Viral Nervous Necrosis Disease

C H A P T E R

30 Viral Nervous Necrosis Disease Mohammad Jalil Zorriehzahra Aquatic Animal Health & Diseases Department, Iranian Fisheries Science Research Institute (IFSRI), Agricultural Research Education and Extension Organization (AREEO), Tehran, Iran

ABBREVIATIONS BFNNV BSE CNS ELISA FHV IFAT IHC NoV PCR RGNNV SJNNV TPNNV VER VNN

barfin flounder nervous necrosis virus bovine spongiform encephalitis central nervous system enzyme-linked immunosorbent assay flock house virus immunofluorescence antibody technique immunohistochemistry Nodamura virus polymerase chain reaction red-spotted grouper nervous necrosis virus striped jack nervous necrosis virus tiger puffer nervous necrosis virus viral encephalopathy and retinopathy viral nervous necrosis

INTRODUCTION Viral encephalopathy and retinopathy (VER)/viral nervous necrosis (VNN) is a neuropathogenic disease; so it was named first VER (Munday et al., 1992), which names sea bass encephalopathy (Bellance and Gallet de Saint-Aurin, 1988), VNN (Yoshikoshi and Inoue, 1990) encephalomyelitis turbot (Bloch et al., 1991), and fish encephalopathy (Comps and Raymond, 1996). Also it is known that the neuropathological condition is supposed to be different for fish species. Influence of

Emerging and Reemerging Viral Pathogens DOI: https://doi.org/10.1016/B978-0-12-819400-3.00030-2

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different fish species occurs by a limited number of viral agents that belong to the family Nodaviridae. Also, VNN disease is renamed as VER. In the recent decades the causative agent of VNN (VER) disease has become a serious threat of marine finfish and mariculture, and in recent decade the disease has also been associated with farmed freshwater fish. The virus has been classified as a Betanodavirus within the family Nodaviridae, and the fact that Betanodaviruses are known to affect more than 120 different farmed and wild fish and invertebrate species highlights the risk that Betanodaviruses pose to global aquaculture production (Costa and Thompson, 2016). VNN disease could be considered a serious viral disease of larvae and juvenile fish that especially susceptible with up to 100% mortality and in some cases infecting some adult marine fish (Chi et al., 2003; Zorriehzahra et al., 2005). A number of vaccine preparations have been tested in the laboratory and in the field, for example, inactivated the virus, recombinant proteins, virus-like particles (VLPs), and DNA-based vaccines, and their efficacy, based on relative percentage survival, has ranged from medium to high levels of protection to little or no protection. Meanwhile, the first commercial vaccine against VNN was applied recently for sea bass in the Mediterranean region (Zorriehzahra et al., 2019). Therefore a combination of effective prophylactic measures, including vaccination, is needed to control and prevent VNN and should also target larvae and broodstock stages of production to help the industry deal with the problem of vertical and horizontal transmission (Costa and Thompson, 2016).

VIRUS MORPHOGENESIS AND GENOMIC CHARACTERISTICS Some researchers suggested that there is a single virus, piscine neuropathy virus (Frerichs et al., 1996), but some researchers do not believe in this concept (Munday and Nakai, 1997). The causative agent of VER or VNN was first identified as a member of the family Picornaviridae (Maltese and Bovo, 2007), but on the basis of its nucleic acid, genomic structure, protein properties, and serological relationships (Lin et al., 2007), and following virus purification from affected striped jack larval (Pseudocaranx dentex) classified as a new member of the family Nodaviridae [Office Internationales Epizooties (OIE), 2013]. Subsequently, other agents of VNN were isolated from diseased fish species. The family name is derived from the Nodamura virus (NoV), the type species of Alphanodavirus genus, which was first discovered in a Japanese village called Nodamura (Scherer and Hurlbut, 1967). These RNA

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viruses comprise two genera alphanodavirus and betanodavirus. Predominantly, the alphanodaviruses can infect insects, and the betanodaviruses infect fish and referred as nervous necrosis virus (NNV) (Choi et al., 2013; Maltese and Bovo, 2007; Thiery et al., 2003). In the other hand, current taxonomy classifies the VNNV into the genus Betanodavirus (Thiery et al., 2003). Alphanodavirus contain five species including NoV, black beetle virus, Boolarra virus, flock house virus, and Pariacoto virus. NoV is the only alphanodavirus that also was seen in vertebrates (Vega and Kaya, 2012). Earlier studies classified VNNV into four major genotypes, based on their nucleotide sequence of RNA2 [Choi et al., 2013; Ma et al., 2012; Office Internationales Epizooties (OIE), 2013]. But recent studies were proposed the fifth genotype after a natural outbreak of VER in Norway (Johansen et al., 2004; Olveira et al., 2013). These betanodavirus particles are small, 25 30 nm in diameter, spherical or icosahedral morphology, nonenveloped with a single coat protein (Kokawa et al., 2008; Mazelet et al., 2011; Nazari et al., 2011; Tanaka et al., 2004). The genome is segmented and consists of two [Office Internationales Epizooties (OIE), 2013] or three (Kuo et al., 2011) molecules of linear positive-sense, single-stranded, nonpolyadenylated RNA. RNA1 (3.1 kb) encodes protein A, the replicase, which is the viral part of the RNA-dependent RNA polymerase with 110 kDa. RNA2 (1.4 kb) encodes the capsid protein with 42 kDa [Kuo et al., 2011; Office Internationales Epizooties (OIE), 2013]. RNA3 (0.4 kb) (Furusawa et al., 2006) encodes protein B2, which is a subgenomic transcript of RNA1 and only seen in the nucleus of a virus-infected cell but not in the viron (Kuo et al., 2011) and has a suppressor function for posttranscriptional gene silencing (Iwamoto et al., 2001). Complete nucleotide sequences of RNA1 and RNA2 were reported for striped jack NNV (SJNNV) and others [Office Internationales Epizooties (OIE), 2013].

