV 33R 100 AGK44962

34 41240-41560 107 SSME 34R 100 AHM26113

35 41652-42113 154 SSME 35L 99 AHM26114

65


ORF

Position (nt

range)

Product


Best BLAST hit


36 42311-42742 144 RUK13 36L 100 AIX94599

37 43137-43772 212 NIF/NLI interacting factor SSME 37R 100 AHM26116


39 45716-46066 117 FV3 39R 100 YP_031617

40 46155-46703 183 SSME 40R 99 AHM26119

41 47085-50582 1166 SSME 41R 100 AHM26120

42 51078-51335 86 FV3 42L 100 YP_031620

43 51334-51921 196 ATV 80R 95 ALN36670

44 52233-52643 137 FV3 45L 100 YP_031623


46 53223-53639 139 ToRV 47L 100 AJR29281

47 53642-53893 84 FV3 48L 100 YP_031626

48 54002-55573 524 SSME 49L 100 AHM26127

49 55653-57338 562 FV3 51R 100 YP_031629






55 62418-62855 146 FV3 56R 100 YP_031635


57 64790-65503 238 FV3 58R 100 YP_031637

58 66054-67112 353 FV3 59L 100 YP_031638


60 70324-70506 61 FV3 61L 100 YP_031640


62 74991-75485 165 dUTPase FV3 63R 100 YP_031642


64 76305-76469 55 FV3 65L 96 YP_031644

65 76466-77017 184 FV3 66L 99 YP_031645


67 78518-78805 96 FV3 68R 100 YP_031647

68 78941-79207 89 FV3 69R 100 YP_031648

69 79225-79599 125 FV3 70R 99 YP_031649

70 79639-79872 78 FV3 71R 100 YP_031650

71 79929-80645 239 FV3 72L 100 YP_031651

66


ORF

Position (nt

range)

Product


Best BLAST hit



73 82255-83523 423 ATV 29R 96 ALN37132



FV3 75L 100 YP_031654

75 83872-84093 74 FV3 76R 100 YP_031655

76 84090-84437 116 FV3 77L 100 YP_031656

77 85013-85651 213 FV3 78L 100 YP_031657








85 93447-93632 62 SSME 86L 100 AHM26162

86 93985-95784 600 SSME 87L 100 AHM26163


88 96337-97419 361 SSME 89R 100 AHM26165



91 100533-100778 82 SSME 92R 100 AHM26168

92 100960-101127 56 FV3 93L 100 YP_031672

93 101237-101704 156 FV3 94L 100 YP_031673


95 103692-104363 224 FV3 96R 100 YP_031675



67

Table 2-12. Predicted open reading frames for the Russian sturgeon ranavirus isolate (RSRV). RUK13 = Ranavirus United Kingdom

13. See Table 2-3 for other taxa abbreviations.

ORF

Position (nt

range)

Product


Best BLAST hit



2 1378-2349 324 Myristylated membrane protein SSME 2L 99 AHM26081

3 2387-3226 280 RGV 3L 100 AFG73045

4 3156-4472 439 FV3 3R 100 YP_031581

5 4513-4695 61 FV3 4R 100 YP_031582

6 5525-5953 143 STIV 9L 96 ACF42228



9 13145-13558 138 FV3 10R 100 YP_031588

10 13908-14120 71 FV3 11R 100 YP_031589

11 14186-15079 298 FV3 12L 100 YP_031590

12 15620-15826 69 FV3 13R 100 YP_031591

13 15841-16197 119 STIV 15R 100 ACF42234

14 16294-17262 323 ATPase FV3 15R 100 YP_031593

15 17542-18369 276 FV3 16R 100 YP_031594

16 18610-20118 503 FV3 17L 100 YP_031595

17 20156-20392 79 FV3 18L 99 YP_031596

18 20444-23086 881 Serine/threonine protein kinase SSME 10L 99 AHM26098

19 23134-23580 149 PPIV 22 100 ANZ57004

20 23803-24462 220 SSME 21L 100 AHM26100


22 27891-29039 383 STIV 23R 98 ACF42246

23 29421-30518 366 RUK13 24R 99 AIX94589

24 30711-31493 261 ToRV 63L 100 AJR29294



27 36052-36540 163 FV3 28R 100 YP_031606

28 36719-37015 99 FV3 29L 100 YP_031607

29 37217-37369 51 FV3 30R 100 YP_031608

30 37431-37850 140 FV3 31R 100 YP_031609

31 37900-39975 692 Neurofilament triplet H1-like protein CGSIV 82L 100 AHA80927

32 40058-40249 64 CGSIV 33R 100 AGK44962

33 40393-40713 107 SSME 34R 98 AHM26113

34 40806-41267 154 SSME 35L 100 AHM26114

35 41465-41896 144 RUK13 36L 100 AIX94599

68


ORF

Position (nt

range)

Product


Best BLAST hit




38 44871-45221 117 FV3 39R 100 YP_031617

39 45310-45858 183 SSME 40R 100 AHM26119

40 46240-49737 1166 SSME 41R 100 AHM26120

41 50233-50490 86 FV3 42L 100 YP_031620

42 50489-51212 241 ATV 80R 95 ALN36670

43 51524-51934 137 FV3 45L 99 YP_031623


45 52514-52930 139 ToRV 47L 100 AJR29281

46 52933-53184 84 FV3 48L 100 YP_031626

47 53293-54825 511 SSME 49L 100 AHM26127

48 54905-56590 562 FV3 51R 100 YP_031629






54 61750-62187 146 FV3 56R 100 YP_031635

55 62301-63797 499 Phosphotransferase PPIV 52 99 ANZ56968

56 64251-64805 185 CGSIV 37L 100 AGK44973

57 65386-66024 213 US22 family protein ADRV 49L 99 AGV20580

58 66423-67481 353 RUK13 59L 99 AIX94616


60 70749-70874 42 RGV 64L 100 AFG73106


62 75361-75855 165 dUPTase FV3 63R 100 YP_031642


64 76645-76809 55 FV3 65L 100 YP_031644

65 76806-77357 184 FV3 66L 100 YP_031645


67 78858-79145 96 FV3 68R 99 YP_031647

68 79281-79547 89 FV3 69R 100 YP_031648

69 79565-79939 125 FV3 70R 99 YP_031649

70 79979-80212 78 FV3 71R 100 YP_031650

71 80269-80985 239 FV3 72L 100 YP_031651

69


ORF

Position (nt

range)

Product


Best BLAST hit



73 82568-83773 402 ADRV 34R 100 AGV20565



FV3 75L 100 YP_031654

75 84122-84343 74 FV3 76R 100 YP_031655

76 84340-84687 116 FV3 77L 99 YP_031656

77 85287-85925 213 FV3 78L 100 YP_031657








85 93708-93893 62 FV3 44R 98 YP_031665

86 94246-96045 600 SSME 87L 100 AHM26163


88 96598-97881 428 SSME 89R 100 AHM26165



91 101003-101242 80 FV3 92R 100 YP_031671

92 101424-101591 56 FV3 93L 100 YP_031672

93 101701-102168 156 FV3 94L 99 YP_031673


95 104153-104824 224 FV3 96R 100 YP_031675



70

Table 2-13. Predicted open reading frames for the white sturgeon ranavirus isolate (WSRV). NA = ORF number was not assigned.

RUK13 = Ranavirus United Kingdom 13. See Table 2-3 for other taxa abbreviations.

ORF

Position (nt

range)

Product


Best BLAST hit


1 1-771 257 Replicating factor ADRV 1R 100 AGV20532

2 1551-2582 344 Myristylated membrane protein ADRV 2L 100 AGV20533

3 2620-3459 280 CGSIV 71R 100 AGK44995

4 3489-4703 405 ADRV 4R 100 AGV20535

5 4744-4926 61 CMTV_NL 5R 100 AIW68496

6 5360-6025 222 US22 family protein CGSIV 6R 100 AHA80850

7 5997-6215 73 ADRV 7R 100 AGV20538

8 6976-7404 143 CGSIV 66R 100 AGK44994

9 7485-11369 1295 DNA-dependent RNA polymerase II largest subunit ADRV 9R 100 AGV20540

10 11732-14578 949 NTPase/helicase CGSIV 64R 100 AGK44993

11 14594-15007 138 ADRV 11R 100 AGV20542

12 15398-15616 73 CGSIV 9L 100 AHA80857

13 15647-16738 364 DNA repair enzyme RAD2 RGV NA 100 AAY43134

14 16831-17298 156 ADRV 13R 99 AGV20544

15 17408-17575 56 ADRV 14L 100 AGV20545

16 17765-18028 88 ADRV 15L 100 AGV20546

17 18331-19518 396 Immediate early protein ICP-46 CGSIV 58R 100 AGK44991

18 19642-21033 464 Major capsid protein ADRV 17L 100 AGV20548

19 21230-22243 338 CGSIV 20L 100 AHA80864

20 22311-22763 151 Thiol oxidoreductase ADRV 19L 100 AGV20550

21 22796-24613 606 CH8/96 17R 100 AJR29100

22 24966-25247 94 ADRV 21R 100 AGV20552

23 25594-26181 196 Thymidine kinase CGSIV 52R 100 AGK44988

24 26256-26993 246 Proliferating cell nuclear antigen (PCNA) CGSIV 51R 100 AGK44987

25 27395-28039 215 Cytosine DNA methyltransferase CGSIV 50R 100 AGK44986

26 28406-28693 97 Thymidylate synthase ADRV 25L 100 AGV20556

27 28991-29464 158 Immediate early protein ICP-18 ADRV 26L 100 AGV20557

28 29593-29871 93 Transcription elongation factor S-II CGSIV 48R 100 AGK44984

29 29930-31048 373 Ribonuclease III ADRV 28R 100 AGV20559

30 31672-33348 559 ATPase-dependent protease ADRV 29L 100 AGV20560

31 33484-34122 213 ADRV 30R 98 AGV20561

32 34822-35169 116 FV3 77L 99 YP_031656

33 35166-35387 74 CGSIV 43R 100 AGK44982



ADRV 33R 100 AGV20564

71


ORF

Position (nt

range)

Product


Best BLAST hit


35 37079-38053 325 CH8/96 29R 100 AJR29112

36 38513-39229 239 ADRV 36R 100 AGV20567

37 39286-39519 78 CGSIV 38R 100 AGK44980

38 39560-39934 125 CMTV 37L 100 AFA44942

39 39952-40218 89 ADRV 39L 100 AGV20570

40 40350-40619 90 ADRV 41L 100 AGV20572

41 40655-41818 388 Ribonucleotide reductase beta subunit CMTV 40R 100 AFA44945

42 42221-42508 96 Caspase recruitment domain protein CMTV_NL 43L 98 AIW68535

43 42603-43097 165 dUTPase CGSIV 32R 99 AGK44976

44 43477-47142 1222 DNA-dependent RNA polymerase II second largest subunit ADRV 46R 100 AGV20577

