1 University of Washington Friday Harbour Laboratories, 620 University Road, Friday Harbour, WA 98250, USA
2 School of Aquatic and Fishery Sciences, University of Washington, Box 355100, Seattle, WA 98195, USA
3 Western Fisheries Research Center, 6505 NE 65th Street, Seattle, WA 98115, USA
Correspondence
James Winton
jim_winton{at}usgs.gov
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ABSTRACT |
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The GenBank accession number of the sequence reported in this paper is AY450644.
Present address: Department of Microbiology, Boston University, 715 Albany Street, Boston, MA 02118, USA.
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INTRODUCTION |
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Salmonids have been shown to harbour these viruses, especially in freshwater hatcheries or in marine net pens where the viruses can spread rapidly, sometimes resulting in high mortality. However, little is known about the host range of these viruses among various species of marine fish. In addition to serving as possible reservoirs of viruses that can be transmitted to salmonids, some of these marine fish species may themselves be highly susceptible to infection (Kocan et al., 1997), leading to natural outbreaks in the marine environment (Meyers & Winton, 1995
; Meyers et al., 1992
, 1994
, 1999
; Hershberger et al., 1999
; Takano et al., 2000
; Kocan et al., 2001
; Hedrick et al., 2003
). For these reasons it is important to sample a range of marine fish species to determine the prevalence of these viruses in the wild, and to assess the susceptibility of various marine fish species to these infectious agents.
In late March of 2000, a survey of marine fish captured in the San Juan Archipelago of northern Puget Sound, WA, USA, was conducted to determine the natural prevalence of these or other viruses in a variety of marine fish species, including English sole (Parophrys vetulus) and starry flounder (Platichthys stellatus). During the course of the survey, a new rhabdovirus of marine fish was isolated. Detection of this virus, tentatively termed starry flounder rhabdovirus (SFRV), during a limited survey, highlights the void of knowledge regarding viruses that naturally infect marine fish species in the North Pacific Ocean.
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METHODS |
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Cell culture.
Epithelioma papulosum cyprini (EPC; Fijan et al., 1983) cells were grown in tissue culture flasks using Eagle's minimum essential medium (MEM) supplemented with 10 % fetal bovine serum (FBS) and adjusted to pH 7·4 by the addition of 7·5 % sodium bicarbonate (MEM-10-SB). Cells were grown at 25 °C for the first week and then incubated at 15 °C until used.
Virus isolation.
Kidney and spleen tissues from each fish were removed and diluted 1 : 4 in MEM containing 140 mM Tris and supplemented with 100 IU penicillin, 100 µg streptomycin, 100 µg gentamicin sulfate and 2·5 µg amphotericin B ml-1 (MEM-AF). Excised tissue samples were stored at -80 °C until virus assays were performed. Samples were thawed, homogenized by mortar and pestle and clarified by low-speed centrifugation for 35 min. Serial 10-fold dilutions of the supernatant were made in MEM containing 140 mM Tris and supplemented with 5 % FBS (MEM-5-T), and the dilutions were inoculated onto EPC monolayers in 24-well plates. After 30 min incubation at room temperature to allow virus adsorption, 1 ml of an overlay composed of 0·75 % methylcellulose in MEM-5-T was added to each well and the plates were incubated at 15 °C. Plates were observed regularly for cytopathic effect (CPE) for 7 days, then fixed and stained with a crystal violet and formalin solution. The initial virus titre in the tissues was reported as plaque-forming units (p.f.u.) (g tissue)-1.
Stock viruses.
Medium from cell cultures showing CPE was stored in aliquots at -80 °C for use as stock virus. In addition, reference strains of IHNV, VHSV, Hirame rhabdovirus (HIRRV), Spring viremia of carp virus (SVCV), IPNV and an aquareovirus isolated from adult chinook salmon (Oncorhynchus tshawytscha) in the Green River of Washington, USA, were used as controls in various assays.
Plaque assays.
Treated tissue culture 24-well plates (Costar) were seeded with EPC cells in MEM-5-T and incubated at 25 °C for 24 h. Serial 10-fold dilutions were prepared in MEM-5-T and inoculated into wells of the 24-well plates from which the medium had been drained. After 30 min incubation at room temperature to allow virus adsorption, 1 ml of an overlay composed of 0·75 % methylcellulose in MEM-5-T was added to each well. The cultures were incubated at 15 °C for 7 days, then fixed and stained with a crystal violet and formalin solution. Virus titre was reported as p.f.u. (ml culture fluid)-1.
Growth temperature.