HOSTS AND GEOGRAPHICAL DISTRIBUTION The first record of the disease was first reported in barramundi (Lates calcarifer) farmed in Australia (Glazebrook et al., 1990; Munday et al., 2002), Japanese parrotfish Oplegnathus fasciatus (Yoshikoshi and Inoue, 1990) followed a year later in turbot Scophthalmus maximus (Bloch et al., 1991), European sea bass Dicentrarchus labrax (Breuil et al., 1991), red-spotted grouper Epinephelus akaara (Mori et al., 1992), and striped jack P. dentex (Mori et al., 1992). Also, some unofficial reports have documented the occurrence of VNN in some African countries (Personal Communications with Prof. G. Bovo from Italy, 2013 and Dr. Abderrafik Meddour from Algeria, 2015). All of them

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revealed that early unknown mortality occurred in fry and juveniles of some marine fish in the decade of 1980 90 in the world (Caveriviere and Toure, 1990). Also, the disease has been reported in marine water, freshwater, and brackish water fish. Fish species affected by VNN are included Acipenser gueldenstaedti (Athanassopoulou et al., 2004), Anguilla anguilla (Chi et al., 2001), Chanos chanos (Gomez et al., 2006), Parasilurus asotus (Ucko et al., 2004), Tandanus tandanus (Bovo et al., 2011), Gadus morhua (Johanson et al., 2002; Starkey et al., 2004), Melanogrammus aeglefinus (Gagne´ et al., 2004), Poecilia reticulata (Ransangan and Manin, 2010), Acanthurus triostegus (David et al., 2010), Apogon exostigma (David et al., 2010), Anarhichas minor (Thie´ry et al., 2003), P. dentex (Mori et al., 1992), Seriola dumerili (Muroga, 2001), Trachinotus falcatus (Bandı´n and Dopazo, 2011; Chi et al., 2001), L. calcarifer (Le Breton et al., 1997), Lateolabrax japonicus (Nishizawa et al., 1997), Oreochromis niloticus (Bigarre´ et al., 2009), Oxyeleotris lineolata (Ransangan et al., 2011), Platax orbicularis (David et al., 2010), Latris lineate (Bandı´n and Dopazo, 2011), Lutjanus erythropterus (Maeno et al., 2004), Branchiostegus japonicus (Nakai et al., 2009), Mugil cephalus (Gomez et al., 2004), Liza saliens and Liza auratus (Zorriehzahra et al., 2016), Mullus barbatus (Giacopello et al., 2013), O. fasciatus (Munday and Nakai, 1997), D. labrax (Le Breton et al., 1997), Rachycentron canadum (David et al., 2010), Sciaenops ocellatus (Oh et al., 2002), Umbrina cirrosa (Le Breton et al., 1997), Atractoscion nobilis (Curtis et al., 2001), Thunnus orientalis (Yanong, 2010), E. akaara (Le Breton et al., 1997), Epinephelus awoara (Munday et al., 2002), Epinephelus fuscoguttatus (Nagasawa and Cruz-Lacierda, 2004), Epinephelus malabaricus (Curtis et al., 2001), Epinephelus marginatus (Vendramin et al., 2013), Epinephelus moara (Munday and Nakai, 1997), Epinephelus tauvina (Hegde et al., 2002), Epinephelus lanceolatus (Nagasawa and CruzLacierda, 2004), Cromileptes altivelis (Maeno et al., 2004), Epinephelus aeneus (Le Breton et al., 1997), Epinephelus coioides (Maeno et al., 2004), Sparus aurata (Castric et al., 2001), O. niloticus (Bigarre´ et al., 2009), Stephanolepis cirrhifer (Gomez et al., 2004), Takifugu rubripes (Nishizawa et al., 1997), Verasper moseri (Watanabe et al., 2000), Solea solea (Dalla Valle et al., 2005), Psetta maxima (Park, 2009), Paralichthys olivaceus (Nguyen et al., 1994), Hippoglossus hippoglossus (Le Breton et al., 1997), and Sebastes oblongus (Gomez et al., 2004).

MOLECULAR EPIDEMIOLOGY AND PHYLOGENETIC ANALYSIS Based on similarities and phylogenetic analysis of the variable region of RNA2 sequences encoding the C-terminal halves of the coat proteins,

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called T4, preliminarily betanodaviruses have been classified into four major genotypes: SJNNV, barfin flounder NNV (BFNNV), tiger puffer NNV (TPNNV), and red-spotted grouper NNV (RGNNV) [Furusawa et al., 2006; Kim et al., 2012; Olveira et al., 2013; Office Internationales Epizooties (OIE), 2013; Thiery et al., 2003; Wang et al., 2015]. The fifth genotype including a turbot betanodavirus strain proposed after a natural outbreak of VER in Norway with sequencing the polymerase chain reaction (PCR) product obtained [Johansen et al., 2004; Kim et al., 2012; Olveira et al., 2013; Office Internationales Epizooties (OIE), 2013]. Table 30.1 shows the major genotypes, target host species, and growth temperature. Also, three different serotypes complete this genotype identifying by virus neutralization with polyclonal antibodies [Office Internationales Epizooties (OIE), 2013]. A numerical nomenclature contains clusters I, II, III, and IV, proposed by Thiery et al. (2003) to avoid confusion the taxonomy. This mentioned nomination is independent from the host species [Office Internationales Epizooties (OIE), 2013]. Often, each host species affected by single viral agents, but in some cases reported that one host can be infected by more than one viral agent, such as in D. labrax (Maltese and Bovo, 2007). SJNNV and TPNNV, respectively, infected striped jack and tiger puffer and have limited host ranges, but BFNNV and RGNNV have a broad host range

TABLE 30.1

Genotypic and Phenotypic Variants of Betanodavirus.

Genotype

Serotype

Target host fish

Optimum growth temperature (in vitro) ( C)

SJNNV

A

Striped jack (Pseudocaranx dentex)

20 25

TPNNV

B

Tiger puffer (Takifugu rubripes)

20

BFNNV

C

Cold-water fish: Atlantic halibut, Atlantic cod, flounder, etc.

15 20

RGNNV

C

Warm-water fish: Asian sea bass, European sea bass, groupers, etc.

25 30

TNV

Turbot (Scophthalmus maximus)

BFNNV, Barfin flounder nervous necrosis; RGNNV, red-spotted grouper nervous necrosis virus; SJNNV, striped jack nervous necrosis virus; TNV, turbot betanodavirus strain; TPNNV, tiger puffer nervous necrosis virus. Data from: 1) (Iwamoto, T., Nakai, T., Mori, K.I., Arimoto, M., Furusawa, I., 2000. Cloning of the fish cell line SSN-1 for piscine nodaviruses. Dis. Aquat. Org. 43 (2), 81 89.). 2) (Olveira, J.G., Souto, S., Dopazo, C.P., Bandin, I., 2013. Isolation of betanodavirus from farmed turbot Psetta maxima showing no signs of viral encephalopathy and retinopathy. Aquaculture 406 407, 125 130.). 3) Office Internationales Epizooties (OIE), 2013. Viral encephalopathy and retinopathy. In: Diagnostic Manual for Aquatic Animal Diseases. Paris, France, p. 136.

