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CHARACTERIZATION OF A NOVEL PICORNAVIRUS ISOLATED FROM MORIBUND AQUACULTURED CLOWNFISH By ELIZABETH C. SCHERBATSKOY A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIRMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2020

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Page 1: CHARACTERIZATION OF A NOVEL PICORNAVIRUS ISOLATED …

CHARACTERIZATION OF A NOVEL PICORNAVIRUS ISOLATED

FROM MORIBUND AQUACULTURED CLOWNFISH

By

ELIZABETH C. SCHERBATSKOY

A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF

FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIRMENTS FOR THE DEGREE OF

MASTER OF SCIENCE

UNIVERSITY OF FLORIDA

2020

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© 2020 Elizabeth Catherine Scherbatskoy

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To my parents, for teaching me to pursue my passions unwaveringly,

and to Granny, for lighting the way

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ACKNOWLEDGMENTS

This thesis was made possible by the cumulative effort of a great many individuals and

establishments, and it is with great pleasure that I extend my formal gratitude to them. First and

foremost, I would like to express my profound gratitude to my family, particularly my parents

Dr. Timothy Scherbatskoy and Carin Cooper, my brother Alexander Scherbatskoy and sister-in-

law Meghan Scherbatskoy, and my grandparents Ruth (Granny) Cooper, Abraham Cooper, Mary

Ellen Scherbatskoy, and Serge Scherbatskoy Sr. Thank you for inspiring me to do my best each

day. This gratitude extends to the rest of my unwavering support system of family, friends, and

animals, without whom these last four years would’ve seen far less laughter and far more panic

attacks.

I would like to thank the University of Florida (UF) for supporting my work through their

generous four-year UF Alumni Fellowship. I would also like to thank the UF Graduate Student

Council (GSC) and the Veterinary Graduate Student Association (VGSA) for providing travel

grant opportunities, enabling me to present my research at both domestic and international

conferences.

I would like to express appreciation for my major professor, Dr. Thomas Waltzek, for his

assistance with this degree, as well as all of my advisory committee members, Drs. Salvatore

Frasca Jr., Terry Fei Fan Ng, Kuttichantran Subramaniam, and Roy Yanong for their insight and

expertise which helped to shape both my thesis and my graduate school experience.

I am grateful to all of the faculty, staff, and students who contributed so much to my M.S.

and to my education in molecular biology. It has been an honor and a blessing getting to work

with such talented, hard-working, and humorous individuals during my time at UF.

Finally, I am thankful for the collaborations that I was able to be a part of with the

following individuals: Melissa Brown and the UF CVM diagnostic laboratories for their

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assistance processing histological samples, Dr. Andrew Kane & Ross Brooks for their assistance

around the UF Aquatic Pathobiology Laboratory, Deborah Pouder at the UF Tropical

Aquaculture Laboratory for her laboratory expertise and valuable water quality lessons

(“Niagra”); Dr. Vsevolod Popov of the University of Texas Medical Branch at Galveston, Texas

and Dr. Karen Kelley at the UF Electron Microscopy Core for their skillful processing of

electron microscopy samples, and Dr. Jeffrey Wolf of the Experimental Pathology Laboratories

for his expertise and assistance interpreting histology slides.

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TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...............................................................................................................4

LIST OF TABLES ...........................................................................................................................8

LIST OF FIGURES .........................................................................................................................9

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

CHAPTER

1 PICORNAVIRUSES ..............................................................................................................12

Introduction .............................................................................................................................12 Picornaviruses in Fish .............................................................................................................14

Transmission and Pathology ............................................................................................16 Detection Methods ...........................................................................................................17

Polymerase chain reaction ........................................................................................18 Real-time polymerase chain reaction .......................................................................18 In situ hybridization .................................................................................................19 Immunofluorescence ................................................................................................20

Rivers’ Postulates ............................................................................................................21

2 CHARACTERIZATION OF A NOVEL PICORNAVIRUS ISOLATED FROM

AQUACULTURED CLOWNFISH .......................................................................................23

3 METHODS .............................................................................................................................29

Parasitology, Bacteriology, and Histopathology ....................................................................29 Virus Isolation ........................................................................................................................31 Transmission Electron Microscopy ........................................................................................32 Genomic Characterization and Phylogenetic Analysis ...........................................................34 RNA Extraction and Development of a CFPV RT-PCR Assay .............................................35 Testing Archived Clownfish Tissue Samples by RT-PCR .....................................................36

4 RESULTS ...............................................................................................................................40

Parasitology, Bacteriology, and Histopathology ....................................................................40 Virus Isolation ........................................................................................................................40 Transmission Electron Microscopy ........................................................................................41 Genomic Characterization and Phylogenetic Analysis ...........................................................41 Development of a CFPV RT-PCR Assay ...............................................................................42

5 DISCUSSION .........................................................................................................................50

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LIST OF REFERENCES ...............................................................................................................56

BIOGRAPHICAL SKETCH .........................................................................................................63

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LIST OF TABLES

Table page

3-1 Sequences used for phylogenetic analysis .........................................................................37

3-2 CFPV conventional RT-PCR primer set ............................................................................39

4-1 Predicted genome organization of the clownfish picornavirus ..........................................43

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LIST OF FIGURES

Figure page

2-1 Clownfish species and their host anemones.......................................................................28

4-1 Histologic sections of branchial cavity and alimentary tracts of a clownfish sampled

from a CFPV-positive population ......................................................................................44

4-2 In vitro growth characteristics of the CFPV-2015 isolate in SSN-1 cells .........................45

4-3 Ultrastructural features of the CFPV-2015 isolate in SSN-1 cells ....................................46

4-4 Annotated CFPV polyprotein with sequence identity matrices for each of the P1, 2C,

3C & 3D regions ................................................................................................................47

4-5 Phylogenetic analysis of picornavirus 3Dpol gene sequences ............................................48

4-6 CFPV alignment with six other fish picornaviruses ..........................................................49

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Abstract of Thesis Presented to the Graduate School

of the University of Florida in Partial Fulfillment of the

Requirements for the Degree of Master of Science

CHARACTERIZATION OF A NOVEL PICORNAVIRUS ISOLATED

FROM MORIBUND AQUACULTURED CLOWNFISH

By

Elizabeth Catherine Scherbatskoy

December 2020

Chair: Thomas Waltzek

Major: Veterinary Medical Sciences

Over the last decade, a number of U.S. aquaculture facilities have experienced periodic

mortality events of unknown etiology in clownfish (Amphiprion ocellaris). Clinical signs of

affected individuals included lethargy, altered body coloration, reduced body condition,

tachypnea, and abnormal positioning in the water column. Samples from outbreaks were

processed for routine parasitological, bacteriological, and virological diagnostic testing, but no

consistent parasitic or bacterial infections were observed. Histopathological evaluation revealed

individual cell necrosis and mononuclear cell inflammation in the branchial cavity, pharynx,

esophagus, and/or stomach of four examined clownfish, and large basophilic inclusions within

the pharyngeal mucosal epithelium of one fish. Homogenates from pooled external and internal

tissues from these outbreaks were inoculated onto striped snakehead (SSN-1) cells for virus

isolation and cytopathic effects were observed, resulting in monolayer lysis in the initial

inoculation and upon repassage. Transmission electron microscopy of infected SSN-1 cells

revealed small round particles (20.0 – 21.7 nm in diameter) within the cytoplasm, consistent with

the ultrastructure of a picornavirus. Full genome sequencing of the purified virus revealed a

novel picornavirus most closely related to the bluegill picornavirus and other members of the

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genus Limnipivirus. Additionally, pairwise protein alignments between the clownfish

picornavirus (CFPV) and other known members of the genus Limnipivirus yielded results in

accordance with the current International Committee on Taxonomy of Viruses criteria for

members of the same genus. Thus, the clownfish picornavirus represents a proposed new

limnipivirus species. Future experimental challenge studies are needed to determine the role of

the CFPV in disease.

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CHAPTER 1

PICORNAVIRUSES

Introduction

The superfamily of picorna-like viruses is comprised of many different RNA virus

families that infect everything from plants to animals to unicellular eukaryotic organisms. All of

these viral families appear to be evolutionarily related to picornaviruses and are thought to have

likely evolved in a “Big Bang” diversification event prior to the dispersion of the five major

eukaryotic supergroups (Unikonta, Plantae, Chromalveolata, Rhizaria, and Excavata). This

hypothesis is supported by phylogenetic analyses which indicate the presence of picorna-like

viruses in hosts belonging to two or three different eukaryotic supergroups, and it is thus

believed that early picorna-like viruses invaded ancestral eukaryotic hosts, rather than a

concomitant virus-host co-evolution following the divergence of the major eukaryotic

supergroups that we know today (Koonin et al. 2008).

The family Picornaviridae is currently comprised of more than 100 different species,

grouped within 47 separate genera (Aalivirus, Ailurivirus, Ampivirus, Anativirus, Aphthovirus,

Aquamavirus, Avihepatovirus, Avisivirus, Bopivirus, Cardiovirus, Cosavirus, Crohivirus,

Dicipivirus, Enterovirus, Erbovirus, Gallivirus, Harkavirus, Hepatovirus, Hunnivirus,

Kobuvirus, Kunsagivirus, Limnipivirus, Livupivirus, Malagasivirus, Megrivirus, Mischivirus,

Mosavirus, Orivirus, Oscivirus, Parechovirus, Pasivirus, Passeriviris, Poecivirus,

Potamipivirus, Rabovirus, Rafivirus, Rosavirus, Sakobuvirus, Salivirus, Sapelovirus,

Senecavirus, Shanbavirus, Sicinivirus, Teschovirus, Torchiviris, Tottorivirus and Tremovirus).

