spring viraemia of carp
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- spring viraemia of carp
International Common Names
- English: spring viremia of carp
OverviewTop of page
Spring viraemia of carp (SVC) is an acute, systemic, contagious disease caused by a rhabdovirus, which primarily affects common carp. Several other cyprinids and some other species are also susceptible. Spring viraemia of carp typically occurs at water temperatures below 18°C, and predominantly in the spring. The term SVC and the name Rhabdovirus carpio (RVC) for the causal agent were introduced by Fijan et al. (1971). However, the virus name approved by the International Committee on Taxonomy of Viruses is spring viraemia of carp virus (SVCV) and it is classified as a tentative species in the Vesiculovirus genus of the virus family Rhabdoviridae. For many years the disease was only recognized in certain countries in Europe, and in states of the former USSR. In recent years, however, the range of the disease has spread to countries in North and South America, and to Asia. Prior to the description of SVC as a separate aetiological and clinical entity, this disease was part of the condition known as infectious dropsy of carp (IDC) or infectious ascites, haemorrhagic septicaemia, red contagious disease or rubella. Infectious dropsy of carp was initially considered to be a bacterial disease. Distinction of two forms of IDC - an acute or ascitic and a chronic or ulcerative one - was also to some extent a matter of controversy. Mixed infections are common in fish. Fijan (1972) indicated that IDC and its synonyms cover several distinct aetiological and pathological entities. It is clear that the acute or ascitic form of IDC encompassed SVC, and some of the chronic IDC corresponded to carp erythrodermatitis (Fijan, 1972; Bootsma et al., 1977). Swim-bladder inflammation (SBI), Pseudomonas fluorescens infection, motile aeromonad infection and some cases of columnaris disease with skin lesions were also covered in IDC. The viruses isolated from cases of SBI and SVC were indistinguishable by serological tests. Reviews of SVC have been produced by Bootsma and Ebregt (1983), Wolf (1988) and Fijan (1999).
[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|
|Aristichthys nobilis (bighead carp)||Domesticated host|
|Carassius auratus auratus (goldfish)||Domesticated host||Aquatic: Adult|Aquatic/Fry||Enclosed systems/Aquaria (marine / freshwater ornamentals)|
|Carassius carassius (crucian carp)||Domesticated host|
|Ctenopharyngodon idella (grass carp)||Domesticated host|
|Cyprinus carpio (common carp)||Domesticated host, Experimental settings, Wild host||Aquatic: Adult|Aquatic/Broodstock|Aquatic/Fry|
|Danio rerio (zebra danio)||Experimental settings|
|Esox lucius (pike)||Domesticated host|
|Hypophthalmichthys molitrix (silver carp)||Domesticated host|
|Lepomis gibbosus (pumpkinseed)||Experimental settings|
|Leuciscus idus (ide)||Domesticated host|
|Notemigonus crysoleucas (golden shiner)||Experimental settings|
|Oncorhynchus mykiss (rainbow trout)||Domesticated host, Wild host|
|Poecilia reticulata (guppy)||Experimental settings|
|Rutilus rutilus (roach)||Domesticated host|
|Silurus glanis (wels catfish)||Domesticated host|
|Tinca tinca (tench)||Domesticated host|
Hosts/Species AffectedTop of page
Carp is the main species affected by SVC. All four scale-distribution varieties as well as koi carp can be infected. Hill (1977) found that feral carp were more susceptible to SVC than farmed ones. Both young fish and adults are susceptible to the virus. As a result of the seasonality of the disease, the principal age groups affected are 9-12 and 21-24 months old. Although the majority of cases of SVC occur in cultured carp, the disease also occurs in feral carp (Munoz et al., 1994; Dikkeboom et al., 2004).
Other species of fish from which the virus has been isolated are listed in Table 1 and experimentally-susceptible species are listed in Table 2. There have been reports of SVCV in other host species, but as those reports were based on clinical signs alone, they are not listed here. Additional unspecified young pond fishes are also susceptible (OIE Manual, 2003). The rhabdovirus isolated from bighead carp (Aristichthys nobilis) by Rudikov et al. (1975) is serologically unrelated to SVCV (P.R. Pichugina, cited in Shchelkunov and Shchelkunova, 1989). There are no further details of the isolation of SVCV from rainbow trout; the isolate was listed in a table of isolates that were compared by nucleotide sequencing of their G-genes, which demonstrated that the virus tested was SVCV. Under experimental conditions rainbow trout were refractory to an SVCV isolate (Haenen and Davidse, 1993). However, fish such as crucian carp and goldfish that are now recognised as hosts of SVCV were found to be refractory to experimental infection with SVCV isolates (Bachmann and Ahne, 1974; Ahne cited in Wolf, 1988). Wizigmann et al. (1983) reported that SVCV had been isolated from pike fry, but as the virus was identified by the fluorescent antibody test (FAT), which does not discriminate between SVCV and pike fry rhabdovirus (PFR) (see Diagnosis section), it is not certain that the identification was accurate. Likewise a rhabdovirus isolated from the catfish Ictalurus melas [Ameiurus melas] reacted in the FAT using polyclonal SVCV antiserum, but not with a monoclonal antibody against SVCV (Selli et al., 2002); the final identification of that virus awaits further study. The virus can replicate in the fruit fly (Drosophila melanogaster) under experimental conditions (Bussereau et al., 1975), but the involvement of that species in transmission of the disease is improbable. The virus has been isolated from Penaeus stylirostris and P. vannamei in Hawaii (Lu et al., 1991) but there were no reports that finfish there were affected by SVC. The virus was initially called rhabdovirus of penaeid shrimp (RPS), but subsequently it was shown to be serologically related to SVCV (Lu et al., 1994; Lu and Loh, 1994). The nucleotide sequences of the glycoprotein genes of RPS and SVCV were practically identical and hence they should be considered the same virus (Johnson et al., 1999).
