vps_report

1









SCIENTIFIC REVIEW OF DISEASES OF QUARANTINE SIGNIFICANCE

CARRIED BY BIVALVE MOLLUSCS









JUNE 19, 2000







Prepared for the Australian Quarantine and Inspection Service



By



Veterinary Pathology Services Pty. Ltd.

33 Flemington Street, Glenside, South Australia 5065

Telephone (08) 8372 3700

Facsimile (08) 8372 3766

Email: ruthreuter@vps.com.au









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

2









TABLE OF CONTENTS



TERMS OF REFERENCE 3



PERSONNEL INVOLVED IN THE REVIEW 4



EXECUTIVE SUMMARY 5



INTRODUCTION 6



A. SIGNIFICANT DISEASES OF KNOWN AETIOLOGY 10



1. VIRAL DISEASES 10

1.1 Viral diseases of bivalves in Australia 10

1.2 Significant viral diseases of bivalves not known in Australia 11



2. PROKARYOTE DISEASES 28

2.1 Rickettsia-like and Chlamydia-like diseases in bivalves in Australia 28

2.2 Significant prokaryote diseases of bivalves not known in Australia 30



3. EUBACTERIAL DISEASES 37

3.1 Eubacterial diseases in bivalves in Australia 37

3.2 Significant eubacterial diseases of bivalves not known in Australia 38



4. MYCOTIC DISEASES 45

4.1 Mycotic diseases in bivalves in Australia 45

4.2 Significant mycotic diseases of bivalves not known in Australia 46



5. PROTOZOAN DISEASES 50

5.1 Protozoan diseases of bivalves in Australia 50

5.2 Significant protozoan diseases of bivalves not known in Australia 52



6. SHELL BORING SPONGES 79

6.1 Shell boring sponges in bivalves in Australia 79

6.2 Shell boring sponges in bivalves not known in Australia 80



7. HELMINTH DISEASES 82

7.1 Turbellarids in bivalves in Australia 82

7.2 Turbellarids in bivalves not known in Australia 84



8. PARASITIC CRUSTACEANS 91

8.1 Parasitic crustaceans of bivalves in Australia 91

8.2 Parasitic crustaceans of bivalves not known in Australia 91



B. SIGNIFICANT DISEASES OF UNKNOWN AETIOLOGY 94



C. BIVALVE MOLLUSCS AS PASSIVE VECTORS OF NOXIOUS

ORGANISMS 96







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

3





TABLE OF CONTENTS CONTINUED



D. OVERVIEW AND GENERAL COMMENTS 96



E. APPENDICES 99





TERMS OF REFERENCE AS PROVIDED BY AQIS





To document and analyse information reported in the scientific literature on disease

agents and pests affecting or carried by bivalve molluscs. The report should also

include those agents which do not produce significant pathogenic effects in bivalve

molluscs, but which are known to cause significant pathogenic effects in other hosts.

The review should not include those agents that occur in Australia and are primarily

of public health significance.



Information in the report should be supported by scientific references. The following

information should be provided for each pathogen:



1. Name and taxonomy of the disease agent



2. Agent stability and inactivation data (chemical/thermal processes)



3. Epidemiological features of the disease



 Geographic range and features of distribution (international spread)

 Host range (including prevalence and incidence, resistant strains/species,

lifestage susceptibility and course of infection, habitat (wild or cultured) and

seasonality)

 Morbidity/mortality rates

 Transmission (including route and infectious dose)



4. Host impact

 Tissue tropism

 Brief description of major pathological and biological effects



5. Diagnostics and Disease Control

 Key diagnostic features

 Overview of diagnostic methods, including sensitivity and specificity

 Disease management activities in major producing countries



The study will be carried out in consultation with AQIS. The draft report should be

submitted by 3 May 1999 and a hard copy of the final report, with an electronic

version, should be submitted two weeks after receipt of final comment from AQIS.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

4









PERSONNEL INVOLVED IN THE REVIEW





Dr. Michael Hine, BSc., PhD

Shellfish Consultant, NIWA, Wellington, New Zealand





Dr. Robert Lester, BSc, MSc, PhD

Specialist Parasitologist, University of Queensland.





Dr. Barry Munday, BVSc (Hons), MVSc, DVSc, MACVSc

Specialist Fish Health Consultant, Veterinary Pathology Services





Dr. Ruth Reuter, DVM, Dip Vet Path, PhD, Dip ACVP

Specialist Veterinary Pathologist, Veterinary Pathology Services









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

5









EXECUTIVE SUMMARY





The object of this review was to identify and describe agents and pests carried by

bivalve molluscs, excluding those that are present in Australia and those of public

health significance. The review was conducted by a search of the literature available

and included access to databases held by the team and personal observations of the

members. Where information was available on the points examined, it was entered

into the review. If information was not available on a particular point, this was

recorded.



The review includes 11 viral diseases, 5 eubacterial diseases, 3 mycotic diseases, 16

protozoan diseases, 1 group of shell boring sponges, 6 groups of helminth diseases

and 2 crustacean diseases. One disease of unknown aetiology, Malpeque disease, was

also included as significant.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

6





INTRODUCTION



This review is on the pathogens, pests and parasites of non-viable bivalves that may at

some time be considered for importation into Australia. Non-viable bivalves include

chilled, frozen or brined shucked meat, or chilled meat in the half shell. Imported

species for consumption are oysters, mussels, clams, cockles and scallops. Viable

whole chilled oysters, mussels, clams, cockles and scallops are not included in this

review. Chilled or frozen oysters are traded in the half shell, but not mussels, clams,

cockles or scallops. All groups may be imported chilled or frozen, whole or as

shucked meat. The meat of such product may contain viable viruses, bacteria, fungi,

protozoans, metazoan parasites, crustaceans and turbellarians. The shell may be

infested with boring sponges and polychaetes. Frozen product is likely to be less

dangerous than chilled product because the bacteria and fungi causing disease in

molluscs are not all resistant to freezing and thawing. However frozen product,

especially mussel meat, is likely to be used for fish bait or fed to aquarium or

aquaculture species and thus may introduce pathogens into the aquatic environment.

The effects of brining are too poorly understood to be able to assess the risk of such

product. In view of the overall lack of information on this aspect, a conservative

approach is appropriate.



The countries from which Australia imports molluscs are given in Table 1. This

review will examine which diseases might enter Australia with importation of non-

viable molluscs.



Table 1. Imports of fish products, including molluscs, into Australia (1994-5).*



Country of Origin Quantity (kg) Value ($A)

Argentina 1 10

Canada 800 2824

China 13,630 20,831

Denmark 22,020 16,518

Indonesia 89,994 356,709

Japan 1,604,928 4,269,712

Malaysia 3,471,186 1,768,704

Netherlands 4,533,270 2,890,729

Norway 32 707

New Zealand 5,131,254 3,639,797

Peru 6,319,310 3,008,548

Philippines 482 1,787

Singapore 202,200 284,149

Solomon Islands 20 100

South Africa 91,761 20,172

Taiwan 203,104 613,636

Thailand 1,387,753 1,206,646

United Kingdom 276,141 143,510

United States of America 10,305,104 5,493,881

Vietnam 69,650 275,488

Total All Countries 33,722,640 24,014,458

* Taken from the Report of the National Task Force on Imported Fish and Fish Products, December

1996.





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

7





Bearing in mind that non-viable product is involved, and that pathogens already

reported from Australia are not to be included, the selection of each group was as

follows:-



Viral infections: All serious or potentially serious viral diseases that are unknown in

Australia are included, as viruses are not only likely to remain viable in chilled

product, but may be preserved in frozen product. As the review includes organisms in

molluscs that may cause disease in other groups, infectious pancreatic necrosis virus

(IPNV) which causes serious disease in some salmonids, is included. Viruses which

do not actually infect molluscs, but which may be present in their tissues as a result of

their filtration activities, are not included. However, it should be recognised that

molluscs harvested from, or in close proximity to, finfish farms may carry important

viral pathogens originating from those farms.



Bacterial infections: The bacterial diseases of bivalves reviewed here include the

Rickettsial-like organisms (R-LOs) and chlamydia-like organisms (C-LOs) that cause

serious disease in scallops. Chlamydiosis of bay scallops (Argopecten irradians) has

been associated with mortalities in hatcheries on the east coast of the USA (Leibovitz

1989). The giant extracellular R-LOs of Pacific oysters in Spain, and intranuclear R-

LOs in clams (Siliqua) in western North America, are included as they are unusual R-

LO infections and cause disease. The many R-LOs and C-LOs of other bivalves are

excluded as they are almost always benign, and the taxonomy is confused. A

Mycoplasma-like disease of scallops is also included. Of the eubacterial infections of

bivalves, vibriosis causing brown ring disease, and juvenile oyster disease are

reviewed, but not the many studies reporting opportunistic vibriosis resulting from

poor husbandry or adverse environmental conditions. Summer mortality due to

Nocardia crassostreae infection of Pacific oysters is also reviewed. As with viruses,

it is conceivable that molluscs could act as mechanical vectors for a number of

important bacterial pathogens of fish. This is particularly likely with Aeromonas

salmonicida, where massive numbers of bacteria are released into the water column.



