viral haemorrhagic septicaemia
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- viral haemorrhagic septicaemia
International Common Names
- English: viral hemorrhagic septicemia
OverviewTop of page
Viral haemorrhagic septicaemia (VHS) is an infectious disease of rainbow trout (Oncorhynchus mykiss), brown trout (Salmo trutta), grayling (Thymallus thymallus), whitefish (Coregonus sp.), pike (Esox lucius), largemouth bass (Micropterus salmoides), Japanese flounder (Paralichthys olivaceus) and turbot (Scophthalmus maximus). The disease is caused by viral haemorrhagic septicaemia virus (VHSV, synonym: Egtved virus) (Benmansour et al., 1997), a virus belonging to the newly approved genus Novirhabdovirus within the family Rhabdoviridae (Walker et al., 2000).
Until the mid-1980s VHS was regarded as a disease affecting only rainbow trout and a few other freshwater fish species in aquaculture in continental Europe. In the past decades, however, VHSV has been isolated from a large range of free-living marine fish species in the North American part of the Pacific and Atlantic Oceans, the North Atlantic, the North Sea and the Baltic Sea, and in the waters around Japan. The number of susceptible wild marine species is still growing and, to date, at least 45 different species (both freshwater and marine) have been found to test positive for the virus. Susceptible fish species are found among the Salmoniformes (seven species), Esociformes (one species), Clupeiformes (four species), Gadiformes (eleven species), Pleuronectiformes (seven species), Osmeriformes (three species), Perciformes (six species), Scorpaeniformes (two species), Anguilliformes (one species), Cyprinodontiformes (one species) and Gasterosteiformes (two species).
At present, the isolates from wild marine fish are not distinguishable from normal freshwater isolates by serological means. Isolations of VHSV from fish from the North American waters and Japan, however, have so far been associated with genetically characteristic virus strains (Jensen, 1965; Einer-Jensen et al., 1995; Nishizawa et al., 2002). Four major genotypes have been identified (Snow et al., 1999; Einer-Jensen et al.,2004): Genotype I - European freshwater VHSV isolates and a group of marine isolates from the Baltic Sea, Genotype II - a group of marine isolates circulating in the Baltic Sea, Genotype III - isolates from the North Sea, Skagerrak and Kattegat and Genotype IV - North American VHSV isolates. The genetic differences appear to be more strongly related to geographical location than to year of isolation or host species (Basurco et al., 1995; Benmansour et al., 1997; Snow et al., 1999; Stone et al., 1997). However, phylogenetic anyalysis of Japanese isolates shows significant overlap of 2 genogroups, the American genogroup I and the European genogroup III (Nishizawa et al., 2002). The genetic characteristics of VHSV isolates highly pathogenic for rainbow trout have not yet been defined as to the precise gene loci responsible for virulence, but significant progress has been made in sequence analysis of virulent and avirulent strains. All examined isolates from wild marine fish have shown low or no mortality in infection trials by immersion with rainbow trout (Dixon et al., 1997; Follett et al., 1997; Winton et al., 1991). Several marine isolates are pathogenic to turbot fry (King et al., 2001a). Japanese flounder are highly susceptible to Japanese VHSV isolates (Isshiki et al., 2000). The Pacific isolates are highly pathogenic to Pacific herring (Kocan et al., 1997; Meyers et al.,1999). This newly discovered large host range and the significant differences in pathogenicity in different host species cause some problems for VHS control programmes, which are based mainly on the protection of the significant rainbow trout production industry. The main problem is whether the finding of marine VHSV in free-living fish in an approved VHSV-free area should lead to the withdrawal of that status. It has been observed in a large part of Europe that VHS in free-living fish in the marine environment affects an approved VHS-free status in only very few cases.
VHS occurs in continental Europe and is important because of its clinical and economic consequences to rainbow trout farming. For more detailed reviews of the condition, see Wolf (1988) and Jorgensen (1992). Natural infections with VHSV have caused significant mortality in turbot and Japanese flounder in aquaculture, and significant natural mortality in Pacific herring and pilchard along the Pacific coast of Alaska and Washington, United States of America (USA), and Canada.
The infection of susceptible fish species is often lethal, due to the impairment of the osmotic balance, and occurs within a clinical context of oedema and haemorrhages. Virus multiplication in endothelial cells of blood capillaries, leukocytes, haematopoietic tissues and nephron cells, underlies the clinical signs.
Three neutralising subtypes of VHSV have been recognised using a panel of polyclonal and monoclonal antibody preparations (Olesen and Jorgensen, 1992). Apart from the above variation, VHSV seems to share a VHS-group neutralising epitope, and several non-neutralising epitopes located on the viral glycoprotein (G protein). Variations in virus strain virulence have been recorded in both natural cases of disease and infection trials.
The reservoirs of VHSV are clinically infected fish and covert carriers among cultured, feral or wild fish. Virulent virus is primarily shed in the urine (Neukirch 1985), whereas kidney, spleen and heart are the sites in which virus is most abundant. Once VHSV is established in a farm stock and therefore in the water catchment system, the disease becomes enzootic due to the virus carrier fish.
Several factors influence susceptibility to VHS. Among each fish species, there is individual variability in susceptibility, and the age of the fish appears to be of some importance - the younger the fish the higher the susceptibility. In highly susceptible fish stocks, however, overt infection is seen in all sizes of fish.
Water temperature is an important environmental factor. Disease generally occurs at temperatures between 4°C and 14°C. Low water temperatures (1-5°C) generally result in an extended disease course with low daily mortality but high accumulated mortality. At high water temperatures (15-18°C), the disease generally takes a short course with a modest accumulated mortality. VHS outbreaks occur during all seasons, but are most common in spring when water temperatures are rising or fluctuating.
The screening procedure for VHS is based mainly on virus isolation in cell culture. Confirmatory testing is by immunological virus identification, e.g. neutralisation, immunofluorescence, enzyme-linked immunosorbent assay (ELISA), and immunoperoxidase staining or by reverse-transcription polymerase chain reaction (RT-PCR)-based techniques. However, more rapid diagnostic methods for presumptive evidence of viral antigen in infected organ imprints or homogenates (fluorescence, ELISA, immunohistochemistry, RT-PCR) may be suitable for fish with overt disease. Fish serology (neutralisation, ELISA) could be of importance to the detection of the carrier state among fish stocks, but has yet to be validated.