Table 1. Fish species from which SVCV has been isolated
|Shchelkunov and Shchelkunova, 1989|
|Jörgensen et al., 1989|
Grass carp, white amur
|Shchelkunov and Shchelkunova, 1989|
|Fijan et al., 1971|
|Koutná et al. (2003)|
|Stone et al., 2003|
Sheatfish, European catfish, wels
|Fijan et al., 1984|
|Dixon et al., 1994|
|Dixon et al., 1994|
Table 2. Fish species experimentally-susceptible to SVCV
|Sanders et al., 2003|
Lebistes reticulatus (=Poecilia reticulata)
|Bachmann and Ahne, 1974|
As in almost all fish diseases, temperature is the main factor determining the course and outcome of SVCV infection. Under experimental conditions, high mortality occurs within 10-15 days at 16-17°C but later at 11-15°C (Fijan et al., 1971). The disease does not develop in infected fish kept at 20-22°C or higher (Fijan, 1976) or at a constant temperature of 18°C (Baudouy et al., 1980a,b,c) or 20°C (Ahne, 1980). In several long-term laboratory experiments, Baudouy et al. (1980a,b,c) investigated the influence of increasing and decreasing temperature on the infection. A gradual increase from 11 to 16°C will lead to faster disease development than a decrease in temperature. The gradual decrease of temperature from 11 to 5°C for 60 days, followed by a slow increase to 20°C during the next 140 days, led to low mortality during the decreasing temperature and massive mortality during the increase from 7 to 14°C (Baudouy et al., 1980c). These findings are in agreement with field observations about the predominant occurrence of SVC outbreaks in the spring. The disease is first registered from March to June (Fijan et al., 1971), but it can also occur in the autumn (Pfeil-Putzien and Baath, 1978). In Bavaria, most cases are diagnosed in April and May and several in June, while single or low numbers occurred in March, July, November and December (Wizigmann et al., 1983). In the Ukraine, the majority of cases appeared in April and May; a single case in carp brood stock was diagnosed in June and a few affected young fish in July (Osadcaja and Rudenko, 1981). In summary, SVC can be expected from November to July, with a peak in April-June (Fijan, 1988). Tropical and subtropical climates are unfavourable for the development of SVC
The immaturity of the defence system in up to 1-month-old carp and sheatfish (Silurus glanis) permits the development of SVC under rearing and laboratory conditions at 23°C, although older fish resist the infection at this temperature (Fijan et al., 1984; Pasco et al., 1987).
The age of fish influences the resistance to SVC. Shchelkunov and Shchelkunova (1989) ascertained that young fish were susceptible and 1.5-year-old fish were resistant. Sheatfish up to 25 days old are susceptible, but older fish become resistant (Pasco et al., 1987). Farmed carp are more susceptible than grass carp, which in turn are more susceptible than the bighead × silver carp hybrid (Shchelkunov and Shchelkunova, 1989).
The causes of differences in susceptibility of groups and individuals of the same species, scale-distribution variety and age to SVC are not known. Field observations indicate that the crowding of fingerlings in winter or in spring and long transportation prior to stocking favour outbreaks and high mortalities. Physiological status of the fish, water quality and stress caused by handling also influence the course and outcome of infection. Munoz et al. (1994) suggested that dehydroabiotic acid, that was present in water following a pollution event, affected the defence mechanism of carp and made them more susceptible to SVCV.
DistributionTop of page
Spring viraemia of carp has been diagnosed in the following countries with low water temperatures during winter: Austria, Bosnia and Hercegovina, Bulgaria, Croatia, Czech Republic, France, Germany, Great Britain, Hungary, Italy, Poland, Romania, Serbia, Slovakia, Spain, Switzerland, Yugoslavia and several countries which were part of the former Soviet Union (Belarus, Georgia, Lithuania, Moldova, Russian Federation and Ukraine). Serological surveys demonstrated virus-neutralizing antibodies in 95% of carp farms in Austria (Kölbl and Kainz, 1977), 86% of farms in Bavaria (Wizigmann et al., 1983) and in Schleswig-Holstein (Hoppe and Werner, 1985). Spring viraemia of carp virus infection is mostly clinically unapparent (Wizigmann et al., 1980); for instance, some farms in Germany with serologically positive fish had no record of SVC outbreaks. In Croatia, the percentage of serologically positive carp varies from one pond to another and usually reaches a peak at the end of the summer (N. Fijan, D. Sulimanovic and Z. Petrinec, 1972-1976, unpublished). Over the 10-year period 1992-2002, 12 of 38 hatcheries in Serbia were positive for SVC by virus isolation and the enzyme-linked immunosorbent assay (ELISA) (Jeremic et al., 2004).
