© 2017 natalie katherine stilwell
TRANSCRIPT
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
© 2017 Natalie Katherine Stilwell
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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
2-9 Predicted open reading frames for the 2009 pallid sturgeon ranavirus isolate
(PSRV09)…. ......................................................................................................................58
2-10 Predicted open reading frames for the 2013 pallid sturgeon ranavirus isolate
(PSRV13)…. ......................................................................................................................61
2-11 Predicted open reading frames for the 2015 pallid sturgeon ranavirus isolate
(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
4-8 Hematoxylin and eosin (H&E) and RNAscope® in situ hybridization (ISH) results from
the gill of a warmwater exposed sturgeon sampled on day 7 of study 2 .........................126
4-9 Hematoxylin and eosin (H&E) and RNAscope® in situ hybridization (ISH) results from
the posterior kidney of a warmwater exposed sturgeon sampled on day 7 of study 2. ...127
9
4-10 Hematoxylin and eosin (H&E) and RNAscope® in situ hybridization (ISH) results from
the heart, including pericardial lymphomyeloid tissue, from a warmwater exposed
sturgeon sampled on day 7 of study 2 ..............................................................................128
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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
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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
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
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
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
25La Unknown function
26Ra,c
Unknown function
28La Unknown function
29Ra Unknown function
30Ra Unknown function
32Ra 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
45Ra Unknown function
48L Phosphotransferase
53L Myristylated membrane protein
55Ra Unknown function
56Ra Unknown function
57Ra Unknown function
58La Unknown function
59La Unknown function
62R Tyrosine kinase
67Ra Unknown function
68Ra Neurofilament triplet H1-like protein
69Ra Unknown function
44
Table 2-1. Continued
Gene Predicted function
70Ra Unknown function
77R Unknown function
78La Unknown function
81La Unknown function
85L D5 family NTPase/ATPase
86R Unknown function
89L Serine/threonine protein kinase
90Ra Unknown function
91Ra Unknown function
92L ATPase
95R Unknown function
96La Unknown function
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
Pallid sturgeon ranavirus 2009 PSRV09 Unpublished WAVDLa
Pallid sturgeon ranavirus 2013 PSRV13 Unpublished WAVDLa
Pallid sturgeon ranavirus 2015 PSRV15 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)
PSRV09 Pallid sturgeon (Scaphirhynchus albus) EPC 48 143.3 (6.5) 189.9 (18.9)
PSRV13 Pallid sturgeon (Scaphirhynchus albus) EPC 24 125.9 (5.2) 162.0 (8.9)
PSRV15 Pallid sturgeon (Scaphirhynchus albus) EPC 24 148.6 (4.3) 216.7 (13.1)
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
Table 2-6. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
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
Table 2-6. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
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
73 80333-81307 325 NTPase/helicase FV3 73L 100 YP_031652
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
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 100 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 100 YP_031582
6 5118-5732 205 US22 family protein FV3 5R 97 YP_031583
7 5735-5962 76 STIV 8R 100 ACF42227
8 6723-7151 143 STIV 9L 96 ACF42228
9 7231-11112 1294 DNA-dependent RNA polymerase II largest subunit RGV 9R 100 AFG73051