Control methods for VHS currently lie in official health surveillance schemes coupled with control policy measures, such as stamping-out and fallowing procedures, and have resulted in the eradication of the disease from several parts of Europe (Benmansour et al., 1997; Olesen, 1998). At present, genetic approaches using selection and intergeneric hybridisation and vaccination are both at experimental stages.
Viral haemorrhagic septicaemia (VHS) is the most serious viral disease of farmed rainbow trout, Oncorhynchus mykiss, in Europe and responsible for significant losses in many continental European countries. The economic loss due to VHS in Europe has been estimated at £40 million sterling a year (N.J. Olesen, Århus, 1997, personal communication). The disease still poses a threat to farmed trout because of the carrier state of VHS virus (VHSV) in wild marine stocks in water systems close to fish farms (Meier et al., 1994).
The pathogen responsible is a rhabdovirus belonging to the family Rhabdoviridae, the same virus family as rabies virus of dogs and foxes. Some pronounced features of the virus are the ability to cause marked kidney necrosis, especially of the head region, which causes loss of haematopoietic cells important for constitutive and adoptive immunity. In the later stages of the disease, there can be tropism of the brain tissue, a feature shared with other rhabdoviruses. The disease was first described in rainbow trout by Schaperclaus (1938) and a viral aetiology was postulated. It was not until 1963 that Jensen isolated a European strain of the virus and over 20 years before a Pacific marine strain of VHS was reported from the USA (Brunson et al., 1989). Characterization of the virus and its effects has proceeded rapidly, particularly with pathogenicity and vaccine studies in France and Denmark (de Kinkelin, 1988; Wolf, 1988).
Viral haemorrhagic septicaemia is still the most important disease of cultured rainbow trout in the European Union (EU) member states, and no universal vaccine has been developed to date, despite considerable effort. The most effective control therefore remains avoidance.
Topics for further study
A key question in current VHSV research is the true importance of the marine VHSV isolates. These are clearly very close in genetic sequence to the virulent freshwater strains of VHSV from European trout farms. Given that approved zone status of VHS as a disease in farmed fish stocks within continental land masses and water systems is decided by policy-makers from the known prevalence of the virus, how should the marine isolations from sea fish be viewed? The considered answer to this question can be made only when the marine isolates have been thoroughly tested for virulence in freshwater species, their potential for reversion to virulence by in vivo passage determined and comparisons at the gene level made.
[Based upon material originally published in Woo PTK, Bruno DW, eds., 1999. Fish diseases and disorders, Vol. 3 Viral, bacterial and fungal infections. Wallingford, UK: CABI Publishing.]
Host AnimalsTop of page
|Animal name||Context||Life stage||System|
|Clupea pallasii (pacific herring)||Wild host||Aquatic: Adult|
|Coregonus||Experimental settings||Aquatic: Fry|
|Ctenopharyngodon idella (grass carp)||Aquatic: Adult|Aquatic/Fry||Enclosed systems/Ponds|
|Dicentrarchus labrax (European seabass)||Experimental settings|
|Esox lucius (pike)||Domesticated host||Aquatic: Fry|
|Gadus macrocephalus||Wild host||Aquatic: Adult|
|Gadus morhua (Atlantic cod)||Wild host||Aquatic: Adult|
|Melanogrammus aeglefinus (haddock)||Wild host||Aquatic: Adult|
|Micropterus salmoides (largemouth bass)|
|Oncorhynchus aguabonita (golden trout)||Experimental settings|
|Oncorhynchus kisutch (coho salmon)||Wild host||Aquatic: Adult|
|Oncorhynchus mykiss (rainbow trout)||Domesticated host||Aquatic: Fry|
|Oncorhynchus tshawytscha (chinook salmon)||Domesticated host, Wild host||Aquatic: Adult|Aquatic/Fry|
|Paralichthys olivaceus (bastard halibut)||Domesticated host||Aquatic: Adult|
|Psetta maxima (turbot)||Domesticated host||Aquatic: Adult|Aquatic/Broodstock|Aquatic/Fry||Enclosed systems/Tanks|
|Salmo salar (Atlantic salmon)||Domesticated host, Experimental settings, Wild host||Aquatic: Adult|Aquatic/Fry||Enclosed systems/Cages|
|Salmo trutta (sea trout)||Experimental settings||Aquatic: Fry|
|Salvelinus fontinalis (brook trout)|
|Salvelinus namaycush (lake trout)|
|Thymallus thymallus (grayling)||Aquatic: Adult|Aquatic/Fry|
Hosts/Species AffectedTop of page
Rainbow trout are especially susceptible to VHSV, and the classical signs are seen in first-feeding fry. This species is an imported species to Europe and the UK, being shipped from the USA in the last century. Speculating whether the disease would have been seen is interesting, if rainbow trout had not been imported from the USA first into France in 1879 and then later farmed in Denmark. Rainbow trout in sea-water cages are also susceptible; Castric and de Kinkelin (1980) reported an outbreak where 85% mortality was observed 80 days after sea-water transfer.
Other species of trout are susceptible to VHSV under natural conditions, including brook trout, Salvelinus fontinalis (Rasmussen, 1965), and lake trout, Salvelinus namaycush (Ghittino, 1973). Within the genus Salmo, Atlantic salmon, Salmo salar (Rasmussen, 1965), brown trout, Salmo trutta, and golden trout, Salmo aguabonita (=Oncorhynchus aguabonita), are susceptible to VHSV. Wild freshwater inhabitants of streams, rivers and lakes in Germany and Switzerland are susceptible, notably pike, Esox lucius, grayling, Thymallus thymallus, and whitefish, Coregonus sp. (Reichenbach-Klinke, 1959; Ahne and Thomsen, 1985; Meier et al., 1994).
While it is stated that species such as Atlantic salmon are susceptible to experimental infection by intraperitoneal infection (de Kinkelin and Castric, 1982), an important distinction must be made between natural hosts of VHSV and experimental hosts. Currently, Atlantic salmon in European coastal waters is not considered a natural host of VHSV and, to date, represents only an experimental model. This was confirmed by the data of King et al. (2001a) who showed that juvenile Atlantic salmon were not susceptible by water bathing to a range of VHSV genotypes. However, only time will tell whether any marine strains of VHSV can be pathogenic to Atlantic salmon.