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and, respectively, cause disease among some cold-water species and a variety of warm-water fish species, especially, in D. labrax. A phylogenetic study by Dalla Valle et al. (2001) collected from the Mediterranean area showed the probability of the existence of an ancestral strain hosted in D. labrax. Analysis showed TPNNV and SJNNV have derived from BFNNV and RGNNV around 100 150 years ago (Maltese and Bovo, 2007). Also in the past 10 years, each major genotype suffered a minor divergence, parallel with the growth of aquaculture activities (Maltese and Bovo, 2007). All the isolates from four different fish species in Korea fell within one subgroup, RGNNV. The phylogenetic analysis showed no relations between the position of strain variants host-dependent evolution and geography (Cha et al., 2007). Also, Japanese viral particles isolated originally from Japanese flounder (P. olivaceus) and belonging to the genotype RGNNV are hypothesized as one ancestral strain (Maltese and Bovo, 2007). Nishizawa et al. (1997) suggested that the Japanese Flounder must have played a key role in the spread of VNN in Japan that supported by obtaining isolates from the same species belongs to the TPNNV. Fish transportation between Europe and the Pacific affected on the geographic distribution of nodavirus (Skliris et al., 2001). Immunofluorescence antibody test (IFAT) test showed antigenic similarities between striped jack nodaviruses and other fish nodaviruses but not identical antigenic relationships (Munday and Nakai, 1997; Skliris et al., 2001). Some findings showed that the temperature effects significantly, on the molecular evolution of betanodaviruses (Totland et al., 1999; Thiery et al., 1999). For example, one isolate caused the disease in sea bass farmed on the Atlantic coast, and the second one isolated from the same species but farmed in the Mediterranean, with higher temperatures (Thiery et al., 1999). It has been hypothesized that BFNNV and RGNNV particles have a European origin, whereas a Pacific origin has been suggested for isolates belonging to the genotypes TPNNV and SJNNV (Maltese and Bovo, 2007). Probably, SJNNV isolates after reached Europe (through the exported ornamental fish) and adapted to both the local warm- and cold-water species, returned to the Pacific (through the whitefish and salmonid) (Maltese and Bovo, 2007; Skliris et al., 2001). Nevertheless, BFNNV was reported from North America, Norway, and Japan as well as sea bass (D. labrax) in France, TPNNV was isolated from T. rubripes in Kagawa, Japan. Similarly, SJNNV was observed originally in Japanese waters but also detected in Spain and Portugal, and finally, RGNNV spread in the large range of warm-water species and has observed throughout Asia, the United States, Australia, and the Mediterranean Basin (Munday et al., 2002; Panzarin et al., 2012). The first description of RGNNV in Iran with similarity .95% was reported in the Mullet fish of Caspian Sea (Zorriehzahra et al., 2005). EMERGING AND REEMERGING VIRAL PATHOGENS

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CLINICAL SIGNS VNN is a disastrous fish disease and one of the main reasons for the great economic loss in the aquaculture industry. Unfortunately, this disease has been reported from almost all regions of the globe. The clinical signs of VNN are linked with the neuroinvasive nature of Nodaviradae family of viruses, causing this disease, as well as the consequences of the disease in the form of lesion present in the retina and brain of the infected fish, ultimately leading to abnormal movement, coloration, sight, and swim bladder control (Munday et al., 2002). In general, the clinical signs of VNN are observed in the specific behavior of affected fish individuals. These behavioral changes include loss of appetite, erratic swimming patterns such as whirling, spiral, looping swimming, and belly up, loss of equilibrium, minimized nervous coordination, uncoordinated swimming, and alterations in pigmentations (Nopadon et al., 2009) (Figs. 30.1 and 30.2).

FIGURE 30.1 Clinical signs of VNN: inflation of swim bladder (left) and abdominal extension (right) in infected Liza aurata in Caspian Sea (Zorriehzahra et al., 2013). VNN, Viral nervous necrosis.

FIGURE 30.2 Spiral swimming in affected larva in VNN (Zorriehzahra et al., 2013). VNN, Viral nervous necrosis.

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These signs are accompanied by some general signs, such as anemia, lethargy, and anorexia (Munday and Nakai, 1997). The infected individuals also adopt a peculiar stationary position, take vertical position keeping caudal fin, and head above the water surface. Some VNNinfected fish swim straight so swiftly that they became unable to discontinue their speed and get smacked to tanks’ walls, and they experience traumatic and harrowing lesions on their jaws (Maltese and Bovo, 2007). Other changes that coexist are hyperinflation of the swim bladder (Mori et al., 1992). The presence or absence of any of these signs or deviation in pigmentation change may be due to change in temperature (Fukuda et al., 1996).

ANATOMO-PATHOLOGICAL LESIONS Changes in anatomy of different body organs and tissues have been reported in different species on account of VNN such as swim bladder’s hyperinflation in O. fasciatus, P. dentex, C. altivelis, L. calcarifer, L. aurata, and D. labrax (Breuil et al., 1991; Munday and Nakai, 1997; Munday et al., 2002; Zorriehzahra et al., 2005; Yuwanita et al., 2013). These changes are more prominent at the larval stage. There is a very little literature available describing any significant work on anatomopathological lesions yet some changes reported include reddening of head areas, jaws’ lesions, and depigmentation of the area in cranial skin swinging in the brain, and eclectic exposed opercula (Bovo et al., 1996; Sweetman et al., 1996; Le Breton et al., 1997; Pavoletti et al., 1998) (Fig. 30.3).

FIGURE 30.3 Exophthalmia and swollen abdomen in affected Poecilia retriculata (left) and hemorrhage of head and body surface (right) (Zorriehzahra et al., 2013).

