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infectious pancreatic necrosis

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infectious pancreatic necrosis

Summary

  • Last modified
  • 14 July 2018
  • Datasheet Type(s)
  • Animal Disease
  • Preferred Scientific Name
  • infectious pancreatic necrosis
  • Overview
  • Infectious pancreatic necrosis (IPN) is a highly contagious viral disease of young fish of salmonid species held under intensive rearing conditions (

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PictureTitleCaptionCopyright
Frequency of isolation of aquatic birnaviruses in the USA.
TitleIsolation of aquatic birnaviruses in the USA
CaptionFrequency of isolation of aquatic birnaviruses in the USA.
CopyrightP. W. Reno
Frequency of isolation of aquatic birnaviruses in the USA.
Isolation of aquatic birnaviruses in the USAFrequency of isolation of aquatic birnaviruses in the USA.P. W. Reno
Frequency of isolation of aquatic birnaviruses in Canadian Provinces.
TitleFrequency of isolation of aquatic birnaviruses in Canadian Provinces
CaptionFrequency of isolation of aquatic birnaviruses in Canadian Provinces.
CopyrightP. W. Reno
Frequency of isolation of aquatic birnaviruses in Canadian Provinces.
Frequency of isolation of aquatic birnaviruses in Canadian ProvincesFrequency of isolation of aquatic birnaviruses in Canadian Provinces.P. W. Reno
Frequency of isolation of aquatic birnaviruses in other countries.
TitleFrequency of isolation of aquatic birnaviruses in other countries
CaptionFrequency of isolation of aquatic birnaviruses in other countries.
CopyrightP. W. Reno
Frequency of isolation of aquatic birnaviruses in other countries.
Frequency of isolation of aquatic birnaviruses in other countriesFrequency of isolation of aquatic birnaviruses in other countries.P. W. Reno

Identity

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Preferred Scientific Name

  • infectious pancreatic necrosis

International Common Names

  • English: aquabirnavirus infection; aquatic birnavirus infection; infectious pancreatic necrosis and associated aquatic birnaviruses

English acronym

  • IPN

Overview

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Infectious pancreatic necrosis (IPN) is a highly contagious viral disease of young fish of salmonid species held under intensive rearing conditions (Hill, 1982; Wolf, 1988; Wolf et al., 1960). The disease most characteristically occurs in rainbow trout (Oncorhynchus mykiss), brook trout (Salvelinus fontinalis), brown trout (Salmo trutta), Atlantic salmon (Salmo salar), and several Pacific salmon species (Oncorhynchus spp.). Susceptibility generally decreases with age, with resistance to clinical disease in salmonid fish usually being reached at about 1500 degree-days (value obtained by multiplying the age in days by the average temperature in degrees centigrade during the lifespan) (Dorson and Torchy, 1981), except for Atlantic salmon smolts, which can be affected normally within the first weeks after transfer from fresh water to seawater, but also before seawater transfer (Smail et al., 1989). The first sign of an outbreak in salmonid fry is frequently a sudden and usually progressive increase in daily mortality, particularly in the faster growing individuals. Clinical signs include darkening pigmentation, a pronounced distended abdomen and a corkscrewing/spiral swimming motion. Cumulative mortalities in a tank may vary from less than 10% to more than 90% depending on the combination of several factors, such as virus strain (McAllister and Owens, 1995) and infectious dose (Okamoto et al., 1984), host and environment (Dobos and Roberts, 1983). For further details see reviews by Hill (1982), Reno (1999) and Wolf (1988).

The disease is transmitted horizontally via the water route. Vertical transmission has been demonstrated for brook trout and rainbow trout, but this has never been successfully demonstrated in Atlantic salmon (Ahne et al., 1989; Ahne and Negele, 1985; Dorson and Torchy, 1981). Surface disinfection of eggs is not entirely effective in preventing vertical transmission (Bullock et al., 1976).

The disease has a wide geographical distribution, occurring in most major salmonid-farming countries of North and South America, Europe, Asia and Oceania (Anderson, 1996; Crane et al., 2000).

IPN virus (IPNV), or viruses showing serological relatedness to IPNV, have been reported to cause diseases in some farmed marine fish species, such as yellowtail (Seriola quinqueradiata) (Nakajima et al., 1993), turbot (Scophthalmus maximus) (Castric et al., 1987; Mortensen et al., 1990; Novoa et al., 1993b), dab (Limanda limanda) (Olesen et al., 1988), halibut (Hippoglossus hippoglossus) (Mortensen et al., 1990, Rodger and Frerichs, 1997) and Atlantic cod (Gadus morhua). Subclinical covert infections have been detected in a wide range of estuarine and freshwater fish species, such as loach (Misgurnus anguillicaudatus) (Chou et al., 1993), pike (Esox lucius) (Ahne, 1978) and numerous other species in the families Anguillidae, Atherinidae, Bothidae, Carangidae, Catostomidae, Cichlidae, Clupeidae, Cobitidae, Coregonidae, Cyprinidae, Esocidae, Moronidae, Paralichthydae, Percidae, Poecilidae, Sciaenidae, Soleidae and Thymallidae (Reno, 1999). It has been suggested to reserve the term IPNV for when the virus is isolated from a salmonid fish or has been shown to be able to produce IPN in salmonids through challenge experiments. Otherwise the term aquatic birnaviruses or aquabirnavirus is used.

The causative agent, IPNV, is the type species of the genus Aquabirnavirus of the Birnaviridae virus family. Birnaviruses have a bi-segmented (called segment A and B), double-stranded RNA genome, which is contained within a medium-sized, non-enveloped, single-shelled, icosahedral capsid (see review by Dobos 1995). Isolates display wide antigenic diversity (Dorson and Torchy, 1981; Hill and Way, 1995; Melby et al., 1994; Okamoto et al., 1983) and fall into two serogroups that do not cross-react in serum neutralisation tests (Ahne et al., 1989; Olesen et al., 1988; Underwood et al., 1977), with the majority of strains belonging to serogroup A, which comprises at least nine serotypes (Hill and Way, 1995). Isolates also show marked differences in degrees of virulence (Hill, 1982; Hill, 1992; McAllister and Owens, 1995).

Control methods rely on the implementation of control policies and of hygiene practices in salmonid husbandry, through the avoidance of the introduction of fertilised eggs originating from IPNV-carrier broodstock, and the use of a protected water supply (e.g. spring or borehole pond) where the ingress of fish, particularly possible virus carriers, is prevented. In outbreaks, a reduction in the population density (‘thinning out’) may help to reduce the overall mortality. There are vaccines available aimed to induce protection of Atlantic salmon smolts after seatransfer (Christie 1997) but there are few publication of results allowing for reliable judgement about vaccine efficacy, and little information is provided to validate the protection under field conditions.

Infectious pancreatic necrosis (IPN) is a disease which causes high mortalities in young salmonid fishes; the aetiological agent of the disease (IPNV) was the first virus to be isolated from teleosts (Wolf et al., 1960). In the time since that work, agents biochemically and serologically identical to IPNV - the aquatic birnaviruses - have been detected in a wide variety of both diseased and non-diseased fishes and other aquatic animals.

The early work on IPNV was mainly responsible for the development of fish virology. Techniques for the isolation of viruses from teleosts were modified from methods used for the isolation of mammalian viruses.

The virus itself has sufficiently unusual biophysical characteristics to warrant its place as the archetype of a new family of viruses, the birnaviruses (Dobos et al., 1979).

Serologically and biochemically related viruses induce a variety of disease syndromes in taxonomically diverse host groups, which also include aquatic invertebrates. The worldwide distribution of these viruses in cultured as well as wild fishes and shellfish provides significant potential for economic damage.

Infectious pancreatic necrosis and aquatic birnavirus-induced diseases can cause a significant economic impact on cultured fishes, especially salmonids. The cost can be as a consequence of lethal disease in young-of-the-year fish, or the destruction of infected stocks even in the absence of disease, or in Atlantic salmon smolts after sea transfer. The virus may be vertically transmitted; therefore, the detection of virus in broodstock often means the destruction of these animals. In the late 1980s, there were instances in the USA where 7 million young salmonids were destroyed to eliminate IPNV from a single hatchery (P. Walker, Colorado, 1988, personal communication) and others where detection of the virus in returning coho salmon resulted in the destruction of 2 million eggs (Olson et al., 1994); similar occurrences were also reported in Canada (B. Larson, Alberta, 1988, personal communication). In addition, regulations currently in place to reduce the dissemination of diseases of salmonids often result in restriction of the movement of IPNV carrier fish between local, regional and/or national boundaries. These restrictions have attendant adverse economic consequences to aquaculturists. In many instances, the slaughter of infected fish and/or the restriction of movement of fish may be more economically detrimental than the loss associated with direct mortalities.

In some countries there are mandatory destruction of salmonid stocks and restriction of movement after detection of IPNV. This is controversial. Since swim-up fry and fingerlings are generally affected, the impact of the disease can be ameliorated by spawning additional brood fish to compensate for the potential loss caused by the disease. Another perspective is the dissemination into watersheds of a virus unresponsive to therapeutic agents

which can readily infect susceptible native or cultured stocks in adjacent drainages. Given the myriad of variables which come into play in the generation of IPN and the potential for extensive, irreparable damage to stocks, it would seem prudent to adhere to more stringent rather than lax requirements. Resolution of the problem of IPNV management will be difficult and achieved only through a thorough knowledge of the epizootiology of the disease and an adequate database on both the disease and the virus.

Infectious pancreatic necrosis virus may also have an economic impact on aquaculture because of fish health inspections mandated prior to the movement or sale of fish across local, regional and national boundaries. Tests required for the detection of IPNV and other teleost viruses often represent an expense for aquaculturists. Again, this can be a contentious issue because the regulations are not often applied uniformly, even within regions regulated by one agency.

Control of the disease worldwide will also be difficult, given the ubiquitous nature of the agent.

Table. Natural diseases caused by, or associated with, aquatic birnaviruses. Included are isolations of aquatic birnaviruses in the absence of substantive proof of causal association with disease.