45 47769-50810 1014 DNA polymerase CGSIV 29R 100 AGK44974

46 50969-52027 353 ADRV 48R 100 AGV20579

47 52427-53065 213 US22 family protein ADRV 49L 100 AGV20580

48 53648-54202 185 CGSIV 37L 100 AGK44973

49 54656-56152 499 Phosphotransferase CGSIV 26R 100 AGK44972

50 56634-56783 50 Rmax 55 100 YP_009272766

51 56791-58086 432 Helicase-like protein ADRV 54R 100 AGV20585


53 58521-60089 523 Myristylated membrane protein CGSIV 21R 100 AGK44970

54 60425-61483 353 3beta-hydroxy-delta 5-C27 steroid oxidoreductase-like protein CGSIV 59L 100 AHA80903

55 61740-63425 562 ADRV 61L 100 AGV20592

56 63505-65058 518 ADRV 62R 100 AGV20593

57 65166-65417 84 ADRV 63R 100 AGV20594

58 65420-65836 139 ToRV 50R 100 AJR29281

59 65961-66227 89 Neurofilament triplet H1-like protein FV3 46L 100 YP_031624

60 66390-66800 137 ADRV 66R 100 AGV20597

61 66928-67773 282 STIV 47L 100 ACF42265

62 68236-71733 1166 CGSIV 11R 100 AGK44968

63 71823-71987 55 CMTV_NL 68 95 AIW68559

64 72240-72875 212 CGSIV 51L 98 AGK44967

65 72988-73314 109 CH8/96 52L 100 AJR29135

66 73420-75117 566 Ribonucleoside reductase alpha subunit CH8/96 53L 100 AJR29136

67 75256-75897 214 NIF/NLI interacting factor PPIV 72 100 ANZ56987

68 76291-76581 97 ADRV 73R 100 AGV20604

69 77019-77417 133 PPIV 75 95 ANZ57022

70 77430-77750 107 CGSIV 4R 100 AGK44963

72


ORF

Position (nt

range)

Product


Best BLAST hit


71 77892-78083 64 CGSIV 3R 100 AGK44962

72 78166-79770 535 Neurofilament triplet H1-like protein PPIV 79 100 ANZ56993

73 79820-80239 140 TFV 32R 99 ABB92294

74 80301-80453 51 PPIV 81 94 ANZ56994

75 80835-81131 99 ADRV 81R 100 AGV20612

76 81310-81798 163 CGSIV 82L 100 AHA42350



79 86209-86994 262 ToRV 63L 100 AJR29294

80 87188-88285 366 RUK13 24R 100 AIX94589

81 88668-89816 383 FV3 23R 99 YP_031601

82 90194-93121 976 D5 family NTPase/ATPase CH8/96 65L 100 AJR29148

83 93251-93910 220 FV3 21L 100 YP_031599

84 94147-94593 149 FV3 20R 100 YP_031598

85 94641-97181 847 Serine/threonine protein kinase CGSIV 96L 99 AHA80941

86 97234-97470 79 ADRV 92R 100 AGV20623

87 97507-99015 503 ADRV 93R 100 AGV20624

88 99256-99906 217 CGSIV 102L 100 AHA80947

89 100362-101309 316 ATPase CGSIV 80R 100 AGK44999

90 101406-101765 120 CGSIV 79R 100 AGK44998

91 101780-101986 69 CGSIV 105L 100 AHA80950

92 102076-102270 65 CGSIV 106R 100 AHA80951

93 102531-103424 298 CoIV 96 100 ANZ57123

94 103490-103702 71 CMTV 99L 100 AFA45004

95 104182-104868 229 CH8/96 74R 100 AJR29157

96 104934-105347 138 Myeloid cell leukemia protein ADRV 101R 100 AGV20632

73

Figure 2-1. Cytopathic effect (CPE) typical of the 12 ranavirus isolates characterized within this

chapter. A) An uninfected control flask confluent with EPC cells. B) Advanced CPE

characterized by rounded, refractile cells and the formation of plaques.

A

B

74

Figure 2-2. Transmission electron microscopy photomicrographs illustrating typical ranavirus

virion morphogenesis. A) Characteristic ranaviral assembly site and adjacent

paracrystalline array within the cytoplasm of the host cell. B) Ranaviral nucleocapsids

budding through host plasma membrane to obtain an envelope. C) Characteristic

hexagonal ranaviral nucleocapsids with electron-dense cores. Photos courtesy of Dr.

Vsevolod Popov.

A

B

C

75

A.

Figure 2-3. Maximum likelihood (ML) cladogram and phylogram depicting the relationships of

28 ranaviruses based on their aligned genomes. All nodes were supported by posterior

probability values of 1 from the Bayesian analysis, and 24/25 nodes were supported

by bootstrap values >70% from the ML analysis. A) Cladogram indicates bootstrap

support and posterior probability values above and below each node, respectively. B)

Phylogram branch lengths represent the number of inferred substitutions as indicated

by the scale. Refer to Table 2-3 for taxa abbreviations.

76

B.

Figure 2-3. Continued

0.02

ATV

Rmax

PSRV09

TFV

PSRV15

RGV

WSRV

FHMRV

EHNV

RCV-Z

BIV

GGRV

ToRV

STIV

SSME

FV3

PSRV01

CH8-96

PSRV13

SERV

RSRV

ADRV

CMTV_NL

NPRV

PPIV

CoIV

CMTV_E

ESV

77

Figure 2-4. Whole-genome alignments of 28 ranaviruses displaying 6 locally collinear blocks to

indicate genome arrangement. Inverted regions relative to short-finned eel ranavirus

(SERV) are set below those matching in the forward orientation. Connecting lines

collate aligned blocks, while crossing lines between taxa indicate inversion events.

Refer to Table 2-3 for taxa abbreviations.

78

CHAPTER 3

VALIDATION OF A TAQMAN REAL-TIME QUANTITATIVE PCR FOR THE

DETECTION OF RANAVIRUSES

Introduction

Since the isolation of Frog virus 3 from a Northern leopard frog (Lithobates pipiens) in

the late 1960s, detections of ranaviruses have expanded to include a range of species across three

classes of ectothermic vertebrates (Osteichthys, Amphibia, Reptilia) inhabiting temperate and

tropical environments worldwide (Duffus et al. 2015). The impact of ranaviruses on cultured and

wild populations of endangered ectothermic vertebrates is recognized as an important

conservation concern (Geng et al. 2011, Miller et al. 2011, Waltzek et al. 2014, Cunningham et

al. 2015). As a result, ranaviral disease in amphibians and disease due to the fish ranavirus

Epizootic hematopoietic necrosis virus (EHNV) are notifiable to the World Organization for

Animal Health (OIE 2016a,b).

According to the OIE diagnostic manual for ranaviral infection in amphibians (OIE

2016a), a case is suspect for ranaviral infection if the skin and/or parenchymal tissues of an

apparently healthy, moribund, or dead individual contain histological evidence of necrosis with

or without the presence of cytoplasmic basophilic inclusion bodies. A case is confirmed when

the suspect animal’s tissues or cell culture test positive via: 1) immunoperoxidase test/stain, 2)

antigen-capture ELISA, 3) PCR followed by restriction enzyme analysis (REA) or sequencing,

and/or 4) immunoelectron microscopy (tissue only). For EHNV infections in fish (OIE 2016b), a

suspect case is defined when one or more animals demonstrates characteristic histopathology

(e.g., liquefactive or coagulative necrosis) with or without the presence of cytoplasmic inclusion

bodies. Suspect EHNV cases are confirmed via PCR (Hyatt et al. 2000) with sequencing or REA,

plus one or more of the following: 1) immunoperoxidase test/stain (Reddacliff and Whittington

1996); 2) antigen-capture ELISA; 3) immunoelectron microscopy.

79

End-point PCR (conventional) and real-time quantitative PCR (qPCR) assays have been

developed for the detection of ranaviruses (Table 3-1). Most target the major capsid protein

(MCP) gene (Tidona et al. 1998, Mao et al. 1997, Hyatt et al. 2000, Marsh et al. 2002, Pallister et

al. 2007). The MCP gene is ideal for molecular diagnostic assays because it is highly conserved

and maintains stable base mutations that allow for the differentiation of species (Mao et al.

1997). Two conventional PCR assays targeting the MCP are recommended by the OIE (2016a,b)

for diagnosis of ranaviruses. One assay allows for the differentiation of Australian ranaviral

species (EHNV and BIV) from American (FV3) and European (ECV) species by performing

REA of PCR amplicons (Marsh et al. 2002). The second conventional assay targets a 580 bp

region of the MCP which can be sequenced for ranaviral species identification (Hyatt et al.

2000). Real-time qPCR assays have been designed to detect specific ranaviruses (Goldberg et al.

2003, Getchell et al. 2007, Allender et al. 2013a) or a range of ranaviruses (Pallister et al. 2007,

Holopainen et al. 2011, Jaramillo et al. 2012) (Table 3-1). Of these assays, only one SYBR

Green qPCR assay has been rigorously validated against a number of ranaviral taxa (Jaramillo et

al. 2012).

The documented increase in ranaviral epidemics negatively impacting ectothermic

vertebrate populations globally, the dissemination of previously geographically restricted

ranaviruses (e.g., BIV) through the unregulated international trade in exotic animals (Hick et al.

2016), and the recent characterization of novel ranaviral lineages all underscore the need for

diagnostic tools capable of detecting genetically diverse ranaviruses infecting a range of hosts

from disparate regions of the world (Duffus et al. 2015, Hick et al. 2016). Currently available

ranavirus-specific assays have not been validated for the full diversity of ranaviruses; thus, we

80

developed and partially validated a TaqMan qPCR assay capable of detecting most genetic

lineages of ranaviruses.


The following steps to develop and validate a ranavirus TaqMan qPCR assay were

carried out at the University of Florida’s Wildlife and Aquatic Animal Veterinary Disease

Laboratory (WAVDL) in Gainesville, FL : 1) in silico primer and probe design; 2) estimation of

the analytical sensitivity and specificity, slope, y-intercept, correlation coefficient (R2),

efficiency, and dynamic range; and 3) estimation of the repeatability and reproducibility.

Diagnostic sensitivity and specificity were evaluated at the OIE Reference Laboratory (OIERL)

for EHNV and amphibian ranaviruses at the University of Sydney in Camden, Australia. The

OIERL also tested a subset of ranaviral isolates as part of determining the analytical specificity

of the qPCR assay.