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HISTOPATHOLOGICAL LESIONS There is a huge literature available demonstrating histopathological lesions due to VNN. In most typical cases, these lesions are detected in different parts of the brain including medulla oblongata, telencephalon, metencephalon, mesencephalon, retina’s granular layers, spinal cord, germinal epithelium, and rod and cones cells (Munday et al., 1992; Grotmol et al., 1995; Comps and Raymond, 1996; Grove et al., 2003). These changes are also accompanied by typical findings of consistent necrosis and vacuolation of CNS (central nervous system) for different species (Hegde et al., 2002; Maltese and Bovo, 2007) (Fig. 30.4). Research has shown that anterior brain is more vulnerable to VNN while compared to the posterior brain and spinal cord, and larval are more susceptible to VNN than adult stages (Munday and Nakai, 1997). Inverted microscopy of VNN-infected C. altivelis showed different histopathological alterations in intestine, kidney, liver, gill, brain, and eyes such as vacuolation, fibrosis, ruptured epithelial tissues, hemorrhage, goblet proliferation, inclusion bodies, occlusion bodies, necrosis, vacuoles, atrophy, cloudy swelling, edema, hyperplasia, and hypertrophy (Yuwanita et al., 2013). These changes, such as necrosis, hyperemia, inflammation, necrosis, edema, and vacuolation of brain and eye, have also been observed by other researchers in different marine and freshwater fish species (Nguyen et al., 1996; Peducasse et al., 1999; Zorriehzahra et al., 2005, 2010; Nazari et al., 2011).

FIGURE 30.4 Vacuolation of brain in golden grey mullet in the Caspian Sea (Zorriehzahra et al., 2013).

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Different alterations reported in O. fasciatus were including spinal ganglia escorted by intracytoplasmic brains’ gray matter, basophilia, and shrinkage of affected cells and pyknosis (Yoshikoshi and Inoue, 1990). Different other studies have shown karyorrhexis and focal pyknosis of neural cells, eosinophilic materials accumulation in blood vessels and macrophages, as well as the presence of mononuclear cells’ infiltrates (Munday et al., 1992; Grotmol et al., 1995).

ACUTE INFECTION Betanodavirus pathogenesis is varied widely (Mori et al., 2003) and related to the fish age, water temperature, and the route of infection. In young fish, VNN develops because of their immature and weak immune system. Also, their neural cells have a high level of proliferation. Thus VNN can show acute or chronic infection in same species in different conditions [Munday et al., 2002; Office Internationales Epizooties (OIE), 2013]. Clinical signs include the brain, spinal cord, and retina lesions. The infected fish shows abnormal manner, such as whirling swimming and darting movements, as they lose their sight and motor control, swelling of the swim bladder, and belly-up position as their ability to control their swim bladder is lost (Johansen et al., 2003, 2004; Starkey et al., 2001). Swelling can also be seen in the liver and spleen (Oh et al., 2002; Johansen et al., 2002). The fish growth rate is reduced and changes in pigmentation also are observed. VNN is associated with acute mortality about 100% (Hick et al., 2011; Johansen et al., 2003). Because the virus attacks to the neural tissues, juvenile and larval fish with prolific neural cells are most severely affected, often suffering mortalities of 100% at about the 10 30 days of posthatching. If larval stages infected, highest mortality, often reaching 100%, is observed, but in juveniles and older fish lower losses have been generally reported. In striped jack, mortalities in high levels observed within 10 days after hatching, and European sea bass show mortality after about 30 days of posthatching [Office Internationales Epizooties (OIE), 2013]. Mass mortality in Asian sea bass (L. calcarifer) was reported after 18 21 days of posthatching with clinical manifestations, such as anorexia, blackening, lack of swimming coordination, and settling to bottom by Azad et al. (2006). Adult fish are also affected, but the disease mostly shows subclinical signs. Johansen et al. (2004) reported high rates of mortality in farmed turbot, S. maximus, after 13 16 days of posthatching, with any clinical nervous signs. Many researchers reported outbreaks associated with

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FIGURE 30.5 Hemorrhage in eye and reddening of head in acute infection of golden grey mullet in the Caspian Sea (Zorriehzahra et al., 2013).

almost 100% death occur within a very short time (Kuo et al., 2012; Murali et al., 2002; Johansen et al., 2002, 2004; Ma et al., 2012). Disease development has a direct correlation with high water temperatures (Skliris et al., 2001) even the disease was originally called “summer disease” (Bellance and Gallet de Saint-Aurin, 1988; Maltese and Bovo, 2007). A study by Mori et al. (2003) was showed that there was a greater mortality rate in temperatures above 25 C. Also, the genotypic variants have different optimal growth temperatures: 15 C 20 C for BFNNV, 20 C for TPNNV, 20 C 25 C for SJNNV, and 25 C 30 C for RGNNV (Mori et al., 2003) (Fig. 30.5).

SUBCLINICAL INFECTION There is little information about subclinical infection. Many studies showed that several fish species can be infected by nodavirus without showing signs of disease (Barker et al., 2002; Castric et al., 2001; Johansen et al., 2004; Skliris and Richards, 1999). In the acute stage of the disease, clinical signs and histopathologic lesions were observed. In larval stages, highest mortality was reported, and older fish showed lower losses, and surviving fish became subclinical carriers. In these fish, CNS cells contain nodavirus particles (Johansen et al., 2004). Adult fish are also affected, and viruses continue to proliferate in neural cells throughout their lives. The disease usually has no signs, and it is mostly subclinical, but these fishes show loss of appetite and poor weight being the only indicator of the presence of VNNV.

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However, these fish in stressful environments are showed mortalities because their immune responses reduced. Subclinical infections can show hidden nodavirus replication sites, and recent researchers focused on the possibility of transmission of nodavirus from such fish (Castric et al., 2001; Barker et al., 2002; Johansen et al., 2002, 2003, 2004). However, subclinically infected fish without signs, immunohistochemistry (IHC) test can be positive for nodavirus within the CNS (Johansen et al., 2002). Johanson et al. (2004) reported large amounts of nodavirus-like particles in neural cells of subclinical infections by electron microscopy. In this study, nodavirus was isolated in cell culture from subclinically infected fish 1 year after the acute outbreak. A study of Atlantic halibut resulted in the presence of VNNV in a subclinical infection revealed by reverse transcription (RT)PCR, enzyme-linked immunosorbent assay (ELISA), and IHC (Grove et al., 2003; Johansen et al., 2004). These results showed that H. hippoglossus can be a carrier for a long time after infection, thus asymptomatic carriers are considered the main source of past infections. The presence of the virus in gonads and intestine, stomach, kidney, and liver of adult striped jack (P. dentex) was reported (Nguyen et al., 1997), but no nodavirus particles in CNS were observed. Betanodavirus was detected in neural cells of asymptomatic Sciaena umbra and S. aurata by Comps and Raymond. (1996), Castric et al. (2001) and Dalla Valle et al. (2000). These observations show that this species could play a main role as healthy carriers. Several studies have proved that some species can act as asymptomatic carriers (Glazebrook et al., 1990; Johansen et al., 2003; Skliris and Richards, 1999).