Diseases for which Rivers' postulates have been fulfilled

Disease Species Mortalities Behavioural signs Gross signs Microscopic lesions Ref
Infectious pancreatic necrosis Salmonids 0–95% Whirling, anorexia Blackening, abdominal distension, petechia in viscera, yellow exudate in gut Focal coagulative necrosis of exocrine and endocrine pancreas, kidney, intestine 1
Turbot haemopoietic necrosis Scophthalmus maximus Not reported Whirling, anorexia, lethargy, surface swimming Muscle haemorrhage, anaemic gills and liver, no food in gut Coagulative necrosis of haematopoietic elements of kidney, focal necrosis of liver 2
Yellowtail ascites disease Seriola quinqueradiata High None reported Ascites, catarrh, haemorrhage of liver, stomach and pyloric caeca; pale spleen, kidney, gills Coagulative necrosis of pancreatic acinar cells, kidney tubular epithelia, and hepatocytes 3
Eel nephritis Anguilla anguilla and Anguilla japonica 50–75% Body spasms and rigidity Retracted abdomen, reddening of anal fin, abdomen, gills; ascites, kidney enlargement Proliferative, exudative glomerulonephritis, tubular degeneration; focal necrosis of liver and spleen 4
Spinning disease of menhaden Brevoortia tyrranus Natural, unknown. Experimental, 100% Spinning Haemorrhage at fin base, gills, eyes; darkening Degeneration of basal layer of epidermis with pyknosis 5
Japanese flounder ascites Paralichthys olivaceus 5–60% None Ascites, cranial haemorrhage None reported 6

Diseases for which birnaviruses have been isolated from moribund animals

Disease Species Mortalities Behavioural signs Gross signs Microscopic lesions Ref
Seabass nephritis Dicentrarchus labrax 90% Spiral swimming Exophthalmia, distension of swim-bladder and gallbladder Delamination of intestinal epithelium 7
Striped bass mortality Morone saxatilis High Darting swimming; head up at rest None Degeneration of epidermal basal layer; pan-necrosis 8
Milkfish ulcer disease Chanos chanos 0 None Ulcerative necrosis of skin None described 9
Dab ascites Limanda limanda 0 None Ascites None described 10
Unnamed epizootic Paralichthys lethostigma Unknown Not reported Not reported Not reported 11
Eel stomatopapilloma Anguilla anguilla 0 None Stomatopapilloma Papilloma 12
Kumura shrimp disease Penaeus japonicus 26–43% Lethargy, difficulty in moulting Erosive necrosis of thoracic limbs and uropods None reported 13
Clam gill necrosis Meretrix lusoria 0 None Darkened gills Necrosis of gill tissue 14
Unnamed Tellina tenuis 0 None Chalky, brittle shell None reported 15

1, Wood et al., 1995; 2, Castric et al., 1987; 3, Sorimachi and Hara, 1985; 4, Sano et al., 1981; 5, Stephens et al., 1981; 6, Kusuda et al., 1989; 7, Bonami et al., 1988; 8, Schutz et al., 1984; 9, Chen et al., 1990; 10, Diamant et al., 1988; 11, McAllister et al., 1984; 12, Nagabayashi and Wolf, 1979; 13, Giorgetti, 1990; 14, Lo et al., 1991; 15, Hill et al., 1982.

Host Animals

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Animal nameContextLife stageSystem
Abramis brama (carp bream)Subclinical
Anguilla anguilla (European eel)Subclinical
Anguilla japonica (Japanese eel)Subclinical
Anguilla rostrata (american eel)Subclinical
Ardea cinereaExperimental settings
Astacus astacus (European crayfish)Experimental settings
Barbonymus schwanenfeldii (tinfoil barb)Subclinical
Barbus barbusSubclinical
Blicca bjoerknaSubclinical
Brachionus plicatilisSubclinical
Brevoortia tyrannusSubclinical
Bubo virginianusExperimental settings
Carassius auratus auratus (goldfish)Subclinical
Carassius carassius (crucian carp)Subclinical
Carcinus maenas (European shore crab)Subclinical
Channa striata (snakehead murrel)Subclinical
Chanos chanos (milkfish)
Chondrostoma nasusSubclinical
Corbicula fluminea (Asian clam)Subclinical
Crassostrea gigas (Pacific oyster)Subclinical
Crassostrea virginica (eastern oyster)Subclinical
Cyprinus carpio (common carp)Subclinical
Danio rerio (zebra danio)Subclinical
Dicentrarchus labrax (European seabass)Subclinical
Dorosoma cepedianumSubclinical
Esox lucius (pike)Subclinical
Esox niger (chain pickerel)Subclinical
Gadus morhua (Atlantic cod)Domesticated hostEnclosed systems/Cages|Enclosed systems/Tanks
Gallus gallus domesticus (chickens)Experimental settings
Gobio gobio (gudgeon)Subclinical
Hampala disparSubclinical
Hippoglossus hippoglossus (Atlantic halibut)Domesticated hostEnclosed systems/Cages|Enclosed systems/Tanks
Hucho hucho (huchen)Subclinical
Lampetra fluviatilisSubclinical
Larus ridibundusExperimental settings
Leiostomus xanthurusSubclinical
Limanda limandaDomesticated host
Littorina littorea (common periwinkle)Subclinical
Macropipus depuratorSubclinical
Marsupenaeus japonicus (kuruma shrimp)Subclinical
Menidia menidiaSubclinical
Mercenaria mercenaria (northern quahog)Subclinical
Meretrix lusoria (common oriental clam)Subclinical
Misgurnus anguillicaudatus (oriental weatherloach)Subclinical
Morone saxatilis (striped sea-bass)Subclinical
MustelaExperimental settings
Mytilus edulis (common blue mussel)Subclinical
Oncorhynchus clarki clarki (cutthroat trout)Domesticated hostAquatic: FryEnclosed systems/Cages|Enclosed systems/Pens
Oncorhynchus gorbuscha (pink salmon)Domesticated hostAquatic: FryEnclosed systems/Cages|Enclosed systems/Pens
Oncorhynchus keta (chum salmon)Domesticated hostAquatic: FryEnclosed systems/Cages|Enclosed systems/Pens
Oncorhynchus kisutch (coho salmon)Domesticated hostAquatic: FryEnclosed systems/Cages|Enclosed systems/Pens
Oncorhynchus mykiss (rainbow trout)Domesticated hostAquatic: All StagesEnclosed systems/Cages|Enclosed systems/Pens|Enclosed systems/Raceways / running water ponds
Oncorhynchus tshawytscha (chinook salmon)Domesticated host, Wild hostAquatic: FryEnclosed systems/Cages|Enclosed systems/Pens
Oreochromis mossambicus (Mozambique tilapia)Subclinical
Ostrea edulis (European oyster)Subclinical
Oxyeleotris marmorataSubclinical
Paralichthys lethostigma (southern flounder)Subclinical
Patella vulgataSubclinical
Perca fluviatilis (perch)Subclinical
Phoxinus phoxinus (European minnow)Subclinical
Prosopium williamsoniSubclinical
Psetta maxima (turbot)Domesticated hostEnclosed systems/Other enclosed systems
Rutilus rutilus (roach)Subclinical
Salmo salar (Atlantic salmon)Domesticated host, Wild hostAquatic: Adult|Aquatic/FryEnclosed systems/Cages|Enclosed systems/Pens
Salmo trutta (sea trout)Domesticated hostAquatic: FryEnclosed systems/Cages|Enclosed systems/Pens
Salvelinus alpinus (Arctic charr)Domesticated hostAquatic: FryEnclosed systems/Cages|Enclosed systems/Pens
Salvelinus fontinalis (brook trout)Domesticated hostAquatic: FryEnclosed systems/Cages|Enclosed systems/Pens
Salvelinus namaycush (lake trout)Domesticated hostAquatic: FryEnclosed systems/Cages|Enclosed systems/Pens
Scardinius erythrophthalmus (rudd)Subclinical
Seriola quinqueradiata (japanese amberjack)Domesticated hostEnclosed systems/Cages
Symphysodon discusSubclinical
Tellina tenuisSubclinical
Thymallus thymallus (grayling)Subclinical
Xiphophorus xiphidiumSubclinical

Hosts/Species Affected

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Early reports of IPNV were limited to epizootics in cultured brook trout, Salvelinus fontinalis (Wood et al., 1955; Snieszko et al., 1957, 1959; Wolf et al., 1960). With the development of several continuous cell lines from teleost fishes (Wolf and Quimby, 1962; Wolf and Mann, 1980) and as more laboratories became more proficient in the use of cell lines, it was found that IPNV was responsible for disease in a variety of salmonid species, including members of the genera Salmo, Salvelinus and Oncorhynchus. During the early 1970s, IPN caused high mortalities in European and Japanese rainbow and brown trout (Ball et al., 1971; Sano, 1971; Vestergård-Jørgensen and Kehlet, 1971).

The first report of birnavirus-induced disease in non-salmonids was in Japanese eels (Anguilla japonica), first sampled in 1969 (Sano et al., 1981). This disease of cultured eels was characterized as branchionephritis. Experimental infection with the virus (eel virus European (EVE)) indicated that the branchial lesions were probably due to supervening bacterial infections and that the disease should be referred to as eel nephritis. Neutralization studies indicated that the aetiological agent was a birnavirus which is serologically related to the Sp (from Spjarup, Denmark) serotype of IPNV. Another birnavirus-induced disease which caused mortalities in Japanese fish caused ascites in yellowtail, Seriola quinqueradiata (Sorimachi and Hara, 1985). The condition was in feral fry used as seed stock for mariculture. Because clinical signs occurred in fry, the disease is similar to IPN. In addition, mortalities occurred in another flat-fish species from Japan, the Japanese flounder (Paralichthys olivaceus). Fish held in culture facilities died from ascites and cranial haemorrhage. An aquatic birnavirus was isolated from moribund fish and an ascitic disease was experimentally induced by intraperitoneal injection of the virus from moribund fish showing either disease syndrome (Kusuda et al., 1989).

Stephens et al. (1983) isolated an aquatic birnavirus from the brain and other tissues of menhaden (Brevoortia tyrranus) with ‘spinning disease’, a perennial high-mortality disease in the Chesapeake Bay region of the eastern USA. The virus was isolated in a cell line from the same species (Stephens, 1981). The disease was experimentally induced in susceptible menhaden after intraperitoneal injection of the virus. Subsequent attempts to isolate the same virus from diseased menhaden in other teleost continuous cell lines were unsuccessful. Since the original menhaden cell line was lost, there have been no further isolations of the virus from diseased or non-diseased fish.

In each of these cases, Rivers’ postulates have been fulfilled, indicating that the aetiological agents of the diseases were aquatic birnaviruses which cross-reacted serologically with IPNV. In several other instances, birnaviruses have been isolated from aquatic animals undergoing epizootic disease, but as yet there has not been irrefutable evidence that the isolated birnaviruses were causal agents.

A birnavirus infection was associated with haematopoietic necrosis and caused high mortalities in turbot (Scophthalmus maximus) held at 18°C in sea water on the coast of France (Castric et al., 1987). Within 15 days after transfer from fresh water to sea water, cumulative mortalities reach 25%, but no mortalities occurred in the same population of fish held at the freshwater site. No definitive studies have been done to confirm Rivers’ postulates for this disease and a birnavirus is suspected.

In addition to a birnavirus isolated from Japanese eels with nephritis (Sano et al., 1981), a birnavirus was also isolated in Japan from eels exhibiting stomatopapillomas (Nagabayashi and Wolf, 1979). Although the virus was isolated from moribund fish, there was no indication that the virus was capable of producing the papillomatous lesions in homologous animals.

Transmitting papillomas in homeotherms is often difficult, and the strong association of the papilloma group of viruses with this disease in other animals makes it unlikely that a birnavirus was the causal agent in eels. In retrospect, it is possible that the eels affected with the stomatopapillomas were adventitious carriers of the virus (Sano et al., 1981) and the isolated aquatic birnavirus had no involvement in the development of papillomas.