10 11481-14327 949 NTPase/helicase FV3 9L 100 YP_031587
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
23 25731-28652 974 D5 family NTPase/ATPase SSME 22R 100 AHM26101
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
28 33471-36383 971 Tyrosine kinase FV3 27R 99 YP_031605
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
Table 2-7. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
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
38 42358-42993 212 NIF/NLI interacting factor FV3 37R 100 YP_031615
39 43134-44831 566 Ribonucleoside reductase alpha subunit FV3 38R 100 YP_031616
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
51 56964-58031 356 3-beta-hydroxysteroid dehydrogenase RGV 52L 100 ABI36881
52 58369-59937 523 Myristylated membrane protein FV3 53R 100 YP_031631
53 60184-60414 77 Nuclear calmodulin-binding protein FV3 54L 100 YP_031632
54 60452-61747 432 Helicase-like protein STIV 57L 99 ACF42275
55 60596-61735 380 40 kDa protein FV3 55R 98 YP_031634
56 61835-62272 146 FV3 56R 99 YP_031635
57 62386-63882 499 Phosphotransferase FV3 57R 100 YP_031636
58 64197-64910 238 FV3 58R 100 YP_031637
59 65461-66519 353 FV3 59L 100 YP_031638
60 66681-69722 1014 DNA polymerase SSME 60R 100 AHM26138
61 69731-69913 61 FV3 61L 100 YP_031640
62 70364-74029 1222 DNA-dependent RNA polymerase II second largest subunit FV3 62L 100 YP_031641
63 74408-74902 165 dUTPase FV3 63R 100 YP_031642
64 75042-75329 96 Caspase recruitment domain protein FV3 64R 100 YP_031643
65 75722-75886 55 FV3 65L 100 YP_031644
54
Table 2-7. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
66 75883-76434 184 FV3 66L 100 YP_031645
67 76489-77652 388 Ribonucleoside reductase beta subunit FV3 67L 100 YP_031646
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
73 80466-81440 325 NTPase/helicase FV3 73L 100 YP_031652
74 81615-82727 371 FV3 74L 100 YP_031653
75 82759-83013 85 LITAF/PIG7 possible membrane associated motif in LPS-induced
tumor necrosis factor alpha factor
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
79 85015-86733 573 ATPase-dependent protease RUK13 79R 100 AIX94632
80 87356-88471 372 Ribonuclease III FV3 80L 100 YP_031659
81 88527-88805 93 Transcription elongation factor S-II FV3 81R 100 YP_031660
82 88934-89407 158 Immediate early protein ICP-18 FV3 82R 100 YP_031661
83 89857-90501 215 Cytosine DNA methyltransferase FV3 83R 100 YP_031662
84 90873-91610 246 Proliferating cell nuclear antigen (PCNA) PPIV 84R 100 ANZ56939
85 91685-92272 196 Thymidine kinase FV3 85R 100 YP_031664
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
90 96760-98151 464 Major capsid protein SSME 90R 100 AHM26166
91 98275-99462 396 Immediate early protein ICP-46 SSME 91R 100 AHM26167
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
95 101029-102120 364 DNA repair enzyme RAD2 STIV 100R 100 ACF42318
96 102921-103592 224 FV3 96R 100 YP_031675
97 103675-104088 138 Myeloid cell leukemia protein STIV 103R 100 ACF42321
98 104652-105317 222 US22 family protein STIV 104R 100 ACF42322
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
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 100 YP_031579
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
6 5128-5742 205 US22 family protein FV3 5R 96 YP_031583
7 5745-5942 66 STIV 8R 100 ACF42227
8 6733-7161 143 STIV 9L 96 ACF42228
9 7241-11122 1294 DNA-dependent RNA polymerase II largest subunit RGV 9R 100 AFG73051
10 11491-14337 949 NTPase/helicase FV3 9L 100 YP_031587
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
20 21654-24272 873 Serine/threonine protein kinase SSME 19R 99 AHM26098
21 24320-24766 149 PPIV 92 99 ANZ57004
22 24989-25648 220 SSME 56L 100 AHM26100
23 25778-28699 974 D5 family NTPase/ATPase SSME 22R 100 AHM26101
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
28 33516-36428 971 Tyrosine kinase FV3 27R 100 YP_031605