However, many of these species await formal classification, as the number of species and genera

has more than doubled over the past few years as a result of increased use of next-generation

sequencing (NGS) technologies (Zell et al. 2017, King et al. 2018). Members of the

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Picornaviridae family possess non-enveloped, round to icosahedral nucleocapsids of

approximately 30 nm in diameter (Racaniello 2013, Jiang et al. 2014). Within the nucleocapsid

lies a single-stranded positive-sense RNA genome, ranging from 6.7 to 10.1 kb in length, which

typically contains a single long open reading frame (ORF), with the exceptions of canine

picodicistrovirus and hedgehog dicipivirus in the genus Dicipivirus which contain two (Woo et

al. 2012, Reuter et al. 2018). The 3’ end of the genome is polyadenylated, while the 5’ end is

covalently linked to a small viral protein, VPg. The 5’ and 3’ ends of the genome have

untranslated regions (UTRs), both of which include functionally important secondary structures,

such as the internal ribosome entry site (IRES) within the 5’ UTR, essential for ribosome binding

and cap-independent translation. The typical picornavirus genome encodes a single polyprotein

that is divided into three regions, P1, P2, and P3. The P1 region contains smaller structural

proteins, 1A, 1B, 1C, and 1D, which encode four viral proteins (VP1-VP4) responsible for

capsid formation and initiation of the virus into host cells through receptor binding. An N-

terminus leader protein (L) is also present in the P1 region of some picornavirus genera (e.g.,

Aphthovirus and Kobuvirus), preceding the P1 region. The P2 and P3 proteins encode non-

structural proteins necessary for viral replication, 2A, 2B, 2CATPase and 3A, 3BVPg, 3Cpro, 3Dpol,

respectively. 2A and 2B interfere with host cell functions, while 2CATPase is associated with

vesicle formation. The 3A protein is involved with membrane protein presentation and cellular

protein transport inhibition, while 3BVPg, a genome-linked protein, acts as a primer for RNA

synthesis during viral replication. The 3Cpro protein is a protease responsible for the cleavage of

the P1 precursor protein, and the 3Dpol protein is an RNA-dependent RNA polymerase required

for viral replication (Lin et al. 2009, Racaniello 2013, Jiang et al. 2014, Zell et al. 2017).

However, these functions may vary among different picornaviruses (TFF Ng pers. obs. 2019).

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Picornaviruses are a diversified group of pathogens of which infections range in severity

from very mild, such as the common cold, to extremely serious, such as poliomyelitis, hepatitis,

and encephalitis. The Picornaviridae family contains a number of well-known diseases of

humans and animals, among which are poliovirus and rhinovirus (genus Enterovirus), foot-and-

mouth disease virus (genus Aphthovirus), and hepatitis A virus (genus Hepatovirus) (Jiang et al.

2014). As such, picornaviruses have played a significant role in the history of virology.

Poliovirus is thought to have existed since 1400 BCE, due to the discovery of ancient Egyptian

artworks depicting deformities consistent with paralytic poliomyelitis (Mehndiratta et al. 2014).

Foot-and-mouth disease virus was the first identified animal virus, discovered in 1898 by

Friedrich Loeffler and Paul Frosch, closely followed by the isolation of poliovirus by Karl

Landsteiner and Erwin Popper a decade later (Skern 2010, Racaniello 2013). These two viral

diseases have been the most studied of all the picornaviruses and were two of the first viruses to

have vaccines developed against them (Tuthill et al. 2010, Racaniello 2013).

Picornaviruses in Fish

While picornaviruses have been known to infect a wide variety of animals, confirmed

reports in fish have been limited until recently. Over the last thirty years, fish picorna-like

viruses have been isolated in cell cultures and/or visualized by electron microscopy in infected

cultures or tissues without sequence confirmation. For example, picorna-like viruses were

reported in rainbow and European smelt Osmerus mordax and O. eperlanus (Moore et al. 1998,

Ahne et al. 1990), Atlantic salmon Salmo salar and other salmonids (Hedrick et al. 1990,

Hedrick et al. 1991, Eaton et al. 1992, Iwanowicz et al. 2017), barramundi Lates calcarifer

(Glazebrook et al. 1990, Munday et al. 1992), turbot Scophthalmus maximus (Bloch et al. 1991),

sea bass Dicentrarchus labrax (Breuil et al. 1991), redspotted grouper Epinephelis akaara (Mori

et al. 1991), sandbar shiners Notropis scepticus (Iwanowicz et al. 2000), rainbowfish

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Melanotaenia lacustris (Petty and Fraser 2005), and common carp Cyprinus carpio (Reuter et al.

2014). However, many of these picorna-like viruses were later genetically characterized as

hepeviruses (Batts et al. 2011) or betanodaviruses (reviewed in Bovo and Florio 2008 and

Walker and Winton 2010).

More recently, four fish viruses were isolated and confirmed by genome sequencing to be

picornaviruses. One, a picornavirus found in bluegill Lepomis macrochirus, was isolated

following a bluegill fish kill in Wisconsin (Barbknecht et al. 2014) while another, a carp

picornavirus, was discovered when a pond of common carp Cyprinus carpio were accidentally

killed as a result of a liquid manure spill in Germany (Lange et al. 2014). A third picornavirus

was isolated from fathead minnows Pimephales promelas, collected during routine surveillance

of apparently healthy stock in the northcentral United States (Phelps et al. 2014) while another,

the eel picornavirus, was isolated from European eels Anguilla anguilla collected following a

morbidity and mortality event in the Lake Constance area of Europe (Fichtner et al. 2013). The

bluegill, common carp, and fathead minnow picornaviruses all belong to the Limnipivirus genus

in the Picornaviridae family, while the eel picornavirus belongs to the Potamipivirus genus (Zell

et al. 2017 and King et al. 2018). More recently, the genome of a novel potamipivirus was

discovered in intestinal tissue samples derived from apparently healthy threespine stickleback

Gasterosteus aculeatus in Alaska (Hahn and Dheilly 2019). Similarly, the genome of a

picornavirus derived from the gut tissues of asymptomatic zebrafish Danio rerio was discovered

in lab fish in North America, Europe, and Asia, and was found to be divergent enough from other

fish picornaviruses to constitute the type species of a newly proposed genus, “Cyprivirus” (Altan

et al. 2019).

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In 2018, two studies employing metagenomic approaches resulted in the discovery of

twenty-nine new fish picornaviruses, a great leap forward from the handful of recognized fish

picornavirus species (Geoghegan et al. 2018 and Shi et al. 2018). The sequences of these 29

unclassified fish picornaviruses were determined from specimens that included both wild

freshwater and marine species with representatives from lobe-finned (Sarcopterygii), ray-finned

(Actinopterygii) and cartilaginous (Chondrichthyes) fishes. These studies show that

picornaviruses in fish are a rapidly expanding group of viruses, with more and more species

being discovered each year. As a result, there is a need for fish picornavirus research to further

our understanding of their phylogenetic relationships, their pathogenicity and transmission, their

detection, and their prevention.

Transmission and Pathology

While picornaviruses have been found in a number of different fish species as discussed

above, picornavirus infections in fish are relatively new discoveries and the role that they play in

disease has yet to be determined in many cases. Very little is known about the transmission of

fish picornaviruses, although infection trials have been performed with a few fish picornaviruses.

The eel picornavirus was used to experimentally infect European eels (A. anguilla) using a water

bath, which yielded mortalities in 7 of the 16 challenged fish and successful re-isolation of the

virus from all of those infected (Fichtner et al. 2013). The carp picornavirus was used to infect

common carp (C. carpio) using both a water bath and intracoelomic injection, and while none of

the fish showed clinical signs, the virus was successfully re-isolated from all of the fish

inoculated via injection. The bath challenge yielded no clinical signs nor re-isolation (Lange et

al. 2014). Laboratory infection trials using the bluegill picornavirus yielded morbidity, mortality,

and re-isolation of the virus following intracoelomic injection into bluegill (L. macrochirus), and

while a survey of waters in Wisconsin showed widespread prevalence of the virus in bluegill

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populations (Barbknecht et al. 2014), the route of transmission in the wild remains to be seen, as

is the case for all of the above picornaviruses.

The pathology associated with picornavirus infection in fishes ranges from clinical

insignificance to severe morbidity and mortality. The carp, zebrafish, threespine stickleback, and

baitfish picornaviruses have all been found in asymptomatic fishes, although the baitfish

picornavirus has also been found in a small number of fish with ocular and dermal hemorrhages

(Lange et al. 2014, Phelps et al. 2014, Hahn and Dheilly 2019, Altan et al. 2019). Gross

pathology associated with the European eel picornavirus included increased mucus production,

erythema, ulcers, and mortality (Fichtner et al. 2013), while that of the bluegill picornavirus

included inflammation, erythema, exophthalmia, coelomic distention, ascites, and internal

hemorrhaging (although the virus has also been isolated from asymptomatic bluegill)

(Barbknecht et al. 2014). Microscopically, the European eel picornavirus was shown to be

associated with single cell necrosis and necrotic foci (Fichtner et al. 2013), while no microscopic

pathology was reported to be associated with the bluegill picornavirus (Barbknecht et al. 2014).

Detection Methods

The detection and characterization of fish picornaviruses has been facilitated by

laboratory techniques including cell culture, electron microscopy, immunofluorescence, and

various molecular approaches. While many of the more recently discovered fish picornaviruses

were determined through metagenomic approaches using NGS technologies, the discovery of

many of the fish picornaviruses relied on virus isolation through cell culture, electron

microscopy, virus-specific end-point reverse-transcription polymerase chain reaction (RT-PCR)

and real-time reverse-transcription quantitative PCR (RT-qPCR) assays, and hybridization

assays.

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Polymerase chain reaction

End-point, or conventional, PCR (and RT-PCR) technology was developed by Kary

Mullis in 1983 and allows for the simple and sensitive detection of a particular nucleic acid

sequence through the use of deoxyribonucleotides and polymerase enzymes, along with specific

oligomers (i.e., primers) and template DNA (or RNA, in the case of picornaviruses) (Mullis

1990, Garibyan and Avashia, 2013). The primers are short complementary fragments of DNA

designed to flank the specific nucleic acid sequence to be amplified, allowing for its extension

via the polymerase enzyme and nucleotides. Following amplification of the target nucleic acid,

the PCR products must be visualized, through chemical or fluorescent dyes and the use of

agarose gel electrophoresis (Garibyan and Avashia, 2013).

