Annexe 1

REVIEW OF PATHOGENS OF PRAWNS

ContentsSection 1 Disease agents which will be considered further in the IRA

VirusesYellow-head virus (YHV)White Spot Syndrome (WSSV)Taura Syndrome Virus (TSV)Infectious Hypodermal and Hematopoietic Necrosis Virus (IHHNV)Baculovirus penaei (PvSNPV)Baculoviral Midgut Gland Necrosis Virus (BMNV)Monodon Baculovirus (MBV)Infectious pancreatic necrosis virus (IPNV)Rhabdovirus of Penaeid Shrimp (RPS)

BacteriaNecrotizing Hepatopancreatitis (NHP)Vibrio species (vibriosis)RickettsiaAerococcus viridans var. homari

ParasitesMicrosporidiaHematodinium-like organismParauronema spp.

Section 2 Disease agents which will not be further considered in the IRAViruses

Lymphoid Organ Vacuolization Virus (LOVV)REO-III AND REO-IVHepatopancreatic Parvo-Like Virus (HPV)Lymphoidal Parvo-Like Virus (LPV)Lymphoid Organ Virus (LOV)Gill Associated Virus (GAV)Spawner-isolated Mortality Virus (SMV)Penaeid Haemocytic Rod-shaped Virus (PHRV)

BacteriaMycobacteria (mycobacteriosis)Chitinoclastic Bacteria (other than vibrios) Associated with Shell DiseaseAeromonas sp. and Pseudomonas sp. (Necrosis And Septicemias)Epibiont Bacteria which Cause Fouling (Principally Leucothrix mucor)

FungiFusarium solani (Fusariosis)Lagendium and Sirolpidium Species (Larval Mycosis)

ParasitesHaplosporidiaGregarinesOther Miscellaneous Parasites

REVIEW OF PATHOGENS OF PRAWNS

The following information is drawn from Chapter 3 of the Scientific Review Of Prawn Diseases report to AQIS by Ausvet Animal Health Services in 1997 (Ausvet report). This paper only considers the scientific evidence relevant to a discussion of the epidemiological features of prawns pathogens. Ausvet’s report contained other information on the evaluation of quarantine risk associated with prawn pathogens and the identification of risk management options. That information will be considered later in the IRA process.

Section 1 of this paper describes disease agents which are identified for further consideration in the IRA. RAP comments appear in italics. Several disease agents were not covered in the Ausvet report and information on these has been included in italics, eg. infectious pancreatic necrosis.

Section 2 of this paper contains information from the Ausvet report on the disease agents that have been classified as not requiring further consideration in this IRA. RAP comment is not provided on this information.

Section 1 Disease agents which will be considered further in the IRA

Viruses

Yellow-head virus (YHV)

Yellow-head disease (YHD) was first noted by Limsuwan (1991) in cultured Penaeus monodon adults in central Thailand. It appears to be widespread in cultured stocks of P. monodon and is a serious disease of cultured P. monodon in South-East Asia and India. There is also some evidence to suggest that YHD may have been associated with the P. monodon industry crash in Taiwan in 1986-1987 and also with epizootics in Indonesia, Malaysia, China, India and the Philippines (Lightner, 1996). In 1995 cultured P. setiferus production in Texas, USA decreased dramatically, allegedly due to the introduction of yellow-head virus (YHV) and white-spot baculovirus (nonoccluded bacilliform virus) with raw and frozen prawn products imported from Thailand (Lightner et al., 1997).

YHV is an RNA virus (Wongteerasupaya et al., 1995) with a number of properties in common with plant and crab rhabdoviruses (Nadala et al., 1997). YHV has now been shown to be a coronavirus based on sequence information (Peter Walker, personal communication ).

YHD effects primarily juvenile to subadult prawns (Boonyaratpalin et al., 1993). The American penaeids P. setiferus, P. aztecus and P. duorarum developed disease when infected experimentally with YHV (Flegel et al., 1995) as did P. vannamei and P. stylirostris (Lu et al., 1994a). High health P. vannamei, produced in Hawaii, are being tested for susceptibility to YHV. P. merguiensis and Metapenaeus ensis were infected successfully in laboratory challenge studies, although they were resistant to YHV in ponds (Flegel et al., 1995b). YHD has not been reported in Australia.

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Two viruses, one pathogenic (gill-associated virus, GAV) and the other benign (lymphoid organ virus, LOV) were found in cultured P. monodon in Australia (Spann & Lester, 1997). GAV and LOV have approximately a 1% difference over a 400 base pair PCR sequence of a highly conserved RNA polymerase gene (Peter Walker, personal communication). Thai YHV has 15% difference to the Australian viruses for the same targeted gene sequence (Peter Walker, personal communication). However this 400 base segment represents only a small portion of the ~20,000 base genome, and because much larger cDNA probes made to Thai YHV react quite efficiently to both GAV and LOV, these viruses appear to be too closely related to merit three different names. This is especially true for LOV and GAV which have virtually identical sequences, geographic distribution, and viral morphologies (Don Lightner,personal communication).

P. merguiensis is refractory to infection with YHV (Loh et al., 1997).

Clinical signsPrawns with YHD display yellow colouration of the dorsal cephalothorax caused by the underlying yellow hepatopancreas showing through the translucent carapace. Within the ponds, infected animals, usually between 5 and 15 g (Limsuwan, 1991), begin consuming feed at an abnormally high rate for several days then cease feeding entirely. One day after cessation of feeding, moribund prawns may be seen swimming slowly near the edges of the pond. By the third day, mass mortality occurs and the entire crop is typically lost (Chantanachookin et al., 1993). Gross PathologyThere is very little gross pathology associated with YHD. Infected prawns usually have a pale yellow hepatopancreas and may also have pale yellow to brown gills (Boonyaratpalin et al., 1994). Moribund prawns collected from ponds afflicted with yellow-head disease in Thailand do not always display yellow colouring of the cephalothorax, indicating that this may not be a reliable sign of YHV infection (Flegel et al., 1992).

HistopathologyMoribund prawns suffering YHD usually have extensive abnormalities in the lymphoid organ. These include foci of necrotic cells which resemble degenerate tubules with occluded lumens and contain cells with hypertrophied nuclei, pyknotic nuclei, large vacuoles and cytoplasmic, basophilic, Feulgen-positive inclusions. Similar inclusions may also be found in the interstitial tissues of the hepatopancreas, connective tissues underlying the midgut, cardiac tissues, haematopoietic tissues, haemocytes and gill tissues ( Flegel et al., 1992; Chantanachookin et al., 1993). Earlier cellular changes in infected cells may include nuclear hypertrophy, chromatin margination and lateral displacement of the nucleolus (Lightner, 1996).

The viral agent responsible for YHD was detected in 1992 (Chantanachookin et al., 1993). Electron microscopy of lymphoid organ and hepatopancreatic interstitial cells, haemocytes and epithelial gill cells revealed rod-shaped virions measuring 173 13 nm by 44 6 nm within the cytoplasm of abnormal cells. Very long filaments, up to 800 nm in length, also were observed in the cytoplasm and appeared to be precursors of enveloped virions. Similar precursors and virions were found in

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apparently healthy broodstock. Viral envelopes appeared to be acquired by passage through the host endoplasmic reticulum resulting in the collection of virions within vesicles. Often virions were packed densely into paracrystalline arrays. Acquisition of capsids and envelopes often proceeded fragmentation of the long filaments into shorter rod-shaped virions (Chantanachookin et al., 1993; Boonyaratpalin et al., 1994; Flegel et al., 1995a).

DiagnosisPresumptive diagnosis of yellow-head disease is based on the presence of clinical signs and the history of disease in the culture facility, region or species (Lightner, 1996). A haemocyte staining method has been developed for the rapid diagnosis of the early stages of YHD (Anon, 1992). This involves taking a sample of haemolymph from the ventral or cardiac sinuses, diluting the prawn haemolymph in 10% seawater formalin, fixing it on a slide in methanol and staining with Wright’s stain and Giemsa. Haemocytes may then be inspected for nuclear pyknosis and karyorhexis by brightfield microscopy (Nash et al., 1995). A drop of undiluted haemolymph may also be investigated for YHD using phase contrast microscopy. During the later stages of infection, when the haemocytes have been depleted, YHD may be diagnosed by identifying characteristic basophilic pyknotic nuclei in rapidly stained gill mounts (Flegel et al., 1995a and b).

When investigating a suspected outbreak of YHD, profiles of moribund prawn gills and the haemolymph of non-symptomatic prawns from the same pond should be composed and bacteria should be absent from the haemolymph smears. YHV infection may be confirmed by TEM demonstration of rod-shaped enveloped virions and filamentous nucleocapsids in the cytoplasm of infected cells (Chantanachookin et al., 1993).

A diagnostic PCR for YHV has been developed (Wongteerasupaya, et al., 1997). PCR of nucleic acid using primers designed from the highly conserved sequence of the RNA Polymerase gene (L Protein gene) of insect rhabdoviruses gave a predicted 450 bp PCR product (Flegel et al., 1996). Forty-five recombinant clones have been obtained from this product, 2 of which hybridised with YHV RNA and not with P. monodon genomic DNA (Wongteerasupaya et al., 1996). A diagnostic DNA probe has been prepared (Wongteerasupaya, 1996) and is being evaluated.

Transmission and potential carriersYHV in Thailand may be transmitted to cultured penaeids in the ponds from wild crustaceans, introduced to the ponds with incoming water. Flegel et al. (1995) reported the use of P. monodon as a bioassay to demonstrate that the shrimp Palaemon styliferus and Acetes sp., which are often found in prawn ponds, were susceptible to YHV and acted as carriers for the virus (Charoen Pokphand CP and Department of Fisheries, Thailand, unpublished data). Wild-caught P. monodon broodstock do not appear to be the primary source of YHV as few broodstock screened since the beginning of the epizootic in 1992 have been found infected (Flegel et al., 1997a). Within ponds, YHV is transmitted by water and by cannibalisation of moribund prawns and infected carcasses. Birds have been associated with the spread of YHD, however, CP (unpublished data) have found that YHV cannot be transmitted via the faeces of common pond birds that had eaten YHV-infected prawns.

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There is evidence that the spread of YHV to US may have resulted from frozen imported ‘commodity’ prawns (Lightner et al., 1997).

ViabilityYHV remains viable in aerated seawater for 3 to 4 days, depending on the amount of virus present (Flegel et al., 1995a). YHV is transmitted experimentally by bathing prawns in water from infected ponds, in water containing membrane-filtered extracts of YHV, and in water containing infected individuals. Clinical symptoms of YHD are expressed within 7 to 10 days of exposure, depending on the amount of virus extract added to the water (Flegel et al., 1995a). The time YHV remains viable in prawn carcasses is not known as dead and moribund prawns are usually cannibalised by other prawns or removed from the pond. It is expected that the lifetime of YHV in a rotting prawn carcass would be short due to bacterial growth and the release of digestive enzymes (Tim Flegel, personal communication). The viability of YHV in frozen prawns is not known, however Dr. T. Flegel has successfully infected healthy prawns with YHV using tissues stored at –60oC for 6 months. YHV purification has proven difficult due to the labile nature of the viruses, therefore it is highly likely that the viability of YHV would decline rapidly once uncooked prawns are thawed.

On the other hand, Lightner et al. (1997) suggest that outbreaks of YHV in the US can be traced to waste from imports of frozen prawn; indicating that the virus may remain viable after thawing. Further, the YHV- like viruses that occur in Australia are known to survive three freeze-thaw cycles (Leigh Owens, personal communication).

PreventionYHV in Thailand is controlled using closed and semi-closed systems (Limsuwan, 1996). In these systems, in-take water is treated before use with calcium hypochlorite at a rate of 300 kg/ha to kill wild crustaceans which may carry YHV. In semi-closed systems, no water exchange takes place within the ponds until 30-60 days post-stocking while in closed systems there is no water exchange during the culture cycle. Additional preventative measures, such as excluding potential carriers, not using fresh feeds and not exchanging water for 4 days when it is known that an infected pond in the area is discharging water, have proven effective against YHD (Flegel et al., 1995a).

Present status of yellow-head diseaseIn Thailand during 1992 and 1993, pond side losses caused by YHV were estimated at about US$ 40 Million (Flegel et al., 1997b). Mortalities caused by YHV in Thailand decreased in 1996, yet the prevalence of the virus remains high (Pasharawipas et al., 1997), indicating that YHV may have mutated and adapted to P. monodon as a host. In the Songkhla and Nakornsrithammarat districts of southern Thailand, diseases of cultured penaeids are controlled by pumping pond discharge waters one km off-shore beyond a sand bar which was previously restricting the discharge of contaminated waste water. Although white-spot virus is now considered the primary cause of mortality of cultured prawns in Thailand, YHD remains a serious problem. YHV causes a significant decrease in prawn growth rates (Timothy Flegel, personal communication). When epizootics due to YHV(and WSSV) occur, emergency harvests are commonly employed in Asia to salvage marketable prawn crops (Lightner et al., 1997).

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ReferencesAnonymous, 1992. Routine and rapid diagnosis of yellow-head disease in Penaeus

monodon. Asian Shrimp News, October. Issue No. 12: 2-3.Chantanachookin, C. Boonyaratpalin, S. Kasornchandra, J., Sataporn, D.,

Ekpanithanpong, U., Supamataya, K., Sriurairatana, S. and Flegel, T.W. 1993. Histology and ultrastructure reveal a new granulosis-like virus in Penaeus monodon affected by yellow-head disease. Dis. Aquat. Org. 17: 145-157.

Boonyaratpalin, S., Supamattaya, K., Kasornchandra, J., Direkbusaracom, S., Aekpanithanpong, U. and Chantanachookin, C. 1994. Non-occluded baculo-like virus, the causative agent of yellow-head disease in the black tiger shrimp (Penaeus monodon). Gyobyo Kenkyu 28(3): 103-109.

Federici, B.A. 1986. Chapter 3: Ultrastructure of baculoviruses. In: R.R. Granados and B.A. Federici (eds.) The Biology of Baculoviruses, Vol. 1. CRC Press Inc., Boca Raton, Florida. pp. 61-88.

Flegel, T.W., Fegan, D.F., Kongsom, S., Vuthikomudomkit, S., Sriurairatana, S., Boonyaratpalin, S., Chantanachookin, C., Vickers, J. and MacDonald, O.D. 1992. Occurrence, diagnosis and treatment of shrimp diseases in Thailand. In W. Fulks and K. Main (eds.). Diseases of Cultured Penaeid Shrimp in Asia and the United States. The Oceanic Institute, Hawaii. pp. 57-112.

Flegel, T.W., Sriurairtana, S., Wongteerasupaya, C., Boonsaeng, V., Panyim, S. and Withyachumnarnkul, B. 1995a. C.L. Browdy and J.S. Hopkins (eds.). Swimming Through Troubled Water, Proceedings of the Special Session on Shrimp Farming, Aquaculture '95. The World Aquaculture Society, Baton Rouge, LA. pp. 76-83.

Flegel, T.W., Fegan, D.F. and Sriurairatana, S. 1995b. Environmental control of infectious shrimp diseases in Thailand. In: M. Shariff, J.R. Arthur and R.P. Subasinghe, R.P. (eds.) Diseases in Asian Aquaculture II, Fish Health Section, Asian Fisheries Society, Manila. pp. 65-79.

Flegel, T.W., Boonyaratpalin, S. and Withyachumnamkul. 1996. Current status of research on yellow-head virus and white-spot virus in Thailand. World Aquaculture '96 Book of Abstracts. World Aquaculture Society, Baton Rouge, LA. p. 126.

Flegel, T.W., S. Sriurairatana, D.J. Morrison and Napaa Waiyakrutha. 1997a. Penaeus monodon captured broodstock surveyed for yellow-head virus and other pathogens by electron microscopy. In T.W. Flegel, P. Menasveta and S. Paisarnrat (eds). Shrimp Biotechnology in Thailand. National Center for Genetic Engineering and Biotechnology, Thailand, pp. 37-43.

Flegel, T.W., Sitdhi Boonyaratpalin and Boonsirm Withyachumnarnkul. 1997b. Current status of research on yellow-head virus and white-spot virus in Thailand. In T.W. Flegel and I. MacRae (eds.) Diseases in Asian Aquaculture III. Asian Fisheries Soc. In press.

Lightner, 1996 (Ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, Louisiana, USA.

Limsuwan, C. 1991. Handbook for cultivation of black tiger prawns. Tansetakit Co. Ltd, Bangkok.

Lightner, D.V., Redman, R.M., Nunan, L.N., Mohney, L.L., Mari, J.L. and Poulos, B.T. 1997. Occurrence of WSSV, YHV and TSV in Texas shrimp farms in 1995: Possible mechanisms for introduction. World Aquaculture ’97 Book of Abstracts,

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World Aquaculture Society, Baton Rouge, LA. p. 288.Lightner, D.V., Redman, R.M., Poulos, B.T., Nunan, L.M., Mari, J.L. and Hasson,

K.W. 1997. Risk of spread of penaeid shrimp viruses in the Americas by the international movement of live and frozen shrimp. Rev. sci. tech. Off. int. Epiz. 16: 146-160

Limsuwan, C. 1996. Intensive shrimp pond management in Asia. World Aquaculture '96, Book of Abstracts. World Aquaculture Society, Baton Rouge, LA. p. 229.

Lu, Y., Tapay, L.M., Loh, P.C., Brock, J.A. and Gose, R.B. 1994. Distribution of yellow-head virus in selected tissues and organs of penaeid shrimp Penaeus vannamei. Dis. Aquat. Org. 23: 67-70.

Nadala, E.C.B. Jr, Tapay, L.M. and Loh, P.C. 1997. Yellow-head virus: a rhabdovirus-like pathogen of penaeid shrimp. Dis. Aquat. Org. 31: 141-146

Nash, G., Arkarjamon, A. abd Withyachumnarnkul, B. 1995. Histological and rapid haemocytic diagnosis of yellow-head disease in Penaeus monodon. In: M. Shariff, J.R. Arthur and R.P. Subasinghe, R.P. (eds.) Diseases in Asian Aquaculture II, Fish Health Section, Asian Fisheries Society, Manila. pp. 89-98.

Pasharawipas, T., Flegel, T.W., Sriurairatana, S. and Morrison, D.J. 1997. Latent yellow-head infections in Penaeus monodon and implications regarding disease resistance or tolerance. In: T.E. Flegel, P. Menasveta and S. Paisarnrat (eds.) Shrimp Biotechnology in Thailand, National Center for Genetic Engineering and Biotechnology, Bangkok. pp. 45-53.

Spann, K.M., Cowley, J.A., Walker, P.J. and Lester, R.J.G. 1997. A yellow-head-like virus from Penaeus monodon cultured in Australia. Dis. Aquat. Org. 31: 169-179

Wagner, R.R. 1987. Chapter 2: Rhabdovirus biology and infection: an overview. In: Wagner, R.R. (ed.) The Rhabdoviruses. Plenum Press, New York. pp. 9-74.

Wagner, R.R. 1990. Chapter 31: Rhabdoviridae and their replication. In: B.N. Fields, D.M. Knipe, R.B. Chanock, M.S. Hirsch, J.L. Melnick, T.P. Monath and B. Roizman (eds.) Fields Virology, Vol. 1., 2nd Edition, Raven Press, New York. pp. 867-929.

Wongteerasupaya, C. 1996. Viral characterisation and development of specific detection for yellow-head and white-spot diseases in Penaeus monodon. Ph.D. thesis, Mahidol University, Bangkok, Thailand.

Wongteerasupaya, C., Sriurairatana, S., Vickers, J.E., Akrajamorn, A., Boonsaeng, V., Panyim, S., Tassanakajon, A., Withyachumnarnjul, B. and Flegel, T.W. 1995. Yellow-head virus of Penaeus monodon is an RNA virus. Dis. Aquat. Org. 22: 45-50.

Wongteerasupaya, C., Tongchuea, W., Boonsaeng, V., Panyim, S., Withyachumnarnkul, B. and Flegel, T.W. 1996. Polymerase chain reaction detection of yellow head virus in the black tiger prawn, Penaeus monodon. World Aquaculture '96, Book of Abstracts, World Aquaculture Society, Baton Rouge, LA. p. 442.

White Spot Syndrome (WSSV)

Five baculoviruses have been reported to cause white spot syndrome in cultured Penaeus monodon, P. japonicus, P. chinensis, P. indicus, P. merguiensis and P.

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setiferus stocks world-wide. These are: hypodermal and haematopoietic necrosis baculovirus (HHNBV; Huang et al., 1994) in China; rod-shaped nuclear virus of P. japonicus (RV-PJ; Inouye et al., 1994) in Japan, China and Korea; systemic ectodermal and mesodermal baculovirus (SEMBV; Wongteerasupaya et al., 1995) in Thailand; white spot baculovirus (WSBV; Wang et al., 1995) in Indonesia, Vietnam, Malaysia, India, South Carolina and Texas; and Penaeus monodon non-occluded baculovirus (PMNOB; Lo et al., 1995) in Taiwan. SEMBV has recently been identified in cultured P. monodon in Bangladesh (Ahmed, 1996).

All viruses in this group are reported to be very similar in morphology and replicate in the nuclei of infected cells. Lightner et al. (1997a) consider them to be similar, if not the same virus. White Spot Syndrome Virus is not a baculovirus (Volkmann et al., 1995) so it is preferable to refer to it as “White Spot Syndrome Virus” or WSSV (Don Lightner, personal communication).

White spot syndrome was first recognised in 1992-1993 in North East Asia (Takahashi et al., 1994; Chou et al., 1995), and has spread throughout most prawn culture areas of the Indo-Pacific. SEMBV first appeared in Thailand in 1994 where it surpassed yellow-head virus (YHV) as the primary cause of stock losses. In 1995 WSBV was observed in pond-reared P. setiferus in Texas. The virus was apparently introduced with raw and frozen prawns from Thailand which had been processed at nearby plants (Lightner, et al., 1997). Most mortalities occur in young juvenile prawns weighing 3-5 g (Takahashi et al., 1994). WSBV causes mortalities in P. vannamei, P. stylirostris, P. aztecus, P. duorarum and P. setiferus when experimentally infected (Lightner, 1996). The wild penaeids Parapenaeopsis spp., P. semisulcatus, Metapenaeus spp. and Macrobrachium spp. (a caridean not a penaeid) from Taiwan developed disease following experimental infection with WSBV (Chang et al., 1996). Larvae of the freshwater shrimp, Macrobrachium rosenbergii may be infected experimentally and suffer some mortality, however, survivors can carry an infection without mortality as adults. Resistance to WSBV has not been reported for any penaeid species (Lightner, 1996). WSBV has not been reported in Australia.

Clinical signsInfected juvenile and adult prawns become lethargic, cease feeding and have a loose cuticle with white calcium deposits embedded in the cuticle (Takahashi et al., 1994). Infected prawns may display pink to red colouration of the body surface and appendages (Takahashi et al., 1994; Wang et al., 1995). Cumulative mortalities in infected populations may reach 100% within 2 to 10 days of the onset of clinical signs (Chou et al., 1995; Lightner, 1996). Gross PathologyThere is very little gross pathology associated with WSBV. Abnormal deposits of calcium, the accumulation of vacuoles and lysed debris and the necrosis of cuticular pore canals produce white spots, 0.5 to 2.0 mm in diameter, on the cuticular epidermis (Lightner, 1996; Wang et al., 1995). Not all prawns infected with WSBV display white spots on the carapace (Lightner, 1996). Red body discolouration is also common (Inouye et al., 1996). The lymphoid organ of diseased prawns may be swollen or shrunken (Takahashi et al., 1994). Infiltration of haemolymph in the enlarged hemal sinuses and interstitial spaces may cause the hepatopancreas to become swollen, fragile and pale yellow in colour (Wang et al., 1995).

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HistopathologyWSBV infects cells of mesodermal and ectodermal origin, such as the subcuticular epithelium, lymphoid organ, haemocytes, haematopoietic tissue, stomach cuticular epidermis and connective tissue (Momoyama et al., 1995; Lightner, 1996). Infected tissues display widespread focal necrosis (Wongteerasupaya et al., 1995). Degenerate cells are characterised by hypertrophied nuclei with marginated chromatin and eosinophilic to basophilic intranuclear inclusions (IB’s; Chou et al., 1995; Wongteerasupaya et al., 1995). Haemocytic encapsulation of necrotic cells as small brown masses in the stomach may be associated with infection (Momoyama et al., 1995).

The average virion size for baculoviruses from the WSBV complex is 70-150 nm x 250-380 nm (Wongteerasupaya et al., 1995). Replication appears to occur in the nucleus and protective occlusion bodies are not formed.

DiagnosisDiagnosis of white spot syndrome depends mainly on the demonstration of eosinophilic to basophilic IB’s in stained fresh squashes or impression smears of ectodermal and mesodermal tissues. Feulgen-positive intranuclear IB’s may be identified in cuticular epithelial cells and connective tissue cells. A rapid field test for WSBV has been developed. The gills and epithelium under the carapace are excised, stained with haemotoxylin and eosin, mounted and then viewed as squash preparations (Flegel and Sriuriairatana, 1993; K. Supamattaya, pers. comm). WSBV infection may be confirmed by the demonstration of rod-shaped, non-occluded virions in the intranuclear IB’s of affected cells using electron microscopy. The history of disease within the cultured facility, region and species, and the presence of clinical signs are also considered (Lightner, 1996).

Diagnostic DNA probes have been developed and published primers are available for PCR assays from Japan (Kimura et al., 1996) and Taiwan (Lo et al., 1996a and b). Probes have also been developed in Thailand (Wongteerasypaya et al., 1996) and through cooperation between France and the USA (Durand et al., 1996) from prawn tissues infected with SEMBV from Thailand. The Thai probe is being marketed by DiagXotics Co. Ltd and has positively identified WSBV in six penaeids from China, India, Indonesia, Malaysia and Thailand (Wongteerasupata et al., 1996). Diagnostic PCR is used routinely in Thailand to screen postlarvae, broodstock and potential carrier animals (Flegel et al., 1997). PCR primers from Thailand, Taiwan and Japan are being used successfully for the diagnosis of WSBV in penaeids and other crustacea throughout Asia.

Asymptomatic infection of wild-caught Metapenaeus ensis with WSSV was detected using in situ hybridization and PCR (Wang et al., 1997).

Transmission and potential carriersRecent experiments and surveys using diagnostic PCR have shown that approximately forty arthropods, including penaeids, crabs, lobsters, Macrobrachium spp, and possibly copepods and insects can act as carriers (Chou et al., 1996; Lo et al., 1996b; Flegel, 1997; Maeda et al., 1997). Many of these arthropods, such as the wild crab, Portunus pelagicus, and wild krill, Acetes sp., are common in prawn culture areas and

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may transmit the virus to penaeid culture systems with the in-take water (Supamattaya et al., 1996). Within the culture system WSBV is transmitted by cannibalisation of moribund prawns and carcasses or via contaminated water (Chang et al., 1996). Crustacean carriers which enter prawn ponds may transmit WSBV when they die and are eaten be prawns. Birds may mechanically transmit the virus between ponds by releasing captured prawns over neighbouring ponds. Unpublished work done by Charoen Pokphand (CP) and Aquastar Co. Ltd. in Thailand (Tim Flegel, personal communication) showed that there was a clear correlation between some postlarval (PL) batches used to stock ponds and subsequent WSBV outbreaks. This has led to the general practice of testing PL batches for WSBV by PCR assay before stocking. It suggests that unrestricted transportation of live PL from areas known to be infected by WSBV to uninfected areas would be very hazardous.