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
Table 2-8. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
36 41266-41727 154 SSME 35L 100 AHM26114
37 41925-42347 141 FV3 36L 100 YP_031614
38 42744-43379 212 NIF/NLI interacting factor FV3 37R 100 YP_031615
39 43520-45217 566 Ribonucleoside reductase alpha subunit FV3 38R 100 YP_031616
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
46 52359-52778 140 Neurofilament triplet H1-like protein BIV 46L 98 ANK57973
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
51 57284-58351 356 3-beta-hydroxysteroid dehydrogenase RGV 52L 100 ABI36881
52 58689-60257 523 Myristylated membrane protein FV3 53R 100 YP_031631
53 60456-60686 77 Nuclear calmodulin-binding protein FV3 54L 100 YP_031632
54 60724-62019 432 Helicase-like protein FV3 55L 100 YP_031633
55 60868-61557 230 40 kDa protein RGV 57R 100 AFG73099
56 62107-62544 146 FV3 56R 100 YP_031635
57 62658-64154 499 Phosphotransferase FV3 57R 100 YP_031636
58 64479-65192 238 FV3 58R 100 YP_031637
59 65743-66801 353 FV3 59L 100 YP_031638
60 66963-70004 1014 DNA polymerase SSME 60R 100 AHM26138
61 70013-70195 61 FV3 61L 100 YP_031640
62 70634-74299 1222 DNA-dependent RNA polymerase II second largest subunit FV3 62L 100 YP_031641
63 74678-75172 165 dUTPase FV3 63R 100 YP_031642
64 75282-75569 96 Caspase recruitment domain protein FV3 64R 100 YP_031643
65 75962-76126 55 FV3 65L 94 YP_031644
66 76123-76674 184 FV3 66L 99 YP_031645
67 76729-77892 388 Ribonucleoside reductase beta subunit FV3 67L 100 YP_031646
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
Table 2-8. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
72 79580-80296 239 FV3 72L 100 YP_031651
73 80702-81676 325 NTPase/helicase FV3 73L 100 YP_031652
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
79 85236-86954 573 ATPase-dependent protease RUK13 79R 100 AIX94632
80 87577-88692 372 Ribonuclease III FV3 80L 100 YP_031659
81 88748-89026 93 Transcription elongation factor S-II FV3 81R 100 YP_031660
82 89155-89628 158 Immediate early protein ICP-18 FV3 82R 100 YP_031661
83 90078-90722 215 Cytosine DNA methyltransferase SSME 83R 100 AHM26159
84 91107-91844 246 Proliferating cell nuclear antigen (PCNA) PPIV 84R 100 ANZ56939
85 91919-92506 196 Thymidine kinase FV3 85R 100 YP_031664
86 92896-93081 62 SSME 86L 100 AHM26162
87 93434-95269 612 FV3 87L 100 YP_031666
88 95302-95754 151 Thiol oxidoreductase GGRV 65R 99 AJR29223
89 95822-96904 361 SSME 89R 100 AHM26165
90 96997-98388 464 Major capsid protein SSME 90R 100 AHM26116
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
95 101320-102411 364 DNA repair enzyme RAD2 STIV 100R 100 ACF42318
96 103214-103885 224 FV3 96R 100 YP_031675
97 103968-104381 138 Myeloid cell leukemia protein STIV 103R 100 ACF42321
98 104877-105542 222 US22 family protein STIV 104R 100 ACF42322
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
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 100 YP_031579
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
6 5127-5741 205 US22 family protein FV3 5R 96 YP_031583
7 5744-5971 76 STIV 8R 100 ACF42227
8 6732-7160 143 STIV 9L 96 ACF42228
9 7240-11121 1294 DNA-dependent RNA polymerase II largest subunit RGV 9R 100 AFG73051
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
20 21653-24319 889 Serine/threonine protein kinase SSME 19R 99 AHM26098
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
Table 2-9. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
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
38 43000-43635 212 NIF/NLI interacting factor FV3 37R 100 YP_031615
39 43776-45473 566 Ribonucleoside reductase alpha subunit FV3 38R 100 YP_031616
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
51 57593-58660 356 3-beta-hydroxysteroid dehydrogenase RGV 52L 100 ABI36881
52 58998-60566 523 Myristylated membrane protein FV3 53R 100 YP_031631