An aquatic birnavirus belonging to the Sp (European) serotype was isolated from visceral organs of snakehead fish (Ophicephalus striatus) and eyespot barb (Hampala dispar) undergoing episodes of ulcerative disease in Thailand and Laos (Wattanavijarn et al., 1988). Again, there is no further information to indicate that this agent was responsible for the syndrome.

Another birnavirus was isolated from sea bass (Dicentrarchus labrax) off the coast of France (Bonami et al., 1983) during a severe epizootic, in which mortalities reached 95%. There was no evidence of bacterial infection or parasites in the moribund fish, but again it is uncertain whether the virus is merely associated with the mortality or was the aetiological agent.

With the rather large exception of IPN disease, the disease syndromes that have been strongly associated with aquatic birnaviruses (yellowtail ascites, Japanese flounder disease syndromes, eel nephritis, spinning disease of menhaden and turbot renal necrosis) occur in marine animals or in those that spend the bulk of their life in seawater. In the intensive marine farming of Atlantic salmon there has likewise been an increase of the occurrence of IPN-virus and IPN outbreaks in post-smolts in sea farms through the second half of the 1980s (Willumsen 1988, Bruheim 1991). Both Jarp (1999) and Brun (2001) have shown that the median time from sea transfer to IPN outbreak in post smolts is about 35 days (range 3-75, Brun 2001). Melby (1991) found that 63.7% out of 1939 individual post-smolts from 608 sea farms of Atlantic salmon in Norway were positive for IPNV.

It is likely that diseases caused by aquatic birnaviruses are more extensive in nature because reports of this family of viruses are primarily in cultured animals, which have economic importance. Relatively little information is available regarding birnavirus infections in wild populations of aquatic animals that are not of commercial importance and this is unlikely to change unless some of these animals develop commercial importance.

Table. Species of aquatic animals from which birnaviruses have been isolated.


ClassFamilyGenus and speciesCommon nameRef
Monogonta(Order) PloimaBranchionus plicatilisRotifer27
MolluscaVeneridaeMeretrix lusoriaEnamel venus shell1
 VeneridaeCorbicular fluminouAsian clam30
 AcmaeidaePatella vulgataScallop2
 LittorinidaeLittorina littoreaPeriwinkle2
 MytilidaeMytilus edulisMussel2
 OstreidaeCrassostrea virginicaAmerican oyster2
 OstreidaeC. gigasJapanese oyster2
 OstreidaeOstrea edulisEuropean oyster2
 TellinidaeTellina tenuisTellina3
 VeneridaeMercenaria mercenariaSurf clam2
CrustaceaPortunidaeCarcinus maenasShore crab2
 PortunidaeMacropipus depuratorHarbour crab2
 PenaeidaePenaeus japonicusJapanese shrimp4
AgnathaPetromyzontidaeLampetra fluviatilisLamprey5
TeleosteiClupeidaeBrevoortia tyrranusMenhaden6
 ClupeidaeDorosoma cepedianumGizzard shad25
 AnguillidaeAnguilla japonicaJapanese eel7
 AnguillidaeA. rostrataAmerican eel25
 AnguillidaeA. anguillaEuropean eel8
 EsocidaeEsox luciusNorthern pike9
 EsocidaeE. nigerChain pickerel10
 SalmonidaeHucho huchoGrayling10
 SalmonidaeOncorhynchus mykissRainbow trout11
 SalmonidaeO. ketaChum salmon12
 SalmonidaeO. kisutchCoho salmon13
 SalmonidaeO. clarkiCutthroat trout12
 SalmonidaeO. tshawytchkaChinook salmon12
 SalmonidaeO. gorbuschaPink salmon12
 SalmonidaeO. rhodurusAmago trout14
 SalmonidaeSalvelinus fontinalisBrook trout15
 SalmonidaeS. naymacushLake trout16
 SalmonidaeS. alpinusArctic char12
 SalmonidaeSalmo salarAtlantic salmon16
 SalmonidaeS. truttaBrown trout17
 SalmonidaeThymallus thymallusGrayling18
 SalmonidaeProsopium williamsoniWhitefish28
 ChannidaeOphicephalus striatusSnakehead fish18
 CyprinidaeAbramis bramaBream2
 CyprinidaeBarbodes schwanefeldiBarb10
 CyprinidaeBlicca bjoerknaDace17
 CyprinidaeCarassius auratusGoldfish20
 CyprinidaeC. ceratisinisCarp10
 CyprinidaeC. carassiusCarp2
 CyprinidaeChondrostoma nasusNase10
 CyprinidaeCyprinus carpioCommon carp2
 CyprinidaeGobio gobioGoby10
 CyprinidaeOxyeleotris marmoratusSand goby29
 CyprinidaeHampala disparEyespot barb18
 CyprinidaePhoxinus phoxinusDace5
 CyprinidaeRutilus rutilusRoach2
 CyprinidaeScardinius erythrophthalmusRudd10
 CyprinidaeBarbus barbusBarbel10
 CyprinidaeBrachydanio rerioZebra danio10
 CobitidaeMisgurnus anguillicaudatusRoach20
 AtherinidaeMenidia menidiaSilversides22
 SciaenidaeLeiostomus xanthurusDrum22
 CarangidaeSeriola quinqeradiataYellowtail21
 CichlidaeSymphysodon discusDiscus22
 CichlidaeTilapia mossambicaTilapia10
 PercidaePerca fluviatilisYellow perch5
 PercichthyidaeDicentrarchus labraxSea bass29
 PercichthyidaeMorone saxatilisStriped bass24
 PoeciliidaeXiphophorus xiphidiumPlaty10
 GadidaeGadus morhuaAtlantic cod2
 PleuronectidaeLimanda limandaCommon dab26
 PleuronectidaeParalichthys lethostigmaSouthern flounder25
 PleuronectidaePleuronectes fluviatisSole2
 PleuronectidaeHippoglossus hippoglossusHalibut29
 PleuronectidaeScophthalmus maximusTurbot25

1, Lo et al., 1991; 2, Hill et al., 1982; 3, Hill, 1976; 4, Bovo et al., 1984; 5, Munro et al., 1976; 6, Stephens et al., 1983; 7, Sano et al., 1981; 8, Hudson et al., 1981; 9, Ahne, 1979; 10, Ahne et al., 1989b; 11, Parisot et al., 1963; 12, Wolf, 1988; 13, Wolf and Pettijohn, 1970; 14, Sano, 1973; 15, Hedrick et al., 1986a; 16, MacKelvie and Artsob, 1969; 17, Wolf et al., 1960; 18, Wattanavijarn et al., 1988; 19, Ahne, 1980; 20, Hill, 1982; 21, Chen et al., 1984; 22, Sorimachi and Hara, 1985; 23, Adair and Ferguson, 1981; 24, Wechsler et al., 1986b; 25, McAllister et al., 1984; 26, Castric et al., 1987; 27, Comps et al., 1991; 28, Yamamoto and Kilistoff, 1979; 29, Bonami et al., 1983; 30, T. Håstein, personal communication.

Distribution

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Disease

While the disease was evidently first described from the Maritime Provinces, Canada (M’Gonigle, 1940), and thereafter in the eastern USA (Wood et al., 1955), it was later noted in many areas of the western USA (Yasutake et al., 1965), especially in rainbow trout and brook trout transferred from the eastern USA. Epizootics in rainbow trout with signs of IPN disease reported in France (Besse and de Kinkelin, 1965) were caused by an aetiological agent which was serologically distinct from the North American serotype (Wolf and Quimby, 1971). The detection of IPN in France was followed by diagnosis of disease in rainbow trout in Denmark (Vestergård-Jørgensen and Bregnballe, 1969), Norway (Håstein and Krogsrud, 1976), Sweden (Ljungberg and Vestergård-Jørgensen, 1973), the UK (Ball et al., 1971), Germany (Schlotfeldt et al., 1975) and Italy (Ghittino, 1972). Outside Europe, the disease has been documented in the Far East, usually in fish imported from enzootic areas or in fish cohabiting the same water source as infected imported fish. The virus has been isolated from several countries like Japan (Sans, 1971), Korea (Hah et al., 1984), Taiwan (Hedrick et al., 1983), China (Jiang et al., 1989), Thailand and Laos (Wattanavijarn et al., 1988), Australia (Crane et al., 2000), New Zealand (Anderson, 1996) and Turkey (Candan, 2002).

Virus

The isolation of aquatic birnaviruses from various geographical areas has increased markedly since the isolation of IPNV from the eastern USA (Wolf et al., 1960). In a few instances, there has been sufficient epizootiological evidence to support the orderly ‘spread’ of the virus from region to region. When the viruses were serotyped, they have in most cases proved to be serologically identical to serotypes of IPN from epizootics (Caswell-Reno et al., 1986; Hill and Way, 1995).

The aquatic birnaviruses have been isolated from regions where salmonids are reared and from areas without salmonids. The data are often difficult to access and summarize easily because of the inconsistent nature of the published literature and the variety of sources from which samples have been taken. The presence of these viruses in feral animals is a confounding factor. Since little current information about the distribution of aquatic birnaviruses is available in the literature, an attempt has been made in this review to update the information.

Prior to the late 1980s, the virus was found to be present in most areas of North America. In many areas, the prevalence levels were low, but in some areas (Maritime Canada, Pennsylvania, West Virginia and some north-eastern US states) a large number of hatcheries were contaminated. However, because of increased awareness of vertical transmission of the virus and regulations on movement of fish eggs, the majority of states and provinces have significantly reduced or eliminated the virus from hatcheries. There has been a marked reduction in reports of the virus, especially in the western USA and eastern Canada. This may be due to the reduction in rearing of brook trout, which serve as reservoirs for the virus in the western states.

Unfortunately, the situation in Europe and Asia is not as positive. The virus has been found in virtually every European country (de Kinkelin, 1989, personal communication), with almost total contamination of facilities in some countries (Melby et al., 1991). The pervasive dispersal of the virus in Europe may be due to the transfer of contaminated stocks and an inability to start with virus-free stocks without destroying the industry. The presence of IPNV in trout is often considered an impediment to the industry, not because of the consequences of disease but rather because of the restriction of fish movements (Ghittino, 1982).

The virus and epizootic outbreaks of the disease are widespread in salmonids in Asia, where importations from other countries, especially the USA and Europe, have been extensive. Most probably, the increased detection of IPNV in this region reflects an increased dispersal of the virus, an increased ability of fish health workers to isolate the viruses, and the examination of a wider variety of aquatic animals for birnavirus infection.

Iceland is one of the few countries which raise salmonids on a large scale and where extensive fish health inspections are routinely performed that have no reports of IPN. The country has very strict importation policy for fin-fish. However, birnavirus serogroup B has been isolated from Icelandic farmed halibut (Hippoglossus hippoglossus) (Skall et al., 2000).