Medium was decanted from twelve 25 cm2 flasks containing monolayers of EPC cells and each was inoculated with 0·25 ml stock virus, incubated at room temperature for 30 min and then rinsed three times with MEM to remove unattached virus. MEM-10-SB (5 ml) was added to each flask and sets of triplicate flasks were incubated at 10, 15, 20 or 25 °C. At 1, 2, 3, 4, 5 and 6 days post-inoculation, a 0·2 ml aliquot of culture fluid was removed from each flask, pooled by temperature and the virus titre was determined by plaque assay.
Stability to freezethaw.
To determine the stability of the virus to repeated freezethaw cycles, 1·0 ml aliquots of original stock virus were held at -80, -20 and 5 °C. After 3, 7 and 14 days of storage, the frozen aliquots were thawed and the virus titre in the aliquot stored at each temperature was determined by plaque assay. The aliquots were then refrozen until the next assay period.
Chloroform sensitivity.
Presence of a lipid-containing envelope was determined by mixing 0·5 ml cell culture fluid containing virus with an equal volume of chloroform. The mixture was shaken for 10 min, then centrifuged at 200 g for 5 min to separate the aqueous phase from the chloroform. Control viruses included VHSV as an enveloped (positive) control and an aquareovirus as a non-enveloped (negative) control. Titres of infectious virus in the aqueous phases of treated and untreated preparations were determined by plaque assay.
Electron microscopy.
Monolayer cultures of EPC cells were inoculated at a relatively high m.o.i. and the cultures were incubated at 15 °C. At 2, 3 and 7 days post-inoculation, control and virus-infected cultures were fixed for 24 h using 4 % glutaraldehyde in 0·1 M cacodylate buffer (pH 7·4). Cells were dislodged into the medium that was then centrifuged at 1000 g for 10 min, and 0·1 M sodium cacodylate was added to the pellet. Samples were recentrifuged for 5 min at 1000 g and the pellet was placed in 2 ml cold 1 % OsO4 for 1·5 h. Following three 10 min rinses in 1 ml 0·1 M sodium cacodylate, 2 ml 1 % aqueous uranyl acetate was added. The uranyl acetate was removed after 1 h and the samples were dehydrated in a graded ethanol series, then embedded in Spurr's epoxy resin. Thin sections were cut using a diamond knife in an ultramicrotome and the sections were placed on grids. The grids were examined using an electron microscope and sections were photographed at various magnifications.
Staining of infected cells.
Monolayers of EPC cells were grown on sterile cover glasses in 6-well plates. Cells were infected with 0·1 ml 10-3 and 10-4 dilutions of stock virus to provide approximately 10100 p.f.u. per cover glass. After 23 days incubation at 15 °C, sets of cover glasses were transferred to Petri dishes and fixed in either MEM containing 10 % formalin or Carnoy's fixative. After storage at 5 °C overnight, the fixative was removed and replaced with MEM. Formalin-fixed cover glasses were stained with either MayGrunwald Giemsa or with haematoxylin and eosin by standard methods (Rovozzo & Burke, 1973), then mounted onto slides using Permount and stored at 25 °C to dry. The stained cover glasses were examined by light microscopy and photographed. Acridine orange staining was performed by standard methods (Rovozzo & Burke, 1973
) using cover glasses fixed in Carnoy's fixative. Cover glasses were kept moist with McIlvaine's buffer while mounted on the slide and immediately examined using a fluorescence microscope fitted with a mercury lamp and appropriate filter blocks.
Analysis of structural proteins.
Virus grown in 150 cm2 flasks of EPC cells was harvested and the culture fluid clarified by centrifugation at 4 °C for 20 min at 1000 g. The supernatant was placed into ultracentrifuge tubes containing 0·2 ml glycerol, then centrifuged at 82 700 g at 4 °C for 1 h in an SW28 rotor (Beckman). The pellet was resuspended and layered on top of a step gradient composed of 50, 35 and 20 % sucrose in 0·01 M Tris buffer (pH 7·4). The gradient was centrifuged at 82 700 g at 4 °C for 1·5 h in an SW28 rotor. The virus band was then removed with a syringe, diluted in 0·01 M Tris and the virions were pelleted by centrifugation at 115 000 g at 4 °C for 1 h in an SW 50.1 rotor (Beckman). SDS-PAGE was used to analyse the structural proteins of the new virus. The pellet of purified virus was resuspended in sample buffer, heated to 95 °C for 2 min and stored at -20 °C until used. A 10 % SDS-PAGE gel was prepared and run using standard methods. The gel was stained with Coomassie blue (Sigma). For comparison, three other fish rhabdoviruses, IHNV, SVCV and VHSV, were purified as indicated and included in some of the gels.
