infectious haematopoietic necrosis
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IdentityTop of page
Preferred Scientific Name
- infectious haematopoietic necrosis
International Common Names
- English: infectious hematopoietic necrosis
Local Common Names
- USA: Chinook salmon virus disease; Coleman disease; Columbia River sockeye disease; Cultus lake virus disease; Oregon sockeye virus; Sacramento River Chinook disease; sockeye salmon virus diease
OverviewTop of page
Infectious haematopoietic necrosis (IHN) is an infectious disease of Pacific salmonid fish including rainbow or steelhead trout (Oncorhynchus mykiss), Chinook (O. tshawytscha), sockeye (O. nerka), chum (O. keta), masou (O. masou), and coho (O. kisutch), and Atlantic salmon (Salmo salar). The original geographical range of IHN was limited to the western parts of North America, but the disease has spread to continental Europe and Asia via the importation of infected fish and eggs. For more detailed reviews of the disease, see Bootland & Leong (1999), Pilcher and Fryer (1980), and Wolf (1988).
The principal clinical and economic consequences of IHN occur at farms rearing rainbow trout in freshwater; however, both Pacific and Atlantic salmon reared in freshwater or seawater can be severely affected. Large mortalities have also been recorded among some wild stocks of Pacific salmon. Infection is often lethal due to osmotic imbalance leading to oedema and haemorrhage. The causative agent, the rhabdovirus IHNV, multiplies in the endothelial cells of blood capillaries, haematopoietic tissues and nephron cells. High levels of virus are shed from infected juvenile fish. Older fish are increasingly resistant to infection, but adult fish at spawning may shed virus in sexual products. Survivors of IHNV infection demonstrate a strong protective immunity with the synthesis of circulating antibodies to the virus (LaPatra et al., 1993a) and, in certain individuals, a covert carrier state (Drolet et al., 1995; Kim et al, 1999).
On the basis of antigenic studies conducted with neutralising polyclonal rabbit antisera, IHNV isolates form a single serogroup (Engelking et al., 1991a). However, mouse monoclonal antibodies have revealed a number of neutralising epitopes on the glycoprotein (Huang et al., 1994; Ristow and Arnzen De Avila, 1991; Winton et al., 1988; Xu et al.,1991), as well as the existence of a non-neutralising, group epitope borne by the nucleoprotein (Ristow and Arnzen, 1989). Variations in the virulence of IHNV strains have been recorded during both natural cases of disease and in experimental infections (LaPatra et al., 1993a).
The reservoirs of IHNV are clinically infected fish and covert carriers among cultured, feral or wild fish. Virus is shed via faeces, urine, sexual fluids and external mucus, whereas kidney, spleen, encephalon and the digestive tract are the sites in which virus is most abundant during the course of overt infection. The transmission of IHNV between fish is primarily horizontal; however, cases of vertical or ‘egg-associated’ transmission have been recorded. Horizontal transmission is typically by direct exposure, but invertebrate vectors have been proposed to play a role in some cases. Egg-associated transmission is significantly reduced by the now common practice of surface disinfection of eggs with an iodophor solution, but is the only mechanism accounting for the appearance of IHN in new geographical locations among alevins originating from eggs that were incubated and hatched in virus-free water. Once IHNV is established in a farmed stock or in a watershed, the disease may become established among carrier fish.
Among individuals of each fish species, there is a high degree of variation in susceptibility to IHNV. The age of the fish appears to be extremely important: the younger the fish, the more susceptible to disease. As with viral haemorrhagic septicaemia virus, good overall fish health condition seems to decrease the susceptibility to overt IHN, while co-infections with bacterial diseases (e.g. bacterial coldwater disease), handling and other types of stress frequently cause subclinical infections to become overt.
The most prominent environmental factor affecting IHN is water temperature. Clinical disease occurs between 8°C and 15°C under natural conditions.
The screening procedure for IHNV is based on virus isolation in cell culture. Confirmatory identification may be achieved by use of immunological (neutralisation, indirect fluorescent antibody test or enzyme-linked immunosorbent assay), or molecular (DNA probe or polymerase chain reaction) methods (Arakawa et al., 1990; Arnzen et al., 1991; Deering et al., 1991; Dixon and Hill, 1984; Jorgensen et al., 1991; LaPatra et al., 1989; Winton and Einer-Jensen, 2002).
Control methods for IHN currently rely on avoidance of exposure to the virus through the implementation of strict control policies and sound hygiene practices (Winton, 1991). The thorough disinfection of fertilised eggs and the incubation of eggs and rearing of fry and alevins on virus-free water supplies in premises completely separated from those harbouring possible virus carriers and free from possible contact with inanimate objects, are critical for preventing the occurrence of IHNV in a defined fish production site. At present, vaccination is at an experimental stage; however, several new vaccine preparations have shown substantial promise in both laboratory and field trials (Winton, 1997).
[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.]
Hosts/Species AffectedTop of page
Infectious haematopoietic necrosis was first observed in cultured sockeye salmon (Oncorhynchus nerka) on the west coast of North America (Rucker et al., 1953; Watson et al., 1954) and IHNV was first isolated in cell culture by Wingfield et al. (1969). Initially, the host range of IHNV was thought to be limited to the genus Oncorhynchus. Infectious haematopoietic necrosis is often reported in rainbow trout, in steelhead trout and in the Pacific salmon - sockeye, kokanee, Chinook and chum salmon (Wolf, 1988). However, in the last 20 years, IHNV has been isolated from several new fish species, and the host range of the virus has enlarged. Cutthroat trout (O. clarki) has been found to be susceptible to IHNV infection and disease (Parisot, 1962; Groberg, 1983). Caution must be used before concluding whether a fish species is susceptible to IHNV infection and mortality. There are several examples where a fish species was not considered susceptible to IHNV, but later studies demonstrated susceptibility under different experimental conditions. For example, the coho salmon (O. kisutch) and pink salmon (O. gorbuscha) are considered to be refractory to IHN. No natural epizootics have been reported and experimental infection with a high viral dose resulted in only low coho fry mortalities (Wingfield and Chan, 1970; Wingfield et al., 1970; Chen et al., 1990) and no pink salmon fry mortalities (Follett et al., 1997). However, coho salmon are susceptible to IHNV infection - virus has been isolated from adult fish (LaPatra et al., 1989b; Eaton et al., 1991) and from fry naturally or experimentally exposed to IHNV (Hedrick et al., 1987; LaPatra et al., 1989b; Chen et al., 1990).
Several species of Salmo have been shown to be susceptible to IHNV infection and disease. Atlantic salmon (Salmo salar) fry have suffered at least one epizootic in North America (Mulcahy and Wood, 1986), and sporadic disease outbreaks have occurred in Japan (Yamazaki and Motonishi, 1992). Traxler et al. (1991, 1993) have demonstrated that Atlantic salmon can be infected with IHNV in salt water by immersion, by injection or, most importantly, by cohabitation with infected sockeye salmon. Recently, natural IHNV infections have been diagnosed in Atlantic salmon adults during the marine phase of their life cycle (Traxler et al., 1997). Brown trout (S. trutta) are moderately susceptible to experimentally induced IHN (LaPatra and Fryer, 1990), but natural infections and disease outbreaks have occurred in Oregon (Engelking and Kaufman, 1994a,b) and occasional IHN outbreaks have been reported in Japan (Yamazaki and Motonishi, 1992). In France, brown trout may be refractory to IHN (Hill, 1992). Since brown trout and Atlantic salmon are important species in aquaculture and sports fisheries in several countries, and Atlantic salmon is also a valuable commercial species, the potential for IHNV to cause devastating disease in these two species is of serious concern (Hill, 1992).
Salvelinus spp. vary in their susceptibility to IHNV. Lake trout (S. namaycush) can be experimentally infected but appear to be relatively resistant to disease (Yamamoto and Clermont, 1990) and Arctic char (S. alpinus) may be completely resistant to IHNV (T.R. Meyers as cited in LaPatra et al., 1992; Follett et al., 1997). Brook trout (S. fontinalis) have had sporadic outbreaks of IHN in Japan (Yamazaki and Motonishi, 1992) and the virus has been isolated from apparently healthy adult brook trout in Oregon (Pilcher and Fryer, 1980a,b). The success of experimentally infecting brook trout fry with the induction of disease is variable and depends on the water temperature, viral isolate and viral dose. Yamamoto and Clermont (1990) stated that 1-month-old brook trout were infected after immersion in IHNV, but unfortunately no data were presented. In a second study, brook trout fry appeared to be completely resistant to infection after immersion in a type 2 Idaho IHNV isolate (LaPatra et al., 1992). The 15°C water temperature used in the latter study may have enhanced resistance of the brook trout to disease since several researchers have shown that temperatures above the optimum for IHN epizootics (10-12°C) resulted in decreased fish mortalities (Watson et al., 1954; Amend, 1970a). Goldes and Mead (1992) found that 2-month-old brook trout immersed at 4°C in a type 1 IHNV isolate had a combined morbidity-mortality of 6.7% and 93% of the dead fish were infected. The virus was isolated from surviving fish 2 months postimmersion. A higher mortality (35%) occurred in brook trout fry immersed at 12°C in a type 1 IHNV isolate and the virus was isolated for 3 weeks postimmersion (Bootland et al., 1994). A similar immersion of brook trout in a type 2 IHNV resulted in only 5% mortality. Thus, brook trout fry are at least moderately susceptible to IHN disease and infection, but the viral isolate and dose influence the degree of susceptibility. The studies with brook trout clearly demonstrate that, when examining the susceptibility of a fish species to IHNV, it is important to test several viral doses and isolates at the optimal water temperature for IHN disease. Resistance to one IHNV isolate does not guarantee that the fish will not be susceptible to other IHNV isolates.
The susceptibility of non-salmonid fishes to IHNV has not been studied in depth. Castric and Jeffroy (1991) found two marine fish species, sea bream (Sparus aurata) and turbot (Scophthalmus maximus), suffered mortalities of 43% and 87%, respectively, after intraperitoneal (i.p.) injection of 106 plaque-forming units (pfu) of a French IHNV isolate. Sea bass (Morone labrax [Dicentrarchus labrax]) were refractory to disease but were infected. LaPatra et al. (1995) examined the susceptibility of white sturgeon (Acipenser transmontanus) cell lines, larvae, juveniles and adults to IHNV. Only one of three cell lines was susceptible, and the viral titre was only 17% of that produced in Chinook salmon embryo (CHSE-214) cells. Infectious haematopoietic necrosis virus replicates in larval fish for at least 9 days, but juveniles fed infected rainbow trout or immersed in virus are refractory to mortality and infection. Adult fish that cohabit with infected rainbow trout are not infected, but there are neutralizing antibodies. It was suggested that sturgeon should be considered as potential sources of virus (LaPatra et al., 1995). Michel Dorson and his colleagues tested the susceptibility of 1-2 month old pike fry (Esox lucius) to IHNV and found haemorrhage with frequent exophthalmia with mortality (Dorson et al., 1987).
In Oregon, adult northern squawfish (Ptychocheilus oregonensis), adult largescale suckers (Catostomus columbianus) and lamprey (Entosphenus tridentatus) ammocoetes appear to be refractory to disease and infection after injection of a type 1 IHNV isolate. However, adult mountain whitefish (Prosopium williamsoni), a member of the Salmonidae family, can be infected by injection and the virus persists for at least 3 weeks (L.M. Bootland, H.V. Lorz and J.C. Leong, unpublished data). A cell line derived from embryonic tissue of the inconnu or sheefish (Stenodus leucichthys), a member of the Salmonidae, is susceptible to IHNV (Follett and Schmitt, 1990), but it appears that in vivo susceptibility tests have not been done. Arctic grayling (Thymallus arcticus) fry are refractory to infection with a type 1 IHNV isolate, and this species is not likely to be a reservoir of IHNV (Follett et al., 1997). In wild fish surveys, IHNV has been isolated from Pacific herring (Clupea pallasi), shiner perch (Cymatogasteraggregata), and tubesnout (Aulorhynchus flavidus) off the cost of British Columbia (Kent et al., 1998).
Infectious haematopoietic necrosis virus has been isolated from a few invertebrates. The salmon leech (Piscicola salmositica) (Mulcahy, 1986; Yamamoto et al., 1989; Mulcahy et al., 1990) and ectoparasitic copepods (Salmincola sp.) removed from sockeye salmon (Mulcahy et al., 1990) harbour IHNV. The third invertebrate from which IHNV has been isolated is the common mayfly (Callibaetis sp.). Virus was detected in adult mayflies in Idaho by growth in cell culture after two blind passages and by Western blots (Shors and Winston, 1989b). Invertebrate species may play a role in the life cycle of IHNV by acting as vectors or reservoirs of infection, but this has to be clarified.