In Silico TaqMan qPCR Primer and Probe Design

Sequences for the 26 iridovirus core genes (Eaton et al. 2007) were obtained from

GenBank for 18 fully sequenced ranaviruses. Each gene was aligned in MAFFT (Katoh and Toh

2008) using default settings. Perusal of the initial alignments revealed the Santee-Cooper

ranaviruses (doctor fish virus DFV, guppy virus 6 GV6, largemouth bass virus, LMBV) and

grouper iridoviruses (grouper iridovirus GIV, Singapore grouper iridovirus SGIV) are highly

divergent from other ranaviruses and thus, they were excluded from all gene alignments. The

alignments were imported into Geneious 7.0 to generate a consensus sequence with the threshold

set to 100% (Kearse et al. 2012). The consensus sequences for each gene were then imported into

PrimerExpress v2.0 (Applied Biosystems) to design pan-ranaviral primers and hydrolysis probes

using default settings. The consensus sequence of the major capsid protein returned the only

suitable primers (RanaF1 and RanaR1) and probe (RanaP1) combination (Table 3-2). The

81

homologous MCP sequences were then obtained for an additional 18 partially sequenced

ranaviruses and the resulting 36 sequences were then aligned and visualized in Bioedit v7.2.5

using default settings (Figure 3-1, Table 3-3).

Detection of Ranaviral DNA Using the qPCR Assay

At the WAVDL, qPCR assays were prepared using TaqMan® Fast Universal PCR

Master Mix 2X (Applied Biosystems). The master mix consisted of 20 μL per reaction

containing 0.36 μM of each primer, 0.1 μM of probe, 4 μL of nucleic acid template, 10 μL of

universal qPCR mix (TaqMan® Fast Universal PCR Master Mix 2X, Applied Biosystems), and

3 μl of molecular grade water. Reactions were run on a 7500 Fast Real-Time PCR System

(Applied Biosystems) using the standard Fast protocol thermocycling conditions: 95°C for 20 s

followed by 45 cycles at 95°C for 3 s and 60°C for 30 s. A threshold cycle (Ct) was calculated

and interpreted as a positive result for samples if the ROX normalised FAM signal exceeded the

threshold assigned by the Applied Biosystems software

Polypropylene plates (Olympus Plastics, Genesee Scientific) sealed with 50 µm

polyolefin films (ThermalSeal RTS, Excel Scientific) included at least 2 no template negative

controls (molecular grade water). Each sample was run in triplicate and singly with a 18s rRNA

exogeneous control assay (Applied Biosystems Assay ID Hs99999901_s1) with a final reaction

concentration of 200 nM of each primer and 250 nM of probe. An MGB quencher was added to

the probe to increase specificity, reduce background fluorescence and increase signal-to-noise

ratio. A 10-fold dilution series of Frog virus 3 (FV3) MCP linearized plasmid DNA (107-10

copies) isolated from a 2015 hatchery epidemic in pallid sturgeon (Scaphirhynchus albus)

(Chapter 2) was amplified in triplicate on each plate. Efficiency was calculated as 10-1/slope

-1

(Bustin et al. 2009).

82

The qPCR assay was also evaluated in the OIERL. Samples were tested in duplicate 20

µl reactions using Path-ID qPCR Master Mix (ThermoFisher Scientific), 0.9 µM of each primer,

0.25 µM probe, and 4 µl of nucleic acid template. Thermocycling was performed on a Mx3000

qPCR system (Strategene) with the following conditions: 95°C for 10 min followed by 40 cycles

of 95°C for 30 s and 60°C for 45 s. A threshold cycle (Ct) was calculated and interpreted as a

positive result for samples if the ROX normalised FAM signal exceeded the threshold assigned

by the MxPro software (Stratagene).

Estimation of the qPCR Assay Slope, Y-Intercept, Correlation Coefficient (R2), Efficiency,

Dynamic Range, Analytical Sensitivity, Repeatability, Reproducibility, and Analytical

Specificity

Triplicate 10-fold dilutions of the FV3 plasmid standard were used in each of the 12

experiments (plates) to estimate the slope, y-intercept, correlation coefficient (R2), efficiency,

dynamic range, analytical sensitivity, repeatability, and reproducibility of the qPCR assay. The

qPCR assay limit of detection (analytical sensivitiy) was defined as the lowest dilution at which

50% of positive samples (wells) were detected (OIE 2016c). The percent coefficient of variation

(CV% = (Std/mean) x 100%)) for intra-assay (repeatability) and inter-assay (reproducibility)

variability were calculated as the mean and standard deviation of the Ct values within

(repeatability) or among (reproducibility) the 12 experiments.

To estimate the qPCR assay analytical specificity 36 reference ranaviruses were cultured

either at the WAVDL (25 isolates) or the OIERL (11 isolates) (Table 3-3). Ranaviruses were

amplified in either epithelioma papulosum cyprini (EPC) or bluegill fry (BF-2) cell lines

maintained in Minimal Essential Medium with 10% fetal bovine serum at 23°C until complete

destruction of the cell monolayer was observed. DNA was extracted from tissue culture

supernatants using a QIAcube with the DNeasy Blood and Tissue Extraction protocol (Qiagen).

Extracted DNA was quantified using a Qubit® 3.0 Fluorometer (LifeTechnologies). DNA from

83

the 36 reference libraries was used to determine the detectability of each isolate. The TaqMan

qPCR assay was also tested at the WAVDL against DNA from tissues infected with related

iridoviruses in the genera Lymphocystivirus (LCDV) and Megalocytivirus (ISKNV) and three

alloherpesviruses in the genus Cyprinivirus (CyHV1-3) (Table 3-3). The LCDV infected tissue

DNA was extracted from the fin of a longnose butterflyfish (Forcipiger flavissimus) displaying

the stereotypical gross proliferative lesions typical for the disease and was confirmed by PCR

and Sanger sequencing of the partial viral major capsid protein gene using the primers

LymphoF1 (5’-TGGTTCAGTAAATTRCCRGT-3’) and Lympho R1 (5’-

CCCATYAAWCGACGTTCYTC-3’) (unpublished). The ISKNV and CyHV1-3 infected tissue

DNA samples have been described by Subramaniam et al. (2016a) and Viadanna et al. (2017),

respectively.

Estimation of the qPCR Assay Diagnostic Sensitivity and Specificity

DNA was extracted and purified from tissue homogenates of pooled kidney-liver-spleen

from EHNV-exposed redfin perch (Perca fluviatilis) (Becker et al. 2016), Murray–Darling

rainbowfish (Melanotaenia fluviatilis), eastern mosquitofish (Gambusia holbrooki), freshwater

catfish (Tandanus tandanus), Macquarie perch (Macquaria australasica), and silver perch

(Bidyanus bidyanus) and corresponding EHNV free control fish as previously described (Becker

et al. 2013) (Table 3-4). Briefly, samples were determined to be positive or negative for EHNV

based on isolation of the virus in BF-2 cells. Cytopathic effect (CPE) was confirmed to be due to

EHNV by PCR (Marsh et al. 2002) or quantitative PCR (Jaramillo et al. 2012) on infected tissue

culture supernatant. Samples were determined to be negative for EHNV if virus isolation was

negative after three passes in cell culture. The purified nucleic acid samples from tissue

homogenates had been archived at -80°C prior to this study. In total, 106 known positive and 80

84

negative fish tissue samples were used to estimate the diagnostic sensitivity and specificity of the

qPCR assay assay (Table 3-4).

Results

In Silico TaqMan qPCR Primer and Probe Design

Our initial examination of the in silico specificity of the RanaF1 primer revealed two or

three mismatches in nine ranaviral isolates (Cum5, Cum30, BUK2, BUK4, GV, Mat2, Mg1,

RUK11, TEV) originally sequenced by Hyatt and colleagues (2000). The deposited Sanger

sequences for these isolates (Table 3-3) all display a CG at nucleotide positions four and five in

Figure 3-1. However, comparison of the Sanger sequencing reported in Hyatt et al. (2000) with

complete genome sequencing of some of the same isolates (BIV, EHNV, TEV) suggest the

aforementioned CG is actually a GC as observed in every other ranavirus reported in Figure 3-1

with the exception of the divergent ranaviruses DFV, GV6, and LMBV that display a GT at

positions four and five (Jancovich et al. 2010, Hick et al. 2016, Waltzek unpublished). In

addition to the aforementioned mismatches, RUK11 and RUK13 both possess a third primer

mismatch at position fifteen in Figure 3-1 encoding an adenine (Hyatt et al. 2000). However,

genomic sequencing of the RUK13 genome (GenBank accession no. KJ538546) does not

support the previously reported Sanger sequencing that would result in primer mismatches in

positions four, five, or fifteen (Hyatt et al. 2000). Thus, positions 4 and 5 were changed to GC in

all nine ranaviral isolates mentioned above and position 15 was changed to a G in RUK11 in the

final presented version of Figure 3-1. Thus, no mismatches in the ranavirus qPCR assay primers

or probe were detected in 33/36 ranaviruses analyzed (Figure 3-1).

85

Estimation of the qPCR Assay Slope, Y-Intercept, Correlation Coefficient (R2), Efficiency,

Dynamic Range, Analytical Sensitivity, Repeatability, Reproducibility, and Analytical

Specificity

The mean parameters (+ SE) for the qPCR assay averaged over the 12 experiments

(plates) were as follows: slope = -3.42 + 0.02, Y-intercept = 40.57 + 0.26, R2 = 0.998 + 0.0006,

and efficiency = 96.82% + 0.55 (Figure 3-2a). The dynamic range of the qPCR assay was

determined to be 107-10 copies of plasmid DNA per reaction (Figure 3-2b). Amplification of the

100 standard was inconsistent, and a 10

8 standard sample produced an abnormally shaped

standard curve, presumably due to high DNA concentration. The limit of detection of the assay

(analytical sensitivity) was determined to be 10 plasmid copies of FV3 DNA (positive in 93% of

the reactions). The coefficient of variation of intra-assay and inter-assay mean CT values ranged

from <0.1-3.5% and 1.1-2.3%, respectively (Table 3-5).

The analytical specificity confirmed the in silico design of the assay (Figure 3-1), with

ranaviral DNA amplifying 33 ranaviral DNA samples belonging to 5/6 tested ranaviral species as

well as taxonomically unclassified ranaviruses (Table 3-3). The three tested divergent

ranaviruses (DFV, GV6, LMBV), iridoviruses from other genera (i.e., ISKNV in the genus

Megalocytivirus and LCDV in the genus Lymphocystivirus), and alloherpesviruses in the genus

Cyprinivirus (CyHV1-3) were all negative. The 18s rRNA internal controls were positive for all

samples.

Estimation of the qPCR Assay Diagnostic Sensitivity and Specificity

Testing of 106 known EHNV-infected samples and 80 negative samples indicated a

diagnostic sensitivity of 95% (95% confidence limits: 89.3-98.5%) and diagnostic specificity of

100% (95.5-100%) when compared to viral isolation as a standard test (Table 3-4).

86

Discussion

The availability of a validated real-time qPCR assay capable of detecting all members of

the genus Ranavirus has become an increasingly important goal given the expansion of the

geographic and host ranges of known ranaviruses (Duffus et al. 2015). The qPCR assay reported

here was optimized and partially validated to detect all known ranaviruses excluding the most

divergent species (SCRV and SGIV) and members of the other iridovirus genera

Lymphocystivirus and Megalocytivirus. SCRV and SGIV were deliberately excluded during our

initial assay design as their inclusion in the genus has been questioned (Whittington et al. 2010,

Jancovich et al. 2015) and because specific SCRV qPCR assays have previously been developed

(Goldberg et al. 2003, Getchell et al. 2007). The qPCR assay detected 33 ranaviruses including 5

of 6 recognized species (ATV, BIV, EHNV, FV3, ECV) as well as unclassified ranaviruses from

fish, amphibians, and reptiles inhabiting varied ecosystems in North and South America, Europe,

Asia, and Australia (Table 3-3). Several species and/or genetic clades were represented by more

than one isolate in testing (e.g. FHMRV, NPRV, RSRV, PSRV, and SSME isolates have been

confirmed via genomic sequencing as strains of FV3) (Chapter 2).