The recorded geographic range of the disease has enlarged in recent years, to include Brazil, the USA and China (Alexandrino et al. 1998; Goodwin, 2002; Stone et al., 2003; Dikkeboom et al., 2004; Liu et al., 2004). The disease was recorded in goldfish in Brazil, and in common and koi carp in the USA. The virus was isolated from apparently healthy common and koi carp from two separate sites in China following a survey of 65 ornamental fish farms (Liu et al., 2003). There had been an earlier report of the occurrence of SVCV in the USA (Inchausty and Heckmann, 1996), based solely on the observation of an enlarged swim bladder in a single carp and virus particles in both nucleus and cytoplasm by electron microscopy, with no confirmation by serology or other means. The supposed virus particles were indistinct, but their presence in both nucleus and cytoplasm would suggest that they were not rhabdoviruses, which are restricted to the cytoplasm.
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.Last updated: 10 Jan 2020
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Burundi||Absent, No presence record(s)||OIE Handistatus (2005)|
|Cabo Verde||Absent, No presence record(s)||OIE Handistatus (2005)|
|Cameroon||Absent, No presence record(s)||OIE Handistatus (2005)|
|Central African Republic||Absent, No presence record(s)||OIE Handistatus (2005)|
|Congo, Democratic Republic of the||Absent, No presence record(s)||OIE Handistatus (2005)|
|Lesotho||Absent, No presence record(s)||OIE (2009)|
|Sudan||Absent, No presence record(s)||OIE (2009)|
|Tunisia||Absent, No presence record(s)||OIE (2009)|
|Armenia||Absent, No presence record(s)||OIE (2009)|
|Azerbaijan||Absent, No presence record(s)||OIE (2009)|
|Bahrain||Absent, No presence record(s)||OIE (2009)|
|Bangladesh||Absent, No presence record(s)||OIE (2009)|
|Brunei||Absent, No presence record(s)||OIE Handistatus (2005)|
|-Beijing||Present, Localized||Stone et al. (2003); Liu et al. (2004)|
|-Hebei||Present||Liu et al. (2004)|
|Georgia||Absent, No presence record(s)||OIE Handistatus (2005)|
|Hong Kong||Absent, No presence record(s)||OIE (2009)|
|Indonesia||Absent, No presence record(s)||OIE (2009)|
|Iran||Absent, No presence record(s)||OIE (2009)|
|Iraq||Absent, No presence record(s)||OIE (2009)|
|Israel||Absent, No presence record(s)||OIE (2009)|
|Japan||Absent, No presence record(s)||OIE (2009)|
|Kazakhstan||Absent, No presence record(s)||OIE (2009)|
|Kuwait||Absent, No presence record(s)||OIE (2009)|
|Kyrgyzstan||Absent, No presence record(s)||OIE (2009)|
|Malaysia||Absent, No presence record(s)||OIE (2009)|
|-Peninsular Malaysia||Absent, No presence record(s)||OIE Handistatus (2005)|
|North Korea||Absent, No presence record(s)||OIE Handistatus (2005)|
|Singapore||Absent, No presence record(s)||OIE (2009)|
|Taiwan||Absent, No presence record(s)||OIE Handistatus (2005)|
|Turkmenistan||Absent, No presence record(s)||OIE Handistatus (2005)|
|Uzbekistan||Absent, No presence record(s)||OIE Handistatus (2005)|
|Andorra||Absent, No presence record(s)||OIE Handistatus (2005)|
|Belarus||Absent, No presence record(s)||OIE (2009)|
|Bosnia and Herzegovina||Absent, No presence record(s)||Fijan (1999); OIE Handistatus (2005)|
|Croatia||Absent, No presence record(s)||OIE (2009)|
|Cyprus||Absent, No presence record(s)||OIE (2009)|
|Denmark||Absent, No presence record(s)||OIE (2009)|
|Finland||Absent, No presence record(s)||OIE (2009)|
|Germany||Absent, No presence record(s)||2005||OIE (2009); Bachmann and Ahne (1973)||Last reported: 200512|
|Hungary||Absent, No presence record(s)||2006||OIE (2009); Békési and Szabó (1977); Békési and Szabó (1979)|
|Iceland||Absent, No presence record(s)||OIE (2009)|
|Ireland||Absent, No presence record(s)||OIE (2009)|
|Isle of Man||Absent, No presence record(s)||OIE Handistatus (2005)|
|Italy||Absent, No presence record(s)||2004||OIE (2009); Ghittino et al. (1980)|
|Jersey||Absent, No presence record(s)||OIE Handistatus (2005)|
|Latvia||Absent, No presence record(s)||OIE (2009)|
|Lithuania||Absent, No presence record(s)||OIE (2009)|
|Moldova||Present||CABI (Undated)||Original citation: USA, Center for Food Security & Public Health and USA, Institute for International Cooperation in Animal Biologics (2007)|
|Netherlands||Absent, No presence record(s)||OIE (2009)|
|Norway||Absent, No presence record(s)||OIE (2009)|
|Poland||Present||OIE (2009); Fijan (1999); Antychowicz and Kozińska (2000)|
|Portugal||Absent, No presence record(s)||OIE (2009)|
|Romania||Absent, No presence record(s)||OIE (2009); Fijan (1999)|
|Serbia||Absent, No presence record(s)||OIE (2009)|
|Slovakia||Absent, No presence record(s)||1999||OIE (2009); Kočiš et al. (1983)|