PCR.
RT-PCR assays were used to determine if the new virus was similar to one of several other fish rhabdoviruses for which genomic information was available. Nine sets of PCR primers for IHNV (1 set), VHSV (1 set), HIRRV (3 sets) and SVCV (4 sets) were synthesized from published sequences. Culture fluids containing the new virus or the corresponding virus controls were heated at 95 °C for 2 min to release viral RNA, then cooled on ice, a procedure used routinely in our laboratory to release nucleic acids from poikilotherm viruses grown in cell culture (Huang et al., 1996), but which is not recommended for extraction of nucleic acids from tissue samples. Our standard RT-PCR consisted of 50 pmol each primer pair in a separate single-tube (50 µl reaction) containing 5 µl viral RNA, 10 mM Tris/HCl (pH 8·3), 50 mM KCl, 2·5 mM MgCl2, 2·5 units AmpliTaq (Applied Biosystems), 4·5 units reverse transcriptase (Promega), 10 units RNasin ribonuclease inhibitor (Promega) and 200 µM dNTP. Reverse transcription and annealing temperatures were varied during this study to allow primers to bind to homologous regions of SFRV genes that may not have been identical. Four reverse transcription and annealing temperatures were tested in separate experiments: 45, 40, 35 and 30 °C. After a 30 min reverse transcription reaction and initial denaturation at 95 °C for 2 min, 35 cycles of amplification were performed using the following conditions: 95 °C for 30 s, 4530 °C (same as reverse transcription temperature) for 30 s, 72 °C for 1 min. For final extension, tubes were incubated at 72 °C for 7 min. The PCR products were then held at 4 °C until loaded for electrophoresis on a 1·5 % agarose gel and visualized by ethidium bromide staining.
Sequencing of a region of the viral polymerase gene.
Relatively conserved primer sites were identified in the rhabdovirus polymerase (L) gene by comparing published sequences of SVCV, IHNV, VHSV, Vesicular stomatitis New Jersey virus (VSNJV), Rabies virus (RABV) and Mokola virus (MOKV). Three degenerate primers, homologous to positions 15921611, 19381919 and 21572141 of the L gene of Vesicular stomatitis Indiana virus (VSIV; GenBank accession no. K02378), were synthesized and used to amplify a region of the SFRV polymerase gene in a semi-nested PCR reaction using RT and annealing temperatures of 40 and 30 °C, respectively. The first round of PCR used degenerate sense primer 5'-AARGTCAAGGCGATGGARYT-3' and antisense primer 5'-TGATTGTCCCCCTGNGC-3' in standard RT-PCR reaction mixes with SFRV RNA extracts to amplify a 577 bp product. A semi-nested PCR was conducted on the first round PCR product by using the same sense primer, but with reverse primer 5'-TCAAAAGTCTCGTGAACTCT-3' to amplify a 348 bp region. For both amplifications, DNA was analysed on a 1·5 % agarose gel and visualized by ethidium bromide staining. The visible PCR product from the second round was purified with a StrataPrep kit (Stratagene) and labelled for automated sequencing by BigDye terminator cycle sequencing (Applied Biosystems) on an Applied Biosystems 310 genetic analyser. A 243 nt region of authentic SFRV sequence was obtained, consisting of a portion of an intact ORF. This authentic SFRV sequence, when compared with other known rhabdovirus sequences, allowed the creation of primers for obtaining additional sequence information. Because the sequence was located near the 5' end of the L gene mRNA, a 5' RACE kit (Invitrogen) was used with three gene-specific primers (GSP) to gain additional sequence information. Primer GSP1 was 5'-AAAGATCAAYTCDCGGGAWG-3', GSP2 was 5'-AAYTCDCGGGAWGGTTSTG-3' and GSP3 was 5'-GAGAGMTCKAYGAATTAGA-3'. Additional primers were used to sequence toward the 5' end of the mRNA of the L protein gene. Sequence information was also obtained later by using conserved primers toward the 3' end of the L mRNA combined with known SFRV primer sequences.
Phylogenetic analysis.
To perform a phylogenetic comparison of SFRV with other fish rhabdoviruses, the polymerase gene sequence of SFRV and selected L gene sequences from GenBank were truncated to align with the 456 aa of the amino terminus of the polymerase protein. Sequences were aligned with CLUSTAL W followed by analysis with PAUP 4.0. Both neighbour-joining and parsimony analyses were performed using the homologous region of the polymerase sequence of Human parainfluenza virus 1 (HPIV-1), a member of the family Paramyxoviridae, as the outgroup. Phylogenetic trees were generated containing 1000 bootstrapped samples and values shown as percentages were transferred to the nodes of the branches.