Evidence suggests that outbreaks of WSBV in the USA were probably due to introduction of the virus in frozen prawns from Asia (Lightner et al., 1997).Kou et al. (1997) detected WSBV by in situ hybridization in oogonia and follicle cells in P. monodon ovarian tissues. Mohan et al. (1997) observed intranuclear viral inclusions in the gonads of P. monodon and concluded that WSBV could be transmitted vertically. However Lo et al. (1997) in their studies of WSSV tissue tropism were unable to find any infected mature eggs and suggested that infected egg cells were killed by the virus before maturation.

ViabilityExperiments indicate that WSBV can remain viable in seawater for 4 to 7 days, although data is not yet available (Flegel, 1997). No published data is available on the effect of heat and desiccation on the viability of WSBV. However, unpublished work by CP and Aquastar Co. Ltd. in Thailand showed that there was no correlation between feed batches and pond outbreaks of WSBV. The viability of WSBV in frozen prawns is not known.

Chou et al. (1995) observed 100% mortality in prawns fed frozen tissues infected with WSBV.

WSSV (PRDV in the paper) is inactivated after 50 min at 50 C, but after only 1 min at 60 C. The virus is sensitive to low levels of exposure to UV and is inactivated by desiccation (to 3.7% water remaining) after only 3 hours (Nakano et al., 1998).

Prevention As for YHV, SEMBV is being controlled in Thailand by the use of closed and semi-closed systems (Limsuwan, 1996) involving the pre-treatment of water with formalin or chlorine and storage of any water to be exchanged. Unpublished aquarium trials from CP (Boonsirm Withayachumnarnkul, personal communication) indicated that 70 ppm formalin treatment at 6 hour intervals could prevent the transmission of WSBV from infected to non-infected shrimp. By contrast, similar unpublished work from Aquastar Co. Ltd. (Vithaya Thammavit, personal communication) and field experience (Chalore Limsuwan, personal communication) indicate that formalin administered at 20 to 40 ppm at 5 to 7 day intervals is sufficient to prevent the spread of infection. It is likely that effectiveness of this treatment would depend upon the quantity of virus present (ie, concentration in the water, number of infected shrimp and severity of infection). The elimination of fresh feed from the diet, the exclusion

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of potential carriers from prawn culture ponds and PCR screening of postlarvae prior to stocking are also recommended as control measures (Flegel et al., 1996; Boonsirm Withayachumnarnkul, personal communication).

The present status of white spot syndromeSEMBV infection in P. monodon from Thailand alone, resulted in a US$ 600 million loss in 1996 (Dr. Lin, CP, personal communication). The epidemic of WSBV in Thailand, China and India appears to be abating due to the widespread use of the recommended preventative measures and better farming practices. WSBV continues to cause massive stock losses in other affected prawn culture countries. ReferencesAhmed, A.T.A. 1996. Disease problems of cultured tiger shrimp (Penaeus monodon)

in Bangladesh. 2nd Int. Conference on the Culture of Penaeid Prawns and Shrimps, book of abstracts, SEAFDEC/AQD, Iloilo City, Philippines. p. 111

Chang, P.S., Wang, Y.C., Lo, C.F. and Kou, G.H. 1996. Infection of white spot syndrome associated with non-occluded baculovirus in cultured and wild shrimps in Taiwan. 2nd Int. Conference on the Culture of Penaeid Prawns and Shrimps, book of abstracts, SEAFDEC/AQD, Iloilo City, Philippines. p. 90.

Chou, H.Y., Huang, C.Y., Wang, C.H., Chiang, H.C. and Lo, C.F. 1995. Pathogenicity of a baculovirus infection causing white spot syndrome in cultured penaeid shrimp in Taiwan. Dis. Aquat. Org. 23: 165-173.

Chou, H.Y., Huang, C., Kou, G.H. and Durand, S., Lightner, D.V., Nunan, L.M., Redman, R.M., Mari, J. and Bonami, J-R. 1996. Application of gene probes as diagnostic tools for white spot baculovirus (WSBV) of penaeid shrimp. Dis. Aquat. Org. 27: 59-66.

Flegel, T.W. 1997. Major viral diseases of the black tiger prawn (Penaeus monodon) in Thailand. World. J. Microbiol. Biotech., in press.

Flegel, T.W. and Sriurairatana, S., 1993. Black tiger prawn diseases in Thailand. In. D.M. Akiyama (ed.) Technical Bulletin AQ39 1993/3, American Soybean Association, Singapore. p. 16

Flegel, T.W., Sitdhi Boonyaratpalin and Boonsirm Withyachumnarnkul. 1997. Current status of research on yellow-head virus and white-spot virus in Thailand. In T.W. Flegel and I. MacRae (eds.) Diseases in Asian Aquaculture III. Asian Fisheries Soc. In press.

Kou, G.H., Chen,C.H., Ho, C.H. and Lo, C.F. 1997 White spot sydrome virus (WSSV) in wild-caught black tiger shrimp: WSSV tissue tropism with a special emphasis on reproductive organs World Aquaculture ’97 Book of Abstracts, World Aquaculture Society, Baton Rouge, LA. p. 262

Huang, J. Song, X-L., Yu, J. and Yang, C.H. 1994. Baculoviral hypodermal and hematopoietic necrosis - pathology of the shrimp explosive epidemic disease. Abstract, Yellow Sea Fishery Research Institute, Qingdao, P.R. China. Cited by Lightner, 1996.

Inouye, K., Miwa, S., Oseko, N. Nakano, H. Kimura, T. Momoyama, K. and Hiraoka, M. 1994. Mass mortalities of cultured Kuruma shrimp Penaeus japonicus in Japan in 1993: electron microscope evidence of the causative virus. Fish Pathol. 29: 149-158.

Kimura, T., Yamano, K., Nakano, H., Momoyama, K., Hiraoka, M. and Inouye, K. 1996. Detection of penaeid rod-shaped DNA virus (PRDV) by PCR. Fish Pathol. 31(2): 93-98.

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Lan, J., Pratanpipat, P., Nash, G., Wongwisansri, S., Wongteerasupaya, C., Withyachumnamkul, B., Thammasert, S. and Lohawattanakul, C. 1996. Carrier and susceptible host of the systemic ectodermal and mesodermal baculovirus, the causative agent of white spot disease in penaeid shrimp. World Aquaculture ’96, Book of Abstracts. World Aquaculture Society, Baton Rouge, LA. p.213.

Lightner, 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, Louisiana, USA.

Lightner, D.V., Redman, R.M., Poulos, B.T., Nunan, L.M., Mari, J.L .and Hasson, K.W. 1997 Risk of spread of penaeid shrimp viruses in the Americas by the international movement of live and frozen shrimp. Rev. sci. tech. Off. int. Epiz. 16: 146-160

Lightner, D.V., Redman, R.M., Nunan, L.N., Mohney, L.L., Mari, J.L. and Poulos, B.T. 1997. Occurrence of WSSV, YHV and TSV in Texas shrimp farms in 1995: Possible mechanisms for introduction. World Aquaculture ’97 Book of Abstracts, World Aquaculture Society, Baton Rouge, LA. p. 288.

Limsuwan, C. 1996. Intensive shrimp pond management in Asia. World Aquaculture '96, Book of Abstracts. World Aquaculture Society, Baton Rouge, LA. p. 229

Lo, C.F., Wang, C.H. and Kou, G.H. 1995. Purification and genomic analysis of white spot syndrome associated non-occluded baculovirus (PmNOB) isolated from Penaeus monodon. Abstract, European Association of Fish Pathologists, 7th International Conference on Diseases of Fish and Shellfish, Spain. p. 77.

Lo, C.F. 1996. Studies on the transmission of white spot syndrome-associated baculovirus (WSBV) in Penaeus monodon and P. japonicus via waterborne contact and oral ingestion. 2nd Int. Conference on the Culture of Penaeid Prawns and Shrimps, book of abstracts, SEAFDEC/AQD, Iloilo City, Philippines. p. 55

Lo, C.F., Leu, J.H., Ho, C.H., Chen, C.H., Peng, S.E., Chen, Y.T., Chou, C.M., Yeh, P.Y., Huang, C.J., Chou, H.Y., Wang, C.H. and Kou, G.H. 1996a. Detection of baculovirus associated with white spot syndrome (WSBV) in penaeid shrimps using polymerase chain reaction. Dis. Aquat. Org. 25: 133-141.

Lo, C.F., Ho, C.H., Peng, S.E., Chen, C.H., Hsu, H.C., Chiu, Y.L., Chang, C.F., Liu, K.F., Su, M.S., Wang, C.H. and Kou, G.H. 1996b. White spot syndrome baculovirus detected in cultured and captured shrimp, crabs and other arthropods. Dis. Aquat. Org. 27: 215-225.

Lo, C.F., Ho, C.H., Chen, C.H., Liu, K.F., Chiu, Y.L., Yeh, P.Y., Peng, S.E., Hsu, H.C., Liu, H.C., Chang, C.F., Su, M.S., Wang,C.H. and Kou, G.H. (1997) Detection and tissue tropism of white spot syndrome baculovirus (WSBV) in captured brooders of Penaeus monodon with a special emphasis on reproductive organs. Dis. Aquat. Org. 30: 53-72.

Maeda, M., Itami, T., Kondo, M., Hennig, O., Takahashi, Y., Hirono, I. And Aoki, T. 1997. Characteristics of penaeid rod-shaped DNA virus of Kuruma shrimp. New Approaches to Viral Diseases of Aquatic Animals. Proceedings of the National Research Institute of Aquaculture International Workshop, Japan. pp. 218-228.

Mohan, C.V., Sudha, P.M., Shankar, K.M and Hegde, A. 1997. Vertical transmission of white spot baculovirus in shrimps - a possibility? Current Science (Bangalore) 73: 109-110

Momoyama, K., Hiraoka, M., Inouye, K., Kimura, T. and Nakano, H. 1995. Diagnostic techniques of the rod-shaped nuclear virus infection in the kuruma shrimp, Penaeus japonicus. Fish. Pathol. 30(4): 263-269.

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Nakano, H., Hiraoka, M., Sameshima, M., Kimura, T. and Momoyama, K. 1998 Inactivation of penaeid rod-shaped DNA virus (PRDV), the causative agent of penaeid acute viremia (PAV), by some chemical and physical treatments. Fish Pathol.33: 65-71

Supamattaya, K., Hoffman. R.W. and Boonyaratpalin, S. 1996. Transmission of red and white spot disease (bacilliform virus) from black tiger shrimp Penaeus monodon to portunid crab Portunus pelagicus and krill Acetes sp. 2nd Int. Conference on the Culture of Penaeid Prawns and Shrimps, book of abstracts, SEAFDEC/AQD, Iloilo City, Philippines. p. 97.

Takahashi, Y. Itami, T., Kondo, M., Maeda, M., Fujii, R., Tomonaga, S., Supamattaya, K. and Boonyaratpalin, S. 1994. Electron microscope evidence of a bacilliform virus infection in Kuruma shrimp (Penaeus japonicus). Fish Pathol. 29: 121-125.

Volkman, L.E., Blissard, G.W., Friesen, P., Keddie, B.A., Possee, R., Theilmann, D.A. 1995. Baculoviridae. In: Murphy, F.A., Fauquet, C.M., Bishop, D.H.L., Ghabrial, S.A., Jarvis, A.W., Martelli, G.P., Mayo, M.A. and Summers, M.D. (eds.) Virus Taxonomy, Sixth Report of the International Committee on Taxonomy of Viruses., Springer-Verlag, Wein, New York. pp. 104-113.

Wang, C.H., Lo, C.F., Leu, J.H., Chou, C.M., Yeh, P.Y., Chuo, H.Y., Tung, M.C., Chang, C.F., Su, M.S. and Kou, G.H. 1995. Purification and genomic analysis of baculovirus associated with white spot syndrome (WSBV) of Penaeus monodon. Dis. Aquat. Org. 23: 239-242.

Wang, C.S., Tsai, Y.J,. Kou, G.H. and Chen, S.N. 1997. Detection of white spot disease virus infection in wild-caught greasy back shrimp, Metapenaeus ensis (de Haan) in Taiwan. Fish Pathol. 32: 35-41

Wongteerasupaya, C. Vickers, J.E., Sriurairatana, S., Nash, G.L., Akarajamorn, A., Boonsaeng, V., Panyim, S., Tassanakajon, A., Withyachumnarnkul, B. and Flegel, T.W. 1995. A non-occluded, systemic baculovirus that occurs in the cells of ectodermal and mesodermal origin and causes high mortality in the black tiger prawn, Penaeus monodon. Dis. Aquat. Org. 21: 69-77.

Wongteerasupaya, C., Wongwisansri, S., Boonsaeng, V., Panyim, S., Withyachumnarnkul, B. and Flegel, T.W. 1996. Sensitive and rapid detection of systemic ectodermal and mesodermal baculovirus by DNA amplification. World Aquaculture ’96, Book of Abstracts. World Aquaculture Society, Baton Rouge, LA. p. 443.

Taura Syndrome Virus (TSV)

Natural infections of TSV have been documented in cultured Penaeus vannamei, P. stylirostris, P. aztecus and P. setiferus in the Americas (Lightner et al., 1997). Taura syndrome (TS) was first recognised in Ecuador in 1991 and was believed to be caused by pesticide run-off from nearby banana farms. It is now known that TS is caused by a virus, TSV (Lightner et al., 1995b). Mortalities have been reported in cultured penaeids from Ecuador, Peru, Colombia, El Salvador, Guatemala, Brazil, Nicaragua, Hawaii, Florida and Mexico (Lightner et al., 1994; Lightner, 1996). More recently TSV has been introduced into Texas (Johnson, 1995) with wild-caught P. vannamei postlarvae and broodstock. TS has not yet been reported in Panama or South

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Carolina (Brock et al., 1996). TS has been reported in South Carolina (Brock 1997), Costa Rica and Panama (Don Lightner, personal communication). Experimental infection have been produced in P. aztecus and P. chinensis. P. stylirostris and P. aztecus appear resistant to TS disease, although they are susceptible to TSV. P. dourarum may also be resistant to TS. A report of TSV from New Caledonia has not been substantiated (Don Lightner, personal communication). YHV was also suspected as being the cause of Syndrome 93. TSV and YHV were considered by pathologists working on Syndrome 93 because affected shrimp showed a diffuse necrosis of connective tissues, hemocytes and the lymphoid organ that mimicked that presented by shrimp in the acute phase of infections due to YHV, and to a lesser extent, TSV. Neither was shown to be involved, and the disease has been induced experimentally (filling the requirements of Koch's postulates) with certain strains of Vibrio penaeicida (Don Lightner, personal communication).

Experimental exposure to TSV by ingestion produced infection in P. schmitti and P. dourarum and mortalities in P. setiferus (Overstreet et al., 1997). Therefore these prawn species can serve as carriers of TSV without necessarily exhibiting disease. These studies also showed that P. chinensis developed disease after injection of TSV. P. vannamei, P. schmitti, P. setiferus and P. stylirostris show clinical signs of disease when exposed to TSV, while P. monodon, P. japonicus, P. duorarum, and P. aztecus are disease-resistant (Brock, 1997).

Brock (1997) states that TSV has no known impact on nauplii through mysis stage, but P. vannamei from post-larval to the adult stage are highly susceptible. Reports are lacking which demonstrate that TSV is infectious to other groups of decapods or crustaceans.

Clinical signs and gross pathologyTS primarily affects prawns in the nursery phase when they are 0.1 to 5 g (Lightner et al., 1994). During the preacute/acute phase of infection, prawns appear pale red while their tail fans become bright red (TS is also referred to a red tail disease). They are soft shelled, lethargic and anorexic. Those with severe infections die during moult (Chamberlain, 1994) and cumulative mortalities may reach 80-95%. Recovering, chronically infected prawns generally display multifocal, melanised cuticular lesions and may also have soft cuticles and red body colouration. They may behave and feed normally. Survivors of TS epizootics may have survival rates of 60% to harvest size (Lightner, 1996).

HistopathologyTS lesions appear as multifocal areas of necrosis in the cuticular epithelium of the body surface, appendages, gills, hindgut, stomach and oesophagus ( Lightner et al., 1995). Feulgen-negative IB’s, which may first appear eosinophilic than change to basophilic, may be present in the cytoplasm of cells in areas of necrosis. When abundant, these IB’s give TS lesions a characteristic peppered appearance.

TSV particles are cytoplasmic, icosohedral and 30-32 nm in diameter (Hasson et al., 1995). TSV has been tentatively classified as a picornavirus based on its morphology, site of replication, ssRNA genome of 9 kb and polypeptide capsid structure (Brock et al., 1995; Hasson et al., 1995).

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DiagnosisDiagnosis of TSV infection is based on clinical signs, history of infection within the culture facility, region and species and the demonstration of multifocal lesion in the cuticular and subcuticular tissues (Brock et al., 1995; Lightner, 1996). Recovering prawns may shed characteristic lesions and appear healthy while carrying the virus (Brock et al., 1996). TSV infection may be confirmed by the identification of icosohedral viral particles in areas of necrosis or by bioassay of suspected infected prawns with Specific Pathogen Free (SPF) juvenile P. vannamei as the indicator host (Brock et al., 1995). A diagnostic DNA probe for TSV has recently been developed (Brock 1997) and marketed.

RT-PCR, in situ hybridization and immunoblot ELISA techniques have been developed and successfully used to detect TSV in infected tissues (Poulos et al., 1998). The lymphoid organ is the tissue of choice for detection of chronically infected TSV survivors (Ken Hasson, personal communication).

Transmission and potential carriersIt is not certain how many penaeid species besides P. stylirostris and P. aztecus are potential carriers of TSV (Lightner et al.,1997). Within ponds TSV is transmitted ‘per os’. TSV is not transmitted vertically from broodstock to offspring (Lotz and Ogle, 1997). Numerous crustaceans native to Texas have been tested for susceptibility to TSV. The results of these experiments indicate that there are few potential carriers of TSV (Erickson et al., 1997). Migratory birds, aquatic insects and humans have been implicated as mechanical vectors of TSV between ponds (Johnson, 1995).

Garza et al. (1997) suggest that sea gulls are probable transport vectors of TSV within and between nearby prawn farms.

A probable means of dissemination of TSV is with shipments of post-larvae. Movement of TSV with infected broodstock of P. vannamei has been reported (Brock et al.,1997 as cited in Brock, 1997). Transfer of TSV between geographic locations in frozen shrimp products should not be overlooked as a means of dissemination of this virus (Lightner, 1995 as cited in Brock, 1997).

P. schmitti and P. dourarum can be TSV carriers (Overstreet et al., 1997).TSV has been detected by bioassay in clinically healthy survivors following a disease outbreak (Lotz & Ogle, 1997). Clinical signs of TS were observed in P. vannamei after they were fed experimentally infected P. setiferus (Erickson et al., 1997). Saline extracts of TSV infected tissues diluted 10-4 retained pathogenicity on experimental inoculation of P. vannamei (Brock et al., 1995).

ViabilityTSV in prawn tissue remains active after freezing and storage at 0oC (Brock et al., 1995). Experimental transmission trials, using autoclaved TS-infected tissues, indicates that TSV is inactivated at 121oC (Brock et al., 1995).

Hasson et al. (1995) suggested that one of the possible reasons for the rapid dissemination of TSV is the highly stable nature of the virus. This is endorsed by the recovery of viable, infectious virions from dead shrimp showing advanced post

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mortem changes and the apparent capacity of the virus to endure long term freezing and multiple freeze-thaw cycles.

PreventionEffective control measures for TS have not yet been established. Many farmers have switched to SPF P. stylirostris, which is less susceptible to TSV than P. vannamei (Chamberlain, 1994; Brock et al., 1995). Reducing stress by lowering stocking densities is recommended (Brock et al., 1995). Standard disinfection methods are not effective, although treating drained ponds with lime has reduced mortalities in some instances (Brock et al., 1996).

Comprehensive management practices have been reported to have eliminated TSV following disease outbreaks on prawn farms in Belize (Dixon, 1997).

Present status of Taura SyndromeSince 1991, TS has cost the prawn culture industries of the USA and Latin America over US$1 billion (Brock et al., 1996). Although production improved during 1995 on some farms, TS continues to be a problem throughout the Americas. The impact of TS on wild fishery stocks has not been documented (Brock et al., 1996).

ReferencesBrock, J.A., Gose, R. Lightner, D.V. and Hasson, K.W. 1995. An overview on Taura

syndrome, an important disease of farmed Penaeus vannamei. In: C.L. Browdy and J.S. Hopkins (eds.). Swimming through Troubled Waters, Proceeding of the special session on shrimp farming, Aquaculture ’95. World Aquaculture Society, Baton Rouge, LA. pp. 84-89.

Brock, J.A. , Lightner, D.V., Hasson, K. and Gose, R. 1996. An update on Taura syndrome of farmed shrimp in the Americas. World Aquaculture ’96, Book of Abstracts, World Aquaculture Society, Baton Rouge, LA. p. 50.

Brock, J.A. 1997. Special topic review: Taura syndrome, a disease important to shrimp farms in the Americas. World J. Microbiol & Biotechnol. 13:415-418

Chamberlain, G.W. 1994. Taura syndrome and China collapse caused by new shrimp viruses. World Aquaculture 25(3): 22-25.

Dixon, H., Dorado, J. and Hyde, C. 1997. Managing Taura syndrome virus in Penaeus vannamei production ponds in Belize, Central America: a case study. World Aquaculture ’97, book of abstracts. The World Aquaculture Society, Baton Rouge, LA. p. 139.

Erickson, H.S., Lawrence, A.L., Gregg, K.L., Lotz, J and McKee, D.V. 1997. Sensitivity of Penaeus vannamei, P.vannamei TSV survivors and Penaeus setiferus to Taura syndrome virus infected tissue and TSV infected pond water; and, sensitivity of P. vannamei to TSV bioassays with P. setiferus and Penaeus aztecus. World Aquaculture ’97, book of abstracts. The World Aquaculture Society, Baton Rouge, LA. p. 139.

Erickson, H.S., Lawrence, A.L., Gregg, K.L., Frelier, P., Lotz, J and McKee, D.V. 1997. Sensitivity of Penaeus vannamei, Sciaenops ocellatus, Cynoscion nebulosus, Palaemonetes sp. and Callinectes sapidus to Taura syndrome virus infected tissues. World Aquaculture ’97, book of abstracts. The World Aquaculture Society, Baton Rouge, LA. p. 140.

Garza, J.R., Hasson, K.W., Poulos, B.T., Redman, R.M., White, B.L., Lightner, D.V. 1997. Demonstration of infectious Taura syndrome virus in the feces of seagulls

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collected during an epizootic in Texas. J. Aquatic Anim. Health 9: 156-159Hasson, K.W., Lightner, D.V., Poulos, B.T., Redman, R.M., White, B.L., Brock, J.A.

and Bonami, J.R. 1995. Taura syndrome in Penaeus vannamei: demonstration of a viral etiology. Dis. Aquat. Org. 23: 115-126.

Johnson, S.K. Taura virus hits Texas. World Aquaculture 26(3): 82-83.Lightner, D.V., Jones, L.S. and Ware, G.W. 1994. Proceedings of the Taura

syndrome workshop executive summary. University of Arizona, Tucson, USA.Lightner, D.V., Redman, R.M., Hasson, K.W. and Pantoja, C.R. 1995. Taura

syndrome in Penaeus vannamei (Crustacea: Decapoda): gross signs, histopathology and ultrastructure. Dis. Aquat. Org. 21: 53-59.

Lightner, D.V., Redman, R.M., Poules, B.T., Nunan, L.M., Mari, J.L., Masson, K.W. and Bonami, J.R. 1997. Taura Syndrome: etiology, pathology, hosts and geographic distribution, and detection methods. In: New Approaches to Viral Diseases of Aquatic Animals. National Research Institute of Aquaculture, Japan. pp. 190-225.

Lightner, 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA, USA.

Lotz, J.M. and Ogle, J.T. 1997. Taura syndrome and reproduction of Penaeus vannamei. World Aquaculture ’97 Book of Abstracts, World Aquaculture Society, Baton Rouge, LA. p. 294.

Overstreet, R.M., Lightner, D.V., Hasson, K.W., McIlwain, S. and Lotz, J.M. 1997. Susceptibility to Taura syndrome virus of some penaeid shrimp species native to the Gulf of Mexico and the Southeastern United States. J. Invert.Pathol. 69: 165-176

Poulos, B.T., Nunan,L.M., Mohney, L.L. and Lightner, D.V. 1998. Detection of Taura syndrome virus in penaeid shrimp: comparison of testing methods employing gene probes, monoclonal antibodies and PCR. Abstract, The Triennial Meeting of World Aquaculture Society, Las Vegas Feb 15-19, 1998.

Infectious Hypodermal and Hematopoietic Necrosis Virus (IHHNV)

Infectious hypodermal and haematopoietic necrosis virus (IHHNV) is distributed widely in penaeid culture facilities in Asia and the Americas. Countries which have reported epidemics of IHHNV include: south-east USA, Mexico, Ecuador, Peru, Brazil, Carribean, Central America, Hawaii, Guam, Tahiti, New Caledonia, Singapore, Malaysia, Thailand, Indonesia and the Philippines (Lightner, 1996). Natural infections have been reported from Penaeus stylirostris, P. vannamei, P. occidentalis, P. californiensis, P. monodon, P. semisulcatus and P. japonicus. P. setiferus, P. dourarum and P. aztecus have been infected experimentally with IHHNV and P. indicus and P. merguiensis appear to be refractory to infection (Brock and Lightner, 1990; Lightner, 1996). IHHNV is believed to be enzootic in wild reservoir hosts such as P. monodon (Brock and Lightner, 1990). An IHHNV-like virus has been reported from a hybrid penaeid, P. monodon x P. esculentus, bred in Australia (Owens et al., 1992). It is not known if distinct geographic strains of IHHNV exist.

IHHNV is widely distributed in culture facilities in the Americas and Asia including southeast U.S., Mexico, Central America, Ecuador, Peru, Brazil, and numerous

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Caribbean Islands, Hawaii, Guam, Tahiti, New Caledonia, Singapore, Philippines, Thailand, Malaysia, Indonesia (Lightner, 1996) and China (Zhang & Sun, 1997).

Species like P. indicus and P. merguiensis may be infected with the virus but do not show signs of the disease; they appear to be refractory to IHHNV (Lightner, 1996). IHHNV infection has been reported in P. chinensis (Zhang & Sun, 1997).

Owens et al. (1992) reported that IHHNV had been found in Australia and that samples were sent to the USA to be tested with a monoclonal antibody to IHHNV in an ELISA. The tissue gave values of 38 to 78% intensity when compared to the known positive control (Don Lightner, personal communication). The American ELISA referred to was apparently still in the developmental stage when the Australian tissue was tested (Poulos et al., 1994). Subsequent work with a commercial IHHNV probe suggested limited genetic similarity (Owens, 1997). The term "IHHNV-like" is moreappropriate for describing the Australian agent. (Don Lightner, personal communication).

Bonami et al. (1990) describe the IHHNV ssDNA genome and the basis for classification of the virus as a probable parvovirus.