|Slovenia||Absent, No presence record(s)||OIE (2009)|
|Sweden||Absent, No presence record(s)||OIE (2009)|
|Switzerland||Absent, No presence record(s)||2003||OIE (2009); Bernet and Wahli (2002)||Last reported: 200305|
|Ukraine||Absent, No presence record(s)||OIE (2009); CABI (Undated)|
|United Kingdom||Absent, No presence record(s)||2008||OIE (2009); Bucke and Finlay (1979)||Last reported: 200803|
|-Northern Ireland||Absent, No presence record(s)||OIE Handistatus (2005)|
|Barbados||Absent, No presence record(s)||OIE Handistatus (2005)|
|Belize||Absent, No presence record(s)||OIE (2009)|
|Bermuda||Absent, No presence record(s)||OIE Handistatus (2005)|
|British Virgin Islands||Absent, No presence record(s)||OIE Handistatus (2005)|
|Canada||Absent, Unconfirmed presence record(s)||OIE (2009)|
|Cayman Islands||Absent, No presence record(s)||OIE Handistatus (2005)|
|Costa Rica||Absent, No presence record(s)||OIE (2009)|
|Cuba||Absent, No presence record(s)||OIE (2009)|
|Dominica||Absent, No presence record(s)||OIE Handistatus (2005)|
|Dominican Republic||Absent, No presence record(s)||OIE Handistatus (2005)|
|Greenland||Absent, No presence record(s)||OIE (2009)|
|Guatemala||Absent, No presence record(s)||OIE (2009)|
|Jamaica||Absent, No presence record(s)||OIE (2009)|
|Mexico||Absent, No presence record(s)||OIE (2009)|
|Nicaragua||Absent, No presence record(s)||OIE (2009)|
|Saint Kitts and Nevis||Absent, No presence record(s)||OIE Handistatus (2005)|
|Saint Vincent and the Grenadines||Absent, No presence record(s)||OIE Handistatus (2005)|
|Trinidad and Tobago||Absent, No presence record(s)||OIE Handistatus (2005)|
|United States||Present, Localized||OIE (2009); Goodwin (2002); Dikkeboom et al. (2004)|
|-North Carolina||Present, Localized||Goodwin (2002); Dikkeboom et al. (2004)|
|-Wisconsin||Present, Localized||Dikkeboom et al. (2004)|
|Australia||Absent, No presence record(s)||OIE (2009)|
|French Polynesia||Absent, No presence record(s)||OIE (2009)|
|New Caledonia||Absent, No presence record(s)||OIE (2009)|
|New Zealand||Absent, No presence record(s)||OIE (2009)|
|Vanuatu||Absent, No presence record(s)||OIE Handistatus (2005)|
|Argentina||Absent, No presence record(s)||OIE (2009)|
|Brazil||Present||CABI (Undated a)||Present based on regional distribution|
|-Sao Paulo||Present, Localized||Alexandrino et al. (1998)|
|Chile||Absent, No presence record(s)||OIE (2009)|
|Colombia||Absent, No presence record(s)||OIE (2009)|
|Falkland Islands||Absent, No presence record(s)||OIE Handistatus (2005)|
|French Guiana||Absent, No presence record(s)||OIE (2009)|
|Guyana||Absent, No presence record(s)||OIE Handistatus (2005)|
|Paraguay||Absent, No presence record(s)||OIE Handistatus (2005)|
|Uruguay||Absent, No presence record(s)||OIE (2009)|
|Venezuela||Absent, No presence record(s)||OIE (2009)|
PathologyTop of page
Histopathological changes in some organs of experimentally infected carp were described by Negele (1977). Blood vessels in the liver show a varying degree of oedematous perivasculitis, which leads to necrosis. The liver parenchyma is hyperaemic, with multiple focal necroses and degeneration. The pancreas is inflamed and shows multifocal necrosis. The hyperaemic spleen shows hyperplasia of the reticuloendothelium and enlarged melanomacrophage centres. In the kidney, both the excretory and haematopoietic tissue is damaged. Tubules are clogged with casts and the cells undergo hyaline degeneration and vacuolation. The peritoneum is inflamed and the lymph vessels are filled with detritus and macrophages. Changes in the intestine are dominated by perivascular inflammation, desquamation of the epithelium and atrophy of the villi. In the swim-bladder, the epithelial lamina changes from a monolayer into a discontinuous multilayer; vessels in its submucosa are dilated, while lymphocytes infiltrate nearby areas. The heart is affected by pericarditis and by infiltration of the myocardium, which is followed by focal degeneration and necrosis. Osadcaja and Rudenko (1981) found similar changes in both natural and experimentally induced SVC. They also noted encephalitis, haemorrhage and inflammation in the spleen, as well as acute catarrhal enteritis, with necrosis and desquamation of the epithelium. Sulimanovic et al. (1986) encountered two types of severe changes in the premortal stage of experimentally infected carp: acute, necrotic alterations with weak inflammatory reactions or prolonged host response with hydrops and pronounced inflammation
DiagnosisTop of page
Mortality of fingerlings and older fish can be expected at temperatures below 18-20°C, i.e. in the spring and at the beginning of the summer, but less frequently in the autumn and winter.