Although a number of qPCR assays to detect ranaviruses have been developed (Table 3-

1), some were designed to specific ranaviruses and none have been rigorously validated for

diagnostic application with the exception of a SYBR Green qPCR assay designed by Jaramillo

and colleagues (2012). Their assay was partially validated for diagnostic application and detected

all 20 ranaviruses tested. While no primer or probe mismatches were permitted in the design of

our TaqMan qPCR assay primer (Figure 3-1), mismatches were noted when mapping the SYBR

Green qPCR assay primers of Jaramillo et al. (2012) onto the ranaviral MCP sequences assessed

in this study (Figure 3-3). Primer mismatches could negatively impact assay sensitivity (Green et

al. 2012). On balance, the authors reported high assay efficiency, analytical/diagnostic sensitivity

87

and specificity, and repeatability (Jaramillo et al. 2012). Thus, the partially validated TaqMan

qPCR assay presented here and the aforementioned SYBR Green qPCR assay both lend

themselves to diagnostic applications.

The qPCR assay reported here detected a wide range of ranaviruses from different hosts

and continents; however, the assay cannot discriminate between ranaviral species or strains.

Species discrimination can be achieved using OIE recommended conventional PCR assays

targeting the MCP gene (Hyatt et al. 2000, Marsh et al. 2002). More recently, Next Generation

Sequencing technologies to sequence ranaviral genomes facilitate phylogenomic analyses to

discriminate ranaviral species or strains (Ariel et al. 2016, Hick et al. 2016, Holopainen et al.

2016, Subramaniam et al. 2016b).

In this report, we completed stages 1 (analytical characteristics) and 2 (diagnostic

characteristics) of the OIE criteria for assay development and validation (OIE 2016c).

Experiments using plasmid standards demonstrated high assay efficiency, correlation coefficient,

repeatability, reproducibility, and analytical sensitivity and specificity. The limit of detection

(LOD) of the assay was 10 copies of FV3 plasmid DNA (Table 3-5). The assay performed well

against a panel of known EHNV positive and negative fish tissue samples indicating its high

diagnostic sensitivity (95%) and specificity (100%) (Table 3-4). The five positive fish tissue

samples that tested negative (false negatives) were all samples with low EHNV viral loads that

may have been below the assay LOD. Reduced sample quality following long term storage or

PCR inhibition are possible alternative explanations for these false negative results (Jaramillo et

al. 2012). Regardless, future optimization of the assay mastermix (primer and probe

concentrations) and thermocycling conditions are planned in an effort to improve the assay

88

sensitivity with the goal of detecting 10 plasmid copies in >95% of the tested samples (93%

reported here).

In partial fulfillment of stages 3 (reproducibility) and 4 (implementation) in the OIE

criteria for assay development and validation, the qPCR assay has routinely been by the

WAVDL for the last two years, providing insights into the pathogenesis of FV3 in pallid

sturgeon Scaphirhynchus albus following experimental challenges (Chapter 4) and FV3

epidemics within cultured and wild herpetofauna. The OIERL has implemented the assay in

surveillance of cane toad populations in Australia for approximately one year, and more recently,

in other U.S. university laboratories that routinely conduct ranaviral diagnostics. A ring trial is

planned among collaborating laboratories organized through the Global Ranavirus Consortium

(www.ranavirus.org) in support of stages 3 and 4.

The reported qPCR assay offers an expedient and cost-effective test for the diagnosis of

ranaviruses in both tissue culture supernatants and tissues, making this tool particularly useful for

understanding ranaviral pathogenesis and epidemiology. As this assay was created and validated

using a diverse group of fish, amphibian, and reptile ranaviruses from around the world, it serves

as an important tool in ranaviral surveillance programs aimed at the management and mitigation

of these globally emerging pathogens.

89

Table 3-1. Quantitative PCR assays developed for the detection of ranaviruses. See Table 3-3 for taxa abbreviations.

Assay Purpose of assay

Gene

target Method Viruses evaluated

Validation for

diagnostic use Reference 1 Quantification MCP TaqMan qPCR LMBV No Goldberg et al. 2003

2 Quantification MCP TaqMan qPCR LMBV Yes Getchell et al. 2007

3 Detection and

differentiation

MCP TaqMan qPCR BIV, ECV, ESV, EHNV No Pallister et al. 2007

4 Quantification DNA pol SYBR Green qPCR EHNV, ECV, ESV, FV3, BIV, DFV, GV6,

PPIV, REV, SERV

No Holopainen et al. 2011

5 Detection and

differentiation

MCP SYBR Green qPCR BIV, Cum5, Cum6, Cum30, EHNV, ECV,

ESV, FV3, GVB3, Mat2, Mg1, MHRV,

TEV, UKIV1, WV

Yes Jaramillo et al. 2012

6 Detection and

quantification

MCP TaqMan qPCR FV3 Partial (sensitivity

and specificity)

Allender et al. 2013a

90

Table 3-2. Primers and probes designed against the ranavirus major capsid protein (MCP) gene for development of the plasmid

standard and for use in the diagnostic assay. MCP gene positions 891 and 1231 correlate to positions 98235 and 98575

in the Frog virus 3 genome (Genbank accession no. AY548484).

Primer/probe

name Primer/probe sequence

Melting

temperature (°C)

Positions in MCP

gene (5’ to 3’)

Amplicon size (nt)

including primers

RanaMCPstdF GTTCTCACACGCAGTCAAGG 53.8 891-910 359

RanaMCPstdR CGGACAGGGTGACGTTAAG 53.2 1231-1249

RanaF1 CCAGCCTGGTGTACGAAAACA 54.4 1040-1060 97

RanaR1 ACTGGGATGGAGGTGGCATA 53.8 1136-1117

RanaP1 6FAM-TGGGAGTCGAGTACTAC- MGB 47.1 1079-1095

91

Table 3-3. Panel of ranaviruses used for primer and probe design and/or validation of the TaqMan qPCR assay. Positive (+) and

negative (-) symbols indicate expected specificity with the primer/probe design or the actual qPCR TaqMan test result.

Viral species accepted by the International Committee on Taxonomy of Viruses are italicized. NE = sample not evaluated

in this study.

Virus species/isolate

Abbrevia-

tion Geographic origin

Genbank

accession

Primer/probe

design

qPCR

result

Reference

Andrias davidianus ranavirus ADRV China KC865735 + NE Chen et al. 2013

Ambystoma tigrinum virus ATV Arizona AY150217 + + Jancovich et al. 2003

Bohle iridovirus BIV Australia KX185156 + + Hick et al. 2016

Bufo bufo United Kingdom iridovirus 1 b

BUK2 United Kingdom AF157653 + + Hyatt et al. 2000

Bufo bufo United Kingdom iridovirus 3 b

BUK4 United Kingdom AF157657 + + Hyatt et al. 2000

Bufo marinus Venezuelan iridovirus 1 b

GV Venezuela AF157649 + + Hyatt et al. 2000


Mg1 Venezuela AF157677 + + Hyatt et al. 2000


Mat2 Venezuela AF157675 + + Hyatt et al. 2000


Cum5 Venezuela AF157663 + + Hyatt et al. 2000


Cum6 Venezuela NE + Hyatt et al. 2000


Cum30 Venezuela AF157661 + + Hyatt et al. 2000

Epizootic hematopoietic necrosis virus EHNV Australia FJ433873 + + Jancovich et al. 2010

European sheatfish virus ESV Germany JQ724856 + + Mavian et al. 2012b

Fathead minnow ranavirus FHMRV Arkansas NE + Waltzek et al. 2014

Frog virus 3 FV3 Wisconsin/ Minnesota AY548484 + + Tan et al. 2004

Grouper iridovirus a GIV Taiwan AY666015 - NE Tsai et al. 2005

Guppy virus 6 a GV6 USA (imported from

Asia)

FR677325 - - Ohlemeyer et al. 2011

Infectious spleen and kidney necrosis

virus a

ISKNV USA NE - Subramaniam et al. 2016a

Largemouth bass virus a

LMBV Mississippi FR682503 - - Ohlemeyer et al. 2011

Lymphocystis disease virus a

LCDV USA NE - Waltzek unpublished

Mahaffey Road virus MHRV Australia GU292010 + + Weir et al. 2012

Marbled sleepy goby virus MSGV Thailand NE + Prasankok et al. 2005

Northern pike ranavirus NPRV Ohio NE + Waltzek et al. 2014

Pike-perch iridovirus PPIV Finland KX574341 + + Holopainen et al. 2016

Pallid sturgeon ranavirus PSRV Missouri KF646249 + + Waltzek et al. 2014

Rana esculenta virus REV Italy FJ358611 + + Holopainen et al. 2009

Rana grylio virus RGV China JQ654586 + NE Lei et al. 2012

92


Virus species/isolate

Abbrevia-

tion Geographic origin

Genbank

accession

Primer/probe

design

qPCR

result

Reference

Rana temporaria United Kingdom

iridovirus 1 b

RUK11 United Kingdom

AF157645 + + Hyatt et al. 2000

Russian sturgeon ranavirus RSRV Georgia NE + Waltzek et al. 2014

Short-finned eel ranavirus SERV Italy NC_030394 + + Subramaniam et al. 2016b

Singapore grouper iridovirus a SGIV Singapore NC_006549 - NE Song et al. 2004

Softshell turtle iridovirus STIV China EU627010 + NE Huang et al. 2009

Spotted salamander Maine virus SSME Maine KJ175144 + + Morrison et al. 2014

Tadpole edema virus b

TEV North America AF157681 + + Hyatt et al. 2000

Tiger frog virus TFV China AF389451 + NE He et al. 2002

Wamena virus b

WV Australia (imported

from Indonesia)

NE + Hyatt et al. 2002

White sturgeon ranavirus WSRV California + + Waltzek et al. 2014

Zoo ranavirus ZRV Iowa KF699143 + + Cheng et al. 2014 a Divergent virus not expected to amplify with the Ranavirus TaqMan qPCR primers and probe

b Sample tested by the OIE Reference Laboratory for EHNV and Amphibian Ranaviruses

93

Table 3-4. Results for the TaqMan qPCR assay on fish tissue homogenates with EHNV infection status determined by virus isolation

and confirmed by PCR. Samples were originally determined EHNV(+) or (-) by Jaramillo et al (2012).

