CHARACTERIZATION OF A NOVEL PICORNAVIRUS ISOLATED …
Transcript of 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
24
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
25
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.
26
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
27
(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
28
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.
29
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
30
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
31
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
32
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
33
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.
34
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
35
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
36
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.
37
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
38
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
39
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
40
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
41
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
42
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%
43
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)
44
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.
45
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.
46
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.
47
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.
48
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.
49
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.
50
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
51
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
52
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
53
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.
54
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
55
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.
56
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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.