Fungal infections: Three mycoses (Ostracoblabe implexa, Sirolpidium zoophthorum

and a chytrid disease of clams), all of which are primary pathogens, are reviewed.

Other fungal infections reported are usually opportunist saprophytes invading

moribund hosts.



Protozoan infections: The 6 OIE listed notifiable diseases of molluscs that have not

been reported from Australia (Marteilia refringens, Bonamia ostreae, Mikrocytos

mackini, Haplosporidium nelsoni, Haplosporidium costale, Perkinsus marinus), are

reviewed. Four other species, two Marteilia spp., Perkinsus qugwadi, and Perkinsus

atlanticus, associated with large-scale mortalities, are also reviewed. Although there

is molecular evidence that P. atlanticus is closely related to, and may be conspecific

with, Perkinsus olseni from Australia, until the taxonomic status is clarified P.

atlanticus is regarded as a separate species, not reported from Australia. It is now

considered that the organism formerly called Perkinsus karlssoni (McGladdery et al.

1991, 1993, Whyte et al. 1993a, 1993b, 1994) is a composite of two unidentified

species (Goggin et al. 1996), and here it is included as a Perkinsus-like infection of

scallops. As some species of the genera Haplosporidium and Minchinia may cause

disease in their hosts, those infecting oysters, mussels and clams are included.





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

8







The specific identity of other protozoans (Ancistrocoma-like ciliates, Sphenophyra-

like ciliates, gill trichodinids, kidney coccidians [Klossia, Pseudoklossia], gregarines

[Nematopsis spp.], flagellates [Hexamita], intracellular ciliates of mussels), that occur

in and on oysters in Australia (Hine: unpublished information) is unknown. However,

they cause little or no disease, appear to be ubiquitous in ostreid oysters globally, and

therefore are not included here.



Microalgae: Bivalve molluscs ingest microalgae, some of which are toxic to fish. It

is possible that exotic, toxic microalgae could be introduced with non-viable

molluscs. However the danger from this source, in comparison to ballast water, is so

minute that microalgae are not included in this review.



Helminth infections: Digenean trematodes, cestodes and nematodes all infect

bivalves, which usually serve as intermediate hosts. The larval stages of helminths are

unlikely to survive in non-viable product unless chilled for a short time. Even then,

for helminths to establish they must come into contact with the other hosts of their life

cycles, and most are specific to few hosts. They do not multiply in molluscs. However

some of the helminth diseases of quarantine significance in bivalve molluscs are

included in the review.



Polychaetes: Polychaetes of the family Spionidae are major pests in bivalve culture,

as the tubes of these mudworms extend through the shells of bivalves, causing

discolouration and blistering. This group, which includes genera such as Polydora and

Boccardia, is already well represented in Australia (Blake & Kudenov 1978, Wisely

et al. 1979, Nell 1993, Platell & Potter 1996). However recent information indicates

that an unknown sabellid has become a serious pest in the culture of abalone in

California. A review of this group is included here.



Sponges: Sponges of the genus Cliona burrow through dead shells as part of the

natural process of decomposition. Some clionid species will also burrow through the

shells of living bivalves. This does not ordinarily result in high mortalities. However,

some species burrow through the shells of living pearl oysters, causing blemishes in

the nacre, and loss of value to the mother-of-pearl trade. As Australia has a large pearl

farming industry, clionids that attack pearl oyster shells and may be introduced in the

shells of edible oysters but have not yet been reported from Australia, are reviewed.



Turbellarians: Some turbellarians (Urastoma, Paravortex, Stylochus) live in the

digestive tracts or valve chambers of bivalves. They are free living opportunists and

most of the species living with bivalves traded for consumption have no effect on the

health of the host. As several genera of Polycladida have been associated with

epizootics in oysters in other countries, a brief description of these is included.



At the beginning of each group of potential pathogens, the known members of that

group of pathogens already in Australia are briefly summarized.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

9





References



Blake, J.A. & Kudenov, J.D. (1978). The Spionidae (Polychaeta) from southeastern

Australia and adjacent areas with a revision of the genera. Mem. Natl mus. Victoria,

Melb. 39: 171-280.



Goggin, C.L., McGladdery, S.E., Whyte, S.K. & Cawthorn, R.J. (1996). An assessment

of lesions in bay scallops Argopecten irradians attributed to Perkinsus karlssoni

(Protozoa, Apicomplexa). Dis. Aquat. Org. 24: 77-80.



Leibovitz, L. (1989). Chlamydiosis: a newly reported serious disease of larval and

postmetamorphic bay scallops, Argopecten irradians (Lamarck). J. Fish Dis. 12: 125-

136.



McGladdery, S.E., Cawthorn, R.J. & Bradford, B.C. (1991). Perkinsus karlssoni n.sp.

(Apicomplexa) in bay scallops Argopecten irradians. Dis Aquat. Org. 10: 127-137.



McGladdery, S.E., Bradford, B.C. & Scarratt, D.J. (1993). Investigations into the

transmission of parasites of the bay scallop, Argopecten irradians (Lamarck, 1819),

during quarantine introduction to Canadian waters. J. Shellfish. Res. 12: 49-58.



Nell, J.A. (1993). Farming the Sydney rock oyster (Saccostrea commercialis) in

Australia. Rev. Fish. Sci. 1: 97-120.



Platell, M.E. & Potter, I.C. (1996). Influence of water depth, season, habitat and estuary

location on the macrobenthic fauna of a seasonally closed estuary. J. Mar. Biol. Assoc.,

U.K. 76: 1-21.



Whyte, S.K., Cawthorn, R.J., MacMillan, R.J. & Despres, B. (1993). Isolation and

purification of developmental stages of Perkinsus karlssoni (Apicomplexa: Perkinsea), a

parasite affecting bay scallops Argopecten irradians. Dis. Aquat. Org. 15: 199-205.



Whyte, S.K., Cawthorn, R.J. & McGladdery, S.E. (1994). Co-infection of bay scallops

Argopecten irradians with Perkinsus karlssoni (Apicomplexa, Perkinsea) and an

unidentified coccidian parasite. Dis. Aquat. Org. 18: 53-62.



Whyte, S.K., Cawthorn, R.J., McGladdery, S.E., MacMillan, R.J. & Montgomery, D.M.

(1993). Cross-transmission studies of Perkinsus karlssoni (Apicomplexa) from bay

scallops Argopecten irradians to native Atlantic Canadian shellfish species. Dis. Aquat.

Org. 17: 33-39.



Wisely, B., Holliday, J.E., Reid, B.L. (1979). Experimental deepwater culture of the

Sydney rock oyster (Crassostrea commercialis). 3. Raft cultivation of trayed oysters.

Aquaculture 17: 25-32.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

10





A. SIGNIFICANT DISEASES OF KNOWN AETIOLOGY



1. VIRAL DISEASES



1.1. VIRAL DISEASES OF BIVALVES IN AUSTRALIA



The viral diseases of molluscs have been subject to little study throughout the world.

Intranuclear inclusions have been reported from pearl oysters (Pinctada maxima) in

Western Australia (Pass et al. 1988), and occur in other bivalves (Saccostrea cuccullata,

Isognomon isognomum, Pinna bicolor) along the northern coast of Western Australia

(Hine: unpublished information). Pearl oysters in Queensland have intracellular

inclusions thought to be due to papovavirus-like infection, in the mantle epithelium,

(Norton et al. 1993). Herpesviruses have been reported in the haemocytes of adult flat

oysters (Ostrea angasi) in Western Australia (Hine & Thorne 1997), and infect larval

clams (Katelysia) in Tasmania (Judith Handlinger: pers. comm.), and ostreid oysters in

the eastern United States (Farley et al. 1972), France (Nicolas et al. 1992, Comps &

Cochennec 1993) and New Zealand (Hine et al. 1992, 1998). Small RNA viruses

associated with degeneration of the digestive tubule epithelium during digestion, occur

in greenshell mussels (Jones et al. 1996), scallops and toheroa (Hine & Wesney 1997) in

New Zealand. Similar lesions present in scallops (Pecten alba) from Port Phillip Bay,

Victoria, and pearl oysters (P. maxima) in northern Western Australia have not been

associated with mortalities (Hine: unpublished information), and are not included here..

Haematopoietic neoplasms, similar to those reported overseas as being possibly viral

induced, have been seen in Australian oysters (Elston et al, 1992).



As the review covers parasites or diseases that may affect other host groups, infectious

pancreatic necrosis virus, causing disease in juvenile cultured salmonids, is included

here.