The serotypes of aquatic birnavirus serogroup A are called A1 to A9, but are also known as West Buxton, Sp, Ab, He, Te, Can 1, Can 2, Can 3 and Jasper. There are antigenic cross-reactions between the viruses in serogroup A, and the criterion for each of these 9 serotypes being a serotype is suggestions based on calculations of results in neutralization tests (Hill and Way, 1995).The serotype A1 (WB archetype from brook trout in West Buxton, Maine, USA) is found extensively throughout the USA and on many occasions in Canada. However, the Canadian serotypes have rarely been isolated in the USA (Caswell-Reno et al., 1990). The isolations of A1 viruses from other countries have almost always been traced to the importation of specific batches of contaminated eggs or fish from the USA (Sano, 1971; McAllister and Reyes, 1984; Bragg and Combrink, 1987a). Interestingly, the A1 serotype has only been isolated from Europe in a single instance - from turbot in Spain (Novoa et al., 1993a) - although trout have been extensively transferred to Europe from the USA. Serotypes A2 (Sp) and A3 (Ab from Abild, Denmark) have been disseminated widely not only throughout Europe, but also to many countries in Asia. Serotype A2 has been detected only once in feral fish in marine waters off the coast of the USA (McAllister et al., 1984). The four serotypes Canada 1, 2 and 3 and Jasper, Alberta, are restricted to Canada, except for a few recent isolations from facilities in watersheds that are contiguous in the USA and Canada (Caswell-Reno et al., 1990). Although fish have been exported from Canada to many other countries, none of the Canadian 1, 2 or 3 serotypes have been found outside of Canada or its borders with the USA. Thus, the distribution of serotypes does not seem to support the hypothesis that the aquatic birnaviruses are spread from continent to continent with the importation of fish or fish eggs. It is likely that the virus had a global distribution prior to the widespread dissemination of salmonids in the nineteenth and the early twentieth century.

Distribution Table

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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/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Sea Areas

Atlantic, NorthwestPresentM'Gonigle, 1940; Wood et al., 1955
Pacific, Eastern CentralPresentYasutake et al., 1965
Pacific, NorthwestPresentSano, 1971; Hedrick et al., 1983; Hah et al., 1984; Jiang et al., 1989
Pacific, Western CentralPresentWattanavijarn et al., 1988

Asia

ChinaPresentJiang et al., 1989
JapanPresentSano, 1971
Korea, Republic ofPresentHah et al., 1984
LaosPresentWattanavijarn et al., 1988
TaiwanPresentHedrick et al., 1983
ThailandPresentWattanavijarn et al., 1988
TurkeyPresentCandan, 2002

Africa

South AfricaPresentReno, 1999

North America

CanadaPresentM'Gonigle, 1940
-AlbertaPresentReno, 1999
-British ColumbiaPresentReno, 1999
-ManitobaPresentReno, 1999
-New BrunswickPresentReno, 1999
-Newfoundland and LabradorPresentM'Gonigle, 1940
-Northwest TerritoriesPresentReno, 1999
-Nova ScotiaPresentReno, 1999
-OntarioPresentReno, 1999
-Prince Edward IslandPresentReno, 1999
-QuebecPresentReno, 1999
USAPresentWood et al., 1955; Yasutake et al., 1965
-AlabamaPresentReno, 1999
-ArizonaPresentReno, 1999
-ArkansasPresentReno, 1999
-CaliforniaPresentReno, 1999
-ColoradoPresentReno, 1999
-ConnecticutPresentReno, 1999
-FloridaPresentReno, 1999
-GeorgiaPresentReno, 1999
-IdahoPresentReno, 1999
-IllinoisPresentReno, 1999
-MainePresentReno, 1999
-MarylandPresentReno, 1999
-MassachusettsPresentReno, 1999
-MichiganPresentReno, 1999
-MinnesotaPresentReno, 1999
-MontanaPresentReno, 1999
-NevadaPresentReno, 1999
-New HampshirePresentReno, 1999
-New JerseyPresentReno, 1999
-New MexicoPresentReno, 1999
-New YorkPresentReno, 1999
-North CarolinaPresentReno, 1999
-North DakotaPresentReno, 1999
-OregonPresentReno, 1999
-PennsylvaniaPresentReno, 1999
-Rhode IslandPresentReno, 1999
-South CarolinaPresentReno, 1999
-South DakotaPresentReno, 1999
-TennesseePresentReno, 1999
-TexasPresentReno, 1999
-UtahPresentReno, 1999
-VermontPresentReno, 1999
-VirginiaPresentReno, 1999
-WashingtonPresentReno, 1999
-West VirginiaPresentReno, 1999
-WisconsinPresentReno, 1999

Europe

BelgiumPresentReno, 1999
Czechoslovakia (former)PresentReno, 1999
DenmarkPresentVestergård-Jørgensen and Bregnballe, 1969
FinlandPresentReno, 1999
FrancePresentBesse and de, 1965
GermanyPresentSchlotfeldt et al., 1975
ItalyPresentGhittino, 1972
LuxembourgPresentReno, 1999
NorwayPresentHastein and Krogsrud, 1976
SpainPresentReno, 1999
SwedenPresentLjungberg and Vestergård-Jørgensen, 1973
UKPresentBall et al., 1971
Yugoslavia (Serbia and Montenegro)PresentReno, 1999

Oceania

AustraliaPresentReno, 1999
-TasmaniaPresentCrane et al., 2000
New ZealandPresentTisdall and Phipps, 1987; Reno, 1999

Diagnosis

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Clinical Diagnosis

Infectious pancreatic necrosis virus causes mortality of fry and fingerling salmonids and Atlantic salmon smolts, and is characterized by behavioural changes and gross external, internal and histopathological lesions. There are no specific pathognomonic signs of IPN disease. Behavioural changes (Wood et al., 1955) include anorexia and an agonal corkscrew swimming motion interspersed with ataxia. Nonspecific external signs include hyperpigmentation, exophthalmia and petechial haemorrhage on the ventral surfaces. Internal gross lesions are visceral petechia and an empty gut containing a yellow exudate. The disease may be manifested with only a few, or even none, of these signs. Microscopically, there is focal necrosis of the acinar and islet cells of the pancreas and of the haematopoietic cells of the kidney. There are typical icosahedral virus particles in the cytoplasm of pancreatic acinar cells (Lightner and Post, 1969; Hedrick et al., 1985). Viral titres in the tissues of infected fish are usually quite high, usually in the range of 107-1010 (TCID50) g-1 (Wolf et al., 1969; Castric et al.,1987).

Survivors of epizootics or those fish with no disease have few effects of residual infection, although high titres of virus can be isolated from their viscera. In carrier Atlantic salmon there are interstitial cells in the lamina propria of the gut which appear necrotic, the so-called ‘McKnight cells’ (McKnight and Roberts, 1976). Studies examining the effect of IPNV infection on Atlantic salmon which evince no clinical signs of IPN indicate that there is no adverse effect on growth rate or susceptibility to other diseases (Smail et al., 1986).

Several studies have demonstrated that there are markedly variable virus titres in carrier fish. Yamamoto (1975b) found that rainbow trout had IPNV titres in the viscera ranging from 100.85 to 104.2 TCID50 g-1, while they ranged from 100.85 to 106.5 TCID50 g-1 in brook char (Reno, 1976).

The virulence of IPNV isolates is quite variable. Most have been isolated from aquatic animals with no evidence of disease. Only a few isolates from nondiseased animals have been tested for virulence.

Table. Virulence of aquatic birnaviruses in experimental infections of homologous and heterologous hosts. Serotype of virus, if reported, in parentheses.


Isolate fromSpecies infectedSize of hostRoute of exposureDose per fish (TCID50)Specific mortality (%)Duration of observationRef
Homologous species infected       
Salvelinus fontinalisS. fontinalis'Fry'IMMNot reported7-9814 d1
S. fontinalisS. fontinalis (NB)0.5 gIMM/15 min106.9/ml31 y2
S. fontinalisS. fontinalis (Penn-2)5 moIP106.3025 d3
S. fontinalisS. fontinalis (VR-299)6 moIP106.3025 d3
S. fontinalisS. fontinalis (VR-299)14 moIP106.3025 d3
Oncorhynchus mykissO. mykiss0.11 gIMM/1 h104.81590 d4
O. mykissO. mykiss (Sp)5-7 cmIMM105.02021 d5
O. mykissO. mykiss (Ab)5-7 cmIMM105.0021 d5
O. mykissO. mykiss (Ab)0.2 gIMM104935 d6
O. mykissO. mykiss (Sp)0.2 gIMM1044235 d6
Salmo salarS. salarFryIMM105-1080-1530 d7
Brevoortia tyrranusB. tyrranus5-25 cmIP1051007 d8
Anguilla japonicaA. japonica10 gIP107.85.5-75NR4
Paralichthys olivaceusP. olivaceus5 gIPNot reported6021 d9
Misgurnus anguillicaudatusM. anguillicaudatus10-15 gIP104757 d11
Morone saxatilisM. saxatilis5 d-6 moOral, IMM102-106033 w12
Seriola quinqueradiataS. quinqueradiata      
Oncorhynchus mykissSalvelinus fontinalis (Sp)25 mmIMMNot reported42-5120 d12
O. mykissSalmo salar (Sp)Fry-parrIMM105-080-1.530 d7
O. mykissO. tshawytscha2-3 inSC103100< 6 d13
O. clarkiO. nerka'Juvenile'Not reportedNot reported0Not known14
O. clarkiO. kisutch'Juvenile'Not reportedNot reported0Not known14
O. clarkiO. tshawytscha'Juvenile'Not reportedNot reported0Not known14
Anguilla japonicaO. mykiss0.11 gIMM/1 h105.1090 d4
Esox nigerS. fontinalisNot reportedIMM/6 h10616.6-4014 d15
E. nigerS. namaycushNot reportedIMM/6 h1061.614 d15
E. luciusO. mykiss (Hecht)0.2 gIMM104035 d6
Morone saxatilisS. fontinalis20 dIMM1059460 d16
Rutilus rutilusO. mykissFryIMM1041030 d17
Perca fluviatilisO. mykissFryIMM104630 d17
Cyprinus carpioO. mykissFryIMM1046030 d17
Salvelinus fontinalisO. mykiss100 gIP106084 d1
Paralichthys lethostigmaS. fontinalisFryNot reportedNot reported'Slightly virulent'Not reported10
A. japonicaMisgurnus anguillicaudatus (EVE)10-15 gIP104207 d11
Ostrea edulisCrassostrea gigas1-2 cmIMM1041850 d18
Crassostrea virginicaOstrea edulis5-6 cmIMM1043450 d18
Tellina tenuisCrassostrea gigasNot reportedIMMNot reported090 d19
S. fontinalisHippocampus erectusNot reportedIP105034 d20
S. fontinalisHaemulon flavolineatumNot reportedIP105022 d20

NR, not reported; IMM, immersion; IP, intraperitoneal; SC, subcutaneous; d, days; w, weeks; mo, months; y, years.

1, Wolf and Quimby, 1969; 2, Reno et al., 1978; 3, Swanson et al., 1982; 4, Sano et al., 1981; 5, Vestergård-Jørgensen and Kehlet, 1971; 6, Kohlmeyer et al., 1986; 7, Smail et al., 1986; 8, Stephens et al., 1983; 9, Kusuda et al., 1989; 10, McAllister et al., 1984; 11, Chen et al., 1984; 12, Wechsler et al., 1987b; 13, Klontz et al., 1965; 14, Parisot et al., 1963; 15, Silim et al., 1982; 16, McAllister and McAllister, 1988; 17, Hill, 1982; 18, Hill et al., 1982; 19, Hill, 1976; 20, Moewus-Kobb, 1965.