Factors affecting transmission and virulence
Fish typically become more resistant to IHNV as they increase in age and weight (Amend, 1974; Amend and Nelson, 1977; Leong and Turner, 1979; Wolf, 1988; LaPatra et al. 1994a). Yolk-sac fry and fish up to 2 months of age are highly susceptible, with mortality often over 90%. Older fish up to 6 months of age typically have less than 50% mortality. After experimental IHNV exposures, fry mortality generally increases as the viral dose increases from 102 to 105-6 pfu ml-l (Chen et al., 1990; LaPatra et al., 1990a, 1993a). Infectious haematopoietic necrosis virus can kill juvenile to 2-year-old sockeye salmon (Yasutake, 1978; Burke and Grischkowsky, 1984), kokanee salmon (Traxler, 1986; Banner et al., 1991) and rainbow trout (Busch, 1983; Roberts, 1986), but mortality is often very low.
Although the viral electropherotype and the host fish species are not definite indicators of IHNV virulence (LaPatra et al., 1989a, 1990a; Chen et al., 1990; Traxler et al., 1993). The practice of using electropheroptyes to distinguish different virus isolates was first introduced by Hsu et al. (1986). Based on the electrophoretic migration pattern of the viral glycoprotein and nucleoproteins in polyacrylamide gels, several investigators observed some correlation between electropherotype and host range. In general, type 1 IHNV isolates are most virulent in kokanee and sockeye salmon and type 2 isolates are more virulent in rainbow trout and steelhead trout than types 1 or 3 (LaPatra et al., 1990a,b, 1993a). For type 2 IHNV, susceptibility of kokanee and sockeye salmon increases, but susceptibility of rainbow trout decreases, with increasing fish age and weight (Amend and Nelson, 1977; LaPatra et al., 1990a, 1991a). Chinook salmon are more susceptible to type 3 isolates, but virulence of isolates within the same electropherotype is variable (Chen et al., 1990; LaPatra et al., 1993a). Viral virulence for a fish species appears to be dependent on the geographical location of virus isolation. Type 3 isolates from California and southern Oregon are more virulent for Chinook salmon than for steelhead trout, but type 3 isolates from the Columbia River are more virulent for steelhead trout than for Chinook salmon (LaPatra et al., 1993a). In addition to electropherotyping of IHNV, isolates can be divided into antigenic groups, based on reactivity with monoclonal antibodies (MAbs) (Winton et al., 1988; LaPatra et al., 1991a; Ristow and Arnzen-de Avila, 1991). LaPatra et al. (1994a) divided 106 type 2 isolates from the Hagerman Valley, Idaho, into ten antigenic groups. Virulence of isolates from the first seven antigenic types induced a wide range (14-92%) of mortality in rainbow trout. Forecasting the virulence of an isolate based on IHNV type or geographical location of isolation would not be completely accurate. Each IHNV isolate should be typed and evaluated for virulence in different fish species.
Fish genetics has a large influence on IHNV susceptibility. Wertheimer and Winton (1982) demonstrated that Chinook salmon fry from a Washington stock were more susceptible to two IHNV isolates than two stocks from Alaska. Amend and Nelson (1977) also noted distinct differences in IHNV susceptibility in families of sockeye salmon fry. In highly susceptible families, mortality was 98% or more, while in other families, the mortality was 52%. McIntyre and Amend (1978) found that the heritability of resistance to IHNV was about 30%. Family-level differences in IHNV susceptibility have also been observed in rainbow trout (Yamamoto et al., 1991; Kasai et al., 1993).
Interspecific hybrids have been made between resistant and susceptible species to enhance resistance to IHNV. Hybrids of coho salmon × rainbow trout (Parsons et al., 1986; Chen et al., 1990; LaPatra et al., 1993c), brook trout × rainbow trout (Dorson et al., 1991; LaPatra et al., 1993c) and cutthroat trout × rainbow trout (LaPatra et al., 1994b) have increased resistance to IHNV. However, it is also possible to obtain hybrids that have equal or higher susceptibility than the parental species. Such was the case for coho salmon × Chinook salmon (Hedrick et al., 1987). The most recent cross of brown trout females × lake trout males produced ‘brake trout’ and these triploid hybrid progeny were significantly more resistant to IHNV than rainbow trout after immersion or injection of IHNV (LaPatra et al., 1996). Brake trout immersed in IHNV had a weaker antibody response than rainbow trout, and it was postulated that protection was occurring at the cellular level. After i.p. injection of IHNV, the hybrids mounted a strong antibody response and had less extensive and severe necrosis of the haematopoietic tissue compared with rainbow trout. Hybrids generally have poorer performance in culture compared with parental species; however, hybrids can provide a model system for examining resistance mechanisms (LaPatra et al., 1996).
High fish density is correlated with outbreaks of IHN, possibly due to a rapid horizontal spread of the virus. The density of adult fish in a spawning channel may be a better predictor of outbreaks in fry than finding IHNV in adults (Traxler and Rankin, 1989). When artificial spawning beaches are stocked with two densities of sockeye salmon, IHN occurs only in the fry that have the higher density of adult spawners (Olson and Thomas, 1994). In addition to adult density, the egg density is also an important determinant of IHN outbreaks. Sockeye salmon eggs at a medium or high density in incubation boxes can result in IHN epizootics, but fry from the low-egg-density boxes are not infected (Mulcahy and Bauersfeld, 1983). In natural spawning, egg and alevin densities in the gravel are very low. There is one report of IHN epizootics in emergent sockeye salmon fry in Chilko Lake, British Columbia (Williams and Amend, 1976), which suggests that factors other than fish density may be involved. In Oregon and Washington, fry, smolts and adult salmonids are often moved around dams by truck or barge. The high fish density during transport would provide an excellent opportunity for horizontal virus transmission. There appears to be only one study where water in transport trucks was tested for IHNV. Using tangential flow filtration, an IHNV concentration of 1000 pfu ml-1 was found in the water from a truck transporting sockeye salmon fry (Batts and Winton, 1989b), compared with 20 pfu ml-1 in the spawning beach water in which the fry hatched (Olson and Thomas, 1994). These fry were infected with IHNV prior to shipping and, 10-14 days after shipping, fry mortality reached 100%. It is not certain whether fish density in the transport truck or IHNV in the water exacerbated fish mortality. Under experimental conditions, small differences in fry density do not affect IHN mortality. Groups of 25 or 50 rainbow trout fry immersed in IHNV did not have a significantly different mortality or mean day to death (LaPatra et al., 1991b). However, a wider range of fish densities must be evaluated to clarify the importance of fry density on mortality.
The most important environmental factor affecting IHNV virulence is temperature. Infectious haematopoietic necrosis epizootics most frequently occur during the spring and autumn when water temperatures are 10-12°C (Nicholson, 1982). Although it is generally accepted that disease outbreaks do not occur above 15°C, IHN killed rainbow trout fry from 3°C to 18°C. The mean day to death was directly related to temperature in that the higher the temperature, the shorter the survival of the fish (Hetrick et al., 1979a). In contrast, rainbow trout and sockeye salmon held at temperatures above 15.5°C before infection or moved to a higher temperature within 24 h after exposure had significantly reduced mortality. However, if virus was detectable in the fish, then a shift to a higher water temperature had little effect on mortality (Amend, 1970a, 1976). Results may have varied between studies because of differences in experimental design and fish size.
The nutritional status of the host and other environmental factors, such as water salinity and the presence of pollutants, may influence susceptibility to IHNV. Although the majority of IHN outbreaks occur in fresh water, IHN epizootics have been reported in Atlantic salmon held in ocean net pens (Traxler et al., 1991). Stress, caused either by physical handling or acute environmental changes, is known to decrease fish resistance to disease. Exposure of rainbow trout to copper increases IHN mortality. The copper may cause an increase in cortisol levels, and this in turn may cause a depression of the immune response (Hetrick et al., 1979b). In contrast, exposure of rainbow trout to polychlorinated biphenyls or 2,3,7,8-tetrachlorodibenzo-p-dioxin, which are immunosuppressive in mammals, did not significantly affect mortality or mean day to death due to IHNV (Spitsbergen et al., 1988).
One factor that does not appear to affect IHNV virulence is the cell line used for viral propagation. A type 2 IHNV isolate grown in rainbow trout gonad (RTG-2), Chinook salmon embryo (CHSE-214) or epithelioma papulosum cyprini (EPC) cells had equivalent virulence for rainbow trout (LaPatra et al., 1991b). It was also suggested that the IHNV glycoprotein is not responsible for virulence, because the nucleic acid sequence is highly conserved (LaPatra et al., 1993a). However, small differences in the amino acid sequence of the glycoprotein can affect virulence. Neutralization-resistant (escape) mutants, selected by incubation in neutralizing anti-glycoprotein MAb, often had decreased virulence and changes in tissue tropism in comparison with the homologous wild-type virus (Roberti et al., 1991; Kim et al., 1994). In addition to detecting differences in the glycoprotein, MAbs have detected heterogeneity in the nucleoprotein (Ristow and Arnzen, 1989; Ristow and de Avila, 1991). The relation between virulence and heterogeneity in the nucleoprotein has not been tested.
DistributionTop of page
Infectious haematopoietic necrosis virus primarily causes disease in the genus Oncorhynchus and was first identified in the Pacific North West of the USA. During the 1950s, IHNV caused severe losses of sockeye salmon (O. nerka) at hatcheries in Washington (Rucker et al., 1953) and Oregon (Wingfield et al., 1969). The virus might have been introduced into Oregon from Washington in unpasteurized sockeye salmon viscera that were fed to young fish, a practice that was soon stopped (Amend and Wood, 1972). A similar disease was next reported in hatchery-reared Chinook salmon (Oncorhynchus tshawytscha) in California (Ross et al., 1960; Wingfield and Chan, 1970). In 1967, IHN occurred in young sockeye salmon in British Columbia and was reported for the first time in rainbow trout (Amend et al., 1969).
With the realization that IHNV could devastate hatchery populations of young sockeye salmon, Chinook salmon and rainbow trout, several surveys were carried out to determine the distribution and prevalence of IHNV in the Pacific North West. Prior to the 1970s, IHNV was not a problem in Washington stocks of resident Chinook or coho salmon (O. kisutch), but two stocks of sockeye salmon were infected (Amend and Wood, 1972). During the 1970s, IHN epizootics increased in Oregon stocks of rainbow trout, steelhead trout (anadromous O. mykiss), Chinook salmon and kokanee salmon (land-locked O. nerka) (Groberg et al., 1980; Mulcahy et al., 1980). The source of the virus was not clear but it was suggested that the import of infected eggs from Washington and wild adult kokanee salmon or rainbow trout harbouring the virus might have introduced the virus into the area. In 1977, IHNV began causing severe losses of rainbow trout in Idaho (Busch, 1983). During the early 1980s, IHNV was isolated from additional salmonid species and fish losses due to IHN increased dramatically. By 1982, IHNV had been isolated in cutthroat trout (O. clarki) and there were very high IHN losses in rainbow and steelhead trout in the Columbia River basin (Groberg, 1983; Groberg and Fryer, 1983). The first reported natural IHN epizootic in Atlantic salmon fry occurred in 1984 in Washington (Mulcahy and Wood, 1986) and, in 1986, IHNV was isolated for the first time from adult chum salmon (O. keta) with no clinical disease (Hopper, 1987). In Idaho, IHNV became enzootic in rainbow trout (Busch, 1983) and was isolated from adult steelhead trout and kokanee salmon (Groberg, 1983). Currently, IHNV can be isolated from salmonid fish in Oregon, Washington, Idaho, California, Alaska and British Columbia and is considered endemic in the Pacific North West (Pilcher and Fryer, 1980a,b; Wolf, 1988).
In Alaska, IHNV began causing epizootics in sockeye salmon in 1973 and is now enzootic (Grischkowsky and Amend, 1976). Infectious haematopoietic necrosis was a major cause of failure in the culture of sockeye salmon until 1981, when a statewide policy devised by the Fisheries Rehabilitation, Enhancement and Development Division (FRED) proved successful in minimizing losses (Meyers et al., 1990). However, outbreaks of IHN are still occurring in enhanced populations of sockeye salmon smolts in at least one lake system (Follett and Burton, 1995). The virus has also been isolated in Alaska from diseased Chinook salmon and chum salmon (Follett et al., 1987).