References



Comps, M. & Cochennec, N. (1993). A herpes-like virus from the European oyster

Ostrea edulis L. J. Invertebr. Pathol. 62: 201-203.



Elston, R.A., Moore, J.D. & Brooks, K (1992). Disseminated neoplasia of bivalve

molluscs. Reviews in Aquatic Sciences 6:405 – 466.



Farley, C.A., Banfield, W.G., Kasnic, G. & Foster, W.S. (1972). Oyster herpes-type

virus. Science 178: 759-760.



Hine, P.M. & Thorne, T. (1997). Replication of herpes-like viruses in haemocytes of

adult flat oysters Ostrea angasi (Sowerby, 1871): an ultrastructural study. Dis. aquat.

Org. 29: 189-196.



Hine, P.M. & Wesney, B. (1997). Virus-like particles associated with cytopathology in

the digestive gland epithelium of scallops Pecten novaezelandiae Reeve, 1853 and

toheroa Paphies ventricosum (Gray, 1843). Dis. aquat. Org. 29: 197-204.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

11





Hine, P.M., Wesney, B. & Besant, P. (1998). Replication of herpes-like viruses in larvae

of the flat oyster Tiostrea chilensis at ambient temperatures. Dis. Aquat. Org. 32: 161-

171.



Hine, P.M., Wesney, B. & Hay, B.E. (1992). Herpesviruses associated with mortalities

among hatchery-reared larval Pacific oysters Crassostrea gigas. Dis. aquat. Org. 12:

135-142.



Jones, J.B., Scotti, P.D., Dearing, S.C. & Wesney, B. (1996). Virus-like particles

associated with marine mussel mortalities in New Zealand. Dis. Aquat. Org. 25: 143-

149.



Nicolas, J.L., Comps, M. & Cochennec, N. (1992). Herpes-like virus infecting Pacific

oyster larvae. Bull. Eur. Assoc. Fish Pathol. 12: 11-13.



Norton, J.H., Shepherd, M.A. & Prior, H.C. (1993). Papovavirus-like infection of the

golden-lipped pearl oyster, Pinctada maxima, from the Torres Strait, Australia. J.

Invertebr. Pathol. 62: 198-200.



Pass, D.A., Perkins, F.O. & Dybdahl, R. (1988). Viruslike particles in the digestive

gland of the pearl oyster Pinctada maxima. J. Invertebr. Pathol. 51: 166-167.







1.2 SIGNIFICANT VIRAL DISEASES OF BIVALVES NOT KNOWN IN

AUSTRALIA



1.2.1 OYSTER VELAR VIRUS DISEASE (OVVD)



1. Name and taxonomy of disease agent: Oyster velar virus disease, OVVD, blister

disease; icosahedral DNA virus, possibly an iridovirus.



2. Agent stability and inactivation data: Not known.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Reported from Washington

State, U.S.A. (Elston 1979, 1993), but current distribution unknown. Has not

been observed in recent years.



 Host range: Larvae of Crassostrea gigas.



 Morbidity/mortality rates: Mortality rate of 100% has been reported in

hatcheries.



 Transmission: Thought to be horizontal, direct.



4. Host impact:



 Tissue tropism: The epithelium of the velum.





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

12







 Brief description of major pathological and biological effects: Occurs over

March to May, and sometimes into the Northern summer (July to August) in

hatchery-reared larvae 150 um shell length and 10 days old. Hypertrophy and

loss of cilia on infected cells, during which lesions may appear as blisters.



5. Diagnostics and disease control



 Key diagnostic features: There are no specific clinical signs that can be used

diagnostically. Mortalities begin among larvae at 10 days post-spawn. Infected

larvae become unable to swim normally, and drop in the water column. Dying and

dead larvae are seen at the bottom of the tank. The course of the disease is

approximately 4 days. Similar signs are present in larval mortalities due to

herpesvirus infection, although the latter may start 6 days after spawning.



 Overview of diagnostic methods, including sensitivity and specificity: Primary

diagnosis is by histology, with confirmatory diagnosis by electron microscopy.

Under the light microscope, intracytoplasmic inclusion bodies, that are spherical,

dense and basophilic, 1.2-2.4 m in diameter, occur in velar cells, and

occasionally in velar-supporting esophageal and oral epithelia. They are rarely

seen in mantle epithelium. They become less basophilic as virions form.

Definitive diagnosis is made on the basis of electron microscopy showing

icosahedral particles 228 ± 7 nm in diameter with a dense core separated from the

capsid by a moderately dense zone.



 Disease management activities in major producing countries: Destruction of

infected stocks, safe disposal of water, disinfection of tanks and equipment.



References



Elston, R. (1979). Virus-like particles associated with lesions in larval Pacific oysters

(Crassostrea gigas). J. Invertebr. Pathol. 33: 71-74.



Elston, R. (1993). Infectious diseases of the Pacific oyster, Crassostrea gigas. Ann. Rev.

Fish Dis. 3: 259-276.





1.2.2 GILL NECROSIS VIRUS DISEASE (GNV)



1. Name and taxonomy of disease agent: Gill disease of Portuguese oysters, Gill

necrosis virus disease (GNV), Maladie des branchies. Attributed to an iridovirus on

morphological grounds, but identity has yet to be confirmed.



2. Agent stability and inactivation data: Not known



3. Epidemiological features of the disease:



 Geographic range and features of distribution: France, Portugal, Spain and the

U.K.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

13





 Host range: Crassostrea gigas and Crassostrea angulata (C. angulata may be a

subspecies of C. gigas).



 Morbidity/mortality rates: From 1966-1973, gill necrosis occurred in 70-80%

of oysters, and mortalities occurred in  40% of oysters (Comps 1988).



 Transmission: Unknown, but as the gill is the site of infection, horizontal direct

transmission is probable..



4. Host impact:



 Tissue tropism: Haemocytes infiltrating the gills.



 Brief description of major pathological and biological effects: Extensive gill

erosion. Initial signs of yellow spots on the gills progress to brown discolouration

with associated necrosis and degeneration, leaving a perforation or V-shaped

indentation, if the lesion occurs at the edge of the gill. Yellow or green pustules

may also occur on the mantle or adductor muscle.



5. Diagnostics and disease control



 Key diagnostic features: Gill indentations, yellow to brown spots on the gills, V-

shaped perforations. Similar perforations occur in infection with the protist

pathogen, Bonamia ostreae.



 Overview of diagnostic methods, including sensitivity and specificity:

Preliminary diagnosis may be made on the presence of two or more of the gross

signs given above. Histology on the gill or labial palp reveals massive haemocytic

infiltration around lesions containing polymorphic hypertrophic cells, and

hypertrophic globular cells with basophilic cytoplasmic inclusions. Such

haemocytoses are non-specific, and frequently occur in a variety of infections.

More definitive diagnosis is made by electron microscopy showing

intracytoplasmic inclusions in the haemocytes in the gills, or in gill cells. These

inclusions contain virogenic stroma and icosahedral virions 380 nm in diameter.

However the virus cannot be readily distinguished from that associated with

haemocyte infection virus disease.



 Disease management activities in major producing countries: Controls were

exercised on movements, but this disease caused farmers to switch from C.

angulata to C. gigas, which appeared to be more resistant to infection. The disease

has not been seen for ~20 years, and it has been recommended that it be removed

from the OIE list of Significant Diseases.



Reference



Comps, M. (1988). Epizootic diseases of oysters associated with viral infections. Am.

Fish. Soc. Spec. Publ. 18: 23-37.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

14





1.2.3 HAEMOCYTIC INFECTION VIRUS DISEASE



1. Name and taxonomy of disease agent: Haemocytic infection virus disease.

Attributed to an iridovirus on morphological grounds, but identity has yet to be

confirmed.



2. Agent stability and inactivation data: Not known.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Caused epizootics among C.

angulata in France from 1970 to 1973. A similar disease caused mortalities

among C. gigas in France in 1977. Was reported from Spain in 1974. An outbreak

occurred among C. angulata spat in a hatchery in northern France in 1983. The

disease has not been seen since.



 Host range: Crassostrea angulata and Crassostrea gigas.



 Morbidity/mortality rates: Described as mass mortalities (Comps 1988).



 Transmission: Presumed to be horizontal and direct.



4. Host impact:



 Tissue tropism: Haemocytes infiltrating the gills..



 Brief description of major pathological and biological effects: Atrophy and

weakness of the adductor muscle, leading to gaping and death.



5. Diagnostics and disease control



 Key diagnostic features: Atypical hemocytes with pycnotic nuclei and round

basophilic intracytoplasmic inclusion bodies (2 – 3 m in diameter) in the

connective tissue.



 Overview of diagnostic methods, including sensitivity and specificity:

Presumptive diagnosis is based on histology showing an acute inflammatory

response of atypical haemocytes containing inclusion bodies. More definitive

diagnosis is confirmed by electron microscopy, showing icosahedral particles 380

nm in diameter, associated with cytoplasmic viral protein. At present there is no

means of differentiating this virus from that causing Gill disease (GNV).