Clinical signsIHHNV disease has been studied closely in P. vannamei and P. stylirostris in the Americas (Lightner et al., 1983; Bell and Lightner, 1984). The clinical signs of IHHNV disease in P. stylirostris are nonspecific and include anorexia, lethargy and erratic swimming. Early larvae and postlarvae, which have been vertically infected do not become diseased until they are older and within the size range 0.05 to 1 g (Lightner et al., 1983). Infected prawns have been observed to rise to the water surface, remain motionless for a few moments then roll over and sink to the bottom. This behaviour may be repeated until mortality occurs. Mortality may exceed 90% within several weeks of onset of infection in juvenile P. stylirostris (Bell and Lightner, 1987). In P. vannamei IHHNV is typically a chronic disease linked to runt deformity syndrome and infected populations of juvenile shrimp typically display a wide distribution of sizes (Kalagayan et al., 1991). For the hybrid prawns bred in Australia, mortality from IHHNV infection occurred when the prawns reached 3 to 4 g (Owens et al., 1992).

P. monodon may appear clinically normal when heavily infected by IHHNV (Flegel, 1997).Gross PathologyGross signs of infection include white to buff mottling of the cuticle, opacity of striated muscle and melanised foci within the hypodermis (Bell and Lightner, 1987). In the later stages of infection P. stylirostris and P. monodon may appear bluish in colour. Infected P. vannamei display deformed rostrums, cuticle and antennal flagella (Lightner et al., 1983; Lightner, 1996). Australian hybrids infected with IHHNV became weak and lethargic and there was no noticeable change in colouration (Owens, et al., 1992).

Histopathology IHHNV is an unenveloped, icosahedral virus, 17-27 nm in diameter (Lightner et al., 1983b), which replicates in the cytoplasm of cells of ectodermal origin (epidermis,

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gills, fore and hind gut, antennal gland and neurons) and mesodermal origin (haematopoietic tissue, haemocytes, striated muscle, heart, lymphoid organ and connective tissues). Infection of the midgut epithelium is rare (Lightner et al., 1983b). IHHNV forms Cowdry Type A intranuclear inclusion bodies (IB’s) associated with widespread cytopathological changes including hypertrophy of the nucleus and margination of the chromatin (Lightner et al., 1983b). IHHNV has been tentatively assigned to the Parvoviridae (Lightner, 1996). Cowdry Type A IB’s were observed in cells of ectodermal and mesodermal origin in Australian hybrid prawns. The hearts of some prawns investigated had focal haemocyte infiltrations and in some prawns melanised nodules were observed in the connective tissues (Owens et al., 1992).

DiagnosisIHHNV may be diagnosed by the demonstration of Cowdry Type A IB’s using routine histochemical techniques for light microscopy and electron microscopy. Bioassays may be used to detect asymptomatic carriers of the virus, using specific pathogen free P. stylirostris as the indicator host. IHHNV-specific gene probes have been developed from naturally infected P. stylirostris juveniles to use for in situ and dot blot hybridisation (Lightner et al., 1992; Mari et al., 1993). These probes are commercially available and severe to low grade infections may be detected. Non-lethal screening of broodstock may be carried out by removing an appendage, such as a pleopod or gill process, or sample of haemolymph and processing it for routine histology or to test with the probe by in situ hybridisation (Bell et al., 1990). Polymerase Chain Reaction (PCR) primers have also been developed which allow IHHNV to be detected in fresh, frozen or fixed samples of tissue or haemolymph (Lightner, 1996). Murine monoclonal antibodies to IHHNV have been developed for an ELISA detection system (Poulos et al., 1994). However, further work is required before this system can be used for reliable routine diagnosis.

The Australian strain of IHHNV gave negative results when tested with a probe for IHHNV developed in the USA (Leigh Owens, personal communication) and it is diagnosed using routine histochemical techniques.

TransmissionIt is believed that IHHNV may be transmitted vertically from broodstock to their progeny (Lightner et al., 1983). However, this has not been proven. IHHNV-resistant penaeid species and early life stages carry the virus latently and transfer it to more susceptible species and life stages. The virus is transmitted either via the water or is ingested with infected prawns (Bell and Lightner, 1984).

ViabilityIHHNV in prawn tissues will survive storage at -5oC to -10oC (Bell and Lightner, 1984). The survival of IHHNV after exposure to high temperatures is not known.

PreventionEffective control measures for IHHNV disease are not known. Avoidance of the virus through quarantine is strongly recommended (Brock and Lightner, 1990). The disease impact of IHHNV may be reduced by improving farm management practices, such as lowering stocking densities, using nutritionally balanced feeds and stocking ponds with more resistant prawn species.

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Present status of IHHNV Disease caused by IHHNV infection continues to be a chronic problem of cultured prawns in a number of countries. However, reports of serious epidemics have been rare in the last couple of years. IHHNV has occurred in multiple infections with other, more pathogenic viruses and is considered, in most cases, to be a chronic infection which represses the prawns’ defence system, allowing infection by other disease-causing agents. IHHNV has not been recorded in Australia since the death of the diseased hybrid prawns originally described.

ReferencesBell, T.A. and Lightner, D.V. 1984. IHHN virus: infectivity and pathogenicity studies

in Penaeus stylirostris and Penaeus vannamei. Aquaculture 38: 185-194.Bell, T.A. and Lightner, D.V. 1987. IHHN disease of Penaeus stylirostris: effects of

shrimp size on disease expression. J. Fish. Dis. 10: 165-170.Bell, T.A., Lightner, D.V. and Brock, J.A. 1990. A biopsy procedure for the non-

destructive determination of IHHN virus infection in Penaeus vannamei. J. Aquat. Anim. Health 2: 151-153.

Bonami,J.R., Trumper, B., Mari, J., Brehelin, M. and Lightner, D.V. 1990 Purification and characterisation of the infectious hypodermal and haematopoeitic necrosis virus of penaeid shrimps J. Gen. Virology 71, 2657-2664

Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne (ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Flegel, T.W. 1997. Special topic review: Major viral diseases of the black tiger prawn (Peneaus monodon) in Thailand. World Journal of Microbiology & Biotechnology 13: 433-442

Kalagayan, G. Godin, D., Kanna, R., Hagino, G., Sweeney, J., Wyban, J. and Brock, J. 1991. IHHN virus as an etiological factor in runt deformity syndrome of juvenile Penaeus vannamei cultured in Hawaii. J. World Aquaculture Soc. 22: 235-243.

Lightner, D.V., Redman, R.M. and Bell, T.A. 1983a. Infectious hypodermal and hematopoietic necrosis a newly recognised virus disease in penaeid shrimp. J. Invertebr. Pathol. 42: 62-70.

Lightner, D.V., Redman, R.M., Bell, T.A. and Brock, J.A. 1983b. Detection of IHHN virus in Penaeus stylirostris and Penaeus vannamei imported into Hawaii. J. World Maricult. Soc. 14: 212-225.

Lightner, D.V., Poulos, B.T., Bruce, L. Redman, R.M., Mari, J. and Bonami, J.R. 1992. New developments in penaeid virology: application of biotechnology in research and disease diagnosis for shrimp viruses of concern in the Americas. In: W. Fulks and K. Main (eds.) Diseases of Cultured Penaeid Shrimp in Asia and the United States. The Oceanic Institute, Makapuu Point, Honolulu. pp. 233-253.

Lightner, 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA.,USA.

Mari, J., Bonami, J.R. and Lightner, D.V. 1993. Partial cloning of the genome of infectious hypodermal and hematopoietic necrosis virus, an unusual parvovirus pathogenic for penaeid shrimps; diagnosis of the disease using a specific probe. J. Gen. Virol. 74: 2637-2643

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Owens, L., Anderson, I.G., Kenway, M., Trott, L. and Benzie, J.A.H. 1992. Infectious hypodermal and hematopoietic necrosis virus (IHHNV) in a hybrid penaeid prawn from tropical Australia. Dis. Aquat. Org. 14: 219-228.

Owens, L. 1997. Special topic review: the history of the emergence of viruses in Australian prawn aquaculture. World J. Micro. & Biotechnol. 13: 427-431

Poulos, B.T., Lightner, D.V. and Trumper, B. 1994. Monoclonal antibodies to a penaeid shrimp parvovirus, infectious hypodermal and hematopoietic necrosis virus (IHHNV). J. Aquat. Anim. Health. 6: 149-154.

Zhang, J. and Sun, X. 1997 A study on pathogens of Chinese prawn (Penaeus chinensis) virus diseases in The fourth Asian Fisheries Forum ed. Zhou Yingqi, Zhou Hongqi, Yao Chaoqi, Lu Yi, Hu Fuyuan, Cui He and Din Fuhui Ocean Press, Bejing, China

Baculovirus penaei (PvSNPV)

Baculovirus penaei (BP) has recently been designated “Penaeus vannamei Singular Nucleopolyhedrosis Virus” (PvSNPV; Bonami et al., 1995) and often appears under this name in recent literature. However, until PvSNPV is accepted by the International Committee on the Taxonomy of Viruses (ICTV), we will use the term BP.

Several strains of BP are likely to exist (Lightner, 1996). Genomic probes have been developed and used to detect BP strains (Durand et al., 1998). Overstreet (1994) states that BP probably constitutes at least 3 different strains. Virus from natural infections of P. marginatus in Hawaii differs significantly in virulence from two closely related, but different, strains of BP from the Gulf of Mexico and from the Pacific coast of Latin America. The basis for differences in the virulence of strains is not well understood.

BP was first discovered in captured P. duorarum in the USA (Couch, 1974a) and has since been described from cultured and captured P. vannamei, P. stylirostris, P. setiferus, P. schmitti, P. penicillatus, P. brasiliensis, P. paulensis, P. subtilis , P. aztecus and P. marginatus from the Americas (Lightner et al., 1989; Brock and Lightner, 1990, LeBlanc et al., 1991). BP has also been observed in wild Trachypenaeus similis from Florida and Mississippi and in wild Protrachypene precipua from Ecuador. BP infects larval, juvenile and adult prawns and may cause disease in the larval stages of P. vannamei (Akamine and Moores, 1989). Disease does not apparently occur in wild populations infected with BP (Brock and Lightner, 1990). BP has not been reported outside of the Americas and Hawaii.

Clinical signsEpizootics of BP are characterised by sudden, high mortality rates among larval, and postlarvae prawns. Disease is most severe in the mysis stages. Gross signs of infection include reduced feeding and growth rates and increased fouling (Lightner, 1988). Infected prawns may also display a milky-white midgut. BP may exist at subclinical levels in juvenile and adult populations.

Gross pathologyThere is little gross pathology associated with BP infection as mortalities occur very

21

quickly in early life stages.

HistopathologyBP infections are limited to the hepatopancreas and the anterior mid-gut epithelium. BP is an enveloped baculovirus which forms a characteristic, intranuclear, eosinophilic tetrahedral occlusion body (OB) in infected cells (Couch, 1974a and b). The average size of the BP nucleocapsid is 270 x 50 nm, however the size of the enveloped virions varies between regions, indicating that several strains of BP are likely to exist (Lightner, 1996). Infected nuclei show hypertrophy and chromatin margination (Couch, 1974a and b).

DiagnosisDiagnosis is based on the presence of clinical signs of disease, the history of infection within a culture facility, region and species, and the demonstration of tetrahedral OB’s in the epithelial cell nuclei in preparations of the hepatopancreas, midgut or faeces. BP OB’s may be observed by phase contrast or bright field microscopy in squash preparations of the hepatopancreas or by using histological techniques for light and electron microscopy (Lightner, 1983). Prawn stocks suspected of carrying BP may be tested using a bioassay of P. vannamei as the indicator host (Overstreet et al., 1988).

DNA probes for BP have been developed (Bruce et al., 1993) and some of these are commercially available from DiagXotics (Don Lightner, personal communication).A PCR-based diagnostic test has been developed to detect BP in experimentally-infected P. vannamei (Wang et al., 1996, Stuck and Wang, 1996) but is not yet commercially available. A diagnostic enzyme-linked immunosorbent assay (ELISA) using rabbit antibody to BP has also been developed (Lewis, 1986).

TransmissionBP OB’s are passed in the faeces of infected prawns. Eggs and newly hatched nauplii may be exposed to the virus when contaminated with spawner faeces (Lightner, 1988). Older prawns may become infected by ingesting waterborne virus or virus in moribund prawns and prawn carcasses.

ViabilityBP is inactivated within 10 min at 60oC to 90oC. Desiccation for 48 hours also inactivates the virus (LeBlanc and Overstreet, 1991b). The effect of freezing and thawing on the viability of BP is unknown. However, it seems likely that BP would survive in frozen prawns given the protective function of the occlusion body ( Granados and Williams, 1986). It is considered that the occluded baculoviruses can persist in putrefying host bodies and are not inactivated between -20oC and +40oC (Longworth, 1975).

An infectivity experiment comparing four samples of viral material, from different sources, frozen at -70ºC for various periods of time (up to 3.5-years) was conducted (Overstreet, 1988;1994). The results indicate that there is no relationship between virulence and length of time the virus was frozen at this temperature. At -20ºC infectivity does not persist more than a few weeks (Don Lightner, personal communication).

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PreventionControl of BP involves adopting stringent management procedures such as: the use of spawners which are not passing BP OB’s in the faeces, avoidance of cross-contamination of larval batches, avoidance of contaminating eggs with spawner faeces, and the use alkaline disinfectants (Lightner, 1988). The protective OB formed by BP protects the virus in the external environment (Federici, 1986), allowing it to remain viable for a long period of time. Calcium hypochlorite inactivates BP within 1 hr at 200mg/l and within 20 sec at 1,600 mg/l. BP is also inactivated within 30 min when exposed to pH 3 and within 40 min under ultraviolet (UV) light at a wavelength of 254 nm.

ReferencesAkamine, A.Y. and Moores, J.L. 1989. A preliminary study on disinfection methods

of penaeid shrimp hatcheries contaminated with Baculovirus penaei. Abstract, J. World. Aquacult. Soc., 20; 11A.

Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne (ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp.245-424.

Bonami, J.R., Bruce, L.D., Poulos, B.T., Mari, J. and Lightner, D.V. 1995. Partial charactization and cloning of the genome PvSNPV (=BP-type baculovirus) pathogenic for Penaeus vannamei. Dis. Aquat. Org. 23: 59-66.

Bruce, L.D., Redman, R.M., Lightner, D.V. and Bonami, J.R. 1993. Application of gene probes to detect a penaeid shrimp baculovirus in fixed tissues using in situ hybridization. Dis. Aquat. Org. 17: 215-221.

Couch, J.A. 1974a. Free and occluded virus similar to baculovirus in hepatopancreas of pink shrimp. Nature 247: 229-231.

Couch, J.A., 1974b. An enzootic nuclear polyhedrosis virus of the pink shrimp: ultrastructure, prevalence and enhancement. J. Invertebr. Pathol. 24(3): 311-331.

Durand, S., Lightner, DV. and Bonami, JR. 1998. Differentiation of BP-type baculovirus strains using in situ hybridization. Dis. Aquat. Org. 32:237-239

Federici, B.A. 1986. Ultrastructure of baculoviruses, In: R.R. Granados and F.A. Frederici (eds.) The Biology of Baculoviruses, Vol. 1. CRC Press Inc., Boca Raton, Florida. p. 37.

Granados, R.R. and Williams, K.A. 1986. Chapter 4 - In vivo infection and replication of baculoviruses. In: R.R. Granados and F.A. Frederici (eds.) The Biology of Baculoviruses, Vol. 1. CRC Press Inc., Boca Raton, Florida. pp. 89-108.

LeBlanc, B.D. and Overstreet, R.M. 1991a. Efficacy of calcium hypochlorite as a disinfectant against the shrimp virus Baculovirus penaei. J. Aquat. Anim. Health 3: 141-145.

LeBlanc, B.D. and Overstreet, R.M. 1991b. Effect of desiccation, pH, heat and ultraviolet irradiation on viability of Baculovirus penaei. J. Invertebr. Pathol. 57: 277-286.

LeBlanc, D.H., Overstreet, R.M. and Lotz, J.M. 1991. Relative susceptibility of Penaeus aztecus to Baculovirus penaei. J. World Aquaculture Society 22: 173-177.

Lewis, D.H. 1986. An enzyme-linked immunosorbent assay (ELISA) for detecting penaeid baculovirus. J. Fish Dis. 9: 519-522.

Lightner, D.V. 1983. Diseases of cultured penaeid shrimp. In: J.R. Moore (ed. in chief) CRC Handbook of Mariculture, Vol. 1. J.P. McVay (ed.), Crustacean

23

Aquaculture, CRC Press, Boca Raton, Florida. pp. 289-320.Lightner, D.V. 1988. Diseases of cultured penaeid shrimp and prawns. In: C.J.

Sindermann and D.V. Lightner (eds.) Disease Diagnosis and Control in North American Marine Aquaculture, 2nd. ed. Elsevier, New York. pp. 8-127.

Lightner, 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA., USA.

Lightner, D.V., Redman, R.M. and Almada Ruiz, E.A. 1989. Baculovirus penaei in Penaeus stylirostris (Crustacea: Decapoda) cultured in Mexico: Unique cytopathology and a new geographic record. J. Invertebr. Pathol. 53: 137-139.

Longworth, J.F. 1975. Viruses and Lepidoptera. In: A.J. Gibbs (ed.), Viruses and invertebrates. Elsevier Publ. Co., N.Y. pp. 429-441.

Overstreet, R.M., Stuck, R.A., Krol, R.A. and Hawkins, W.E. 1988. Experimental infections with Baculovirus penaei in the white shrimp Penaeus vannamei as a bioassay. J. World Aquacult. Soc. 29: 175-187.

Overstreet, R.M. 1994. BP (Baculovirus Penaei) in penaeid shrimps. USMSFP 10th Anniversary Review, GCRL Special Publication No. 1, 97-106, 1994

Stuck, K.C. and Wang, S.Y. 1996. Establishment and persistence of Baculovirus penaei infections in cultured pacific white shrimp, Penaeus vannamei. J. Invertebr. Pathol. 68: 59-64.

Wang, S.Y., Hong, C. and Lotz, J.M. 1996. Development of a PCR procedure for the detection of Baculovirus penaei in shrimp. Dis. Aquat. Org. 25: 123-131.

Baculoviral Midgut Gland Necrosis Virus (BMNV) BMNV was accepted by the International Committee on Taxonomy of Viruses (ICTV; Franki et al., 1991) and designated PjNOB (Penaeus japonicus non occluded baculovirus; Arimoto et al., 1995). The non-occluded baculoviruses, including PjNOB have been omitted from the latest publication of the ICTV (Volkman et al., 1995). Therefore, we have referred to this virus as BMNV.

BMNV as a nonoccluded baculovirus was classified as a “type-C” baculovirus under the ICTV 1991 classification. However, the ICTV 1997 classification removed this group from the baculoviruses and suggested that they should be regarded as unclassified bacilliform viruses. Additional nonoccluded baculoviruses in prawn species other than P. japonicus, have been grouped as BMNV-like viruses (Brock, 1991; Lightner, 1996), but this is misleading as it gives the impression that they are similar to BMNV when the necessary information is lacking. The term nonoccluded bacilliform viruses is more correct at this point in time.

Baculoviral midgut gland necrosis virus (BMNV) is known from hatchery-reared P. japonicus in southern Japan (Sano et al., 1984) and Korea (Lightner, 1996). In Japan, epizootics of BMNV have occurred since 1971 (Sano et al., 1984). Despite repeated introduction of P. japonicus to Hawaii, Italy, Spain, France, Brazil and elsewhere, BMNV epizootics have not been reported outside Japan and Korea (Lightner, 1993). BMNV-type infections have been observed in P. monodon in East and South-East Asia. BMNV has been experimentally transmitted to P. chinensis and P. semisulcatus, whereas Metapenaeus ensis and the crab Portunus pelagicus appear to

24

be refractory to experimental infection. P. monodon is highly susceptible to experimental infection with BMNV (Momoyama and Sano, 1996).

The geographic distribution of the BMN-like agents includes Japan, Australia, Indonesia and the Philippines (Brock, 1991; Lightner, 1996)

Clinical signsMortalities in hatcheries occur in mysis through to 20 day old postlarvae (PL) and may reach up to 98% in PL9-10 (Sano et al., 1981). The onset of mortality is usually rapid. The first gross sign of infection is the white, turbid appearance of the hepatopancreas. Severely affected postlarvae may float inactively on the surface of the water and display a white midgut line (Lightner, 1988).

Infection with BMNV is reported to be subclinical in juvenile to subadult P. japonicus (Momoyama and Sano, 1989)

Gross PathologyApart from the white, turbid appearance of the hepatopancreas and/or midgut, there is little gross pathology associated with BMNV as mortalities occur rapidly in the early life stages.

HistopathologyBMNV infects the nuclei of hepatopancreocytes and causes margination of chromatin, hypertrophy, nucleolar dissociation and ultimately the collapse of the hepatopancreas. BMNV virions average 310 x 72 nm and occur within eosinophilic to basophilic intranuclear inclusion bodies (Sano et al., 1984).

DiagnosisDiagnosis of BMNV is based on the presence of clinical signs, the disease history of the culture facility, region and species and the demonstration of hypertrophied hepatopancreatic cell nuclei in Giemsa-stained impression smears (Momoyama, 1983). Hypertrophied hepatopancreatic cell nuclei appear white under darkfield microscopic illumination of unstained squash preparations (Momoyama, 1983). Labelled polyclonal antibodies have been raised against BMNV for direct diagnosis using fluorescent antibodies (Sano et al., 1984). BMNV infection is confirmed by the demonstration of rod-shaped, enveloped virions, 72 x 310 nm in hepatopancreatic cell nuclei. Asymptomatic carriers of BMNV may be identified using bioassays in which mysis stage P. japonicus are used as the indicator host (Momoyama and Sano, 1988). Molecular probes and PCR primers for BMNV are being developed in Japan (Arimoto et al., 1995).

Transmission and potential carriersBMNV is introduced to hatcheries with wild-caught broodstock and transmission to the larvae is thought to occur when the virus is shed with the faeces during spawning (Momoyama, 1988). Water-borne BMNV has been successfully transmitted experimentally to zoea and postlarvae (Momoyama and Sano, 1989). BMNV may be transmitted per os within populations of postlarvae. Wild penaeid species, which may be carriers of this virus have not been identified.

Viability

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Water-borne transmission of BMNV has been successful, indicating that the virus, although non-occluded, can remain viable in the external environment for a period of time. An increase in temperature results in a decrease in the time BMNV remains viable in seawater. BMNV was inactivated within 3 hr at 30oC (Momoyama, 1989).

PreventionBMNV can be controlled by avoiding contamination of the eggs and nauplii with broodstock faeces by washing them in clean seawater prior to stocking (Momoyama and Sano, 1989). BMNV is inactivated within 10hr in 25% NaCl and within 18 hr in ethyl ether at 4oC. BMNV is also inactivated by exposure to UV irradiation for 20 min (Momoyama, 1989 a and b).

Present status of BMNVDiseases caused by BMNV continue to be a problem in P. japonicus hatcheries in Japan and Korea.

According to recent reviews on shrimp/prawn virus diseases in Japan, BMNV is not currently a significant problem in Japan and Korea. This is because the egg/nauplii rinsing and disinfection methods are effective in reducing faecal (containing BMNV)contamination of spawns (Don Lightner, personal communication).

ReferencesArimoto, M., Yamazaki, T., Mizuta, Y., Furusawa, I. 1995. Characterization and

partial purification of the genomic DNA of a baculovirus from Penaeus japonicus (PjNOB = BMNV). Aquaculture 132: 213-220.

Brock, J.A. 1991. An overview of diseases of cultured crustaceans in the Asia Pacific region. In: Fish Health Management in Asia-Pacific . Report on a regional study and workshop on fish diseases and fish health management.

Lightner, D.V. 1988. Diseases of cultured penaeid shrimp and prawns. In: C.J. Sindermann and D.V. Lightner (eds.) Disease Diagnosis and Control in North American Marine Aquaculture, 2nd. ed. Elsevier, New York. pp. 8-127.

Lightner, D.V. 1993. Diseases of cultured penaeid shrimp. In: J.R. Moore (ed. in chief) CRC Handbook of Mariculture, Second Edition, Vol. 1. J.P. McVay (ed.), Crustacean Aquaculture, CRC Press, Boca Raton, Florida. pp. 393-486.

Lightner, 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA.,USA.

Momoyama, K. 1983. Studies on baculoviral mid-gut gland necrosis of Kuruma shrimp (Penaeus japonicus) III. Presumptive diagnostic techniques. Fish Pathol. 17: 263-268.

Momoyama, K. 1988. Infection source of baculoviral mid-gut gland necrosis (BMN) in mass production of Kuruma shrimp larvae, Penaeus japonicus. Fish Pathol. 23: 105-110.

Momoyama, K. 1989a. Tolerance of baculoviral mid-gut gland necrosis virus (BMNV) to ether, NaCl concentration and pH. Fish Pathol. 24: 47-49.

Momoyama, K. 1989b. Inactivation of baculoviral mid-gut gland necrosis (BMN) virus by ultraviolet irradiation, sunlight exposure, heat and drying. Fish Pathol. 24: 115-118.

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Momoyama, K. and Sano, T. 1988. A method of experimental infection of kuruma shrimp larvae, Penaeus japonicus Bate, with baculoviral mid-gut necrosis (BMN) virus. J. Fish Dis. 11: 105-111.

Momoyama, K. and Sano, T. 1989. Developmental stages of kuruma shrimp, Penaeus japonicus Bate, susceptible to baculovirus mid-gut gland necrosis (BMN) virus. J. Fish Dis. 12: 585-589.

Momoyama, K. and Sano, T. 1996. Infectivity of baculovirus midgut gland necrosis virus (BMNV) to larvae of 5 crustacean species. Fish Pathol. 31(2): 81-85.

Sano, T. Nishimura, T., Oguma, K., Momoyama, K. and Takeno, N. 1981. Baculovirus infection of cultured Kuruma shrimp, Penaeus japonicus in Japan. Fish Pathol. 25: 185-191.

Sano, T., Nishimura, T., Fukuda, H. and Hayashida, T. 1984. Baculoviral mid-gut gland necrosis (BMN) of Kuruma shrimp (Penaeus japonicus) larvae in intensive culture systems. Helgolander Meeresunters. 37: 255-264.

Volkman, L.E., Blissard, G.W., Friesen, P., Keddie, B.A., Possee, R., Theilmann, D.A. 1995. Baculoviridae. In: Murphy, F.A., Fauquet, C.M., Bishop, D.H.L., Ghabrial, S.A., Jarvis, A.W., Martelli, G.P., Mayo, M.A. and Summers, M.D. (eds.) Virus Taxonomy, Sixth Report of the International Committee on Taxonomy of Viruses., Springer-Verlag, Wein, New York. pp. 104-113.

Wilson, M. 1991. Baculoviridae. In: Francki, R.I.B., Fauquet, C.M., Knudson, D.L. and Brown, F. (eds.) Classification and Nomenclature of Viruses. Fifth Report of the International Committee on Taxonomy of Viruses, Archives Virology Supplementum 2. pp 117-123.