Several other states have had sporadic IHN outbreaks in rainbow trout, and this was usually associated with the import of infected eggs or fry. The first epizootics outside the Pacific North West occurred in 1969 in Minnesota (Plumb, 1972) and South Dakota (Wolf et al., 1973). Disease outbreaks have also occurred in West Virginia (Wolf et al., 1973), Montana (Holway and Smith, 1973), New York (Carlisle et al., 1979), Colorado (Janeke, 1984) and Utah (Amos, 1985).
IHNV has spread by the movement of infected eggs and/or fry outside North America (Hill, 1992). In Japan, IHNV was apparently introduced in 1968 with eggs imported from Alaska (Sano et al., 1977) and yearly IHN outbreaks occur in nearly all areas that rear salmonid fish (Fukuda et al., 1992). Rainbow trout, yamame (O. masou), and amago trout (O. masou macrostomus) are most severely affected but sporadic outbreaks have been reported in sockeye and kokanee salmon, masu salmon (O. masou masou), chum salmon, cherry salmon (O. masou rhodurus), Atlantic salmon, brown trout (S. trutta), brook trout (S. fontinalis) and Japanese char (S. leucomaenis) (Kimura and Awakura, 1977; Sano et al., 1977; Yamazaki and Motonishi, 1992). In 1985, IHNV was spread to northeast China by importation of infected eggs from Japan (Niu and Zhao, 1988; Zhao and Niu, 1994). Importation of infected rainbow trout eggs into Italy resulted in the first diagnosis of IHNV in 1987 (Bovo et al., 1987) and IHNV has spread to at least six regions within the country (Bovo et al., 1991). Other countries in which IHNV has been identified include Taiwan (Chen et al., 1985; Wang et al., 1996), France (Laurencin, 1987; Hattenberger-Baudouy et al., 1989, 1995a,b), Belgium (Hill, 1992) and Korea (John et al., 1992; Park et al., 1993), The first confirmed finding of IHNV has also occurred in Russia (Shchelkunov et al, 2001) and the virus has also been found in Austria (Office International des Epizooties, 2004), the Czech Republic (Office International des Epizooties, 2004), Germany (Enzmann et al., 1992), and Switzerland (Office International des Epizooties, 2004).
It is likely that the geographical range of IHNV will continue to increase. The migration of infected anadromous fish species may spread IHNV (Hill 1992) and the movement of infected eggs and fish within and between countries will probably continue to occur, despite attempts in several countries to avoid this type of transfer by strict fish health certification requirements.
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|
|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||Absent, reported but not confirmed||NULL||Niu and Zhao , 1988; OIE, 2009|
|-Hong Kong||Disease never reported||OIE, 2009|
|Georgia (Republic of)||Disease never reported||OIE Handistatus, 2005|
|India||No information available||NULL||Office International des Epizooties, 2004; OIE, 2009|
|Indonesia||Disease not reported||OIE, 2009|
|Iran||Disease not reported||2007||Fallahi et al., 2003; OIE, 2009|
|Iraq||Disease never reported||OIE, 2009|
|Israel||Disease never reported||OIE, 2009|
|Japan||Present||NULL||Sano and et al. , 1977; 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||NULL||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||Chen and et al. , 1985; OIE Handistatus, 2005|
|Tajikistan||No information available||OIE, 2009|
|Thailand||No information available||OIE, 2009|
|Turkey||No information available||OIE, 2009|
|Turkmenistan||Disease not reported||OIE Handistatus, 2005|
|United Arab Emirates||No information available||OIE, 2009|
|Uzbekistan||Disease never 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||No information available||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||No information available||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||No information available||OIE Handistatus, 2005|
|Senegal||No information available||OIE, 2009|
|Seychelles||No information available||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||Absent, reported but not confirmed||OIE, 2009|
|-British Columbia||Amend and et al. , 1969|
|Greenland||Disease never reported||OIE, 2009|
|Mexico||Disease not reported||OIE, 2009|
|USA||Restricted distribution||OIE, 2009|
|-Alaska||Grischkowsky and Amend , 1976|
|-California||Ross and et al. , 1960|
|-Colorado||Janeke , 1984|
|-Minnesota||Plumb , 1972|
|-Montana||Holway and Smith , 1973|
|-New York||Carlisle and et al. , 1979|
|-Oregon||Wingfield and et al. , 1969|
|-South Dakota||Wolf and et al. , 1973|
|-Utah||Amos , 1985|
|-Washington||Rucker and et al. , 1953|
|-West Virginia||Wolf and et al. , 1973|
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||No information available||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||No information available||OIE, 2009|
|Nicaragua||No information available||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||Disease never reported||OIE, 2009|
|Albania||No information available||OIE, 2009|
|Andorra||Disease never reported||OIE Handistatus, 2005|
|Austria||Disease not reported||OIE, 2009|
|Belarus||Disease never reported||OIE, 2009|
|Belgium||No information available||NULL||Hill , 1992; OIE, 2009|
|Bosnia-Hercegovina||Disease not reported||OIE Handistatus, 2005|
|Bulgaria||No information available||OIE, 2009|
|Croatia||Disease not reported||OIE, 2009|
|Cyprus||Disease never reported||OIE, 2009|
|Czech Republic||Disease not reported||OIE, 2009|
|Denmark||Disease never reported||OIE, 2009|
|Estonia||No information available||OIE, 2009|
|Finland||Disease never reported||OIE, 2009|
|France||Disease not reported||200805||Laurencin , 1987; OIE, 2009|
|Germany||Present||NULL||Enzmann et al., 1992; OIE, 2009|
|Greece||No information available||OIE, 2009|
|Hungary||Disease never reported||OIE, 2009|
|Iceland||Disease never reported||OIE, 2009|
|Ireland||Disease never reported||OIE, 2009|
|Isle of Man (UK)||Disease never reported||OIE Handistatus, 2005|
|Italy||Present||NULL||Bovo and et al. , 1987; OIE, 2009|
|Jersey||Disease never reported||OIE Handistatus, 2005|
|Latvia||Disease never reported||OIE, 2009|
|Liechtenstein||No information available||OIE, 2009|
|Lithuania||Disease never 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|
|Norway||Disease never reported||OIE, 2009|
|Poland||Disease not reported||1994||Antychowicz et al., 2001; OIE, 2009|
|Portugal||Disease not reported||OIE, 2009|
|Romania||No information available||OIE, 2009|
|Russian Federation||No information available||OIE, 2009|
|Serbia||No information available||OIE, 2009|
|Slovakia||Disease not reported||OIE, 2009|
|Slovenia||Present||NULL||Office International des Epizooties, 2004; OIE, 2009|
|Spain||Disease not reported||OIE, 2009|
|Sweden||Disease never reported||OIE, 2009|
|Switzerland||Disease not reported||200407||Office International des Epizooties, 2004; OIE, 2009|
|UK||Disease not reported||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
Clinical signs and histopathology
The clinical signs and general histopathology of IHN in young salmonid fish are well documented (Amend et al.,1969; Amend, 1970b, 1974; Yasutake, 1970, 1975; Pilcher and Fryer, 1980a,b; Nicholson, 1982; Wolf, 1988). In acute disease there is a sudden increase in fish mortality, but the fish may not show clinical signs and may die without apparent cause. More typically, at the start of an epizootic, there are moribund fish that are lethargic, with periods of sporadic whirling or hyperactivity. Moribund fry also may have a dark coloration, a distended abdomen, exophthalmia, pale gills and mucoid, opaque faecal casts. Petechial haemorrhages may be observed at the base of the fins and vent and occasionally in the gills, mouth, eye, skin and muscle. In Chinook salmon, but not in sockeye salmon, the fry may have a subdermal haemorrhagic area immediately behind the head (Yasutake, 1970; Amend, 1974). Many of the above clinical signs are similar to those of infectious pancreatic necrosis (IPN) and viral haemorrhagic septicaemia (VHS). In older fish there are fewer external clinical signs. Two-year-old kokanee salmon have erratic swimming and haemorrhages near the base of the fins (Traxler, 1986), and sockeye salmon smolts have gill and eye haemorrhages, clubbed and fused lamellae and cutaneous lesions (Burke and Grischkowsky, 1984).
The liver, spleen and kidney of fry are pale due to anaemia; there may be ascites; and the stomach is filled with a milky fluid but no food. The intestine contains a watery, yellowish fluid and there may be petechial haemorrhages in the visceral mesenteries, adipose tissue, swim-bladder, peritoneum, meninges and pericardium (Rucker et al., 1953; Ross et al., 1960; Wolf, 1988). Older fish may have empty stomachs, intestines filled with yellowish mucus and lesions in the musculature near the kidney (Traxler, 1986).
The haematopoietic tissues of the kidney and spleen of young fish are the most severely affected and are the first tissues to show extensive necrosis (Amend et al., 1969; Yasutake, 1970, 1975). In recent studies, the gills, oesophagus/cardiac stomach region (OCSR), small intestine and pyloric caeca of rainbow trout and coho salmon fry were examined within 24 h after immersion in IHNV (Helmick et al., 1995a, b). No pathological changes were observed in the gills, small intestine or pancreatic acinar cells. However, pathological changes were observed in the OCSR mucus-secreting serous cardiac glands (MSSG) and epithelial cells of both species. In a study with steelhead trout fry, IHNV was transiently detected using immunohistochemistry in the epithelial cells of most organs, but it had a propensity for the connective tissue (Drolet et al., 1994). Virus was detected in the anterior kidney 1 day prior to detection in the posterior kidney. This was also observed by Yasutake and Amend (1972) in sockeye salmon. The first changes in the anterior kidney are small, lightly stained, focal areas, consisting of what appear to be macrophages and degenerating lymphoid cells. As the disease progresses, degenerative changes throughout the kidney become more noticeable. Macrophages increase in number and may have a vacuolated cytoplasm and chromatin margination of the nuclei (Klontz et al., 1965; Yasutake and Amend, 1972). There may also be a decrease in the number of non-differentiated blast cells, and pyknotic and necrotic lymphoid cells may be present. Focal areas of cells in the spleen, pancreas, liver, adrenal cortex and intestine show nuclear polymorphism and margination of the chromatin, with eventual necrosis (Amend et al., 1969; Yasutake, 1975; Wolf, 1988). Necrosis may be so severe that the kidney tissue consists primarily of necrotic debris (Yasutake, 1975; Wolf, 1988). Extensive necrosis in all organs is accompanied by pyknosis, karyorrhexis and karyolysis (Yasutake and Amend, 1972). A pathognomonic feature of IHN is degeneration and necrosis of granular cells in the lamina propria, stratum compactum and stratum granulosum of the alimentary tract (Yasutake, 1970; Wolf, 1988) and it is postulated that sloughing of intestinal mucosa may give rise to faecal casts (Amend et al., 1969; Yasutake, 1970, 1975). In Chinook and sockeye salmon fry there may also be hepatic deposits of ceroid (Wood and Yasutake, 1956; Yasutake, 1970). In the final stages of disease, necrosis is not only in the haematopoietic tissue of the kidney, but also in the glomeruli and kidney tubules. There is little effect on the corpuscles of Stannius tissue (Yasutake, 1975; Wolf, 1988). Smolts and yearlings tend to show less severe histopathological changes. The kidney, spleen, pancreas and liver may show necrosis, but there is only moderate sloughing of the intestinal mucosa and no faecal casts (Yasutake, 1978; Burke and Grischkowsky, 1984; Traxler, 1986).
In spawning sockeye salmon, Yamamoto et al. (1989) found that the spleen and kidney did not show the massive necrosis observed in fry. Instead, the kidney had no tissue destruction and the spleen had small focal infections near the periphery or at the outer cell layer. The connective tissues surrounding the spleen were also infected. Gill tissue had highly localized lesions, with some gill filaments having infection limited to the non-differentiating serosal cell layer, which lies internal to the epithelial pavement cell layer.