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RESULTS |
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Growth temperature and physical stability
Using cultures of EPC cells incubated at selected temperatures, the optimal growth temperature was determined to be 15 °C (Fig. 2). The virus also grew well at 10 °C. The virus replicated poorly at 20 °C and there was no evidence of virus growth at 25 °C. Aliquots of tissue homogenate stored at 5, -20 or -80 °C showed little loss in titre over the 14-day period that included three freezethaw cycles (data not shown). No plaques appeared when the virus isolated from starry flounder was mixed with chloroform before inoculation of EPC cells, indicating the virus possessed a lipid-containing envelope. The enveloped control virus (VHSV) was also inactivated, while the non-enveloped aquareovirus, used as a negative control, showed no loss of infectious titre following treatment with chloroform.
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PCR
Control virus templates produced RT-PCR products of the expected size, while no products were obtained using the unknown virus template. Reduction of the annealing temperature to lower the stringency of the reaction did not alter the results, indicating the new virus was not an isolate of IHNV, SVCV, HIRRV or VHSV.
Sequence analysis of SFRV polymerase gene
The SFRV polymerase gene sequence obtained was 2678 nt long and had a single large ORF starting with an AUG codon in strong initiation context at nt 1113. The first 13 nt of the gene were identical between SFRV and SVCV, while 12 of the 13 nt were identical to SSTV and 903/87. The SFRV ORF encoded an 889 aa portion of the 5' end of the putative polymerase protein and this region was most homologous to that of species and tentative species of the genus Vesiculovirus, having 53 % amino acid identity to VSIV and 53 % amino acid identity to SVCV. The next closest isolates, 4546 % identical to SFRV, were Bovine ephemeral fever virus (BEFV), Flanders virus (FLAV) and the European lake trout isolate (903/87). Less related (38 % identity) was RABV, the type species of the genus Lyssavirus. Identities of 21 % or lower were obtained between SFRV and representatives of the genera Novirhabdovirus, Cytorhabdovirus and Nucleorhabdovirus, and for HPIV-1, a member of the family Paramyxoviridae.
Within the polymerase gene of members of the Mononegavirales, a consistent pattern of conserved sequence domains and subdomains has been described (Kamer & Argos, 1984; Poch et al., 1990
). These include major domains IVI and more highly conserved subdomains AD within domain III. Rhabdoviruses, in contrast to paramyxoviruses, do not contain a variable hinge region between conserved domains II and III. Our partial sequence of SFRV L protein contained the first three conserved domains: I, II and III. Pairwise alignments of the SFRV L protein in these regions with other rhabdoviruses and with HPIV-1, a paramyxovirus, showed levels of identity that generally conformed to this previously described pattern (Table 1
). Domains II and III were the most conserved of the major domains, while domain I was the least conserved, showing little identity above the overall level seen throughout the region of the L portion for which sequence was determined. Within domain III, subdomain IIIA was the most conserved, showing 100 % aa identity between SFRV and VSIV, SVCV and FLAV (Fig. 5
). One conserved stretch of 35 aa, encompassing domain IIIA, was identical between SFRV and SVCV from nucleotide position 1808 to 1912 of the putative SFRV polymerase gene. Virus isolate 903/87 from European lake trout had the highest identity to SFRV in domain I. Unfortunately, the sequence available for that virus is limited and could not be compared for the other domains.
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DISCUSSION |
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Electron microscopy and the physical characteristics of the new isolate confirmed that it was a member of the Rhabdoviridae. Only two rhabdoviruses are known to be endemic among freshwater or marine fish species on the west coast of North America, IHNV and VHSV, while several other fish rhabdoviruses have been isolated from freshwater and marine species in Europe and Asia (Table 2). Results from biological, physical, chemical and molecular assays convincingly demonstrated that the new virus was not a strain of either IHNV or VHSV. These assays also showed that SFRV was not related to three other fish rhabdoviruses known to occur in the Pacific Ocean for which limited sequence data are available: SVCV, reported from penaeid shrimp (Penaeus stylirostris and Penaeus vannamei) cultured in the Hawaiian Islands (Johnson et al., 1999
); HIRRV, reported from Japanese flounder (Paralichthys olivaceus) ayu (Plecoglossus altivelis), reared on the Japanese islands of Hokkaido and Honshu, (Kimura et al., 1986
); and snakehead rhabdovirus (SHRV) isolated from striped snakehead (Ophicephalus striatus) cultured in Southeast Asia (Ahne et al., 1988
).