Farmed marine species, such as sea bass (Dicentrarchus labrax) and turbot (Scophthalmus maximus) from the Mediterranean area are susceptible. This was shown experimentally by Castric and de Kinkelin (1984). Virus multiplication was demonstrated in both species and histopathology described. Clinical disease in VHS outbreaks on turbot farms has also been described in Germany by Schlotfeldt et al. (1991) and in Scotland on Gigha Island by Ross et al. (1995). In the latter case it was the largest tank-reared 3-year-old turbot that showed the highest mortality, suggesting that the infection had been dormant for some months during growth.
The last 6 years have seen many further reports on the extension of the host range of VHSV. In the north west Atlantic Ocean, Dopazo et al. (2002) reported VHSV isolation from apparently healthy Greenland halibut (Reinhardtius hippoglossoides) in the Flemish cap, a deep water fishing ground off Newfoundland. In the northern North Sea, Smail (2000) reported the isolation of VHSV from cod (Gadus morhua) with the ulcus syndrome and also from haddock (Melanogrammus aeglefinus) with skin haemorrhages (Fig. Haddock (Melanogrammus aeglefinus) tail showing skin haemorrhages from which VHS virus was isolated, east of Shetland, Scotland, June 1995). A survey of the North Sea, north east Atlantic Ocean and Irish Sea over 2 years reported isolations from healthy Norway pout (Trisopterus esmarkii), cod with a skin lesion, herring (Clupea harengus), whiting (Merlangius merlangus) and poor cod (Trisopterus minutus) (King et. al., 2001b). Monitoring for VHSV in the southern marine waters around the UK reported one isolation from herring (Clupea harengus) in the English Channel and positive RT-PCR signals in herring and cod in other years (Dixon et al., 2003).
Around Japan, isolation of VHSV has been reported in wild Japanese flounder (Paralichthys olivaceus) and sand lance (Ammodytes personatus) (Watanabe et al., 2002; Takano et al., 2001). In Alaska, following the report of VHSV in pacific herring (Meyers et al., 1994) with mortality, 2 new species were reported as susceptible with high mortalities: pacific hake (Merluccius productus) and walleye pollock (Theragra chalcogramma) (Meyers et al., 1999) Mass mortality was described simultaneously in the pacific herring, pacific hake and walleye pollock.
In common with most fish virus diseases, stress plays a role in the induction of disease, inducing recurrence or recrudescence of the infection from a carrier state. A classic example of this was described by Hørlyck et al. (1984) in marine-cultured rainbow trout in Denmark. Overt disease was seen in rainbow trout growers in sea water that were previously screened virus negative in fresh water, and mortality varied from 12 to 50% in four cases. This event proved that VHSV can lie dormant, escaping standard tissue-culture detection and virus isolation methods, and can reactivate later after the stress of handling or difficulty with osmoregulation in sea water. De novo induction of VHS as an apparent recurrence of disease points to a modulating role for the stress hormone cortisol and other corticosteroids in the induction of VHSV replication in the kidney and spleen, the principal leucopoietic tissues.
Since the date of the above isolation, the existence of marine isolates of VHSV has been discovered and an alternative explanation for the new isolation in sea-water-cultured rainbow trout was de novo infection from a marine source.
Rainbow trout fry of 0.3-3.0 g are the most susceptible to disease, with target organs that represent most of the body weight. Mortality of 80-100% due to virulent strains is normal at optimum temperatures of 9-12°C. Fingerlings and growers are also susceptible but mortality is lower, in the range 10-50% for virulent strains, with the nervous chronic state more commonly seen in these fish.
Viral haemorrhagic septicaemia is a cold-water disease, with a temperature range of 2-12°C, although the optimum is 9-12°C, where replication of the virus is greatest. Temperatures above 15°C are inhibitory to virus growth. Vestergård-Jørgensen (1982) showed that temperature was inversely correlated with the longevity of the infection in rainbow trout. In groups of 11 g trout transferred to 5, 10, 15 or 20°C after infection at 10°C, persistence was 14 weeks at 5°C, in contrast to 2 weeks at 10°C and undetectable at 20°C. Neukirch (1985) showed that extended persistent infection of 300-400 days could be established in rainbow trout at 4°C.
DistributionTop of page
Looking to the wider distribution of the VHSV and its host range, a significant finding was the discovery of virus in anadromous salmonids in 1988 off the western Pacific coast of the USA, i.e. in migrating coho salmon (Oncorhynchus kisutch) and chinook salmon (Oncorhynchus tshawytscha). The first Pacific type strain of VHSV, termed Makah, was identified and named after the Makah National Fish Hatchery in Washington State, USA, where the migratory fish were sampled (Brunson et al., 1989). The isolation of this first Pacific strain of virus was unexpected and led regulatory authorities to ask whether VHSV had a marine source and a global distribution, and whether the marine Pacific strains were different in genetic constitution from the established European strains from freshwater trout.
The disease is currently seen widely in continental Europe in trout farms. It has been seen in turbot at a marine fish farm in Scotland, UK (Ross et al. 1995), and Ireland (J. McArdle, Dublin, 1997, personal communication). Marine forms of the virus are known from Alaska and Washington State, USA, in the Pacific Ocean. Marine forms of the virus have also been isolated from the North Sea in cod, Gadus morhua, with skin ulcers and haddock, Melanogrammus aeglefinus, with skin haemorrhages (Smail, 2000).
Viral haemorrhagic septicaemia virus has now been found in other areas of the world where salmonid aquaculture is practised, i.e. in Japan and the west coast of USA (Hedrick et al., 2003), extending the distribution down to the Californian coast.