Monodon Baculovirus (MBV)

Monodon baculovirus (MBV) exists in Australia and is a recurring problem in P. monodon hatcheries. Following characterisation, MBV was designated PmSNPV (P. monodon singular Nucleopolyhedrovirus; Mari et al., 1993). However, until this virus is accepted as a baculovirus by the International Committee on Taxonomy of Viruses (ICTV) we will refer to it as MBV.

Monodon baculovirus (MBV) was first described by Lightner and Redman (1981) from P. monodon prawns cultured in Taiwan. MBV-like baculoviruses have been described for P. merguiensis, P. penicillatus, P. plebejus, P. esculentus, P. semisulcatus, P. kerathurus and P. vannamei and occur in most areas of the Indo-Pacific where penaeid prawns are cultured (Brock and Lightner, 1990). In Australia, MBV has been reported in cultured P. monodon and wild P. merguiensis (Doubrovsky et al., 1988). An MBV-like virus, Plebejus baculovirus (PBV; Lester et al., 1987) was described from cultured P. plebejus from Australia. Another MBV-like virus is reported from Metapenaeus ensis cultured in Taiwan (Chen et al., 1989b). It is believed MBV-like viruses exist as a complex, made distinct by geographic location. MBV has been detected in Peneaus monodon in Taiwan, the Philippines, Malaysia, French Polynesia, Hawaii, Kenya, Mexico, Singapore, Indonesia, Israel, Thailand (Fulks and Main, 1992). Except for possibly Puerto Rico and the Dominican Republic (where P. monodon may still be cultured), MBV has been eradicated from the

27

Americas (i.e the USA {SC, TX, HI}, Ecuador, Mexico, Brazil, etc.) where it was once found in imported stocks of P. monodon (Don Lightner, personal communication).

Clinical signsMortalities occur primarily among postlarvae in the hatchery, although disease may also occur in juvenile and adult prawns (Johnson and Lightner, 1988). Cumulative mortality among postlarvae (PL) may reach over 90%. Disease decreases from (PL16 to 25 (Paynter et al., 1992; Natividad et al., 1992). In contrast to the situation in Australia, hatchery and nursery infections of P. monodon in Thailand are extremely common and result in no abnormal mortality of larvae and PL under good rearing conditions (Fegan et al. 1991). This may reflect differences in geographical strains of the virus or the host prawns.

Gross PathologyDue to the sudden onset of mortality in the early life stages, there are few gross signs of disease apart from reduced feeding and growth rates and an increase in gill and surface fouling (Lightner 1988). Severely infected prawns may display a white hepatopancreas and midgut.

HistopathologyMBV forms large, roughly spherical, eosinophilic, polyhedral occlusion bodies (OB’s) within the nuclei of hepatopancreatic cells. OB’s may occur singularly or in multiples. Early infection may be detected by the presence of hypertrophied nuclei with marginated chromatin and displaced nucleolus. In heavy infections the anterior midgut epithelium may also be infected (Lightner et al., 1983a). The average size of an MBV nucleocapsid is 246 x 42 nm (Brock and Lightner, 1990). The nucleocapsids of MBV from Australia have been reported to be longer (260-300 nm) and wider (45-52 nm) that those from other Pacific areas (Doubrovsky et al., 1988). PBV virions and nucleocapsids are similar in size to those of MBV (Lester et al., 1987).

DiagnosisDefinitive diagnosis is based on the histological demonstration of eosinophilic OBs within the nuclei of hepatopancreocytes. OB’s may also be detected in fresh squash preparations of the hepatopancreas stained with 0.05% aqueous malachite green. DNA probes for MBV have been developed in numerous countries, including Australia (Vickers et al., 1993) and are the most reliable method of detecting MBV infection. A commercial diagnostic probe is available from DiagXotics Co. Ltd., Wilton CT and primers for PCR diagnosis have been published by Chang et al. (1993).

A new method, MBV-reactive PCR assay, has been developed (Belcher, 1997). This detection assay also provides information about each MBV isolate at the DNA level. This information can then be used to distinguish isolates and allow genetically similar isolates to be grouped according to virulence or geographic origin.

TransmissionMBV is transmitted by ingestion of free virus and OB’s and by cannibalism (Paynter et al., 1992). It is also believed to be transmitted vertically from broodstock to offspring (Bonami et al., 1986), but this has not yet been proven. Postlarvae have

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been experimentally infected with MBV via waterborne transmission (Paynter et al., 1992, Natividad et al., 1992), indicating that MBV may remain viable in the external environment for some time due to the protective nature of the polyhedral occlusion body (Federici, 1986). It is presumed that wild P. monodon and other species within the geographical range of the virus are carriers (Brock and Lightner, 1990).

ViabilityMBV remains viable after freezing and thawing as well as at 4oC for 24 hours (Payntner et al., 1992). MBV has been inactivated within 30 min at 60oC (Jan Payntner, unpublished).

Prevention MBV is controlled in the hatchery by avoiding contamination and by strict disinfection regimes. Infected animals should be eradicated and removed from the facility (Lightner, 1988). All equipment and tanks should be disinfected routinely between batches of larvae and equipment used in the spawning area should be segregated from the hatchery (Wyban and Sweeney, 1991). Eggs should be separated from spawner faeces in which MBV OB’s may be passed and washed in clean seawater, iodophore and/or formalin (Chen et al., 1992). Spann et al. (1993) found that 24 hr exposure to 1000 mg/l calcium hypochlorite was necessary to inactivate MBV.

Present status of MBV infectionMBV continues to be a problem in P. monodon hatcheries. A number of MBV epidemics occurred in Australian prawn hatcheries during the late 1980’s. However few outbreaks of MBV have been reported in recent years.

Some States impose movement controls with regard to controlling the spread of MBV. As example, samples of postlarval P. monodon and P. japonicus entering NSW for growout must test negative for MBV and other declared diseases (NSW Fisheries Management Act 1994) prior to stocking. Similarly, progeny, produced in NSW hatcheries, of broodstock sourced from interstate must also test negative prior to stocking. The restrictions relating to virus diseases are stipulated in each prawn farmer's aquaculture permit, which in turn is authorised by the Fisheries Management Act (1994) and related Regulations.References

Belcher, C.R. 1997. Monodon baculovirus (MBV) and its detection by DNA-technologie. Abstract In: Australian Prawn Farmers Association, Annual Conference, 26-27 July 1997.

Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne (ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Bonami, J.R., Brehelin, M and Weppe, M. 1986. Observations sur la pathogenicite, la transmission et la resistance du MBV (Monodon Baculovirus). Abstract, 2nd Int. Coll. Pathol. Mar. Aquac. p. 119.

Chen, S.N., Chang, P.S., Kou, G.H. and Lightner, D.V. 1989a. Studies on virogenesis and cytopathology of Penaeus monodon Baculovirus (MBV) in the great tiger

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prawn (Penaeus monodon) and in the red tail prawn (Penaeus penicillatus). Fish Pathol. 24(2): 89-100.

Chang, P.S., C.F. Lo, G.H. Kou, C.C. Lu and S.N. Chen. 1993. Purification and amplification of DNA from Penaeus monodon-type baculovirus (MBV). Journal of Invertebrate Pathology 62: 116-120.

Chen, S.N., Lo, C.F., Lui, S.M. and Kou, G.H. 1989b. The first identification of Penaeus monodon baculovirus (MBV) in cultured sand shrimp, Metapenaeus ensis. Bull. EAFP 9(3): 62-64.

Chen, S.N., Chang, P.S. and Kou, G.H. 1992. Infection route and eradication of Penaeus monodon Baculovirus (MBV) in larval giant tiger prawns, Penaeus monodon. In: W. Fulks and K.L. Main (eds.) Diseases of Cultured Shrimp in Asian and the United States. The Oceanic Institute, Hawaii. pp. 177-184.

Doubrovsky, A. Paynter, J.L., Sambhi, S.K., Atherton, J.G. and Lester, R.J.G. 1988. Observations on the ultrastructure of baculovirus in Australian Penaeus monodon and Penaeus merguiensis. Aust. J. Mar. Freshwater Res. 39: 743-749.

Fegan, D.F., T.W. Flegel, Siriporn Sriurairatana and Manuschai Waiakrutra. 1991. The occurrence, development and histopathology of monodon baculovirus in Penaeus monodon in Southern Thailand. Aquaculture. 96: 205-217.

Federici, B.A. 1986. Chapter 3: Ultrastructure of baculoviruses. In: R.R. Granados and B.A. Federici (eds.) The Biology of Baculoviruses, Vol. 1. CRC Press Inc., Boca Raton, Florida. pp. 61-88.

Fulks, W. and Main, K.L. (1992). Introduction: 3-33. In Diseases of Cultured Penaeid Shrimp in Asia and the United States. Proceedings of a Workshop in Honululu, Hawaii April 27-30,1992

Granados, R.R. and Williams, K.A. 1986. Chapter 4 - In vivo infection and replication of baculoviruses. In: R.R. Granados and F.A. Frederici (eds.) The Biology of Baculoviruses, Vol. 1. CRC Press Inc., Boca Raton, Florida. pp. 89-108.

Johnson, P.T. and Lightner, D.V. 1988. Rod-shaped nuclear viruses of crustaceans: gut-infecting species. Dis. Aquat. Org. 5: 123-141.

Lester, R.J.G., Doubrovsky, A. Paynter, J.L., Sambhi, S.K. and Atherton, J.G. 1987. Light and electron microscope evidence of baculovirus infection in the prawn Penaeus plebejus. Dis. Aquat. Org. 3: 217-219.

Lightner, D.V. 1988. Diseases of cultured penaeid shrimp and prawns. In: C.J. Sindermann and D.V. Lightner (eds.) Disease Diagnosis and Control in North American Marine Aquaculture, 2nd. ed. Elsevier, New York. pp. 8-127.

Lightner, D.V. and Redman, R.M. 1981. A baculovirus-caused disease of the penaeid shrimp, Penaeus monodon. J. Invertebr. Pathol. 38: 299-302.

Lightner, D.V., Redman, R.M. and Bell, T.A. 1983. Observations on the geographic distribution, pathogenesis and morphology of the baculovirus from Penaeus monodon Fabricius. Aquaculture 32: 209-233.

Mari, J., Bonami, J.R., Poulos, B. and Lightner, D.V. 1993. Preliminary characterization and partial cloning of the genome of a baculovirus from Penaeus monodon (PmSNPV = MBV). Dis. Aquat. Org. 16: 207-215.

Natividad, J.M. and Lightner, D.V. 1992. Susceptibility of the different larval and postlarval stages of the black tiger prawn, Penaeus monodon Fabricius, to monodon baculovirus (MBV). Diseases in Asian Aquaculture I. pp. 111-124.

Paynter, J.L., Vickers, J.E. and Lester, R.J.G. 1992. Experimental transmission of

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Penaeus monodon-type baculovirus (MBV). In: Diseases in Asian Aquaculture 1. Shariff, M., Subasinghe, R.P. and Arthur, J.R. (eds.). Fish Health Section, Asian Fisheries Society, Manila. pp. 97-109.

Spann, K.M., Paynter, J.L. and Lester, R.J.G., 1993. Efficiency of chlorine as a disinfectant against monodon baculovirus (MBV). Asian Fish. Sci. 6: 295-301.

Vickers, J.E., Bonami, J.R., Flegel, T.W., Ingham, A.B., Kidd, S.P., Lester, R.J.G., Lightner, D.V., Mari, J. Pemberton, J.M., Spradbrow, P.B., Wang, J.H., Wong, F.Y.K. and Young, P.R., 1993. A gene probe for monodon baculovirus. Abstract, Conference on Marine Biotechnology in the Asian Pacific Region, Bangkok. p. 76.

Wyban, J.A. and Sweeney, J.N. 1991. The Oceanic Institute Shrimp Manual, Intensive Shrimp Production Technology. The Oceanic Institute, Makapuu Point, Honolulu.

Infectious pancreatic necrosis virus (IPNV)

Infectious pancreatic necrosis is primarily an acute, clinical disease in young freshwater salmonids (Wolf, 1988), with an increasing number of outbreaks in Atlantic salmon post-smolts (Jarp et al., 1995; Smail et al., 1995). The aetiological agent, IPN virus (IPNV), is a member of the Birnaviridae family (Dobos et al., 1979) which also contains numerous IPN-like viruses that have been isolated from a broad range of other fish species (Wolf, 1988), molluscs (Underwood et al., 1977) and crustaceans (Hill, 1982). IPNV and IPN-like virus have been isolated from adult P. japonicus (Bovo et al., 1984; Georgetti, 1989). Geographic distribution of the virus is probably world-wide (OIE, 1995), but to date the virus has not been isolated in Australia.

IPN is listed under ‘Other significant diseases’ in the OIE International Aquatic Animal Health Code (1995). IPN is also included in List III of the European Union Directive 93/54 (1993).

Clinical signsAn IPN-like virus was associated with high mortalities in up to 42.8% of laboratory bred adult P. japonicus. Other clinical signs in these infected animals included lethargy and erosive-necrotic lesions on the thoracic limbs and uropods (Bovo et al., 1984). Giorgetti (1989) observed weakness but no mortalities in adult P. japonicus with natural IPNV infection. Experimentally infected post-larvae displayed locomotor ataxia without mortalities (Giorgetti, 1989). The virus causes high mortalities in young salmonids, especially in rainbow and brook trout in fresh water.

Gross PathologyGross signs of serious degeneration of the hepatopancreas may be evident in juvenile P. japonicus (Giorgetti, 1989).

HistopathologyDegeneration of the hepatopancreas with slight vacuolization of the tubuli cells may be observed in post-larvae and juveniles infected with IPNV (Giorgetti, 1989).

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DiagnosisIPNV can be isolated using a number of established fish cell lines (Wolf & Mann, 1980; Bovo et al., 1985) and is cytopathic. Following isolation, identification can be accomplished using any of a variety of immunodiagnostic techniques such as serum neutralisation (Ishiguro et al., 1984), enzyme-linked immunosorbent assay (ELISA) (Hattori et al., 1984), immunodotblot (Hsu et al., 1989), Western blotting (Williams et al., 1994), immunofluorescence (Swanson & Gillespie, 1981) and immunoperoxidase (Nicholson & Henchal, 1978) using either polyclonal or monoclonal antibodies. Virus isolation and identification can take up to two weeks to complete. Detection of IPNV by hybridisation using either oligonucleotide DNA probes (Rimstad et al., 1990) or cloned cDNA probes (Dopazo et al., 1994) has been reported but in both reports it was shown that virus isolation in cell cultures was more sensitive (up to 105-fold in some cases) and hybridisation assisted in virus identification only following isolation in cell culture. In addition, a PCR assay (Blake et al., 1994) has been reported which appears to be as sensitive as virus isolation. It was not serotype specific and not necessarily IPNV-specific but could be used for the identification of aquatic birnaviruses in general.

Disease agent predilection sitesIPN-like virus was isolated from the hepatopancreas of P. japonicus (Bovo et al., 1984).

Transmission and potential carriersNo information is available on transmission of IPNV in prawns.

IPNV transmission in salmonids occurs both vertically (via eggs and semen; Wolf et al., 1963) and laterally (via water, equipment, birds, blood sucking parasites, faeces, urine and sex products of infected fish, and bivalve molluscs (Billi & Wolf, 1969; Peters & Neukirch, 1986; Halder & Ahne, 1988; Mortensen et al., 1992; Mortensen, 1993).

ViabilityWhipple & Rohovec (1994) found that IPNV was highly resistant to low pH, surviving over 14 days at pH 4 for 22°C. In fish silage (pH 3.8–4.3), survival time was equally long. IPN is stable for several months at 4°C (Malsberger & Cerini, 1963). The longest record of survival at 4°C is four years. IPNV remains viable for several years at –70°C (Malsberger & Cerini, 1963). Gosting and Gould (1981) found IPNV was inactivated after 16 hours at 60°C but low levels of infectivity were found after five hours. High levels of virus can still be detected after 22 hours at 50°C (Malsberger & Cerini, 1963). Whipple and Rohovec (1994) found the virus survived for 8h at 60°C, 3.5h at 65°C, 2h at 70°C and ten minutes at 80°C. There is no significant loss in infectivity at room temperature over a period of 27 days (Tisdall & Phipps, 1987). IPNV is not inactivated by an acidic pH unless the sample (silage) is heated for at least two hours to at least 60°C (Smail et al., 1993). Humphrey et al. (1991) found the virus titre dropped five orders of magnitude after four hours at 60°C and dropped three orders of magnitude after 15 minutes at 70°C. This data indicates that IPNV is a robust virus with long survival in the environment.

PreventionThere are no effective vaccines or chemotherapeutics are available.

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Present status of IPN diseaseNo reports of IPNV in prawns have occurred subsequent to that of Giorgetti (1989).

ReferencesBilli, J.L. and 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., Schill, W., McAllister, P., Lee, M.K., Singer, J. and Nicholson, B. 1994. Detection and identification of aquatic birnaviruses by a PCR assay. International Symposium on Aquatic Animal Health, Seattle, Washington (USA), September 4–8, 1994. Program and Abstracts. Davis, California (USA) University of California, School of Veterinary Medicine 1994 p. W-1.1.

Bovo, G., Ceschia, G., Giorgetti, G. and Vanelli, M. 1984. Isolation of an IPN-like virus from adult kuruma shrimp (Peneaus japonicus). Bulletin of the European Association of Fish Pathologists, 4: 21.

Bovo, G., Giorgetti, G. and Ceschia, G. 1985. Comparative sensitivity of five fish cell lines to wild infectious pancreatic necrosis viruses isolated in northeastern Italy. In Fish and Shellfish Pathology. Ed A. E. Ellis Academic Press, London pp. 289–293.

Dobos, P., Hill, B.J., Hallett R., Kells, D.T.C., Becht, H. and Teninges, D. 1979. Biophysical and biochemical characterization of five animal viruses with bisegmented double-stranded RNA genomes. Journal of Virology 32, 593–605.

Dopazo, C.P., Hetrick, F.M. and Samal, S.K. 1994. Use of cloned cDNA probes for diagnosis of infectious pancreatic necrosis virus infections. Journal of Fish Diseases 17, 1–16.

Giorgetti, G. 1989. Disease problems in farmed penaeids in Italy in Advances in Tropical Aquaculture, Tahiti, AQUACOP, IFREMER, Actes de Colloque 9: 75-87

Gosting, L.H. and Gould, R.W. 1981. Thermal inactivation of infectious hematopoietic necrosis and infectious pancreatic necrosis viruses. Applied and Environmental Microbiology 41, 1081–1082.

Halder, M. and Ahne, W. 1988. Freshwater crayfish Astacus astacus–a vector for infectious pancreatic necrosis virus (IPNV). Diseases of Aquatic Organisms 4, 205–209.

Hattori, M., Kodama, H., Ishiguro, S., Honda, A., Mikami, T. and Izawa, H. 1984. In vitro and in vivo detection of infectious pancreatic necrosis virus in fish by enzyme-linked immunosorbent assay. American Journal of Veterinary Research 45, 1876–1879.

Hill, B.J. 1982. Infectious pancreatic necrosis virus and its virulence. In: Microbial Diseases of Fish. Ed R. J. Roberts. Academic Press, London pp. 91–114.

Hsu, Y.L., Chiang, S.Y., Lin, S.T. and Wu, J.L. 1989. The specific detection of infectious pancreatic necrosis virus in infected cells and fish by the immuno dot blot method. Journal of Fish Diseases 12, 561–571.

Humphrey, J.D., Smith, M.T., Gudkovs, N. and Stone, R. 1991. Heat susceptibility of selected exotic viral and bacterial pathogens of fish–A report of a study undertaken for the Australian Quarantine Inspection Service. Australian Fish Health Reference Laboratory, CSIRO, Australian Animal Health Laboratory, Geelong.

Ishiguro, S., Izawa, H., Kodama, H., Onuma, M. and Mikami, T. 1984. Serological relationships among five strains of infectious pancreatic necrosis virus. Journal of Fish Diseases 7, 127–135.

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Jarp, J., Gjevre, A.G., Olsen, A.B. and Bruheim, T. 1995. Risk factors for furunculosis, infectious pancreatic necrosis and mortality in post-smolts of Atlantic salmon, Salmo salar L. Journal of Fish Diseases 18, 67–78.

Malsberger, R.G. and Cerini, C.P. 1963. Characteristics of infectious pancreatic necrosis virus. Journal of Bacteriology 86, 1283–1287.

Mortensen, S.H. 1993. Passage of infectious pancreatic necrosis virus (IPNV) through invertebrates in an aquatic food chain. Diseases of Aquatic Organisms 16, 41–45.

Mortensen, S.H., Bachere, E., Le Gall G. and Mialhe E. 1992. Persistence of infectious pancreatic necrosis virus (IPNV) in scallops Pecten maximus. Diseases of Aquatic Organisms 12, 221–227.

Nicholson, B.L. and Henchal, E.A. 1978. Rapid identification of infectious pancreatic necrosis virus in infected cell cultures by immunoperoxidase techniques. Journal of Wildlife Diseases 14, 465–469.

OIE 1995. Diagnostic Manual for Aquatic Animal Diseases. 1st edition. Office International des Epizooties, Paris 195 pp.

Peters, F. and Neukirch, M. 1986. Transmission of some fish pathogenic viruses by the heron, Ardea cinerea. Journal of Fish Diseases 9, 539–544.

Rimstad, E., Krona, R., Hornes, E., Olsvik, Ø. and Hyllseth, B. 1990. Detection of infectious pancreatic necrosis virus (IPNV) RNA by hybridization with an oligonucletotide DNA probe. Veterinary Microbiology 23, 211–219.

Smail, D.A., McFarlane, L., Bruno, D.W. and McVicar, A.H. 1995. The pathology of an IPN-Sp sub-type (Sh) in farmed Atlantic salmon, Salmo salar L., post-smolts in the Shetland Isles, Scotland. Journal of Fish Diseases 18, 631–638.

Smail, D.A., Huntly, P.J. and Munro, A.L.S. 1993. Fate of four fish pathogens after exposure to fish silage containing fish farm mortalities and conditions for the inactivation of infectious pancreatic necrosis virus. Aquaculture 113, 173–181.

Swanson, R.N. and Gillespie, J.H. 1981. An indirect fluorescent antibody test for the rapid detection of infectious pancreatic necrosis virus in tissues. Journal of Fish Diseases 4, 309–315.

Tisdall, D.J. and Phipps, J.C. 1987. Isolation and characterisation of a marine birnavirus from returning quinnat salmon (Oncorhynchus tshawytscha) in the south island of New Zealand. New Zealand Veterinary Journal 35, 217–218.

Underwood, B.O., Smale, C.J., Brown, F. and Hill, B.J. 1977. Relationship of a virus from Tellina tenuis to infectious pancreatic necrosis virus. Journal of General Virology 36, 93–109.

Whipple, M.J. and Rohovec, J.S. 1994. The effect of heat and low pH on selected viral and bacterial fish pathogens. Aquaculture 123, 179–189.

Williams, L.M., McRae, C.L., Crane, M.S. and Gudkovs, N. (1994). Identification of Fish Viruses by Western Blot Technique. Aust. Soc. Microbiol. Ann. Sci. Mtg., Melbourne, 25–30 Sept., 1994. Australian Microbiology 15. A-129.

Wolf, K. 1988. Infectious pancreatic necrosis. In Fish Viruses and Fish Viral Diseases (K. Wolf).Cornell University Press, Ithaca, New York pp. 115–157.

Wolf, K. and Mann, J.A. 1980. Poikilotherm vertebrate cell lines and viruses: a current listing for fishes. In Vitro 16, 168–179.

Wolf, K., Quimby, M.C. and Bradford, A.D. 1963. Egg-associated transmission of IPN virus of trouts. Virology 21, 317–321.

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Rhabdovirus Of Penaeid Shrimp (RPS)

Rhabdovirus of Penaeid Shrimp (RPS) was the first rhabdovirus to be recorded from a cultured penaeid (Lu et al., 1991). The virus has been isolated from P. stylirostris and P. vannamei from Hawaii and Ecuador simultaneously infected with IHHNV. The interaction between RPS and IHHNV is not clear but it has been suggested that RPS affects the natural defence system of prawns, rendering them more susceptible to infection by other agents (Nadala et al., 1992). RPS has not been recorded outside the Americas. RPS replicates in the established fish cell line, epithelioma papulosum cyprini (EPC, Lu et al., 1991). It has been suggested that RPS may be a fish rhabdovirus which uses prawns as a carrier host because it is very similar to fish rhabdoviruses in morphology and can replicate in a fish cell line (Lightner, 1996).

Lu & Loh (1994) concluded that RPSV was only partially related to the fish rhabdoviruses, infectious hematopoeitic necrosis virus (IHNV), viral haemorrhagic septicaemic virus (VHSV) but was closely related to spring viraemia of carp virus (SVCV, Rhabdovirus carpio). SVCV is listed as a notifiable disease in the OIE International Aquatic Animal Health Code.

Clinical signsInfected prawns show no signs of clinical disease and mortalities are not common (Lu et al., 1991), even among prawns infected experimentally (Nadala, et al., 1992).

Gross PathologyExperimental infection has resulted in hypertrophy of the lymphoid organ (Nadala et al., 1992).

HistopathologyRPS virions are cytoplasmic, bullet-shaped, contain ssRNA, and measure 65-77 nm x 115-138 nm (Lu et al., 1991). Extensive studies of the infectivity of RPS in P. stylirostris by Nadala et al. (1992) demonstrated that the virus replicated in the lymphoid organ and caused cytopathic changes. Infected lymphoid organs contained numerous, large hyperplastic nodules which contained hypertrophic nuclei, cytoplasmic vacuoles and basophilic cytoplasmic inclusions. Foci of necrosis and inflammation may also be observed in infected lymphoid organs.

DiagnosisRPS is diagnosed by the demonstration of viral particles by electron microscopy and/or the demonstration of cytopathic effects, such as necrosis, in carp EPC cell monolayers (Lightner, 1996). The cytopathic changes caused in the lymphoid organ cannot be used diagnostically as they are similar to those caused by other penaeid viruses such as yellow-head virus, lymphoidal parvo-like virus, Taura syndrome virus and lymphoid organ vacuolization virus. Polyclonal antibodies against RPS can be used with immunofluorescence to diagnose virus in a lymphoid organ smear (Nadala et al., 1992).

TransmissionMethods of transmission are unknown. It is possible that the virus may be transmitted between prawns by cannibalism.

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ViabilityThe viability of RPS in the external environment is unknown. RPS is sensitive to 20% ethyl ether, low pH (3.0) and 12 hr exposure to 37oC heat. Infectivity is lost after repeated freezing and thawing and by storage at -10oC (Lu and Loh, 1992).

ReferencesLightner, D.V. 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic

Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA.,USA.

Lu, Y., Nadala, E.C.B., Brock, J.A. and Loh, P.C. 1991. A new virus isolated from infectious hypodermal and hematopoietic necrosis virus (IHHNV)-infected penaeid shrimps. J. Virol. Methods 31: 189-196.

Lu, Y and Loh, P.C. 1992 Some biological properties of a rhabdovirus isolated from penaeid shrimps. Arch. Virol. 127: 339-343.

Lu, Y. and Loh, P.C. 1994. Viral structural proteins and genome analyses of the rhabdovirus of penaeid shrimp (RPS). Dis. Aquat. Org. 19: 187-192.

