Download review of pathogens of prawns - Australian Prawn Farmers Association

Annexe 1
REVIEW OF PATHOGENS OF PRAWNS
Contents
Section 1 Disease agents which will be considered further in the IRA
Viruses
Yellow-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)
Bacteria
Necrotizing Hepatopancreatitis (NHP)
Vibrio species (vibriosis)
Rickettsia
Aerococcus viridans var. homari
Parasites
Microsporidia
Hematodinium-like organism
Parauronema spp.
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Section 2 Disease agents which will not be further considered in the IRA
Viruses
Lymphoid Organ Vacuolization Virus (LOVV)
REO-III AND REO-IV
Hepatopancreatic 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)
Bacteria
Mycobacteria (mycobacteriosis)
Chitinoclastic Bacteria (other than vibrios) Associated with Shell Disease
Aeromonas sp. and Pseudomonas sp. (Necrosis And Septicemias)
Epibiont Bacteria which Cause Fouling (Principally Leucothrix mucor)
Fungi
Fusarium solani (Fusariosis)
Lagendium and Sirolpidium Species (Larval Mycosis)
Parasites
Haplosporidia
Gregarines
Other Miscellaneous Parasites
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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
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YHV in ponds (Flegel et al., 1995b). YHD has not been reported in Australia.
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 signs
Prawns 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 Pathology
There 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).
Histopathology
Moribund 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
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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 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).
Diagnosis
Presumptive 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 carriers
YHV 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
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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.
There is evidence that the spread of YHV to US may have resulted from frozen
imported ‘commodity’ prawns (Lightner et al., 1997).
Viability
YHV 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).
Prevention
YHV 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 poststocking 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 disease
In 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).
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When epizootics due to YHV(and WSSV) occur, emergency harvests are commonly
employed in Asia to salvage marketable prawn crops (Lightner et al., 1997).
References
Anonymous, 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.
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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.
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: 4550.
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.
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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.
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 signs
Infected 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 Pathology
There 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
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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).
Histopathology
WSBV 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.
Diagnosis
Diagnosis 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).
9
Transmission and potential carriers
Recent 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
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.
Viability
Experiments 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 semiclosed 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
10
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
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 syndrome
SEMBV 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.
References
Ahmed, 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.
11
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: 149158.
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.
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.
12
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.
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.
13
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 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 pathology
TS 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).
14
Histopathology
TS 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).
Diagnosis
Diagnosis 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 carriers
It 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
15
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).
Viability
TSV 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
mortem changes and the apparent capacity of the virus to endure long term freezing
and multiple freeze-thaw cycles.
Prevention
Effective 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 Syndrome
Since 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).
References
Brock, 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.
16
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
collected during an epizootic in Texas. J. Aquatic Anim. Health 9: 156-159
Hasson, 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: 165176
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).
17
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
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 more
appropriate 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 signs
IHHNV 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).
18
Gross Pathology
Gross 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,
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).
Diagnosis
IHHNV 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. Nonlethal 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.
Transmission
It 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
19
ingested with infected prawns (Bell and Lightner, 1984).
Viability
IHHNV 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.
Prevention
Effective 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.
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 diseasecausing agents. IHHNV has not been recorded in Australia since the death of the
diseased hybrid prawns originally described.
References
Bell, 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 nondestructive 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.
20
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
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
21
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 signs
Epizootics 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 pathology
There is little gross pathology associated with BP infection as mortalities occur very
quickly in early life stages.
Histopathology
BP 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).
Diagnosis
Diagnosis 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 experimentallyinfected 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).
Transmission
BP 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.
Viability
BP 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
22
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).
Prevention
Control 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 crosscontamination 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.
References
Akamine, 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.
23
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
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,
24
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
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 signs
Mortalities 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 Pathology
Apart 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.
Histopathology
BMNV 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).
Diagnosis
Diagnosis 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
25
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 carriers
BMNV 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
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).
Prevention
BMNV 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 BMNV
Diseases 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).
References
Arimoto, 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.
26
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.
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.
27
semisulcatus, P. kerathurus and P. vannamei and occur in most areas of the IndoPacific 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
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 signs
Mortalities 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 Pathology
Due 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.
Histopathology
MBV 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).
Diagnosis
Definitive 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.,
28
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.
Transmission
MBV 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
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).
Viability
MBV 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 infection
MBV 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.
29
References
Belcher, C.R. 1997. Monodon baculovirus (MBV) and its detection by DNAtechnologie. 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
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.
30
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
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 signs
An 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).
31
The virus causes high mortalities in young salmonids, especially in rainbow and
brook trout in fresh water.
Gross Pathology
Gross signs of serious degeneration of the hepatopancreas may be evident in juvenile
P. japonicus (Giorgetti, 1989).
Histopathology
Degeneration of the hepatopancreas with slight vacuolization of the tubuli cells may
be observed in post-larvae and juveniles infected with IPNV (Giorgetti, 1989).