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Atlantic, Northeast||Smail , 1995|
|Pacific, Northeast||Present||Brunson and et al. , 1989|
|Afghanistan||No information available||OIE, 2009|
|Armenia||Disease never reported||OIE, 2009|
|Azerbaijan||Disease never reported||OIE, 2009|
|Bahrain||Disease never reported||OIE, 2009|
|Bangladesh||Disease never reported||OIE, 2009|
|Bhutan||No information available||OIE, 2009|
|Brunei Darussalam||Disease not reported||OIE Handistatus, 2005|
|Cambodia||No information available||OIE, 2009|
|China||Disease never reported||OIE, 2009|
|-Hong Kong||Disease never reported||OIE, 2009|
|Georgia (Republic of)||Disease never reported||OIE Handistatus, 2005|
|India||No information available||OIE, 2009|
|Indonesia||Disease not reported||OIE, 2009|
|Iran||Disease not reported||OIE, 2009|
|Iraq||Disease never reported||OIE, 2009|
|Israel||Disease never reported||OIE, 2009|
|Japan||Disease not reported||OIE, 2009|
|Jordan||No information available||OIE, 2009|
|Kazakhstan||Disease not reported||OIE, 2009|
|Korea, DPR||Disease not reported||OIE Handistatus, 2005|
|Korea, Republic of||Present||OIE, 2009|
|Kuwait||Disease not reported||OIE, 2009|
|Kyrgyzstan||Disease not reported||OIE, 2009|
|Laos||No information available||OIE, 2009|
|Lebanon||No information available||OIE, 2009|
|Malaysia||Disease never reported||OIE, 2009|
|-Peninsular Malaysia||Disease never reported||OIE Handistatus, 2005|
|-Sabah||No information available||OIE Handistatus, 2005|
|-Sarawak||No information available||OIE Handistatus, 2005|
|Mongolia||No information available||OIE, 2009|
|Myanmar||No information available||OIE, 2009|
|Nepal||No information available||OIE, 2009|
|Oman||No information available||OIE, 2009|
|Pakistan||No information available||OIE, 2009|
|Philippines||No information available||OIE, 2009|
|Qatar||No information available||OIE, 2009|
|Saudi Arabia||No information available||OIE, 2009|
|Singapore||Disease never reported||OIE, 2009|
|Sri Lanka||No information available||OIE, 2009|
|Syria||No information available||OIE, 2009|
|Taiwan||Disease never reported||OIE Handistatus, 2005|
|Tajikistan||No information available||OIE, 2009|
|Thailand||No information available||OIE, 2009|
|Turkey||Disease not reported||OIE, 2009|
|Turkmenistan||Disease not reported||OIE Handistatus, 2005|
|United Arab Emirates||No information available||OIE, 2009|
|Uzbekistan||Disease not reported||OIE Handistatus, 2005|
|Vietnam||No information available||OIE, 2009|
|Yemen||No information available||OIE, 2009|
|Algeria||No information available||OIE, 2009|
|Angola||No information available||OIE, 2009|
|Benin||No information available||OIE, 2009|
|Botswana||No information available||OIE, 2009|
|Burkina Faso||No information available||OIE, 2009|
|Burundi||Disease never reported||OIE Handistatus, 2005|
|Cameroon||Disease never reported||OIE Handistatus, 2005|
|Cape Verde||Disease not reported||OIE Handistatus, 2005|
|Central African Republic||Disease not reported||OIE Handistatus, 2005|
|Chad||No information available||OIE, 2009|
|Congo||No information available||OIE, 2009|
|Congo Democratic Republic||Disease not reported||OIE Handistatus, 2005|
|Côte d'Ivoire||No information available||OIE Handistatus, 2005|
|Djibouti||No information available||OIE, 2009|
|Egypt||No information available||OIE, 2009|
|Eritrea||No information available||OIE, 2009|
|Ethiopia||No information available||OIE, 2009|
|Gambia||No information available||OIE, 2009|
|Ghana||No information available||OIE, 2009|
|Guinea||No information available||OIE, 2009|
|Guinea-Bissau||No information available||OIE, 2009|
|Kenya||No information available||OIE, 2009|
|Lesotho||Disease never reported||OIE, 2009|
|Libya||No information available||OIE Handistatus, 2005|
|Madagascar||No information available||OIE, 2009|
|Malawi||No information available||OIE, 2009|
|Mali||No information available||OIE, 2009|
|Mauritius||No information available||OIE, 2009|
|Morocco||No information available||OIE, 2009|
|Mozambique||Disease not reported||OIE, 2009|
|Namibia||No information available||OIE, 2009|
|Nigeria||No information available||OIE, 2009|
|Réunion||No information available||OIE Handistatus, 2005|
|Rwanda||No information available||OIE Handistatus, 2005|
|Sao Tome and Principe||Disease not reported||OIE Handistatus, 2005|
|Senegal||No information available||OIE, 2009|
|Seychelles||Disease not reported||OIE Handistatus, 2005|
|Somalia||No information available||OIE Handistatus, 2005|
|South Africa||No information available||OIE, 2009|
|Sudan||Disease never reported||OIE, 2009|
|Swaziland||No information available||OIE, 2009|
|Tanzania||No information available||OIE, 2009|
|Togo||No information available||OIE, 2009|
|Tunisia||Disease not reported||OIE, 2009|
|Uganda||No information available||OIE, 2009|
|Zambia||No information available||OIE, 2009|
|Zimbabwe||No information available||OIE, 2009|
|Bermuda||Disease not reported||OIE Handistatus, 2005|
|Canada||Restricted distribution||OIE, 2009|
|Greenland||Disease never reported||OIE, 2009|
|Mexico||Disease not reported||OIE, 2009|
|USA||Restricted distribution||OIE, 2009|
|-Alaska||Present||Meyers and et al. , 1994|
|-Georgia||Disease never reported||OIE, 2009|
|-Washington||Present||Brunson and et al. , 1989|
Central America and Caribbean
|Barbados||Disease never reported||OIE Handistatus, 2005|
|Belize||Disease never reported||OIE, 2009|
|British Virgin Islands||Disease never reported||OIE Handistatus, 2005|
|Cayman Islands||Disease never reported||OIE Handistatus, 2005|
|Costa Rica||Disease never reported||OIE, 2009|
|Cuba||Disease never reported||OIE, 2009|
|Curaçao||Disease not reported||OIE Handistatus, 2005|
|Dominica||Disease not reported||OIE Handistatus, 2005|
|Dominican Republic||Disease never reported||OIE Handistatus, 2005|
|El Salvador||No information available||OIE, 2009|
|Guadeloupe||No information available||OIE, 2009|
|Guatemala||Disease never reported||OIE, 2009|
|Haiti||No information available||OIE, 2009|
|Honduras||No information available||OIE, 2009|
|Jamaica||Disease never reported||OIE, 2009|
|Martinique||Disease not reported||OIE, 2009|
|Nicaragua||Disease never reported||OIE, 2009|
|Panama||No information available||OIE, 2009|
|Saint Kitts and Nevis||Disease never reported||OIE Handistatus, 2005|
|Saint Vincent and the Grenadines||Disease not reported||OIE Handistatus, 2005|