Nadala, E.C.B., Lu, Y., Loh, P.C. and Brock, J. 1992. Infection of Penaeus stylirostris (Boone) with a rhabdovirus isolated from Penaeus spp. Gyobyo Kenkyu 27(3) 143-147.

Bacteria

Necrotizing Hepatopancreatitis (NHP) NHP was first identified in cultured Penaeus vannamei from Texas, USA (Frelier, et al., 1992) where it has caused annual disease problems since 1985 (Johnson, 1990). NHP-like infections have also been identified in P. aztecus, P. setiferus, P. stylirostris and P. californiensis and have caused serious epizootics in Peru, Ecuador, Venezuela, Brazil, Panama and Costa Rica (Lightner, 1996; Jimenez, 1996). NHP has not been identified in Australian penaeids. NHP has also been named Texas necrotizing hepatopancreatitis (TNHP), Texas pond mortality syndrome (TPMS), Peru necrotizing hepatopancreatitis (PNHP) and Ecuador necrotizing hepatopancreatitis (ENHP).

Environmental factors appear to play an important role in the development of NHP. Lengthy periods of high temperature (29oC to 31oC) and elevated salinities (20 ppt to 40 ppt) have preceeded epizootics in all countries where NHP has been reported (Lightner, 1996).Clinical signsPrawns affected with NHP suffer lethargy, reduced growth, increased food conversion ratios, anorexia, soft shells, heavy surface fouling and black gills. Mortality may reach up to 99% of affected stock within 30 days of the onset of symptoms (Frelier et al., 1992; Lightner, 1993). If prawns are heavily infected, mortalities usually occur half way through the grow-out period (Frelier et al., 1992).

Gross PathologyThe hepatopancreas of affected prawns is typically atrophied and may appear pale and whitish (Krol et al., 1991), pale with black streaks or soft and watery (Lightner,

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1996).

Associated bacterial shell disease may result in melanised appendage erosion and/or cuticle lesions (Sindermann, 1990). Mortalities only occur if hepatopancreatic necrosis is extensive (Frelier et al., 1992).

HistopathologyNHP is caused by a Gram-negative, pleomorphic bacterium which infects hepatopancreatic epithelial cells. The bacterium may represent a new genus of alpha Proteobacteria (Frelier et al., 1994). There are two morphological variants of the NHP bacterium: a rod-shaped form, 0.3 m x 9 m, which lacks a flagella, and a helical form, 0.2 x 2.6-2.9 m, which possesses eight flagella on the basal apex and one or two flagella on the crest of the helix (Lightner et al., 1992).

The rod shaped form of the bacterium plays a dominant role in the pathogenesis of the disease (Frelier et al., 1992). The NHP bacterium is most closely related to bacterial endosymbionts of protozoa and more distantly related to rickettsia of the typhus and spotted fever groups (Loy et al., 1996a).

Infected hepatopancreatic epithelial cells are hypertrophied and contain large basophilic masses of bacteria in the cytoplasm. Infected cells may appear cuboidal, contain little stored lipid and have reduced or no secretory vacuoles (Lightner, 1996). Infected cells become necrotic, cease to function and evoke a host inflammatory response which results in the formation of multiple granulomatous lesions in affected hepatopancreata (Lightner, 1993).

DiagnosisDiagnosis of NPH is based on clinical signs and gross pathology of the hepatopancreas. Lesions and other cellular changes may be demonstrated using histochemical techniques. Giemsa and modified Steiner’s silver stain aid in the demonstration of bacteria within infected cells (Frelier et al., 1992). Some cytopathological changes may be observed in wet mounts of the hepatopancreas. The two morphological variants of the NHP bacterium may be distinguished by TEM. A DIG-labelled DNA probe for NHP bacterium is commercially available (Lightner, 1996).

The aetiologic agent of necrotizing hepatopancreatitis may be detected by PCR (Loy et al., 1996b).

TransmissionTransmission of the NHP bacterium appears to rely on direct ingestion of bacteria and a reservoir host may be involved. Cannibalism also plays a major role in transmission of the disease (Frelier et al., 1994). Specific environmental conditions, such as long periods of high temperature and elevated salinity, may also be required for the syndrome to become obvious (Frelier et al., 1993).

ViabilityNHP in tissue was not able to be transmitted by bioassay when subjected to sun drying for 3 weeks followed by storage at 20-22C for 2 months (Frelier et al., 1993).

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TreatmentNHP may be treated by using medicated feed containing oxytetracycline (Lightner, 1993). Metaphalatic therapy gave the highest survival and growth rates when medicated feeds were used (Bell and Lighter, 1991).

Present status of NHPThe incidence of mortality resulting from NHP has decreased in recent years due to the use of effective and readily accessible treatments.

ReferencesBell, T.A. and Lightner, D.V. 1991. Chemotherapy in aquaculture today – current

practices in shrimp culture: available treatments and their efficiency. In: C. Michel and D.J. Alderman (eds.) Chemotherapy in Aquaculture: from Theory to Reality. Office International des Epizooties, Paris. pp. 45-57.

Frelier, P.K., Sis, R.F., Bell, T.A. and Lewis, D.H. 1992. Microscopic and ultrastructural features of necrotizing hepatopancreatitis in Texas cultured shrimp (Penaeus vannamei). Vet. Pathol. 29: 269-277.

Frelier, P.F., Loy, J.K. and Kruppenbach, R. 1993. Transmission of necrotizing hepatopancreatitis in Penaeus vannamei. J. Invertebr. Pathol. 61: 44-48.

Frelier, P.F., Loy, J.K., Lawrence, A.L., Bray, W.A. and Brumbaugh, G.W. 1994. U.S. Marine Shrimp Farming Program 10th Anniversary Review. Gulf Coast Research Laboratory Special Publication No. 1. Ocean Springs, MI. pp. 55-58.

Jimenez, R. 1996. An epizootic of intracellular bacterium in cultured penaeid shrimp (Crustacea: Decapoda) in the gulf of Guayaquil, Ecuador. World Aquaculture ’96, book of abstracts. The World Aquaculture Society, Baton Rouge, LA. p. 186.

Johnson, S.K. 1990. Digestive gland manifestations. In: S.K. Johnson (ed.) Handbook of Shrimp Diseases. Sea Grant Publication No. TAMU-SG-90-601, Texas A&M University, Galveston.

Krol, R.M., Hawkins, W.E., and Overstreet, R.M. 1991. Rickettsial and mollicute infections in hepatopancreatic cells of cultured pacific white shrimp (Penaeus vannamei). J. Invertebr. Pathol. 57: 362-370.

Lightner, D.V. 1993. Diseases of cultured penaeid shrimp. In: J.P. McVey (ed.) CRC Handbook of Mariculture, Second edition, Volume 1, Crustacean Aquaculture. CRC Press Inc., Boca Raton, FL. p. 393-486.

Lightner, D.V. (ed.). 1996. A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA, USA.

Lightner, D.V., Redman, R.M. and Bonami, J.R. 1992. Morphologic evidence for a single bacterial epiology in Texas necrotizing hepatopancreatitis in Penaeus vannamei (Crustacea: Decapoda). Dis. Aquat. Org. 13:321-328.

Loy, J. K., Dewhirst, F.E., Weber, W., Frelier, P. F., Garber, T.L. Tasca, S. I. and Templeton, J. W. 1996a. Molecular phylogeny and in situ detection of the etiologic agent of necrotizing hepatopancreatitis in shrimp. Appl. Environ. Microbiol. 62:3439-3445.

Loy, J. K., Frelier, P.F., Varner, P. and Templeton, J .W. 1996b. Detection of the etiologic agent of necrotizing hepatopancreatitis in cultured Penaeus vannamei from Texas and Peru by polymerase chain reaction. Dis. Aquat. Org. 25: 117-122.

Sindermann, C.J. 1990. Principal Diseases of Marine Fish and Shellfish, Vol. 2, 2nd

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edition. Academic Press, New York.

Vibrio species (vibriosis)

Vibriosis is ubiquitous throughout the world and all marine crustaceans, including prawns, are susceptible. Epizootics occur in all life stages, but are more common in hatcheries. Major epizootics of vibriosis have been reported for P. japonicus from Japan, P. monodon from the Indo-Pacific region and P. vannamei from Ecuador, Peru, Colombia and Central America (Lightner, 1996). Vibriosis may be expressed as a number of syndromes. These include: oral and enteric vibriosis, appendage and cuticular vibriosis, localised vibriosis of wounds, shell disease, systemic vibriosis and septic hepatopancreatitis (Lightner, 1996). Systemic vibriosis of P. vannamei from Ecuador is known as “Sindroma gaviota” and caused massive stock losses during 1989 and 1990 (Mohney et al., 1991). V. alginolyticus and V. harveyi have caused mortalities associated with “Syndrome 93” of P. stylirostris from New Caledonia since 1993 (Costa et al., 1996).

V. penaeicida was one of two dominant bacterial species recovered from prawns with “Syndrome 93” in New Caledonia (Costa et al., 1996b). The syndrome has been induced experimentally (filling the requirements of Koch's postulates) with certain strains of Vibrio penaeicidia (AM23) (Don Lightner, personal communication).

Vibriosis is caused by a number of Vibrio species of bacteria, including: V. harveyi, V. vulnificus, V. parahaemolyticus, V. alginolyticus, V. penaeicida and Vibrio sp. (Brock and Lightner, 1990; Ishimaru et al., 1995). There have been occasional reports of vibriosis caused by V. damsela, V. fluvialis and other undefined Vibrio species. (Lightner, 1996). Vibrio species are part of the natural microflora of wild and cultured prawns (Sinderman, 1990) and become opportunistic pathogens when natural defence mechanisms are suppressed (Brock and Lightner, 1990). They are usually associated with multiple etiological agents. However, some Vibrio species, or strains of certain species, have been identified as primary pathogens (Owens and Hall-Mendelin, 1989; Lavilla-Pitogo et al., 1990; de la Peña a et al., 1995). Pathogenic strains of V. harveyi, V. vulnificus and V. parahaemolyticus have caused massive epidemics in Thailand (Nash et al., 1992) and the Philippines (Lavilla-Pitogo et al., 1990).

Most Vibrio spp. associated with vibriosis exist in Australia, although few major epidemics have been reported. V. penaeicida has not been reported from Australia. Luminescent V. harveyi appears to release exotoxins (Liu et al., 1996) and may cause 80-100% mortality in Australian P. monodon hatcheries (Harris, 1995). V. damsela is a primary pathogen of P. monodon larvae in Australia and causes a septicaemia (Owens and Hall-Mendelin, 1989). A virulent strain of V. harveyi was identified from moribund P. esculentus broodstock held in captivity in Northern Australia (Owens et al., 1992).

V. anguillarum, V. campbelli, V. nereis, V. cholerae (non 01) and V. splendidus have also been reported in association with disease outbreaks in prawns (Chen 1992; Lavilla-Pitoga, 1990; Esteve & Quijada, 1993; Sahul-Hameed et al., 1996).

Clinical signs

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Mortalities due to vibriosis occur when prawns are stressed by factors such as: poor water quality, crowding, high water temperature and low water exchange (Lewis, 1973; Lightner and Lewis, 1975; Brock and Lightner, 1990). High mortalities usually occur in postlarvae and young juvenile prawns. P. monodon larvae suffered mortalities within 48 hr of immersion challenge with strains of V. harveyi and V. splendidus from the Philippines (Lavilla-Pitogo, et al., 1990). Mortalities involving vibriosis have been reported in market sized P. monodon from Malaysia (Anderson et al., 1988). Adult prawns suffering vibriosis may appear hypoxic, show reddening of the body with red to brown gills, reduce feeding and may be observed swimming lethargically at the edges and surface of ponds (Anderson et al., 1988; Nash et al., 1992).

In China Vibrio spp. cause red-leg disease, characterised by red colouration of the pleopods, periopods and gills, in juvenile to adult prawns and may causes mortalities of up to 95% during the warm season (Chen, 1992). Eyeball necrosis diseases also occurs in China and is caused by V. cholerae. The eyeballs of infected prawns turn brown and fall away and mortality occurs within a few days (Chen, 1992). In Japan V. penaeicida is considered to be the most important pathogen of cultured P. japonicus (de la Peña et al., 1995).

Six Vibrio species, including V. harveyi and V. splendidus cause luminescence, which is readily visible at night, in infected postlarvae, juveniles and adults (Ruby et al., 1980; Lightner, et al., 1992). Infected postlarvae may also exhibit reduced motility, reduced phototaxis and empty guts (Chen, 1992).

Gross PathologyPrawns suffering vibriosis may display localised lesions of the cuticle typical of bacterial shell disease, localised infections from puncture wounds, loss of limbs, cloudy musculature, localised infection of the gut or hepatopancreas and/or general septicemia (Lightner, 1993). Lesions of bacterial shell disease are brown or black and appear on the body cuticle, appendages or gills (Sinderman, 1990). Affected postlarvae may display cloudy hepatopancreata (Takahashi et al., 1985a). Gills often appear brown (Anderson et al., 1988). Septic hepatopancreatitis is characterised by atrophy of the hepatopancreas with multifocal necrosis and haemocytic inflammation.

HistopathologyRod-shaped Vibrio spp., 1.5-4.0 x 0.5-1.0 m, are observed in infected organs using histological techniques and usually appear basophilic. (Lavilla-Pitogo et al., 1990). Sloughing of hepatopancreatic and midgut epithelial cells into the gut lumen is common. Cuticular colonisation typically results in the necrosis of the cuticular epithelium and the formation of melanised lesions. Systemic vibriosis typically results in the formation of septic haemocytic nodules in the lymphoid organ, heart and connective tissues of the gills, hepatopancreas, antennal gland, nerve cord, telson and muscle (Anderson et al., 1988; Mohney et al., 1991; Jiravanichpaisal et al., 1994). Infected hepatopancreocytes may appear poorly vacuolated, indicating low lipid and glycogen reserves (Anderson et al., 1988). Vibriosis in P. monodon in Thailand was associated with the formation of “spheroids” in the lymphoid organ (Nash et al., 1992).

Diagnosis

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Diagnosis of vibrio infection is based on clinical signs and the histological demonstration of rod-shaped Vibrio bacteria in lesions, nodules or haemolymph. Excised organs and haemolymph may be streaked on a Vibrio-selective or general marine agar plate. When investigating postlarvae, the whole animal may be crushed and then streaked onto an agar plate. Luminescent colonies may be observed after 12 to18 hr if incubated at room temperature or 25 to 30oC. Vibrio isolates may be identified by a number of methods, including: Gram stain, motility, an oxidase test, mode of glucose utilisation, growth in the presence of NaCl, nitrate reduction and luminescence. Vibrio species may be identified rapidly in the field using the API-20 NFT system which involves culturing vibrio colonies on API-NFT strips and scoring the colonies according to the kit directions (Lightner, 1996) or BIOLOG (a miniaturised bacterial identification system which is an alternative to the API system). Antimicrobial sensitivity tests may be used to identify vibriosis and can be run using the Kirby-Bauer disk method (DIFCO, 1986) or the Minimum Inhibitory Concentration (MIC) method (Lightner, 1996)

TransmissionVibrio species exist in the water used in prawn culture facilities (Lavilla-Pitogo, et al., 1990). Bacteria enter prawns via wounds or cracks in the cuticle and are ingested with food (Paynter 1989; Lavilla-Pitogo et al., 1990). The primary source of V. harveyi in Filipino hatcheries appears to be the midgut contents of female broodstock, which are shed during spawning (Lavilla-Pitogo et al., 1992).

ViabilityNumerous studies have been undertaken concerning the effect of freezing on vibrios which contaminate harvested shellfish. V. vulnificus in harvested oysters (Crassostrea virginica) survived storage at –20oC for 70 days (Parker et al., 1994). V. parahaemolyticus, isolated from homogenates of oyster meat was inactivated within 16 days at –15oC when the bacterial load was very high (10 cfu/gm; Muntada-Garriga et al., 1995). There is recent evidence to suggest that V. harveyi can survive in pond sediment even after chlorination or treatment with lime (Karunasagar et al., 1996).

TreatmentVibriosis is controlled by rigorous water management and sanitation to prevent the entry of vibrios in the culture water (Baticados, et al., 1990) and to reduce stress on the prawns (Lightner, 1993). Good site selection, pond design and pond preparation are also important (Nash et al., 1992). An increase in daily water exchanges and a reduction in pond biomass by partial harvesting are recommended to reduce mortalities caused by vibriosis. Draining, drying and administering lime to ponds following harvest is also recommended (Anderson et al., 1988).

Luminescent vibriosis may be controlled in the hatchery by washing eggs and avoiding contamination by spawner faeces. V. harveyi in the water column may be inactivated by a 30 min exposure to 10 ppm chlorine (Karunasagar et al., 1996). Antibacterials may be administered directly into the water or via medicated feeds (Monhey and Lightner, 1990), although their efficiency is limited because of the possible development of resistant strains and the limited tolerance of prawns larvae (Baticados et al., 1990). Oxytetracycline-medicated feeds were found to be effective in controlling V. penaeicida in Japan (Takahashi et al., 1985b).

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Formalin killed V. penaeicida and other Vibrio spp. have been reported to successfully vaccinate P. japonicus and P. monodon and may be administered by injection, immersion, spraying (Itami and Takahashi, 1989; Teunissen et al., 1996) or by incorporation into micro-encapsulated feeds (Itami et al., 1991).

Immunostimulants have had some success in reducing prawn mortalities associated with vibriosis (Itami, 1996).

Present status of vibriosisVibriosis is a common problem world-wide, particularly in the Philippines where severe epizootics continue (Lavilla-Pitago et al., 1996). V. harveyi continues to cause chronic mortalities of up to 30% among Australian P. monodon larvae and postlarvae under stressful conditions. A highly pathogenic strain of Vibrio sp. (AM 23) has recently been identified in association with Syndrome 93 from New Caledonia and continues to cause mortalities among cultured P. stylirostris (Le Groumellec et al., 1996). Problems caused by secondary vibriosis are common, but are considered minor compared to viral epidemics.

ReferencesAnderson, I.G., Shamsudin, M.N. and Shariff, M. 1988. Bacterial septicemia in

juvenile tiger shrimp, Penaeus monodon, cultured in Malaysian brackishwater ponds. Asian Fis. Sci. 2: 93-108.

Baticados, M.C.L., Lavilla-Pitogo, C.R., Cruz-Lacierda, E.R., de la Pena, L.D. and Sunaz, N.A. 1990. Studies on the chemical control of luminous bacteria Vibrio harveyi and V. splendidus isolated from diseased Penaeus monodon larvae and rearing water. Dis. Aquat. Org. 9: 133-139.

Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne (ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Chen, D. 1992. An overview of the disease situation, diagnostic techniques, treatments and preventatives used on shrimp farms in China. In: W. Fuls and K.L.Main (eds.) Diseases of Cultured Penaeid Shrimp in Asia and the Unites States. The Oceanic Institute, Hawaii. pp. 47-55.

Costa, R., Mermoud, I., Koblavi, S., Haffner, P., Berthe, F., Le Groumellec, M. and Grimont, P. 1996a. Isolation and characterisation of bacteria associated with a Penaeus stylirostris disease (“Syndrome 93”) in New Caledonia. SICCPPS book of abstracts, SEAFDEC, Iloilo City, Philippines. p. 44.

Costa, R., Mermoud, I., Koblavi, S., Haffner, P., Berthe, F., Le Groumellec, M. and Grimont, P. 1996b. Investigation on a disease of Penaeus stylirostris (“Syndrome 93”) in New Caledonia, exploring the hypothesis of a mixed bacterial and viral infection in WAS book of abstracts, Seattle, US p.90.

de la Peña, L.D., Kakai, T., Muroga, K. 1995. Dynamics of Vibrio sp PJ in organs of orally infected kuruma prawn, Penaeus japonicus. Fish. Pathol. 30: 39-45.

Esteve, M. and Quijada, R. 1993. Evaluation of three experimental infection techniques with Vibrio anguillarum in Penaeus brasiliensis in Carillo et al., (ed.).,“From discovery to commercialization” ’93 World Aquaculture, European Aquaculture Society Special publication #19 Torremolinos, Spain p 129

Ishimaru, K., Akarawa-Matsushita, M., Muroga, K. 1995. Vibrio penaeicida sp., nov., a pathogen of kuruma prawns (Penaeus japonicus). Int. J. Syst. Bacteriol. 43:

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8-19.Harris, L. 1995. The involvement of toxins in the virulence of Vibrio harveyi strains

pathogenic to the black tiger prawn Penaeus monodon and the use of commercial probiotics to reduce prawn hatchery disease outbreaks caused by V. harveyi strains. CRC for Aquaculture, Scientific Conference abstract, Bribie Island, Australia.

Itami, T., Takahashi, Y. and Nakamura, Y. 1989. Efficiency of vaccination against vibriosis in cultured kuruma prawns Penaeus japonicus. J. Aquatic Anim. Health 1: 238-242.

Itami, T. and Takahashi, Y. 1991. Survival of larval giant tiger prawns, Penaeus monodon after addition of killed Vibrio cells to a microencapsulated diet. J. Aquat. Anim. Health 3: 151-152.

Itami, T. 1996. Vaccination and immunostimulation in shrimps. SICCPPS book of abstracts, SEAFDEC, Iloilo City, Philippines. p. 50

Jiravanichpaisal, P and Miyazaki, T. 1994. Histopathology, biochemistry and pathogenicity of Vibrio harveyi infecting black tiger prawn Penaeus monodon. J. Aquat. An. Health 6: 27-35.

Karunasagar, I., Otta, S.K. and Karunasagar, I. 1996. Effect of chlorination on shrimp pathogenic Vibrio harveyi. World Aquaculture ’96, book of abstracts. The World Aquaculture Society, Baton Rouge, LA. p. 193.

Lavilla-Pitogo, C.R., Baticados, C.L., Cruz-Lacierda, E.R. and de la Pena, L. 1990. Occurrence of luminous bacteria disease of Penaeus monodon larvae in the Philippines. Aquaculture 91: 1-13.

Lavilla-Pitogo, C.R., Albright, L.J., Paner, M.G. and Sunaz, N.A. 1992. Studies on the sources of luminescent Vibrio harveyi in Penaeus monodon hatcheries. In: M. Shariff, R.P. Subasinghe and J.R. Authur (eds.) Diseases in Asian Aquaculture 1. Fish Health Section, Asian Fisheries Society, Manila, Philippines. pp. 157-164.

Lavilla-Pitogo, C.R., Leano, E.M. and Paner, M.G. 1996. Mortalities of pond-cultured juvenile shrimp, Penaeus monodon, associated with dominance of luminescent bacteria, Vibrio harveyi in the rearing environment. SICCPPS book of abstracts, SEAFDEC, Iloilo City, Philippines. p. 40.

Le Groumellec, M., Goarant, C., Haffner, P., Berthe, F., Costa, R. and Mermoud, I. 1996. Syndrome 93 in New Caledonia: Investigation of the bacterial hypothesis by experimental infections, with reference to stress-induced mortality. SICCPPS book of abstracts, SEAFDEC, Iloilo City, Philippines. p. 46.

Lewis, D.H. 1973. Response of brown shrimp to infection with Vibrio sp. Proc. Wld. Maricult. Soc. 4: 333-338.

Lightner, D.V. 1993. Diseases of cultured penaeid shrimp. In: J.P. McVey (ed.) CRC Handbook of Mariculture, Second edition, Volume 1, Crustacean Aquaculture. CRC Press Inc., Boca Raton, FL. p. 393-486.

Lightner, D.V. (ed.). 1996. A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA, USA.

Lightner, D.V. and Lewis, D.H. 1975. A septicemic bacterial disease syndrome of penaeid shrimp. Mar. Fish. Rev. 37(5-6): 25-28.

Lightner, D.V., Bell, T.A., Redman, R.M., Mohney, L.L., Natividad, J.M., Rukyani, A. and Poernomo, A. 1992. A review of some major diseases of economic significance in penaeid prawns/shrimps of the Americas and Indo-Pacific. In: M. Shariff, R. Subasinghe and J.R. Arthur (eds.) Proceedings 1st Symposium on

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Diseases in Asian Aquaculture. Fish Health Section, Asian Fisheries Society, Manila, Philippines. pp. 57-80.

Liu, P.C., Lee, K.K. and Chen, S.N. 1996. Pathogenicity of different isolates of Vibrio harveyi in tiger prawn, Penaeus monodon. Letters in Applied Microbiology 22: 413-416.

Mohney, L.L. and Lightner, D.V. 1990. Bioencapsulation of therapeutic quantities of the antibacterial Pomet 30 in the nematode Panagrellas redivivus and in nauplii of Artemia salina. J. World. Aquacult. Soc. 21(3): 186-191

Mohney, L.L., Lightner, D.V. and Bell, T.A. 1991. An epizootic due to Vibrio spp. in pond-reared Penaeus vannamei in Ecuador. World Aquaculture Meeting, Book of Abstracts, Puerto Rico, p. 45.

Muntada-Garriga, J.M., Rodriguez-Jerez, J.J., Lopez-Sabater, E.I. and Mora-Ventura, M.T. 1995. Effect of chill and freezing temperatures on survival of V. parahaemolyticus inoculated in homogenates of oyster meat. Letters in Applied Microbiology 20: 225-227.

Nash, G. Nithimathachoke, C., Tungmandi, C., Arkarjamorn, A., Prathanpipat, P. and Ruamthaveesub, P. 1992. Vibriosis and its control in pond-reared Penaeus monodon in Thailand. In: M. Shariff, R.P. Subasinghe and J.R. Authur (eds.) Diseases in Asian Aquaculture 1. Fish Health Section, Asian Fisheries Society, Manila, Philippines. pp. 143-155.

Owens, L. and Hall-Mendelin, 1989. Recent Advances in Australian prawns (sic) diseases and pathology. Advances in Tropical Aquaculture, Tahiti, AQUACOP, IFREMER, Actes de Colloque 9: 103-112.

Owens, L., Muir, P., Sutton, D. and Wingfield, M. 1992. The pathology of microbial diseases in tropical Australian Crustacea. In: M. Shariff, R.P. Subasinghe and J.R. Authur (eds.) Diseases in Asian Aquaculture 1. Fish Health Section, Asian Fisheries Society, Manila, Philippines. pp. 165-172.

Parker, R.W., Maurer, E.M., Childers, A.B. and Lewis, D.H. 1994. Effect of frozen storage and vacuum packing on survival of V. vulnificus in Gulf Coast Oysters (C. virginica). J. Food Protection 57(7): 604-606.

Paynter, J.L. 1989. Invertebrates in Aquaculture. Refresher Course for Veterinarians, Proceedings 117. The University of Queensland.

Pizzutto, M and Hirst, R.G. 1995. Classification of isolates of Vibrio harveyi virulent to Penaeus monodon larvae by protein profile analysis and M13 DNA fingerprinting. Dis. Aquat. Org. 21: 61-68.

Ruby, E.G., Greenberg, E.P. and Hastings, J.W. 1980. Planktonic marine luminous bacteria: species distribution in the water column. Applied and Environmental Microbiology 39: 302-306.

Sahul Hameed, A.S., Rao, P.V., Farmer, J.J., Hickman-Brenner, W. and Fanning, G.R. 1996. Characteristics and pathogenicity of a Vibrio cambelli-like bacterium affecting hatchery-reared Penaeus indicus (Milne Edwards, 1837) larvae. Aquacult. Res. 27, 853-863.