 Disease management activities in major producing countries: Controls were

exercised on movements, but this disease caused farmers to switch from C.

angulata to C. gigas, which appeared to be more resistant to infection. The close

similarities in cellular tropism, size and appearance of virions, and occurrence in

the same hosts at about the same time of gill necrosis virus and haemocytic

infection virus, suggest that they were two variants of the same disease.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

15





Reference



Comps, M. (1988). Epizootic diseases of oysters associated with viral infections. Am.

Fish. Soc. Spec. Publ. 18: 23-37.





1.2.4 VIRAL GAMETOCYTIC HYPERTROPHY



1. Name and taxonomy of disease agent: Viral gametocytic hypertrophy. Thought to

be caused by a papillomavirus-like papovavirus (Bower et al. 1994).



2. Agent stability and inactivation data: Not known.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Eastern Canada and the eastern

and western coasts of the U.S.A., Japan, Korea. Papovavirus-related changes of

this type have been recorded in Pacific oysters C. gigas in Australia (Wilson et al,

1993, Wilson, 1993).



 Host range: Crassostrea virginica, Crassostrea gigas, Saccostrea commercialis,

Crassostrea rhizophorae, Ostreola conchaphila.



 Morbidity/mortality rates: Does not cause mortalities, or have a significant

effect on the host.



 Transmission: Unknown, but as the virus replicates in the nucleus of ova, it is

likely transmission is vertical.



4. Host impact:



 Tissue tropism: Ovacytes.



 Brief description of major pathological and biological effects: Massive

hypertrophy of the nucleus of individual ova.



5. Diagnostics and disease control



 Key diagnostic features: Massively enlarged ovacytes.



 Overview of diagnostic methods, including sensitivity and specificity:

Histology readily reveals the enlarged ovacytes.



 Disease management activities in major producing countries: None.



Reference



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994). Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4: 1-199.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

16





Wilson, J., Jandlinger, J. & Sumner, C.E. (1993). The health status of Tasmania’s

bivalve shellfish. Sea Fisheries Division Technical Report No. 47, Tasmania.



Wilson, J. (1993). The health status of two species of Tasmanian farmed shellfish,

Crassostrea gigas (Thunberg, 1793) and Ostrea angasi (Sowerby, 1871). Masters

thesis, University of Tasmania at Launceston, 189 pp.





1.2.5 AKOYA OYSTER DISEASE



1. Name and taxonomy of disease agent: Akoya oyster disease, thought to be a

viral disease of the akoya oyster (Pinctada fucata martensii).



2. Agent stability and inactivation data: The mortalities are greatest at >250C, and

decline significantly at 200C (Miyazaki et al 1998, 1999).



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Western regions of Japan

and Hainan Dao, off the coast of China.



 Host range: The temperate pearl oyster, Pinctata fucata martensii.



 Morbidity/mortality rates: In 1996 and 1997 the annual mortality was 400

million akoya oysters in the western regions of Japan, constituting 50% of

Japanese stocks (Miyazaki et al 1998, 1999). Morbidity is unreported.



 Transmission: Cohabitation of infected and uninfected oysters results in

transmission of the disease, suggesting direct horizontal transmission.



4. Host impact:



 Tissue tropism: The principal organ infected is the adductor muscle, which

shows loss of myofibrils and weakening, with concomitant haemocyte

infiltration.



 Brief description of major pathological and biological effects: Relatively

little is known about this disease, and the aetiology is far from certain. When

infected oysters are cohabited with healthy oysters, the previously healthy

oysters show browning in the area of the heart, in the adductor muscle, lips of

the mantle lobe and body. The degree of browning is greatest at high

temperatures, and least at low temperatures. The adductor muscle weakens

and the oysters gape. It is unclear whether death is due to progression of the

disease, or invasion of the gaping oyster by micropredators. Histologically the

oyster appears normal, except for accumulations of brown cells in the heart

area, and thinning and loss of myofibrils from the adductor muscle. There is a

moderate haemocytosis, with infiltration of the muscle by haemocytes.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

17





5. Diagnostics and disease control:



 Key diagnostic features: Browning of the heart region, adductor muscle,

mantle lobe and body; reduced growth, gaping and haemocyte infiltration into

the adductor muscle. The myofibrils show atrophy and decrease in number or

show swelling and vacuolation. The epithelia of the digestive tubules are

frequently flattened, enlarging the lumen, suggesting lack of digestion.



 Overview of diagnostic methods, including sensitivity and specificity: The

single research group to report studies to date claim they have grown the

aetiological agent in EK-1 (eel kidney) and EPC (epithelioma papillosum

cyprini) cell lines. The inocula were ultrafiltrates (450 nm) of homogenised

affected oyster tissues. At 250C, CPE (karyopyknosis and cytoplasmic

vacuolation) began after 7-10 days, and was completed by 14-20 days. The

virions observed by transmission electron microscopy (TEM) were 25-33 nm

in diameter and rounded, with a spiked surface, and occurred in the

cytoplasmic vacuoles. They were negative for DNA, and it was concluded

that they were associated with a small RNA virus. To fulfil Koch’s Postulates,

0.5 ml of culture medium was inoculated into apparently healthy oysters.

Inoculation was used rather than exposure to the virus, because the research

was carried out in winter, when the disease is not normally active. The

inocula contained 105-6 virions/ml, and 5 strains of the virus were inoculated.

The basis on which the strains were distinguished was not given. The

inoculated oysters showed sluggish contraction of the adductor muscle after 7

days, and after 10 days, mortality was 50-100%. Clinical signs were the same

as those in naturally affected oysters. Moribund oysters were fixed and the

virions visualised under TEM. The virus was re-isolated in the fish cell lines.



 Disease management activities in major producing countries: Neither

Japan nor China have any legislation that can be used to control movements,

and movements of live bivalves occur daily between Japan, Korea and China.

As an interim measure while laws are promulgated, attempts are being made

to educate the fish farmers on the risks of moving live animals for relaying in

water.



Note:



1. A paper has recently been published by Chinese scientists, reporting that the

mortalities in another species of pearl oyster, the silverlip Pinctada maxima, in

Hainan Province, are due to a Rickettsia-like organism (R-LO) (Wu & Pan 1999).

However, it appears that they may have mis-identified normal eosinophilic

granulocytic cells in the epithelia as R-LO inclusions. The latter are characteristically

basophilic inclusions in bivalves. Although from TEM it is clear that R-LOs are

present, it appears that they are at the same low levels that are normally seen in other

bivalves.



2. Japanese pearl seeding technicians visit Australia every year, and bring their

surgical instruments with them Western Australia is now requiring that the

instruments be disinfected before use.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

18





References:



Miyazaki, T., Goto, K., Kobayashi, T. & Miyata, M. (1998) An emergent virus

disease associated with mass mortalities in Japanese pearl oysters Pinctada futaka

martensii. In: Proceedings of the VIIth International Colloquium on Intervertebrate

Pathology and Microbial Control. Sapporo, Japan, August 23-28th 1998, pp. 154-159.



Miyazaki, T., Goto, K., Kobayashi, T., Kageyama, T. & Miyata, M. (1999). Mass

mortalities associated with a virus disease in Japanese pearl oysters Pinctada fucata

martensii. Dis. Aquat. Org. 37:1-121.



Wu, X & Pan, J (1999) Studies on Rickettsia-like organism disease of the tropical

marine pearl oyster I: The fine structure and morphogenesis of Pinctada maxima

pathogen Rickettsia-like organism. J.Invertebr.Pathol. 73:162-172.





1.2.6 MASS MORTALITY OF JAPANESE OYSTER CRASSOSTREA GIGAS



1. Name and taxonomy of disease agent: Unspecified virus (Miyazaki, 1999)



2. Agent stability and inactivation data: Unknown. Characteristics under study.



3. Epidemiological features of the disease;



 Geographic range: Japan.



 Host range: Crassostrea gigas.



 Morbidity/mortality rates: Mass mortality of C. gigas reported.



 Transmission: Reproduced by injection of cultured virus.



4. Host impact:



 Tissue tropism: Glycogen-storing cells and adductor muscle.



 Brief description of major pathological and biological effects: Extensive

necrosis and haemocyte infiltration of glycogen-storing cells. Slight necrosis

of adductor muscle.



5. Diagnostics and disease control:



 Key diagnostic features: Necrosis of adductor muscle and glycogen-storing

cells.



 Overview of diagnostic methods, including sensitivity and specificity:

Cytopathic effect noted in EPC cells at three weeks of incubation at 250 C.

Disease reproduced by injection of cultured virus.



 Disease management activities in major producing countries: None.





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

19







Reference



Miyazaki, T (1999). A new viral disease causing mass mortality in Japanese oyster

Crassostrea gigas. Abstr OP 36. 4th Symposium on Diseases in Asian Aquaculture.