53 60765-60995 77 Nuclear calmodulin-binding protein FV3 54L 100 YP_031632
54 61033-62328 432 Helicase-like protein FV3 55L 100 YP_031633
55 61177-62316 380 40 kDa protein FV3 55R 99 YP_031634
56 62416-62853 146 FV3 56R 100 YP_031635
57 62967-64463 499 Phosphotransferase FV3 57R 100 YP_031636
58 64788-65501 238 FV3 58R 100 YP_031637
59 66052-67110 353 FV3 59L 100 YP_031638
60 67272-70313 1014 DNA polymerase SSME 60R 100 AHM26138
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
64 75612-75899 96 Caspase recruitment domain protein FV3 64R 100 YP_031643
65 76158-76247 30 PPIV 42 88 ANZ56958
66 76302-77465 388 Ribonucleoside reductase beta subunit FV3 67L 100 YP_031646
67 77748-78035 96 FV3 68R 100 YP_031647
68 78171-78437 89 FV3 69R 100 YP_031648
60
Table 2-9. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
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
74 82702-82956 85 LITAF/PIG7 possible membrane associated motif in LPS-induced
tumor necrosis factor alpha factor
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
78 84934-86652 573 ATPase-dependent protease RUK13 79R 100 AIX94632
79 87275-88390 372 Ribonuclease III FV3 80L 100 YP_031659
80 88446-88724 93 Transcription elongation factor S-II FV3 81R 100 YP_031660
81 88853-89326 158 Immediate early protein ICP-18 FV3 82R 100 YP_031661
82 89776-90420 215 Cytosine DNA methyltransferase SSME 83R 100 AHM26159
83 90805-91542 246 Proliferating cell nuclear antigen (PCNA) PPIV 84R 100 ANZ56939
84 91617-92204 196 Thymidine kinase FV3 85R 100 YP_031664
85 92594-92779 62 SSME 86L 100 AHM26162
86 93132-95030 633 FV3 87L 100 YP_031666
87 95063-95515 151 Thiol oxidoreductase GGRV 65R 99 AJR29223
88 95583-96644 354 SSME 89R 100 AHM26165
89 96737-98128 464 Major capsid protein SSME 90R 100 AHM26166
90 98252-99439 396 Immediate early protein ICP-46 RUK13 91R 100 AIX94643
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
94 101038-102129 364 DNA repair enzyme RAD2 STIV 100R 100 ACF42318
95 102932-103603 224 FV3 96R 100 YP_031675
96 103686-104099 138 Myeloid cell leukemia protein STIV 103R 100 ACF42321
97 104595-105260 222 US22 family protein STIV 104R 100 ACF42322
61
Table 2-10. Predicted open reading frames for the 2013 pallid sturgeon ranavirus isolate (PSRV13). 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 100 YP_031579
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
6 5130-5744 205 US22 family protein FV3 5R 96 YP_031583
7 5747-5974 76 STIV 8R 100 ACF42227
8 6735-7163 143 STIV 9L 96 ACF42228
9 7243-11124 1294 DNA-dependent RNA polymerase II largest subunit RGV 9R 100 AFG73051
10 11493-14339 949 NTPase/helicase FV3 9L 100 YP_031587
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
20 21656-24268 871 Serine/threonine protein kinase SSME 19R 99 AHM26098
21 24316-24762 149 PPIV 22 99 ANZ57004
22 24985-25644 220 SSME 56L 100 AHM26100
23 25774-28695 974 D5 family NTPase/ATPase SSME 22R 100 AHM26101
24 29073-30221 383 SSME 23R 100 AHM26102
25 30631-31701 357 SSME 24R 100 AHM26103
26 31895-32680 262 ToRV 63L 100 AJR29294
27 33004-33705 234 eIF-2alpha-like protein ADRV 84L 100 AGV20615
28 34261-37173 971 Tyrosine kinase CGSIV 83L 99 AHA42351
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
Table 2-10. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
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
39 44293-45990 566 Ribonucleoside reductase alpha subunit FV3 38R 100 YP_031616
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
46 53092-53493 134 Neurofilament triplet H1-like protein BIV 46L 98 ANK57973
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
51 57951-59018 356 3-beta-hydroxysteroid dehydrogenase RGV 52L 100 ABI36881
52 59356-60924 523 Myristylated membrane protein FV3 53R 100 YP_031631
53 61123-61353 77 Nuclear calmodulin-binding protein FV3 54L 100 YP_031632
54 61391-62686 432 Helicase-like protein FV3 55L 100 YP_031633