|Trinidad and Tobago||Disease never reported||OIE Handistatus, 2005|
|Argentina||Disease never reported||OIE, 2009|
|Bolivia||No information available||OIE, 2009|
|Brazil||Disease never reported||OIE, 2009|
|Chile||Disease never reported||OIE, 2009|
|Colombia||Disease never reported||OIE, 2009|
|Ecuador||No information available||OIE, 2009|
|Falkland Islands||Disease never reported||OIE Handistatus, 2005|
|French Guiana||Disease not reported||OIE, 2009|
|Guyana||Disease never reported||OIE Handistatus, 2005|
|Paraguay||Disease never reported||OIE Handistatus, 2005|
|Peru||No information available||OIE, 2009|
|Uruguay||No information available||OIE, 2009|
|Venezuela||No information available||OIE, 2009|
|Albania||No information available||OIE, 2009|
|Andorra||Disease never reported||OIE Handistatus, 2005|
|Belarus||Disease never reported||OIE, 2009|
|Bosnia-Hercegovina||Disease not reported||OIE Handistatus, 2005|
|Croatia||Disease never reported||OIE, 2009|
|Cyprus||Disease never reported||OIE, 2009|
|Czech Republic||Present||OIE, 2009|
|Denmark||Restricted distribution||NULL||Olesen and et al. , 1993; OIE, 2009|
|Estonia||No information available||OIE, 2009|
|Finland||Restricted distribution||OIE, 2009|
|France||Disease not reported||200803||Kinkelin and Le Berre , 1977; OIE, 2009|
|Germany||Present||NULL||Schlotfeldt and et al. , 1991; OIE, 2009|
|Greece||No information available||OIE, 2009|
|Hungary||Disease never reported||OIE, 2009|
|Iceland||Disease never reported||OIE, 2009|
|Ireland||Disease not reported||OIE, 2009|
|Isle of Man (UK)||Disease never reported||OIE Handistatus, 2005|
|Jersey||Disease never reported||OIE Handistatus, 2005|
|Latvia||Disease not reported||OIE, 2009|
|Liechtenstein||No information available||OIE, 2009|
|Lithuania||Disease not reported||OIE, 2009|
|Luxembourg||No information available||OIE, 2009|
|Macedonia||No information available||OIE, 2009|
|Malta||No information available||OIE, 2009|
|Moldova||Disease never reported||OIE Handistatus, 2005|
|Montenegro||No information available||OIE, 2009|
|Netherlands||Disease not reported||OIE, 2009|
|Portugal||Disease not reported||OIE, 2009|
|Romania||Disease not reported||OIE, 2009|
|Russian Federation||No information available||OIE, 2009|
|Serbia||No information available||OIE, 2009|
|Slovenia||Disease not reported||OIE, 2009|
|Spain||Disease not reported||OIE, 2009|
|Sweden||Disease not reported||OIE, 2009|
|Switzerland||Disease not reported||200805||Meier and et al. , 1994; Knuesel et al., 2003; OIE, 2009|
|UK||Present||NULL||Ross and et al. , 1995; OIE, 2009|
|-Northern Ireland||Disease never reported||OIE Handistatus, 2005|
|Ukraine||Disease not reported||OIE, 2009|
|Yugoslavia (former)||No information available||OIE Handistatus, 2005|
|Yugoslavia (Serbia and Montenegro)||No information available||OIE Handistatus, 2005|
|Australia||Disease never reported||OIE, 2009|
|French Polynesia||Disease never reported||OIE, 2009|
|New Caledonia||Disease not reported||OIE, 2009|
|New Zealand||Disease never reported||OIE, 2009|
|Samoa||No information available||OIE Handistatus, 2005|
|Vanuatu||Disease never reported||OIE Handistatus, 2005|
|Wallis and Futuna Islands||No information available||OIE Handistatus, 2005|
PathologyTop of page
The pathology in rainbow trout is the best described. A common feature in all hosts, salmonids or flatfish, is the widespread haemorrhaging seen internally, over the liver, in adipose tissue (see Fig. Rainbow trout (Oncorhynchus mykiss) fingerling with clinical VHS, showing oedema, pale liver and haemorrhages over the adipose tissue) and especially within the muscle.
Affected rainbow trout fry are lethargic and darker in pigmentation, swim erratically or have difficulty in orientation and may congregate along the sides of tank and pond outlets. Flashing, corkscrewing motion and surface swimming is also seen in the later nervous-form-affected trout fingerlings. Exophthalmia may be seen in one or both eyes (see Fig. Rainbow trout (Oncorhynchus mykiss) fingerling with clinical VHS, showing exophthalmia and pale necrotic kindney, and Fig. Turbot (Scophthalmus maximus) with VHS at a farm outbreak, showing exophthalmia) and there can be haemorrhage around the eye orbit (Fig. Turbot (Scophthalmus maximus) experimental VHS infection with a virulent freshwater strain, showing haemorrhaging around the eye orbit). The gills are markedly pale, reflecting generalized anaemia.
The most pronounced changes are in the kidney and liver. The kidney is swollen and darker red in the early stages, but the head and midsection are often totally necrosed and pale in dead trout fry and fingerlings (see Fig. Rainbow trout (Oncorhynchus mykiss) fingerling with clinical VHS, showing exophthalmia and pale necrotic kindney). The liver is paler than normal, yellowish with areas of mottled haemorrhage. A systemic haemorrhage may be evident in the ocular tissues and skin and in the viscera, including the intestinal submucosa and skeletal musculature (see Fig. Rainbow trout (Oncorhynchus mykiss) fingerling with clinical VHS, showing oedema, pale liver and haemorrhages over the adipose tissue). There is no food in the gastrointestinal tract and a stringy mucus can be observed in the lumen.
Viral haemorrhagic septicaemia virus is most pathogenic to the kidney, and early necrosis of the haematopoietic tissue rather than the excretory tubules is a marked feature of the disease. The melanomacrophages, distributed at high density throughout the head kidney, lyse, with release of granules and necrosis. Viral haemorrhagic septicaemia virus has a marked leucotropism, as shown by the fact that mitogen-stimulated leucocytes are lysed by VHSV in in vitro culture (Estepa and Coll, 1991). In contrast the birnavirus infectious pancreatic necrosis virus (IPNV) sets up a persistent or non-lytic infection (Knott and Munro, 1986). In rainbow trout the liver shows widespread focal necrosis, degeneration of hepatocyte nuclei, granulation of chromatin and the accumulation of hyaline material in the tubular lumen. In other hosts, especially pike fry, there may be extravasation or bloody swelling, deposition of blood in the muscle and pancreatic necrosis (Meier and Vestergård-Jørgensen, 1980). The liver sinusoids become engorged with blood and show widespread necrosis and many pyknotic and karyolytic nuclei without other diagnostic inclusions. Many foci of erythrocytes occur within the musculature.