Clinical diagnosis is based on behavioural and external and internal signs. Haemorrhage in the skin, pale gills, ascites and protruding vent, as well as enteritis, peritonitis, oedema and varying degrees of petechial haemorrhage in the swim-bladder, muscles and other organs, warrant sampling of fish for laboratory examination. The wide variation of disease signs among individual fish in a population or the simultaneous presence of fish with other disease signs will require the selection of separate sample groups. Diagnosis of SVC in clinically affected fish may be achieved by virus isolation or more rapidly by direct immunofluorescence tests (Faisal and Ahne, 1984) or ELISAs (Way, 1991; Rodák et al., 1993) on infected tissues. Ideally, direct diagnosis by IF or ELISA should be confirmed by virus isolation followed by a virus neutralisation test. However, virus isolation may not be possible from decomposed clinical samples (Way, 1991), so the presence of signs of SVC disease and a positive direct IF test or ELISA may be considered sufficient to initiate control measures.
At least ten diseased or moribund fish from each species or group should be selected for laboratory confirmation of SVCV in suspected outbreaks. Live or sacrificed specimens are transported under refrigeration or on ice but never frozen. The OIE Manual (2003) suggests the immediate collection and dispatch of all internal organs and the encephalon for fish of 4-6 cm or the kidney, spleen and encephalon for specimens above 6 cm. Such material from up to five fish is combined to form a tissue pool of 1-1.5 g. Each pool is placed into a sterile vial with a fivefold volume of transport medium containing antibiotics. Vials are transported and stored at 4°C. Virus extraction should be carried out within 24 h and not later than 48 h
Isolation of SVCV from diseased fish is readily accomplished by inoculation of susceptible, young (24 h) and active cell cultures with homogenized, decontaminated, clarified and diluted (three serial 10 × dilutions) organ samples. The OIE Manual (2003) recommends Epithelioma papulosum cyprini (EPC) cells for virus isolation. At least 2 cm2 of cell monolayer should be inoculated with 100 ml of each dilution. After adsorption for 0.5-1 h at 10-15°C, the inoculum is not withdrawn and the cell culture medium is added (buffered at pH 7.6 and supplemented with 2% fetal calf serum); pH is maintained at 7.3-7.6 for 7 days. Positive controls have to be prepared simultaneously. Cultures should be incubated at 15 (OIE Manual, 2003) or 20°C and monitored daily for 7 days. The cytopathic effect develops within 1-2 days but more time may be required. Initially, areas of cell rounding spread over the sheet and finally all cells are detached and lysed. Depending on the stage of disease, the virus concentration can vary from 102 to over 108 TCID50 g-1 tissue. If a sample remains negative for cytopathic effect for 7 days, the inoculated cell culture must be subcultured. Shchelkunov and Shchelkunova (1989) noted virus inhibitors in tissues with high amounts of SVCV, and these may cause absence of cytopathic effect when cultures are inoculated with low dilutions.
Identification of isolated virus is carried out using serological techniques, i.e. virus neutralization (Petrinec, 1973; de Kinkelin and Le Berre, 1974; Fijan, 1976; OIE Manual, 2003), IFAT (Faisal and Ahne, 1983, 1984; Rodak et al. 2003) or immunoperoxidase (Faisal and Ahne, 1980, 1983, 1984; Rodak et al., 1993). Virus neutralization is the preferred assay. The titre of neutralizing antibody solution to SVCV for the virus neutralization must be at least 2000 for 50% plaque reduction (OIE Manual, 2003). When a suspected SVCV isolate remains unaffected by neutralizing antibody to SVCV in a virus neutralization test, it is mandatory to conduct an IFAT (OIE Manual, 2003). Virus isolation accompanied by simultaneous virus neutralization of SVCV accelerates the obtaining of a diagnosis. Kits for IFAT and ELISA to identify SVCV in cell cultures are commercially available. Ariel and Olesen (2001) assessed an IFAT kit that used a monoclonal antibody against SVCV, and although the kit successfully detected all four SVCV isolates tested, it also detected one of three pike fry rhabdovirus (PFR) isolates tested. An SVCV isolate from Wisconsin, USA, was only weakly neutralized by SVCV antiserum, and did not react in an immunocytochemical test (Dikkeboom et al., 2004).
However, SVCV is a poor antigen for rabbits and none of the immunization programmes tested by Hill et al. (1981) gave entirely satisfactory and predictable results. Serum with the highest neutralizing antibody titre of 1:6500 was obtained by a single intramuscular (i.m.) inoculation of concentrated virus, combined with complete Freund's adjuvant, followed by two intravenous 'booster' injections. However, antisera from some rabbits had titres of less than 100. Similarly, Jørgensen et al. (1989) obtained relatively low (1:1280) but highly specific neutralizing antibody titres from rabbits immunized with purified SVCV after 22 weeks. Dixon and Hill (1984) could not develop a satisfactory ELISA for SVCV as a result of the low titre (1:3500) of antiserum. Way (1991) had good SVCV antibodies for ELISA by purification of gamma globulin from rabbit antiserum.