CHRONIC FORM Sea bass developed clinical signs of the disease after over 8 months in freshwater, possibly demonstrating that even asymptomatic carriers can develop clinical disease in real farm situations (Athanassopoulou et al., 2003). The fish survived VNN acute infection, usually become persistently infected, and the viral persistence can extend as long as 1 year (Johansen et al., 2004; Kai and Chi, 2008). The chronic form of VNN was observed seldom in fish species. This form of the disease was reported in L. aurata from Caspian Sea (Zorriehzahra et al., 2016). This disease was known as thalassemia disease between fishermen in the north of Iran (Zorriehzahra et al., 2019) (Fig. 30.6). This form of disease takes may be several years to progress (4 5 years) from the asymptomatic state to clinical form. In this form, clinical signs, such as emaciation, poor growth, abdominal distention and dermal edema, color change, lethargic behavior, floating impassively on

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FIGURE 30.6 VNN mortality in golden grey mullet in the Caspian Sea (Zorriehzahra et al., 2005).

the surface of the water and cachexia, were observed (Zorriehzahra et al., 2013). The chronic form of nodavirus infection with any clinical signs was reported in several cultured and wild marine fish in Japan (Gomez et al., 2004), brown meagre (S. umbra), and Atlantic salmon (Salmo salar) (Dalla Valle et al., 2000; Baeck et al., 2007; Shetty et al., 2012).

VIRAL DETECTION Virus Isolation Cell culture has high sensitivity and specificity. Samples for cell culture are, ideally, live whole fish; otherwise, chilled or frozen tissues can be used. Betanodavirus isolation in cell culture has been found difficult in the past because of the lack of susceptible cell lines (Roongkamnertwongsa et al., 2005). A number of cell lines are now available for the culture of betanodaviruses. The striped snakehead cell line (SSN-1) originally developed by Frerichs et al. (1996) has been shown to be permissive for 17 isolates of fish nodaviruses, encompassing the RGNNV, SJNNV, TPNNV, and BFNNV types (Iwamoto et al., 1999), and Iwamoto et al. (2000) reported that six cell clones were derived from the SSN-1 cell line, which is composed of a mixed cell population and persistently infected with a C-type retrovirus (SnRV). These clones were susceptible to four piscine nodavirus strains

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belonging to different genotypes, such as SJNNV, RGNNV, TPNNV, and BFNNV (striped jack, red-spotted grouper, tiger puffer, and barfin flounder NNVs). Three clones, designated A-6, E-9, and E-11, were highly permissive to nodavirus infection and production. The virus-induced cytopathic effects (CPEs) appeared as cytoplasmic vacuoles and intensive disintegration at 3 5 days of postincubation. These observations were highly reproducible and formed the basis for a successful virus titration system. Quantitative analysis using the cloned E-11 cell line clearly revealed differences in the optimal growth temperatures among the four genotypic variants: 25 C 30 C for strain SGWak97 (RGNNV), 20 C 25 C for strain SJNag93 (SJNNV), 20 C for strain TPKag93 (TPNNV), and 15 C 20 C for strain JFIwa98 (BFNNV). Electron microscopy demonstrated SnRV retrovirus particles only in A-6 and E-9 cells, but PCR amplification for the pool gene and LTR region of the proviral DNA indicated the presence of the retrovirus in the other clones, including E-11. Further susceptible cell cultures (GF-1) have been developed from the groupers Epinephelus coiodes (Chi et al., 1999) and may be used for research and diagnostic purposes provided in which sensitivity is regularly monitored [Office Internationales Epizooties (OIE), 2013]. Lai et al. (2001) have developed other cell line from E. awoara, also Qin et al. (2006) have made other cell line (GS) from the groupers E. coiodes that support grouper NNVs, but the lines have not been tested for other types of fish nodaviruses. Permissive cell lines have also been developed from barramundi/ Asian sea bass tissues (Chong et al., 1990; Chang et al., 2001). In the past, cell culture was used mainly for research, but it has shown promise as a means of expediting diagnosis of betanodavirus infections in a combined cell culture PCR technique (Iwamoto et al., 2001). In SSN-1 cells the CPE appears on the third day post infection and is characterized by the appearance of intracellular vacuolar lesions unevenly distributed throughout the cell monolayer. These vacuolar lesions initially are isolated and began assuming the form of vacuolized cellular aggregates after the passage of hours. About 72 hours post infection, their number and size increase considerably, and the cellular monolayer is gradually replaced by cellular lysis until a complete destruction happens (Maltese and Bovo, 2007) (Fig. 30.7).

Enzyme-Linked Immunosorbent Assay ELISA is the rapid and sensitive test in order to detect specific nodavirus antibodies from blood and body fluids of infected fish (Vatanabe et al., 2000). This method can be employed to identify infected different hosts especially from vectors and broodstock in order to control and

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FIGURE 30.7 SSN-1 cell line inoculated with retina and brain filtrate that not showing CPE. CPE, Cytopathic effect.

prevention of vertical transmission of disease (Arimoto et al., 1992; Arthur, 2005). The efficacy of this assay was confirmed by Vatanabe et al. (2000) to the identification of nodavirus antibodies from barfin flounder broodstock and Arimoto et al. (1992) from striped jack. Fenner et al. (2006) could detect 103 104 TCID50 units of betanodavirus by antigen capture ELISA from infected tissue of juvenile barramundi (L. calcarifer, Bloch). By this assay, 17% and 18% sera of wild and farmed European sea bass broodstock were positive for nodavirus antibodies, respectively (Breuil et al., 2000). In a similar study, 9% sera of commercial barramundi were positive for this virus (Huang et al., 2001) by ELISA method.