Sindermann, C.J. 1990. Principal Diseases of Marine Fish and Shellfish, Vol. 2, 2nd edition. Academic Press, New York.

Takahashi, Y. Shimoyama, Y and Monoyama, K. 1985a. Pathogenicity and characteristics of Vibrio sp. isolated from diseased postlarvae of kuruma prawn, Penaeus japonicus Bate. Bull. Jpn. Soc. Sci. Fish. 51: 721-730.

Takahashi, Y. Itani, T., Nakagawa, A., Nishimura, H. and Abe, T. 1985b.

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Therapeutic effects of oxytetracycline trial tablets against vibriosis in cultured kuruma prawns Penaeus japonicus Bate. Bull. Jp. Soc. Sci. Fish. 51: 1639-1644.

Teunissen, O.S.P., Boon, J.H., Latscha, T. and Faber, R. 1996. Effect of vaccination on vibriosis resistance in the giant black tiger shrimp Penaeus monodon (Fabricius). SICCPPS book of abstracts, SEAFDEC, Iloilo City, Philippines. p. 51.

Rickettsia

The importance of rickettsia and rickettsia-like bacteria as prawn pathogens is not fully known as they usually occur in association with other disease agents such as Gram-negative bacteria, viruses and algal and protozoan epicommensal fouling organisms. Rickettsial infections have been reported in wild-captive P. marginatus from Hawaii (Brock et al., 1986), wild and cultured P. monodon from Malaysia and Indonesia (Anderson, et al., 1987; Lightner, et al., 1992) and cultured P. merguiensis from Singapore (Chong and Loh, 1984). One case of rickettsial infection has been reported in cultured P. vannamei from Mexico (Lightner, 1996). Experimental infection of P. stylirostris with rickettsia from P. marginatus resulted in disease and mortality (Brock et al., 1986). ). Stained prawn disease (SPD) of wild Pandalus platyceros from British Colombia is caused by a rickettsia-like organism (Bower et al., 1996). Rickettsias infect juvenile to adult prawns.Rickettsial infections have not been reported from Australian penaeids.

Rickettsial infection of the connective tissues of freshwater crayfish has been reported in Australia (Owens et al., 1992).

Clinical signsRickettsial infections are usually asymptomatic, however prawns with heavy infections may appear lethargic, stunted, dark in colour, feed poorly and may congregate along the edges of ponds (Anderson et al., 1987). Prawns affected by SPD show black colouration of the cuticle (Bower, et al., 1996). Rickettsias may cause moderate mortalities among populations of cultured prawns (Chang and Loh, 1984; Anderson et al., 1987).

Gross PathologyThe hepatopancreas of infected prawns may appear white or stippled black (Brock et al., 1986; Bower, et al., 1996). The abdominal muscles of P. monodon infected with a systemic rickettsia appeared opaque and white nodules were seen on the midgut wall (Anderson et al., 1987).

HistopathologyRickettsial bacteria are intracellular, rod-shaped and 0.2-0.7 um x 0.8-1.6 um in size (Brock et al., 1986). The rickettsia which occurs in P. monodon is systemic and infects the connective tissues, fixed phagocytes, antennal gland epithelium cells, lymphoid organ sheath cells, hepatopancreas, gill filaments, heart, nervous tissue, tegmental gland and the outer layers of the fore and mid-gut (Anderson et al., 1987). Within these tissues, inflammatory lesions, melanised granulomas and areas of cellular necrosis may be formed. The most severe changes are in the lymphoid organ where normal tissue structure is totally disrupted.

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The rickettsias which infect other penaeid species are restricted to the hepatopancreas. Within hepatopancreatic epithelial cells they form microcolonies which replace the cytoplasm and cause hypertrophy of the epithelial cells (Brock et al., 1986). The nuclei of infected cells may be pyknotic or karyorrhectic. Severe destruction of the hepatopancreas may cause haemocyte infiltration and encapsulation. In P. monodon microcolonies occur within large cytoplasmic vacuoles in the connective tissue cells of the lymphoid organ, gills, hepatopancreas, nerve cord, antennal gland and muscle (Anderson et al., 1987).

DiagnosisDiagnosis is based on the demonstration of rickettsial microcolonies, 5-50 m in diameter, within the cytoplasm of target cells (Brock, 1988). These may be observed in wet mounts of tissue or in Giemsa-stained impression smears. Microcolonies are Feulgen’s positive, basophilic and Gram-negative (Anderson et al., 1987). Steiner’s silver stain and Machiavello’s stain also enable rickettsias to be observed (Brock et al., 1986; Bower, et al., 1996). Infection may be confirmed by the demonstration of rickettsias by TEM.

TransmissionAspects of the biology of rickettsias and rickettsial-like organisms, such as transmission, viability in the environment and host range are not really known. The rickettsial-like organism which causes SPD is transmitted by cannibalism and via the water (Bower et al., 1996).

TreatmentRickettsias may be treated using medicated feeds (Anderson et al., 1987). The spread of rickettsial infection may be controlled by sound management practices, such as destroying infected stocks and disinfecting contaminated ponds, tanks and equipment.

ReferencesAnderson, I.G., Shariff, M., Nash, G., Nash, M. 1987. Mortalities of juvenile shrimp,

Penaeus monodon, associated with Penaeus monodon baculovirus, cytoplasmic reo-like virus, and rickettsial and bacterial infections, from Malaysian brackishwater ponds. Asian Fish. Sci 1: 47-64.

Bower, S.M., Meyer, G.R. and Boutillier, J.A. 1996. Stained prawn disease (SPD) of Pandalus platyceros in British Columbia, Canada, caused by a rickettsial infection. Dis. Aquat. Org. 24: 41-54.

Brock, J.A. 1988. Rickettsial infection in penaeid shrimp. In: C.J. Sindermann and D.V. Lightner (eds.) Disease Diagnosis and Control in North American Marine Aquaculture. Elsevier, Amsterdam. pp. 38-41.

Brock, J.A., Nakagawa, L.K., Hayashi, T., Teruya, S. and van Campen, H. 1986. Hepatopancreatic rickettsial infection of the penaeid shrimp, Penaeus marginatus Randall from Hawaii. J. Fish. Diseases 9: 73-77.

Chong, Y.C. and Loh, H. 1984. Hepatopancreas chlamydial and parvoviral infections of farmed and marine prawns in Singapore. Singapore Veterinary Journal 9: 51-56.

Lightner, D.V. (ed.). 1996. A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society,

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Baton Rouge, LA, USA. Lightner, D.V., Bell, T.A., Redman, R.M., Mohney, L.L., Natividad, J.M., Rukyani,

A. and Poernomo, A. 1992. A review of some major diseases of economic significance in penaeid prawns/shrimps of the Americas and Indo-Pacific. In: M. Shariff, R. Subashnghe and J.R. Arthur (eds.) Proceedings 1st Symposium on Diseases in Asian Aquaculture. Fish Health Section, Asian Fisheries Society, Manila, Philippines. pp. 57-80.

Owens, L., Muir, P., Sutton, D. and Wingfield, M. 1992. The pathology of microbial diseases in tropical Australian crustacea. In: M. Shariff, R.P. Subasinghe and J.R. Authur (eds.) Diseases in Asian Aquaculture 1. Fish Health Section, Asian Fisheries Society, Manila, Philippines. pp. 165-172

Aerococcus viridans var. homariGaffkemia, caused by Aerococcus viridans var. homari, is an acute or chronic, almost invariably fatal disease of impounded American and European lobsters (Homarus americanus and H. gammarus). The causative agent is a Gram-positive, non-motile, catalase negative coccus. The tetrad forming coccus is beta-haemolytic and facultative anaerobic. Strain differences occur and not all strains are lethal to lobsters (Steenberger et al., 1977). The disease has caused substantial economic loss to the lobster trade in the US, Canada, Holland, France, Ireland and England. The impact of Aerococcus viridans var. homari on wild lobster populations is undefined.

Lobster populations along the North American and European Atlantic coastlines and other decapod crustaceans are natural reservoirs for Aerococcus viridans var. homari. The bacterium is apparently not part of the epiflora of lobsters (Stewart, 1980).

Clinical signsLobsters dying of gaffkemia are extremely weak and die in a ‘spread-eagle’ position or lateral recumbency. Lobsters at an advanced stage of the disease may show pink discolouration of of the ventral abdomen and the haemolymph is thin and pink. The time course of gaffkemia in lobsters is strongly temperature dependent. The bacterium has also been reported as the cause of a low-incidence infection of other decapods in nature, including P. aztecus (Stewart and Rabin, 1970).

Experimental infection of other decapods, incuding Pandalus platyceros, resulted in only mild or no disease; where death occurred it was after a prolonged incubation period (Rabin and Hughes, 1968).

Gross PathologySmall black specks due to haemocyte aggregations may be noticeable in the gills and other tissues late in the course of infection.

HistopathologyThere is an absence of specific cytopathology.

DiagnosisThe coccus grows on a range of media at an optimal incubation temperature of 300C.

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ViabilityAerococcus viridans var. homari grows on media at 6 to 440C. The bacterium survives well in seawater and can be recovered from marine sediments (Stewart and Rabin, 1970) and the surfaces of lobster tanks (Wood, 1965).

TransmissionInfection occurs through breaks in the cuticle. Aerococcus viridans var. homari is not transmitted by ingestion as the stomach acidity destroys the bacteria. Injection of virulent Aerococcus viridans var. homari into lobsters is usually fatal within 14 days. The minimum infectious dose via injection into the haemocoel is 5 bacteria.

TreatmentAn effective vaccine to protect the American lobster from gaffkemia has been developed (Keith, 1992). The bacterium is susceptible to tetracycline, penicillin, erythromycin, novobiocin, vancomycin.

Present status of diseaseManagement of gaffkemia in captive lobsters is primarily a matter of good husbandry. The devastating outbreaks of the past should be preventable by ensuring that wounding and crowding of lobsters does not occur.

ReferencesBrock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne

(ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Keith, I.R., Paterson, W.D., Airdrie, D. and Boston, L.D. 1992. Defence mechanisms of the American lobster, (Homarus americanus): vaccination provided protection against gaffkemia infections in laboratory and field trials. Fish Shellfish Immunol. 2: 109-119.

Rabin, H. and Hughes, J.T. 1968. Studies on host-parasite relationships in gaffkemia J. Invert. Pathol. 10; 335-344.

Steenbergen JF, Kimball, H.S., Low, D.A. Scapiro, H.C. and Phelps, L.N. 1977. Serological grouping of virulent and avirulent strains of the lobster pathogen Aerococcus viridans. J. Gen. Microbiol. 99: 425-30.

Stewart, J.E. 1980 Diseases Lobsters, fungal and microbial infections, gaffkemia, parasites, shell disease. in Biol. Manage. Lobster. New York, Academic Press. v. 1 p. 301-342..

Stewart JE, and Rabin, H 1970 Gaffkemia, a bacterial disease of lobsters (genus Homarus) In SF Snieszko (Ed.) A symposium on diseases of fishes and shellfishes American fisheries society Washington DC .pp.. 431-439

Wood, P.C. 1965 A preliminary note on gaffkemia investigations in England. Rapp. P. v. Reun. Cons. perm. int. Explor. Mer. 156; 30-34.

Parasites

MicrosporidiaCotton shrimp, or milk shrimp is caused by three genera of microsporidians:

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Agmasoma (Thelohania), Amesoma (Nosema) and Pleistophora (Plistophora). These microsporidians are ubiquitous in wild and cultured penaeids and infect juveniles and adults. Most microsporidian infections have been reported from the Americas (Iversen and Manning, 1959; Baxter et al., 1970; Feigenbaum, 1975). However, microsporidians have also been reported in cultured Penaeus monodon from the Philippines (Enriques, 1982), Malaysia (Anderson et al., 1989) and south-eastern Thailand (Flegel et al., 1992a) in Australia (Bergin, 1986; Owens and Glazebrook, 1988).

Agmasoma (Thelohania) penaei infects P. mergiuensis as well as P. monodon from south-eastern Thailand (Flegel et al., 1992a). In northern Australia, Agmasoma sp. infects wild juvenile and adolescent P. esculentus, P.semisulcatus and P. merguiensis, while Thelohania. sp. infects wild P. latisculatus, P. longistylus and P. semisulcatus (Owens and Glazebrook, 1988). Although primarily a problem in wild prawns in Australia, microsporidiosis has stopped production on at least one farm in north Queensland (Bergin, 1986).

Clinical signsMost microsporidians infect and replace striated muscle, causing a characteristic opaque white abdomen. The more common species also infect several other tissue types including gonad, connective tissues, and hepatopancreatic epithelial cells (Lightner, 1996). The cuticle of infected prawns may appear dark blue/black (Brock and Lightner, 1990). Microsporidians are not considered to be of great economic significance in comparison to viruses and bacteria, as prevalence of infection typically reaches only 10-20% in cultured populations and mortality is not typical (Lightner, 1993). Infected prawns are more prone to predation and vulnerable to environmental stresses (Lightner, 1988). Infected prawns do not alter their behaviour, however, wild penaeids tend to remain in estuaries to reproduce rather than migrate off shore (Overstreet, 1973).

Gross PathologyAgmasoma penaei infects blood vessels, heart, gonads, gills, hepatopancreas, gut and connective tissues, as well as muscle. Infected gonads appear white and hypertrophied while multiple, white tumour-like swellings may be formed in the gills and subcuticular tissues (Rigdon et al., 1975; Kelly, 1979). Wild-caught broodstock infected with A. penaei become sterile when the ovaries are ultimately destroyed (Kelly, 1979). The other species of microsporidians which infect prawns (Ameson nelsoni, Nosema sp., Thelohania duorara and Thelohania sp.) are largely restricted to striated muscle fibres. Pleistophora sp. infection does not necessarily result in widespread lysis of muscle fibres as occurs in infection with other species and may also infect the heart, gills, foregut and hepatopancreas (Kelly, 1979)

HistopathologyMicrosporidians multiply to produce spores within the cytoplasm of infected cells (Anderson et al., 1989). The tubules of infected hepatopancreata become dilated and necrotic. The lumen of the hepatopancreatic tubule may contain cellular debris and shed spores and the epithelium of the hepatopancreas may be replaced by hemocytic encapsulation. Infections in striated muscle do not invoke a host inflammatory response (Lightner, 1996). The same species of microsporidian may infect different tissues in different species of prawn (Overstreet, 1973).

49

DiagnosisMicrosporidian spores (1-8 m) may be demonstrated in unstained wet mounts or impression smears of infected tissues using light microscopy. Microsporidians may be distinguished by the size of spores, the number of spores produced per sporont and the number of turns made by the polar filament (Iversen et al., 1987; Lightner, 1996) (polar filament coil numbers can only be determined by TEM). Impression smear and histological sections must be stained with Giemsa or acid-fast stains to effectively observe these distinguishing features (Lightner, 1996). A DNA probe and PCR detection method have been developed for a Thai strain of Agmasoma (Thelohania) sp. (Pasharawipas and Flegel, 1994; Pasharawipas et al., 1994).

TransmissionMicrosporidiosis was transmitted experimentally to P. duorarum postlarvae by feeding prawns on the faeces of spotted sea trout (Cynoscion nebulosis) which had been fed infected prawns (Iversen and Kelly, 1976). This suggests that prawns may be infected by microsporidian spores released in the faeces of marine animals such as finfish which act as “conditioning” or intermediate hosts. It has been suggested that Agmasoma penaei occurs only in farms on the south-west Gulf of Thailand due to the absence of intermediate hosts on the south-eastern side of the Gulf (Flegel et al., 1992b). Environmental conditions, such as high rainfall may play a role in the epidemiology of this disease (Flegel et al., 1992a).

Transmission was unsuccessful when prawns were fed directly on infected prawn muscle and when they were exposed to water-borne spores (Iversen and Kelly, 1976; Flegel et al., 1992a). Flegel et al. (1992a) observed that Agmasoma penaei was not transmitted horizontally in adolescent P. monodon and that vertical transmission from broodstock to eggs was unlikely. Owens and Glazebrook (1988) found that freezing prawns damages the microsporidian sporocyte wall.

TreatmentEffective methods of treating microsporidiosis have not been developed. The exclusion of finfish from prawn culture ponds may prevent infection (Iversen and Kelly, 1976). However this is difficult on a commercial scale (Flegel et al., 1992a).

Present status of diseaseMicrosporidiosis is not considered a major disease of cultured prawns. However opacity of abdominal musculature is a marketing problem (Owens and Glazebrook, 1988).High prevalence rates of microsporidiosis in wild prawn populations have been reported and linked to serious impacts on commercial fisheries (Lightner, 1996).

ReferencesAnderson, I.G., Shariff, M. and Nash, G. 1989. A hepatopancreatic microsporidian in

pond-reared tiger shrimp, Penaeus monodon, from Malaysia. J. Invertebr. Pathol. 53: 278-280.

Baxter, K.N., Rigdon, R.H. and Hanna, C. 1970. Pleistophora sp. (Microsporidia: Nosematidae): a new parasite of shrimp. J. Invertebr. Pathol. 16: 289-291.

Bergin, T.J. 1986. An overview of aquaculture and disease control . In: J.D. Humphrey and J.S. Langdon (eds.) Proc. Workshop Dis. Aust. Fish Shellfish. pp.

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3-9.Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne

(ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Fiegenbaum. D.L. 1975. Parasites of the commercial shrimp Penaeus vannamei Boone and Penaeus brasiliensis Latreille. Bull. Mar. Sci. 25: 491-514.

Flegel, T.W., Fegan, D., Kongsom, S., Vuthikornudomkit, S., Sriurairatana, S., Boonyaratpalin, S., Chantanachookin, C., Vickers, J. and MacDonald, O. 1992. Occurrence, diagnosis and treatment of shrimp diseases in Thailand. In: W. Fulks and K. Main (eds.). Diseases of Cultured Penaeid Shrimp in Asian and the United States. The Oceanic Institute, Honolulu, HI. pp. 57-112.

Fulks,W. and Main, K.L. 1992. Diseases in Cultured Penaeid Shrimp in Asian and the United States. The Oceanic Institute, Honolulu, HI. (Preface).

Iversen, E.S. and Manning, R.B. 1959. A new microsporidian parasite from the pink shrimp (Penaeus dourarum). Transaction Am. Fish. Soc. 88: 130-132.

Iversen, E.S. and Kelly, J.F. 1976. Microsporidiosis successfully transmitted experimentally in pink shrimp. J. Invertebr. Pathol. 27: 407-408.

Iversen, E.S., Kelly, J.F. and Alzamora, D. 1987. Ultrastructure of Thelohania dourara. J. Fish Dis. 10: 299-307.

Kelly, J.F. 1979. Tissue specificities of Thelohania dourara, Agmesoma penaei and Pleistophora sp., microsporidian parasites of pink shrimp, Penaeus douraram. J. Invertebr. Pathol. 33: 331-339.

Lightner, D.V. 1988. Diseases of cultured penaeid shrimp and prawns. In: C.J. Sindermann and D.V. Lightner (eds.) Disease Diagnosis and Control in North American Marine Aquaculture. 2nd edition. Elsevier, New York. pp. 8-127.

Lightner, D.V. 1993. Diseases of penaeid shrimp. In: McVey, J.P. (ed.) CRC Handbook of Mariculture: Crustacean Aquaculture. 2nd edition. CRC Press, Boca Raton, FL. pp. 393-486.

Lightner, D.V. (ed.) 1996. A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA., USA .

Owens, L. and Glazbrook, J.S. 1988. Microsporidian infections in commercial prawns from northern Australia. Aust. J. Mar. Freshwat. Res. 39: 301-305.

Overstreet, R.M. 1973. Parasites of some penaeid shrimp with emphasis on reared hosts. Aquaculture 2: 105-140.

Pasharawipas, T. and Flegel, T.W. 1994. A specific DNA probe to identify the intermediate host of a common microsporidian parasite of Penaeus merguiensis and P. monodon. Asian Fisheries Science 7: 157-167.

Pasharawipas, T., Flegel, T.W., Chaiyaro, S., Mongkolsuk, S. and Sirisinha, S. 1994. Comparison of amplified gene sequences from microsporidian parasites (Agmasoma or Thelohania) in Penaeus merguiensis and P. monodon. Asian Fisheries Science 7: 169-178.

Rigdon, R.H., Baxter, K.N. and Benton, R.C. 1975. Hermaphroditic white shrimp Penaeus setiferus, parasitized by Thelohania sp. Trans. Am. Fish. Soc. 104: 292-295.

Hematodinium-like organismA hematodinium-like protozoan infects wild Pandalus platyceros and Pandalus

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borealis in British Colombia and Alaska. Infected prawns have chalky tail musculature and a white fluid which contains the vegetative stages of the parasite (Meyers et al., 1994). This hematodinium-like parasite of prawns is considered to be very different to Hematodinium sp. which causes serious disease in Tanner crabs in the USA. The distribution and prevalence of Hematodinium-like organism in prawns is not fully known. However, prevalence within wild populations is relatively low. This parasite has not been reported in Australia and is not considered an important pathogen of prawns (Meyers et al., 1994).

Hematodinium-like organisms cause clinical signs in up to 10% of wild populations of Pandalus species and subclinical infections in up to 27% of prawns from these same populations (Bower & McGladdery, 1998; Bower et al., 1994). Infected prawns do not survive capture (Bower et al., 1994).

ReferencesBrock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne

(ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Bower, S.M., McGladdery, S.E. and Price, I.M. 1994. Synopsis of diseases and parasites of shellfish. In: M. Faisal and F.M. Hetrick (eds.) Annual Review of Fish Diseases. Vol. 4.

Bower, S.M., and McGladdery, S.E. 1998. Synopsis of diseases and parasites of shellfish. Fisheries and Ocean Canada (http://www.pac.dfo.ca/pac/sealane/aquac/pages/hemorgsp.htm).

Meyers, T.R., Lightner, D.V. and Redman, R.M. 1994. A dinoflagellate-like parasite in Alaskan spot shrimp Pandalus platyceros and pink shrimp P. borealis. Dis. Aquat. Org. 18: 71-76.

Overstreet, R.M. 1973. Parasites of penaeid shrimp with emphasis on reared hosts. Aquaculture 2: 105-140.

Owens, L. 1990. Maricultural considerations of the zoogeography of parasites from prawns in tropical Australia. J. Aqua. Trop. 5: 35-41.

Parauronema spp.The ciliate Parauronema sp. invades the haemocoel of protozoeal, mysid and juvenile stages of the brown shrimp (P. aztecus) and has been reported in association with high mortality at a commercial hatchery (Couch, 1978). Ciliates in haemolymph causes mechanical injury by replacing and dislodging tissues and may become numerous enough to fill entire haemocoel and abdomen (Bower & McGladdery, 1996).

ReferencesBower, S.M., McGladdery, S.E. and Price, I.M. 1994. Synopsis of diseases and

parasites of shellfish. In: M. Faisal and F.M. Hetrick (eds.) Annual Review of Fish Diseases. Vol. 4.

Bower, S.M., and McGladdery, S.E. 1998. Synopsis of diseases and parasites of shellfish. Fisheries and Ocean Canada (http://www.pac.dfo.ca/pac/sealane/aquac/pages/cildsp.htm).

Couch, J.A. 1978. Diseases, parasites, and toxic responses of commercial penaeid

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shrimps of the Gulf of Mexico and south Atlantic coasts of North America Fish. Bull. 76: 1-44.

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Section 2 Disease agents which will not be further considered in the IRA

Viruses

Lymphoid Organ Vacuolization Virus (LOVV)

LOVV is a togavirus that occurs in cultured P. vannamei in the Americas and Hawaii (Bonami et al., 1992). LOVV may also infect P. stylirostris (Lightner, 1996). It is not known if LOVV infects Asian and Australian penaeids.

Gross PathologyLOVV does not cause serious disease and there is no gross pathology associated with infection (Bonami et al., 1992).

HistopathologyLOVV causes necrosis of the sheath cells of the lymphoid organ and the formation of “spheroids” or lesions. Cells within these lesions may contain hypertrophied nuclei with diminished chromatin, pyknotic nuclei, cytoplasmic inclusions and a highly vacuolated cytoplasm. LOVV virus particles average 30 nm in diameter and form cytoplasmic masses which tend towards paracrystalline arrays (Bonami et al., 1992).

DiagnosisDiagnosis of LOVV is based on histopathology of the lymphoid organ and the demonstration of viral particles by electron microscopy.

TransmissionLittle is known of the biology of LOVV, including mode of transmission, potential carrier hosts and the viability of virus in the external environment

Considerations for risk assessmentLOVV appears to have a restricted distribution and does not cause disease. The susceptibility of Australian penaeids to infection is not known.

If LOVV were to behave the same way in Australia as overseas, it is unlikely to cause significant production loss in prawns if it were introduced and became established.

Removal of prawn heads prior to import would reduce the chance of introduction of LOVV to Australia, as infection is restricted to the lymphoid organ.

ReferencesBonami, J.R., Lightner, D.V., Redman, R.M. and Poulos, B.T. 1992. Partial

characterization of a togavirus (LOVV) associated with histopathological changes of the lymphoid organ of penaeid shrimps. Dis. Aquat. Org. 14: 145-152.

Lightner, D.V. 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA.,USA.

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REO-III AND REO-IV

REO-III has been reported in cultured P. japonicus from Japan, France (Tsing and Bonami, 1986) and Hawaii (Lightner et al., 1984); cultured P. monodon from Malaysia (Nash et al., 1988); and cultured P. vannamei from Mississippi (Krol et al., 1990) and Ecuador.

REO-IV has been reported from cultured and wild P. chinensis from the Yellow Sea region of Asia (Lightner, 1996).

Clinical signs and gross pathologyREO infections commonly occur in prawns with multiple infection by other viral, bacterial, parasitic and/or fungal pathogens. Therefore gross signs of REO infection are not known and the significance of REO viruses as pathogens is speculative. Prawns infected with REO display non-specific signs of poor health, such as poor growth rate, anorexia, lethargy, eroded and melanised appendages, opaque musculature, shell disease lesions and gill and surface fouling. The hepatopancreas may appear pale and atrophied (Anderson et al., 1987; Nash et al., 1988; Krol et al., 1990). Infected P. japonicus may appear red in colour and cease burying in the sand substrate (Tsing and Bonami, 1986). P. monodon concurrently infected with MBV, rickettsia and gram negative bacteria, gathered around the edges of ponds and the water surface (Nash et al., 1988). Mortalities of 5 to 95% accompanied these signs.

HistopathologyThe histopathology of REO is unclear due to concurrent infection with other agents. REO forms cytoplasmic inclusion bodies in the F or R cells of the hepatopancreas. REO virions are unenveloped, paraspherical and average 50 nm in diameter (Krol et al., 1990). REO is associated with gut and nerve syndrome (GNS), which is characterised by hypertrophy of the basement membrane of the anterior midgut mucosa, atrophy of the hepatopancreas and hyperplasia of the epineurium of the ventral nerve cord in the gnathothorax (Lightner, 1988; 1996).

DiagnosisConclusive diagnosis is base on the demonstration of REO virus particles by transmission electron microscopy. Clinical signs and histopathology are generally confused by multiple infection by other agents.

TransmissionThe biology of REO-III and IV is unknown. Mode of transmission, range of susceptible species, viability of virus in the external environment and significance as a pathogen have yet to be determined.

PreventionAdequate treatment and control measures for REO have not been established.