The heart is also a target organ. While heart pathology has not been described in salmonids, in turbot from the VHS outbreak at Gigha Island (Ross et al. 1995) a very marked necrosis of the ventricular fibres was seen. Heart failure is probably a prime cause of death in this bottom-living species, as the rate of opercular movement increases in moribund VHSV-infected turbot.
DiagnosisTop of page
A variety of established fish cell lines have proved susceptibility to VHSV. Bluegill fry (BF-2) and rainbow trout gonad (RTG-2) cells are the most sensitive cell lines for a serogroup I freshwater virulent isolate from rainbow trout and also a serogroup III isolate (Olesen and Vestergård-Jørgensen, 1992; Lorenzen E. et al., 1999a). Other possible cell lines for use are epithelioma papulosum cyprini (EPC), fathead minnow (FHM), chinook salmon embryo (CHSE-214) or pike gonad (PG) (Wolf, 1988). Cell lineage and passage number can have a great effect on virus replication as evidenced by titres of several fish viruses including VHSV on CHSE-214 cells (McAllister 1997). The cytopathic effect (CPE) consists of pronounced cell shrinkage with some cell rounding. In RTG-2 cells a partial differential diagnosis of virus type can be made depending on the plaque type; VHSV causes ‘staggered edge’ plaques in RTG-2 cells, whereas IPNV causes more even-edged plaques.
Once the viral CPE is recognized, the supernatant virus is harvested and the virus identified by one of several techniques. The virus can be identified by a serum neutralization test on any of the above susceptible cell lines. This is carried out either by a plaque-neutralization test (PNT) or by a microtitre neutralization test, using a constant dose of virus and varying antiserum dilutions. The PNT for VHSV is highly dependent on heat-sensitive serum factors in normal serum, i.e. complement. These normal serum proteins assist and stabilize the antigen-antibody cross-linking bonds. Detection of antibodies is a good indicator of previous infection. Such antibodies can be detected by either an enzyme-linked immunosorbent assay (ELISA), an immunofluorescent antibody test (IFAT) or a PNT, as described by Olesen et al. (1991). The ELISA is regarded as the most sensitive technique, although different types of antibody are clearly involved. For example, high values of antibody by one assay may be contrasted with low or nil values by another, e.g. ELISA-positive and PNT negative in one fish and ELISA-negative and PNT positive in another.
Immunofluorescent antibody test
An IFAT was developed by Vestergård-Jørgensen and Meyling (1972) to detect viral antigen in infected tissue cells up to 26 h postinfection. Maximum staining is detected from 18 to 26 h at peak protein synthesis and virus assembly. Staining works well with either a cross-reacting specific MAb to the virus or a polyclonal antibody. A secondary antibody labelled with fluorescein is used which tags the primary antibody and labels infected cells which fluoresce in blue light. The technique of immunofluorescent staining for a viral antigen and viral antibody in the same tissue preparation was perfected by Vestergård-Jørgensen (1974) with clarity and brilliance. Antibody detection by IFAT involves using fixed cells with viral antigen stained with test sera, and then an antispecies antibody labelled with fluorescein is added. Positive slides will show foci of fluorescence positivity, which must be compared with appropriate negative and positive controls.
Enzyme-linked immunosorbent assay
The antigen-capture ELISA is widely practised for the detection of virus in culture supernatants. The first layer in the ELISA is the catching antibody, normally a polyclonal antibody to VHSV but alternatively this may be a MAb. The second step is a blocking reaction to coat unadsorbed solid-phase sites, typically bovine serum albumin or milk protein, and the third step is the addition of virus. The fourth step is the addition of a MAb, normally to G protein, and the fifth is the addition of an antispecies antibody to the monoclonal, which is tagged with an enzyme. Way and Dixon (1988) described VHSV ELISA for a polyclonal direct antigen-capture system. Mourton et al. (1990) reported on three modes of the ELISA for virus detection: indirect ELISA, direct ELISA and antigen-capture ELISA, using a variety of MAbs to the viral glycoprotein G. This resulted in virus detection at a protein concentration as low as 1 ng ml-1 of total proteins where 1.5 fmol ml-1 of envelope glycoprotein was present.
Way (1996) described a rapid dipstick immunosorbent assay (RDIA) for VHSV as a practical fast field method for virus detection from clinical fish samples. Dauber et al. (2001) described development of new MAbs for German VHSV isolates: these authors stressed that the diagnostic use of any MAb may be quite limited diagnostically and needs validating in practice.
Virus gene probes
Deoxyribonucleic acid (DNA) probes to VHSV were developed at the Northwest Biological Science Centre, Seattle, USA (Batts et al., 1993). The nucleotide sequences of the nucleoprotein gene of the Makah strain of VHSV and the virulent French 07-71 strain were determined by Bernard et al. (1992). Comparison showed that the two strains differed by around 13% in sequence homology, sufficient for the Pacific Makah strain and the European 07-71 strain to have diverged and evolved separately over a significant period. This sequence information was of practical use, since Batts et al. (1993) designed three complementary-DNA (cDNA) probes for three different uses. The first probe was universal to all VHSV isolates. It was 29 nucleotides long and was synthesized to be antisense to the messenger RNA sequence from nucleotides 430 to 458 in the open reading frame of the N gene. The second probe was specific to the Makah Pacific strain. It was synthesized to be antisense to a unique 28-base sequence that occurred after the true coding sequence in the N gene of the Makah strain but not in the same region of the 07-71 European strain. The third probe was made to be specific to the European strain only. It was designed and made to be antisense to a 22-base sequence (nucleotides 990 to 1011) within the N gene of the 07-71 strain, where six mismatches occurred with the Makah strain. Each probe was labelled with three biotin molecules at the 5' end of the primer. These three probes were confirmed to detect the two main groups of VHSV by a dot-blotting procedure as follows.