Cebu City, Philippines. Paper to be published in Asian Fisheries Science, Vol. 13.





1.2.7 HERPES VIRUS INFECTION OF LARVAL C. GIGAS



1. Name and taxonomy of disease agent: Herpes virus of larval Crassostrea gigas.



2. Agent stability and inactivation data: Unknown.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: France and New Zealand.

Agent may be widespread but disease is only reported at relatively high water

temperatures.



 Host range: Larval C. gigas.



 Morbidity/mortality rages: High,  100 %.



 Transmission: Method unknown.



4. Host impact:



 Tissue tropism: Connective tissue of digestive gland and mantle.



 Brief description of major pathological and biological effects: Macroscopic

lesions non-specific. Rapid onset of mass mortality.



5. Diagnostics and disease control



 Key diagnostic features: Diagnosis is based on the presence of intranuclear

inclusion bodies in haemocytes in the connective tissues. Electron microscopy

can be used to demonstrate typical herpes virus particles 7-100 nm in

diameter.



 Disease control: As the virus is probably widespread, disease control is

difficult if it is not possible to relocate hatcheries to cooler localities.





References



Hine, P.M., Wesney, B. & Hay, B.E. (1992). Herpes-virus associated with mortalities

among hatchery-reared larval Pacific oysters, Crassostrea gigas. Dis.Aquat.Org.

12:135-142.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

20





Nicholas, J.L., Comps, M. & Cochennec, N. (1992). Herpes-like virus infecting

Pacific oyster larvae, Crassostrea gigas. Bull.Eur.Ass.Fish Pathol. 12:11-13.







1.2.8 INFECTIOUS PANCREATIC NECROSIS VIRUS (IPNV)



1. Name and taxonomy of disease agent: Infectious Pancreatic Necrosis Virus

(IPNV) (Birnaviridae), causing sometimes fatal disease in salmonids. Although a low-

virulence birnavirus serologically related to the classical IPN viruses has been isolated

from fish in Tasmanian waters, this review will use the OIE definition of IPN as “a

highly contagious viral disease of young fish of salmonid species” and therefore,

IPNV will be considered.



2. Agent stability and inactivation data:



Temperature: Survives 3-4 h at 65°C to ~10 min at 80°C (Whipple & Rohovec 1994),

Inactivated after 16 h at 60°C (MacKelvie and Desautels 1975, Gostling & Gould

1981), and in silage at 60°C (Smail et al. 1990). Well preserved at -80°C, resistant to

drying (MacKelvie & Desautels 1975).



Salinity: Viable in freshwater, estuarine water, and seawater, longer at 15°C than

20°C (Toranzo and Hetrick 1982). Best survival in estuarine water but slowly

inactivated, probably by microflora (Toranzo et al. 1983), in aquatic environments

(Wolf et al. 1969, Desautels & MacKelvie 1975, Yoshimizu et al. 1986, Smail &

Munro 1989).



pH: Viable for 14 days at pH 3.8-4.3, survival reduced by heat. Slightly resistant to

acid pH (2.5) but labile to alkaline pH (12.2) (Vestergard-Jørgensen 1974). Survives

70% in wild clams (Paillard et al. 1994). Mortalities tend

to occur in winter and in spring (5-20oC) (Ford & Paillard 1994).



 Transmission: Horizontal, direct.



4. Host impact:



 Tissue tropism: Surface of clam meat (~90% of V. tapetis), mantle, periostracal

lamina (Allam et al. 1996).



 Brief description of major pathological and biological effects: Brown ring

disease is a disease caused by strains (Castro et al. 1997a) of Vibrio tapetis

(Borrego et al. 1996, Castro et al. 1997b), which disrupt the shell calcification

process. Tissue lesions are not systematically observed in diseased clams.

Alterations of the digestive gland and the mantle are detected in the more severe





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

39





stages of the disease. In all diseased clams, however, the periostracal lamina

shows alterations. It is invaded by cell debris and bacteria, and areas of darker

melanin-like pigmentation are observed (Paillard et al. 1994). Experimental

inoculation has shown that, one week after inoculation of V. tapetis, two types of

deposit are visible: small pustules generally localized on the surface of the inner

shell layer and a continuous film observed on the surface of the prismatic shell

layer. These first deposits are made up of periostracal and degraded fibrous layers.

From 1-4 weeks after inoculation the deposit becomes progressively wider,

thicker and is invaded by many bacteria (Paillard & Maes 1995).



5. Diagnostics and disease control



 Key diagnostic features: As the common name suggests, the most characteristic

feature of this disease is a brown ring laid down on the shell nacre in association

with abnormalities of the conchiolin. However, a Vibrio that causes juvenile

oyster (Crassostrea virginica) disease on the eastern coast of the USA (see

below), and V. harveyi infection that causes mortalities in pearl oysters (Pinctada

maxima) in Western Australia, both cause conchiolin abnormalities and, in pearl

oysters, similar brown stains on the nacre (Perkins 1996). Therefore brown

staining alone cannot be regarded as pathognomonic for this disease.



 Overview of diagnostic methods, including sensitivity and specificity:

Monoclonal antibodies to V. tapetis have been developed (Noel et al. 1991), and

used to develop a nitrocellulose membrane immunoassay (colony-blot ELISA)

which shows 100% specificity and sensitivity (Noel et al. 1996).



 Disease management activities in major producing countries: Controls have

been placed on movements of infected stocks to minimise the risk of spread.

Furazolidone administered at 10 mg/L for 3 days gives 100% protection against V.

tapetis (Noel et al. 1991).



References



Allam, B., Paillard, C & Maes, P. (1996). Localization of the pathogen Vibrio P1 in

clams affected by brown ring disease. Dis. Aquat. Org. 27: 149-155.



Borrego, J.J., Castro, D., Luque, A., Paillard, C., Maes, P., Garcia, M.T. & Ventosa, A.

(1996). Vibrio tapetis sp. nov., the causative agent of the brown ring disease affecting

cultured clams. Int. J. Syst. Bacteriol. 46: 480-484.



Castro, D., Romalde, J.L., Vila, J., Magarinos, B., Luque, A. & Borrego, J.J. (1997a).

Intraspecific characterization of Vibrio tapetis strains by use of pulsed-field gel

electrophoresis, ribotyping, and plasmid profiling. Appl. Environ. Microbiol. 63: 1449-

1452.



Castro, D., Santamaria, J.A., Luque, A., Martinez-Manzarenares, E. & Borrego, J.J.

(1997b). Determination of the etiological agent of brown ring disease in southwestern

Spain. Dis. Aquat. Org. 29: 181-188.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

40





Ford, S.E. & Paillard, C. (1994). A comparison of juvenile oyster disease in the USA

and brown ring disease of Manila clams in Europe. J. Shellfish Res. 13: 314.



Noel, T., Aubree, E., Blateau, D., Mialhe, E. & Grizel, H. (1992). Treatments against

the Vibrio P1, suspected to be responsible for mortalities in Tapes philippinarum.

Aquaculture 107: 171-174.



Noel, T., Boulo, V., Mialhe, E., Nicolas, J.L., DePauw, N & Joyce, J. (1991).

Diagnosis of the “brown ring” disease in Tapes philippinarum with monoclonal

antibodies. Spec. Publ. Eur. Aquacult. Soc. 14: 234-235.



Noel, T., Nicolas, J-L., Boulo, V., Mialhe, E. & Roch, P. (1996). Development of a

colony-blot ELISA assay using monoclonal antibodies to identify Vibrio P1

responsible for “brown ring disease” in the clam Tapes philippinarum. Aquaculture

146: 171-178.



Paillard, C. & Maes, P. (1995). The brown ring disease in the Manila clam, Ruditapes

philippinarum: 2. Microscopic study of the brown ring syndrome. J. Invertebr.

Pathol. 65: 101-110.



Paillard, C., Maes, P. & Oubella, R. (1994). Brown ring disease in clams. Ann. Rev.

Fish Dis. 4: 219-240.



Perkins, F.O. (1996). Shell disease in the gold lip pearl oyster, Pinctada maxima and

the eastern oyster, Crassostrea virginica. Aquat. Living Resour. 9: 159-168.







3.2.2 JUVENILE OYSTER DISEASE (JOD)



1. Name and taxonomy of disease agent: Juvenile Oyster Disease (JOD), Vibrio sp.

(Eubacteria: Vibrionaceae). There is no general agreement as to the cause of this

disease, some claiming it is caused by a Vibrio sp. (Lee et al. 1996). Others claim

that there is no evidence of a bacterial aetiology, and that a protist may be involved

(Lewis et al. 1996a, 1996b). A broad overview of the published data suggests that the

disease is due to a Vibrio, but that there are complex contributory factors, including

stock genetics and environmental conditions.



2. Agent stability and inactivation data: Disease transmission declines with

decrease in salinity below 20‰ (Lewis & Farley 1994). Temperatures below 220C

and salinities below 18 ‰ inhibit transmission (Lewis et al. 1996b).