ReferencesAnderson, I.G., Shariff, M., Nash, G. and Nash, M. 1987. Mortalities of juvenile

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shrimp, Penaeus monodon, associated with Penaeus monodon baculovirus, cytoplasmic reo-like virus and rickettsial and bacterial infections, from Malaysian brackishwater ponds. Asian Fisheries Society 1: 47-64.

Krol, R.M., Hawkins, W.E. and Overstreer, R.M. 1990. Reo-like virus in white shrimp Penaeus vannamei (Crustacea: Decapoda): co-occurrence with Baculovirus penaei in experimental infections. Dis. Aquat. Org. 8: 45-49.

Lightner, D.V. 1988. Diseases of cultured penaeid shrimp and prawns. In: C.J. Sindermann and D.V. Lightner (eds.) Disease Diagnosis and Control in North American Marine Aquaculture, 2nd. ed. Elsevier, New York. pp. 8-127.

Lightner, 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA.,USA

Lightner, D.V., Redman, R.M., Bell, T.A. and Brock, J.A. 1994. An idiopathic proliferative disease syndrome of the midgut and ventral nerve in the Kuruma prawn, Penaeus japonicus Bate, cultured in Hawaii. J. Fish Diseases 7: 183-191.

Nash, M., Nash, G., Anderson, I.A. and Shariff, M. 1988. A reo-like virus observed in the tiger prawn Penaeus monodon Fabricius, from Malaysia. J. Fish Diseases 11: 531- 535.

Tsing, A. and Bonami, J.R. 1987. A new virus disease of the tiger shrimp Penaeus japonicus Bate. J. Fish Diseases 10: 139-141.

Hepatopancreatic Parvo-Like Virus (HPV)

HPV was identified simultaneously in four cultured populations of P. semisulcatus and P. merguiensis in Asia (Lightner and Redman, 1985). HPV has been documented from 6 other species: P. monodon, P. esculentus, P. indicus, P. chinensis, P. penicillatus and P. vannamei. HPV occurs in cultured and wild penaeids in Australia (Paynter et al., 1985; Roubal et al., 1989), Africa, the Americas (Lightner and Redman, 1992), Israel (Colorni et al., 1987), Thailand (Flegel, 1997) and Kuwait (Lightner, 1996) and well as in Asia (Lightner and Redman, 1985). An HPV-like virus has been described for Macrobrachium rosenbergii from Malaysia (Anderson et al., 1990).

Clinical signs and gross pathologyHPV infections have been linked to disease, however, they are often accompanied by other hepatopancreatic pathogens. HPV may be a serious pathogen of younger life stages of prawns, where difficulty in diagnosis has caused HPV to be overlooked (Lightner et al., 1993). Signs of disease in individual prawns are not specific to HPV and include reduced growth, reduced preening, muscle opacity and hepatopancreas atrophy. HPV appears to be directly associated with runted Penaeus monodon in culture ponds in Thailand (Tim Flegel, personal communication). Cumulative HPV-associated mortality was reported to be 50-100% after 4-8 weeks in juvenile P. merguiensis (Lightner and Redman, 1985).

HistopathologyHPV infects the epithelial cells of the hepatopancreas. Virus particles are 22-24 nm in diameter and occur within intranuclear inclusion bodies (IB’s) composed of electron dense, finely granular material. IB’s are basophilic when fully formed and cause

56

lateral displacement of the nucleolus and margination of chromatin. Cells most commonly infected are the distal E or F-cells (Lightner, 1988; Brock and Lightner, 1990).

DiagnosisDefinitive diagnosis is dependant on the demonstration of basophilic IB’s within cells of the hepatopancreas, using histochemical techniques for light and electron microscopy. A rapid field test for HPV has been developed and involves fixing fresh smears of hepatopancreas and staining with Giemsa (Lightner et al., 1993). A diagnostic DNA probe and PCR primers for HPV are available from DiagXotics, Wilton CT and may be used to detect asymptomatic infections. These were developed by Mari et al. (1995).

TransmissionThe mode of transmission of HPV is not fully understood as it has not been transmitted experimentally. Evidence exists that HPV is transmitted vertically from broodstock to progeny and horizontally during the postlarvae stages (Brock and Lightner, 1990). In two studies on captured Thai broodstock specimens (Flegel et al. 1992; Flegel et al. 1997), none were found to show the characteristic histopathology of HPV. This suggests that the virus may not originate with the broodstock but with some other carrier in the cultivation system.

ViabilityThe viability of HPV in the external environment is not known.

Prevention HPV can be controlled by management practices involving avoidance (Lightner, 1988) such as: avoiding contaminating eggs with spawner faeces; separating equipment used in the hatchery from that used in the spawning area; and disinfecting equipment and tanks between batches of postlarvae.

References

Anderson, I.G., Law, A.T., Shariff, M. and Nash, G. 1990. A parvo-like virus in the giant freshwater prawn, Macrobrachium rosenbergii. J. Invertebr. Pathol. 55: 447-449.

Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne (ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Colorni, A., Samocha, T. and Colorni, B. 1987. Pathogenic viruses introduced into Israeli mariculture systems by imported penaeid shrimp. Bamidgeh 39: 21-28.

Flegel, T.W., D.F. Fegan, Sumana Kongsom, Sompoach Vuthikornudomkit, Siriporn Sriurairatana, Sitdhi Boonyaratpalin, Chaiyuth Chantanachookhin, Joan E. Vickers and O.D. MacDonald 1992. Occurrence, diagnosis and treatment of shrimp diseases in Thailand. Diseases of penaeid shrimp. In: W. Fulks and K.L. Main (eds.), Diseases of cultured penaeid shrimp in Asia and the United States, Oceanic Institute, Honolulu, Hawaii, p. 57-112.

Flegel, T.W., Siriporn Sriurairatana, D.J. Morrison and Napaa Waiyakrutha. 1997. Penaeus monodon captured broodstock surveyed for yellow-head virus and other pathogens by electron microscopy. In: T.W. Flegel and P. Menasveta (eds) Shrimp

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biotechnology. National Center for Genetic Engineering and Biotechnology, Bangkok. 37-43.

Lightner, D.V. 1988. Diseases of cultured penaeid shrimp and prawns. In: C.J. Sindermann and D.V. Lightner (eds.) Disease Diagnosis and Control in North American Marine Aquaculture, 2nd. ed. Elsevier, New York. pp. 8-127

Lightner, 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA.,USA

Lightner, D.V. and Redman, R.M. 1985. A parvo-like virus disease of penaeid shrimp. J. Invertebr. Pathol. 45: 47-53.

Lightner, D.V. and Redman, R.M. 1992. Geographic distribution, hosts and diagnostic procedures for the penaeid virus diseases of concern to shrimp culturists in the Americas. In: A.W. Fast and L.J. Lester (eds.) Culture of Marine Shrimp: Principals and Practices. Elsevier, Amsterdam. pp. 573-592.

Lightner, D.V., Redman, R.M., Moore, D.W. and Park, M.A. 1993. Development and application of a simple and rapid diagnostic method to studies on hepatopancreatic parvovirus of penaeid shrimp. Aquaculture 116: 15-23.

Mari, J., D.V. Lightner, B.T. Poulos and J.R. Bonami. 1995. Partial cloning of the genome of an unusual shrimp parvovirus (HPV): use of gene probes in disease diagnosis. Diseases of aquatic Organisms 22: 129-134.

Paynter, J.L., Lightner, D.V. and Lester, R.J.G. 1985. Prawn virus from juvenile Penaeus esculentus. In: P.C. Rothlisberg, B.J. Hill and D.J. Staples (eds.) Second Australian National Prawn Seminar. NPS2, Cleveland, Queensland. pp. 61-64.

Roubal, F.R., Paynter, J.L. and Lester, R.J.G. 1989. Electron microscopic observation of hepatopancreatic parvo-like virus (HPV) in the penaeid prawn, Penaeus merguiensis de Man from Australia. J. Fish. Dis. 12: 199-201.

Lymphoidal Parvo-Like Virus (LPV) LPV has only been observed in Australia and is reported from cultured P. monodon, P. merguiensis, P. esculentus and a P. monodon x P. esculentus hybrid (Owens et al., 1992).

Clinical signs and gross pathologyLPV does not cause disease or mortality in infected populations.

HistopathologyLPV primarily infects lymphoid organ cells and causes nuclear hypertrophy, marginated chromatin and increase in the proportion of cytoplasm (Owens et al., 1991). Areas of cellular transformation in the lymphoid organ are discrete, are referred to as "spheroids" and are formed from lymphoid organ cords in which the sheath cells have become hypertrophied and hyperplastic, and the central vessel obliterated (Lightner, 1996). Spheroids may also contain pyknotic and karyorrhetic nuclei. LPV is an intranuclear virus, 18-20 nm in diameter which occasionally forms eosinophilic to basophilic inclusion bodies (IB’s) in infected cells of the lymphoid organ, haematopoietic tissues, gills and the connective tissues of various organs. The virus is believed to be a parvovirus based on the size of the virion and inclusions that stain positive for DNA with acridine orange fluorescence (Owens et al., 1991). LPV-

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infected tissue does not react with the BS4.5 Probe for IHHNV (Leigh Owens, personal communication).

DiagnosisDiagnosis of LPV is based on the histological demonstration of eosinophilic IB’s in cells within “spheroids” formed in the lymphoid organ. LPV IB’s are spherical and may also be detected histologically in other tissues. Definitive diagnosis may be made by identifying LPV in lymphoid organs cells by electron microscopy (Owens et al., 1992).

TransmissionAspects of the biology of LPV, such as the mode of transmission, range of susceptible hosts and viability in the external environment are not known

PreventionLPV is not considered a serious risk to wild or cultured prawns. However, its effect on prawn defence mechanisms is not known.

ReferencesLightner, 1996 (ed.) A Handbook of Shrimp Pathology and Diagnostic Procedures for

Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA.,USA.

Owens, L., De Beer, S. and Smith, J. 1991. Lymphoidal parvovirus-like particles in Australian penaeid prawns. Dis. Aquat. Org. 11: 129-134.

Owens, L., Anderson, I.G., Kenway, M., Trott, L. and Benzie, J.A.H. 1992. Infectious hypodermal and hematopoietic necrosis virus (IHHNV) in a hybrid penaeid prawn from tropical Australia. Dis. Aquat. Org. 14: 219-228.

Lymphoid Organ Virus (LOV)

LOV is endemic to cultured P. monodon from Queensland, Australia. It has been identified in the lymphoid organs of early juvenile to adult prawns. The wild penaeids, P. esculentus, P. plebejus, M. bennettae and M. ensis were investigated and found free of LOV as was cultured P. japonicus

Clinical signs and gross pathologyLOV is not associated with disease and does not appear to be a significant pathogen of penaeid prawns (Spann et al., 1995).

HistopathologyLOV infected the lymphoid organ and is associated with LPV-like spheriods, resembling tubules lacking a central haemolymph vessel. Cells within the spheriods are hypertrophied and there is an increase in the proportion of cytoplasm. Nuclei are also hypertrophied and may be pyknotic. Cytoplasmic inclusion may also be present. LOV virions are enveloped, rod-shaped particles, 163-200 nm x 36-63 nm and are often packed into paracrystalline arrays within the cytoplasm of infected nuclei (Spann et al., 1995).

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DiagnosisThe histopathology of lymphoid organs infected with LOV is indicative of numerous prawn viruses and therefore cannot be used as a diagnostic characteristic. Definitive diagnosis of LOV is by the demonstration by TEM of rod-shaped virus particles within spheroids of the lymphoid organ (Spann et al., 1995).

TransmissionLOV has been identified in wild-caught P. monodon broodstock and in postlarvae raised in the laboratory and it has been suggested that LOV, as a systemic virus, is transmitted vertically from broodstock. Within ponds, LOV would probably be transmitted via cannibalism.

ViabilityThe viability of LOV in the external environment is not known.

ReferencesSpann, K.M., Vickers, J.E. and Lester, R.J.G. 1995. Lymphoid organ virus of

Penaeus monodon from Australia. Dis. Aquat. Org. 23: 127-134.

Gill Associated Virus (GAV)

GAV was found to be associated with mortalities of cultured adult Penaeus monodon from farms in Queensland, Australia during 1996. It occurred in the lymphoid organ and gills of infected prawns. The host range of GAV is not known.

Clinical signs and gross pathologyInfected prawns are lethargic, anorexic and swim near the surface and edges of ponds. They display degrees of pink to red colouration of the appendages and body surface. The gills may be yellow to pink. Gill fouling and tail rot are common among infected animals.

DiagnosisDiagnosis is based on the demonstration by TEM of rod-shaped, enveloped virions and filamentous nucleocapsids in the cytoplasm of infected cells of the lymphoid organ and gills.

TransmissionWithin infected ponds transmission is thought to occur via cannibalism. The viability of GAV in the external environment is not known.

ReferencesSpann, K.M., Cowley, J.A., Walker, P.J. and Lester, R.J.G. 1997. Gill-associated

virus (GAV), a yellow head-like virus, from Penaeus monodon cultured in Australia. Dis. Aquat. Org. in press.

Spawner-isolated Mortality Virus (SMV)

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SMV occurred in P. monodon spawners held at a research station in northern Queensland, Australia in 1993 (Fraser and Owens, 1996). SMV is believed to be associated with Mid-crop Mortality Syndrome (MCMS), which caused mortalities among young adult, cultured P. monodon in Australia from 1994 to 1996 (Anderson, 1996).

Clinical signs and gross pathologyPrawns suffering MCMS appear dark red and may produce red faeces (Fraser and Owens, 1996). Spawners infected with SMV are lethargic, do not feed and may also appear red in colour.

Histopathology Young adult P. monodon, infected experimentally with extracts from prawns suffering MCMS displayed haemocytic infiltration, necrosis and the sloughing of cells into the lumens of the gut and hepatopancreas. The hepatopancreatic tubules were shrunken. Eosinophilic refractile material was observed in the subcuticular epithelium and in the capsule surrounding the hepatopancreas. Haemocytic infiltration was also observed in the subcuticular epithelium and underlying muscle. Small icosahedral virions resembling those of the Parvoviridae were observed in cells of the gut (Fraser and Owens, 1996). Partial characterisation and treatment of prawn extracts with DNase and RNase also indicate that SMV is a parvo-like virus (Fraser and Owens, 1996).

DiagnosisDefinitive diagnosis of SMV is by the demonstration by TEM of icosahedral particles, approximately 20 nm in diameter, in the cytoplasm of gut cells (Fraser and Owens, 1996). A probe for SMA is being developed (Owens, 1997).

TransmissionSuccessful feeding trials conducted by Fraser and Owens (1996) indicate that the primary mode of transmission of SMV is by cannibalism.

ReferencesAnderson I. 1996. Overview of Mid-crop Mortality Syndrome and subsequent prawn

mortalities – studies completed and future work. Australian Prawn Farmers, annual meeting, Cairns.

Fraser, C.A. and Owens, L. 1996. Spawner-isolated mortality virus from Australian Penaeus monodon. Dis. Aquat. Org. 27: 141-148.

Owens, L. 1997. Probe and bioassay analysis of mid crop mortality syndrome. Abstract. Australia Prawn Farmers Association, annual meeting, Brisbane. p. 43

Penaeid Haemocytic Rod-shaped Virus (PHRV)

A haemocytic rod-shaped virus was isolated in 1992 from the gills of hybrid P. esculentus x P. monodon prawns bred in Australia. These prawns were also infected with a IHHNV-type virus and were dying (Owens, 1993)Gross PathologyHybrid prawns infected with IHHNV and PHRV showed no gross signs of disease.

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HistopathologyRod-shaped virions, either free in the cytoplasm or enclosed within vesicles, occur in gills haemocytes. The nuclei of infected haemocytes may be slightly marginated and display darkening of the chromatin. PHRV virions are enveloped and may be bent in a V- or U-shape. The nucleocapsids are longer and wider than any other haemocytic rod-shaped virus and are 542-888 nm x 86 nm (Owens, 1993).

DiagnosisPHRV may be diagnosed by the demonstration by TEM of unusually long, rod-shaped virus particles in gill haemocytes.

TransmissionAspects of the biology of PRHV are not known (Owens, 1993).

ReferencesOwens, L. 1993. Description of the first haemocytic rod-shaped virus from a penaeid

prawn. Dis. Aquat. Anim. 16: 217-221.

Bacteria

Mycobacteria (mycobacteriosis)

This condition is also referred to as shrimp tuberculosis (Lightner, 1996). Mycobacterium spp. are widely found in vertebrates, including humans (Howard et al., 1987) and fish (Humphrey et al., 1987; Wada et al., 1993; Lansdell et al., 1993). However infection in penaeids and other invertebrates is rare. Mycobacterium species are ubiquitous and potentially infectious to all prawn species but have not been reported from penaeids cultured in Australia. Infections of a Mycobacterium species have been reported in wild, adult P. vannamei from Panama and Ecuador (Lightner and Redman, 1986; Lightner, 1993) and cultured juvenile P. vannamei from Mississippi (Krol et al., 1989). A Mycobacterium species was identified in a captive, female Macrobrachium rosenbergii spawner (Brock et al., 1986). Mycobacterial granulomas have been found in a captive Macrobrachium rosenbergii prawn from Australia (Owens et al., 1992)

Clinical signsMycobacterium spp. are not known to be associated with disease, however infections pose a marketing problem due to the formation of unsightly lesions in the muscle and cuticle (Lightner, 1993).

Gross PathologyMultifocal haemocytic nodules may be visible in the hepatopancreas and appear dark due to melanisation (Lightner et al., 1986). Nodules may also occur in the connective tissues of the hepatopancreas, haemocytes, ovary, lymphoid organ, heart, cuticle, antennal gland, gills and mandibular organ (Lightner et al., 1986; Lightner, 1993).

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HistopathologyMycobacteriosis is caused by the Gram-positive, acid fast bacteria Mycobacterium marinum, M. fortuitum and M. sp. (Lightner, 1996). These bacteria are rod-shaped, 1.1 0.5 m wide and 4.9 0.5 m long (Krol et al., 1989). Nodules are composed of haemocytes in concentric layers around a core of rod-shaped bacteria and degenerate, necrotic cells (Krol et al., 1989).

DiagnosisDiagnosis is based on clinical signs of infection and the demonstration of bacteria either in impression smears made from melanised lesions or in tissue sections. Bacteria appear basophilic when sections are stained with haemotoxylin and eosin and red when sections are stained with Kinyoun’s carbol fuchsin and the Ziehl-Neelsen method (Luna, 1968; Lightner, 1996).

TransmissionThe significance of Mycobacterium spp. as prawn pathogens and their biology when infecting prawns are not really known.

TreatmentTreatment and control measures are not known.

ReferencesBrock, J.A., Nakagawa, L.K. and Shimojo, R.J. 1986. Infection of a cultured

freshwater prawn, Macrobrachium rosenbergii de Man (Crustacea: Decapoda), by Mycobacterium spp., Runyon Group II. J. Fish Dis. 9: 319-324.

Howard, J.B., Klaas, J., Rubin, S.J., Weissfield, A.S. and Tilton. 1987. Clinical and Pathogenic Microbiology. C.V. Mosby, St. Louis, Missouri.

Humphrey, J.D., Lancaster, C.E., Gudkovs, N. and Copland, J.W. 1987. The disease status of Australian salmonids: bacteria and bacterial diseases. J. Fish Dis. 10: 403-410.

Krol, R.M., Hawkins, W.E., Vogelbein, W.K. and Overstreet, R.M. 1989. Histopathology and ultrastructure of the hemocytic response to an acid-fast bacterial infection in cultured P. vannamei. J. Aquat. Anim. Health 1: 37-42.

Lansdell, W., Dixon, B., Smoth, N and Benjamin, L. 1993. Isolation of several Mycobacterium species from fish. J. Aquat. Anim. Health 5: 73-76.

Lightner, D.V. 1993. Diseases of penaeid shrimp. In: McVey, J.P. (ed.) CRC Handbook of Mariculture: Crustacean Aquaculture. 2nd edition. CRC Press, Boca Raton, FL. pp. 393-486.

Lightner, D.V. and Redman, R.M. 1986. A probable Mycobacterium sp. infection of the marine shrimp Penaeus vannamei (Crustacea: Decapoda). J. Fish Diseases 9: 357-359.

Owens, L. and Hall-Mendelin, 1989. Recent Advances in Australian prawns (sic) diseases and pathology. Advances in Tropical Aquaculture, Tahiti, AQUACOP, IFREMER, Actes de Colloque 9: 103-112.

Wada, S., Hatai, K., Tanaka, E. and Kitahara, T. 1993. Mixed infection of an acid-fast bacterium and an imperfect fungus in a Napoleon fish (Cheilinus undulatus). J. Wildlife Dis. 29(4): 591-595.

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Chitinoclastic Bacteria (other than vibrios) Associated with Shell Disease

Shell disease may also be referred to as black spot, brown spot or spot disease and was recognised early this century as a problem in impounded populations of aquatic animals (Sindermann, 1991). The chitinoclastic bacteria which cause shell disease are ubiquitous and infect a wide range of crustaceans, including prawns. All wild and cultured prawns are susceptible to infection, however disease rarely occurs in wild crustaceans due to a lack of overcrowding and less mechanical damage (Cook and Lofton, 1973). The expression of shell disease as a chronic condition or as an acute condition associated with mortality, depends on host susceptibility, the pathogen involved and environmental conditions (Brock and Lightner, 1990). Numerous other bacterial and fungal organisms invade shell disease lesions as secondary pathogens.

Clinical signsShell disease causes visible lesions on the body cuticle, appendages or gills of affected crustaceans (Sindermann, 1990). Lesions are soft, cratered, often melanised and may progressively enlarge to cover large areas of the cuticle (Getchell, 1989). Segments of affected appendages may be lost. Lesions are typically lost when the animal moults (Gopalana and Young, 1975). Death may occur at the time of ecdysis when the old and new exoskeletons fail to separate or may occur as a result of secondary infection (Fisher et al., 1976; Lightner, 1983).

Gross PathologyShell disease lesions may affect any surface of the body and appendages. Lesions are typically restricted to the cuticle, however in severe cases they may extend into deeper layers and become systemic (Gopalana and Young, 1975).

HistopathologyShell disease is caused by chitinoclastic bacteria or those with lipolytic properties, such as Vibrio spp., Altermonas sp., Beneckea spp. and Spirillum sp. (Cook and Lofton, 1975; Delves-Broughton and Poupard, 1976; Paynter, 1989). There is little histopathology associated with this infection. Rod-shaped bacteria are observed in histological sections of infected tissues. Haemocytic infiltration occurs when lesions extend into the endocuticle.

DiagnosisDiagnosis of based on the demonstration of brown to black cuticular lesions and/or the loss of appendage segments (Paynter, 1989).

TransmissionWounds, abrasions or chemical degradation of the cuticle are required to initiate infection (Brock and Lightner, 1990). Poor culture conditions, such as crowding, poor water exchange, elevated temperature and poor diet result in an increase in the incidence of shell disease (Brock and Lightner, 1990). Physical wounding is required for infection by chitinoclastic bacteria (Cook and Lofton, 1973). Shell disease may be contagious under poor environmental conditions (Getchell, 1989).

Treatment

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High levels of organic matter provide ideal conditions for the growth of chitinoclastic bacteria (Gopalana and Young, 1975). Therefore, shell disease may be controlled in culture facilities by implementing proper husbandry practices, such as avoiding overcrowding, system sterilisation, minimising handling, improving water flow, keeping organics at a low level, culling affected individuals and providing a nutritionally balanced diet (Sindermann, 1991). Antibiotics, such as erythromycin, streptomycin and formalin, or malachite green (0.9 ppm) may be added to the water during larval rearing (Paynter, 1989; Sindermann, 1991).

ReferencesBrock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne

(ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Cook, D.W. and Lofton, S.R. 1973. Chitinoclastic bacteria associated with shell disease in Penaeus shrimp and the blue crab. J. Wildl. Dis. 9: 154-159.

Delves-Broughton, J. and Poupard, C.W. 1976. Disease problems of prawns in recirculating systems in the U.K. Aquaculture 5: 201-217.

Fisher, W.S., Rosemark, T.R. and Nilson, E.H. 1976. The susceptibility of cultured American lobsters to a chitinolytic bacterium. Proc. Wld Maricult. Soc. 7: 511-520.

Getchell, R.G. 1989. Bacterial shell disease in crustaceans: a review. J. Shellfish Res. 8: 1-6.

Gopalana, U.K. and Young, J.S. 1975. Incidence of shell disease in shrimp in the New York Bight. Mar. Pollut. Bull. 6: 149-153.

Lightner, D.V. 1983. Diseases of cultured penaeid shrimp In: J.R. Moore (ed. in chief) CRC Handbook of Mariculture Vol. 1. J.P. McVay (ed.) Crustacean Aquaculture. CRC Press, Boca Raton, FL. pp. 289-320.

Paynter, J.L.1989. Invertebrates in Aquaculture. Refresher course for Veterinarians, Proceedings 177. The University of Queensland.

Sindermann, C.J. 1990. Principal Diseases of Marine Fish and Shellfish, Vol. 2, 2nd edition. Academic Press, New York.

Sindermann, C.J. 1991. Shell disease in marine crustaceans-a conceptual approach. J. Shellfish Res. 10(2): 491-494.

Aeromonas sp. and Pseudomonas sp. (Necrosis And Septicemias)

Aeromonas sp., Pseudomonas sp. are part of the normal microflora of wild and cultured crustaceans and are opportunistic pathogens (Lightner, 1993). They are associated with mortality less frequently than Vibrio spp. and are not considered primary pathogens. Aeromonas sp. is associated with soft-shell syndrome of Penaeus monodon from the Philippines (Baticados et al., 1986). Aeromonas sp. and Pseudomonas sp. usually occur in mixed infections with other bacteria, particularly Vibrio spp., viruses and/or fungi. All species of penaeids and life stages are susceptible to infection by these bacteria.

Clinical signsClinical signs of infection by these bacteria are similar to those of vibriosis, which may also be manifest as bacterial necrosis and/or septicemia. Mortalities may occur

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when prawns are stressed. Lightner and Lewis (1975) demonstrated that Aeromonas sp. is pathogenic to prawns when injected at high doses and infected prawns swim erratically or appear lethargic (Lightner and Lewis, 1975).

Gross PathologyBrown spots may be observed on the gills and abdominal muscle may appear opaque. The general body surface may appear red or dark and cuticular lesions may be formed. Bacterial septicemia may result in slow blood clotting and the turbid appearance of the blood. The hepatopancreas is typically atrophied and melanised granulomas may be formed in the lymphoid organ (Lightner and Lewis, 1975; Anderson et al., 1988; Paynter, 1989).

HistopathologyAeromonas sp., Pseudomonas sp. are Gram-negative, rod-shaped bacteria which may be observed in sections of infected organs using histochemical techniques. Infected cells are typically hypertrophied. Haemocyte nests may be observed in the heart and hepatopancreas. Cuticular colonisation associated with lesions may result in the necrosis of the cuticular epithelium and the formation of lesions (Lightner and Lewis, 1975; Paynter, 1989).

DiagnosisDiagnosis of bacterial necrosis and septicemias is based on clinical signs and the demonstration of Gram-negative, rod-shaped bacteria in tissues and hemolymph. A series of tests are performed to identify different Gram-negative bacteria. These include: motility, growth on specific agar, growth in the presence of NaCl, glucose utilisation, urea hydrolysis, nitrate reduction and luminescence (Lightner, 1993).