The number of monocytes is not affected, but there is leucopenia, with degenerating leucocytes and thrombocytes. Neutrophils are decreased or absent, the number of immature erythrocytes is increased and the cell may be bilobed (Watson et al., 1954; Wood and Yasutake, 1956; Holway and Smith, 1973; Amend, 1974; Amend and Smith, 1975). Amend (1973) suggested that the fish had a normocytic aplastic anaemia. Cellular debris, termed necrobiotic bodies, observed in blood smears or kidney imprints is pathognomonic for IHN (Holway and Smith, 1973; Amend and Smith, 1974, 1975; Wolf, 1988). The blood haematocrit, haemoglobin content, osmolarity and levels of bicarbonate, calcium, phosphorus, chlorides and bilirubin are decreased in infected fish. Lactic acid dehydrogenase levels may increase (Amend, 1974). However, there are no changes in kidney ascorbate levels, total plasma protein, mean corpuscular volume, mean corpuscular haemoglobin or levels of plasma glucose, esterase, glutamic oxylate transaminase or peptidase isozymes (Amend and Smith, 1974, 1975).
DiagnosisTop of page
If there is a previous history of IHN at the facility and the fish are showing characteristic clinical signs, IHNV may be the cause of a disease outbreak. However, the preliminary diagnosis should be confirmed by isolation and identification of IHNV. Recommendations for fish sampling, processing and diagnosis have been outlined by fish health officials in the USA and Canada (Department of Fisheries and Oceans, 1984; Ganzhorn and LaPatra, 1994; Thoesen, 1994). Fish tissues to be tested are dependent on fish size and disease status. The preferred tissues are the kidney and spleen; mucus has also been used in non-lethal sampling (LaPatra et al., 1989c). For testing brood stock, the ovarian fluid is preferred, since the virus is less frequently detected in the milt. Sampling of postspawning females, storage of ovarian fluid or incubation of ovarian fluid cells enhances the sensitivity of viral detection (Mulcahy et al. 1984b; LaPatra and Groberg, 1985; Mulcahy and Pascho, 1986; Mulcahy and Batts, 1987; LaPatra et al., 1990c). If milt is tested for virus, it should be centrifuged and, after the pellet is incubated in water, the water is assayed for virus. This procedure enhanced IHNV detection by at least fourfold (Batts, 1987; Meyers et al.,1990).
Methods used for diagnosing IHNV should be rapid, simple, sensitive, specific, inexpensive and able to test a large number of samples. Ideally, they should also be suitable for field use. The only method initially available for the diagnosis of IHNV was to detect virus in cell culture and then identify the virus by a serum neutralization test (Amend, 1970b). This classical method is widely accepted and is still commonly used. The discovery that rabbit antiserum has higher levels of IHNV-binding antibodies than IHNV-neutralizing antibodies (Hsu and Leong, 1985) allowed the development of several more rapid serological methods. There are also non-serological methods available for diagnosing IHNV. Each of the available methods are discussed below.
Virus isolation by cell culture
Samples are typically added to susceptible cell lines, such as EPC, CHSE-214 or FHM. Methods used for detecting and determining the quantity of IHNV in the sample are end-point titrations and plaque assays. The plaque assay is more sensitive than end-point titrations (Fendrick et al., 1982; Leong et al., 1983a). The American Fisheries Society ‘Blue Book’ (Amos, 1985; Ganzhorn and LaPatra, 1994) and the 'Manual of Compliance' of the Department of Fisheries and Oceans (1984) in Canada recommend that 2 cell lines (pH 7.6) are inoculated when 80-90% confluent and incubated at 15°C for a minimum of 14 days for end point dilutions or 7 days for plaque assays.
For plaque assays, the inoculum volume should be adjusted to reflect the size of the cell-culture well, since using too high a volume can reduce the efficiency of plaquing (Burke and Mulcahy, 1980; Batts et al., 1991). Viral adsorption is optimal after 1 h; shorter times reduce the detectable viral titre and longer times have no effect on the number of plaques (Burke and Mulcahy, 1980; Brunson et al., 1988; Batts and Winton, 1989a,b; Batts et al., 1991). Plaques are larger when cells are overlaid with methyl cellulose, but gum tragacanth or agarose at a pH of 7.2-8.0 can be used (Wolf and Quimby, 1973; Burke and Mulcahy, 1980; Okamoto et al., 1985; Batts et al., 1991).
Cells should be examined daily for cytopathic effect (CPE). If no CPE occurs, then the sample is virus negative, especially if the sample has been passed in tissue culture several times (blind passaged). If there is CPE characteristic of IHNV, then a presumptive diagnosis of IHNV is made. Typical CPE in cell cultures consists of grape-like clusters of rounded cells, with margination of the chromatin of the nuclear membrane (Wolf, 1988). Typical IHNV plaques consist of a cell sheet that retracts or piles up at the inner margins of the opening and the centre contains granular debris (Wolf and Quimby, 1973).
Attempts have been made to enhance the sensitivity of viral detection in cell cultures. Pretreatment of cell monolayers with polybrene, a polycation that enhances the ionic attraction of viruses to cells, increased titres of a laboratory strain of IHNV by five- to tenfold (Leong et al., 1981b) but did not affect titres of field isolates of IHNV assayed by plaque assays or end-point dilution (Fendrick et al., 1982). Pretreatment of cell monolayers with 7% polyethylene glycol (PEG, 20,000 Mr) improved the speed and sensitivity of plaque assays for the quantification of low levels of IHNV and resulted in the production of larger plaques (Batts and Winton, 1989a,b; Batts et al., 1991). Polyethylene glycol can also be added directly to the antibiotic solution that samples are incubated in, prior to inoculation on to cells (Brunson et al., 1988). The discovery of the effectiveness of PEG pretreatment is an important advance in the detection of IHNV in cell culture. Once a virus is detected in cell culture, its identity must be confirmed.
Serum neutralization test
An approved method of viral identification is the serum neutralization test, using either rabbit antisera or MAbs (Department of Fisheries and Oceans, 1984; Amos, 1985; LaPatra, 1994). Rabbit anti-IHNV serum typically has low neutralizing antibody titres (1:250 or less), but binding antibody titres as high as 1:50,000 have been reported (Leong et al., 1983a; Hsu and Leong, 1985; McAllister and Schill, 1986). The best schedule for producing polyclonal serum is to give rabbits an intramuscular injection of concentrated virus in Freund’s complete adjuvant, followed by intravenous boosts at 4 and 5 weeks with purified virus (Hill et al., 1981). Monoclonal antibodies are less toxic than rabbit antiserum, have neutralization titres of up to 1:1000, and can be used to distinguish IHNV variants (Roberti, 1987; Winton et al., 1988; Eaton et al., 1991; Roberti et al., 1991; Kamei et al., 1991; Huang et al., 1996).
Accurate detection of IHNV in cell cultures and identification by serum neutralization tests is expensive and time consuming. A 50% plaque reduction assay is more sensitive than a 50% end-point titration (Ahne, 1981), but neither assay is rapid. It can take from 7 to 10 days before results are obtained from neutralization tests and the total time to make a diagnosis can take from 2 to 8 weeks. Since IHN can kill the majority of fish within 2-3 weeks, it becomes moot to determine whether the fish died from IHN (Mulcahy et al., 1980; Leong et al., 1983a). Obviously, more rapid diagnostic methods had to be developed so that infected fish could be quickly destroyed or quarantined to prevent further spread of the virus.
In addition to the serum neutralization test, the fish health ‘Blue Book’ (LaPatra, 1994) lists immunofluorescence, staphylococcal coagglutination, enzyme-linked immunosorbent assay (ELISA), immunoblots and DNA probes as acceptable methods of identifying IHNV.
Immunofluorescence has been used to study fish viruses since 1972 (Jørgensen and Meyling, 1972) and is widely accepted for the detection and identification of IHNV in cell cultures. Either direct (DFAT) or indirect (IFAT) fluorescent antibody tests can be used. Routinely, DFAT uses fluorescein isothiocyanate (FITC)-labelled anti-IHNV polyclonal antibody or MAb and IFAT uses unlabelled primary antibody, which is detected using FITC-conjugated secondary antibody. Leong et al. (1983a) first reported IFAT for detecting IHNV in cell cultures and this method is rapid, specific and sensitive and can detect all 5 electropherotypes of IHNV (LaPatra et al., 1989a). A diagnosis can be made in 24 h using 50% end-point titration, if ovarian fluid inoculated on to cells has a virus titre of at least 102.9 pfu ml-1, and pretreatment of the cells with PEG improves the limit of detection to 102.5 pfu ml-1. A plaque assay is slightly more sensitive, detecting as low as 10 pfu ml-1, but requires up to 8 days to observe plaques and only provides a presumptive diagnosis (LaPatra et al., 1989a). Polyclonal and monoclonal antinucleoprotein and antiglycoprotein antibodies used in IFAT have equivalent sensitivity and time requirements for making a diagnosis, but preadsorption of polyclonal serum is required to prevent non-specific fluorescence (LaPatra et al., 1989b; Arnzen et al., 1991).
Used on cell culture samples, IFAT and DFAT have similar sensitivity (LaPatra et al., 1989a; Arnzen et al., 1991), but DFAT may be preferred because it is more rapid (Ristow and Arnzen, 1989; Arnzen et al., 1991). Cells should ideally be incubated for 16-24 h before using IFAT or DFAT, but, when rapid decisions are required, DFAT can be performed after only 10-12 h incubation (LaPatra et al., 1989b; Arnzen et al., 1991). Monoclonal antibody-based IFAT or DFAT can be used to type viral isolates, identify new viral types and study viral variants (Ristow and Arnzen, 1989; Ristow and Arnzen-de Avila, 1991; Roberti et al., 1991).
The 2 potential disadvantages of using fluorescent antibody tests (FATs) on cell cultures have been the use of cover glasses and detachment of cells after acetone fixation (LaPatra et al., 1989a). However, these problems have been overcome since LaPatra et al. (1989a) demonstrated that IFAT could be used directly on plaque assay plates and cells could be fixed with other fixatives. Kamei et al. (1991) confirmed that cover glasses were not needed by showing that IHNV could be detected in formalin-fixed cells in 96-well plates. Other disadvantages of FATs are that trained personnel are required and fluorescent microscopes are fairly expensive.
Direct detection of IHNV in fish tissues using IFAT or DFAT could substantially shorten the time for a diagnosis from several days to a few hours. Indirect FAT detected IHNV in blood smears and kidney imprints from clinically infected juvenile fish, but was unsuccessful in detecting virus in smears of seminal fluid (LaPatra et al., 1989a). The use of DFAT on fish tissues has yet to be investigated.
Staphylococcal coagglutination can specifically detect and identify IHNV grown in cell cultures or in infected fish tissue (Bootland and Leong, 1992). This test is simple and, since it takes only 15 min, it is one of the most rapid methods of diagnosing IHNV. It has the added benefit of being suitable for field use, because it only requires a light microscope, glass slides and one reagent. The test uses formalin-fixed Staphylococcus aureus cells sensitized with unadsorbed polyclonal rabbit anti-IHNV antiserum. When the antibody-coated cells are mixed with samples containing IHNV, the antibody specifically binds to the virus and causes the bacterial cells to coagglutinate.
For viruses grown in cell culture, the assay is specific for IHNV isolates belonging to all 5 electropherotypes, but the test is not very sensitive - 106 pfu ml-1 is generally required. However, this should not affect its use in diagnosing IHNV, since cell cultures showing complete CPE typically have virus titres greater than 106 pfu ml-1. The test is not suitable for use in cell cultures pretreated with PEG, due to non-specific coagglutination.
The coagglutination test identified IHNV directly from fry homogenates, adult organs and ovarian fluids, as long as the virus titre was at least 106 pfu g-1. No false positives were observed. Detection of IHNV in adult carriers may not be as effective due to the low sensitivity of the coagglutination test. Samples with a low viral titre would have to be passaged in cell culture to amplify the virus before a coagglutination test could be used.
The binding of MAbs to staphylococcal cells or latex beads, although not yet successful, may increase the sensitivity of the assay and allow differentiation between different types of IHNV.
Immunoassays (enzyme-linked immunosorbent assay, immunoblots and Western blots)
Several different types of immunoassays have been developed for the identification of IHNV. They all utilize the binding of antigen or antibody to a solid support and detection of the virus directly with labelled anti-IHNV antibodies or indirectly with labelled secondary antibody. These assays are not as rapid as coagglutination, IFAT or DFAT, but they are simple, are usually specific and can be quantitative. As for IFAT, rabbit polyclonal serum has to be preadsorbed with fetal bovine serum and cell lysates to make the assays more specific (Dixon and Hill, 1984). The limit of IHNV detection is generally 103-106 pfu ml-1 for ELISA and dot blots, and virus can be detected once cells begin to show CPE. This level of sensitivity is greater than that of the coagglutination test, but not as high as for IFAT and DFAT, which can detect IHNV prior to cells showing CPE. Unlike IFAT or the coagglutination test, none of the immunoassays are recommended for detecting IHNV directly from fish tissues.