Spring viraemia of carp virus is serologically distinct from all other fish rhabdoviruses except other putative vesiculoviruses from fish as described by Stone et al. (2003) (see below). An IFAT cannot distinguish SVCV from PFR (Jørgensen et al., 1989, Ahne et al., 1998). Virus neutralization with selected antisera and ELISA are more specific. If complement is used in virus neutralization, the distinction between SVCV and PFR is impossible (Clerx et al., 1978; Jørgensen et al., 1989). Ahne (1986) isolated a virus from carp that was neutralized to a degree by both SVCV and PFR antisera, but which could not be identified with certainty as either virus. The virus was also less pathogenic for carp under experimental conditions; this, combined with the neutralization data, led the author to suggest that the isolate was a type intermediate between SVCV and PFR, or a sub-type of SVCV. Identification of SVCV antigen in tissues from moribund and dead infected carp is possible by IFAT (Faisal and Ahne, 1984) and by ELISA using tissue extracts in 1 h (Way, 1991; Rodak et al., 1993). The sensitivity of ELISA for detection of subclinical virus levels is lower than that of isolation in cell cultures, but it is superior for recognizing SVCV antigen in fish carcasses which undergo some degree of decomposition and loss of virus infectivity (Way, 1991; Rodak et al., 1993).
Identification of SVCV by molecular biology methods is more reliable than serology, and such methods have also been used for the direct detection of SVCV in infected fish. A ribonuclease protection assay has been used to differentiate SVCV and PFR (Ahne et al., 1998). A radioactively-labelled hybridization probe was constructed from minus-sense RNA transcripts of the G-gene, and annealed with target mRNA. The products were digested with RNAses A and T1, and the cleavage fragment patterns were analysed. RNA from isolates that were judged to be PFR from neutralization data was completely digested, but RNA from SVCV isolates protected the probe.
Stone et al. (2003) compared the partial nucleotide sequence of the G-gene of SVCV, PFR and other putative fish vesiculovirus isolates, and were able to allocate the isolates to one of four genogroups. Genogroup I comprised SVCV isolates, Genogroup II comprised a single isolate from grass carp, Genogroup III comprised the refrerence PFR isolate and Genogroup IV comprised isolates from a number of species, including isolates previously identified as PFR. Phylogenetic analyses of the SVCV Genogroup divided it into four subgroups (a, b, c and d). The clustering of the isolates in the subgroups reflected to a degree their geographic origins: subgroup a contained isolates from Asia, subgroup b contained isolates from Moldova and Ukraine, subgroup c contained isolates from Ukraine and Russia, and subgroup d mainly contained isolates from Europe, but also included one isolate from Moldova and one from the Ukraine. The SVCV isolates from the USA were subsequently allocated to subgroup 1a (Dikkeboom et al., 2004). Genetic differences between SVCV isolates from Europe and Russia have also been reported (Oreshkova and Koutna cited by Einer-Jensen et al., 2002).
A reverse transcription polymerase chain reaction (RT-PCR) and nested PCR that amplified specific regions of the G gene were described by Koutná et al. (2003) The RT-PCR was specific for SVCV and did not detect PFR, nor certain other fish viruses. SVCV isolates from different countries were detected in cell cultures by the RT-PCR but it did not detect all those isolates in tissue homogenates from experimentally-infected fish. However, all isolates tested were detected by the nested PCR.
Detection of SVCV in cell cultures and infected fish tissues by hybridization with biotinilated DNA probes for M-gene or G-gene sequences has been reported (Oreshkova et al., 1995, 1999). However, as the M-gene probe also hybridized with cell-culture grown viral haemorrhagic septicaemia virus and an eel rhabdovirus (Oreshkova et al., 1995), its use for identification of SVCV is limited.
Non-clinical spring viraemia of carp virus carrier fish
Attempts at virus isolation from survivors of SVC outbreaks and suspected carrier fish have been mostly unsuccessful. Nevertheless, the OIE Manual (2003) specifies sampling and laboratory procedures in surveillance programmes for establishing the SVC-free status of carp production units. Encephalon of any size fish and/or ovarian fluid from brood fish at the time of spawning have to be sampled from a number of fish, specified by Ossiander and Wedemeyer (1973), to detect a prevalence of infection in the examined population equal to or higher than 2% at a 95% confidence level. Samples are processed for virus isolation and identification, as specified above for overtly infected fish.
Cases of isolation of virus from subclinical SVC have been reported. Békési and Csontos (1985) examined reproductive products of carp brood stock that was induced to spawn by hypophysation, and recovered virus from 3 of 491 samples of ovarian fluid. All 211 samples of seminal fluid were virologically negative. Fijan (1988) reported negative findings in 86 samples of seminal and ovarian fluids, and Ahne (1983) could not find virus in seminal fluid of experimentally infected carp. Wolf (1988) recommended using immunosuppression or low-temperature stress on brood stock and subsequent examination for the presence of virus in ovarian fluid and urine and in leucocytes by cocultivation. SVCV was isolated in China from common and koi carp exhibiting no signs of disease (Liu et al., 2004). Gill biopsy was recommended by Baudouy et al. (1980c) for screening carp for SVCV at temperatures below 11°C.