 

Wolf et al. (1969) demonstrated differences in virulence with 15 serotype A1 (WB) isolates by infecting brook trout fry. Mortalities ranged from 7 to 98%. The viral titre in the mortalities was approximately 109 g-1; there was no correlation between the titre and mortality levels (regression analysis of data of Wolf et al., 1969). Unfortunately, the titre of virus in the more resistant or in surviving fish was not reported in rainbow trout infected with three European strains: Sp, Ab and Hecht (He). Hecht was avirulent, while the two trout isolates were causing significant mortality, albeit at different levels; virulence was directly correlated with plaque size (Kohlmeyer et al., 1986). Only a few passages of virulent Sp serotype virus in cell culture may cause attenuation of the virus (Santi et al., 2004) This variation in virulence is a reflection of the complex nature of the disease, which is not well understood. The two European serotypes were originally classified as virulent for trout (Sp) or avirulent for trout (Ab) (Vestergård-Jørgensen and Kehlet, 1971).

Predisposing factors for IPN include temperature, age and stress (Frantsi and Savan, 1971a; LaPierre et al., 1988). Six-month-old brook trout at 3 g are resistant to clinical IPN disease; however, it is not known whether the host had cleared the virus entirely or whether residual virus was present, as in typical IPN carriers. Temperatures at which maximum mortalities occur vary, but temperature effects may be obscured by the strain of virus and host species used: e.g. brook trout infected with a Canadian isolate had maximal mortality at 10°C, while those infected with IPNV VR-299 succumbed most readily at 15.5°C (Frantsi and Savan, 1971a). In the epizootic affecting turbot, mortalities were markedly higher in juvenile fish held in sea water at 18°C than at 11°C (Castric et al., 1987). Similar results were obtained by Okamoto et al. (1987b) in rainbow trout fry.

Nephritis in eels, caused by a birnavirus termed eel virus European (EVE), occurs primarily during the winter, when water temperatures are lowest (Sano et al., 1981). The compromised physiological condition of the fish during this period may be a factor in the disease process (Egusa et al., 1971; Oka et al., 1976).

The severity of the disease is dependent not only on the species of fish, but also on the strain of fish within a species. Silim et al. (1982) demonstrated that strains of brook trout from four genetic pools varied in their susceptibility to IPN, with mortalities ranging from 40 to 85%. Similarly, Okamoto et al. (1987a) found that rainbow trout from one hatchery (strain RT-101) were consistently more susceptible than fish from two other hatcheries to lethal IPNV infection over a course of 6 years. Additionally, these authors demonstrated that the quantity of virus to which fish were exposed influenced mortalites. No mortalities occurred in 0.26 g fish exposed to less than 102 TCID50 ml-1 of IPNV-Buhl, whereas 61% of fish died when exposed to 1000-fold higher virus concentrations. The relationship between infection of salmonids and severity of clinical disease is complicated and the IPNV carrier state has been difficult to elucidate.

The diseases known as yellowtail ascites, Japanese flounder ascites and eel nephritis are caused by aquatic birnaviruses. All are severe and cause high mortalities in cultured flat-fish and eels in Japan. In addition, three more diseases have been found to be associated with birnaviruses, although unequivocal proof of their involvement in the promulgation of the diseases is lacking. These diseases cause sea-bass mortalities (Bonami et al., 1983), turbot haematopoietic necrosis (Castric et al., 1987) and spinning disease of menhaden (Stephens et al., 1983).

Diagnostic methods

The diagnosis of IPN has historically been predicated on clinical signs of the disease, isolation and identification of the aetiological agent by cell culture methods and confirmation using serological methods. Consistent detection of aquatic birnaviruses has proved to be simple using cell culture systems. There are at least five reasons for this.

1. The virus is usually present in relatively high titres in tissues.

2. Unlike other fish viruses, such as infectious haematopoietic necrosis (IHN) and channel catfish virus (CCV), isolations of aquatic birnaviruses are made most frequently from non-diseased animals.

3. There is no latent stage, in which the virus cannot be isolated in cell culture.

4. The time required for isolation and identification of the agent is usually 2-3 weeks.

5. Easily available continuous teleost cell lines are generally highly sensitive to the virus and undergo a characteristic, readily observable cytopathic effects (CPE) when infected.

Thus, isolation of virus from visceral samples in standardized teleost cell lines has remained the regular diagnostic method.

Cell culture methods: primary isolation

Many teleost cell lines are susceptible to IPNV infection, with the consequent production of characteristic CPE, usually manifested as a ‘stringy’ appearance of the cells.

Table Continuous teleost cell lines in which aquatic birnaviruses have been propagated.


Cell lineSpeciesVirus yieldRef
RTG-2Oncorhynchus mykiss108.51
RFO. mykissNot reported2
RTH-149O. mykiss109.33
RTF-1O. mykissNot reported4
STE-137O. mykiss109.53
CHSE-214O. tshawytscha109.73
CHSE-114O. tshawytscha109.23
CSE-119O. kisutch108.83
CHH-1O. keta1010.03
SSE-5O. nerka108.83
SSE-30O. nerka108.83
KO-6O. nerka108.83
BBIctalurus nebulosusNot reported4
ASSalmo salar108.05
CCTCyprinus carpio109.06
LFMisgurnus anguillicaudatus107.336
TO-2Tilapia mossambica1010.336
INEM-1Stenodus leucichthys109.97
SWTXiphophorus helleri108.08
GF-1Haemulon sciurus107.09
FHMPimephales promelas0–107.71
OMAKACaranx mate10210
BF-2Lepomis macrochirusNot reported11
PGEsox luciusNot reported12
SJU-1Carassius auratus109.513
CLOphicephalis luciusNot reported14
WC-1Stizostedion vitreumNot reported15
MKBrevoortia tyrannusNot reported16
EPCCyprinus carpio108.517
EK-1Anguilla japonica109.518

1, Kelly et al., 1978; 2, Wolf and Mann, 1980; 3, Lannan et al., 1984; 4, Wolf and Quimby, 1969; 5, Piper et al., 1973; 6, Chen et al., 1990; 7, Follett and Schmitt, 1990; 8, Kelly and Loh, 1972; 9, Moewus-Kobb, 1965;10, Lee and Loh, 1973; 11, Wolf and Quimby, 1966; 12, Ahne, 1978; 13, Lee and Loh, 1975; 14, Wattanavijarn et al., 1988; 15, Kelly et al., 1980; 16, Stephens et al., 1981; 17, Fijan et al., 1983; 18, Kusuda et al., 1989.

 

The ‘standard’ cell lines for aquatic birnavirus isolation are: RTG-2 (rainbow trout gonad; Wolf and Quimby, 1962), CHSE-214 (chinook salmon embryo; Lannan et al., 1984), FHM (fathead minnow; Gravell and Marlsberger, 1965), BF-2 (bluegill fry; Wolf and Quimby, 1966) and EPC (epithelioma papulosum cyprini; Fijan et al., 1983). For diagnostic purposes, variations in the susceptibility of the teleost cell lines to infection become important and, for this reason, attempts at the primary isolation of virus are usually made on at least two different, well characterized continuous cell lines (Hill, 1976; Gillespie et al., 1977; Amos, 1985). Hill suggested that the BF-2 cell line, derived from tissues of bluegill (Lepomis macrochirus), is the most sensitive for the isolation of aquatic birnaviruses in molluscs and invertebrates (Hill, 1982). Kelly et al. (1978) compared the sensitivities of three cell lines to infection with IPNV from fish tissues and found that EPC was the most sensitive. In many instances, if the fish or shellfish to be examined is a non-salmonid, it is prudent to include a cell line from the homologous or closely related species. For example, in attempting primary isolation of the aetiological agent of spinning disease of menhaden (B. tyrranus), the birnavirus could only be isolated in cells from this fish species (Stevens, 1981).

While the in vitro host range of aquatic birnaviruses is wide, it is also erratic. In one instance, only one of four cell lines derived from carp was susceptible to a birnavirus from milkfish (Chanos chanos) (Chen et al., 1990). In vitro host range variants of IPNV also occur: some strains do not replicate in FHM or EPC cells, while others produce a high yield (Scherrer and Cohen, 1975; Kelly et al., 1978; Nicholson et al., 1979; Castric et al., 1987). Furthermore, Kelly et al., (1978) found that the RTG-2 cell line was more sensitive than the FHM cell line in detecting IPNV from tissues of carrier fish (50% vs. 12.5% of carriers detected). The use of multiple cell lines is recommended and adopted by national and international regulatory agencies for the detection of IPNV (Hill, 1976; Gillespie et al., 1977; Amos, 1985).

Other methods which have been developed to enhance the sensitivity of birnavirus detection in cell cultures are based on increasing the efficiency of the sample preparation rather than on increasing the susceptibility of cells. Two main methods of sample preparation are utilized. In both, homogenization of the sample is followed by low speed centrifugation. In one method, the supernatant was filtered through a bacteria-retaining filter, while in the others antibiotics were added to inhibit bacterial growth (Gillespie et al., 1977; Okamoto and Sano, 1984; Amos, 1985; Hedrick et al., 1986); the two methods yield comparable results. Cocultivation is another method; the separated host cells (usually kidney or spleen) are allowed to settle on a preformed monolayer of continuous cells, thereby infecting them by direct contact. This method has been reported to provide greater sensitivity than homogenization techniques (Agius et al., 1982, 1983).

The screening procedure for IPN is based on IPNV isolation tests in cell culture (Agius et al., 1982) followed by immunological identification, either by serum neutralisation (Hill and Way, 1995) or enzyme-linked immunosorbent assay (ELISA) (Dixon and Hill, 1983), of virus isolated. Diagnosis of clinical cases is normally based on histology (McKnight and Roberts, 1976) and/or immunological demonstration of IPNV antigen in infected tissues (Evensen and Rimstad, 1990), confirmed by isolation and immunological identification of IPNV in tissue culture, as for screening.

Serological and biochemical methods: detection and identification

Serological techniques have been used extensively in the identification and classification of aquatic birnaviruses. The serological identification methods have increased in sensitivity over the years, but the neutralization test remains the benchmark for other tests due to its inherent sensitivity and its use for classifying viruses. Neutralization tests utilizing polyvalent antisera are used to identify serogroup A aquatic birnaviruses (Lientz and Springer, 1973; Amos, 1985). A cautionary note, however, was raised by Vestergård-Jørgensen and Grauballe (1971), who found that the method used to immunize rabbits in the production of anti-IPNV antiserum could affect the specificity of the reaction.