A multiplex RT-PCR assay was developed by Mor et al. (2015) for the detection of three

fish picornaviruses: the bluegill picornavirus, the fathead minnow picornavirus, and the

European eel picornavirus. Primers for the assay were designed to target the RdRp gene of each

of the three viruses (Mor et al. 2015). Conventional RT-PCR technology was also used to detect

the threespine stickleback picornavirus, using primers which targeted the RNA-dependent RNA

polymerase region of the genome (Hahn and Dheilly 2019), and the bluegill picornavirus, using

primers targeting the 3’ UTR (Barbknecht et al. 2014). A nested conventional RT-PCR assay

using two sets of primers was used to fill in gaps in the zebrafish picornavirus genome sequence

after deep sequencing, as well as to detect the virus in zebrafish (Altan et al. 2019). Additionally,

RT-PCR technology was also used to determine information about the genome sequences of the

baitfish picornavirus (Phelps et al. 2014) and the carp picornavirus (Lange et al. 2014).

Real-time polymerase chain reaction

Real-time, or quantitative, polymerase chain reaction (qPCR) assays take this technology

a step further by allowing the amount of nucleic acids in a given sample to be quantified

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throughout the PCR process, typically through the use of a fluorescent probe. This probe,

included in the reaction mixture, increases as the target product increases, allowing for the

amount of target to be measured (Klein, 2002; Peirson & Butler, 2007). During the qPCR

process, a 6-carboxy-X-rhodamine (ROX)-normalized 6-carboxyfluorescein (FAM) signal is

emitted as the sequence-specific probe binds to its target (Garibyan & Avashia, 2013), crossing a

theoretical cycle threshold (Ct) which enables the user to determine the nucleic acid

concentration of the sample (Valasek & Repa, 2005). The viral copy number present in each

sample can be obtained by comparing results to known standards in the form of a standard curve

(e.g., ranging from 10 to 107 copies). This is in contrast to end-point PCR (i.e., conventional

PCR), which is qualitative rather than quantitative and establishes the presence or absence of

nucleic acids but not the amount (Garibyan & Avashia, 2013). Additionally, procedures to

analyze and visualize the data following conventional PCR are required, which can be time-

consuming and lead to cross-contamination (Klein, 2002; Garibyan & Avashia, 2013).

In addition to the nested conventional PCR assay that was developed for the zebrafish

picornavirus, a real-time RT-qPCR assay was designed for the rapid detection of the virus. The

assay used standard primer and probe concentrations (Applied BioSystems™) and the

LightCycler 480 Probes Master master mix (Roche Applied Science, Indianapolis, IN). A

hydrolysis probe-based RT-qPCR assay designed to target a universal bacterial

reference gene (16s rRNA) was used for all samples to rule out PCR inhibition and to verify the

presence of amplifiable DNA, as well as positive and negative controls for the zebrafish

picornavirus-specific assay (Altan et al. 2019).

In situ hybridization

While PCR and qPCR are extremely helpful when searching for nucleic acids present in a

given sample, other techniques become necessary in order to visualize the positive or negative

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results within the context of the tissue itself (Wang et al., 2012). One such technique is in situ

hybridization, or ISH, which allows one to visualize viral nucleic acids within microscopic

lesions of the host (if present). In situ hybridization is a process by which specific nucleic acid

sequences can be localized within a given histologic section through the use of labeled probes.

These sequence-specific probes are complementary to the target DNA or RNA and can be

detected through the use of either fluorescence, radioactivity, or antigen-labeling (Jensen 2014).

RNAscope® is a relatively recent evolution of ISH technology in which single molecules of

target RNA can be visualized within intact cells, while background noise is simultaneously

suppressed through the use of uniquely designed probes. This leads to signal amplification of the

desired sequence with very little interference. Both chromogenic and fluorescent dyes can be

used with an RNAscope® assay, for use with bright-field or epifluorescent microscopy,

respectively (Wang et al., 2012).

RNAscope® technology has been used for the detection of the zebrafish picornavirus. A

zebrafish picornavirus-specific assay was designed to detect the virus in thin (5 μm) sections of

formalin-fixed, paraffin-embedded zebrafish tissue mounted on AutoFrost charged adhesion

slides (Cancer Diagnostics, Inc, Durham, NC), using the chromogenic substrate Fast Red. A

probe targeting the bacterial DapB gene was used as a negative control (Altan et al. 2019).

Immunofluorescence

Immunofluorescence assays can be used to detect viruses by determining the presence of

a specific antigen bound to fluorescently labeled antibodies. There are two main types of these

assays, referred to as direct or indirect immunofluorescence assays. With direct

immunofluorescence (DIF), an antibody conjugated to a fluorescent label binds directly to the

antigen of interest. Indirect immunofluorescence assays (IFA) are two step processes which use

two antibodies. The primary antibody is unlabeled and binds to the antigen of interest, while the

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secondary antibody is fluorescently labeled and binds to the primary antibody. While this

technique is more complicated and takes more time, it is also more sensitive as multiple

secondary antibodies can bind to the primary antibodies, producing a greater signal (Odell 2013).

An indirect immunofluorescence assay was used to detect the presence of the carp

picornavirus in infected monolayers of fathead minnow (FHM) cells. The assay used the

monoclonal antibody 5C9 and a secondary indocarbocyanine-conjugated goat anti-mouse IgG

antibody. The cross-reactivity of 5C9 was tested by using positive reagents on cells infected with

the eel picornavirus, among other viruses (Lange et al. 2014). Similarly, an indirect

immunofluorescence assay was designed to detect the European eel picornavirus, using rabbit

antiserum T51 and fluorescein isothiocyanate-conjugated goat anti-rabbit IgG (Fichtner et al.

2013).

Rivers’ Postulates

In order to definitively confirm that a particular pathogen is the true etiological cause of a

disease, as well as to help characterize the pathology associated with the disease, naïve

individuals must be challenged with the pathogen (Williams 2010). Koch’s postulates, developed

to determine whether a microorganism is the true agent responsible for a particular disease, state

that a pathogen must be (1) found in only diseased hosts, (2) cultured from said diseased hosts,

(3) inoculated into a healthy host and cause comparable disease, and (4) be re-cultured from the

newly diseased host and shown to correspond to the original pathogen (Segre 2013). The

examination of viral diseases necessitates further criteria however, as viruses are obligate

intracellular parasites which must be grown in living cells and thus cannot be grown on artificial

media (e.g., agar plates, nutrient broth) like most bacteria (Williams 2010). Cell culture

propagation of viruses can lead to unintended alterations, such as adaptations of the virus to, or

even attenuation of the virus in, the particular cell lines used (Prescott et al. 2017). In addition to

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this, viruses present further challenges when trying to fulfill criteria for the causation of a

disease, such as asymptomatic or latent viral infections, prolonged viral shedding following an

acute disease episode, or viruses which cause clinical disease in only a small number of infected

hosts (Williams 2010). Koch’s postulates were therefore altered by Thomas Rivers in 1937 for

the examination of viral diseases, and include the following criteria: Similar to Koch’s

postulates, the virus must be (1) isolated from diseased hosts, (2) cultured, (3) re-isolated from

newly infected naïve hosts, and (4) shown to produce comparable disease in said new hosts.

However, Rivers’ postulates include additional criteria necessary when examining viruses, such

as proof of filterability, successful cultivation in host cells in cell culture, and the detection of

specific immune responses to the virus in question (Rivers 1937, Fouchier et al. 2003, Williams

2010). Additional criteria have been proposed over the years to try to accommodate the

aforementioned challenges that viruses can present when trying to establish disease causation,

such as epidemiologic studies, prevention of disease by vaccination, and comparison to known

pathogenic viruses (Williams 2010).

As mentioned previously, of the many fish picornaviruses known, only three have been

used to try to fulfill Rivers’ postulates – the eel, bluegill, and carp picornaviruses. All three of

these viruses were able to be reisolated from infected individuals, but only the eel and bluegill

picornaviruses produced clinical signs in their hosts (Fichtner et al. 2013, Barbknecht et al. 2014,

Lange et al. 2014). However, it should be noted that the carp picornavirus was initially isolated

from a clinically asymptomatic individual to begin with (Lange et al. 2014).

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CHAPTER 2

CHARACTERIZATION OF A NOVEL PICORNAVIRUS ISOLATED FROM

AQUACULTURED CLOWNFISH

Clownfish are members of the Pomacentridae, a large family containing over 300 species

(Fautin and Allen 1992, Sin et al. 1994). Members of the family can be found in marine habitats,

although a few brackish water species can occasionally be found in fresh water. Pomacentrids

primarily inhabit tropical latitudes, with the vast majority residing in the Indo-west and central

Pacific regions. Within the Pomacentridae family are four subfamilies, one of which is the

Amphiprioninae. This subfamily is comprised of the Amphiprion and Premnas genera and

associated species are known informally as the anemonefishes - so called for their symbiotic

relationship with sea anemones (Fautin and Allen 1992) (Fig. 2-1).

Clownfish are among the most popular marine fishes traded in the international

ornamental fish industry, and among them A. ocellaris and A. percula are particularly favored,

along with the maroon clownfish P. biaculeatus (Patkaew et al. 2014). An estimated 90% of all

traded clownfish species are now raised in captivity (King 2019). In the United States alone,

ornamental fish aquaculture is a multimillion-dollar industry (Watson and Shireman 2002), while

the value of the global marine ornamental trade exceeds USD $300 million per year. It is

estimated that 2 million people participate in this trade worldwide each year, either recreationally

or professionally (Wabnitz et al. 2003, Sirajudheen et al. 2014).

The farming of aquatic organisms, otherwise known as aquaculture, is the most rapidly

growing form of agriculture worldwide. Global aquaculture production grew at an average

annual rate of 5.8% between the years 2000-2016 and is only expected to increase in response to

the growing world population. According to FAO projections, it is predicted to reach 109 million

tons by 2030, which represents a 37% increase from 2016 production totals (Ahmed and

Thompson 2019). The international trade of cultured and wild-caught ornamental fishes and

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invertebrates is valued at approximately $278 million USD, while the aquarium industry has

been estimated to be worth over $1,000 million USD according to surveys of the pet industry

(Livengood and Chapman 2007). Ornamental aquaculture is a significant contributor to the

United States aquaculture economy, and the majority of ornamental fish exports from the United

States are raised in the state of Florida (Chapman et al. 2007, Groover et al. 2020). In 2003, the

farm-gate value of tropical fishes produced in Florida totaled approximately $47.2 million USD

(Hill and Yanong 2016). Thus, the successful production of ornamental fish is of great

significance to both the economy of the United States and to the economy of the state of Florida,

emphasizing the importance of biosecurity and the effective prevention of infectious diseases in

aquaculture.