ENZYME-LINKED IMMUNOSORBENT ASSAY
Using polyclonal antibodies or MAbs, ELISA techniques can detect and identify IHNV in cell cultures and have shown some success in detecting IHNV in fish tissues. Two different ELISA systems have been developed: a sandwich or capture ELISA and a direct ELISA. Both assays require 6-22 h to complete, and antibody concentrations, plate type, blocking agent, buffers, incubation times and temperatures each have to be optimized.
The initial sandwich ELISAs used plates coated for 6 h with rabbit anti-IHNV serum (Dixon and Hill, 1984) or 18 h with purified anti-IHNV immunoglobulin (Ig) (Way and Dixon, 1988), and IHNV antigen was detected using labelled rabbit anti-IHNV Ig. The assays were specific and detected IHNV in cell cultures once the virus titre had reached 106-107 pfu ml-1. The above 2 assays were improved by Medina et al. (1992) by use of a double-antibody sandwich ELISA. Plates were coated with chicken anti-IHNV serum and virus was detected by incubation with rabbit anti-IHNV serum, followed by labelled goat antirabbit serum. This assay was very sensitive, since it could detect 500 pfu ml-1, could be completed in 1 day and detected virus belonging to the 4 types tested.
A sandwich ELISA using biotinylated anti-N MAbs was developed and optimized by Ristow and de Avila (1991) to detect IHNV in cell cultures. Monoclonal antibodies had to be carefully chosen because they can recognize the same epitope, lose virus binding ability or have altered reactivity after biotinylation. The sandwich ELISA recognized many virus isolates belonging to the 5 IHNV electropherotypes and 103 pfu ml-1 could be detected with some IHNV isolates, but other isolates at high titres did not react. Using a mixture of MAbs or polyclonal antibodies combined with MAbs may improve the sandwich ELISA technique (Ristow and de Avila, 1991). This method is still not ideal, nor is it as sensitive and reliable as plaque assays, IFAT or dot blots for detecting IHNV in cell culture.
In the direct ELISA, plates are coated with purified virus instead of antibody. Using this method, with unlabelled MAbs to G, N and M2 IHNV proteins, Kamei et al. (1991) detected several isolates of purified IHNV, but the specificity depended on which MAb was used. There were no cross reactions between the IHNV N- or M2-specific MAbs and HIRRV, but one of 3 IHNV G-specific MAbs did cross-react, suggesting that IHNV and HIRRV have a common epitope. Before using G-specific MAbs in ELISA to detect IHNV, it must be ensured that cross-reactions with HIRRV do not occur.
Identification of IHNV in fish tissues using ELISA is not yet optimal. With a sandwich ELISA, Dixon and Hill (1984) and Way and Dixon (1988) found that whole fry homogenates or abdominal extracts gave a high background and weak cross reactions between heterologous viruses. To decrease background and make the assay more specific, Dixon and Hill (1984) suggested using concentrated Ig for plate coating, a more dilute affinity-purified, labelled Ig and diluted fish viscera. In the double antibody sandwich ELISA of Medina et al. (1992), there were low background reactions and IHNV was detectable directly from Atlantic salmon tissue homogenates if the titre exceeded 70 pfu ml-1, but only 6 fish were tested. The use of MAbs in ELISA to detect IHNV in fish tissues has not yet been investigated. Monoclonal antibodies could increase the specificity and sensitivity of the assay, thus improving the ability to detect virus in fish tissues without background. The techniques of ELISA may be well suited for field use, because visual results can be obtained without the need for specialized equipment and the assay is relatively rapid, especially if plates used in sandwich ELISA are precoated (Dixon and Hill, 1984).
IMMUNOBLOTS (DOT BLOTS)
The immunoblot assay or dot blot is a variation of the ELISA technique. Instead of binding antigen to wells, antigen is bound directly to a nitrocellulose or nylon membrane and detected with labelled primary antibody or indirectly with labelled secondary antibody. Immunoblots are faster than ELISA, requiring less than 4 h to complete, and may be more specific for identifying IHNV in cell cultures. Cell incubation time depends on the input multiplicity of infection (MOI), but ranges from 36 to 72 h or until CPE is observed and the virus titre has reached 103-106 pfu ml-1 (McAllister and Schill, 1986; Schultz et al., 1989; Ristow et al., 1991). Dot blots have detected as little as 4 ng of purified virus, and, unlike ELISA, no cross-reactions were observed with heterologous viruses and the primary antibody can be reused several times (McAllister and Schill, 1986; Eaton et al., 1991). Blot assays with labelled MAbs did not significantly enhance the sensitivity of the assay, but could distinguish between IHNV and VHSV, detect IHNV at titres as low as 103 pfu ml-1, recognize all 5 electropherotypes of IHNV and be completed in as little as 2.5 h (Schultz et al., 1989; Ristow et al., 1991).
Immunoblots have not been successful for identifying IHNV in fish samples, because the high protein content of tissues or reproductive products clogged the filters (McAllister and Schill, 1986). Also, an unknown component in ovarian fluids, which had a migration rate similar to the IHNV G and M2 proteins, cross reacted when biotinylated anti-IHNV MAb was used (Schultz et al., 1989). The cross-reacting components in ovarian fluids were not identified and cross-reactions were not due to endogenous peroxidase activity (Schultz et al., 1989). Further work is required to adapt immunoblots for detecting IHNV in fish samples.
In this method, cell culture proteins are separated by SDS-PAGE and transferred on to a membrane. Specific IHNV proteins are detected directly with labelled anti-IHNV antibodies or indirectly with labelled secondary antibody. The indirect method was first used to identify IHNV in cell cultures by Hsu and Leong (1985) and Leong et al. (1985). Using adsorbed polyclonal rabbit serum and 125I-protein A or secondary antibody labelled with horseradish peroxidase, all IHNV proteins were visible when the total viral protein was 2 µmg. The viral G, N and M1 proteins were detectable at 10 ng total viral protein. Cell growth and the Western blot required a total of 51.5-66 h, which included a 9 h cell incubation and 4 h for cell lysis and SDS-PAGE. Radioactivity is no longer required and several steps in the method of Hsu and Leong (1985) have been shortened. When anti-N antibody is used, cell incubation time can be decreased from 9 h to 3 h because viral N protein has already been synthesized in infected cells (Leong et al., 1983b). SDS-PAGE still takes 1-2 h, but protein transfer and the Western blot can now be completed in less than 6 h (Kamei et al., 1991; L.M. Bootland and J.C. Leong, unpublished observations). Western blots are of value in typing IHNV isolates and for comparing immunological relatedness among IHNV isolates or to other viruses. Kamei et al. (1991) used N-, M2- and G-specific MAbs in Western blots to study the antigenic relationship between IHNV and HIRRV and suggested that the 2 viruses have at least one G epitope in common. Chou et al. (1993) showed by Western blots that Japanese IHNV isolates had an N epitope that was only faintly visible in the North American strains tested. This may be of relevance in monitoring the spread of IHNV from area to area.
Western blots have not been commonly used in diagnostic laboratories because they are slower than other assays and specialized equipment is needed. However, Western blots are useful in determining the specificity of other assays (Schultz et al., 1989), provided that the antibodies have been proved to be IHNV-specific. Reports of using a direct technique with labelled primary antibody in Western blots have not been made, but this method may further decrease the time required to diagnose IHNV.
IMMUNOPEROXIDASE AND ALKALINE PHOSPHATASE CELL STAINING
These 2 methods are similar to FAT but use horseradish peroxidase- or alkaline phosphatase-labelled antibodies instead of FITC-labelled antibodies. Direct and indirect methods are rapid, specific and simple, do not require specialized equipment (Ahne, 1981) and may be suitable for field use. For identification of IHNV in cell cultures, immunoperoxidase staining was first suggested by Leong et al. (1983a), but this method is not routinely used. Kamei et al. (1991) used indirect immunoperoxidase staining with MAbs to IHNV N and M2 proteins to distinguish between IHNV and HIRRV. However, one of the 3 anti-G MAbs cross-reacted with HIRRV.
Alkaline phosphatase staining of cells has recently been adapted for confirming IHNV in plaque assays (Drolet et al., 1993). Plaque assay plates showing CPE were formalin-fixed, blocked overnight with 5% non-fat dry milk in PBS, incubated with unlabelled anti-N MAb and then biotinylated secondary antibody and avidin-alkaline phosphatase. If the plaques were caused by IHNV, then the cells around the margin of the plaque were coloured after the addition of substrate. If plates were initially stained with crystal violet, they could easily be destained by washes with 70% ethanol prior to blocking. The test was specific for IHNV, detected all 5 electropherotypes and often resulted in a positive reaction prior to visible CPE.
The immunoperoxidase and alkaline phosphatase cell-staining techniques have been frequently used in immunohistochemistry to study the pathogenesis of IHNV (Yamamoto et al., 1989, 1990, 1992; Drolet et al., 1994; Kim et al., 1994). Thin sections of fish were incubated with unlabelled primary monoclonal anti-G or anti-N antibody and horseradish peroxidase or biotin-labelled secondary antibody. The colour of the IHNV-infected cells depended on the substrate employed. Non-specific reactions have been observed, but could be removed by pretreatment of tissues with hydrogen peroxidase for horseradish peroxidase-based assays (Yamamoto et al., 1989) or by including levamisole in the substrate solution in alkaline phosphatase-based assays (Drolet et al., 1994). It is unlikely that staining of fish tissues will be adapted for the routine diagnosis of IHNV. The fixing, embedding and sectioning of fish tissues is the most time consuming part, with the actual antibody portion of the test being rapid, taking less than 6 h.
Identification of infectious haematopoietic necrosis virus by non-serological methods
In most of the above serological methods of diagnosing IHNV, the virus must be amplified in cell culture until protein antigen concentrations reach detectable levels. Instead of detecting antigens, it is possible to detect viral particles by electron microscopy (EM). Nucleic acids can be detected using the polymerase chain reaction (PCR) and nucleic acid probes. Since viral mRNA is synthesized before proteins in infected cells, detection of viral RNA may result in an earlier diagnosis of IHNV.
Electron microscopy was used for identifying IHNV in water and reproductive products (Leong et al., 1983a). The method is relatively cheap and very rapid, taking only 3-4 h, and detects 102-103 particles IHNV ml-1 of water, or 1 µ × 104 tissue culture infectious dose at 50% end-point (TCID50) ml-1. Diagnosis is based on visualizing viral particles having the characteristic bullet shape of rhabdoviruses. However, EM does not allow easy differentiation of viruses within the rhabdovirus family, especially between IHNV and VHSV, since the 2 viruses are of similar size. This technique provides only a presumptive diagnosis and is of little value in areas where both IHNV and VHSV occur. It also has the disadvantages of requiring trained personnel and an electron microscope.
Immunoelectron microscopy has the advantage of being able to specifically identify IHNV. It was first used by Helmick et al. (1991) to detect IHNV G and N proteins on the surface of infected cells. Using biotinylated MAbs, streptavidin-gold, scanning EM and X-ray analysis, viral proteins can be detected on cells at 8 h postinfection. Immunogold-labelling and EM has also been used to identify IHNV N protein in the kidney tissue of rainbow trout 1 year postinfection (Drolet et al., 1995). Although the method is sensitive and allows differentiation between IHNV and other fish rhabdoviruses, it required over 16 h to complete. It is unlikely that this technique will be used in diagnostic laboratories, since more rapid, simpler and less expensive serological methods have been developed. However, this method is of value for studying the life cycle of IHNV, since viral proteins can be detected in fish when infectious virus is not present.
POLYMERASE CHAIN REACTION AND NUCLEIC ACID PROBES
Polymerase chain reaction is a powerful technique used to amplify specific regions of DNA or RNA to easily detectable levels on agarose gels or with nucleic acid probes. This technique uses 2 primers that span the nucleic acid sequence of interest and a thermostable polymerase to exponentially increase the amount of nucleic acid through repeated cycles of synthesis. Nucleic acid probes consist of pieces of labelled RNA or DNA that specifically bind to complementary sequences in nucleic acids (Winton, 1991).