55 61535-62674 380 40 kDa protein FV3 55R 100 YP_031634
56 62774-63211 146 FV3 56R 100 YP_031635
57 63325-64821 499 Phosphotransferase FV3 57R 100 YP_031636
58 65146-65859 238 FV3 58R 100 YP_031637
59 66410-67468 353 FV3 59L 100 YP_031638
60 67630-70671 1014 DNA polymerase SSME 60R 100 AHM26138
61 70680-70862 61 FV3 61L 100 YP_031640
62 71299-74964 1222 DNA-dependent RNA polymerase II second largest subunit FV3 62L 100 YP_031641
63 75343-75837 165 dUTPase FV3 63R 100 YP_031642
64 75977-76264 96 Caspase recruitment domain protein FV3 64R 100 YP_031643
65 76657-76821 55 FV3 65L 96 YP_031644
66 76818-77369 184 FV3 66L 99 YP_031645
63
Table 2-10. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
67 77424-78587 388 Ribonucleoside reductase beta subunit FV3 67L 100 YP_031646
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
73 81448-82422 325 NTPase/helicase SSME 73L 100 AHM26149
74 82597-83709 371 FV3 74L 100 YP_031653
75 83741-83995 85 LITAF/PIG7 possible membrane associated motif in LPS-induced
tumor necrosis factor alpha factor
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
79 85973-87691 573 ATPase-dependent protease RUK13 79R 100 AIX94632
80 88314-89429 372 Ribonuclease III FV3 80L 100 YP_031659
81 89485-89763 93 Transcription elongation factor S-II FV3 81R 100 YP_031660
82 89892-90365 158 Immediate early protein ICP-18 FV3 82R 100 YP_031661
83 90815-91459 215 Cytosine DNA methyltransferase SSME 83R 100 AHM26159
84 91844-92581 246 Proliferating cell nuclear antigen (PCNA) PPIV 84R 100 ANZ56939
85 92656-93243 196 Thymidine kinase FV3 85R 100 YP_031664
86 93633-93818 62 SSME 86L 100 AHM26162
87 94171-96042 624 FV3 87L 100 YP_031666
88 96075-96527 151 Thiol oxidoreductase GGRV 65R 99 AJR29223
89 96595-97677 361 SSME 89R 100 AHM26165
90 97770-99161 464 Major capsid protein SSME 90R 100 AHM26166
91 99285-100472 396 Immediate early protein ICP-46 RUK13 91R 100 AIX94643
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
95 102055-103146 364 DNA repair enzyme RAD2 STIV 100R 100 ACF42318
96 103950-104621 224 FV3 96R 100 YP_031675
97 104704-105117 138 Myeloid cell leukemia protein STIV 103R 100 ACF42321
98 105613-106278 222 US22 family protein STIV 104R 100 ACF42322
64
Table 2-11. Predicted open reading frames for the 2015 pallid sturgeon ranavirus isolate (PSRV15). 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 100 YP_031579
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
6 5130-5744 205 US22 family protein FV3 5R 96 YP_031583
7 5747-5974 76 STIV 8R 100 ACF42227
8 6735-7163 143 STIV 9L 96 ACF42228
9 7243-11124 1294 DNA-dependent RNA polymerase II largest subunit RGV 9R 100 AFG73051
10 11493-14339 949 NTPase/helicase FV3 9L 100 YP_031587
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
23 29083-30231 383 D5 family NTPase/ATPase SSME 23R 100 AHM26102
24 30641-31711 357 SSME 24R 100 AHM26103
25 31905-32690 262 ToRV 63L 100 AJR29294
26 33014-33715 234 eIF-2alpha-like protein ADRV 84L 100 AGV20615
27 34271-37183 971 Tyrosine kinase CGSIV 83L 99 AHA42351
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
Table 2-11. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
36 42311-42742 144 RUK13 36L 100 AIX94599
37 43137-43772 212 NIF/NLI interacting factor SSME 37R 100 AHM26116
38 43913-45610 566 Ribonucleoside reductase alpha subunit FV3 38R 100 YP_031616
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
45 52697-53098 134 Neurofilament triplet H1-like protein BIV 46L 98 ANK57973
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
50 57595-58662 356 3-beta-hydroxysteroid dehydrogenase RGV 52L 100 ABI36881
51 59000-60568 523 Myristylated membrane protein FV3 53R 100 YP_031631
52 60767-60997 77 Nuclear calmodulin-binding protein FV3 54L 100 YP_031632
53 61035-62330 432 Helicase-like protein FV3 55L 100 YP_031633