Biosecurity is an important concept in any agricultural or animal husbandry endeavor,

seeking to protect against the spread of disease and/or foreign organisms (Meyerson et al. 2002).

Biosecurity in aquaculture involves the implementation and execution of practices that seek to

prevent the introduction and spread of infectious diseases on and off farms, as well as those that

seek to reduce the stress of aquacultured species (Yanong and Erlacher-Reid 2012). Because

aquaculture, like other agriculture forms, results in high densities of a given species within very

close quarters, the risk of diseases being spread among populations is very high. Among these,

viral diseases present the greatest challenges for aquaculture ventures, as a result of limited

therapeutics, inadequate understanding of viral pathogenesis and host resistance, and

vulnerability of young organisms to disease (Kibenge et al. 2012). There are three main goals of

biosecurity, which focus on the management of (1) animals, (2) pathogens, and (3) people.

Animal management includes obtaining healthy animal stocks, excellent husbandry and

preventative medical practices, and proper quarantine procedures. Important quarantine

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components to consider involve all-in-all-out stocking, isolation of separate populations,

adequate observation and adjustment of these populations, and finally, the testing of samples

from these populations for various diseases and implementation of any necessary treatments.

Pathogen management involves the prevention, reduction, and/or elimination of

infectious agents on aquaculture farms. In order to accomplish this, an understanding of the

disease-causing organisms that may exist at a given facility is required. This includes an

understanding of their pathogenicity and diagnosis, as well as the living and non-living reservoirs

in which they may persist. It is also necessary to understand the regulatory importance of

different pathogens, such as OIE reportable diseases, or those regulated by USDA-APHIS or

governments (Yanong and Erlacher-Reid 2012). While progress has been made in pathogen

diagnostics and control, prevention tends to be more cost effective (Tidbury et al. 2018). Thus,

the application of proper sanitation and disinfection protocols are essential to any aquaculture

endeavor, ideally eliminating or inactivating many potentially infectious microorganisms not

typically found on farms from tanks, equipment, and water (Verner-Jeffreys et al. 2009, Mainous

et al. 2010). Three main methods of disinfection exist, in the form of ozonation, ultraviolet

irradiation, and chemicals (Mainous et al. 2010). There are many different types of chemical

disinfectants, such as alcohols, chlorine, formaldehyde, and Virkon® Aquatic, a disinfectant

using potassium peroxymonosulfate and sodium chloride (Yanong and Erlacher-Reid 2012). The

efficacy of disinfectants has been shown to vary with respect to contact time, temperature, and

organic material present (Tidbury et al. 2018), and it is thus of great importance to gain as much

of an understanding as possible of the pathogens present on a farm so that effective prevention

measures may be taken.

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Finally, the management of people in regard to biosecurity includes ensuring all

individuals going to and from aquaculture facilities understand the importance of adhering to

proper biosecurity practices like those outlined above. In addition to education initiatives

outlining the details of the biosecurity program being followed, setting up and encouraging the

use of disinfection stations is of the utmost importance. For example, footbaths at the entrance

and exits of rooms, handwashing stations and showers, net dips, and areas to disinfect vehicles

traveling in between facilities can all help prevent the spread of pathogens onto and off of farms

(Yanong and Erlacher-Reid 2012).

Staying on top of good biosecurity practices is made easier and more effective by

developing a written biosecurity plan. This plan should include risk assessment, risk

management, and risk communication, and should be developed with a fish health expert,

aquacultural engineer, and an aquaculture production specialist. This plan should be easy to

follow and should be impressed upon those working or visiting the facility (Yanong and

Erlacher-Reid 2012).

There are many different pathogens that affect aquacultured clownfish, from large

ectoparasites to very small viruses. One of these pathogens, a ciliated protozoan named

Brooklynella hostilis, is so common in clownfish that it is informally referred to as “Clownfish

Disease.” This parasite of the skin and gills can cause significant sloughing of the skin,

hemorrhage, and increased mucus production (Lom and Nigrelli 1970). Another common

ciliated protozoan known to infect clownfish is Cryptocaryon irritans, otherwise known as

“marine white spot disease” or “marine ich.” This ectoparasite can cause serious injury to the

fins, skin, and gills of fish, leading to changes in appearance such as pale gills, increased mucus

production, or emaciation, as well as changes in behavior such as flashing, lethargy, or tachypnea

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(Yanong 2009, DiMaggio et al. 2017). Clownfish are also often parasitized by the dinoflagellate

Amyloodinium ocellatum, known familiarly as “marine velvet disease.” A. ocellatum is an

ectoparasite that affects the skin and gills of its fish hosts, which can cause severe disease and

high mortalities in a very short period of time (DiMaggio et al. 2017, Francis-Floyd and Floyd,

2011). Other common parasites of aquacultured clownfish include flagellates and monogeneans

(Vorbach et al. 2016). Common bacteria that affect clownfish include species of Vibrio,

Aliivibrio, and Bacillus, as well as intracellular bacteria known to cause the skin and gill disease

Epitheliocystis (Vorbach et al. 2016, Blandford et al. 2018). A common viral agent of disease in

clownfish is lymphocystis disease virus (LCDV) (Vorbach et al. 2016). LCDV is a member of

the viral family Iridoviridae, and results in chronic wart-like growths on the skin, fins, and gills

of fish, as well as in the eyes and mouth. In clownfish, these warty nodules have been reported,

as well as white spots and fin rot (Siva et al. 2014, Yanong 2010). Microscopically, LCDV has

been shown to lead to hypertrophied cells with basophilic intracytoplasmic inclusions (Lam

2020). While the virus does not typically cause severe mortalities, it can be very damaging to

aquaculture ventures as the altered appearance of the fish can compromise their marketability

(Siva et al. 2014, Yanong 2010).

Over the past decade, a number of aquaculture facilities have experienced large-scale

morbidity and mortality events of unknown etiology in aquacultured A. ocellaris. Clinical signs

associated with these outbreaks included lethargy, increased respiration rates, reduced body

condition, altered body coloration, and holding an abnormal position in the water column, in

addition to mass die-offs. These disease episodes have thus resulted in significant economic and

production losses to these producers, threatening their livelihood and the profitable rearing of

clownfish on farms (RPE Yanong pers. obs. 2018). In this investigation, we describe the

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characterization of a novel picornavirus isolated from moribund A. ocellaris, provisionally

named the clownfish picornavirus (CFPV), which we believe to be the etiological agent

responsible for these mass mortality events. Additionally, we report the in vitro growth

characteristics microscopic pathology, and ultrastructural features of the virus. This was

achieved through the use of cell culture, histopathology, transmission electron microscopy

(TEM), reverse transcription polymerase chain reaction (RT-PCR), next-generation sequencing

(NGS), and phylogenetic analyses.

Figure 2-1. Clownfish species and their host anemones. (A) Amphiprion ocellaris hosted by

Stichodactyla gigantea, (B) Amphiprion perideraion hosted by Heteractis magnifica,

(C) Amphiprion sandaracinos hosted by Stychodactyla mertensii, and D) Premnas

biaculeatus hosted by Entacmaea quadricolor. Photos courtesy of Thomas Waltzek.

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CHAPTER 3

METHODS

Parasitology, Bacteriology, and Histopathology

In 2015, an aquaculture facility experienced chronic morbidity and mortality in juvenile

clownfish (A. ocellaris) reared in a recirculating system. Diseased fish exhibited a range of non-

specific clinical signs, including lethargy, altered body coloration, reduced body condition,

tachypnea, and holding an abnormal position in the water column. The water quality of the

recirculating system was assessed on-site using a Model FF-3 Saltwater Aquaculture Test kit

(Hach Co.) and found to be within normal limits including total ammonia: 0 ppm; nitrite: 0 ppm;

total alkalinity: 205 ppm; dissolved oxygen: 6.9 ppm (YSI 550A dissolved oxygen meter);

salinity: 27 ppt (Vital Sine Salinity Refractomer); pH: 7.87 (Pinpoint pH meter PH370); and

water temperature: 26.7°C.

Ten juvenile clownfish from affected tanks were shipped overnight to the Wildlife and

Aquatic Veterinary Disease Laboratory (WAVDL) in Gainesville, FL for virological

examination, and twenty were shipped to the Tropical Aquaculture Laboratory (TAL) in Ruskin,

FL for parasitological, bacteriological, and histopathological examination. The weight of the fish

ranged from 0.48 - 1.84 g and their standard lengths ranged from 2.6 - 4.6 cm in total length. Of

the shipped fish, ten fish arrived alive to the WAVDL and were processed for virus isolation as

described below; thirteen fish arrived alive to the TAL with eight fish processed for parasitology

and bacteriology, and five fish processed for histopathology.

In 2018, the same aquaculture facility experienced a similar chronic morbidity and

mortality event as described above. The water quality was found to be mostly within normal

limits including total ammonia: 0 ppm; nitrite: 0.23 ppm; total alkalinity: 304.3 ppm; dissolved

oxygen: 5.7 ppm; salinity: 22 ppt; pH: 7.86, and water temperature: 30°C. Eight juvenile

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clownfish from affected tanks were shipped overnight to the WAVDL in Gainesville, FL for

virological examination, and an equal number were shipped to the Tropical Aquaculture

Laboratory (TAL) in Ruskin, FL for parasitological, bacteriological, and histopathological

examination. The weight of the juvenile A. ocellaris ranged from 0.7 - 1.8 g and their standard

lengths ranged from 3.7 – 4.6 cm in total length. Of the eight shipped fish, eight arrived alive to

the WAVDL and were processed for virus isolation as described below. Of the eight fish shipped

to TAL seven fish arrived alive, with four fish processed for parasitology and bacteriology and

three fish processed for histopathology.