Diagnosis
IPNV 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 sites
IPN-like virus was isolated from the hepatopancreas of P. japonicus (Bovo et al.,
1984).
Transmission and potential carriers
No 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).
Viability
Whipple & 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
32
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.
Prevention
There are no effective vaccines or chemotherapeutics are available.
Present status of IPN disease
No reports of IPNV in prawns have occurred subsequent to that of Giorgetti (1989).
References
Billi, 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 enzymelinked immunosorbent assay. American Journal of Veterinary Research 45, 1876–
1879.
33
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.
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.
34
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.
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 signs
Infected 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 Pathology
Experimental infection has resulted in hypertrophy of the lymphoid organ (Nadala et
al., 1992).
Histopathology
RPS 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.
Diagnosis
RPS is diagnosed by the demonstration of viral particles by electron microscopy
35
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).
Transmission
Methods of transmission are unknown. It is possible that the virus may be transmitted
between prawns by cannibalism.
Viability
The 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).
References
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.
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) 143147.
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).
36
Clinical signs
Prawns 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 Pathology
The 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, 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).
Histopathology
NHP 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).
Diagnosis
Diagnosis 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).
37
Transmission
Transmission 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).
Viability
NHP in tissue was not able to be transmitted by bioassay when subjected to sun
drying for 3 weeks followed by storage at 20-22C for 2 months (Frelier et al., 1993).
Treatment
NHP 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 NHP
The incidence of mortality resulting from NHP has decreased in recent years due to
the use of effective and readily accessible treatments.
References
Bell, 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.
38
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
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
39
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
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 Pathology
Prawns 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.
Histopathology
Rod-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
40
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
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)
Transmission
Vibrio 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).
Viability
Numerous 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).
Treatment
Vibriosis 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
41
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).
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 vibriosis
Vibriosis 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.
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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. 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 signs
Rickettsial 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 Pathology
The hepatopancreas of infected prawns may appear white or stippled black (Brock et
45
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).
Histopathology
Rickettsial 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.
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).
Diagnosis
Diagnosis 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.
Transmission
Aspects 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).
Treatment
Rickettsias 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.
References
Anderson, 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.
46
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,
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. homari
Gaffkemia, 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 signs
Lobsters 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).
47
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 Pathology
Small black specks due to haemocyte aggregations may be noticeable in the gills and
other tissues late in the course of infection.
Histopathology
There is an absence of specific cytopathology.
Diagnosis
The coccus grows on a range of media at an optimal incubation temperature of 300C.
Viability
Aerococcus 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).
Transmission
Infection 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.
Treatment
An 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 disease
Management 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.
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.
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.
48
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
Microsporidia
Cotton shrimp, or milk shrimp is caused by three genera of microsporidians:
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 signs
Most 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 Pathology
Agmasoma 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
49
(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)
Histopathology
Microsporidians 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).
Diagnosis
Microsporidian 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).
Transmission
Microsporidiosis 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.
Treatment
Effective 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).
50
Present status of disease
Microsporidiosis 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).
References
Anderson, 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.
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.
51
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: 292295.
Hematodinium-like organism
A hematodinium-like protozoan infects wild Pandalus platyceros and Pandalus
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).
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.
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.
52
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).
References
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/cildsp.htm).
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.
53
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 Pathology
LOVV does not cause serious disease and there is no gross pathology associated with
infection (Bonami et al., 1992).
Histopathology
LOVV 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).
Diagnosis
Diagnosis of LOVV is based on histopathology of the lymphoid organ and the
demonstration of viral particles by electron microscopy.
Transmission
Little 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 assessment
LOVV 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.
References
Bonami, 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,
54
Baton Rouge, LA.,USA.
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 pathology
REO 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.
Histopathology
The 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).
Diagnosis
Conclusive 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.
Transmission
The 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.
Prevention
Adequate treatment and control measures for REO have not been established.
References
55
Anderson, I.G., Shariff, M., Nash, G. and 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 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 pathology
HPV 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 HPVassociated mortality was reported to be 50-100% after 4-8 weeks in juvenile P.
merguiensis (Lightner and Redman, 1985).
Histopathology
HPV 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
56
dense, finely granular material. IB’s are basophilic when fully formed and cause
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).
Diagnosis
Definitive 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).
Transmission
The 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.
Viability
The 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: 447449.
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.
57
Penaeus monodon captured broodstock surveyed for yellow-head virus and other
pathogens by electron microscopy. In: T.W. Flegel and P. Menasveta (eds) Shrimp
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 pathology
LPV does not cause disease or mortality in infected populations.
Histopathology
LPV 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
58
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). LPVinfected tissue does not react with the BS4.5 Probe for IHHNV (Leigh Owens,
personal communication).
Diagnosis
Diagnosis 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).
Transmission
Aspects of the biology of LPV, such as the mode of transmission, range of susceptible
hosts and viability in the external environment are not known
Prevention
LPV is not considered a serious risk to wild or cultured prawns. However, its effect
on prawn defence mechanisms is not known.