Previous contact of a fish with SVCV and the presence of virus in a fish population can sometimes be detected indirectly by demonstration of virus-specific neutralizing antibodies in the serum. This approach is not currently accepted as a routine diagnostic method, as a result of insufficient knowledge of the serology of virus infections in fishes (OIE Manual, 2003). However, the validation of serological techniques for certain fish virus infections could arise in the near future, rendering the use of fish serology more widely acceptable for health screening purposes. Circulating antibodies to SVCV in clinically healthy and virologically negative carp survivors were first found in Croatia by an indirect haemagglutination test (Sulimanovic, 1973) and by virus neutralization (Petrinec, 1973). As already mentioned, examination of carp sera for neutralizing antibodies was also used in epizootiological studies on farms in several other European countries (Dangschat, 1979; Hoppe and Wernery, 1985; Kölbl and Kainz, 1977; Wizigmann et al., 1980, 1983). Kretschmar and Dresenkampf (1987) applied a solid-phase ELISA for detecting anti-SVCV antibodies. The method was 100 times more sensitive than virus neutralization in detecting antibodies in hyperimmune carp sera, but detected a lower percentage of antibody-positive carp than virus neutralization in surveys of suspected populations. A similar ELISA method has also been described by Rodák et al. (1986). The competitive immunoassay (Dixon et al., 1994) was more sensitive than virus neutralization, and detected more antibody-positive fish in naturally exposed carp populations.
List of Symptoms/SignsTop of page
|Finfish / Darkened coloration - Skin and Fins||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / 'Dropsy' - distended abdomen, 'pot belly' appearance - Body||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Fish sinking to bottom - Behavioural Signs||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Generalised lethargy - Behavioural Signs||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Haemorrhaging - Body Cavity and Muscle||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Loss of balance - Behavioural Signs||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Mortalities -Miscellaneous||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Paleness - Gills||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Pop-eye - Eyes||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Red spots - Gills||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Red spots: pin-point size (petechiae) - Skin and Fins||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Splenomegaly - spleen swelling / oedema - Organs||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Swelling - Organs||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / Trailing, flocculent or mucous faeces (casts) - Body||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
|Finfish / White-grey patches (necrosis / tissue damage) - Organs||Aquatic:Adult,Aquatic:Broodstock,Aquatic:Larval,Aquatic:Fry||Sign|
Disease CourseTop of page
Multiplication of virus in capillary endothelium, as well as in haematopoietic and excretory kidney tissues, causes an impaired salt-water balance, which is often lethal. Characteristic macroscopic lesions include oedema, haemorrhage, anaemia, enteritis and peritonitis. Mixed and secondary infections can lead to additional clinical signs and increased mortality. The disease signs outlined below are based on uncomplicated natural SVC outbreaks and experimental infections in the laboratory (Fijan et al., 1971; Bachmann and Ahne, 1973, 1974; Fijan, 1972, 1973, 1975).
Fish in the initial stages of disease congregate where there is a slow flow of water and near pond banks or lie on the bottom. The rate of respiration, reaction to sensory stimulation and swimming speed are progressively reduced. Terminally, the fish are sluggish, swim on their side, move slowly and aimlessly or rest in a normal or abnormal position.
External signs of SVC include skin darkening, swollen belly, exophthalmia, petechial haemorrhage and ecchymoses in the skin, gills and anterior eye chamber, anaemia and pale gills, as well as protrusion and inflammation of the vent. Shedding of long, white to yellowish and thick mucoid casts from the vent is readily visible in fish kept in experimental facilities. Internal signs are dominated by oedema in all organs, haemorrhage, peritonitis and catarrhal enteritis. Small to large quantities of a transparent fluid in the peritoneal cavity may be tinged with blood. Fibrinous peritonitis causes adhesions of organs to each other and to the parietal peritoneum. Petechial haemorrhage is evident in internal organs and muscles. Haemorrhage is easily recognizable in the lamina epithelialis of the swim-bladder. The intestine is void of food and appears stretched from the abundant mucus. The spleen is enlarged and sometimes affected by haemorrhagic or ischaemic infarcts.
Pathogenesis and immunity
Uptake and multiplication of SVCV in carp was studied by Ahne (1977, 1978) at 13°C. After infection by immersion, the virus was detectable in gills up to 2 h and then on the fourth day, when it was also present in kidneys, liver, spleen and gut. Viraemia was evident on day 5 and reached 5 × 107 plaque-forming units (pfu) ml-1 on day 6. Virus was detected in faeces on day 11. At 11°C Baudouy et al. (1980c) found the virus in gills on day 2 and and viraemia by day 3. High virus titres in kidney, liver and spleen of naturally and experimentally infected carp (Fijan et al., 1971; Ahne, 1977, 1978) indicate that these organs are primary targets of virus replication. Twenty-four days after infection, kidneys have a much lower (or no) virus titre than liver and spleen (Faisal and Ahne, 1984).
Reduced haematocrit and haemoglobin demonstrated in experimentally infected sheatfish by Jeney et al. (1990) are a consequence of damage to the haematopoietic tissue. Necrosis at this and other sites is accompanied by increased blood transaminase activity.
The defence system is stimulated during SVCV infection, and encompasses neutralizing antibodies, resistance to reinfection (Fijan et al., 1977a,b; Hill, 1977; Ahne, 1980; Baudouy et al., 1980b) and interferon synthesis (Baudouy et al., 1977; de Kinkelin et al., 1982). Nothing is known about the cellular immune response that is probably involved in protective immunity in fish in the absence of neutralizing antibodies.
Temperature has a decisive influence on carp-SVCV interactions. The virus replicates in vitro at a wide temperature range, with an optimum at 20-22°C, but the defence system of immunologically mature fish restricts this activity in vivo to suboptimal temperature. Viraemia, detectable virus shedding and mortality occur mainly below 15°C, when carp are not capable of rapid interferon and neutralizing antibody synthesis (Fijan, 1988).