Description of sample Virus isolation qPCR assay

Expected

EHNV status Species of fish Positive (n) Negative (n)

- Redfin Perch (Perca fluviatilis) - 0 36

Freshwater catfish (Tandanus tandanus) - 0 4

Trout cod (Maccullochella macquariensis) - 0 40

Total, all species 0 80

+ Redfin Perch (Perca fluviatilis) + 70 1

Freshwater catfish (Tandanus tandanus) + 2 0

Mosquito fish (Gambusia holbrooki) + 23 1

Silver Perch (Bidyanus bidyanus) + 0 3

Murray-Darling rainbow fish (Melanotaenia

fluviatilis)

+ 5 0

Macquarie Perch (Macquaria australasica) + 1 0

Total, all species 101 5

94

Table 3-5. Inter-assay variability (reproducibility) of the pan-ranavirus qPCR across twelve

experiments (plates) at the WAVDL. Reactions for each plasmid standard (107-10

copies) were run in triplicate.

Standard

dilution

Mean

Ct

Standard

deviation

Coefficient of variation

(%)

Number of wells

positive (/36)

107

16.57 0.27 1.6 36

106

19.73 0.27 1.4 36

105

23.18 0.31 1.4 36

104

26.57 0.35 1.3 36

103 30.04 0.34 1.1 36

102

33.58 0.37 1.1 36

101

37.09 0.53 2.3 33

95

Figure 3-1. Aligned partial (97 bp) major capsid protein (MCP) sequences for 36 ranaviruses illustrating the in silico specificity of the

qPCR primers (RanaF1 and RanaR1) and TaqMan probe (RanaP1). Divergent ranaviruses are shown below the solid black

line. Refer to Table 3-3 for viral abbreviations and GenBank accessions.

96

A.

Amplification Plot

Figure 3-2. Quantification of a standard curve for Frog virus 3 using the TaqMan real-time polymerase chain reaction (qPCR) assay.

A) Assay amplification plot of 10-fold serial dilutions of standards ranging from 107-10 copies. A typical experiment is

shown with the blue line indicating the automatic threshold assigned by the Applied Biosystems software. The X-axis

indicates the cycle number and the Y-axis indicates the log ΔFn (normalized fluorescence). B) Standard curve generated

using triplicate 10-fold serial dilutions of the FV3 plasmid DNA standards ranging from 107-10 copies. The mean qPCR

assay parameters (+ SE) averaged over the 12 experiments (plates) are provided. The X-axis shows the log plasmid

standard copy number and the Y-axis indicates the corresponding cycle threshold (Ct) value.

97

B.

Figure 3-2. Continued

0

5

10

15

20

25

30

35

40

10 100 1,000 10,000 100,000 1,000,000 10,000,000

CT

Quan( ty

StandardPlot

Slope: -3.42 ± 0.02

Y-intercept: 40.57 ± 0.26

Correlation coefficient: 0.998 ± 0.0006

Efficiency: 96.82 ± 0.55

Copy number

CT

Standard Plot

98

Figure 3-3. Aligned partial (94 bp) major capsid protein (MCP) sequences for 36 ranaviruses illustrating in silico specificity of the

SYBR green qPCR primers developed by Jaramillo and colleagues. Divergent ranaviruses according to Jaramillo et al.

(2012) are shown below the solid black line. Refer to Table 3-3 for viral abbreviations and GenBank accessions.

99

CHAPTER 4

THE EFFECT OF WATER TEMPERATURE ON FROG VIRUS 3 DISEASE IN HATCHERY-

REARED PALLID STURGEON (SCAPHIRHYNCHUS ALBUS)

Introduction

Sturgeon belong to the family Acipenseridae, one of the oldest ray-finned fish families

with fossils dating back to the upper Cretaceous period (Gardiner 1966). They are among the

largest and longest-lived of all known fishes with adults reaching maturity late in life (Bemis et

al. 1997, Berra 2007). Sturgeon have historically been fished for their meat and caviar; however,

overfishing combined with habitat destruction and pollution have decreased wild stocks of many

species (Artyukhin 1997, Debus 1997, Baker and Borgeson 1999). All sturgeon are currently

designated CITES Appendix I or II (CITES 2012) and many are listed as endangered or critically

endangered on the IUCN Red List of Threatened Species (Billard and Lecointre 2001, IUCN

2001). Fishery restrictions have led to decreased pressure on wild sturgeon stocks over the last

two decades and aquaculture is steadily increasing to meet the demand for meat and caviar.

Currently, production of sturgeon occurs predominantly in China, Europe, Russia, and the United

States (Chebanov and Billard 2001, Bronzi et al. 2011). Multiple hatcheries in North America

aim to help replenish dwindling wild stocks of native species, including the endangered pallid

sturgeon (Scaphirhynchus albus).

Infectious agents have negatively impacted sturgeon aquaculture and restoration efforts.

The melanized fungus Veronaea botryosa has negatively impacted the production of Siberian

sturgeon (Acipenser baerii) in Florida and white sturgeon (A. transmontanus) in California

(Steckler et al. 2014). In particular, viral pathogens (e.g., adenovirus, alloherpesviruses, and

iridoviruses) have hindered production of farmed sturgeon (Hedrick et al. 1985, 1990, 1991a,

1991b, Bauer et al. 2002, Kelley et al. 2005, Kurobe et al. 2008, 2010, 2011, Waltzek et al.

2014). Multiple ranavirus epizootics have occurred at three separate North American sturgeon

100

hatcheries (Waltzek et al. 2014, Chapter 2). In 2001, 2009, 2013, and 2015 young-of-year pallid

sturgeon at the Blind Pony State Fish Hatchery in Sweet Springs, MO experienced heavy

mortalities (up to 90-100%) due to a strain of Frog virus 3 (FV3) (Waltzek et al. 2014, Chapter

2). These four FV3 epizootics resulted in economic losses >$400,000 USD and have impeded

efforts to replenish wild pallid sturgeon stocks (Jake Colehour personal communication).

Ranavirus epizootics have also caused high mortality in farmed white (Acipenser transmontanus)

and Russian (Acipenser gueldenstaedtii) sturgeon in California in 1998 and Georgia in 2004,

respectively. Koch’s postulates were fulfilled in the 2009 pallid sturgeon epizootic and the

isolate from the 2004 Russian sturgeon outbreak reproduced the disease following intracoelomic

injection of both juvenile Russian and lake (A. fulvescens) sturgeon (Robert Bakal personal

communication).

The severity of ranavirus epizootics in sturgeon hatcheries is likely the combination of

host, viral, and environmental factors (Chapter 1). Hatcheries rearing young-of-year sturgeon

under intensive conditions often experience increased infectious disease epizootics as

environmental conditions deteriorate (e.g., water quality, inappropriate water temperature,

malnutrition) (Barton et al. 1991, Chua et al. 1994, LaPatra et al. 1996, Buentello et al. 2000,

Geordialis et al. 2000, 2001, Drennan et al. 2005, Savin et al. 2011). High densities facilitate

transmission by increasing contact rates between animals as well as pathogen concentrations

within the water (Woodland et al. 2002, Brunner et al. 2007, Brenes et al. 2014, Brunner et al.

2015). Additionally, many sturgeon stocks have undergone severe genetic bottlenecks,

potentially resulting in decreased immunogenetic diversity and increased susceptibility to

pathogens (LaPatra et al. 1999, Marranca et al. 2015) as has been observed in other species on

the brink of extinction (Pearman and Garner 2005, O’Brien and Evermann 1988).

101

Temperature is a well-known factor influencing ranaviral disease in fish (Wedemeyer et

al. 1976). Whittington and Reddacliff (1995) observed that natural outbreaks of Epizootic

hematopoietic necrosis virus (EHNV) were most severe in juvenile redfin perch congregating in

shallow warm waters, and experimental infections with EHNV in rainbow trout showed highest

mortality at temperatures beyond the host’s upper threshold (Ariel et al. 2009). Watson et al.

(1998) determined that maintaining temperatures either above or below the recommended range

for white sturgeon resulted in increased mortality and/or severity of infection with white

sturgeon iridovirus. FV3 epizootics at the Blind Pony Fish State Hatchery in Missouri have been

most severe during elevated water temperatures experienced in summer months (Waltzek et al.

2014). Although temperature tolerance studies have not been extensively performed in pallid

sturgeon, a small study using 18 juveniles noted stress at temperatures above 30°C with mortality

beginning at 33°C (Blevins 2011). Shovelnose sturgeon (Scaphirhynchus platorynchus), the

closest relative to the pallid sturgeon, grow most efficiently at 22°C with steadily increasing

mortality in individuals raised above that temperature (Kappenman et al. 2009).

In this study, water temperature was examined for its effect on ranaviral disease

following bath challenges of FV3 in young-of-year pallid sturgeon. Study 1 compared the

cumulative mortality of young-of-year pallid sturgeon at two temperatures. Study 2 compared

disease progression of young-of-year pallid sturgeon at two temperatures.


Quarantine and Husbandry

The use of juvenile pallid sturgeon in these studies were approved by and conducted in

compliance with the University of Florida (UF) Institutional Animal Care and Use Committee

(IACUC, protocol 201609405). Young-of-year pallid sturgeon were obtained from the Gavins

Point Fish Hatchery in South Dakota with no prior history of ranavirus infection. Prior to

102

shipment to the UF Wildlife and Aquatic Animal Veterinary Disease Laboratory (WAVDL),

pectoral fin clips from 60 fish were individually tested and confirmed negative for Missouri

River sturgeon iridovirus by the United States Fish and Wildlife Service Bozeman Fish Health

Center using a conventional PCR assay (Kurobe et al. 2010). At the WAVDL, fish were

quarantined for six wk prior to experimentation. Fish were arbitrarily assigned to rectangular

glass holding tanks ranging from 204-455 l in volume. The tanks were supplied with flow-

through dechlorinated municipal water at 17°C and aerated with airstones. Flow-through rate

was set at 10 volume changes day-1

and fish were stocked at a density of 5 fish/ft2 of tank space.

After a 3 d acclimation period, water temperature in half of the tanks was raised to 23°C at a rate

of 1°C day-1

for the remainder of the quarantine period. Fish were fed twice daily to satiation

with a combination of commercial pelleted salmonid crumble (Otohime), frozen brine shrimp,

and frozen bloodworms. Water temperature was recorded twice daily in all tanks and dissolved

oxygen (DO) levels were measured daily using a meter (Hach Co.) until levels were determined

to be stable at ≥85% O2 saturation, after which DO levels were measured weekly. Water

chemistry analysis was performed every other day (total ammonia nitrogen, nitrite, pH) or every

week (total hardness and total alkalinity) using a multi-parameter water quality kit (Hach Co.).

Fish were monitored twice daily for morbidity and mortality.

On d 36 of quarantine, 10 fish were arbitrarily selected and sacrificed for health

assessments, which consisted of gross physical examination, microscopic examination of tissue

biopsies, bacteriology, and ranavirus screening. Immediately following collection of external

tissue (skin, fin, gill) wet mounts, fish were euthanized in 1000 mg l-1

tricaine methane sulfonate

(MS-222®, Argent Laboratories, Finquel®) buffered in an equal weight of sodium bicarbonate

and then pithed. Individuals were weighed (g) and measured (cm) for standard length (SL) and

103

fork length (FL) and a general examination was performed to identify any gross lesions.