3. Epidemiological features of the disease:



 Geographic range and features of distribution: JOD occurs intermittently along

the coast of the eastern USA, particularly along the northeastern states (Maine,

Rhode Island, New England, New York, Maryland).



 Host range: Eastern oysters (Crassostrea virginica).







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

41





 Morbidity/mortality rates: Up to 100% mortalities around New England and

New York in the late 1980’s (Lewis et al. 1996), 50-100% in other stocks (Bricelj

et al. 1992, Lee et al. 1996). Usually 40-80%, but as low as 8% at some sites

(Farley & Lewis 1995). Oysters 30 mm length are affected (Lewis et al. 1996b).



 Transmission: Horizontal, direct.



4. Host impact:



 Tissue tropism: Epithelial cells on the edge of the mantle.



 Brief description of major pathological and biological effects: Infection of the

mantle epithelium by vibrios or ill-defined protists, results in mantle retraction,

possibly due to bacterial exotoxins. Before dying, afflicted oysters also exhibit

generalized signs of stress, such as slow growth and ring-like conchiolin deposits,

which include bacteria and cell debris (Lee et al. 1996). Lesions may occur on the

mantle surface (Bricelj et al. 1992).



5. Diagnostics and disease control



 Key diagnostic features: The most consistent signs appear to be mantle

retraction, ring-like conchiolin deposits, reduced growth, and the small size

groups (30 mm) of oyster infected.



 Overview of diagnostic methods, including sensitivity and specificity: As the

aetiological agent is still debatable, diagnosis depends on recognition of the suite

of signs typical of this disease. Consequently, both sensitivity and specificity of

diagnosis are likely to be low. No presumptive aetiological agent has been

observed under the TEM, and it is assumed that bacterial toxins cause the

pathology.



 Disease management activities in major producing countries: Controls on

movements of affected stocks, and identification of hatcheries from which

affected stocks originate.





References



Bricelj, V.M., Ford, S.E., Borrero, F.J., Perkins, F.O., Rivara, G., Hillman, R.E., Elston,

R.A. & Chang, J. (1992). Unexplained mortalities of hatchery-reared, juvenile oysters,

Crassostrea virginica (Gmelin). J. Shellfish Res. 11: 331-347.



Farley, C.A. & Lewis, E.J. (1995). Juvenile oyster disease studies 1994: Epizootiology,

geographic occurrence. J. Shellfish Res. 14: 241-242.



Lee, M., Taylor, G.T., Bricelj, V.M., Ford, S.E. & Zahn, S. (1996). Evaluation of Vibrio

spp. and microplankton blooms as causative agents of juvenile oyster disease in

Crassostrea virginica (Gmelin). J. Shellfish Res. 15: 319-329.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

42





Lewis, E.J. & Farley, C.A. (1994). Effects of salinity and selected treatments on

juvenile oyster disease. J. Shellfish Res. 13: 316.



Lewis, E.J., Farley, C.A., Baya, A.M. & Small, E.B. (1996a). Juvenile oyster disease

– transmission and bacteriological studies. J. Shellfish Res. 15: 516.



Lewis, E.J., Farley, C.A., Small, E.B. & Baya, A.M. (1996b). A synopsis of juvenile

oyster disease (JOD) experimental studies in Crassostrea virginica. Aquat. Living

Resour. 9: 169-178.





3.2.3 SUMMER MORTALITY



1. Name and taxonomy of disease agent: Summer mortality, Pacific oyster

(Crassostrea virginica) nocardiosis (PON), - Nocardia crassostreae

(Actinomycetales).



2. Agent stability and inactivation data: Unknown, but likely to be similar to other

Nocardia spp.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: West coast of North America

from the Strait of Georgia, British Columbia, to California, and in Japan

(Matsushima Bay) (Bower et al 1994). A problem in shallow muddy embayments

with elevated temperatures and nutrients (Friedman 1991). Peak prevalence in

September (Friedman et al. 1991).



 Host range: Pacific oysters (Crassostrea gigas), particularly males and immature

oysters.



 Morbidity/mortality rates: Mortalities 35 % in some localities.



 Transmission: Unknown, probably horizontal and direct.



4. Host impact:



 Tissue tropism: In tissues of the mantle, gill, adductor muscle, heart.



 Brief description of major pathological and biological effects: Small yellow-

green pustules are seen grossly on the surface of the mantle, gill, adductor muscle

and heart. However the hyphae-like filaments of these actinomycetes ramify

through the tissues of most of the organs. It appears that oysters that are close to

muddy sediments, and that are stressed by elevated temperatures, become infected

by these opportunistic bacteria present in the sediment.



5. Diagnostics and disease control



 Key diagnostic features: Round yellow-to-green pustules 1 cm in diameter

occur on the surface of the mantle, gills, adductor muscle or heart. The hyphae-





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

43





like filaments in underlying tissues are Gram-positive, PAS positive, and acid fast.

The pathogen grows slowly in Lowenstein-Jensen medium.



 Overview of diagnostic methods, including sensitivity and specificity:

Histopathology and specific stains. Sensitivity of all methods probably moderate.

Specificity of culture high, and probably moderate for all direct staining and

histology.



 Disease management activities in major producing countries: Sensitive to

isoniazid and mitomycin C (Friedman 1991). Controls are not possible, since the

bacteria probably occur naturally and widely distributed in mud along the affected

coast.



References



Bower, S.M., McGladdery, S.E., Price, I.M. (1994) Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann.Rev.Fish Dis. 4:1-199.



Friedman, C.S. (1991). Nocardiosis of the Pacific oyster, Crassostrea gigas Thunberg.

Diss. Abst. Int. Pt. B – Sci. & Eng. vol. 51 (11): 135pp.



Friedman, C.S., Beattie, J.H., Elston, R.A. & Hedrick, R.P. (1991). Investigation of the

relationship between the presence of a Gram-positive bacterial infection and summer

mortality of the Pacific oyster, Crassostrea gigas Thunberg. Aquaculture 94: 1-15.





3.2.4. HINGE LIGAMENT DISEASE



1. Name and taxonomy of disease agent: Hinge ligament disease caused by

Cytophaga-like, Flexibacter-like, or Flavobacterium-like bacteria.



2. Agent stability and inactivation data: Unknown, but probably similar to other

Cytophagia-like and Flavobacterium-like bacteria.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Regarded as ubiquitous (Bower

et al. 1994). Absence of reports is more likely due to lack of investigation than

absence of the organism(s). Cytophaga or Flexibacter-like bacteria have been

isolated from Pacific oyster hinge lesions in Australia (Handlinger, unpublished).



 Host range: Juvenile oysters (Crassostrea gigas, Crassostrea virginica, Ostrea

edulis) and clams (Mercenaria mercenaria, Ruditapes philippinarum, Siliqua

patula), probably many more species.



 Morbidity/mortality rates: 90% mortality under crowded hatchery conditions.



 Transmission: Horizontal, direct.



4. Host impact:





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

44







 Tissue tropism: Infects the hinge ligament, mantle and connective tissue (Dungan

& Elston 1988).



 Brief description of major pathological and biological effects: Causes

liquefaction of the hinge ligament, which progresses more rapidly with increasing

temperature. In one report the ligament resilia of 93% of oysters contained erosive

lesions, but only 20% of the ligament tensilia had lesions. Erosive lesions were

associated with clumps of elongated bacteria, sometimes causing perforation of

the hinge ligament (Dungan & Elston 1988). The long axes of bacteria were

oriented at right angles to the lesion margins.



5. Diagnostics and disease control



 Key diagnostic features: Lesions in the hinge filament associated with long (3

m) filamentous bacteria (Dungan & Elston 1988)



 Overview of diagnostic methods, including sensitivity and specificity:

Isolation and growth on low nutrient agars result in mixed colonies. The

Cytophaga-like bacterium that is associated with the lesions is the only species

that is capable of sustained growth with hinge filament matrix as the sole source

of organic carbon and oxygen (Dungan et al. 1989). Diagnosis by histology is

likely to be of moderate specificity but low sensitivity.



 Disease management activities in major producing countries: None. The

aetiological agent is ubiquitous in the marine environment. However, initially

healthy stock in environments with minimal stress (adequate nutrition, good water

quality, no over-crowding) seldom develop the disease.



References



Dungan, C.F. & Elston, R.A. (1988). Histopathological and ultrastructural

characteristics of bacterial destruction of the hinge ligaments of cultured juvenile

Pacific oysters, Crassostrea gigas. Aquaculture 72: 1-14.



Dungan, C.F., Elston, R.A. & Schiewe, M.H. (1989). Evidence for colonization and

destruction of hinge ligaments in cultured juvenile Pacific oysters (Crassostrea gigas)

by cytophaga-like bacteria. Appl. Environ. Microbiol. 55: 1128-1135.