Immunofluorescence Antibody Test IFT is a rapid, economical, potent, and practical method for the screening of Nodaviridae that use fluorescent-labeled antibodies to detect specific antigens from target tissues including brain, spinal cord, and retina (Harlow and Lane, 1988). In this assay by preparing of the histopathological section from brain (CNS) or other tissues such as eye, swim bladder, spleen, kidney, and liver staining with specialized immunofluorescence technique indicates the localized virus in target tissues (Sanz and Coll, 1992). The binding of antibodies to target tissues, cells, or organisms can be visualized if those antibodies are directly coupled to a fluorochrome or indirectly bound by a fluorescent reagent. Indirect fluorescent antibody test showed at least 20% of infected golden grey mullet (L. auratus) fish to VNN disease have a positive reaction to

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betanodavirus antigens in the optic nerve, outer molecular, and granular layers of the brain and inner and outer nuclear layers of retina (Ghasemi et al., 2013). Molecular Diagnostic Techniques Molecular methods have been used in the diagnosis of NNV. These methods were developed for the rapid, convenient, and sensitive diagnosis of the NNV pathogen in the fish and including PCR (Dalla Valle et al., 2000; Grotmol and Totland, 2000; Muroga, 1994), real-time PCR (Dalla Valle et al., 2005; Hodneland et al., 2011; Starkey et al., 2004), and nucleic-acid sequence amplification (NASBA) (Starkey et al., 2004). The most important technique for the amplification of RNA is RT followed by PCR. For the first time, Nishizawa et al. (1995) established NNV RTPCR detection method. With the development of the technology, the nucleic acid extraction technique improvement now allowed an easy, fast, and high-quality RNA preparation, and the availability of more NNV genome sequences facilitated the primer optimization (Mu et al., 2013). More recently, RT-PCR assays with or without nested PCR have been developed as a strong diagnostic tool alone or in combination with cell culture (Dalla Valle et al., 2005; Iwamoto et al., 2001). These PCR protocols have greatly improved test sensitivity, allowing better control of VNN infection through identification and eliminate of infected spawners (Dalla Valle et al., 2005), for example, striped jack (P. dentex) broodstocks screen for NNV to prevent from vertical transmission of this pathogen to the larval offspring (Muroga, 1994). For this matter amplify a portion of the coat protein gene (RNA2) of betanodavirus and is a powerful and sensitive method for identification of the infection (Azad et al., 2005; Barker et al., 2002; Grotmol and Totland, 2000; Nishizawa et al., 1997). Although it has been shown that nested RT-PCR is 10 100 times more sensitive than the previously reported RT-PCR methods (Dalla Valle et al., 2000; Thiery et al., 1999). Moreover, since conventional PCR is a nonquantitative technique, the actual copy number of the viral template in samples cannot be determined (Dalla Valle et al., 2005; Starkey et al., 2004). So Dalla Valle et al. (2005) described the setting up of two real-time, SYBR Green I-based, PCR diagnostic assays targeting both RNA1 and RNA2 of betanodavirus for its quantitative detection in biological samples. The sensitivity of this technique was compared with that of conventional RT-PCR assays previously developed for betanodavirus (Dalla Valle et al., 2000; Grotmol and Totland, 2000; Mu et al., 2013) and with the results of routine virus isolation test (Delsert et al., 1997; Iwamoto et al., 2001), to check for a correlation between measured viral RNA load and virus isolation response. It has been powerful to study transmission and development of this viral infection in juvenile (Hodneland et al., 2011). The

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most important primers that used for the detection of NNV are shown in Table 30.1. NASBA is another useful method that consists of an isothermal method for nucleic acid amplification that is particularly suited to RNA targets (Deiman et al., 2002). The method amplifies a targetspecific product through oligonucleotide primers and the coordinated activity of three enzymes, such as reverse transcriptase, RNase H, and T7 RNA polymerase (Deiman et al., 2002; Starkey et al., 2004). This method has been developed for the detection of betanodavirus by Starkey, and the sensitivity of this procedure was compared to a conventional single-tube RT-PCR assay (Starkey et al., 2004). Currently, real-time PCR technique was recommended by Office Internationales Epizooties (OIE) (2013) as a fast and confidential method for the diagnosis of betanodavirus in the world.

DISEASE TRANSMISSION Virus transmission from a fish to other fish is possible, and potential of cross infections and initial source of infection is important in aquaculture management (Johansen et al., 2003; Munday et al., 2002). Munday et al. (1992) suggested that water could contain virus particles excreted by carriers. Experimental studies by several authors completely support the horizontal transmission route; but the vertical transmission is proposed for some species (Johansen et al., 2002; Maltese and Bovo, 2007). Some researchers have been approved vertical transmission recently and their investigations demonstrate a possibility of vertical transmission of the betanodavirus. So it should emphasize the importance and vital need for the screening of eggs and larvae for evolving suitable preventive and prophylactic health management strategies in broodstocks.

The Entry and Progression of Nervous Necrosis Virus In bath-challenged striped jack larvae, vacuolation was first observed in the spinal cord above the swim bladder, later in the brain, and then in the retina, indicating spinal cord is the initial site for NNV proliferation (Nguyen et al., 1996). In naturally affected young larvae the virus was detected in the epithelial cells of the skin and in the intestinal epithelium, concurrently with the nerve cells of CNS in the early stage of NNV infection. At the early development stage of fish, epithelial cells of the skin may be susceptible to NNV. The neurotropism of NNV indicates that virus might gain access to CNS via peripheral nerves, for example, via the automatic nerves link to the digestive tract via sensory

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and/or motor nerves link to the epithelium of skin (Nguyen et al., 1996; Grotmol et al., 1999). Seven-banded grouper at the grow-out stage was able to be infected with NNV by per nasal challenge. NNV penetrated the nasal epithelium, passed through the olfactory nerve and olfactory bulb, and invaded olfactory lobe (Tanaka et al., 2004). By intramuscular (I.M.) challenge model, NNV passed through the peripheral nervous system in lateral muscular tissue, transported through axon to the spinal cord. NNV may invade the CNS via blood circulation from the injection site (Tanaka et al., 2004).