The N and G mRNA of IHNV have been completely sequenced (Koener et al., 1987; Gilmore and Leong, 1988), making it possible to develop PCR techniques and design specific probes to either IHNV gene. Initially, PCR and probes were developed for detecting N RNA, since the N mRNA is the first and most abundant mRNA produced in infected cells (Rose and Schubert, 1987) and is probably the best target for amplification and detection. Polymerase chain reaction has been used to detect IHNV N RNA in cell cultures, fresh fish tissue and paraffin-embedded fish kidney tissues.
Arakawa et al. (1990) were the first to use PCR for detecting IHNV N RNA in cell cultures. Using 20 mer (position 319-338) and 19 mer (position 570-552) primers, they detected a 252 bp PCR product from RNA extracted from cells infected with the 5 IHNV electropherotypes. No bands were evident after PCR of uninfected cells or with VHSV-infected cells, but 2 bands were observed from HIRRV-infected cells, suggesting that HIRRV and IHNV have some homology at the nucleotide level. Southern blots and dot blots, using a 30 mer (position 427-456) biotinylated antisense oligonucleotide probe, detected only IHNV. Cell cultures were harvested prior to CPE at 24 h and, as long as the extracted RNA template was 1 pg, PCR successfully amplified the target to detectable levels in blots. Deering et al. (1991) confirmed that the probe was specific for IHNV and showed that the cell incubation time was dependent on the input MOI. At a high MOI of 10, IHNV mRNA was detected by the probe at 6- 9 h, but, at a low MOI of 0.0002, the cells had to be incubated 4 days or until small foci of CPE were evident. More recently, reverse transcriptase-dependent PCR (RT-PCR) has been developed for differentiating between VHSV and IHNV, using primer pairs that amplify viral glycoprotein gene fragments (Bruchhof et al., 1995). By amplifying an IHNV 548 bp fragment and a VHSV 443 bp fragment from total RNA extracts of virus-infected RTG-2 cells, these researchers could differentiate the 2 viruses within 8 h.
Polymerase chain reaction has been used to amplify IHNV RNA extracted from fresh fish tissues (Arakawa et al., 1990). Ribonucleic acid extracted from kidney/spleen and leucocytes of rainbow trout killed at 3 days postinfection produced an N gene PCR product when the fish were infected with 104 or 103 pfu ml-1, but not when the viral dose was 102 pfu ml-1. All samples yielding a PCR product were also positive in plaque assays, but only the plaque assay identified IHNV in one fish infected with the lowest virus dose. Polymerase chain reaction is not as sensitive as plaque assays for detecting IHNV in fish tissues, but it is much more sensitive, more rapid and cheaper than several other techniques. At least 400 pfu ml-1 was required before PCR was able to amplify viral RNA to detectable levels in gels or blots.
Polymerase chain reaction has also been used to amplify RNA from formalin-fixed and paraffin-embedded fish tissues (Chiou et al., 1995; Drolet et al., 1995). Tissues could be stored in formalin for up to 2 years before embedding in paraffin, and PCR-amplifiable RNA template was obtained when RNAzol (phenol and guanidinium thiocyanate) was used to extract RNA. However, tissues could not be subjected to proteinase K digestion. Using the same N gene primers as Arakawa et al. (1990) and RT-PCR, a 252 bp product was identified in the kidneys of steelhead trout fry sampled at 7 days postinfection (Chiou et al., 1995) and in 1-year-old survivors (Drolet et al., 1995). Specificity was confirmed using Southern blots and the N-gene-specific probe of Arakawa et al. (1990). Investigators have developed nested PCR assays that can be completed in 7-9 hours (Miller et al., 1998; Alonso et al., 1999) and a multiplex PCR that can detect 3 different viruses in one PCR amplification reaction (Williams et al., 1999).
Polymerase chain reaction and probes to IHNV mRNA are relatively rapid, very sensitive and specific. Twenty-five PCR cycles require approximately 2 h, gels take 1-4 h and dot blots with the probe require over 6 h (Arakawa et al., 1990; Deering et al., 1991). For identifying IHNV in cell cultures infected with a high MOI, there is no clear advantage in using PCR and probes over other methods, when consideration is given to the time required and the need for specialized equipment, primers and oligonucleotide probe. The real advantage of PCR is its ability to amplify very low amounts of IHNV RNA more than a millionfold in a few hours to levels detectable with probes. This would be useful in detecting very low levels of virus in samples such as water, sediment, fish or other animals. Another area where PCR is very useful is the direct detection of IHNV in fish tissues, especially when infectious virus is no longer present. Although oligonucleotide probes are quite specific, their use for diagnosing IHNV in fish sections is not feasible, because of the amount of time involved - approximately 2-4 days. However, probes should prove valuable in studying the progression of virus within fish during and after an epizootic and for determining whether the virus enters a latent state.
Detection of anti-infectious haematopoietic necrosis virus antibodies in fish serum
For all of the above diagnostic methods, fish are usually sacrificed to obtain samples. A serological diagnosis of IHNV, based on the detection of anti-IHNV antibodies in fish serum, could determine the IHNV status of a fish stock without killing the fish.
Anti-IHNV antibodies were first demonstrated in fish serum using a serum neutralization test, in which heat-inactivated rainbow trout serum was incubated for 1 h with a low dose of virus (50-100 TCID50 per well) (Amend and Smith, 1974). Hattenberger-Baudouy et al. (1989) then showed that most fish sera would be negative for anti-IHNV antibodies if complement was absent and incubation was for only an hour. Infectious haematopoietic necrosis virus neutralization by trout antibodies is complement-dependent and is more evident after 16 h incubation. Heat-inactivation or one freeze-thaw of serum readily inactivates complement and the serum neutralization test is inconsistent and inaccurate unless serum containing complement is added.
In the improved serum neutralization technique of Hattenberger-Baudouy et al. (1989), heat-inactivated fish serum is diluted in 2 steps, mixed with 5% virus-free trout serum as a complement source and IHNV (1250 pfu per well) and incubated for 16 h at 4°C in 96-well plates. After adding EPC cells, the plates are incubated for 3-4 days at 15°C, fixed and stained with crystal violet. The antibody titre is defined as the highest serum dilution giving obvious protection of the cell monolayer.
In plaque reduction assays, the antibody titre is the serum dilution that results in a 50% reduction in the number of plaques compared with plates containing only a complement source. Background neutralization varies with the batch of complement and was observed at serum dilutions of less than 64 and with haemolysed serum (Hattenberger-Baudouy et al., 1989). These researchers determined that the plaque reduction technique was approximately ten times more sensitive than the microtechnique, but the latter method was preferred because a large number of samples could be easily analysed. Plaque neutralization tests were used to compare the antibody titre of trout injected with the 5 electropherotypes of IHNV (Basurco et al., 1993). Jørgensen et al. (1991) determined that IFAT was the most sensitive technique for detecting anti-IHNV antibodies in IHN survivors, and that a sandwich ELISA and neutralization tests had equivalent sensitivity. For IFAT, a double-labelling technique allowed detection of both IHNV and anti-IHNV antibody. Cross-reactions of fish serum with VHSV and IHNV were more evident with ELISA than with IFAT, but did not occur in neutralization tests. Ristow et al. (1993) found that a simplified ELISA was more rapid (18 h) and detected a higher number of fish with anti-IHNV antibodies than a plaque neutralization test, which requires over 6 days. Western blots did not detect anti-IHNV antibodies in the majority of the fish, possibly because the conformation of the viral epitopes had been changed, due to denaturing conditions, but this assay is valuable in determining which viral proteins are inducing a humoral immune response.
For determining the virus-free status of a hatchery fish population, Jørgensen et al. (1991) recommended using neutralization assays, because of the specificity of the test. However, this method requires proper storage of the serum used as a complement source, is time-consuming and requires technical expertise. Ristow et al. (1993) suggested that ELISA could replace the plaque neutralization test for detecting anti-IHNV antibodies, because the 2 assays were of similar sensitivity but ELISA required less time to complete. The ELISA is valuable for screening large numbers of samples, is relatively specific and reproducible and is of low cost in time and materials.
Future directions for research: diagnosis
Since IHN is an untreatable disease and control is by avoidance, it is critical that there be a rapid diagnostic technique for IHNV. Current diagnostic assays are too slow to adequately prevent spread of the virus. To decrease the time required, sensitive assays to detect IHNV directly from fish tissue need to be developed. Assays such as coagglutination, DFAT and ELISA, which incorporate MAbs may have improved sensitivity and specificity and ultimately allow detection of IHNV without the use of cell culture.
List of Symptoms/SignsTop of page
|Finfish / Bursts of abnormal activity - Behavioural Signs||Aquatic:Fry||Sign|
|Finfish / Darkened coloration - Skin and Fins||Aquatic:Fry||Sign|
|Finfish / 'Dropsy' - distended abdomen, 'pot belly' appearance - Body||Aquatic:Fry||Sign|
|Finfish / Generalised lethargy - Behavioural Signs||Aquatic:Fry||Sign|
|Finfish / Haemorrhaging - Body Cavity and Muscle||Aquatic:Fry||Sign|
|Finfish / Mortalities -Miscellaneous||Aquatic:Fry||Sign|
|Finfish / Paleness - Gills||Aquatic:Fry||Sign|
|Finfish / Pop-eye - Eyes||Aquatic:Fry||Sign|
|Finfish / Red spots (haemorrhagic lesions) - Eyes||Aquatic:Fry||Sign|
|Finfish / Red spots: pin-point size (petechiae) - Skin and Fins||Aquatic:Fry||Sign|
|Finfish / Trailing, flocculent or mucous faeces (casts) - Body||Aquatic:Fry||Sign|
Disease CourseTop of page
Future directions for research: pathogenicity and epizootiology
Progress has been made in elucidating the life cycle of IHNV, but this remains a critical area for future research. Studies are needed to fully characterize the latent state in rainbow trout and determine if latent infections also occur in other fish species. It must also be determined if viral reactivation occurs and, if so, the conditions that trigger the production of infectious virus need to be elucidated.
Chronic IHNV infections in fish over 6 g often cause less than a 30% mortality, but, because losses occur in market-sized fish and fish may also be scoliotic, high economic losses can occur. It was estimated that discarding scoliotic fish caused losses of $350,000 per year at one facility in Idaho (R.A. Busch in Nicholson, 1982). Infectious haematopoietic necrosis virus isolates show variations in tissue tropism (Kim et al., 1994), but the mechanism causing scoliosis is unknown. Studies are required to elucidate the virulence factors of IHNV, identify IHNV cell receptors and further characterize the correlations between IHNV antigenic types and virulence.
It was recently shown that IHNV could be isolated from sexually immature fish in the marine environment (Traxler et al., 1997). These fish, as well as sexually mature fish in fresh water, may have a reactivation of a latent infection. Alternatively, fish may be infected through contact with a marine or freshwater reservoir or vector of IHNV. Although marine or freshwater organisms could harbour IHNV, sources of IHNV other than salmonid fish have not been identified. The susceptibility of non-salmonid fish to IHNV and the role of ‘wild’ fish in spreading IHNV have not received much attention.
The evidence for vertical transmission is primarily circumstantial. Confirming that vertical transmission occurs will be difficult, but attempts should continue. Another area that needs further investigation is the immune response to IHNV and whether maternal transfer of antibodies to eggs occurs.
EpidemiologyTop of page
The epizootiology of IHNV is not completely understood and the source of virus infecting salmonids is still unknown. However, survival of IHNV in a fish population depends on a close association of the virus with the life cycle of the fish host. Similarly to other salmonid viruses, IHNV typically causes an acute disease in young salmon and trout. Mortalities of fry and fingerlings can be as high as 90%, but occasional epizootics have been reported in smolts and older fish (Yasutake, 1978; Busch, 1983; Burke and Grischkowsky, 1984; Roberts, 1986; Traxler, 1986). The source of virus infecting young salmonids may be from vertical transmission or by direct water-borne exposure to IHNV from an unknown reservoir or host. During an epizootic in young fish, the virus is transmitted horizontally from fish to fish and infectious virus is readily isolated up to approximately 50 days after viral exposure (Amend, 1975; Busch, 1984; Bootland et al., 1995; Drolet et al., 1995). Thereafter, infectious virus is usually not isolated again until the fish near or reach sexual maturity (Amend, 1975; Busch, 1984; Meyers et al., 1990; Hattenberger-Baudouy et al., 1995b). Two mechanisms may explain why infectious virus is isolated at only two stages of the salmonid life cycle. First, IHNV may be entering a latent state in fish surviving an IHN epizootic, with virus reactivation occurring only as fish mature sexually (Amend, 1975). Alternatively, the fish may completely clear the virus but become reinfected prior to or during their spawning migration (Elston et al., 1989; Traxler et al., 1997). Although non-piscine reservoirs have not been identified, there is evidence to support the suggestion that both mechanisms exist.