References
Lightner, 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 pathology
LOV is not associated with disease and does not appear to be a significant pathogen
of penaeid prawns (Spann et al., 1995).
Histopathology
LOV 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
59
et al., 1995).
Diagnosis
The 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).
Transmission
LOV 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.
Viability
The viability of LOV in the external environment is not known.
References
Spann, 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 pathology
Infected 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.
Diagnosis
Diagnosis 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.
Transmission
Within infected ponds transmission is thought to occur via cannibalism. The viability
of GAV in the external environment is not known.
References
Spann, 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.
60
Spawner-isolated Mortality Virus (SMV)
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 pathology
Prawns 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).
Diagnosis
Definitive 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).
Transmission
Successful feeding trials conducted by Fraser and Owens (1996) indicate that the
primary mode of transmission of SMV is by cannibalism.
References
Anderson 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)
61
Gross Pathology
Hybrid prawns infected with IHHNV and PHRV showed no gross signs of disease.
Histopathology
Rod-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).
Diagnosis
PHRV may be diagnosed by the demonstration by TEM of unusually long, rod-shaped
virus particles in gill haemocytes.
Transmission
Aspects of the biology of PRHV are not known (Owens, 1993).
References
Owens, 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 signs
Mycobacterium 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 Pathology
Multifocal 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
62
tissues of the hepatopancreas, haemocytes, ovary, lymphoid organ, heart, cuticle,
antennal gland, gills and mandibular organ (Lightner et al., 1986; Lightner, 1993).
Histopathology
Mycobacteriosis 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).
Diagnosis
Diagnosis 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).
Transmission
The significance of Mycobacterium spp. as prawn pathogens and their biology when
infecting prawns are not really known.
Treatment
Treatment and control measures are not known.
References
Brock, 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: 403410.
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.
63
Wildlife Dis. 29(4): 591-595.
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 signs
Shell 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 Pathology
Shell 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).
Histopathology
Shell 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.
Diagnosis
Diagnosis of based on the demonstration of brown to black cuticular lesions and/or the
loss of appendage segments (Paynter, 1989).
Transmission
Wounds, 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
64
contagious under poor environmental conditions (Getchell, 1989).
Treatment
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).
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.
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 signs
65
Clinical 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
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 Pathology
Brown 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).
Histopathology
Aeromonas 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).
Diagnosis
Diagnosis 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).
Transmission
Opportunistic 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).
Treatment
Bacterial 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).
References
Anderson, 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.
66
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. 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 signs
L. 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 Histopathology
Fouling 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).
Diagnosis
L. 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).
Transmission
Ecdysis 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.
67
Treatment
Malachite green is an effective treatment for prawn eggs, but not larvae (Brock and
Lightner, 1990). Antibiotics, such as streptomycin, chloramphenicol and
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).
References
Bland, 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.).
68
Clinical signs
Behavioural changes have not been reported for prawns infected with F. solani,
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 Pathology
Lesions 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).
Histopathology
Melanised 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).
Diagnosis
Diagnosis 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).
Transmission
Fusarium 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).
Treatment
Fusariosis 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).
References
Baticados, M.C.L.1988. Diseases of prawns in the Philippines. SEAFDEC Asian
69
Aquaculture 10(1): 1- 8.
Bian, B.Z. and Egusa, S. 1981. Histopathology of black gill disease caused by
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: 12531260.
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
70
from the Californian brown shrimp, Penaeus californiensis. J. Invertebr. Pathol.
27: 77-86.
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 signs
There 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 pathology
Larval 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).
Histopathology
The 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).
Diagnosis
Diagnosis 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).
71
Transmission
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).
Treatment
Various 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 disease
Larval mycosis seldom causes disease in prawns hatcheries with sound management
practices.
References
Armstrong, 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.
72
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 signs
No definitive clinical signs of haplosporidian infections have been reported. Infected
individuals may show poor growth (Lightner et al., 1992).
Gross Pathology
None known.
Histopathology
Haplosporidian 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).
Diagnosis
Diagnosis 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).
Transmission
Not known.
Treatment
Not known.
Present status of disease
Haplosporidiosis is rare and not significant economically.
References
Dykova, I., Lom, J. and Fajer, E. 1988. A new haplosporean infecting the
73
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.
Gregarines
Gregarines (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 signs
There 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 Pathology
Gregarine trophozoites may be visible in the midgut of infected larvae and postlarvae
when viewed under a dissecting microscope (Jones et al., 1994).
Histopathology
There 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).
Diagnosis
Gregarine sporozoites, trophozoites and gametocytes may be observed by light
microscopy in wet mount preparations of midgut contents (Overstreet, 1973).
74
Transmission
Prawns 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).
Treatment
Prawn farmers in Ecuador have found that medicated feed containing anticoccidial
drugs are effective in controlling gregarine infections (Bell and Lightner, 1992).
References
Bell, 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 species
Paranophrys 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
75
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.
Isopods
Isopods 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
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