For both the 2015 and 2018 cases, parasite burdens were assessed at the TAL by

examining fin, skin, and gill biopsies. Wet mounts of each tissue biopsy were examined by light

microscopy at 40x, 100x, and 200x magnifications within five min of collection. Immediately

following biopsy collection, clownfish were euthanized in 500 mg/L tricaine methane sulfonate

(MS-222®, Syndel Inc., Tricaine-S®) buffered with an equal concentration of sodium

bicarbonate. After euthanasia, bacterial cultures were obtained using sterilized metal loops to

aseptically sample brain and posterior kidney for inoculation onto plates of tryptic soy agar

(TSA) with 5% sheep blood. Culture plates incubated at 28°C for 48 hours and observed daily

for presence/absence of bacterial growth. Following bacteriology, necropsies were conducted,

and wet mounts of liver, spleen, anterior kidney, posterior kidney, stomach, and intestine were

examined by light microscopy at 40x, 100x, and 200x magnifications.

Small ventral body wall incisions were made into the coelomic cavities of five clownfish

from the 2015 case and three clownfish from the 2018 case to facilitate fixative penetration, and

the fish were placed whole into 10% neutral buffered formalin for 48 hours. The fixed fish were

cut into multiple transverse slabs through the head and trunk regions, and then were processed by

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embedment in paraffin, microtome sectioning at ~5 μm thickness, and staining with hematoxylin

and eosin (H&E). Slides were examined by brightfield microscopy for histopathological changes

at 40x, 100x, 200x and 400x magnifications.

Virus Isolation

For both the 2015 and 2018 cases, virus isolation was attempted at the WAVDL using

three cell lines: Epithelioma papulosum cyprini (EPC), grunt fin (GF), and striped snakehead

(SSN-1). Cell lines were maintained in L-15 medium (Leibovitz; Gibco, USA) containing 10%

Fetal bovine serum (FBS; Gibco, USA), and 1X Antibiotic-Antimycotic (AA; Gibco, USA)

resulting in a final concentration of 100 IP penicillin mL-1, 100 μg streptomycin mL-1, and 0.25

μg Amphotericin B mL-1. EPC and GF cells were incubated at 21°C, and SSN-1 cells were

incubated at 25°C.

The submitted clownfish were euthanized using tricaine methanesulfonate at a

concentration of 500 mg/L (MS-222®, Western Chemical Inc) based on an Animal Use and Care

Procedure (IACUC, Cornell), and divided into pools containing four fish each. Internal (kidney,

liver, spleen, and heart) and external (gill and skin) tissue samples were taken from each fish in

each of the pools and kept separate. Each of the tissue pools was diluted 1:25 in L-15 media and

then homogenized at high speed with a stomacher (Seward stomacher 80, Biomaster Lab system)

for 30 seconds. Two hundred microliters of each homogenate were pipetted into microcentrifuge

tubes and placed on ice for RNA extraction (see below), while the rest of the homogenates were

moved to 15 mL conical tubes and used to inoculate cells for virus isolation. The internal and

external clownfish tissue homogenates were then centrifuged at 3,000 x g for 10 minutes at 4°C

to pellet cellular debris. The clarified supernatant from each sample was then pipetted into new

15 mL conical tubes, and an equal volume of L-15 media containing 2X AA was added to each

tube to make a final dilution of 1:50 and a final concentration of 500 IP penicillin mL-1, 500 μg

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streptomycin mL-1, and 12.5 μg Amphotericin B mL-1. The tubes of supernatant were then

incubated at 4°C overnight.

The following day, the supernatant was clarified once again, and 200 μL of each clarified

tissue homogenate was inoculated onto triplicate wells of confluent monolayers of EPC, GF, or

SSN-1 cells grown within 24-well plates. The plates were rocked every 15 minutes for one hour

at 21°C for EPC and GF cells and 25°C for SSN-1 cells. After this time, the supernatant was

removed from each of the infected wells and fresh L-15 media with 2% FBS and 1X AA was

added. The EPC and GF plates were then moved to an incubator set at 21°C, while the E-11 plate

was incubated at 25°C. Triplicate negative control wells were inoculated with L-15

supplemented with 2% FBS and 1X AA. All cell lines were monitored daily for the development

of cytopathic effects (CPE). Upon the appearance of extensive CPE, the supernatant was

clarified and passaged onto recently split cells to rule out toxicity and confirm that the effects

were the result of a passageable agent. Wells not displaying CPE were left for 14 days, after

which time the clarified supernatant was passaged onto fresh cells and observed for an additional

14 days before the samples were considered negative. Clarified supernatant from cultures

displaying CPE in both the first and second passage were frozen in liquid nitrogen for

downstream transmission electron microscopy and genomic sequencing.

Transmission Electron Microscopy

In the 2015 case, a 75 cm2 flask of SSN-1 cells displaying CPE was processed for

transmission electron microscopy (TEM) at the Electron Microscopy Core, Interdisciplinary

Center for Biotechnology Research, University of Florida. Upon the appearance of CPE, cells

were fixed in 15 mL of modified Karnovsky’s fixative (2P+2G, 2% paraformaldehyde and 2%

glutaraldehyde in 0.1 M cacodylate buffer, pH 7.4) for 1 hour at room temperature, and then

washed in cacodylate buffer, scraped and pelleted at 3,000 x g for 10 minutes at 4°C. The

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following day, the pellet was washed in 0.1 M sodium cacodylate buffer (pH 7.24) twice, post-

fixed in 2% OsO4, washed with water, and dehydrated in ascending ethanol series. The

dehydrated pellet was infiltrated with 50 and 100% LR White resin (Electron Microscopy

Sciences) in 3 repetitions followed by overnight incubation at 4°C. Resin-embedded tissue was

cured at 60°C for 48 h. Semi-thick sections were prepared at 500 nm and stained with toluidine

blue. Ultra-thin sections at 80 to 120 nm were collected on carbon-coated Formvar copper 100

mesh grids, post-stained with 2% aqueous uranyl acetate and Reynold’s lead citrate. Sections

were examined with a Hitachi H-7000 TEM (Hitachi High Technologies America), and digital

images were acquired with a Veleta 2k × 2k camera and iTEM software program (Olympus Soft-

Imaging Solutions).

In the 2018 case, a 75 cm2 flask of SSN-1 cells displaying CPE was fixed in 15 mL of

modified Karnovsky’s fixative for 1 hour at room temperature, and then washed in cacodylate

buffer, scraped and pelleted at 3,000 x g for 10 minutes at 4°C. The pellet was resuspended in

phosphate-buffered saline (PBS) and transferred overnight on ice packs to the University of

Texas Medical Branch at Galveston, Department of Pathology Electron Microscopy Laboratory

(UTMB-EML). At UTMB-EML, the pelleted cells were washed in cacodylate buffer and then

fixed in Karnovsky’s 2P+2G fixative overnight at 4°C. The following day, the pellet was washed

in cacodylate buffer twice before being post-fixed in 1% OsO4 in 0.1 M cacodylate buffer (pH

7.4), en bloc stained with 2% aqueous uranyl acetate, and dehydrated in ascending

concentrations of ethanol. It was then processed using propylene oxide and embedded in Poly-

Bed 812 epoxy plastic (Polysciences). Ultrathin sections were cut with a Leica EM UC7

ultramicrotome (Leica Microsystems), stained with 0.4% lead citrate, and examined using a

JEM-1400 electron microscope (JEOL USA) at 80 kV.

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Electron photomicrographs were taken to examine and measure virion size and structure.

The mean capsid diameter for both cases was determined from 60 virus particles using ImageJ2

software (Schindelin et al. 2015).

Genomic Characterization and Phylogenetic Analysis

For both the 2015 and 2018 isolates (clownfish picornavirus-2015 {CFPV-2015} and

CFPV-2018, respectively), second passage cell lysate from infected flasks of SSN-1 cells was

centrifuged at 3000 x g for 10 minutes at 4°C in a Beckman-Coulter Allegra X-14R centrifuge to

spin down cellular debris. The clarified supernatant was then collected and recentrifuged in a

Beckman JA-14 fixed angle rotor at 100,000 x g for 90 minutes at 4°C to pellet the virus. The

supernatant was pipetted off and replaced with resuspension buffer (10 mM Tris-HCl, pH 7.6, 10

mM KCl, 1.5 mM MgCl2) to resuspend the pelleted virus. Baseline-Zero DNase (Lucigen,

Middleton, WI, USA) was added to digest extraneous DNA into mononucleotides. The viral

RNA (vRNA) was then purified using a RNeasy Mini Kit (Qiagen). The purified vRNA served

as a template to generate a cDNA library using a NEBNext Ultra RNA Library Prep Kit (New

England Biolabs® Inc.), which was then sequenced on an Illumina MiSeq sequencer using a 600-

cycle v3 MiSeq Reagent Kit.

Paired-end sequence reads were then trimmed and de novo assembled using CLC

Genomics Workbench v11.0 for both the 2015 and 2018 isolates (CFPV-2015 and CFPV-2018).

The BLASTX search tool was used to screen the resulting contigs against a proprietary viral

database, built in CLC Genomics Workbench v11.0 from virus protein sequences retrieved from

the UniProt Knowledgebase (https://www.uniprot.org/uniprot/). For CFPV-2015, the 5’ end of

the genome was determined by using a 5’ Rapid Amplification for cDNA End (RACE) PCR Kit

(Roche Diagnostics, Mannheim, Germany) and Sanger sequencing. This was not performed on

CFPV-2018. The cleavage sites of the CFPV-2015 and CFPV-2018 polyproteins were predicted

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by sequence alignment comparisons to the polyproteins of other fish picornaviruses within the

genus Limnipivirus (Table 3-1) using Geneious R10 (Kearse et al. 2012). The same approach

was implemented in order to predict the polyprotein cleavage sites of three other closely related

fish picornaviruses (i.e., Wenling bighead beaked sandfish picornavirus, Guangdong spotted

longbarbel catfish picornavirus, and West African lungfish picornavirus) that had not been

previously annotated (Shi et al. 2018).