3.2.5 BACTERIAL ABSCESS DISEASE (BAD)



1. Name and taxonomy of disease agent: Bacterial abscess disease (BAD), brown

spot caused by unidentified Gram-positive bacteria.



2. Agent stability and inactivation data: Not reported.



3. Epidemiological features of the disease:









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

45





 Geographic range and features of distribution: Northeastern U.S., and Atlantic

Canada.



 Host range: Wild Placopecten magellanicus.



 Morbidity/mortality rates: Prevalence of 5-10%. Disruption of the adductor

muscle leads to death (Bower et al. 1994).



 Transmission: Not reported.



4. Host impact:



 Tissue tropism: Adductor muscle.



 Brief description of major pathological and biological effects: The systemic

bacterial infection results in brown spots on the adductor muscle. Histologically,

focal haemocyte infiltration throughout the tissues is associated with aggregations

of Gram-positive bacteria. In the adductor muscle, muscle fibres show disruption

associated with haemocyte infiltration, necrosis and abscessation. The main

impact is to render the scallops unmarketable.



5. Diagnostics and disease control



 Key diagnostic features: Brown spots (abscesses) up to 3 mm in diameter in the

adductor muscle.



 Overview of diagnostic methods, including sensitivity and specificity:

Histology is the diagnostic method currently used. This is probably of moderate

specificity and low sensitivity.



 Disease management activities in major producing countries: None, but the

disease may be initiated by scallops overlapping their valves (biting) under

crowded conditions.



Reference



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994). Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4: 1-199.





4.0 MYCOTIC DISEASES



4.1 MYCOTIC DISEASES OF BIVALVES IN AUSTRALIA



The only mycoses reported from Australian bivalves were three isolates (Curvularia

sp., Exserohilum rostratum, unidentified sp.) from cultured juvenile clams (Tridacna

crocea) that were thought to be malnourished (Norton et al. 1994). Possible shell

disease has been seen in aged Pacific oysters on collapsed stakes in a shallow

enclosed bay in Tasmania (Munday, unpublished).







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

46





Reference



Norton, J. H., Thomas, A.D. & Barker, J.R. (1994). Fungal infection in the cultured

juvenile boring clam Tridacna crocea. J. Invertebr. Pathol. 64: 273-275.





4.2 SIGNIFICANT MYCOTIC DISEASES OF BIVALVES NOT KNOWN IN

AUSTRALIA



4.2.1 SHELL DISEASE



1. Name and taxonomy of disease agent: Shell disease, Dutch shell disease, Maladie

du Pied, Maladie de la charnière, caused by Ostracoblabe implexa (Phycomycetales).



2. Agent stability and inactivation data: Restricted to waters where temperatures

exceed 22°C for more than two weeks.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: U.K., Ireland, Portugal.



 Host range: Edible oysters (Ostrea edulis, Crassostrea gigas, Crassostrea

angulata, Saccostrea cuccullata), pearl oysters (Pinctada margaritifera) and

probably many other bivalve species.



 Morbidity/mortality rates: Although primarily a shell disease, Ostracoblabe

implexa may kill the host if infesting the shell where the adductor muscle attaches,

causing it to detach and the shell to gape, providing access to the host by other

organisms.



 Transmission: Horizontal, direct.



4. Host impact:



 Tissue tropism: The shell.



 Brief description of major pathological and biological effects: Invasion occurs

at the outer surface of the shell. The fungus penetrates the shell, initially causing

raised white spots with a transparent centre on the inner surface. These coalesce

forming discoloured conchiolin “warts” and thickening of the shell margin.



5. Diagnostics and disease control



 Key diagnostic features: The appearance of the shell (white raised spots,

conchiolin “warts”, abnormal shell growth.



 Overview of diagnostic methods, including sensitivity and specificity: In

squash preparations of shell decalcified in a 5% solution of disodium EDTA, a

dense mycelial network of straight hyphae, 1.5-2.5 m in diameter, with 4-6 m







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

47





dilations at 40-100 m intervals , may be seen. Septa only occur in dying mycelia.

Histologically metaplasia may be apparent in the mantle epithelium. The fungus

may be cultured on shell fragments in sterile seawater containing antibiotics.

Diagnostic methods are likely to have low sensitivity and specificity.



 Disease management activities in major producing countries: In the wild O.

implexa breaks down shells in a natural process of degradation (Mao Che et al.

1996). However, shells should not be moved from areas where O. implexa is

present to areas where the fungus is absent and temperatures of >22°C occur for

more than 2 weeks. Removal of old oysters and accumulations of dead shell

should assist with control.



Reference



Mao Che, L., Le Campion-Alsumard, T., Boury-Esnault, N., Payri, C., Gobulic, S. &

Bezac, C. (1996). Biodegradation of shells of the black pearl oyster, Pinctada

margaritifera var. cumingii, by microborers and sponges in French Polynesia. Mar.

Biol. 126: 509-519.





4.2.2 LARVAL MYCOSIS



1. Name and taxonomy of disease agent: Larval mycosis caused by Sirolpidium

zoophthorum (Sirolpidiaceae, Phycomycetales).



2. Agent stability and inactivation data: Grows well in culture at 20-30°C.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: In hatcheries and natural stocks

along the eastern coast of the U.S.



 Host range: Oysters (Crassostrea virginica), scallops (Argopecten irradians) and

clams (Mercenaria mercenaria, Ruditapes decussatus) (Davis et al. 1954, Martin

et al. 1997, Martin 1998).



 Morbidity/mortality rates: Over 90% of larvae may be killed within 2-4 days of

the infection appearing. Higher mortalities may occur, but there are usually some

uninfected survivors.



 Transmission: Horizontal, direct.



4. Host impact:



 Tissue tropism: Systemic in soft tissues.



 Brief description of major pathological and biological effects: Affects larvae

ranging from early veligers to postmetamorphic juveniles 400 m in diameter.

The fungus spreads through the soft tissues causing them to disintegrate.

Sporangia produce tubes which protrude outside of the shell, releasing zoospores.





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

48







5. Diagnostics and disease control



 Key diagnostic features: Highly aerobic fungus that grows well on nutrient agar

at 20-30°C, with branched septate thallus 10-15 m in diameter when young, and

up to 82 m in diameter when mature. In thalli >40 um diameter, large spherical

vacuoles become visible. Sporangia have a discharge tube 15-142 m long and 5

m thick, formed from swollen terminal cells. Planonts are 2 x 5 m, biflagellate,

heterokont and monoplanetic. Occasionally produced resistant spores are ovoid,

light golden brown, thick walled and ~ 45-90 x 40-80 m.



 Overview of diagnostic methods, including sensitivity and specificity: Culture

on nutrient agar, and appearance in wet mounts, are used for diagnosis. Specificity

is high and sensitivity probably moderate.



 Disease management activities in major producing countries: Destruction of

infected stocks and disinfection of the hatchery.



References



Davis, H.C., Loosanoff, V.L., Weston, W.H. & Martin, C. (1954). A fungus disease in

clam and oyster larvae. Science 120: 36-38.



Martin, C. (1998). Fungal diseases in aquaculture: recent observations in larval

bivalves. J. Shellfish Res. 17: 359-360.



Martin, C., Stiles, S., Choromanski, J., Widman, J.C., Schweitzer, D. & Cooper, C.

(1997). Sirolpidium zoophthorum, lethal fungus parasite of bivalve larvae: recent

observations in bay scallop cultures. J. Shellfish Res. 16: 291.





4.2.3 CHYTRID-LIKE DISEASE (QUAHUAG PARASITE X)



1. Name and taxonomy of disease agent: Chytrid-like disease, quahuag parasite X.

This agent showns similarities to the Thraustochytriales and the Labyrinthulales

(Whyte et al 1994).



2. Agent stability and inactivation data: Not reported.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Wild and hatchery stocks, Gulf

of St Lawrence, northeastern United States.



 Host range: Quahuag (Mercenaria mercenaria)



 Morbidity/mortality rates: Causes 80-90% mortalities in juvenile hatchery

stocks (Bower et al. 1994), and up to 100% mortalities in certain stocks (Whyte et

al. 1994).







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

49







 Transmission: Direct by a uninucleate biflagellate stage (Whyte et al. 1994).



4. Host impact:



 Tissue tropism: Throughout the connective tissue.



 Brief description of major pathological and biological effects: Infected animals

show massive haemocyte infiltration, and connective tissue necrosis in the

digestive gland and foot. Spherical stages, usually multinucleate, are found

throughout the connective tissue. They are usually surrounded by a clear halo-like

area that is thought to indicate production of an exotoxin. Haemocytes infiltrate

the tissues surrounding the halo, but do not occur within it. When several stages

occur close together, the haloes coalesce. When a halo is not present, the organism

is surrounded by haemocytes (Whyte et al. 1994).



5. Diagnostics and disease control



 Key diagnostic features: Connective tissue congestion and necrosis in the

presence of spherical organisms, often surrounded by a clear space or halo.