54 61179-62318 380 40 kDa protein FV3 55R 100 YP_031634
55 62418-62855 146 FV3 56R 100 YP_031635
56 62969-64465 499 Phosphotransferase FV3 57R 100 YP_031636
57 64790-65503 238 FV3 58R 100 YP_031637
58 66054-67112 353 FV3 59L 100 YP_031638
59 67274-70315 1014 DNA polymerase SSME 60R 100 AHM26138
60 70324-70506 61 FV3 61L 100 YP_031640
61 70947-74612 1222 DNA-dependent RNA polymerase II second largest subunit FV3 62L 100 YP_031641
62 74991-75485 165 dUTPase FV3 63R 100 YP_031642
63 75625-75912 96 Caspase recruitment domain protein FV3 64R 100 YP_031643
64 76305-76469 55 FV3 65L 96 YP_031644
65 76466-77017 184 FV3 66L 99 YP_031645
66 77072-78235 388 Ribonucleoside reductase beta subunit FV3 67L 100 YP_031646
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
Table 2-11. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
72 81106-82080 325 NTPase/helicase SSME 73L 100 AHM26149
73 82255-83523 423 ATV 29R 96 ALN37132
74 83555-83809 85 LITAF/PIG7 possible membrane associated motif in LPS-
induced tumor necrosis factor alpha factor
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
78 85787-87505 573 ATPase-dependent protease RUK13 79R 100 AIX94632
79 88128-89243 372 Ribonuclease III FV3 80L 100 YP_031659
80 89299-89577 93 Transcription elongation factor S-II FV3 81R 100 YP_031660
81 89706-90179 158 Immediate early protein ICP-18 FV3 82R 100 YP_031661
82 90629-91273 215 Cytosine DNA methyltransferase SSME 83R 100 AHM26159
83 91658-92395 246 Proliferating cell nuclear antigen (PCNA) PPIV 84R 100 ANZ56939
84 92470-93057 196 Thymidine kinase FV3 85R 100 YP_031664
85 93447-93632 62 SSME 86L 100 AHM26162
86 93985-95784 600 SSME 87L 100 AHM26163
87 95817-96269 151 Thiol oxidoreductase GGRV 65R 99 AJR29223
88 96337-97419 361 SSME 89R 100 AHM26165
89 97512-98903 464 Major capsid protein SSME 90R 100 AHM26166
90 99027-100214 396 Immediate early protein ICP-46 RUK13 91R 100 AIX94643
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
94 101797-102888 364 DNA repair enzyme RAD2 STIV 100R 100 ACF42318
95 103692-104363 224 FV3 96R 100 YP_031675
96 104446-104859 138 Myeloid cell leukemia protein STIV 103R 100 ACF42321
97 105355-106020 222 US22 family protein STIV 104R 100 ACF42322
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
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 100 YP_031579
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
7 6033-9914 1294 DNA-dependent RNA polymerase II largest subunit RGV 9R 100 AFG73051
8 10283-13129 949 NTPase/helicase FV3 9L 100 YP_031587
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
21 24592-27513 974 D5 family NTPase/ATPase SSME 22R 100 AHM26101
22 27891-29039 383 STIV 23R 98 ACF42246
23 29421-30518 366 RUK13 24R 99 AIX94589
24 30711-31493 261 ToRV 63L 100 AJR29294
25 31817-32518 234 eIF-2alpha-like protein ADRV 84L 100 AGV20615
26 33091-36003 971 Tyrosine kinase FV3 27R 100 YP_031605
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
Table 2-12. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
36 42292-42927 212 NIF/NLI interacting factor FV3 37R 100 YP_031615
37 43068-44765 566 Ribonucleoside reductase alpha subunit FV3 38R 100 YP_031616
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
44 51988-52389 134 Neurofilament triplet H1-like protein BIV 46L 98 ANK57973
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
49 56847-57914 356 3-beta-hydroxysteroid dehydrogenase RGV 52L 100 ABI36881
50 58252-59820 523 Myristylated membrane protein FV3 53R 100 YP_031631
51 60099-60329 77 Nuclear calmodulin-binding protein FV3 54L 100 YP_031632
52 60367-61662 432 Helicase-like protein FV3 55L 100 YP_031633
53 60511-61650 380 40 kDa protein FV3 55R 100 YP_031634
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
59 67643-70684 1014 DNA polymerase SSME 60R 100 AHM26138
60 70749-70874 42 RGV 64L 100 AFG73106
61 71317-74982 1222 DNA-dependent RNA polymerase II second largest subunit FV3 62L 100 YP_031641