For phylogenetic analysis, the amino acid (aa) sequences of the 3Dpol of both the CFPV-

2015 and CFPV-2018 isolates were aligned with those of 66 other picornavirus sequences (Table

3-1) using the Multiple Alignment using Fast Fourier Transform (MAFFT) 7.0 server

(https://mafft.cbrc.jp/alignment/software/) with default parameters. A Maximum Likelihood tree

was generated using 1000 bootstrap replicates in IQ-TREE (Nguyen et al. 2015) with default

parameters. Pairwise genetic comparisons of the aa sequences of the P1, 2C, 3C, and 3D regions

of the CFPV-2015 polyprotein were each compared to six other closely related fish

picornaviruses (Table 3-1) using the Sequence Demarcation Tool v1.2 (Muhire et al. 2014), with

the MAFFT alignment option implemented. Additionally, the full polyproteins of these six fish

picornaviruses were aligned and compared with the full polyprotein of the CFPV-2015 using

Geneious R10.

RNA Extraction and Development of a CFPV RT-PCR Assay

Frozen internal and external tissue homogenates generated from the 2015 and 2018 case

material for virus isolation (described above) were subjected to RNA extraction using a RNeasy

Mini Kit following the manufacturer’s instruction (Qiagen). One-step conventional RT-PCR was

performed on the extracted RNA samples using a QIAGEN OneStep RT-PCR Kit and primers

targeting the RNA-dependent RNA polymerase (3Dpol) region of the CFPV-2015 polyprotein

(Table 3-2). These primers, CFPV-F (5’-6130CAGAGAAGAGCACACCCTGG6149-3’) and

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CFPV-R (5’-6385GCTGGTGCTTTGGTCAACTG6366-3’), were used as follows: after the initial

reverse transcription step at 50°C for 30 minutes and the denaturation step at 95°C for 30

minutes, 40 amplification cycles of 94°C for 30 seconds (denaturation), 58°C for 30 seconds

(annealing), and 72°C for 30 seconds (elongation) were carried out, followed by a final

elongation step at 72°C for 5 minutes. Reaction volumes were 30 μL and consisted of 8.4 μL of

molecular grade water, 6 μL of 5X RT-PCR buffer, 6 μL of 5X Q solution, 1.2 μL of 10 mM

dNTPs, 1.2 μL each of 20 μM forward and reverse primers, 1.2 μL of RT-PCR enzyme mix, and

4.8 μL of RNA template. Following 1% agarose gel electrophoresis, bands of the expected size

(256 bp amplicon including primers) were purified using a QIAGEN QIAquick Gel Extraction

Kit and submitted to Eurofins Genomics (USA) to be confirmed by Sanger sequencing.

Testing Archived Clownfish Tissue Samples by RT-PCR

The WAVDL received clownfish specimens from two different facilities experiencing

similar disease episodes in their cultured A. ocellaris in the years 2011, 2012, 2104, 2015, 2017,

and 2018. The CFPV conventional RT-PCR assay described above was used to screen archived

RNA extracts from gill of a representative A. ocellaris from each disease episode from 2011-

2018. A gill RNA extract was also tested in 2019 from an A. ocellaris as part of a healthy

appearing aquacultured stock to serve as a negative control.

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Table 3-1. Sequences used for phylogenetic analysis. GenBank accession numbers for the 66

picornavirus species used in the phylogenetic analysis with the two CFPV isolates.

These 66 species come from at least 35 different genera of picornaviruses. Sequences

annotated with an asterisk (*) indicate those used to generate the sequence identity

matrices (Fig. 4-4) and the genome alignment (Fig. 4-6).

Sequence Genus Species GenBank

1 Ampivirus Ampivirus A KP770140

2 Aphthovirus Foot-and-mouth-disease virus AY593829

3 Aquamavirus Seal picornavirus EU142040

4 Avihepatovirus Duck hepatitis A virus DQ226541

5 Avisivirus Turkey avisivirus KC614703

6 Cardiovirus Encephalomyocarditis virus 1 M81861

7 Cosavirus Cosavirus A1 FJ438902

8 Dicipivirus Canine picodicistrovirus JN819202

9 Enterovirus Coxsackievirus A2 AY421760

10 Erbovirus Equine rhinitis B virus X96871

11 Gallivirus Gallivirus A JQ691613

12 Harkavirus Falcovirus KP230449

13 Hepatovirus Hepatitis A M14707

14 Hunnivirus Bovine hungarovirus 1 JQ941880

15 Kobuvirus Aichi virus A AB040749

16 Kunsagivirus Kunsagivirus A KC935379

17 Limnipivirus Bluegill picornavirus JX134222

18 Limnipivirus Carp picornavirus KF306267

19 Limnipivirus Fathead minnow picornavirus KF183915

20 Megrivirus Duck megrivirus KC663628

21 Mischivirus Minopterus schreibersii picornavirus 1 JQ814851

22 Mosavirus Mosavirus A1 JF973687

23 Oscivirus Turdivirus 2 GU182408

24 Parechovirus Human parechovirus S45208

25 Pasivirus Swine pasivirus JQ316470

26 Passerivirus Turdivirus 1 GU182406

27 Potamipivirus Eel picornavirus KC843627

28 Rabovirus Rabovirus A KP233897

29 Rosavirus Rosavirus A1 JF973686

30 Sakobuvirus Feline sakobuvirus KF387721

31 Salivirus Salivirus A GQ179640

32 Sapelovirus Avian sapelovirus AY563023

33 Senecavirus Seneca Valley virus DQ641257

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Table 3-1 Continued.

Sequence Genus Species GenBank

34 Sicinivirus Sicinivirus KF741227

35 Teschovirus Porcine teschovirus 1 AJ011380

36 Torchivirus Tortoise picornavirus KM873611

37 Tremovirus Avian encephalomyelitis virus AJ225173

38 Unassigned Threespine stickleback picornavirus MK189163

39 Unassigned Zebrafish picornavirus MH368041

40 Unassigned Beihai conger picornavirus MG600065

41 Unassigned Beihai pentapodus picornavirus MG600071

42 Unassigned Beihai wrasse picornavirus MG600073

43 Unassigned Guandong spotted longbarbel catfish picornavirus MG600094

44 Unassigned Wenling banjofish picornavirus 1 MG600070

45 Unassigned Wenling banjofish picornavirus 2 MG600072

46 Unassigned Wenling bighead beaked sandfish picornavirus MG600092

47 Unassigned Wenling brown-lined puffer picornavirus MG600100

48 Unassigned Wenling chelidoperca picornavirus MG600074

49 Unassigned Wenling crossorhombus picornavirus MG600095

50 Unassigned Wenling fish picornavirus 1 MG600078

51 Unassigned Wenling hoplichthys picornavirus MG600101

52 Unassigned Wenling jack mackerels picornavirus MG600075

53 Unassigned Wenling lepidotrigla picornavirus MG600079

54 Unassigned Wenling pleuronectiformes picornavirus MG600098

55 Unassigned Wenling rattails picornavirus MG600077

56 Unassigned Wenling scaldfish picornavirus 1 MG600096

57 Unassigned Wenling scaldfish picornavirus 2 MG600097

58 Unassigned Wenling sharpspine skate picornavirus MG600093

59 Unassigned Wenling thamnaconus septentrionalis picornavirus MG600080

60 Unassigned Wenling triplecross lizardfish pirocnavirus MG600076

61 Unassigned Western African lungfish picornavirus MG600102

62 Unassigned Wuhan carp picornavirus MG600066

63 Unassigned Wuhan sharpbelly picornavirus 1 MG600067

64 Unassigned Wuhan sharpbelly picornavirus 2 MG600068

65 Unassigned Wuhan sharpbelly picornavirus 3 MG600069

66 Unassigned Yancheng osbecks grenadier anchovy picornavirus MG600099

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Table 3-2. CFPV conventional RT-PCR primer set. Primers designed against the CFPV RNA-

dependent RNA polymerase (3Dpol) gene for use in the conventional RT-PCR

diagnostic assay. The CFPV 3Dpol gene occurs at nucleotide positions 6,047-7,513

within the genome, and the amplicon from this primer set occurs at positions 6,130-

6,385.

Primer Name Primer Sequence Tm (°C)

Amplicon size

including primers (nt)

CFPV-F CAGAGAAGAGCACACCCTGG 64.5 256

CFPV-R GCTGGTGCTTTGGTCAACTG 62.4

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CHAPTER 4

RESULTS

Parasitology, Bacteriology, and Histopathology

In both the 2015 and 2018 cases, bacteriological and parasitological examinations did not

yield significant bacterial or parasitic burdens. However, histopathologic lesions potentially

consistent with a viral etiology were evident in four examined fish from the 2015 outbreak (Fig.

4-1). Such findings included minimal to moderate individual cell necrosis and mononuclear cell

inflammation in mucosal epithelia of the branchial cavity, pharynx, esophagus, and/or stomach.

Mucosal epithelial necrosis was characterized by nuclear fragmentation (karyorrhexis),

accompanied by occasional cell loss. In one fish, necrosis of gastric glands was associated with

accumulations of exfoliated cells in the proximal intestine. In another of the four clownfish,

several round to oval basophilic inclusions (10-15 μm diameter) were evident within the mildly

hyperplastic pharyngeal mucosal epithelium, in which low to moderate numbers of lymphocytes

and infrequent necrotic cells were also present. Due to their large size, the precise subcellular

location of these inclusions (i.e., nuclear vs. cytoplasmic) was difficult to determine in H&E

sections. A histopathologic examination of samples from the case material in the 2018 case did

not reveal significant microscopic lesions.

Virus Isolation

Cytopathic effects (CPE) were observed on the SSN-1 cell line within three days post-

inoculation of the 2015 and 2018 pooled external and internal tissue homogenates. Cellular

changes included enlargement and refractility of cells, and the development of round plaques that

eventually coalesced to result in complete destruction of the cellular monolayer by day six (Fig.

4-2). The clarified SSN-1 supernatants in both years were passaged onto fresh SSN-1 cells and

again resulted in complete destruction of the monolayers. No CPE was observed in the EPC or

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GF cell lines over the course of the incubation period (14 days), nor in the subsequent passage

onto confluent monolayers of EPC and GF cells in either year.