Following the gross examination, sterile posterior kidney cultures were inoculated onto

Columbia agar with 5% sheep blood and incubated for 72 hr at 24°C for bacterial screening.

Additional wet mounts of internal organs (i.e., stomach, intestine, liver, posterior kidney, and

spleen) were taken and then the external and internal tissue samples were assessed for tissue

lesions and pathogens (i.e., bacteria, fungi, parasites) via light microscopy at 40x, 100x, and

200x magnifications. The ten individuals were screened for ranavirus using a pan-ranavirus

TaqMan qPCR assay (Stilwell et al. 2017, Chapter 3) and virus isolation in cell culture as

described below.

Internal tissues (liver, kidney, and spleen) were pooled by individual and diluted 1:25 in

Minimal Essential Medium (MEM) with 2% Fetal Bovine Serum (MEM-2). Tissue suspensions

were then homogenized using a stomacher (Seward Stomacher® 80 Biomaster) on high speed

for 2 min. Then, 200 µl of each pooled tissue homogenate was processed for DNA extraction

(described below) and the remainder received further processing for virus isolation. The

remaining homogenate was then clarified by centrifugation at 3000 x g (10 min at 4°C) to

remove cellular debris. An equal volume of clarified tissue homogenate was added to a MEM-2

antibiotic solution resulting in a final concentration of 500 IU penicillin ml−1

, 500 µg

streptomycin ml−1

, and 12.5 µg fungizone ml−1

, and 14 mM HEPES (4-(2-hydroxyethyl)-1-

piperazine ethane sulfonic acid) buffer. After incubating overnight at 4°C, the samples were

again clarified at 3000 x g (10 min at 4°C) and 10 μl of undiluted and serially diluted (10-1

-10-11

)

tissue homogenates were inoculated onto 6 replicate wells of a 96-well plate containing

confluent monolayers of EPC cells. After a 1 hr virus adsorption period, 190 μl of MEM-2 was

added to each well. Two replicate wells per dilution containing only MEM-2 served as negative

104

controls. Cells were incubated at 24°C and observed for cytopathic effect (CPE) daily for 14 d.

The 50% Tissue Culture Infectious Dose (TCID50) was calculated by endpoint analyses at 14 d

postinoculation (Ramakrishnan 2016). Samples not displaying CPE at d 14 were blind passaged

and observed for an additional 10 d before declaring the sample negative.

For ranaviral testing via qPCR, 200 µl of the pooled tissue homogenates were transferred

to lysis bead tubes (Benchmark Scientific) and processed in a TissueLyser II (Qiagen) at 30

cycles s-1

for 1 min. DNA extraction was then performed using a Qiacube (Qiagen) with the

DNeasy Blood and Tissue Extraction protocol (Qiagen). TaqMan qPCR samples were tested in

triplicate using 4 µl of DNA extract with a single 18s internal control per sample (Chapter 3).

Threshold of detection (CT) and number of copies were estimated for samples using the 7500

Fast Real-Time PCR System software program (Applied Biosystems) based on the use of a

linearized plasmid standard curve. Samples were considered positive for ranavirus if 2/3 wells

generated a CT value of ≤40.

Study 1: Effect of Water Temperature on Cumulative Mortality

After the 6 wk quarantine period, 120 sturgeon were arbitrarily selected for the first

challenge study. Forty exposed and 20 unexposed fish were used at each temperature (17°C and

23°C). Virus for the challenge study was prepared from a frozen stock of the 2015 pallid

sturgeon isolate (PSRV15) isolated in Chapter 2 and inoculated (500 µl flask−1

) into four 175

cm2

flasks containing confluent EPC cells with MEM-2 at room temperature (24°C). After 2 d,

CPE was complete and the supernatants were combined and clarified at 3000 × g (10 min at

4°C). The viral titer was determined by TCID50 endpoint analyses at 7 d postexposure (PE) as

described above (Ramakrishnan 2016).

The PSRV15 isolate or sham (negative controls – MEM only) exposures were performed

via 1 hr hour bath challenge in static, aerated systems at the two experimental temperatures (17

105

and 23°C). Following bath exposure, fish were arbitrarily placed into 84 l tanks and stocked at a

density of 10 fish per tank (2.6 fish/ft2 of tank space) for a 28 d observation period. Four exposed

replicate tanks and two control replicate tanks were used at each temperature (Figure 4-1). Tanks

were aerated with airstones and supplied with flow-through dechlorinated municipal water at

either 17 or 23°C with a turnover rate of 10 volumes d-1

. Water quality parameters (total

ammonia nitrogen, nitrate, pH, total hardness, and total alkalinity) were measured weekly

throughout the study. Fish were monitored twice daily for signs of morbidity and mortality. Dead

or moribund fish were processed individually as described above during the quarantine period

including: weight and length measurements, gross and wet mount examination of tissues,

bacteriology, and ranavirus testing of pooled liver, spleen, and posterior kidney homogenates via

TaqMan qPCR and TCID50 endpoint analyses. Moribund fish displaying advanced clinical signs

of ranavirus infection were euthanized as described above. Endpoint criteria for euthanasia

included the display of two or more of the following advanced ranaviral disease signs: external

hemorrhage of skin and/or fin, distended coelom, erratic swimming, and abnormal buoyancy

and/or orientation. To minimize potential pathogen cross-contamination among tanks and fish,

each animal was collected individually with separate nets disinfected in 2% Virkon Aquatic for

at least 2 min prior to use (Bryan et al. 2009) and euthanized in separate containers. At 28 d

postexposure (PE), surviving fish were euthanized. One fish per tank per temperature treatment

(8 exposed and 4 unexposed control fish) was processed as described above to assess the health

status and test for ranaviral infection among survivors.

Study 2: Effect of Water Temperature on Disease Progression

The experimental design of study 2 was identical to study 1 (Figure 4-1). On d 1, 3, 5, 7,

14, and 28 PE, one fish per tank was euthanized as described above, for a total of 12 fish per

sampling period (i.e., 4 exposed and 2 unexposed fish per temperature treatment). To minimize

106

potential pathogen cross-contamination among tanks and fish, each animal was collected

individually with separate nets disinfected in 2% Virkon Aquatic for at least 2 min prior to use

(Bryan et al. 2009) and euthanized in separate containers as described above.

Dead or moribund fish were processed individually as described above during the

quarantine period and study 1 (unless noted otherwise) including: weight and length

measurements, and gross and wet mount examination of tissues. Additionally, sterile instruments

were used to collect pooled samples of skin, fin, gill, and barbel for each individual. After a

ventral midline incision was made to access the coelomic cavity, a new set of autoclaved

instruments was used to collect samples of liver, spleen, posterior kidney, and heart. Using the

methods described in study 1, pooled internal (liver, kidney, spleen, heart, and pericardial

lymphomyeloid tissue) and external (skin, fin, gill, and barbel) tissue homogenates were assessed

separately for viral load by qPCR and viral titer by TCID50 endpoint analyses.

Sections of all above tissues as well as stomach, spiral colon, head, and transverse body

wall at the level of the posterior kidney were placed in 10% neutral buffered formalin for 24-48

hr for histological fixation. Calcified tissues (e.g., head, body wall, fin, and gill) underwent an

additional 48-96 hr decalcification step in 0.5M ethylenediaminetetraacetic acid (EDTA) pH

buffered to 8.0 (Fisher BioReagents). Formalin-fixed, paraffin-embedded sections of tissues

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

(H&E). Slides were examined for histopathological changes consistent with ranavirus including:

hemorrhage; necrosis of hematopoietic tissues, vascular endothelium, and epithelial cells; and

cytoplasmic basophilic to amphophilic inclusions (Reddacliff and Whittington 1996, Waltzek et

al. 2014, Miller et al. 2015). To assist with the histopathological interpretations, a subset of the

fixed tissues was subjected to a semi-validated ranavirus in situ hybridization (ISH) assay (see

107

below). Histological examination (H&E and ISH) was performed on tissue sections from 11 fish

(55 slides per stain technique), consisting of one exposed fish per temperature treatment per

sampling day. Fish with the highest qPCR ranaviral copy number on each day within their

treatment groups were preferentially chosen to have the greatest probability of having positive

labeling within samples.

Preliminary Development Of A Ranavirus In Situ Hybridization Assay

In situ oligonucleotide probes were designed based on the 1392 nt coding sequence for

the ranavirus major capsid gene (MCP). Previously designed assays suggest the MCP is a good

candidate for ranavirus diagnostics as it exhibits 94-100% nucleotide sequence identity (out of

1392 nt) across all ranaviruses, excluding the highly divergent Santee-Cooper ranavirus and

Singapore grouper iridovirus (Jancovich et al. 2015, Chapter 3). The probe was designed by

Advanced Cell Diagnostics based on the full MCP nucleotide sequence for a Frog virus 3

isolated from the 2009 pallid sturgeon epizootic (PSRV09) (Waltzek et al. 2014, Chapter 2).

Formalin-fixed, paraffin-embedded sections of tissues were sectioned at 5 μm, mounted onto

Superfrost™ Plus glass slides (Fisherbrand™) and prepared for the RNAscope® in situ

hybridization (ISH) protocol according to the manufacturer instructions. Mounted tissue slides

were placed in a dry oven for 60°C for 1 hr within one wk prior to the RNAscope® assay. The

ISH assay was performed using the HybEZ II oven and RNAscope® 2.5 HD RED Reagent Kit

(Advanced Cell Diagnostics) according to the manufacturer’s instructions (Wang et al. 2012).

The following steps were performed sequentially: 1) sample deparaffinization and dehydration,

2) tissue section pretreatments, 3) target probe hybridization, 4) signal amplification, 5) signal

detection, 6) counterstaining, and 7) slide mounting. Known FV3 (PSRV09)-positive and -

negative formalin-fixed, paraffin-embedded (FFPE) pallid sturgeon tissue sections from an

experimental exposure study (Waltzek et al. 2014) served as procedural controls (Waltzek et al.

108

2014). Rather than using the stock red chromagen supplied with the RNAscope® 2.5 HD RED

Reagent Kit assay (Advanced Cell Diagnostics), a permanent red chromagen and hematoxylin

(Dako Cytomation) were used for labeling and counterstaining, respectively, to ensure long-term

preservation of sections. Permanent reagents were integrated into the protocol based on methods

developed by Draghi et al. (2010). Additionally a 5 min bath with 5 mM tetramisole

hydrochloride (Sigma-Aldrich Corp.) was performed immediately prior to red chromagen

labeling to reduce endogenous alkaline phosphatase activity and thus minimize background red

chromagen staining.