Tissue Tropism The main histopathological changes observed include degeneration and vacuolation of cells in the brain, retina, and nerve cord, which clearly demonstrate that VNN virus has a neurotropism with replication sites in these sensitive tissues (Johansen et al., 2004), vacuolation of the brain’s gray matter, particularly, in the optic tectum and cerebellum with a lesser involvement of the telencephalon and medulla oblongata. Also, vacuolation of the nuclear layers of the retina intracytoplasmic inclusions (up to 5 µm in diameter) in nervous cells and nervous necrosis of the spinal cord, spinal ganglia brain, and retina were considered important lesions in VNN. These agents have more density and concentration in the CNS (Kokawa et al., 2008; Oh et al., 2002; Parameswaran et al., 2008). In the other hand, CNS and retina are virus targets due to the affinity of nodaviruses for nervous cells and their ability to cause histopathological changes in nervous tissues (Mori et al., 1992; Comps and Raymond, 1996). First replication sites in the larva of P. dentex is the spinal cord, and then the virus could reach to the retina via the brain by traveling up the optic nerve (Nguyen et al., 1996; Maltese and Bovo, 2007). The viral antigens in the gonads, intestine, stomach, kidney, and liver of carriers were observed, which indicate a major difference between the carrier and clinically affected fish (Castric et al., 2001; Comps and Raymond 1996; Nguyen et al., 1997; Dalla Valle et al., 2000). These findings suggest vertical transmission of offspring through the gonadal and intestinal products (Nguyen et al., 1997; Maltese and Bovo, 2007). Also, Mladineo (2003) detected that viral antigens in the olfactory lobes that show like mouth and nasal cavity are a point of viral entry.

Horizontal Transmission The horizontal transmission route completely confirmed and well recognized in many experimental studies in marine fish larva by several

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authors with mortalities ranging from as low as 4% to as high as 100% (Nakai et al., 2009; Pakingking et al., 2009; Parameswaran et al., 2008; Tanaka et al., 2003). Horizontal transmission of betanodavirus also caused several outbreaks (Grotmol et al., 1999; Le Breton et al., 1997; Peducasse et al., 1999). Horizontal transmission of nodavirus from infected sea bass to sturgeon was detected for the first time by (Athanassopoulou et al., 2004). This caused serious pathology and clinical signs and reaching a total mortality of 30% per day. Results revealed a close link between sea bass and sturgeon nodavirus, which indicates that the virus has passed horizontally in the farm from sea bass to sturgeon (Ransangan et al., 2011) reported that Betanodavirus can be transmitted horizontally to healthy fish through contact with water contaminated with infected fish. This study showed virus transmits to other fish species and suggests that the virus has a wide range of fish hosts. Moreover, it suggested that SJNNV in larval striped jack spreads by horizontal transmission from diseased larvae to healthy ones by cohabitation (Arimoto et al., 1993). Also, newly hatched larvae infection via the waterborne route (horizontal transmission) could not be disregarded (Pakingking et al., 2009). Munday et al. (1992) suggested that water could contain virus particles excreted by carriers. Arimoto et al. (1993) showed healthy fish could be infected by bathing with contaminated waters. Bath by infected water and intraperitoneal inoculation E. akaara juveniles showed histopathological damages 10 14 days after exposure with lower mortality rates (Mori et al., 1991). Successful transmission of betanodavirus infection via immersion route associated with clinical signs such as abnormal swimming behavior and high infective virus titers ranging was carried out by Pakingking et al. (2009). Grotmol and Totland (2000) had shown that nodavirus survived in pH 2 9 and seawater at 15 C for over a year. This result increases the likelihood of horizontal transmission. Temperature and virulence of the virus determine the extent of lateral spread occurrence (Castric et al., 2001). About 28% mortality rate following I.M. inoculation of brain homogenate in D. labrax juveniles is reported by Thiery et al. (1997). Intraperitoneal injection of infected materials in E. malabaricus resulted in clinical signs that were comparable to the natural disease and induced 40% 60% mortality (Boonyaratpalin et al., 1996).

Vertical Transmission Vertical transmission thought to be the main route of nodavirus infection from subclinically infected spawners in farmed fish in which

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broodstock acting as the main virus reservoir (Johansen et al., 2002; Johansen et al., 2004). Vertical transmission in striped jack, barfin flounder, and European sea bass has been demonstrated via eggs or genital fluids (Castric et al., 2001). The virus has been detected in fertilized eggs and larvae in experimentally infected broodfish (Breuil et al., 2002). Indirect ELISA test of striped jack and barramundi showed high frequency in the ovaries of spawners. This finding indicates that the virus transmits vertical to larva (Arimoto et al., 1992, 1996; Azad et al., 2006). Pakingking et al. (2009) suggested that vertical transmission of the virus from VNN-infected parent to offspring in grouper and sea bass could be the most feasible route. These data showed that disease might be transmitted from broodfish to their offspring, but it is not clear whether the real intraovarian transmission is involved or an external contamination (mechanical transmission) occurs (Maltese and Bovo, 2007). Nonetheless, the absence of infectious agents in the broodstock was established. Regardless of broodstock, the absence of infection in eggs and larvae was determined with a very high degree of confidence (Hick et al., 2011). Vertical transmission is strongly suspected because of the high prevalence of the infection in the very earliest larval stages provided with water treatment (Arimoto et al., 1992; Comps and Raymond, 1996; Yoshimizu et al., 1987; Grotmol and Totland, 2000).

Other Transmission Routes It is clear that nodaviruses can be transmitted by vertical and horizontal mechanisms, but it is not known whether the virus can be transmitted via a mechanical transmission. Fish can infect by I.M. and intraperitoneal injection or by bathing in infected tissues (intranasal route) experimentally (Skliris and Richards, 1999). Asian sea bass and brown-marbled grouper fingerlings transmitted the virus in the culture tank horizontally via cannibalism (Ransangan et al., 2011). The possibility of live feeds that facilities the disease for larvae, such as Artemia salina and the rotifer Brachionus plicatilis, considered by Skliris and Richards (1998), can play a key role in the transmission of the disease. The absence of VLPs in cell cultures test of these two invertebrates convinced the authors that the risk, in this case, existed only at the level of mechanical carrier following superficial contamination. Also, raw fish mainly used for the broodstock feeding is a possibility for transmission of the disease (Mori et al., 2005).