Transmission Electron Microscopy

Analysis of the infected SSN-1 cells, from 2015 case, using transmission electron

microscopy revealed non-enveloped, round to icosahedral viral particles within cell cytoplasm

(Fig. 4-3). Similar ultrastructural findings were noted in the 2018 case. The mean diameter and

standard deviation (SD) of virus particles from 2015 and 2018 case material were 20.0 nm (n =

60, SD = 1.05 nm) and 21.7 nm (n = 60, SD = 2.64 nm), respectively. Virus particles were

observed individually and as part of paracrystalline arrays in both cases.

Genomic Characterization and Phylogenetic Analysis

The complete CFPV genome sequence from the 2015 isolate was determined to be 8,166

bp and predicted to have a 3-4-4 genome layout: 5′UTR-P1(1AB-1C-1D)-P2(2A1-2A2-2B-2C)-

P3(3A-3B-3C-3D)-3′UTR (Fig. 4-4). The 5’ and 3’ UTRs of the CFPV-2015 were determined to

be 571 and 423 bp, respectively. BLASTN analysis of the 5’ UTR of the CFPV 2015 showed no

homology to other limnipiviruses. A single open reading frame encoding a putative multi-

functional polyprotein of 2,314 aa was identified in CFPV-2015 and CFPV-2018 (Table 4-1).

The P1 region of both isolates was determined to be 671 aa long and encoded structural proteins

(i.e., capsid proteins). Similarly, the P2 and P3 regions were 835 and 808 aa long, respectively

and encoded non-structural proteins. Like limnipiviruses, the CFPV-2015 and CFPV-2018

isolates did not possess a leader peptide and the 2C region included a Walker A GxxGxGKS

(GKPGQGKT; aa 1308-1315) motif. In both isolates, the putative 3C protease region included a

GxCGx10-15GxH (GYCGSLILQKQYGTWKIVAMH; aa 1784-1804) motif. The 3D polymerase

of the CFPV-2015 and CFPV-2018 isolates included the following conserved motifs: KDE (aa

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1994-1996), DxxxxD (DYSKFD; aa 2070-2075), PSG (aa 2126-2128), YGDD (aa 2171-2174),

and FLKR (aa 2219-2222).

The Maximum Likelihood analysis based on the 3Dpol aa sequence alignment among

different picornavirus genera yielded a well-supported and highly resolved tree, with bootstrap

replicate values of 100% for most nodes (Fig. 4-5). The CFPV-2015 and CFPV-2018 were

supported as each other’s closest relative and together they branched as the sister species to the

bluegill picornavirus (BGPV), with bootstrap support of 100%. The CFPV and BGPV clade was

supported as the sister group to a well-supported clade composed of the other two limnipiviruses

(i.e., fathead minnow picornavirus and the carp picornavirus) as well as an unclassified Wenling

bighead beaked sandfish picornavirus. Two other unclassified picornaviruses, the spotted

longbarbel catfish and the West African lungfish picornaviruses, formed well supported basal

branches to the aforementioned limnipivirus clades. The P1, 2C, 3C, and 3D regions of the

CFPV-2015 showed 99%, 99.7%, 99.5%, and 99.5% aa identity to CFPV-2018, respectively.

The P1 region of the CFPV-2015 isolate exhibited greatest (70%) aa identity to that of the

BGPV, while its 2C, 3C, and 3D displayed 54%, 49% and 61% identity to the BGPV,

respectively (Fig. 4-4). Comparison of the full CFPV-2015 polyprotein to the polyproteins of six

other fish picornaviruses revealed similar cleavage products (Fig. 4-6).

Development of a CFPV RT-PCR Assay

The CFPV RT-PCR assay yielded positive results for all of the pooled tissue homogenate

samples generated in the 2015 and 2018 cases. Sanger sequencing of the purified PCR products

resulted in identical sequences to the corresponding CFPV-2015 and CFPV-2018 sequences

generated by the Illumina MiSeq sequencer. The archived samples from diseased A. ocellaris in

the years 2011-2018 all yielded the expected amplicons and yielded sequences with >99%

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nucleotide identity to the 2015 and 2018 sequences. The gill tissue sample collected from a

healthy appearing aquacultured A. ocellaris in 2019 was negative.

Table 4-1. Predicted genome organization of the clownfish picornavirus. Predicted cleavage sites

for genes within the clownfish picornavirus (2015) [CFPV-2015] polyprotein, based

on a MAFFT-alignment with four previously annotated fish picornaviruses showing

cleavage sites. Clownfish picornavirus (2018) [CFPV-2018] possesses identical

cleavage sites as CFPV-2015. The Asparagine (N) near the CFPV-2015 3B/3C

cleavage site (highlighted in yellow) has been replaced by Serine (S) in CFPV-2018.

Nucleotide

Sequence

Amino Acid

Sequence

Predicted Downstream

Cleavage Site

Gene Start End Size Start End Size

5' NTR 1 571 571 - - - -

1AB 572 1300 729 1 243 243 AVLE / GNGN

1C 1301 1990 690 244 473 230 VKFQ / GPGQ

1D 1991 2584 594 474 671 198 YFLQ / SPPS

P1 572 2584 2013 1 671 671

2A1 2585 2932 348 672 787 116 SNPG / PAIF

2A2 2933 3355 423 788 928 141 ENPG / PTFK

2B 3356 4084 729 929 1171 243 PTQQ / GQKE

2C 4085 5089 1005 1172 1506 335 ATFQ / GGPG

P2 2585 5089 2505 672 1506 835

3A 5090 5374 285 1507 1601 95 PEEQ / RAYN

3B 5375 5452 78 1602 1627 26 VEPQ / GGNK

3C 5453 6046 594 1628 1825 198 PQQQ / GVVE

3D 6047 7513 1467 1826 2314 489 ICDD/

P3 5090 7513* 2424 1507 2314 808 -

3' NTR 7517 8166 650 - - - -

*7514-7516: Stop codon (TAG)

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Figure 4-1. Histologic sections of branchial cavity and alimentary tracts of a clownfish sampled

from a CFPV-positive population. A) The branchial cavity mucosa is disrupted and

vacuolated, and necrosis is evidenced by the presence of karyorrhectic nuclear debris

(black arrows) and phagocytized fragments of cellular debris (white arrows). B)

Nuclear debris of necrotic cells (arrows) can be seen frequently in the glandular

stomach mucosa. C) Abundant exfoliated cells in the lumen of the proximal intestine

of a clownfish in which gastric gland necrosis was observed. D) Large basophilic

inclusions (black arrows) in epithelial cells of the pharyngeal mucosa, accompanied

by occasional necrotic cells (white arrows). All images, bar = 25 m.

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Figure 4-2. In vitro growth characteristics of the CFPV-2015 isolate in SSN-1 cells. A) Healthy, uninfected striped snakehead (SSN-1)

cells, day 8 post-inoculation. B) SSN-1 cells infected with the clownfish picornavirus (2015), day 8 post-inoculation. Bar =

50 m.

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Figure 4-3. Ultrastructural features of the CFPV-2015 isolate in SSN-1 cells. A) Transmission electron microscopy of the clownfish

picornavirus (2015) showing non-enveloped icosahedral virions with an average diameter of 20 nm within the cytoplasm of

an infected SSN-1 cell. Arrows to insets provide higher magnification of the virus particles. B) Clownfish picornavirus-

2015 particles arranged in a paracrystalline array (arrowhead) within the cytoplasm of an infected SSN-1 cell.

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Figure 4-4. Annotated CFPV polyprotein with sequence identity matrices for each of the P1, 2C, 3C & 3D regions. Annotated

clownfish picornavirus (2015) polyprotein showing predicted cleavage sites and functional domains (orange), alongside a

map of the CFPV genome showing read coverage from the MiSeq data. Different shades of blue from top to bottom show

the maximum, average, and minimum coverage values as calculated using a window size of 1 bp. Sequence identity

matrices are shown below for P1, 2C, 3C, and 3D regions of the clownfish polyprotein (2015) compared with those of the

bluegill picornavirus (BGPV), fathead minnow picornavirus (FHMPV), carp picornavirus (CPV), sandfish picornavirus,

catfish picornavirus, and lungfish picornavirus.

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Figure 4-5. Phylogenetic analysis of picornavirus 3Dpol gene sequences. Maximum Likelihood

phylogram based on the amino acid sequence of the complete 3Dpol gene of 68

picornavirus species. Taxa shown in blue are fish picornaviruses forming a clade with

the two clownfish picornavirus isolates, while those shown in black represent the type

species from all other known picornavirus genera, as well as all other known fish

picornavirus species. All nodes are supported by bootstrap values >80% (with the

exception of those with nodes marked with “•”). Inferred substitutions are

represented by the lengths of each branch as indicated by the scale bar.

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Figure 4-6. CFPV alignment with six other fish picornaviruses. Seven MAFFT-aligned fish picornavirus polyproteins comparing the

bluegill, fathead minnow, carp, sandfish, catfish, lungfish, and clownfish picornavirus (2015) sequences. A sequence

identity graph is shown above the alignment, illustrating residue identity among sequences across all positions. Green

represents complete amino acid identity for a given position, yellow represents less than total similarity, and red represents

residues with very low similarity.

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CHAPTER 5

DISCUSSION

In this investigation, we characterized a novel clownfish picornavirus (CFPV) isolated

from diseased Amphiprion ocellaris, the first report of a picornavirus isolated from a

maricultured ornamental fish. Growth of CFPV in SSN-1 cells from both the 2015 and 2018 case

material facilitated its downstream ultrastructural and genomic characterization. The CFPV

virion architecture and development were congruous with typical picornavirus virion

morphogenesis including the observation of small, non-enveloped, round to icosahedral virus

particles within the cytoplasm of the infected SSN-1 cells. The size, shape and location of the

CFPV nucleocapsids are consistent with reports of other picornaviruses, including those

previously isolated from fish (Barbknecht et al. 2014, Lange et al. 2014, Phelps et al. 2014,

Fichtner et al. 2013, Hahn and Dheilly 2019, Altan et al. 2019). The genomic and phylogenetic

analyses strongly supported this CFPV as a novel species and closest relative to the BGPV

within the fish picornavirus genus Limnipivirus. Finally, a specific conventional RT-PCR assay

was designed as a rapid screening tool to test A. ocellaris tissues for the presence of the CFPV.