The most frequently employed neutralization test uses a constant viral titre (approximately 100 TCID50) and varies the antibody concentration. The test is confirmatory for the identification of the virus and requires up to 1 week to complete. Other serological techniques used include complement fixation (Finlay and Hill, 1975), fluorescent antibody (Nicholson and Dunn, 1974; Tu et al., 1974; Vestergård-Jørgensen, 1974), immunoperoxidase tests (Reno, 1976; Nicholson and Henchal, 1978), neutralization kinetics (Nicholson and Pochebit, 1981), Staphylococcus coagglutination (Kimura et al., 1984; Bragg and Combrick, 1987b), counterimmunoelectrophoresis (Dea and Elazhary, 1983), enzyme-linked immunosorbent assay (ELISA) (Nicholson and Caswell, 1982; Dixon and Hill, 1983; Hattori et al., 1984), immunodot (McAllister and Schill, 1986; Ramsey et al., 1986; Caswell-Reno et al., 1989; Babin et al., 1991; Ross et al., 1991), and immunoprecipitation (Lipipun, 1988) tests. The tests vary in sensitivity, as well as in specificity and in the complexity and efficiency of the performance of the test.

Table Serological tests used for the detection and identification of aquatic birnaviruses, their sensitivities and time required for assay.


TestDetection limits (in vitro)Detects IPNV in diseased fish?Detects IPNV in carrier fish?Time required for testRef
Neutralizationless than or equal to 102 TCID50 ml-1YesYes4-5 days1
Complement fixation106 TCID50 ml-1YesNo4 h2
ImmunodiffusionNot reportedYesNo24 h3
Counterimmunoelectrophoresis105 TCID50 ml-1YesNo2 h4
Fluorescent antibody test105 TCID50 ml-1YesYes2 h5
ImmunoperoxidaseNot reportedYesYes2 h6, 7, 8
ELISA104-6 pfu ml-1 (10 ng)YesNo7 h9
Immunodot105 pfu ml-1 (10 ng)YesNo8 h10, 11
Western blotNot reportedNot reportedNot reported8 h12
Staphylococcus coagglutination105.9  TCID50 ml-1 (103.1 TCID50 g-1)YesNot reported1 h13

1, Wolf and Quimby, 1969; 2, Finlay and Hill, 1975; 3, Piper et al., 1973; 4, Elazhary et al., 1982; 5, Vestergård-Jørgensen, 1974; 6, Nicholson and Henchal, 1978; 7, Reno, 1976; 8, Hedrick and Fryer, 1981; 9, Nicholson and Caswell, 1982; 10, Caswell-Reno et al., 1989; 11, Ross et al., 1991; 12, Lipipun, 1988; 13, Kimura et al., 1984.

 

The most commonly used tests, ELISA and immunodot, have sensitivity levels that are sufficient to detect viral antigen in the 10-100 ng range (Nicholson and Caswell, 1982; Dixon and Hill, 1983; Caswell-Reno et al., 1986), which corresponds to approximately 106 infectious doses of virus ml-1 of cell culture fluid or organ homogenate (Hattori et al., 1984; Babin et al., 1991). However, viral titres in carrier animals are frequently more than 10,000-fold lower than this (Wolf et al., 1968b; Yamamoto, 1975a; Reno, 1976). The sensitivity of these tests is, therefore, inadequate for confidently detecting virus in carrier animals; hence cell cultures are still used for the primary isolation of the virus. However, it has been demonstrated that viral antigen is widespread in the tissues of carrier fish, even in the absence of high viral titres (Reno, 1976, 1988; Hedrick and Fryer, 1982).

Serological techniques are frequently used to confirm the identity of virus following isolation in cell culture.Rodriguez Saint-Jean et al. (1991) reported the use of flow cytometry to detect IPNV in the blood leucocytes in an IPN epizootic. It was determined that at least 70% of the leucocytes must be infected before a positive could be confirmed, although only small numbers of cells (approximately 10,000) are needed for the test. The test is rapid and can be performed more rapidly than enzyme linked assays. A cocultivation method was used to increase the percentage of leucocytes that were infected, but this increased the time required for the assay.

Vestergård-Jørgenson and Grauballe (1971) reported that the adjuvant used for the immunization of rabbits with IPNV affected the specificity of the response to different strains of the virus: Freund’s complete adjuvant promoted the generation of more specific antibodies than did Freund’s incomplete adjuvant. As with other viral systems, the assay that is used to determine cross-reactivity of antibodies can play a role in the results obtained. For example, Evensen and Rimstad (1990) determined that rabbit antisera against strains Sp, Ab and VR-299 cross-reacted in neutralization and Western blotting assays; however, in paraffin-embedded tissues from fish infected with strain Sp, only the homologous antiserum reacted. Small differences in reaction patterns were noted with neutralization reactions versus solid-phase immunoassays, even when MAbs were used (Caswell-Reno et al., 1986).

An in situ hybridization (ISH) technique has been developed to detect aquatic birnavirus in infected BF-2, EPC, and CHSE-214 cells (Alonso et al., 2004).

Many prototcols for the use of the reverse transcriptase-polymerase chain reaction (RT-PCR) assay have been developed for the detection of aquatic birnaviruses, both in cell culture (Rimstad et al., 1990; Cepica et al., 1991; McAllister et al., 1992; Shankar and Yamamoto, 1994; Wang et al., 1997) and in fish (Taksdal et al., 2001; Candan, 2002). The sensitivity and specificity of the RT-PCR method has improved accordingly with the general technical development and present RT-PCR protocols detect a higher number of IPNV-positive samples than standardized cell culture methods. The RT-PCR methods have also been developed to be able to detect several fish viruses simultaneously (multiplex-RT-PCR) (Williams et al., 1999).

In a comparative evaluation of five serological methods and RT-PCR assay for the detection of IPNV in fish it was concluded that RT-PCR and flow cytometry were the most appropriate and sensitive methods for the routine detection of IPNV from affected fish (Rodriguez Saint-Jean et al., 2001).

There is a continuous improvement of laboratory diagnosis and detection of IPN/IPNV along with the development of new and more sophisticated methods. Detection of IPNV with special properties (i.e. virulent types/strains) requires that the molecular basis for these properties are well and unequivocally described.

An evaluation of the ability of a test to detect IPNV includes evaluation of both the specificity and sensitivity. The gold standard for IPNV detection is virus isolation in cell culture, which is thoroughly described in the OIE manual. It is advisable to select a method with regard to the information required. Is it solely to detect the presence of IPNV or are quantitative measurements wanted? Is it required to relate eventual virus presence to pathological changes? Are qualitative properties of the detected IPNV, i.e. possible virulence markers, present?

List of Symptoms/Signs

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SignLife StagesType
Finfish / Bursts of abnormal activity - Behavioural Signs Aquatic:Fry Sign
Finfish / Cessation of feeding - Behavioural Signs Aquatic:Fry Sign
Finfish / Change in shape (e.g. distension) - Eyes Aquatic:Fry Sign
Finfish / Corkscrewing - Behavioural Signs Aquatic:Fry Sign
Finfish / Darkened coloration - Skin and Fins Aquatic:Fry Sign
Finfish / Mortalities -Miscellaneous Aquatic:Fry Sign
Finfish / Pancreas - white-grey patches (haemorrhage / necrosis / tissue damage) - Organs Aquatic:Fry Sign
Finfish / Red spots: pin-point size (petechiae) - Skin and Fins Aquatic:Fry Sign

Disease Course

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Little has been done with natural IPN disease to determine the portal of entry, dissemination of virus to target organs or effects on the target cells, except histopathologically.

In experimental infections, the disease is manifest by 1 week postinfection and generally has run its course within another week or so (Wood et al., 1955; Reno, 1976; Reno et al., 1978). The acute nature of the disease is characteristic of all aquatic birnavirus-induced diseases, including Japanese flounder and yellowtail ascites, turbot disease, nephritis of eels and spinning disease of menhaden (Sano et al., 1981; Bonami et al., 1983; Stephens et al., 1983; Sorimachi and Hara, 1985).

Swanson and Gillespie (1981, 1982) experimentally infected 6-month-old or yearling brook trout with IPN (strain VR-299) by intraperitoneal injection and followed the distribution of virus in the gastrointestinal tract, kidney and blood. While no clinical disease was produced , high viral titres were found in the gut and kidney, and focal necrotic lesions were detected in the pyloric caeca within 3 days postinfection and in the kidney within 5 days postinfection. Indirect fluorescent antibody tests indicated the presence of foci of IPNV antigen in pyloric caeca, kidney and liver and the virus was detected in both the serum and the mononuclear cell fractions of the blood. Hornich et al. (1986) indicated that fry undergoing acute IPN had the most intense fluorescence in the gut epithelial cells. In contrast, in natural carrier brook trout no virus was detected in the plasma, although virus was detected in the leucocytic fraction (Yu et al., 1982). In addition, McAllister et al. (1987) found that the cellular phase of the ovarian fluid of carriers contained up to 400 times more virus than the fluid portion. The distribution of virus within the viscera is not uniform, with distinct tissue tropisms.

Epidemiology

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Transmission of the disease and epidemiology

In the classic Culture and Diseases of Game Fishes, Davis (1953) makes the following reference to M’Gonigle’s information (1940) on alleviating acute catarrhal enteritis, now assumed to be IPN: "M’Gonigle recommends that affected fish be planted in small streams where they can get natural food such as insect larvae. Since the disease is not due to infection by animal parasites or bacteria, liberation of the fish in natural waters can do no harm." It is unknown how many fish farmers adhered to this practice, which was promulgated before the discovery of virus infections in teleosts. It is possible that the panzootic nature of aquatic birnaviruses may be due in part to this type of remedy for a disease of unknown aetiology.

The epizootiology of IPN is complex and is not thoroughly understood. The disease affects young fish and postsmolts, the virus may be retained in the body after primary infection without causing clinical signs in older fish, and the virus may be transmitted vertically.

IPNV may be introduced to a hatchery by eggs, fry, human, fomites, different animals, and water. The impacts of these possible introductory alternatives may vary.

The prevalence of IPNV and IPN disease is not easily appraised, although the virus appears to be present at rather high levels in some instances.


Table. Reported IPNV carrier rates in various aquatic animals.


YearSpeciesDisease?Pooled samples?Sample testedPrevalence (%)Ref
1976BKTnonoS, K, P, L, I, G. F98.703
1975BKTyes?noS. K. P. G. F50–901
1963BKTyesyesO, F502
1968BKTyes?noF174
1971BKTyes?noS, K. P. L. F15–405
1982BKTnoyesS, K, P. L, I. G606
1967BKTyes?noFE, PW92.77
1968RBTnonoF. PW0.3–54
1975RBTno?noS, K, P, G, F231
1976RBTnoyesS, K, P288
1975RBTyesnoS, K209
1980RBTyesnoB, K, S1510
1981RBTnonoB, K, S011
1979RBTno  1712
1976BNTnonoS, K, P0.68
1976ATSnonoS, K, P0.68

ATS, Atlantic salmon; BKT, brook trout; BNT, brown trout; RBT, rainbow trout. S, spleen; K, kidney; P, pyloric caeca; G, gonads; I, intestine; L, liver; F, faeces; PW, peritoneal wash; OF, ovarian fluid; B, blood.

1, Yamamoto, 1975b; 2, Wolf et al., 1961; 3, Reno, 1976; 4, Wolf et al., 1968b; 5, Frantsi and Savan, 1971b; 6, Hedrick and Fryer, 1982; 7, Billi and Wolf, 1969; 8, Munro et al., 1976; 9, Dorson, unpublished; 10, Dorson 1980, 1982; 11, Dorson, 1981, 1982; 12, Bucke et al., 1979.