TransmissionOpportunistic bacteria invade through wounds and cracks in the cuticle and are ingested with food. They are spread to other organs via the haemolymph (Paynter, 1989).

TreatmentBacterial necrosis and septicemias are controlled primarily by maintaining good husbandry practices, such as ensuring adequate water exchange and adequate, high quality feeds in order to reduce stress on prawns (Baticados et al., 1986; Paynter, 1989). Antibiotics may be added to water or feed, however resistant strains of bacteria may develop (Baticados et al., 1990).

ReferencesAnderson, I.G., Shamsudin, M.N. and Shariff, M. 1988. Bacterial septicemia in

juvenile tiger shrimp, Penaeus monodon, cultured in Malaysian brackishwater ponds. Asian Fis. Sci. 2: 93-108.

Baticados, M.C.L., Lavilla-Pitogo, C.R., Cruz-Lacierda, E.R., de la Pena, L.D. and Sunaz, N.A. 1990. Studies on the chemical control of luminous bacteria Vibrio harveyi and V. splendidus isolated from diseased Penaeus monodon larvae and rearing water. Dis. Aquat. Org. 9: 133-139.

Lightner, D.V. 1993. Diseases of cultured penaeid shrimp. In: J.P. McVey (ed.) CRC Handbook of Mariculture, Second edition, Volume 1, Crustacean Aquaculture. CRC Press Inc., Boca Raton, FL. p. 393-486.

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Lightner, D.V. and Lewis, D.H. 1975. A septicemic bacterial disease syndrome of penaeid shrimp. Mar. Fish. Rev. 37(5-6): 25-28.

Paynter, J.L. 1989. Invertebrates in Aquaculture. Refresher Course for Veterinarians, Proceedings 117. The University of Queensland.

Epibiont Bacteria which Cause Fouling (Principally Leucothrix mucor)

Numerous bacteria, algae and protozoans are involved in fouling diseases of prawns. Most of these agents are free-living organisms and not true pathogens. The filamentous bacterial epibionts of prawns are: Flavobacterium sp., Cytophaga sp., Flexibacter sp., Thiothrix sp. and Leucothrix sp. The most important of these is Leucothrix mucor.

L. mucor is an epiphyte of macroscopic algae (Bland and Brock, 1973) and can cause fouling problems in numerous crustaceans (Brock and Lightner, 1990). L. mucor is ubiquitous and all penaeid species are susceptible. However, geographically distinct strains may exist (Lightner, 1996). Severe fouling occurs when prawns are stressed and decrease preening activity and when water quality is low. Egg and larval stages are more prone to suffer disease and mortality than juveniles and adults. Fouling of gills and chemoreceptor sites is more significant than fouling of general body surface as gas exchange and other vital functions may be impaired (Brock and Lightner, 1990).

Clinical signsL. mucor is a filamentous bacteria and therefore readily visible under a dissecting microscope. Fouling may cause discolouration of the body surface and gills from yellow-brown to brown-black (Lightner, 1977). Fouled prawns may appear “fuzzy” if heavily colonised.

Gross Pathology and HistopathologyFouling by L. mucor does not cause any structural changes to the cuticle and does not invade internal tissues. A mucoid layer may be formed on gill lamellae and interfere with oxygen uptake (Fisher, 1987).

DiagnosisL. mucor can be observed in wet, unstained tissue mounts using a dissecting microscope with transmitted light or a compound microscope at low magnification (Lightner, 1983).

TransmissionEcdysis results in shedding of epibionts. L.mucor will not settle on prawns unless they are stressed and have reduced preening activity (Brock and Lightner, 1990). L. mucor would probably not survive freezing and transport, although no data is available.

TreatmentMalachite green is an effective treatment for prawn eggs, but not larvae (Brock and Lightner, 1990). Antibiotics, such as streptomycin, chloramphenicol and

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oxytetracycline are effective against fouling at all life stages (Lightner, 1983). Surface fouling organisms can be killed by dipping adult prawns in formalin (Lightner, 1996). Copper compounds are also effective when added to pond water (Lightner, 1983).

ReferencesBland, J.A. and Brock, T.D. 1973. The marine bacterium Leucothrix mucor as an

algal epiphyte. Marine Biology 23: 283-292. Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne

(ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Fisher, W.S. 1977. Epibiotic microbial infestations of cultured crustaceans. Proc. Wld. Maricult. Soc. 8: 673-684.

Lightner, D.V. 1977. Shrimp diseases. In: C.J. Sindermann (ed.) Disease Diagnosis and Control in North American Marine Aquaculture. Developments in Aquaculture and Fisheries Science. Vol 6. Elsevier, New York. pp. 10-77 .

Lightner, D.V. 1983. Diseases of cultured penaeid shrimp In: J.R. Moore (ed. in chief) CRC Handbook of Mariculture Vol. 1. J.P. McVey (ed.) Crustacean Aquaculture. CRC Press, Boca Raton, FL. pp. 289-320.

Fungi

Fusarium solani (Fusariosis)

Fusariosis or black gill disease is caused by the imperfect fungi Fusarium solani which has been reported as a pathogen of numerous crustaceans (Lightner, 1981). All penaeid species are potentially susceptible, however Penaeus japonicus and P. californiensis are highly susceptible (Ishikawa, 1968). P. stylirostris and P. vannamei are moderately susceptible while P. monodon and P. mergiuensis are relatively resistant (Lightner, 1988). Fusariosis has also been reported from P. duorarum (Johnson, 1974), P. setiferus and P. aztecus (Solangi and Lightner, 1976). Major epidemics, resulting in 100% mortality have been reported in Japan (Ishikara, 1968), the Philippines (Baticados, 1988) and Mexico (Lightner, 1975).

Fusarium causes disease and mortality in cultured prawns only and is a result of poor pond conditions and stress on animals (Lightner, 1976). Adult prawns are most susceptible to infection (Lightner, 1988). F. solani is ubiquitous and present in soils and detritus. This fungus is an opportunistic pathogen and may be introduced to culture systems from the pond bottom (Lightner et al., 1979). F. solani is present in coastal soils of Queensland from Rockhampton to Bundaberg (Burgess and Summerell, 1992). A single presumptive diagnosis of infection with F. solani has been made in P. monodon in Australia (L. Owens, pers. comm.).

Clinical signsBehavioural changes have not been reported for prawns infected with F. solani,

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however they show lesions on the gills, appendages and/or cuticle. Lesions may be singular or multiple, ulcerated or raised and slow developing (Egusa and Ueda, 1972, Brock and Lightner, 1990). Head and tail appendages may be deformed or missing (Lightner, 1996) and mortality may occur in heavily infected populations. Experimental infection of prawns, where F. solani propagules were inoculated into fresh cuticular wounds, resulted in 100% mortality within 2 weeks (Hose et al., 1984). Mortality may be associated with the production of mycotoxins (Lightner, 1976).

Gross PathologyLesions caused by F. solani are usually melanised. The fungus is restricted to head and tail appendages, gills and muscle adjacent to the cuticle. Invasion of internal organs has not been reported (Brock and Lightner, 1990).

HistopathologyMelanised cuticular lesions appear as granulomatous nodules due to the encapsulation of fungal hyphae by host haemocytes (Bian and Egusa, 1981). In advanced infections, the levels of glucose, protein, alkaline phosphatase and serum glutamic oxaloacetic transaminase in the haemolymph may be altered (Hose et al., 1984). Fungal hyphae and conidia are visible in wet mounts and stained sections of infected tissue (Lightner, 1993). Haemocyte activity, in response to infection by F. solani, varies between penaeid species and is an important factor in the apparent resistance of some species (Solani and Lightner, 1976).

DiagnosisDiagnosis is based on the presence of lesions and dark colouration of gills and on the demonstration of fungal hyphae and conidia within haemocytic nodules using light microscopy. Hyphae appear eosinophilic with haemotoxylin and eosin stain. PAS and PAS-based silver stains clearly demonstrate F. solani hyphae and the characteristic canoe-shaped macroconidia (Lightner, 1996). The macroconidia may also be seen in unstained wet mount smears of material from lesions (Lightner, 1983). Definitive diagnosis is made by isolating and identifying Fusarium spp., following culture on any mycological medium (Lightner, 1988).

TransmissionFusarium conidiospores are present in the soil and water and establish infection in prawns following invasion of slight cuticular wounds (Lightner, 1981). Experimental exposure of wounded and unwounded prawns resulted in infection in wounded prawns only (Solani and Lightner, 1976).

TreatmentFusariosis cannot be control through the use of chemicals (Lightner et al., 1979). It may be controlled in the hatchery by filtering and sterilising water prior to use. The risk of damage to the carapace may be reduced by the avoidance of overcrowding and good nutrition (Paynter, 1989). ReferencesBaticados, M.C.L.1988. Diseases of prawns in the Philippines. SEAFDEC Asian

Aquaculture 10(1): 1- 8.Bian, B.Z. and Egusa, S. 1981. Histopathology of black gill disease caused by

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Fusarium solani (Martius) infection in the Kuruma prawn, Penaeus japonicus Bate. J. Fish Diseases 4: 195-201.

Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne (ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Burgess, L.W. and Summerell, B.A. 1992. Mycogeography of Fusarium: survey of Fusarium spp in subtropical and semi-arid grassland soils from Queensland, Australia. Mycological Research 96(9): 780-784.

Egusa, S. and Ueda, T. 1972. A Fusarium sp. associated with black gill disease of the Kuruma prawn, Penaeus japonicus Bate. Bull. Japanese Soc. Sci. Fish. 38: 1253-1260.

Hose, J.E. Lightner, D.V., Redman, R.M. and Donald, D.A. 1984. Observations on the pathogenesis of the imperfect fungus, Fusarium solani, in the Californian brown shrimp, Penaeus californiensis. J. Invertebr. Pathol. 44: 292-303.

Ishikawa, Y. 1968. Preliminary report on black gill disease of the kuruma prawn, Penaeus japonicus Bate. Fish Pathol. 3: 34-38.

Johnson, S.K. 1974. Fusarium sp. in laboratory-held pink shrimp. Texas A&M University, Texas Agricultural Extension Service, Fish Disease Diagnostic Laboratory, publication no. FDDL- #1.

Lightner, D.V. 1975. Some potentially serious disease problems in the culture of penaeid shrimp in North America. Proc. U.S-Japan Natural Resources Program, Symposium on Aquaculture Diseases, Tokyo. pp. 75-97.

Lightner, D.V. 1976. Epizootiology of two mycotic diseases in the culture of penaeid shrimp. Proc. Int. Colloq. Invertebr. Pathol. 1: 179-183.

Lightner, D.V. 1981. Fungal diseases of marine crustacea. In: E.W. Davidson (ed.) Pathogenesis of Invertebrate Microbial Diseases. Allanheld, Osmund Publishers, Totowa, NJ. pp. 451-484.

Lightner, D.V. 1983. Diseases of cultured penaeid shrimp In: J.R. Moore (ed. in chief) CRC Handbook of Mariculture Vol. 1. J.P. McVey (ed.) Crustacean Aquaculture. CRC Press, Boca Raton, FL. pp. 289-320.

Lightner, D.V. 1988. Diseases of cultured penaeid shrimp and prawns. In: C.J. Sindermann and D.V. Lightner (eds.) Disease Diagnosis and Control in North American Marine Aquaculture. 2nd edition. Elsevier, New York. pp. 8-127.

Lightner, D.V. 1993. Diseases of penaeid shrimp. In: McVey, J.P. (ed.) CRC Handbook of Mariculture: Crustacean Aquaculture. 2nd edition. CRC Press, Boca Raton, FL. pp. 393-486.

Lightner, D.V. (ed.). 1996. A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA., USA

Lightner, D.V., Moore, D. and Danald, D.A. 1979. A mycotic disease of cultured penaeid shrimp caused by the fungus Fusarium solani. In: D.H. Lewis and J.K. Leong (eds.) Proceedings of the Second Biennial Crustacean Health Workshop. Texas A&M Univ. Publication No. TAMU-SG-79-114. pp. 135-158.

Paynter, J.L. 1989. Invertebrates in Aquaculture. Refresher Course for Veterinarians, Proceedings 117. The University of Queensland.

Solani, M.A. and Lightner, D.V. 1976. Cellular inflammatory response of Penaeus aztecus and Penaeus setiferus to the pathogenic fungus, Fusarium sp., isolated from the Californian brown shrimp, Penaeus californiensis. J. Invertebr. Pathol. 27: 77-86.

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Lagendium and Sirolpidium Species (Larval Mycosis)

Larval mycosis is also called fungus disease, Lagenidium or Sirolpidium disease. The phycomycetous fungi which cause this disease are ubiquitous and infect eggs and larvae of most farmed crustaceans including prawns (Brock and Lightner, 1990). Most larval mycoses are caused by members of the Lagenidium and Sirolpidium genera. The eggs and larvae of most penaeid species, including Penaeus monodon are susceptible to Lagenidium spp. (Lio-Po and Sanvictores, 1985). It is unclear if Lagenidium and Sirolpidium fungi from different geographical areas are distinct species or strains of the same species (Lightner, 1996).

Clinical signsThere are few clinical signs associated with larval mycosis as mortalities are typically sudden and mortality rates high. Infected larvae may become immobile and settle on the bottom of the tank when water circulation is stopped (Lightner and Fontaine, 1973). Secondary bacterial infections are common (Lightner 1996). Susceptibility to infection diminishes with age and mortalities among postlarval crustaceans are rare (Lightner and Fontaine, 1973). However, Sirolpidium sp. may infect early postlarval prawns (Brock and Lightner, 1990).

Gross pathologyLarval penaeids do not mount a significant immune response to phycomycetes fungi, hence fungi grow unrestricted and eventually replace most of the prawns muscle and soft tissue (Lightner and Fontaine, 1973). The fungal hyphae and discharge tubes are visible within the body cavity of infected larvae and zoospore discharge tubes may also be observed protruding through the cuticle (Brock and Lightner, 1990).

HistopathologyThe phycomycetous fungi of importance to cultured prawns are Lagenidium callinectes and Sirolpidium sp. Fungal hyphae are irregular, branched, septate, pale yellow-green in colour and possess numerous refractile oil droplets (Lightner and Fontaine, 1973).

DiagnosisDiagnosis of phycomycetes fungi is based on the demonstration of fungal hyphae and discharge tubes projecting from the host’s body. These may be observed in unstained wet mounts of larvae using either a dissecting microscope or compound microscope at low magnification. Lagenidium spp. have long discharge tubes with terminal vesicles containing zoospores. Sirolpidium spp. have short discharge tubes without terminal vesicles (Lightner, 1988). Sporogenesis and the morphology of the discharge tube may be demonstrated in pure fungal cultures grown in agar or broth (Lightner and Fontaine, 1973; Baticados et al., 1977).

Transmission

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L. callinectes and Sirolpidium spp. are introduced to hatcheries by broodstock and/or carrier hosts present in the seawater supply (Lightner, 1993). The fungal zoospores can survive in seawater for long periods of time and readily attach and encyst on the cuticle of an egg or larval prawn (Lightner, 1993).

TreatmentVarious chemicals effectively destroy fungal zoospores in hatchery seawater. These include: trifuralin (Treflan), malachite green, formalin, potassium permanganate and benzalkonium chloride (Lio-Po et al., 1982). The treatment of seawater with UV light (Armstrong et al., 1976) and flushing spawned eggs with clean seawater (Lightner, 1993) are also effective in controlling the number of zoospores in hatchery seawater.

Present status of diseaseLarval mycosis seldom causes disease in prawns hatcheries with sound management practices.

ReferencesArmstrong, D.A., Buchanan, D.V. and Caldwell, R.S. 1976. A mycosis caused by

Lagenidium sp. in laboratory-reared larvae of a Dunganess crab, Cancer magister, and possible chemical treatments. J. Invertebr. Pathol. 28: 329-336.

Baticados, M.C.L., Po, G.L., Lavilla, C.R. and Gucatan, R.Q. 1977. Isolation and culture in artificial media of Lagenidium from Penaeus monodon larvae. SEAFDEC quart. Res. Rep. Aquacult. Dep. 1(4): 9-10.

Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne (ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Lightner, D.V. 1988. Diseases of penaeid shrimp. In: C.J. Sindermann and D.V. Lightner (eds.) Disease Diagnosis and Control in North American Marine Aquaculture. 2nd edition, Elsevier Scientific Publishing Co., Amsterdam. pp. 8-133.

Lightner, D.V. 1993. Diseases of penaeid shrimp. In: McVey, J.P. (ed.) CRC Handbook of Mariculture: Crustacean Aquaculture. 2nd edition. CRC Press, Boca Raton, FL. pp. 393-486.

Lightner, D.V. (ed.). 1996. A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA., USA .

Lightner, D.V. and Fontaine, C.T. 1973. A new fungus disease of the white shrimp, Penaeus setiferus. J. Invertebr. Pathol. 22: 94-99.

Lio-Po, G.D., Sanvictores, M.E.G., Baticados, M.C.L. and Lavilla, C.R. 1982. “In vitro” effect of fungicides on hyphal growth and sporogenesis of Lagenidium spp. isolated from Penaeus monodon larvae and Scylla serrata eggs. J. Fish Diseases 5: 97-112.

Lio-Po, G. and Sanvictores, E. 1985. The tolerance of Penaeus monodon eggs and larval to fungicides against Lagenidium sp. and Haliphthoros sp. In: Y.Taki, J.H. Primavera and J.A. Llobrera (eds.) Proceedings First International Conference on the Culture of Penaeid Prawns/Shrimps. Aquaculture Dept. SEAFEC, Iloilo, Philippines. p. 180.

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Parasites

Haplosporidia

Haplosporidiosis is caused by one or more putative haplosporidian parasites which have not been investigated sufficiently to allow placement within the phylum Haplosporea (Lightner, 1996). Haplosporidiosis has been reported from juvenile P. vannamei imported from Nicaragua to Cuba (Dykova et al., 1988), juvenile P. monodon from Indonesia and the Philippines (Lightner et al., 1992) and P. stylirostris from Mexico (Lightner, 1996). One case of presumptive haplosporidiosis has been reported from P. monodon from northern Australia (L. Owens, pers. comm.).

Clinical signsNo definitive clinical signs of haplosporidian infections have been reported. Infected individuals may show poor growth (Lightner et al., 1992).

Gross PathologyNone known.

HistopathologyHaplosporidian parasites are restricted to the hepatopancreata of infected prawns (Lightner, 1993). They occur in the cytoplasm of tubule epithelial cells and cause cellular hypertrophy as the cytoplasm is replaced by multiplying plasmodia (Dykova et al., 1988). Infected cells are ultimately destroyed, releasing uninucleate stages of the parasite into the lumen. Moderate to heavy haemocytic inflammation and encapsulation may occur around heavily infected tubules (Dykova et al., 1988; Lightner et al., 1992).

DiagnosisDiagnosis is based on the demonstration of multi-nucleate plasmodia in epithelial cells of the hepatopancreas. Plasmodia may be observed in histological tissue sections stained with haemotoxylin and eosin or Wolbach’s Giemsa (Lightner, 1996). Haplosporosomes, 29-140 nm x 130-603 nm may be seen in the parasite using TEM (Dykova et al., 1988). Mature haplosporidian spores have never been observed, hence definitive classification is not possible (Lightner, 1993).

TransmissionNot known.

TreatmentNot known.

Present status of diseaseHaplosporidiosis is rare and not significant economically.

ReferencesDykova, I., Lom, J. and Fajer, E. 1988. A new haplosporean infecting the

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hepatopancreas in the penaeid shrimp, Penaeus vannamei. J. Fish Dis. 11: 15-22.Lightner, D.V. 1993. Diseases of penaeid shrimp. In: McVey, J.P. (ed.) CRC

Handbook of Mariculture: Crustacean Aquaculture. 2nd edition. CRC Press, Boca Raton, FL. pp. 393-486.

Lightner, D.V. (ed.). 1996. A Handbook of Shrimp Pathology and Diagnostic Procedures for

Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA, USA. Lightner, D.V., Bell, T.A., Redman, R.M., Mohney, L.L., Natividad, J.M., Rukyani,

A. and Poernomo, A. 1992. A review of some major diseases of economic significance in penaeid prawns/shrimp of the Americas and Indo-Pacific. In: M. Shariff, R. Subasinghe and J.R. Arthur (eds.) Proceeding 1st Symposium on Diseases in Asian Aquaculture. Fish Health Section, Asian Fisheries Society, Manilla, Philippines. p. 57-80.

GregarinesGregarines (Protozoa, Apicomplexa) have been observed in wild and cultured prawns from every continent and all penaeids and life-stages are potential hosts (Couch, 1978). Three genera infect prawns: Nematopsis spp. Cephalolobus spp. and Paraophioidina spp. (Couch, 1978; Jones et al., 1994). Each genera contains numerous species, although they are difficult to distinguish. Gregarines have been observed in numerous wild penaeid species in Australia (L. Owens and R.J.G. Lester, pers. comms.).

Clinical signsThere are no clinical signs of light infection (Couch, 1978). Severely infected prawns may show reduced growth rates and yellow discolouration of the midgut (Lightner, 1993, 1996). The only adverse effect gregarines may have on prawns in that massive infection may block the hosts filter apparatus or ducts leading to the hepatopancrea (Couch, 1983).

Gross PathologyGregarine trophozoites may be visible in the midgut of infected larvae and postlarvae when viewed under a dissecting microscope (Jones et al., 1994).

HistopathologyThere is little histopathology associated with gregarine infection as ingested spores attach to the walls of the gastric filter, midgut , midgut caecae, primary ducts of the hepatopancreas, posterior stomach and anterior hind gut and rarely invade host cells (Couch, 1978; Lightner, 1993). In heavy infections lesions consisting of necrosis and perforation of the midgut mucosa and hyperplasia of the midgut epithelium to form villus-like folds, may form at sites of attachment (Lightner, 1993).

DiagnosisGregarine sporozoites, trophozoites and gametocytes may be observed by light microscopy in wet mount preparations of midgut contents (Overstreet, 1973).

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TransmissionPrawns become infected after ingestion of an intermediate host, such as a mollusc or polychaete worm, which contains gregarine spores. Once attached to the wall of the gastric filter or midgut, spores develop into feeding trophozoites, which in turn are released and pass to the hindgut, where they lodge in the folds of that organ and develop into gametocytes. The gametocytes rupture to release gametes, which form zygotes. Zygotes (or zygospores) are ingested by an intermediate host and undergo sporogony in the epithelial cells. Spores are released in the pseudofaeces of the mollusc or when the intermediate host is ingested by a prawn (Overstreet, 1973; Couch, 1978, 1983; Lightner, 1993).

TreatmentPrawn farmers in Ecuador have found that medicated feed containing anticoccidial drugs are effective in controlling gregarine infections (Bell and Lightner, 1992).

ReferencesBell, T.A. and Lightner, D.V. 1992. Chemotherapy in aquaculture today – current

practices in shrimp culture: available treatments and their efficiency. In: C. Michel and D.J. Alderman (eds.) Chemotherapy in Aquaculture: from Theory to Reality. Office International des Epizooties, Paris. pp. 45-57.

Couch, J.A. 1978. Diseases, parasites and toxic responses of commercial penaeid shrimps of the Gulf of Mexico and South Atlantic Coasts of North America. Fish. Bull. 76: 1-44.

Couch, J.A. 1983. Diseases caused by protozoa. In: A.J. Provenzano, Jr.(ed.) The Biology of Crusctacea, Vol. 6, Academic Press, New York. pp. 79-111.

Jones, T.J., Overstreet, R.M. Lotz, J.M. and Frelier, P.F. 1994. Paraophioidina scolecoides n. sp., a new aseptate gregarine form cultured Pacific white shrimp Penaeus vannamei. Dis. Aquat. Org. 19: 67-75.

Lightner, D.V. 1993. Diseases of penaeid shrimp. In: McVey, J.P. (ed.) CRC Handbook of Mariculture: Crustacean Aquaculture. 2nd edition. CRC Press, Boca Raton, FL. pp. 393-486.

Lightner, D.V. (ed.). 1996. A Handbook of Shrimp Pathology and Diagnostic Procedures for Diseases of Cultured Penaeid Shrimp. World Aquaculture Society, Baton Rouge, LA, USA.

Overstreet, R.M. 1973. Parasites of some penaeid shrimp with emphasis on reared hosts. Aquaculture 2: 105-140.

Other Miscellaneous Parasites

Paranophrys speciesParanophrys sp. is a holotrich ciliate which infects Penaeus chinensis cultured in China. Disease occurs primarily in larvae and overwintering adults. This ciliate is an opportunistic pathogen which invades wounds (Bower et al., 1994) and has not been recorded in Australia. It is unlikely that Paranophrys sp. would survive shipment to Australia. Affected prawns have obvious wounds.

Sylon species

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Sylon species is a rhizocephalan parasite known from 21 species of Caridea prawns from northern oceans. Sylon sp causes disease primarily in Spirontocaris lilljeborgi from Norway and Pandalus platyceros from Canada (Brock and Lightner, 1990). The rootlet system of the parasite surrounds the nerve cord and invades various connective tissues. The parasite castrates the host and infected prawns usually die. Survivors are marked by obvious brown tissue scars (Bower et al., 1994). Sylon sp. is easily diagnosed and the parasite forms an externa on the ventral surface of the prawn’s abdomen. It is unlikely that a parasite of cold-climate prawn species would survive in Australian waters if released. Affected prawns can be recognised by brown tissue scars or a small sac on the ventral surface of the abdomen.

IsopodsIsopods from the family Bopyridae parasitise prawns from numerous parts of the world (Brock and Lightner, 1990). In Australia bopyrids parasitise numerous prawn species from the north-west of Western Australia to Townsville (Owens, 1990). Bopyrids attach to the brachial chamber of the host prawn and are therefore easily identified. They do not generally kill the host, but are considered a commercial problem as they interfere with the process of grading different sized prawns.

Metacercariae in prawns

Metacercariae of the parasites Microphallus sp. and Opercoeloides fimbriatus have been reported from P. setiferus and P. vannamei in the USA. Adult parasites occur in drum fish in the Mississippi River. These parasites are not associated with disease in either fish or prawns (Overstreet, 1973).

References

Brock, J.A. and Lightner, D.V. 1990. Chapter 3: Diseases of Crustacea. In: O. Kinne (ed.) Diseases of Marine Animals Vol. 3, Biologische Anstalt Helgoland, Hamburg. pp. 245-424.

Bower, S.M., McGladdery, S.E. and Price, I.M. 1994. Synopsis of diseases and parasites of shellfish. In: M. Faisal and F.M. Hetrick (eds.) Annual Review of Fish Diseases. Vol. 4.

Meyers, T.R., Lightner, D.V. and Redman, R.M. 1994. A dimoflagellate-like parasite in Alaskan spot shrimp Pandalus platyceros and pink shrimp P. borealis. Dis. Aquat. Org. 18: 71-76.

Overstreet, R.M. 1973. Parasites of penaeid shrimp with emphasis on reared hosts. Aquaculture 2: 105-140.

Owens, L. 1990. Maricultural considerations of the zoogeography of parasites from prawns in tropical Australia. J. Aqua. Trop. 5: 35-41.

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