Statistical Analyses

For study 1, statistical analysis (unpaired T-test) was performed to determine significance

of mean cumulative mortality (proportion of dead fish to total fish) within and among the two

treatment groups (17°C and 23°C). In study 2, one-way, repeated-measures analysis of variance

(ANOVA) was performed to determine the significance of TaqMan qPCR viral copy numbers

between the 17°C and 23°C temperature treatments. Prior to testing, the residuals of the model

were checked for normality. Analyses were performed using SAS® version 9.3 (SAS Institute

Inc.) Numerical data are represented as mean ± standard error (SE), and statistical differences

were considered significant at P< 0.05.

Results

Quarantine and Husbandry

Water chemistry values were within normal limits during quarantine and both studies.

Parameter range values during the studies were as follows: 17°C DO: 7.65-8.81 mg l-1

, 17°C

oxygen saturation: 78.1-89.9%, 23°C DO: 7.33-9.05 mg l-1

, 23°C oxygen saturation: 84.3-

100.4%, total ammonia nitrogen: 0-0.2 mg l-1

, unionized ammonia: 0-0.006 mg l-1

, nitrite: 0 mg l-

1, pH: 7.75-8.0, total hardness: 136.8-153.9 mg l

-1, total alkalinity: 68.4 mg l

-1. Water

109

temperatures were successfully maintained at 17±1°C and 23±1°C in the cold- and warmwater

tanks, respectively.

Health assessments performed during the quarantine period revealed no abnormalities on

gross or wet mount examinations. The mean weight (n=10) was 17.7 g (range: 14.3-22.3 g) and

mean fork length was 24.4 cm (range: 20.0-27.5 cm). Bacteriology results were negative and all

fish were ranavirus-negative via TaqMan qPCR and virus isolation.

Study 1: Effect of Water Temperature on Cumulative Mortality

The mean weight for the 120 fish used in the cumulative mortality study was 28.7 g

(range 14.4-50 g) and mean fork length was 21.6 cm (range: 16.8-25.5 cm). Infection dose for

each temperature was 60 ml in 60 l water containing 40 exposed fish, for a final dose of 2.14 x

106 TCID50 ml

-1 water. During the 28 d PE observation period, no morbidity or mortality was

observed in the 40 exposed fish held at the cooler water temperature or in the 20 unexposed

control fish at either water temperature. In contrast, 42.5% cumulative mortality (17/40) was

observed in the exposed fish (Figure 4-2) held at the warmer temperature. Of the affected fish, 3

fish were found dead while euthanasia endpoints were utilized for 14 fish. The majority of

mortalities occurred on days 3-6 PE (Figure 4-2). The most frequently observed abnormalities in

affected fish were lethargy (8/17 fish) and hemorrhages of the skin and/or fins (13/17 fish).

Additional findings occurring in fewer than 25% of sampled warmwater fish included: excess

mucus production on the skin and/or fins, sloughing/torn fins, proliferative cottony material on

external lesions, and abnormal orientation (i.e., lying upside down on the tank floor).

Necropsy, qPCR, and virus isolation were performed on 15/17 exposed 23°C fish, as two

dead fish were significantly autolyzed. Necropsy of the 15 examined individuals revealed several

gross lesions consistent with ranavirus infections including: enlarged and/or abnormally pale,

darkened, or mottled color of the posterior kidney (8/15), spleen (6/15), liver (6/15), and intestine

110

(1/15); frank blood within the coelom (5 fish); empty gastrointestinal tract (3/15); and congested

(i.e., abnormally dark) gills (2/15) (Figure 4-3). Bacterial cultures yielded growth in 4/15 fish

after 24 hr that was identified via conventional PCR using universal bacterial primers 27F and

1492R (Yu et al. 2013) followed by Sanger sequencing as Aeromonas spp. The proliferative

cottony material on the external lesions of four fish was identified as fungal hyphae (1/15 fish)

and oomycetes in the genera Saprolegia (2/15 fish) and Aphanomyces (1/15 fish).

Fourteen of 15 moribund or deceased warmwater exposed fish tested positive by qPCR

with mean viral loads ranging from 100.7

-106.4

viral copies 4μl-1

of extracted internal tissue

homogenate DNA (Table 4-1). Virus isolation was positive for 12/15 fish with viral titers

ranging from 101.8

-106 TCID50 ml

-1 of internal tissue homogenates (Table 4-1). One additional

fish was positive after the blind passage.

The 8 exposed coldwater fish and 4 control fish at each temperature sampled at d 28 PE

appeared healthy with no gross or wet mount abnormalities and no bacterial growth observed on

cultures. Additionally, internal tissue homogenates were all negative for ranavirus by TaqMan

qPCR and virus isolation. In contrast, 5/8 exposed warmwater fish sampled at d 28 exhibited torn

fins and 4/8 appeared abnormally underweight. Internal necropsy findings included a few fish

with small liver (3/15 fish) or spleen (1/15). Two of eight fish were ranavirus-positive via qPCR

with 101.3

-101.4

viral copies 4μl-1

while all eight fish were negative by virus isolation.

Study 2: Effect of Water Temperature on Disease Progression

Mean weight of the 120 fish used in Study 2 was 35.6 g (range 18-57 g) and 22.7 cm

(range 18.4-26.8 cm) fork length. Infection dose for each temperature was 113 ml in 40 l water

containing 40 exposed fish, for a final dose of 4.0 x 104 TCID50 ml

-1 water. No natural mortality

occurred throughout the study. Of the 48 virus exposed fish, 54% (13/24) of the warmwater

exposed fish displayed clinical signs consistent with ranavirus infection compared to none of

111

coldwater exposed fish (Table 4-2). Internal and external tissue viral loads were significantly

higher in the warmwater fish on d 3, 5, and 7 PE and additionally on day 14 PE for internal

tissues (Table 4-3, Figure 4-4). Days 1 and 28 yielded low viral loads (<102 copies 4μl

-1 tissue

homogenate DNA) and titers (<101.5

TCID50 ml-1

) at both temperatures. Prevalence of infection

(percentage of fish displaying a viral load or titer) was higher in the warmwater versus coldwater

fish (Figures 4-5 and 4-6).

Histological examination of slides from the 11 examined fish (6 warmwater and 5

coldwater exposed animals) revealed several findings consistent with ranavirus infection,

including necrosis characterized by multifocal to widespread regions containing pyknotic nuclei,

cellular debris, and/or loss of cellular architecture and definition (Figures 4-7 through 4-10).

Necrosis was recognized in the liver, spleen, pericardial lymphomyeloid tissue, gill, and

posterior kidney. Also present was subjectively decreased hematopoietic cell populations in the

spleen and pericardial lymphomyeloid tissue, as well as hemorrhage and/or disruption of the

epithelial architecture of the gill, skin, and fin. Cytoplasmic basophilic to amphophilic inclusions

consistent with those of ranavirus were not observed. Histopathologic findings were more

prevalent and severe in ranavirus-exposed fish from the warmwater treatment compared to the

coldwater fish, in which microscopic lesions were absent or rare.

ISH using RNAscope® technology revealed ranaviral nucleic acid within several tissues

in the warmwater exposed fish, including skin, fin, gill, barbel, olfactory epithelium, heart,

meningeal and pericardial lymphomyeloid tissue, liver, spleen, stomach, intestine, and posterior

kidney (Table 4-4; Figures 4-7 through 4-10). Tissue sections from the coldwater exposed fish

were negative by ISH on all sampling dates.

112

Discussion

The studies described herein suggest temperature has a clear effect on Frog virus 3 (FV3)

disease in young-of-year pallid sturgeon. Study 1 demonstrated that the severity of disease and

cumulative mortality were significantly greater at the elevated water temperature (23°C). The

reported cumulative mortality in study 1 of 42.5% was lower than the previously reported 90%

by Waltzek and colleagues (2014). This difference may be explained by: 1) viral factors such as

the isolate (PSRV09 vs. PSRV15, Chapter 2) and viral titer used in the bath challenges, host

factors such as size and genetic background of the young-of-year pallid sturgeon, and

environmental factors (e.g., tank size, stocking density, and water flow rate).

A slightly higher viral titer was used in study 1 as compared to the Waltzek et al. (2014)

challenge study suggesting viral titer unlikely explains the difference in cumulative mortality

between the studies. Unique genetic differences, often involving genes encoding proteins of

unknown functions, were observed in all of the closely related PSRV isolates (Chapter 2) making

difficult any interpretation of why the PSRV15 isolate used in this study might be less

pathogenic than the PSRV09 isolate used previously. The mean weight of the fish used in study

1 was smaller (28.7 g) compared to the previous study (39.8 g) which unlikely explains the

difference in cumulative mortality. Although the young-of-year pallid sturgeon in both studies

were supplied by the Gavins Point Fish Hatchery in South Dakota, the genetic background of the

parents and resulting offspring may have varied between the studies. Finally, environmental

factors including tank dimension and material, stocking density, and water flow rate varied

between the studies and may have contributed to the observed difference in cumulative mortality.

The clinical signs, viral load, and viral titer data presented in study 2 demonstrated FV3

disease progressed more rapidly and became more pronounced at the elevated water temperature.

Previously, higher FV3 viral titer has been shown to more rapidly induce disease in young-of-

113

year pallid sturgeon; however, by the end of the study cumulative mortality was identical

between the lower and higher titer treatments (Waltzek et al. 2014).

The histological methods used in study 2, including semi-validated ISH method using

RNAscope®, provide novel insight into FV3 tissue tropism in juvenile pallid sturgeon.

Histological examination revealed a greater distribution and severity of microscopic lesions in

fish held at the elevated water temperature. Gross and microscopic lesions of certain tissues were

similar to those previously reported in experimental FV3 sturgeon infections (Waltzek et al.

2014, Miller et al. 2015); however, the more thorough tissue sampling in this study expanded the

known tissue distribution of the virus. Although some tissue distributions were already

documented for ranaviruses in fish (e.g., hematopoietic tissues), other locations (e.g., gill, skin,

sensory tissues) are less well known and may aid in refining future sampling protocols. For

example, gill was RNAscope®-positive in 60% (3/5) of the tested 23°C fish during the first 2 wk

following infection, suggesting this tissue may be an acceptable tissue for non-lethal ranavirus

screening. The presence of infection in several external tissues (e.g., skin, fin, gill, sensory

epithelium) also suggests one or more of these tissues serve as the route of infection and/or viral

shedding.

Study 2 data demonstrated fish held at the lower water temperature (17°C) did become

infected with FV3 despite not showing clinical signs of disease. Viral DNA was detected within

tissues by qPCR (viral load) and more importantly the recovery of virus in cell culture (viral

titer). This finding is important because it suggests fish exposed at lower temperatures harbor the

virus and might serve as viral reservoirs until conditions become favorable for transmission (e.g.,

thermal stress). In fact, some exposed fish at both temperatures remained infected after 28 days

via qPCR and virus isolation as previously reported (Waltzek et al. 2014). Thus, future challenge

114

studies are needed to determine whether survivors harbor the virus over longer periods of time

and whether they might pose a risk if resocked into wild populations. In balance, future studies

are also needed to determine whether survivors mount a cellular and humoral immune response

following exposure. Demonstrating that fish exposed at lower temperatures, at which mortality is

minimal, eventually clear the virus and/or become protected would be an important step toward

developing an effective mitigation strategy. Ultimately, temperature manipulation may be one of

several possible management tools to help minimize ranaviral outbreaks in hatcheries rearing

endangered sturgeon, whether used alone or in combination with other strategies (e.g., reduced

stocking densities, disinfection of incoming water, vaccination).