 

While there is considerable information in the literature on prevalence levels, substantial variations exist within even small geographical locales and, temporally, even at the same facility. This diversity precludes easy analysis or compilation of information. It is generally accepted that, if the virus is present in a facility, the infection will remain for long periods in the absence of active measures to eradicate the agent. IPNV seems to be persistent once introduced into a hatchery.

For example, in one hatchery, the disease was present in the 1940s and mortalities were in excess of 90% in brook trout; by the late 1970s, the disease was still present, but mortality had dropped to around 40% (L. McCullogh, personal communication). Conversely, a brook trout hatchery known to have IPNV present at enzootic levels for many years has had no virus-induced mortalities for more than 25 years (P.W. Reno, personal observations). It is interesting to note that this hatchery has distributed IPNV-infected brook trout to many other aquaculture facilities, but the virus has not necessarily remained with all of the fish that were moved. For instance, one fish farm received fish from the facility in 1975 and inspection of the stocks indicated the presence of the virus. The only measures taken were to house the fish at the tail-end of the impoundments. When these fish were tested annually, starting in 1987, no virus was detected in them. This is one of only a few instances in which IPNV has been eliminated from a population of carrier fish. Bucke et al. (1979) report that in 1975 IPNV was detected in grayling at the outfall of an IPNV-contaminated hatchery, but there was no evidence of virus within 2 years in the same population of fish.

There is accumulating evidence that the mode of transmission within a hatchery may be a combination of vertical and horizontal transmission. It is most likely that the initial source of infection is from previously infected stocks or from homeothermic vertebrates or invertebrates. Horizontal transmission in hatcheries is most probably due to contact with the virus in faeces and urine of infected fish (Billi and Wolf, 1969; Frantsi and Savan, 1971b).

Little is known of the epizootiology of the other diseases caused by aquatic birnaviruses (branchionephritis of eels, whirling disease of menhaden and yamabe ascites of flat-fish). Much more work in this area is needed.

The aquatic birnaviruses are not restricted to hatchery environments or only associated with fish released from contaminated facilities. The virus is in the viscera of wild stocks in remote regions that have had no known plantings or contacts with hatchery-reared fish. Souter et al. (1986) isolated IPNV from nearly half of 229 anadromous Arctic char, Salvelinus alpinus, prespawners collected from the Mackenzie river delta and rivers in the Yukon Territory, where there was no known contact with hatchery-reared fish. The high level enzootic nature of the virus in this location indicates that natural infections may be widespread and that the maintenance of the carrier state in fish is efficient in the absence of the high viral densities found in rearing facilities. In another remote region of Canada (Hudson Bay), IPNV was enzootic in brook trout during the late 1970s and early 1980s (A.J. Sippel, personal communication); likewise, the virus was detected in wild Atlantic salmon in Loch Awe in Scotland (Munro et al., 1976). In addition, aquatic birnaviruses have been isolated from wild brown trout at the outfalls of IPNV-contaminated hatcheries in England and Wales

(Bucke et al., 1979). In Japan, yellowtail ascites virus was also detected in 15% of wild yellowtail fingerlings some distance from mariculture sites; nearly half of infected fish reared in the laboratory developed clinical disease (Isshiki et al., 1989). However, it cannot be assured that these populations were isolated from potential virus carriers in the sea or from fish upstream of the mouth of the bay. Routine culture of yellowtail in Japan employs feral fingerlings as mariculture stocks and these fish are probably the source of virus which causes the epizootics in cultured yamabe. Thus, it appears that the aquatic birnaviruses are indigenous to some fish populations, even wild stocks.

It appears that IPN disease is extremely contagious in hatchery facilities. Fomites and humans are apparently capable of transmitting the virus within and between fish-rearing facilities. The virus is transmitted both vertically and horizontally. In most instances, it appears that if no recourse is taken to eliminate IPNV from a trout culture facility, the virus will remain at the facility for years. Many of the facilities from which IPNV was originally isolated still have virus present (P.W. Reno, unpublished observations). The perennial pattern of infection and disease in facilities that harbour IPNV led to early suspicion that the virus was transmitted vertically. Wolf et al. (1968a) provided evidence that this was the case with brook trout. This was later confirmed (Sano, 1971; Yamamoto, 1975a; Reno, 1976; Hedrick and Fryer, 1982; Luqi and Zhizhuang, 1988). The exact location of the virus during embryonation is unknown, but it is extremely difficult to isolate virus from the egg. Fijan and Georgetti (1978) detected the virus at low levels in the chorion of the fertilized ovum. However, the virus is readily demonstrable after hatching (Reno, 1976; Hedrick and Fryer, 1982).

Other animals may also be involved in transmission of the virus within and between hatcheries. Sonstegård et al. (1972) infected several mammalian and avian species being able to vector IPNV and found infectious virus in the faeces of mink (Mustela sp.), great horned owl (Bubo virginianus) and chickens (Gallus domesticus) after 1 week. The virus was not detected in the faeces of great blue herons, common eider, or common merganser. Similarly, Eskildsen and Vestergård-Jørgensen (1973) isolated IPNV from gulls (Larus ridibundus) administered virus per os. Peters and Neukirch (1986) demonstrated that herons (Ardea cinerea) which were fed virus either by direct intubation or by ingestion of IPNV-injected trout harboured the virus at fairly high levels for about 30 days after the last feeding. While these findings lend credence to the possibility of transferring the virus by other animals, the consensus is that transfer in hatcheries most often occurs either horizontally within the same facility or watershed or through dissemination of contaminated eggs. It was also demonstrated that freshwater crayfish (Astacus astacus) were susceptible to IPNV, retained the virus in their tissues and haemolymph and shed virus constantly into the water (Halder and Ahne, 1988). Transmission from crayfish to trout eggs occurred, as did the infection of the crayfish from ingestion of IPNV-infected trout. There appeared to be no diminution of virus titre in the crayfish haemolymph or tissues for at least 6 months, but there was no indication of viral replication in the crayfish. These findings indicate that crustaceans and other aquatic invertebrates may be involved in the transmission of aquatic birnaviruses. Trout-rearing facilities whose water-supply is from open sources or sources that are not treated with ultraviolet irradiation or ozonation could have aquatic birnaviruses disseminated by both fish and crustacean vehicles, such as crayfish or copepods.

Mixing populations from several hatcheries, mode of transportation, and size of smolts at sea transfer are associated with increased risk of IPN outbreaks in seawater (Jarp, 1999). Intensified rearing conditions in the freshwater phase (superoxygenation, low water supply and high density) are shown to increase risk of IPN outbreaks in seawater (Jarp, 1999).

The movement of fish and especially eggs, in the international market must have contributed to the dissemination of the virus worldwide, although no thorough retrospective analysis has been done. Widespread movement of infected fish may have spread the virus to non-salmonid animals. However, it must be pointed out that non salmonids, especially invertebrates, show little or no evidence of disease when infected with aquatic birnaviruses and it is possible that the virus can be transmitted from these animals to salmonids, which under artificial rearing conditions were susceptible to disease. In this context, it has been reported by Hill (1982) that at least eight of the aquatic birnaviruses isolated from marine shellfish species were capable of producing characteristic signs of clinical IPN in experimentally infected rainbow trout.

Retrospective epizootiological studies of the diseases and infections caused by aquatic birnaviruses are difficult, due to a lack of consistency and accuracy in record keeping. Molecular markers for different virulence variants of IPNV within different serotypes can be revealed by RT-PCR and nucleotide sequencing (Bruslind et al., 2000; Santi et al., 2004; Shivappa et al., 2004). This implies that more precise characterization of IPNV isolates and the distribution of different variants in the cultivated and wild fish populations are needed. Such information is important for further understanding of IPN, risk factors for spread of the infection and for the implementation of proper control measurements.

Viruses belonging to the He and Tellina serotypes (types A4 and A5 of Hill and Way) have been detected exclusively in Europe, and Canada 1, 2, and 3, and Jasper (types A6-A9) have been confined to North America (Hill and Way, 1988b). This is interesting because it is known that trout infected with the WB serotype (A1) were often transferred to Europe from North America in the 1960s, but there is a single report of a WB serotype virus isolated in Europe (Novoa et al., 1993a). There are, however, some instances where the presence of disease in areas previously free from virus could be traced directly to the importation of infected eggs. Such importations occurred in South Africa (Bragg and Combrick, 1987a) and in Japan (Sano, 1971). In both instances, virus accompanied eggs imported from the USA. In other instances, there was circumstantial temporal evidence that the disease arrived with eggs imported into France from the USA (Besse and de Kinkelin, 1965), but subsequent serological evidence indicated that it was probably an indigenous strain (Wolf and Quimby, 1971). A heightened awareness of the disease in France by fish health workers, rather than the importation of virus in fish, was probably responsible for the detection of these strains. In other instances, there have also been indications that aquatic birnaviruses were transmitted to indigenous and cultured species from imported fish or eggs: to Chile from the USA (McAllister and Reyes, 1984; Espinoza et al., 1985), to Japan from Europe with European eels (Sano et al., 1981) and to Taiwan from trout imported from Japan (Hedrick et al., 1983). It is very difficult, however, to accurately differentiate between ‘indiginous’ aquatic birnaviruses and those that were brought into a country or region. The lack of efficient record-keeping at facilities, the lack of universally applied inspection regimes for viruses and the limited usefulness of the serological classification make epidemiological studies less than optimal.

Except for ‘classic’ IPN disease in freshwater trout, all of the diseases associated with aquatic birnaviruses affect either stenohaline or euryhaline animals: yellowtail and Japanese flounder ascites, sea bass, menhaden, eels and possibly clams.

The virus is quite stable, even under conditions that inactivate most other fish pathogens. The infectivity of the virus survives for nearly a year at 4°C and nearly 2 months at 15°C in buffer (Dorson, 1982) and also for long periods in seawater, brackish water and unsterilized fresh water (Moewus-Kobb, 1965; Toranzo and Hetrick, 1982). This stability increases transmission of the virus from carrier or clinically ill fish in rearing facilities to susceptible fish upstream or downstream in the watershed, or even to susceptible hosts in seawater. The ‘spread’ of the virus among freshwater, brackish and seawater environments is facilitated by the stability of the virus, the migration of anadromous and catadromous fishes and the predator-prey relationships. The latter is supported by the finding of a birnavirus-like agent in mass cultures of rotifers, Branchionus plicatilis (Comps et al., 1991), which could potentially be transmitted to larval sea bass (D. labrax), since rotifers are the primary food source for this species in culture. It was suggested that rotifers could be responsible for the epizootics of the disease in maricultured sea bass in France (Bonami et al., 1983).

Vertical transmission and the carrier state

Vertical transmission is usually defined as the transmission of virus from one generation to the next. The virus can either be within the contents of the gametes, often referred to as true vertical transmission, or alternatively on the surface of gametes or in ovarian and seminal fluids and mucus.