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PROTECTIVE AND CONTROL STRATEGIES Some fish species, such as sea bass and groupers, are susceptible to RGNNV in growing out stages and VNN causes high mortality in the open sea (Nakai et al., 2009). The immune system at the very early stage is not mature enough to accept oral or bath immunization, so the prevention of transmission of VNN becomes critical (Kai et al., 2010). The deletion of virus-carrying animals would be the best means of control in vertical transmission route. Thus the elimination or separation of infected spawners is the best way to prevent (Hick et al., 2011; Munday and Nakai 1997; Mushiake et al., 1994). Mushiake et al. (1994) reported a successful control of VNN by the deletion of infected striped jack (P. dentex) broodstock detected by PCR. In the same study by Breuil et al. (2000) disease is controlled by the elimination of European sea bass (D. labrax) broodstock, in which betanodavirus-specific antibodies were detected in blood samples. However, Anderson and Oakey (2008) reported that the exclusion of barramundi broodstock based on the results of a nested RT-PCR assay did not reduce the VNN disease rate. Schipp et al. (2007) developed a new procedure for larval barramundi to prevent vertical and horizontal transmission of betanodavirus infection based on the new production method provided farming and nutrition. However, outbreaks of VNN continued to occur (Hick et al., 2011). Also, the disinfection of fertilized eggs by ozone has been recommended to control the vertical transmission of betanodavirus in Atlantic halibut (H. hippoglossus) (Grotmol and Totland, 2000). Arimoto et al. (1996) reported that 0.2 µg/mL ozone disinfects fertilized eggs in striped jack and also 4 µg/mL was reported for halibut by Grotmol and Totland (2000). These results indicate that NNV transmitted from the maternal sexual fluid was via the surface of the eggs (Kai et al., 2010). The horizontal transmission of VNN infection may be due to via influent and rearing water, utensils, vehicles, and human activity (Nakai et al., 2009), and some effective disinfectants can inactive the virus and prevent spread of diseases, such as ozone, acid peroxygen, sodium hypochlorite, and benzalkonium chloride (Arimoto et al., 1996; Frerichs et al., 2000). A vaccination method is essential to prevent the disease, especially, during primarily stages, and some researcher reported effective procedures in controlling the disease (Nakai et al., 2009). Recombinant viral coat protein expressed in Escherichia coli injected to fish (Husgard et al., 2001; Tanaka et al., 2003). Injection of like particles expressed in a baculovirus expression system carried out by Thiery et al. (2003) and inactivated virus by Yamashita et al. (2005). Injection of the recombinant protein in

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adult striped jack caused releasing virus neutralizing antibodies (Munday and Nakai, 1997), thus, vaccination seems to be a good way for the control of VNN. Vaccinating broodfish could reduce vertical transmission of VNN and will be more acceptable to the farmers (Kai et al., 2010). Strict hygiene can help to control VNN within hatcheries [Munday and Nakai, 1997; Office Internationales Epizooties (OIE), 2013]. No recycling of water and chemical sterilization of seawater during each hatching cycle was successful to reduce VNN disease in a barramundi hatchery (Anderson et al., 1993). General hygiene practices should be the UV treatment, sanitary barriers, regular following and disinfection of tanks and biological filters, disinfection of utensils and decrease of stress factors and density of larvae and juveniles [Office Internationales Epizooties (OIE), 2013]. Test of specific antibody activity with ELISA in each single broodfish, PCR on sexual products and disinfection of eggs with ozone has been also proposed as an integrated control method of VNN in barfin flounder (Watanabe et al., 2000).

CONCLUSION AND SUGGESTION FOR FUTURE RESEARCH At first, VNN occurred was reported in 13 fish species in four families, but at present, this transmissible viral disease was recorded in more 120 different farmed and wild fish and invertebrate species. Also, unlike of other fish viral diseases that can affect special order or family such as IHN or VHS especially can affect on cold water fish. VNN virus can affect a lot of fish species such as cold water fish (BFNNV serotype), warm water fish (RGNNV serotype), and other fish species such as ornamental fish. One of the other aspects of VNN is the neurotropic property of this invasive virus same as rabies and bovine spongiform encephalitis (BSE) in mammals that can be the effect on the CNS. In a comparison of VNN and BSE, some similarities could be considered seriously. Spread in nervous tissue, necrosis, and vacuolation as some important aspects that should be noticed in both diseases. Also, related clinical signs were similar to due to same sources. For example, CNS necrosis effects lead to paralysis in cow and lethargy and uncommon swimming behaviors in affected fish. Also, some aggressive behavior maybe occurred in infected cattle such as pushing the head to maintained box and kicking up to the owner, and this behavior maybe occur as darting movement in affected fish in VNN. Concerning to zoonotic aspect of BSE that have been reported such as Creutzfeldt Jakob disease in some human morbidity, we should

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notice a zoonotic aspect of VNN seriously in future studies. In this regard, some new findings were reported that using of infected meat to BSE virion could affect the brain of consumer fish, and same vacuoles were occurred in the brain of fish affected. Furthermore, the prospect of farmed fish being contaminated with infectious mammalian PrPSc, or of a prion disease developing in farmed fish is alarming and requires further evaluation (Salta et al., 2009). So, it could be concluded that statement of specific host for each virus could be mistrusted such as BSE, avian flu and maybe for VNN in the near future. As a final point, a zoonotic aspect of VNN is a very important attitude that should be investigated in future complementary studies. Furthermore, the production of recombinant vaccines against VNN virus, applying some immunostimulant drugs, epidemiological investigation of pathogen global spreading in the new regions should be considered in future studies.

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Further Reading Bovo, G., Nishizawa, T., Maltese, C., Borghesan, F., Mutinelli, F., Montesi, F., et al., 1999. Viral encephalopathy and retinopathy of farmed marine fish species in Italy. Virus Res. 63, 143 146. Hick, P., Tweedie, A., Whittington, R., 2010. Preparation of fish tissues for optimal detection of betanodavirus. Aquaculture 310, 20 26. Office Internationales Epizooties (OIE), 2006. Viral encephalopathy and retinopathy. In: Manual of Diagnostic Tests for Aquatic Animal. Office International des Epizooties (OIE), Paris, France, 169 175. Zorriehzahra, M.E.J., 2018. VNN Professional Forum in LinkedIn Networking. ,https:// www.linkedin.com/groups/4131847..

EMERGING AND REEMERGING VIRAL PATHOGENS