Infectious haematopoietic necrosis virus in spawning salmonid fish
Natural IHNV infections have been reported in sexually mature kokanee salmon, Chinook salmon, chum salmon, coho salmon, rainbow trout and steelhead trout from feral and hatchery populations (Busch, 1984; Groberg, 1985; Goldes et al., 1986; Follett et al., 1987; LaPatra et al., 1987, 1989b; Engelking and Kaufman, 1994a). However, the majority of studies on IHNV in sexually mature fish have been on female sockeye salmon because of their high susceptibility and because IHNV is enzootic in this species in the Pacific North- West of North America. Populations containing spawning sockeye salmon, Chinook salmon and steelhead trout have an IHNV infection rate that varies from 0 to 100% and females have a higher prevalence of infection than males (Wingfield and Chan, 1970; Mulcahy et al., 1983a, 1984b; LaPatra et al., 1989b; Yamamoto et al., 1989; LaPatra, 1990; Meyers et al., 1990). The differences in prevalence between the sexes may be related to hormone levels or difficulties in detecting IHNV in milt (Meyers et al., 1990). Infectious haematopoietic necrosis virus should be eluted from sperm using water to enhance detection (Batts, 1987), but many studies do not appear to have used this method.
In naturally spawning sockeye salmon, IHNV was not isolated from fish during their migration from the ocean to the spawning area, but virus suddenly appeared in a low percentage of prespawning females in the spawning grounds. As the spawning migration progressed and fish density increased, viral prevalence and titres increased (Mulcahy et al., 1982, 1984b; Mulcahy and Pascho, 1986). A recent study by Traxler et al. (1997) showed that IHNV could be isolated from adult sockeye salmon during the ocean phase of their life cycle. It was postulated, based on the presence of neutralizing anti-IHNV antibodies in the fish at approximately 2 months prior to spawning, that the fish could have been exposed to IHNV 5-6 months prior to sexual maturation. Postspawning fish have a high viral prevalence and titres in the ovarian fluid, spleen, kidney and gill, but results vary from year to year (Meyers et al., 1990). When Meyers et al. (1990) analysed the data from 96 wild and hatchery populations taken over 14 years, there was no significant difference between spawning and postspawning fish for the prevalence of IHNV infection or viral titres. It is evident that infection rates cannot be predicted from year to year for different fish populations and that fish do not necessarily need to be on their spawning grounds to become infected with IHNV.
How adult salmonid fish are becoming infected
What has intrigued biologists most about the life cycle of IHNV is that virus typically can not be reisolated from adult fish until they approach sexual maturity. There are two hypotheses to explain this phenomenon. The first is that fish may become reinfected by a new exposure to IHNV during the spawning migration. This implies that the virus is present in the environment or in another host. The second hypothesis is that fish surviving IHNV as fry may develop a latent infection, which is reactivated when the fish reach sexual maturity. Fish returning to fresh water to spawn are undergoing stress, which may weaken the immune system so that the fish cannot control the virus (Mulcahy et al., 1984b). Hormonal changes may play a role, but injection of sex hormones did not induce or suppress virus infection in spawning fish (Grischkowsky and Mulcahy, 1982). It is possible that a combination of the two hypotheses accounts for the infection in sexually mature fish. A portion of fish may have reactivated latent infections and virus shed from these fish into the water may spread horizontally to other fish.
Water-borne IHNV can result in horizontal transmission between adult salmonids. Field observations showed that yearling fish cohabiting with sockeye salmon adults became infected with IHNV (Mulcahy et al., 1983a) and waterborne virus potentially spread from adult Chinook salmon to adult coho salmon at the Trinity River hatchery in California (LaPatra et al., 1987, 1989b). Laboratory data have confirmed that spawning fish can be infected by waterborne IHNV. Sexually mature kokanee salmon were infected after only 30 min immersion in a low dose of IHNV and viral titres in pooled spleen and gill increased over an 8-day period (Yamamoto et al., 1989). Immersion of sexually mature kokanee salmon in 103 pfu ml-1 of RB-1 (type 1) or 104 pfu ml-1 of WRAC (type 2) IHNV isolates for 1 h resulted in infection of 11 organs per fish and the reproductive products, with mortalities reaching 94% and 72%, respectively. Control fish had a mortality of 60%, attributable to natural causes (H.V. Lorz, L.M. Bootland, J.S. Rohovec and J.C. Leong, unpublished data). Thus, exposure of adult kokanee to IHNV in the water for only a short time can result in a rapid and widespread infection of the organs and cause premature fish mortality. Water taken from the spawning grounds has viral titres that may be sufficient to infect salmonids, but titres vary during the spawning season and from year to year, and IHNV probably does not survive in water over the winter (Mulcahy et al., 1983a). This suggests that there must be an alternate source of virus. Based on the presence of IHNV in adult sockeye salmon and Atlantic salmon in salt water, there may be a marine reservoir of infection (Traxler et al., 1991, 1993, 1997; Armstrong et al., 1993). However, virus reservoirs, except for the fish themselves, have not been identified.
It is now clear that IHN survivors can become latently infected with IHNV. The initial evidence that IHNV enters a latent state was presented by Amend (1975). In his study, IHNV could not be reisolated from rainbow trout fry within 3 weeks after an IHN epizootic or over the next 2 years, but 33% of the sexually mature fish had IHNV in the reproductive products. Similar results were obtained by Busch (1984), who found that virus was not reisolated from rainbow trout after 60 days postinfection or over the next 2 years from fish kept in virus-free water, but, when the fish reached sexual maturity, infectious IHNV was isolated from 2.56% of the males and 2.84% of the females. In contrast, field data from sockeye salmon in Washington, USA, suggested that fish did not have a latent infection but became reinfected upon reaching the spawning grounds. Fish captured over 3 years from a population which normally had a 98- 100% incidence of IHNV did not become infected when held to sexual maturity in pathogen-free water (Amos et al., 1989). Similarly, it was initially reported that sockeye salmon held in freshwater net pens before the fish reached their spawning grounds did not become infected with IHNV, even though 89% of fish removed from the spawning ground were infected (Kent et al., 1988; Elston et al., 1989). However, when this study was continued, at least 25% of the fish held in net pens became infected, and yet groups of spawning fish held in two locations upstream of the net pens were uninfected or had an infection rate of 17% (Elston et al., 1989). These results demonstrate that it is not possible to predict infection rates from one year to the next and that fish do not need to be at the spawning grounds to become infected with IHNV. The failure to detect a reactivation of a latent state may have been due to a small sample size. Typical sample sizes will detect a 5% prevalence of infection, but, based on the results of Busch (1984), this presumed incidence may be too high.
In a long-term laboratory study, over 2800 rainbow trout fry were experimentally infected by immersion and the surviving fish were held in pathogen-free water and sampled for up to 4 years (Bootland et al., 1995; Drolet et al., 1995). Infectious haematopoietic necrosis virus was isolated from fry for 50 days postinfection and, even though 30 fish were sampled every 6 months, including during two spawning cycles, in which over 600 reproductive samples were tested by plaque assays, infectious virus was not reisolated from organs or from reproductive products. These results indicate that IHNV did not enter a latent state that was reactivated as the fish reached sexual maturity. However, examination of the tissues of the survivors proved that IHNV had entered a latent state. Kidney tissues taken from 30 surviving fish at 3, 6, 12 and 24 months postinfection contained both the N and G IHNV proteins by immunohistochemistry. Viral proteins were not seen in any of the other ten tissues, except for the liver of one fish sampled at 90 days. Examination of the kidneys of 1-year-old survivors using PCR indicated that viral N RNA was present (Chiou et al., 1995), and immunogold EM showed that the renal epithelium of four fish had intracytoplasmic inclusions containing truncated IHNV particles that resembled the defective interfering particles (DIPs) of other rhabdoviruses (Drolet et al., 1995). It was subsequently demonstrated that livers of surviving fish produce large quantities of DIPs and that these particles have biological activity (C.H. Kim and J.C. Leong, unpublished data). When explant liver and kidney tissues from survivor or control fish were re-exposed to IHNV, the viral titres in the liver tissues, but not in the kidneys, of survivor fish were significantly decreased compared with control fish tissues. By immunosorbent EM, survivor tissues had truncated IHNV particles, while control fish tissues had only standard IHNV particles. It was postulated that superinfection of survivor liver tissues resulted in the release of a large number of DIPs, with subsequent reduction in standard viral production. Other factors, such as interferon and cell-mediated immunity, might also have played a role in decreasing tissue viral titres, but there is strong evidence that DIPs are involved. It is now clear that at least some IHNV survivors harbour a latent infection, but to date it has not been possible to show a reactivation of the latent state. Conditions that trigger reactivation still have to be identified.
Humoral immune response to infectious haematopoietic necrosis virus
Amend and Smith (1974) showed that fish hyperimmunized with IHNV mounted a humoral immune response, with the production of neutralizing antibodies. It was later shown that viral neutralization by antibodies was complement-dependent (Hattenberger-Baudouy et al., 1989). The humoral immune responses to IHNV and other fish viruses are very similar. Young rainbow trout during an IHN epizootic did not have detectable neutralizing antibody titres, but by 4.5-6 months after infection, when the virus was no longer detectable, over half of the surviving fingerlings tested had a significant humoral immune response. By 8 months, the number of antibody-positive fish and the antibody titre had decreased (Hattenberger-Baudouy et al., 1989). Similar results were obtained by LaPatra and his colleagues in 1993 (LaPatra et al., 1993b). Juvenile rainbow trout immersed in IHNV had a low prevalence and titre of serum anti-IHNV antibodies at 1 week postexposure, but by 6 weeks over 50% of the fish had high neutralizing antibody titres. Neutralizing activity was also observed in the mucus. Cain et al. (1996) noted that juvenile rainbow trout injected or immersed in IHNV produced serum- neutralizing antibodies by 21 and 28 days. Mucus from the skin and gastrointestinal (GI) tract had viral neutralizing activity, but no antibodies were detected using ELISA. They suggested that there are innate mechanisms of viral resistance that may play an important role as the first line of defence.
Antibodies are readily detected in fish exposed to IHNV using FATs, ELISA and Western blots (Vestergard Jørgensen et al., 1991; Zhuang et al., 1992; Ristow et al., 1993; Engelking and LaPatra, 1996; Traxler et al., 1997). Ristow et al. (1993) examined the serum from juvenile rainbow trout that had survived from one to five natural exposures to IHNV. They reported that 92% of the fish had anti-IHNV antibodies and, by Western blot, some fish reacted to all IHNV proteins, but the majority of the sera reacted with only the IHNV M1 and G proteins. Similarly, rainbow trout injected with virus in adjuvant (Mourich and Leong, 1991) or immersed in IHNV (LaPatra et al. 1993b) only responded to the M1 and G proteins by Western blots. It is possible that fish are reacting to other IHNV proteins, but the epitopes may be destroyed by the denaturing conditions used in Western blots.
Adult salmonids may have detectable serum IHNV neutralizing antibodies after a natural or experimental exposure to the virus (Hattenberger-Baudouy et al., 1989, 1995a,b; LaPatra et al., 1993b; Bootland et al., 1995). Basurco et al. (1993) found that individual adult rainbow trout varied in their ability to produce antibody after injection with the five IHNV electropherotypes. Rainbow trout distinguished the type 2 isolate from the other isolates, but it was concluded that the fish did not discriminate with high fidelity between most types of the virus. In Oregon, a survey of anadromous adult salmonids showed that the fish mounted a similar humoral immune response to IHNV to that of rainbow trout (Engelking and LaPatra, 1996). Antibody titres tended to increase with time in Chinook salmon, and, in some cases, Chinook salmon and steelhead trout with antibodies were also infected with IHNV. Traxler et al. (1997) found that sexually maturing sockeye salmon mounted a humoral immune response to IHNV. In one group of fish captured in the autumn, antibody titres increased over time, although IHNV was only detected in one in 19 fish. In a second group, none of the 21 fish had increased antibody titres, but 42% were virus-positive. Similar results were found with fish removed from the spawning grounds - virus was only found in fish that had no detectable antibodies. That sexually mature fish produce antibodies against IHNV indicates that these fish are immunocompetent. Although adults have now been shown to produce anti-IHNV antibodies, an unresolved question is whether there is maternal transfer of antibodies to the eggs. There is one report of isolating IHNV-neutralizing antibodies in steelhead trout eggs (Shors and Winston, 1989b), suggesting that maternal transfer of antibody occurs. However, this study did not indicate if the parents were infected with IHNV or whether the antibody was specific to IHNV. Transfer of antibodies against Vibrio anguillarum to eggs was reported for coho salmon, but protection against the bacteria did not last after the yolk-sac was absorbed (Brown et al., 1994). Whether antibodies transferred from the female to the eggs will be effective in protecting fry against IHNV has not been determined.