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Among the vesiculovirus-like fish rhabdoviruses, growth temperature, results from the PCR assays and sequence data indicated that the SFRV was not an isolate of SVCV nor, presumably, of the closely related pike fry rhabdovirus found in European freshwater fishes (Rowley et al., 2001; Stone et al., 2003
). Also, the inability of SFRV to replicate at temperatures of 25 °C or greater indicated the virus was not similar to the ulcerative disease rhabdovirus (UDRV; Frerichs et al., 1986
), a vesiculovirus-like agent recovered from snakehead fish in Southeast Asia (Kasornchandra et al., 1992
). Similarly, the eel rhabdoviruses, EVA, EVEX, C30, B44 and D13 from Japan (Sano, 1976
; Sano et al., 1977
) or Europe (Ahne et al., 1987
; Castric et al., 1984
) have temperature optima greater than 25 °C and are serologically related to each other (Castric et al., 1984
; Hill et al., 1980
), having been grouped together under the name Rhabdovirus anguilla by Hill et al. (1980)
. Finally, the rhabdoviruses of perch (Perca fluviatilis; Dorson et al., 1984
), pike-perch (Stizostedion lucioperca; Nougayrede et al., 1992
), pike (Esox lucius; Jorgensen et al., 1993
) and lake (brown) trout (Salmo trutta lacustris; Koski et al., 1992
) in Europe do not replicate well in the EPC cell line in which the virus from starry flounder grew to high titre, have been shown to be closely related by serology (Bjorklund et al., 1994
; Dannevig et al., 2001
; Johansson et al., 2001
; Jorgensen et al., 1993
; Nougayrede et al., 1992
), and at least one of which (903/87) was genetically distinguishable from SFRV. In summary, it appears that SFRV, while related to other vesiculovirus-like isolates from fish, should be considered a novel virus. The final placement of SFRV and the other fish vesiculovirus-like viruses within the appropriate genus (or genera) of the family Rhabdoviridae will have to await further resolution.
Compared with other rhabdoviruses of fish, the virus from starry flounder had a relatively low temperature optimum. In addition to indicating the virus was unlike many of the previously described fish rhabdoviruses, the low temperature at which this agent replicates is similar to the mean temperature of Puget Sound and further suggests the agent is probably well adapted to a marine host in this region and not an introduced pathogen. Thus the starry flounder may represent one of the normal hosts for this virus in nature; however, the prevalence of the agent may be low. For example, soon after the initial report of VHSV in North America in 1989, more than 6000 samples were obtained from both marine and freshwater fish species collected from waters of Western Washington State (Winton et al., 1991). This survey included 543 marine fish of 12 species from northern Puget Sound and the Straits of Juan de Fuca. The samples were examined for viruses using standard cell culture assays and no virus of any type, including the SFRV, was isolated from any fish examined (Amos et al., 1998
).
The initial discovery of this virus in wild fish has several important implications. Many of the known fish viruses were first isolated from fish reared in aquaculture facilities and their later discovery among wild fish was considered, by some, as evidence that fish culture practices were affecting wild stocks. Furthermore, the discovery of a new virus in cultured stocks has often led to the mandated destruction of fish thought to be infected with an exotic virus that was later found to be endemic, but previously undiscovered, in the region. Should the virus from starry flounder be isolated in the future from fish cultured in the Puget Sound region, it will be recognized as an endemic agent.
Two young starry flounder were injected with virus and observed for 714 days. One fish was necropsied after 1 week and the other after 2 weeks with no outward signs of infection. Plaque assays did not show evidence of virus infection in either fish; however, fish collected from the wild may have been previously exposed to the virus with resulting immunity. Additionally, pathogen-free young rainbow trout were injected with the virus from starry flounder and observed for 12 days without visible signs of disease; however, a more in-depth study is needed using various species of marine fish.
Rhabdoviruses have a wide host range, including mammals, plants and fish. Among the viruses of fish, those causing disease in freshwater species have tended to receive the most attention, perhaps because of the relative ease of observation and collection compared with marine fish species. This has resulted in the lack of discovery of many fish rhabdoviruses and the perception, recently being re-examined, that many of the rhabdoviruses of fish are of freshwater origin. Most likely in the years to come, many more rhabdoviruses will be recovered from marine fish species.
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ACKNOWLEDGEMENTS |
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Received 25 June 2003;
accepted 12 November 2003.
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