62 75361-75855 165 dUPTase FV3 63R 100 YP_031642
63 75965-76252 96 Caspase recruitment domain protein FV3 64R 100 YP_031643
64 76645-76809 55 FV3 65L 100 YP_031644
65 76806-77357 184 FV3 66L 100 YP_031645
66 77412-78575 388 Ribonucleoside reductase beta subunit FV3 67L 100 YP_031646
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
Table 2-12. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
72 81419-82393 325 NTPase/helicase FV3 73L 100 YP_031652
73 82568-83773 402 ADRV 34R 100 AGV20565
74 83805-84059 85 LITAF/PIG7 possible membrane associated motif in LPS-
induced tumor necrosis factor alpha factor
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
78 86061-87779 573 ATPase-dependent protease RUK13 79R 100 AIX94632
79 88402-89517 372 Ribonuclease III FV3 80L 100 YP_031659
80 89573-89851 93 Transcription elongation factor S-II FV3 81R 100 YP_031660
81 89980-90453 158 Immediate early protein ICP-18 FV3 82R 100 YP_031661
82 90903-91547 215 Cytosine DNA methyltransferase FV3 83R 100 YP_031662
83 91919-92656 246 Proliferating cell nuclear antigen (PCNA) PPIV 84R 100 ANZ56939
84 92731-93318 196 Thymidine kinase FV3 85R 100 YP_031664
85 93708-93893 62 FV3 44R 98 YP_031665
86 94246-96045 600 SSME 87L 100 AHM26163
87 96078-96530 151 Thiol oxidoreductase GGRV 65R 99 AJR29223
88 96598-97881 428 SSME 89R 100 AHM26165
89 97969-99360 464 Major capsid protein SSME 90R 100 AHM26166
90 99484-100671 396 Immediate early protein ICP-46 SSME 91R 100 AHM26167
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
94 102261-103352 364 DNA repair enzyme RAD2 STIV 100R 100 ACF42318
95 104153-104824 224 FV3 96R 100 YP_031675
96 104907-105320 138 Myeloid cell leukemia protein STIV 103R 100 ACF42321
97 105884-106549 222 US22 family protein STIV 104R 100 ACF42322
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
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
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
34 35450-35704 85 LITAF/PIG7 possible membrane associated motif in LPS-
induced tumor necrosis factor alpha factor
ADRV 33R 100 AGV20564
71
Table 2-13. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
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
52 58124-58354 77 Nuclear calmodulin-binding protein FV3 54L 100 YP_031632
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
Table 2-13. Continued
ORF
Position (nt
range)
Product
size (aa) Predicted function and conserved domain or signature
Best BLAST hit
Ranavirus ORF % ID Accession no.
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
77 81847-84759 971 Tyrosine kinase CGSIV 83L 99 AHA42351
78 85317-86018 234 eIF-2alpha-like protein ADRV 84L 100 AGV20615
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.
Materials and Methods
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
Bufo marinus Venezuelan iridovirus 2 b
Mg1 Venezuela AF157677 + + Hyatt et al. 2000
Bufo marinus Venezuelan iridovirus 3 b
Mat2 Venezuela AF157675 + + Hyatt et al. 2000
Bufo marinus Venezuelan iridovirus 4 b
Cum5 Venezuela AF157663 + + Hyatt et al. 2000
Bufo marinus Venezuelan iridovirus 5 b
Cum6 Venezuela NE + Hyatt et al. 2000
Bufo marinus Venezuelan iridovirus 6 b
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
Table 3-3. Continued
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.
Materials and Methods
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.
Figure 4-8. Hematoxylin and eosin (H&E) and RNAscope® in situ hybridization (ISH) results
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.
Figure 4-9. Hematoxylin and eosin (H&E) and RNAscope® in situ hybridization (ISH) results
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.
Figure 4-10. Hematoxylin and eosin (H&E) and RNAscope® in situ hybridization (ISH) results
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.