For cDNA detection of viral RNA, the viral messenger RNA with (+) strand polarity is harvested. Then there are four main stages for the technique: RNA extraction and purification, dot-blotting the RNA to a nitrocellulose membrane, probe hybridization to target RNA and spot development. The purification of RNA is achieved by elution from oligo-(deoxythymidine)-cellulose spin columns, with precipitation in sodium acetate and ethanol. The RNA is bound to a nitrocelluose membrane, using a vacuum with concentrated binding buffer conditions. Once the RNA is permanently linked, a prehybridization solution, containing annealing buffers and salmon sperm DNA as background stabilizer, is introduced to the membrane and the cDNA probe is allowed to hybridize at medium temperature of 53°C overnight. The membranes are then rinsed and the presence of the probe, with attached biotin molecules, is detected by using a streptavidin/alkaline phosphatase (SAP) conjugate, with development of a SAP-specific staining reaction.
Polymerase chain reaction detection of viral haemorrhagic septicaemia ribonucleic acid sequences
Two groups reported the use of a reverse transcriptase-dependent polymerase chain reaction (PCR) not only for specific detection of VHSV sequences but also for differentiation of serologically similar strains of VHSV (Bruchof et al., 1995; Einer-Jensen et al., 1995). The principle of the PCR detection of viral RNA sequences is as follows. A virion RNA harvest is made from freshly grown virus by diethyl pyrocarbonate extraction and the RNA purified. cDNA is made from this RNA, using a reaction mixture of reverse transcriptase and nucleotides in suitable buffers. This specific cDNA is amplified by PCR, using Super Taq polymerase in a temperature cycler. The amplified cDNA products are then electrophoresed in agarose slab gels, stained with ethidium bromide and the DNA bands visualized under ultraviolet (UV) light.
Einer-Jensen et al. (1995) used two primers that amplified sequences from the N gene of European and American (Pacific) strains of VHSV. They used another primer that amplified only the American strains tested, which revealed a unique non-coding intron of 20 nucleotides in close proximity to the N gene. American and European strains of VHSV could therefore be readily distinguished in a half-day’s work.
Bruchof et al. (1995) also described reverse-transcriptase PCR to distinguish VHSV and IHNV, using specific primers in the glycoprotein gene. This was highly specific and fast, allowing differentiation of virus types within 8 h.
Monoclonal antibody capture and polymerase chain reaction
Estepa et al. (1995) described a MAb capture assay combined with PCR detection of VHSV, using sense primers to the G gene in a defined region of 379 base pairs (amino acids 64-195). This method gave virus-specific gel electrophoresis products for IHNV and VHSV with N and G gene primers and was regarded as quick, highly sensitive and specific. There is the possibility that it could also be automated.
Comparison of methods
Different methods can be compared with respect to sensitivity and specificity. Tissue culture can detect cell culture-adapted virus in both diseased and carrier fish in most, but not all, cases. Some presumptive idea of the virus identity is given by the type of CPE observed.
The ELISA antigen test in a double sandwich format can detect virus at a low level, down to log10 4.0 TCID50 g-1 tissue (Mourton et al., 1990), where 1.5 fmol ml-1 of glycoprotein was present. This is a very good sensitivity for an ELISA and a highly specific assay for VHSV.
Tissue culture isolation has been compared with immunohistochemistry by Evensen et al. (1994) and tissue culture was shown to be the more sensitive in rainbow trout infected tissues. Tissue culture has also been compared with PCR detection of virus by Bruchof et al. (1995) and PCR detection shown to be more sensitive in a proportion of rainbow trout carriers.
List of Symptoms/SignsTop of page
|Finfish / Change in shape (e.g. distension) - Eyes||Aquatic:Adult||Sign|
|Finfish / Corkscrewing - Behavioural Signs||Aquatic:Fry||Sign|
|Finfish / Darkened coloration - Skin and Fins||Aquatic:Adult||Sign|
|Finfish / Fish swimming near surface - Behavioural Signs||Aquatic:Fry||Sign|
|Finfish / Flashing - Behavioural Signs||Aquatic:Fry||Sign|
|Finfish / Generalised lethargy - Behavioural Signs||Aquatic:Adult||Sign|
|Finfish / Haemorrhaging - Body Cavity and Muscle||Aquatic:Adult||Sign|
|Finfish / Kidney - white-grey patches (haemorrhage / necrosis / tissue damage) - Organs||Aquatic:Adult||Sign|
|Finfish / Kidney swelling / oedema - Organs||Aquatic:Adult||Sign|
|Finfish / Liver - white / grey patches (haemorrhage / necrosis / tissue damage) - Organs||Aquatic:Adult||Sign|
|Finfish / Mortalities -Miscellaneous||Aquatic:Adult||Sign|
|Finfish / Mucus-filled intestines - Organs||Aquatic:Adult||Sign|
|Finfish / Paleness - Gills||Aquatic:Adult||Sign|
Disease CourseTop of page
In rainbow trout, the disease occurs in three varied forms, i.e. acute, chronic and nervous, followed by a carrier state in surviving fish, where the virus can be isolated from persistently infected tissues, such as kidney and brain. In the acute stage, from day 0 to 30 by experiment at 8-12°C, sick fish are seen lying on the tank bottom or moribund, displaying the preacute signs of flashing or corkscrewing. The chronic stage reflects a lasting persistent infection and is correlated with no obvious external signs. Virus can be isolated from all the major internal organs and there are subacute depressive effects of the virus on red and white blood cell counts. In the nervous form, there are very marked aberrations of swimming behaviour, i.e. constant flashing, tail-chasing and spiralling. This is associated with a most marked tropism of the virus to the brain, and brain virus titres of 1 × 109 TCID50 g-1 can be attained for the 07-71 strain in nervous-form rainbow trout by experimental challenge (D.A. Smail, unpublished observations).
Nervous tropism is a feature of virulent freshwater strains, such as type F1 Voldbjerg or 07-71, but can be observed with marine strains, such as that from turbot. The nervous tropism of the virulent strains is not completely understood but is likely to be closely related to the specific interaction of the glycoproteins in the virus envelope with membrane lipoproteins in target cells of susceptible nervous tissue. In this connection, Bearzotti et al. (1995) have identified two loci for amino acid substitution in the glycoprotein gene (residues 140 and 430) which are uniquely associated with reduced virulence and increased tissue tropism for nervous tissue.
Pathogenesis: spread of infection
Viral haemorrhagic septicaemia virus gains entry to fish by crossing the gill epithelium and thence quickly, via the blood, to the main internal organs. There is also evidence that the skin can be a portal of virus entry. Yamamoto et al. (1992) showed that skin explants could be infected with virus, with increasing titres in time. By implication, this could also mean that VHSV-infected skin cells are a portal of exit for the virus.