115

Table 4-1. Study 1 results summary.

Study result 23°C 17°C

Cumulative mortality 43% (17/40 fish) 0% (0/40 fish)

Clinical signs Mild to severe None

qPCR range (viral copies 4μl-1

) 100.7

-106.4

NA

Viral titer range (TCID50 ml-1

) 101.8

-106.0

NA

Survivors qPCR(+) at d 28 25% (2/8 fish) 0% (0/8 fish)

116

Table 4-2. Study 2 results summary.

Study result 23°C 17°C

Clinical sign prevalence 54% (13/24 fish) 0% (0/24 fish)

100% infection incidence (qPCR) Days 3-28 Day 7

Animals VI (+) (%) 72% (13/18) 17% (3/18)

Viral titer range (TCID50 ml-1

) 100.7

-104.2

100.7

-101.5

117

Table 4-3. Log mean (± SE) qPCR viral copy number for external and internal tissue

homogenates in the warmwater and coldwater treatments over the 28 d study (n=4

fish per treatment per day). Significant differences (P < 0.05) between the two

temperatures are denoted by different letters (x for external tissue and

y for internal

tissue).

Day post-

exposure

External tissue pool Internal tissue pool

17°C 23°C 17°C 23°C

1 0.33 (0.23) 0.42 (0.25) 0 (0) 0 (0)

3x,y

0 (0)

3.87 (0.51)

0 (0) 4.27 (0.57)

5 x,y

1.09 (0.73) 3.76 (0.62) 0.80 (0.80)

4.07 (0.25)

7 x,y

2.00 (0.26) 4.93 (0.55) 2.11 (0.81)

5.49 (0.54)

14y 0.86 (0.86) 3.02 (0.93) 1.55 (0.16)

2.59 (0.37)

28 0.63 (0.63) 0.18 (0.18) 0.25 (0.25) 1.03 (0.37)

118

Table 4-4. RNAscope® ISH results from cold- and warmwater fish corresponding with the highest average TaqMan qPCR viral load

per sampling date. NE = not examined due to all animals being ranavirus(-) via qPCR.

17°C 23°C

Day Positive tissues

External qPCR

copy #

Internal qPCR copy

# Positive tissues

External

qPCR copy #

Internal qPCR

copy #

1 None 10 0 None 9 1

3 NE 0 0 Skin, fin, gill, olfactory

epithelium, heart,

pericardial lymphoid tissue,

liver, spleen, intestine,

posterior kidney

37,500 351,769

5 None 1,214 1,592 Skin, spleen, intestine 329,115 35,726

7 None 202 5,084 Skin, fin, gill, barbel,

olfactory epithelium, heart,

pericerebral and pericardial

lymphoid tissue, liver,

spleen, stomach, intestine,

posterior kidney

1,178,776 2,308,890

14 None 2,807 106 Gill 623,465 4,536

28 None 320 0 Olfactory epithelium 0 134

119

Figure 4-1. Experimental tank design for studies 1 and 2. Sample sizes (n) are indicated. Tx =

treatment. Rep = replicate.

120

Figure 4-2. Study 1 survival curve. Data indicate survival (%) values for young-of-year pallid

sturgeon exposed to FV3 via bath infection at 17°C and 23°C water temperatures.

0

20

40

60

80

100

0 7 14 21 28

Surv

ival

(%

)

Day post-infection

23 degrees

17 degrees

121

Figure 4-3. Representative photos of gross pathology associated with FV3 disease at 23°C in

studies 1 and 2. A) Severe erythema and hemorrhage of the skin and fins primarily

displayed during early infection. B) Splenomegaly and hemorrhage of the coelomic

organs associated with late stage infection. Photos courtesy of author.

A

B

1 cm

1 cm

122

Figure 4-4. Comparison of log mean (±SE) qPCR copy number for external (Ext.) and internal

(Int.) tissue homogenates in the warmwater and coldwater treatments over the 28 d

study.

0

1

2

3

4

5

6

7

1 3 5 7 14 28

Log V

iral

Copie

s 4μ

l-1

Day post-infection

17°C Ext.

17°C Int.

23°C Ext.

23°C Int.

123

A.

B.

Figure 4-5. Prevalence of FV3 infection in study 2 following bath exposure at 17°C. A)

TaqMan qPCR results. B) Virus isolation (VI) results.

0%

20%

40%

60%

80%

100%

1 3 5 7 14 28

Pre

val

ence

of

ranav

irus

infe

ctio

n

(qP

CR

)

Day postexposure

17°C External

17°C Internal

0%

20%

40%

60%

80%

100%

1 3 5 7 14 28

Pre

val

ence

of

ranav

irus

infe

ctio

n

(VI)

Day postexposure

17°C External

17°C Internal

124

A.

B.

Figure 4-6. Prevalence of FV3 infection in study 2 following bath exposure at 23°C. A) TaqMan

qPCR results. B) Virus isolation (VI) results.

0%

20%

40%

60%

80%

100%

1 3 5 7 14 28

Pre

val

ence

of

ranav

irus

infe

ctio

n

(qP

CR

)

Day postexposure

23°C External

23°C Internal

0%

20%

40%

60%

80%

100%

1 3 5 7 14 28

Pre

val

ence

of

ranav

irus

infe

ctio

n

(VI)

Day postexposure

23°C External

23°C Internal

125

A.

B.

Figure 4-7. Hematoxylin and eosin (H&E) and RNAscope® in situ hybridization (ISH) results

from the spleen of a warmwater exposed sturgeon sampled on day 7 of study 2. A)

Spleen. A focus of necrosis is located adjacent to a perivascular lymphoid aggregate.

H&E. B) Spleen. ISH reveals multiple areas where ranavirus-positive cells are

loosely or densely aggregated. Photos courtesy of Drs. Lisa Farina and Salvatore

Frasca Jr.

126

A.

B.

C.


from the gill of a warmwater exposed sturgeon sampled on day 7 of study 2. A) Gill.

There is multifocal lamellar injury with hemorrhage. H&E. B) Gill. Higher

magnification of a lamella reveals disruption of the vascular core with hemorrhage

and necrosis. H&E. C) Gill. ISH shows ranavirus-positive cells within the lamellar

interstitium and epithelium. Photos courtesy of Drs. Lisa Farina and Salvatore Frasca

Jr.

127

A.

B.


from the posterior kidney of a warmwater exposed sturgeon sampled on day 7 of

study 2. A) Kidney. Mild multifocal vacuolation of tubular epithelial cells is present.

B) Kidney. ISH reveals ranavirus-positive cells scattered within the interstitium.

Photos courtesy of Drs. Lisa Farina and Salvatore Frasca Jr.

128

A.

B.


from the heart, including pericardial lymphomyeloid tissue, from a warmwater

exposed sturgeon sampled on day 7 of study 2. A) Heart. There is depletion of

hematopoietic cells from the pericardial lymphomyeloid tissue. H&E. B) Heart. ISH

reveals ranavirus-positive cells scattered within the pericardial lymphomyeloid tissue

and ventricular endothelium. Photos courtesy of Drs. Lisa Farina and Salvatore

Frasca Jr.

129

CHAPTER 5

CONCLUDING STATEMENTS

Since the first discovery of Frog virus 3 in the Northern leopard frog (Rana pipiens) in

1966, ranaviruses have been detected with increasing frequency in several freshwater and marine

fish species in both aquaculture and wild stocks. Notable hatchery-related epizootics in the US

have included critically endangered sturgeon species (e.g., pallid Scaphirhynchus albus, lake

Acipenser fulvescens, and Russian A. gueldenstaedtii).

To examine the impact and spread of fish ranaviruses, viral characterization and

phylogenomic analyses were used to elucidate significant biologic and epidemiologic trends in

aquaculture. Novel methods based on alignments of full ranaviral genomes produced highly

resolved and supported trees providing insight into the genomic arrangements of fish

ranaviruses. Results suggest that ranaviruses originated in fishes before jumping host classes into

amphibians and reptiles. Furthermore, the fish ranviruses sequenced in this study grouped in

theFrog virus 3 (FV3) or common midwife toad virus (CMTV) clades. Thus, strains of both FV3

and CMTV exhibit very low host specificity infecting fish, amphibians, and reptiles on separate

continents. These epidemiological patterns suggest that international trade of ornamental and

food animals likey explains the global emergence of closely related ranaviruses.

Comparative genomic analyses facilitated the design of improved molecular assays for

the diagnosis and characterization of ranaviruses, including a pan-ranavirus quantitative real-

time TaqMan PCR and a pan-ranavirus in situ hybridization (ISH) assay using RNAscope®

technology. After being tested by the OIE reference laboratory for ranaviruses and validated

against an extensive pool of 36 ranavirus isolates, the qPCR assay was found to detect the

majority of isolates and will hopefully serve as a useful, expedient diagnostic tool in future

ranavirus surveillance and research efforts.

130

Finally, bath FV3 challenges were conducted at 17˚C and 23˚C to examine the effect of

water temperature on ranaviral disease in one of the most vulnerable aquacultured host species

(hatchery-reared young-of-year pallid sturgeon). Elevated water temperature resulted in

significant disease as compared to the lower water temperatue treatment in which juvenile pallid

sturgeon became infected but disease was not observed. To date, limited treatment options exist

for ranaviral infection in affected animals particularly food fish species. However, it appears

temperature manipulation may serve as an effective management tool for sturgeon hatcheries

afflicted with ranavirus epizootics. Ultimately, I hope the body of research provided herein has

advanced our understanding of the impact of fish ranaviruses in aquaculture and might ultimately

lead to management strategies.

131

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BIOGRAPHICAL SKETCH

Natalie Stilwell (formerly Steckler) is a veterinarian from Lexington, KY. From an early

age, Natalie knew she wanted to focus her career on animal care and health. After obtaining her

Bachelor of Science in biology from Bellarmine University (Louisville, KY) in 2005, Natalie

completed her Doctor of Veterinary Medicine degree at Auburn University in Auburn, AL

(2006-2010). She then chose to pursue a veterinary specialty career in aquatic animal health. In

2012, Natalie completed a Master of Science degree as part of the Fisheries and Aquatic

Sciences Program at the University of Florida under the mentorship of Dr. Roy Yanong,

followed by a postdoctoral program with Dr. Thomas Waltzek at the University of Florida

Wildlife and Aquatic Animal Veterinary Disease (February to August 2013). She completed her

Doctor of Philosophy degree in Veterinary Medical Sciences at the UF College of Veterinary

Medicine under the advisorship of Dr. Waltzek in summer 2017.

© 2017 natalie katherine stilwell

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