After intraperitoneal (i.p.) injection of the virus, the incubation period and survival times at 16-17°C are 3-7 days and 8-10 days, respectively (Fijan et al., 1971). Incubation after infection by immersion lasts 7-15 days at 16-17°C, and is much longer below 10°C (Baudouy et al., 1980c). Production of antibodies against SVCV is influenced by the age and condition of the fish, route of infection and, most importantly, temperature. Intraperitoneally injected adult carp produce neutralizing antibodies slowly and unevenly at about 14°C, but at 25°C production is more consistent, faster and with higher titres (Fijan et al., 1977a). Most fish that survive infection at 13°C have peak neutralizing antibody titres after 2-2.5 months, which decline at 4 months (Baudouy, 1978). Ahne (1980) found neutralizing antibodies in carp infected by immersion after 7 days at 20°C and after 7 weeks at 13°C. However, Hill (1977) did not locate measurable neutralizing antibodies in fish kept at 15°C 10 weeks after exposure by immersion, while Fijan et al. (1977b) detected neutralizing antibodies in only 7.5% of 118 sera collected 1 and 2 months after oral application of a live vaccine. In both cases the serologically negative carp were refractory to i.p. challenge with virus, indicating that specific resistance was mediated by other mechanisms. Infection of carp by the Asian tapeworm Bothriocephalus gowkongensis or blood flagellates and other parasites may hamper the development of protective immunity against SVCV (Kölbl, 1980; N. Fijan, unpublished data).
EpidemiologyTop of page
Spring viraemia of carp is generally transmitted horizontally. Overtly infected fish shed virus in faecal casts and possibly in urine and gill mucus (Pfeil-Putzien, 1977; Ahne, 1978; Pfeil-Putzien and Baath, 1978; Baudouy et al., 1980a,b,c). Demonstration of a persistent viraemic phase in experimentally infected carp kept at temperatures below 13-14°C (Ahne, 1977, 1978, 1980; Baudouy et al., 1980b,c) before the appearance of visible symptoms (Baudouy et al., 1980b,c) suggests that SVCV shedding by a few fish during winter is the most important factor in transmission.
Virus shedding by survivors of infection has not been demonstrated but appears to be important for survival of virus in fish populations. The virus probably circulates from farm to open water and back by small wild fish and escaping cultivated fish, as well as by water draining and filling ponds (Fijan, 1984).
Bloodsucking parasites, such as the carp louse, Argulus foliaceus, and the leech, Piscicola geometra, are vectors and can transfer the virus from infected to healthy carp (Pfeil-Putzien, 1977; Ahne, 1985a). They do not support replication or long-term survival of the virus (Ahne, 1985a). Fish-eating birds can probably carry the virus. Regurgitated infected fish from heron (Ardea cinerea) fed SVCV contained virus up to 120 min after feeding (Peters and Neukirch, 1986). The fate of virus in digestive systems of other fish-eating birds and of turtles eating fish carcasses has not been investigated. Under experimental conditions SVCV could be transmitted to pike by predation (Ahne, 1985b). There is concern that SVCV could be spread in the USA by baitfish such as the golden shiner and fathead minnow (Pimephales promelas) (Goodwin et al., 2004), particularly as the former species is experimentally susceptible to the virus (Goodwin, 2002), and part of the wild-caught baitfish business is located in an SVC-positive area. However, SVCV was not isolated from baitfish during an extensive virus-testing programme either prior to or following the isolation of SVCV from carp in the USA (Goodwin et al., 2004).
Since SVCV retains infectivity for a long time in water or mud or in a dry state (Ahne, 1982a,b), the pond environment and the fishery equipment on SVCV-positive farms should be regarded as infected with the virus.
Gills are the natural portal of virus entry and primary replication. Fish can be experimentally infected using a virus bath suspension. Higher concentrations of virus in the bath result in higher mortality (Baudouy et al., 1977). Entry by the intestinal route seems to be of minor importance. The application of virus directly into the anterior gut did not induce disease (Varovic and Fijan, 1973) and the bile rapidly inactivated virus in vitro (N. Fijan, unpublished data).
For experimental purposes, SVC can be readily induced by immersion and intraperitoneal (i.p.) injection. Ahne (1977, 1978) found that i.p. infection causes a higher mortality rate (90%) than immersion of carp in the virus (20%) at 13°C. Intracerebral and intrapneumatic (by the swim-bladder) injection of virus also induces disease but less frequently than i.p. injection (Varovic and Fijan, 1973).
Vertical transmission of SVCV seems possible. Békési and Csontos (1985) found virus in about 0.6% of the examined ovarian fluids. However, the scarcity of natural SVC outbreaks among fry and fingerlings indicates that vertical transmission is of minor importance.
Impact SummaryTop of page
|Fisheries / aquaculture||Negative|
Impact: EconomicTop of page
Spring viraemia of carp virus can induce serious mortality and economic losses, which fluctuate from year to year. These irregularities in epizootic patterns are poorly understood. Mortality is usually around 30%, but may exceed 70%. Most outbreaks occur after stocking of ponds in the spring. Ghittino et al. (1980b) estimated the loss of 1-year-old carp in Europe to be 4000 tons annually, i.e. 10-15% of this age-group. The loss is more disastrous when 2-year-old carp are used for stocking. The need for restocking increases the cost of production (cost of additional fingerlings, increased labour).
Zoonoses and Food SafetyTop of page
This species is not a zoonosis.
ReferencesTop of page
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