Vertical transmission of the virus via the reproductive products of salmonids has been substantiated (Wolf et al., 1968a; Sano, 1971; Yamamoto, 1975a; Reno, 1976; Hedrick and Fryer, 1982; Luqi and Zhizhuang, 1988); however, the actual method by which this occurs remains obscure. The perennial nature of the disease in salmonid hatcheries facilities indicates that this is an important transmission mechanism. This makes sense with trout which are reared in freshwater facilities, often with a well-water source, which would preclude infection via a water-borne route, although the possibility of contamination by birds or transfer by aquatic mammals cannot be ruled out. However, this cannot explain the recurring disease problems in salmonid seawater culture, which indicate that transmission from an environmental source is important. This can be compared to infectious haematopoietic necrosis, which was originally postulated to be vertically transmitted but now appears to be horizontally transmitted (Leong and Munn, 1991).

Wolf et al. (1968a) demonstrated that the virus was transmitted via reproductive products. Eggs and milt were collected from IPNV-positive fish and three pairings were made. The virus was isolated from two matings and frank disease was noted in one batch of the offspring. Bullock et al. (1976) found that the virus was transmitted to an uninfected facility via fertilized brook trout eggs transferred from a contaminated hatchery. However, the presence of the virus in the parent’s reproductive products or progeny does not invariably lead to the development of clinical disease. Reno (1976) found low levels (101.4 TCID50 g-1) of IPNV in all eight groups of progeny from matings of naturally infected brook trout which harboured fewer than 102.4 TCID50 ml-1 of IPNV in milt or ovarian fluid. Virus was not detectable during the embryonation period or until feeding commenced, but IPNV was detected in postfeeding fish for only 1 month. Similarly, in other species, vertical infection has been transient and sporadic. In Atlantic salmon, Smail and Munro (1989) detected virus for less than 1 h in eggs from brood fish exposed to IPNV at titres of 106-109 plaque-forming units (pfu) kg-1 (injection) or 107 pfu kg-1 (immersion), and the resultant fry were negative for virus. However, when fry were infected experimentally, the virus was isolated even after 2 years.

Dorson and Torchy (1985) experimentally infected rainbow trout 4.5 months prior to spawning. In addition, at spawning, they infected milt and ova with high doses of IPNV Sp. Three of six infected females had detectable IPNV in ovarian fluid at spawning, whereas none of the males had virus in the milt. There was no evidence of IPN disease in the progeny. When milt was infected, however, IPN disease developed in the fry. Similarly, Ahne and Negele (1985) demonstrated that experimental infection of rainbow trout and Arctic char resulted in consistent isolation of virus from the chorion of the eggs during embryonation and posthatch. In fry, virus was isolated after only 1 month posthatch, but there was no evidence of disease in these fish. This indicates that eggshells may be involved in transmission of virus from one generation to the next and important for the horizontal transmission of IPNV to first feeding fry. The variations in results between the different experiments may be due to the species differences; brook trout are regarded as more susceptible to disease than other salmonids. However, different virulence variants of viruses, especially in experimental infections using cell culture-derived virus may explain differences in disease development. Only a few passages of virus in cell culture may cause attenuation of the virus (Santi et al., 2004).

Survivors of IPN epizootics remain carriers for up to 6 years (Wolf, 1966; Yamamoto,1975a; Reno, 1976). In a given population, the proportion of fish from which virus can be isolated varies drastically. In naturally infected brook trout, the prevalence ranged from 99% to less than 5% (Reno, 1976; unpublished observations). The latter is the prevalence used in fish health inspections for the detection of viruses in salmonids (Amos, 1985). The potential for the dissemination of the virus through the faeces has been demonstrated (Billi and Wolf, 1969; Frantsi and Savan, 1971b; Reno, 1976) and this ‘shedding’ of virus, albeit at low levels, may be responsible for the maintenance of the virus in susceptible fish. Carrier states have also been reported in survivors of epizootics associated with other aquatic birnaviruses: striped bass, turbot, yellowtail, eels and menhaden (Sano et al., 1981; Stephens et al., 1983; Sorimachi and Hara, 1985; Wechsler et al., 1987a).

There are no reported instances of zoonotic infections contracted from any aquatic birnaviruses.

Impact Summary

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CategoryImpact
Biodiversity (generally) Negative
Fisheries / aquaculture Negative
Native flora Negative

Impact: Economic

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Traditionally, IPNV was a disease of fry and fingerlings, with brook and rainbow trout being considered to be more susceptible than brown trout, lake trout (Salvelinus namaycush), coho and Atlantic salmon (Salmo salar). However, mortality varies considerably between outbreaks and has been related genetic susceptibility of the host (Ozaki et al., 2001 and Silim et al., 1982) and differing level of environmental stress (Frantsi and Savan, 1971 and McAllister and Owens, 1986). IPNV also infects non-salmonid species including eels, molluscs and crustaceans, all of which may act as carriers. The economic impact of IPN mortality of marine-reared Atlantic salmon has been of increasing concern, partly because of the size of the fish (Jarp et al., 1995).

Infection of susceptible species in fresh water can result in high mortality. Virus-associated mortality is rapid between 10 and 14°C, and at lower temperatures is prolonged. Water temperature, fish age and the virus strain affect the severity of the disease, as well as the establishment of covert infections in fish. IPNV replicates in kidney, pancreas, gonad, spleen and intestinal epithelium and may be shed from carrier fish via faeces, as well as through the seminal and ovarian fluids. Virus shedding appears to be dependent on stress. Although stress may induce a recurrence of IPN in fish 6-11 months old, older fish show no clinical signs.

The apparent increased incidence of IPN is considered to be the result of widespread fish and egg movements between countries, and because of increased sensitivity of diagnostic methods, resulting in improved surveillance practices.

IPNV is transmitted horizontally and vertically (Reno, 1999). However, vertical transmission has been confirmed only in brook trout and rainbow trout (Ahne et al., 1989; Ahne and Negele, 1985; Dorson and Torchy, 1981). The exact mechanism of egg entry or location of the virus within the egg is still unclear. Bebak et al. (1988) infected rainbow trout with IPNV by immersion challenge. The fish started to excrete the virus within 2 days of infection and shedding increased and then declined in less than 12 days post-exposure. From this study it was estimated that within 14 days, more than 75% of the population can be infected (Bebak et al., 1988). This gives rise to a rapid spread of IPNV.

Ethanol, methanol, iodophore and chlorine inactivate IPAV (Inouye et al., 1990), but it retains more than 90% of its infectivity after treatment with chloroform or ethyl ether, at pH 3.0 for 60 min. IPNV has been shown to be infective for several years at -70°C and for several months at 4°C. Smail et al., (1993) found that IPNV was not deactivated by an acidic pH unless the sample (silage) was heated for at least 2 h at 60°C. This confirms that IPNV is a robust virus with long survival in the environment.

Zoonoses and Food Safety

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This disease is not a zoonosis.

References

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Agius C; Mangunwiryo H; Johnson RH; Smail DA, 1982. A more sensitive technique for isolating infectious pancreatic necrosis virus from asymptomatic carrier rainbow trout, Salmo gairdneri Richardson. Journal of Fish Diseases, 5(4):285-292.

Agius C; Richardson A; Walker W, 1983. Further observations on the co-cultivation method for isolating infectious pancreatic necrosis virus from asymptomatic carrier rainbow trout, Salmo gairdneri Richardson. Journal of Fish Diseases, 6(5):477-480.

Ahne W, 1978. Isolation and characterization of infectious pancreatic necrosis virus from pike (Esox lucius). Brief report. Archives of Virology, 58(1):65-69.

Ahne W; Jorgensen PEV; Olesen NJ; Fischer-Scherl T; Hoffmann R, 1989. Aquatic birnaviruses: virus of the serogroup II isolated from an IPN outbreak in brook trout (Salvelinus fontinalis). Bulletin of the European Association of Fish Pathologists, 9(1):14-16.

Ahne W; Negele RD, 1985. Studies on the transmission of infectious pancreatic necrosis virus via eyed eggs and sexual products of salmonid fish. In: Ellis A, ed. Fish and Shellfish Pathology. London, UK: Academic Press, 262-270.

Alonso MC; Cano I; Castro D; Perez-Prieto SI; Borrego JJ, 2004. Development of an in situ hybridisation procedure for the detection of sole aquabirnavirus in infected fish cell cultures. Journal of Virological Methods, 116(2):133-138.

Amend DF; Pietsch JP, 1972. Virucidal activity of two iodophors to salmonid viruses. Journal of the Fisheries Research Board of Canada, 29(No.1):61-65.

Amos K, 1985. Procedures for the Detection and Identification of Certain Fish Pathogens. Corvallis, Oregon, USA: Fish Health Section, American Fisheries Society, 6-13.

Anderson C, 1996. Distribution of salmonid diseases in New Zealand. Surveillance (Wellington), 23(4):23-24; 11 ref.

Archetti I; Horsfall FL, 1951. Persistent antigenic variation of influenza A viruses after incomplete neutralization in ova with heterologous immune sera. Journal of Experimental Medicine, 92:441-462.

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Babin M; Hernandez C; Sanchez C; Dominguez J, 1991. Immunodot assay for detection of IPN [infectious pancreatic necrosis] virus in organ homogenates. Bulletin of the European Association of Fish Pathologists, 11(2):65-67.

Ball HJ; Munro ALS; Ellis A; Elson KGR; Hodgkiss W; McFarlane IS, 1971. Infectious pancreatic necrosis in rainbow trout in Scotland. Nature, 234(No. 5329):417-418.

Bebak J; McAllister PE; Smith G, 1998. Infectious pancreatic necrosis virus: transmission from infectious to susceptible rainbow trout fry. Journal of Aquatic Animal Health, 10(3):287-293.

Besse P; de Kinkelin P, 1965. Sur l’existence en France de la nécrose pancréatique de la truite arc en ciel (Salmo gairdneri). Bulletin de l’Académie Vétérinaire, 38:185-190.

Billi JL; Wolf K, 1969. Quantitative comparison of peritoneal washes and faeces for detecting infectious pancreatic necrosis (IPN) virus in carrier brook trout. Journal of the Fisheries Research Board of Canada, 26:1459-1465.

Blake S; Ma JY; Caporale DA; Jairath S; Nicholson BL, 2001. Phylogenetic relationships of aquatic birnaviruses based on deduced amino acid sequences of genome segment A cDNA. Diseases of Aquatic Organisms, 45(2):89-102.

Bonami JR; Cousserans F; Weppe M; Hill BJ, 1983. Mortalities in hatchery-reared sea bass fry associated with a birnavirus. Bulletin of the European Association of Fish Pathologists, 3(3):41.

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Links to Websites

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OIE Manual of Diagnostic Tests for Aquatic Animalshttp://www.oie.intManual accessible from homepage
Universal Virus Database of the International Committee on the Taxonomy of Viruseshttp://www.ncbi.nlm.nih.gov/ICTVdb

Contributors

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Main Author
Espen Rimstad
Norwegian School of Veterinary Science, Dept. of Food Safety & Infection Biology, POB 8146, dep 0033 Oslo, Norway

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