The RT-PCR assay detected the CFPV in archived samples from moribund A. ocellaris dating

back as far as 2011.

The CFPV-2015 genome was determined to be 7939 nucleotides in length before the

poly-A tract, while the genomes of the European eel, bluegill, common carp, fathead minnow,

zebrafish, and three-spine stickleback picornaviruses are 7632, 7834, 8050, 8298, 8404 and 7496

nucleotides, respectively (Barbknecht et al. 2014, Lange et al. 2014, Phelps et al. 2014, Fichtner

et al. 2013, Hahn and Dheilly 2019, Altan et al. 2019). The CFPV 5’ UTR (571 bp) and 3’ UTR

(423 bp) are longer than other limnipiviruses (501-712 bp and 38-342 bp, respectively). In

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addition, the serine (S) in the Walker A GxxGxGKS motif of the 2C helicase is replaced by a

threonine (T) as in all limnipiviruses.

Genetic analysis of regions of the CFPV polyprotein revealed it shares closest amino acid

identity to the BGPV, a limnipivirus. Similarly, the phylogenetic analysis based on the RNA-

dependent RNA polymerase (3Dpol) gene yielded a tree in which the CFPV grouped within the

genus Limnipivirus, as the closest relative to the BGPV. The CFPV and BGPV clade was

supported as the sister group to the other limnipiviruses, the fathead minnow picornavirus

(FHMPV) and the carp picornavirus. Unclassified fish picornaviruses, the Guangdong spotted

longbarbel catfish picornavirus and the West African lungfish picornavirus, formed well

supported branches basal to the currently accepted limnipiviruses. According to the current

International Committee on Taxonomy of Viruses (ICTV) guidelines, criteria used for the

picornavirus genus and species demarcations are based on the genetic distances between P1, 2C,

3C, and 3D. Picornaviruses with aa sequence divergence exceeding 66% for P1 and 64% for 2C,

3C, and 3D are considered members of different genera (Zell et al. 2017). Within the genus

Limnipivirus, those with aa sequence divergence ranges of 30-43% for P1 and 49-57% for 3C

and 3D are considered different species (Zell et al. 2017). The observed aa sequence divergences

of the CFPV P1, 2C, 3C, and 3D proteins to accepted limnipiviruses (i.e., BGPV; carp

picornavirus, CPV-1, and fathead minnow picornavirus, FHMPV) ranged from 30-35%, 46-58%,

51-65%, and 39-51%, respectively, supporting its inclusion as a new species in the genus (Fig. 4-

4).

Additionally, the aa sequence divergences of the Wenling bighead beaked sandfish

picornavirus P1, 2C, 3C, and 3D compared to the other known limnipiviruses ranged between

18-39%, 37-52%, 38-64%, and 33-49%, respectively, while those of the Guangdong spotted

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longbarbel catfish picornavirus ranged from 50-65%, 63-71%, 66-70%, and 51-55%, and those

of the West African lungfish picornavirus ranged from 65-67%, 65-70-%, 67-72%, and 61-62%.

From these genetic distances, we posit that the Wenling bighead beaked sandfish picornavirus

should also be included as a new species within the Limnipivirus genus, while both the

Guangdong spotted longbarbel catfish picornavirus and the West African lungfish picornavirus

represent novel species within yet to be defined genera. Consideration of the clownfish

picornavirus and the Wenling bighead beaked sandfish picornavirus as new limnipivirus species

will require formal proposal to and ratification by the ICTV.

To date, most fish picornaviruses have been detected in wild fish including European eel,

fathead and brassy Hybognathus hankinsoni minnows, bluegill, threespine stickleback, and 29

other freshwater and marine fishes (Barbknecht et al. 2014, Phelps et al. 2014, Fichtner et al.

2013, Hahn and Dheilly 2019, Geoghegan et al. 2018, Shi et al. 2018). In contrast,

picornaviruses from carp and zebrafish, along with fathead minnows, have been characterized

from aquacultured or laboratory managed stocks (Lange et al. 2014, Phelps et al. 2014, Altan et

al. 2019). The CFPV represents the first picornavirus characterized from an important

maricultured ornamental species, the clownfish A. ocellaris.

The majority of fish picornaviruses have not been grown in in vitro, and thus, the role of

many of these viruses (if any) in disease remains to be determined (Hahn and Dheilly 2019,

Altan et al. 2019, Geoghegan et al. 2018, Shi et al. 2018. While the clownfish picornavirus was

isolated from both internal (kidney, liver, spleen, heart) and external (gill and skin) tissue pools,

other studies isolated fish picornaviruses from internal tissues (primarily kidney and spleen, as

well as liver, heart and brain) (Barbknecht et al. 2014, Lange et al. 2014, Phelps et al. 2014,

Fichtner et al. 2013). Isolation of fish picornaviruses have typically involved cell lines derived

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from the same host or a closely related host. For example, the carp and fathead minnow

picornaviruses were isolated on cell lines derived from cyprinids (i.e., FHM and EPC), the eel

picornavirus was isolated on the eel embryonic kidney (EK-1) cell line, and the bluegill

picornavirus was isolated on the bluegill fry (BF-2). In contrast, the CFPV grew on a cell line

(SSN-1) derived from a freshwater fish (striped snakehead; Channa striata). The CFPV did not

grow on the EPC cell line or the grunt fin cell line derived from a marine fish (blue-striped grunt;

Haemulon sciurus). Although the bluegill, carp, clownfish, and eel picornaviruses were initially

isolated from moribund wild fish (Barbknecht et al. 2014, Lange et al. 2014, Fichtner et al.

2013), the fathead minnow picornavirus was primarily isolated from seemingly healthy fathead

and brassy minnows from the wild or sold through baitfish wholesalers (Phelps et al. 2014).

The role of picornaviruses as disease-causing agents of fish has only recently received

significant attention. Picornaviruses isolated from wild European eels and bluegill were capable

of inducing disease under controlled laboratory conditions (Barbknecht et al. 2014, Fichtner et al.

2013). However, another recent study failed to reproduce disease in aquacultured common carp

(Lange et al. 2014), while experimental challenges were not performed in studies involving

picornaviruses characterized from wild fathead minnows (Phelps et al. 2014), threespine

stickleback (Hahn and Dheilly 2019), or managed zebrafish (Altan et al. 2019). Recent studies

employing metagenomics detected numerous picornavirus sequences across a range of fishes

irrespective of their health status (Geoghegan et al. 2018, Shi et al. 2018). Although the role of

the CFPV in disease remains undetermined, we confirmed CFPV RNA by RT-PCR in archived

clownfish tissue extracts (A. ocellaris and A. percula) from mass mortality events that had

occurred at two U.S. clownfish production facilities between the years 2011 and 2018, but not

from a single fish collected in 2019 from a population of apparently healthy fish.

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The role of the CFPV as a causative agent of disease was not clearly established in the

present study. Interestingly, the microscopic lesions observed in the 2015 case were not observed

in the 2018 case. The lack of histopathologic findings in the 2018 fish may be attributed to the

sampling of unaffected or mildly affected animals, or fish that had already begun to recover. For

certain viral infections, lesions may only be evident within a narrow window of time during the

natural course of the disease. It is also possible that the branchial and gastrointestinal lesions

observed in the 2015 case were not the result of the CFPV infection. Future challenge studies are

needed to fulfill River’s postulates (Rivers 1937) in order to confirm that CFPV is a true cause of

morbidity and mortality in clownfish, as well as to better understand the pathology and mode of

transmission of the virus and to provide insight into the historical morbidity and mortality events

at these facilities (Williams 2010).

Should future challenge studies confirm the pathogenic nature of the CFPV, these

findings would assist in defining clinical signs as well as gross and microscopic lesions

associated with the disease. The CFPV genomic sequence generated in the present study can be

used to develop diagnostic tools for surveillance and to better define the pathogenesis. A CFPV

specific in situ hybridization (ISH) assay would complement the histopathological examination

of tissues from CFPV-infected A. ocellaris. The ISH guided determination of tissues associated

with significant microscopic lesions would not only assist in confirming the role of the CFPV in

disease but would also help to determine the tissue tropism of the CFPV as well as the most

appropriate tissues for diagnosis by virus isolation or molecular assay (e.g., reverse transcription

conventional or quantitative PCR assays). The future design of a CFPV reverse transcription

quantitative PCR (RT-qPCR) assay could be used to assess the viral load in tissues of infected

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clownfish, as well as serving as a rapid and sensitive diagnostic tool for screening clownfish

populations.

The discovery of the CFPV may have significant implications for the marine ornamental

industry, as clownfish epizootics over the past decade have led to production lapses and

significant economic losses for some facilities in the U.S. (RPE Yanong pers. obs. 2018). If the

CFPV is determined to be the cause of these aquaculture epizootics and spreads, the

development of effective management strategies would be needed to mitigate the disease and

help prevent over-collection of wild fish from coral reefs. In addition to financial consequences

for clownfish culture facilities, wild collection of clownfish can be a very ecologically damaging

process, especially in the case of cyanide fishing (Dee et al. 2014). Given the potential damage

caused by the CFPV, as well as the increasing number of picornaviruses that have been isolated

from wild fish, it seems important to ascertain whether its significance extends beyond

aquaculture. Therefore, future studies should investigate the prevalence and disease potential of

the CFPV to wild populations of A. ocellaris.

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BIOGRAPHICAL SKETCH

Elizabeth Scherbatskoy is from Saratoga Springs, NY. After obtaining her Bachelor of

Science from Eckerd College (St. Petersburg, FL) in Biology and Environmental Science,

Elizabeth worked as a veterinary assistant at a small animal practice before moving to

Gainesville, FL to pursue her Master of Science degree in Veterinary Medical Sciences at the

University of Florida. Her research was conducted in the Infectious Diseases and Immunology

Department of the UF College of Veterinary Medicine, in the Wildlife and Aquatic Veterinary

Disease Laboratory. Elizabeth graduated with her MS from UF in 2020. Post-graduation,

Elizabeth hopes to attend veterinary school to pursue her DVM to further her career in both

research and clinical veterinary medicine.