Vertical transmission - infectious haematopoietic necrosis virus in the reproductive products
Vertical virus transmission is the transmission of virus from one generation to its offspring, regardless of whether the virus is located within or external to the egg (Fenner and White, 1976; Pilcher and Fryer, 1980a,b). Since iodorphor treatment is usually sufficient to stop the transmission of the virus from adults to progeny in most cases, transmission within the egg (Nicholson, 1982) has generally been dismissed. In those cases, where iodophor treatment fails, most managers have attributed the subsequent IHNV outbreak to improper use of the iodophor. Others have suggested that there is a rare but distinct transmission of IHNV within the egg.
The most frequent evidence for vertical transmission is the association between the shipment of infected eggs into new geographical areas and resultant outbreaks of IHN (Plumb, 1972; Holway and Smith, 1973; Sano et al., 1977; Niu and Zhao, 1988). Further evidence of vertical transmission is that IHN has occurred in progeny from eggs disinfected with iodophor and raised in virus-free water (Wingfield and Chan, 1970; Ratliff et al., 1982; Mulcahy and Bauersfeld, 1983; Mulcahy and Pascho, 1985; Meyers et al., 1990; Roberts, 1993). However, vertical transmission may be an infrequent event. There are several reports where IHNV-infected parents did not produce IHNV-infected progeny when the eggs were raised in virus-free water (Amend, 1975; LaPatra, 1990; Engelking et al., 1991b; LaPatra et al., 1991b; Yamazaki and Motonishi, 1992; Traxler et al., 1997). This has raised doubts as to whether vertical transmission really occurs. However, negative results do not eliminate the possibility of vertical transmission of IHNV. Mulcahy and Pascho (1985) indicated it was difficult to demonstrate vertical transmission and they were only successful three times in isolating IHNV from live and dead eggs and fry of infected adult sockeye salmon. They found that only a small portion of sockeye eggs and fry contained IHNV and that not all parents transmitted the virus to their progeny and not every egg from infected females contained the virus. These observations would explain why studies to demonstrate vertical transmission have failed when only a small number of eggs were used, and why IHN epizootics appear intermittently within hatcheries.
It is accepted that salmonid reproductive products can harbour IHNV, but it is controversial whether the virus is on the egg surface or within the egg. Observations that disinfection of eggs does not always prevent IHNV infection of progeny suggest that the virus is within the egg (Amend, 1975). However, the virus may be external. Another fish virus, IPN virus (IPNV), adheres strongly to the chorion of water-hardened eggs, but not to unfertilized eggs (Ahne and Negele, 1985). Infectious pancreatic necrosis virus was isolated from eggshells after fry hatched and the fry may become infected by eating the eggshells. The chorion of unfertilized eggs is smooth, but hardened eggs have a rough, lobed and porous texture, which would provide anchorage for the virus and may prevent disinfectants from reaching the virus. A similar situation may be occurring with IHNV. Examination of eggshells for IHNV has not yet been done.
Fish species may be an important determinant for vertical transmission. When eggs of masu salmon and chum salmon were exposed to IHNV and then fertilized, the eggs and resulting fry were not infected (Yoshimizu et al., 1989). The stage of egg development influenced the susceptibility to IHNV replication. Exposure of unfertilized eggs to virus did not result in viral replication, but when eyed eggs were injected with IHNV, the virus replicated and the resulting fry suffered IHN-induced mortality. Egg-yolk components inhibited viral replication. An increase in IHNV susceptibility as the egg matured was correlated with a decrease in yolk components. Yoshimizu et al. (1989) concluded that direct vertical transmission of IHNV within the egg is doubtful. The anti-IHNV action of the yolk-sac components may be species-specific. Rainbow trout may have similar viral-inhibiting components in the yolk-sac, since fry developed IHN when eyed eggs were immersed in IHNV (Amend, 1975). However, sockeye salmon do not appear to have viral-inhibiting egg components, since Burke and Mulcahy (1983) found that IHNV infectivity was stable in egg homogenates and Mulcahy and Pascho (1985) isolated IHNV from eyed eggs reared in virus-free water. Inhibition of IHNV by egg components of other species does not appear to have been tested.
Infectious haematopoietic necrosis virus is frequently isolated from milt (Wingfield and Chan, 1970; Meyers et al., 1990; Yamazaki and Motonishi, 1992) and male salmonid fish may have a role in vertical transmission. Sockeye salmon and steelhead trout have a lower prevalence of infection and viral titres in milt than in the kidney and spleen (Mulcahy et al., 1987). The prevalence of infection in milt is lower than that in ovarian fluid, but the proportion of males with high milt viral titres may be equivalent to that of females with high virus concentrations in the ovarian fluid (Meyers et al., 1990). Virus strongly and quickly adsorbs to the surface membrane of steelhead trout and Chinook salmon sperm (Mulcahy and Pascho, 1984). Sperm that has adsorbed IHNV from male fish or from infected ovarian fluid could deliver the virus directly into the egg during fertilization (Mulcahy and Pascho, 1984; Meyers et al., 1990). However, the role of sperm in IHNV transmission is still unknown. Contamination of masu salmon or chum salmon milt with IHNV did not result in infection of eggs or fry (Yoshimizu et al., 1989) and there is no direct evidence that sperm can transmit IHNV into eggs.
Additional factors such as hatchery practices, genetics and environmental conditions may influence vertical transmission. Large-scale hatchery operations raising over 20 million eggs annually may have an increased probability of vertical transmission. However, hatcheries rearing less than 7 million eggs may have no IHNV losses, despite females having a 100% prevalence of infection and high viral titres (Meyers et al., 1990). Infectious haematopoietic necrosis epizootics can be prevented by not mixing eggs from several females and by incubating the eggs in virus-free water. Mixing of eggs from many females may contaminate the eggs from females that had a low viral titre with a lethal dose of virus from a few females with high virus titres (Mulcahy et al., 1983b). The roles of fish genetics and environmental factors in vertical transmission are unclear.
Fry to juvenile susceptibility to infectious haematopoietic necrosis virus
Fry and fingerlings are most susceptible to IHNV infection and mortality. Vertical transmission may be the cause of infection in some young fish, but no correlation was found between outbreaks of IHN in progeny and the infection rate or IHNV titres in the parents (Mulcahy et al., 1983b). Horizontal virus transmission from fish to fish through the water is the major mechanism of spread between fry (Amend, 1974). Prior to the onset of an epizootic, viral titres in water may be undetectable or very low (up to 0.07 pfu ml-1), but during the early stages of epizootics viral titres rise and may reach 50 pfu ml-1 to over 103 TCID50 ml-1 during epizootics (Nishimura et al., 1988; Zhang and Congleton, 1991, 1994). Viral titres are progressively amplified by repeated cycles of shedding and infection (Zhang and Congleton, 1994). Hatchery effluent water has had a viral titre of 400 pfu ml-1 (Leong and Turner, 1979) and this is high enough to infect fish.
Young fish show signs of IHN and mortalities within 5-10 days of exposure (Wingfield and Chan, 1970; Amend, 1975; Nishimura et al., 1988: Yamamoto et al., 1990). The general pathology and histopathology of IHNV infections are well characterized, but it is only recently that the route of virus entry has been investigated. Studies with rainbow trout, coho salmon and steelhead trout immersed in IHNV indicated that the gills, the oral region, the OCSR and the skin are the principal sites of virus entry (Yamamoto et al., 1990; Drolet et al., 1994; Helmick et al., 1995a,b). No histological changes or virus was apparent in the gill tissues within 24 h postinfection (Helmick et al., 1995a), but virus was observed at 2 days (Drolet et al., 1994) and 9 days (Yamamoto and Clermont, 1990) in rainbow trout fry. Yamamoto et al. (1990) indicated that the skin was a major portal of entry and was more important than the gills. However, skin epithelial cells were weakly positive by immunohistochemistry at 1 day postinfection and it appeared that the epithelial cells were only transiently infected in steelhead trout fry (Drolet et al., 1994). In coho salmon and rainbow trout fry, another early target area for IHNV attachment and replication is the OCSR, particularly the MSSG (Helmick et al., 1995b). Because of the proximity of the MSSG to the swim-bladder duct, which is adjacent to the anterior kidney, these researchers suggested that the MSSG may be a portal of entry for viral infection. In steelhead trout fry, the progression of the virus was followed for 14 days postinfection (Drolet et al., 1994). It was found that virus infection progressed from two major sites: from the gill into the circulatory system and from the oral region into the GI tract and then the circulatory system. Once IHNV was in the blood, the virus was disseminated to virtually every organ. Although IHN frequently occurs in fry and fingerlings reared in hatcheries, IHN also occurs in feral or wild salmon. There have been at least two instances where sockeye salmon fry emerging from the gravel in spawning channels died of IHN, and natural epizootics have occurred in sockeye salmon fry and smolts (Williams and Amend, 1976; Traxler and Rankin, 1989; Olson and Thomas, 1994).
In older fish, yearling rainbow trout (Busch, 1983; Roberts, 1986), Chinook salmon (LaPatra et al., 1990d), sockeye salmon smolts (Yasutake, 1978; Burke and Grishchkowsky, 1984; Olson and Thomas, 1994) and 2- to 3-year-old kokanee salmon (Amend, 1974; Traxler, 1986; Banner et al., 1991) have succumbed to IHNV in fresh water. In many cases, infected fish within the population or other species of infected fish are thought to be the virus source, and horizontal transmission through the water probably occurred (Burke and Grishchkowsky, 1984; LaPatra et al., 1990d; Olson and Thomas, 1994).
Potential sources of infectious haematopoietic necrosis virus
Sources of IHNV other than salmonid fish have not been identified, but potential sources include freshwater and marine invertebrates, sediment and other fish species. Depending on temperature, microflora and electrolyte concentration, IHNV survives in fresh water for from 3 days to several months (Wedemeyer et al., 1978; Toranzo and Hetrick, 1982; Yoshimizu et al., 1986; Kamei et al., 1988b). Infectious haematopoietic necrosis virus survives longer in salt water than in fresh water (Winton et al., 1991), and, although it is unlikely that fish become infected through the ocean water, because of the high dilution factor, this remains a possibility. Ingestion of marine organisms harbouring IHNV is another possible mechanism of infection, but IHNV has not been isolated from marine organisms (Traxler et al., 1997).
Salmon leeches (P. salmositica) can harbour IHNV; however, there is little relationship between leech viral titres and those of the host sockeye salmon (Mulcahy, 1986; Yamamoto et al., 1989; Mulcahy et al., 1990). Although leeches can have high IHNV titres, it is not clear whether IHNV is actually replicating in the leech or whether the leech is concentrating and storing the virus from fish blood. Infectious haematopoietic necrosis virus has also been isolated from copepods (Salmincola sp.) and the common mayfly (Callibaetis sp.). Copepods always had lower viral titres than those of fish gill tissue (Mulcahy et al., 1990). Transmission from invertebrates to salmonid fish has not been demonstrated, but the role of invertebrate reservoirs cannot be ruled out.
Impact SummaryTop of page
|Fisheries / aquaculture||Negative|
Impact: EconomicTop of page
Economic losses from IHNV can be a direct consequence of fish mortality, or indirect from regulations restricting the movement of IHNV-infected fish or the destruction of infected fish stocks to control the spread of the virus. Infectious haematopoietic necrosis is the most important constraint to the profitability and continued growth of the US commercial salmonid aquaculture industry, with lost revenue in the Idaho trout industry estimated at $3 million annually (Congleton, 1988). At government hatcheries along the Columbia River, upwards of 70 million fish and eggs have been destroyed as a result of IHNV infection since 1981. The value of these fish has been conservatively estimated at $350 million based on the estimates of losses to the sports and commercial fishery (Leong et al., 1995).
Zoonoses and Food SafetyTop of page
This species is not a zoonosis.
ReferencesTop of page
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