By adhering to, penetrating and multiplying in the endothelial cells lining venules and sinusoids, the virus causes the first degree of pathology to cells lining the circulatory system, from 48 h after infection. Then the virus spreads to the head kidney and causes cell degeneration and necrosis of haematopoietic cells of the kidney and of macrophages and melanomacrophages, from 2-4 days postinfection. Next, infection spreads to the liver (4 days), with necrosis of hepatocytes and, later, necrosis of pancreatic acinar cells (6 days) (Evensen et al., 1994). It was shown by Evensen et al. (1994) for the F1 strain infection of rainbow trout that immunohistochemical staining for viral antigen could show the virus distribution. However, at day 2 postinfection, virus could be isolated without positive immunostaining, showing virus isolation to be more sensitive than immunostaining.
Shedding of virus from carrier fish takes place via the urine by day 3, as shown in experimental bath infection of rainbow trout (Neukirch, 1985), but there is no evidence of faecal excretion, since the virus is sensitive to gut acids. It is likely that for sea fish, such as cod and haddock, bearing ulcers with VHSV (see Fig. Haddock (Melanogrammus aeglefinus) tail showing skin haemorrhages from which VHSV was isolated), the skin is also a source of virus shedding.
In marine species, e.g. turbot, a similar pattern of pathogenesis takes place, there is haemorrhaging over the eye orbit and fin margins and marked swelling, due to impaired osmoregulation (see Fig. Turbot (Scophthalmus maximus) with VHS at a farm outbreak, showing exophthalmia, and Fig. Turbot (Scophthalmus maximus) experimental VHS infection with a virulent freshwater strain, showing haemorrhaging around the eye orbit). The sequence of virus entry, via gills to bloodstream and thence to the target organs, head kidney and liver, is also repeated. Necrosis of target haematopoietic cells quickly leads to kidney malfunction and morbidity (Castric and de Kinkelin, 1984).
Rainbow trout exposed to VHS virus that survive infection develop different types of immunity, both non-specific, due to interferon (IFN), and specific, due to antibodies, both neutralizing and non-neutralizing (de Kinkelin, 1988).
De Kinkelin and Dorson (1973) showed that experimentally infected rainbow trout produced IFN and this peaked at day 3, the time when it could have the most inhibitory effect. The characteristics of the molecule were described. It was shown to have broad antiviral activity against VHSV, IHNV and IPNV, was cell-specific, heat-stable, stable at pH 2, trypsin-labile, ribonuclease (RNAse)-resistant and non-dialysable. Renault et al. (1991) later described a spectrophotometric method for the titration of such IFN, which could be especially useful for measuring IFN in young fish, where only small volumes of sera might be available.
The relationship of the production of IFN to virulence was studied by Bernard et al. (1985), in an original paper on VHS immunogenicity and virulence. The virulent strain 07-71 stimulated an IFN response in 10-20 g rainbow trout by 48-72 h postinfection, whereas two attenuated strains, F25 (heat-attenuated/Danish) and 07-71 (passage attenuated/EPC), did not stimulate an IFN response. The significance of this difference could not be interpreted or related to the sensitivity of the different strains to IFN, for no difference was found in the in vitro sensitivity of the 07-71 strain or passage-attenuated 07-71 strain tested on RTG-2 cells.
Neutralizing antibodies have been described for trout recovering from VHS infection. The assay is dependent on trout complement in normal serum for effective neutralization (Dorson and Torchy, 1979). An interesting facet to the assay, as pointed out by de Kinkelin (1988) is that neutralization is progressive and dependent on the time of incubation of antibody and virus. Incubation for 16 h at 4°C increases the serum neutralization end-point by 20 compared with the value obtained at 1 h at 20°C.
Neutralizing antibody responses take a variable time to develop. Olesen et al. (1991) found that 130 g trout exposed to VHSV Voldberg strain by cohabitation developed antibodies 4-10 weeks later. Other assays for antibody, such as immunofluorescence (IF) and ELISA, detected antibody at different starting times and there was no strict correlation between positivity in the three assays.
Temperature plays a profound role in the development of the immune response as reported by Vestergård-Jørgensen (1982) in the trout response to the Reva live vaccine strain. Basically, a low temperature of 5-10°C was optimal in stimulating a good priming of the immune response. In contrast, too rapid clearance of the virus at high temperatures (15-20°C), involving IFN production, mitigated against the antibody response, giving a very low titre of antibody.
EpidemiologyTop of page
Viral haemorrhagic septicaemia virus has a widespread distribution in a variety of wild freshwater lake and river fish and more evidence is coming forward that the virus is also prevalent in the marine environment. The first report of VHSV in a migratory anadromous salmonid by Brunson et al. (1989) was a surprise to US policy-makers, as VHS was previously regarded as only a freshwater disease. Then came isolations from Pacific cod, Gadus macrocephalus, by Meyers et al. (1992) and Pacific herring, Clupea harengus pallasi (Meyers et al.,1994), off Alaska. In the North Sea, isolation of a rhabdovirus was made from cod by Jensen et al. (1979) and identified as VHSV by Vestergård-Jorgensen and Olesen (1987). Smail (1995) then corroborated this isolation with further isolations from cod off the Shetland Isles, Scotland, in 1993. It is now known that there are four hosts of VHSV in the Baltic Sea - herring, sprat (Sprattus sprattus), cod and rockling (Mortensen et al., 1999) - and this evidence lends weight to a likely marine origin for VHSV.
For epizootiology, the reader should also consult the sections on animals affected and distribution as there is much overlap of information.
Transmission of the disease
Shedding and transmission of the virus in European countries is thought to have taken place with lateral spread from infected fish farms and development of the carrier state in wild fish nearby, i.e. in several European states reinfection of farm stocks has taken place after eradication of the VHS infection on the farms. Shedding via the urine from clinically infected rainbow trout has been demonstrated as the means of virus dissemination (Neukirch, 1985). However, the degree of virus shedding from persistently infected fish is more conjectural and has not been documented.
Impact SummaryTop of page
|Fisheries / aquaculture||Negative|
Impact: EconomicTop of page
The economic loss due to VHS in Europe has been estimated at £40 million sterling a year in 1997.
Zoonoses and Food SafetyTop of page
This species is not a zoonosis.
ReferencesTop of page
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