 Overview of diagnostic methods, including sensitivity and specificity:

Histology and electron microscopy are used to visualize the organism and

associated pathology. It can be cultured in sterile artificial seawater (SAS)

containing antibiotics. In SAS vegetative stages, ranging from 8-71 m in

diameter, increase in size over 5 days at 26°C. During this time the stages undergo

karyokinesis to produce multinucleate stages. Cytokinesis occurs, giving rise to

morula-like clusters of daughter cells 28 m in diameter inside the cell wall of the

original stage. This cell wall bursts, releasing motile uninucleate biflagellate

stages. Culture on potato dextrose agar (PDA) results in growth of yellowish

white colonies 1-3 mm in diameter after 3-8 days at 14°C. Some of the stages

produce hyphal nets. Biflagellate stages occur after 8 days in PDA. The parasite

does not grow in Fluid Thioglycollate Medium, or stain with Lugol’s iodine. It

also does not grow in Minimal Essential Medium at 26°C (Whyte et al. 1994).

Specificity of diagnostic methods is probably high but sensitivity is probably

moderate.



 Disease management activities in major producing countries: Thinning out

clam stocks reduces this disease to a negligible level. Control of movement from

infected areas.



References



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994). Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4: 1-199.



Whyte, S.K., Cawthorn, R.J. & McGladdery, S.E. (1994). QPX (Quahuag Parasite X), a

pathogen of northern quahuag Mercenaria mercenaria from the Gulf of St Lawrence,

Canada. Dis. Aquat. Org. 19: 129-136.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

50









5.0 PROTOZOAN DISEASES



5.1 PROTOZOAN DISEASES OF BIVALVES IN AUSTRALIA



There are 5 listed protozoan diseases of bivalves notifiable to the O.I.E. These are

marteiliosis (caused by Marteilia refringens and Marteilia sydneyi), bonamiosis

(caused by Bonamia ostreae and Bonamia sp.), mikrocytosis (caused by Mikrocytos

mackini and Mikrocytos roughleyi), haplosporidiosis (caused by Haplosporidium

nelsoni and Haplosporidium costale), and perkinsosis (caused by Perkinsus marinus

and Perkinsus olseni).



Marteilia sydneyi infects Sydney rock oysters (Saccostrea commercialis) from the

Great Sandy Strait, southern Queensland to McLay River, N.S.W., Georges River,

N.S.W. and in King Bay, near Dampier, Western Australia. It is probably widely

distributed around mainland Australia (Wolf 1979, Roubal et al. 1989). Although

Marteilia lengehi is considered by some to be difficult to distinguish from M.

refringens (Bower et al. 1994), and M. lengehi occurs in Saccostrea cuccullata in

north Western Australia (Hine: unpublished information), M. lengehi is considered to

be a different species from M. refringens in this review. A similar organism,

Marteilioides chungmuensis, infects the ovacytes of rock oysters (Saccostrea

echinata) in Darwin Harbour. M. chungmuensis is currently causing large scale

spawning failures in Pacific oysters (C. gigas) in Korea and Japan. Similarly, a

microsporidian pathogen, Steinhausia mytilovum, infects the ovacytes of blue mussels

(Mytilus galloprovincialis) in Western Australia. It is currently causing large scale

spawning failures in blue mussels (Mytilus edulis) in Korea and Japan.



Bonamia sp. infects flat oysters (Ostrea angasi), in Port Phillip Bay, Victoria, around

Tasmania, and near Albany, Western Australia. Mikrocytos roughleyi infects Sydney

rock oysters (Farley et al. 1988) in the Georges River, N.S.W. and occurs in

hatcheries at Albany and Carnarvon, Western Australia. Molecular and ultrastructural

evidence suggests that M. roughleyi is more closely related to Bonamia than to

Mikrocytos mackini. Neither of the Haplosporidium spp. causing haplosporidiosis has

been reported from Australia. Neither a Haplosporidium sp. infecting pearl oysters

(Pinctada maxima) (Hine & Thorne 1998), nor another species infecting rock oysters

(Saccostrea cuccullata) in north Western Australia (Hine:unpublished data) are

labelled by a probe specific for H. nelsoni (see below).



Perkinsus olseni was originally reported in abalone (Haliotis spp.) from South

Australia (Lester & Davis 1981, O’Donoghue et al. 1991). Perkinsus sp. has since

been reported to be widespread in bivalves from the Great Barrier Reef (GBR)

(Goggin & Lester 1987), and appears to be non-host specific (Goggin et al. 1989).

Perkinsus sp. also occurs in many bivalve species in northern Western Australia

(Hine: unpublished data). Molecular studies suggest that the GBR Perkinsus are P.

olseni (Goggin & Lester 1995), and that Perkinsus atlanticus from Ruditapes

philippinarum and Ruditapes decussatus around Spain and Portugal, are closely

related to P. olseni (Robledo et al. 1997). One interpretation of the close relationship

of P. olseni and P. atlanticus, is that P. olseni was moved from southeast Asia to

Europe with movement of R. philippinarum. This is supported by the recent report of





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

51





Perkinsus in R. philippinarum in Japan, resembling P. olseni and P. atlanticus

(Hamaguchi et al. 1998). Therefore, for this review, P . atlanticus is regarded as a

subspecies of P. olseni.



Ciliates have commonly been found in Australian oysters (Wilson, 1993) and it is

probable that they are ubiquitous commensals. Gregarines and hexamitids are also

probably ubiquitous (Bower et al, 1994).



References



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994). Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4: 1-199.



Farley, C.A., Wolf, P.H. & Elston, R.A. (1988). A long-term study of "microcell"

disease in oysters with a description of new genus Mikrocytos (g.n.), and two new

species, Mikrocytos mackini (sp.n.) and Mikrocytos roughleyi (sp. n.). Fish. Bull. 86:

581-593.



Goggin, C.L. & Lester, R.J.G. (1987). Occurrence of Perkinsus species (Protozoa,

Apicomplexa) in bivalves from the Great Barrier reef. Dis. Aquat. Org. 3: 113-117.



Goggin, C.L. & Lester, R.J.G. (1995). Perkinsus, a protistan parasite of abalone in

Australia: a review. Mar. Freshwat. Res. 46: 639-646.



Goggin, C.L., Sewell, K.B. & Lester, R.J.G. (1989). Cross-infection experiments with

Australian Perkinsus species. Dis. Aquat. Org. 7: 55-59.



Hamaguchi, M., Suzuki, N., Usuki, H. & Ishioka, H. (1998). Perkinsus protozoan

infection in short-necked clam Tapes (=Ruditapes) philippinarum in Japan. Fish Pathol.

33: 473-480.



Hine, P.M. & Thorne, T. (1998). Haplosporidium sp. (Haplosporidia) in hatchery-reared

pearl oysters, Pinctada maxima (Jameson, 1901), in north Western Australia. J.

Invertebr. Pathol 71: 48-52..



Lester, R.J.G. & Davis, G.H.G. (1981). A new Perkinsus species (Apicomplexa,

Perkinsea) from the abalone Haliotis ruber. J. Invertebr. Pathol. 37: 181-187.



O'Donoghue, P.J., Phillips, P.H. & Shepherd, S.A. (1991). Perkinsus (Protozoa:

Apicomplexa) infections in abalone from South Australian waters. Trans. R. Soc. S.

Aust. 115: 77-82.



Robledo, J.A.F., Wright, A.C., Coss, C.A., Vasta, G.R. & Goggin, C.L. (1997). Further

studies of conserved genes from Perkinsus isolates. J. Shellfish Res. 16: 342.



Roubal, F.R., Masel, J. & Lester, R.J.G. (1989). Studies on Marteilia sydneyi, agent of

QX disease in the Sydney rock oyster, Saccostrea commercialis, with implications for its

life cycle. Aust. J. Mar. Freshwater Res. 40: 155-167.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

52





Wilson, J. (1993). The health status of two species of Tasmanian farmed shellfish,

Crassostreae gigas (Thunberg, 1793) and Ostrea angasi (Sowerby, 1871). Masters

Thesis, University of Tasmania, 189 pp.



Wolf, P.H. (1979). Life cycle and ecology of Marteilia sydneyi in the Australian oyster,

Crassostrea commercialis. Mar. Fish. Rev. 41: 70-72.







5.2 SIGNIFICANT PROTOZOAN INFECTIONS OF BIVALVES NOT

KNOWN IN AUSTRALIA



5.2.1 ABER DISEASE (MARTEILIASIS)



1. Name and taxonomy of disease agent: Aber disease, Marteiliasis – Marteilia

refringens (Paramyxea)



2. Agent stability and inactivation data: High salinities (35-37‰) limit

development.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Atlantic Europe, from southern

England to Portugal.



 Host range: Flat oysters (Ostrea edulis, Ostrea angasi, Tiostrea chilensis),

Pacific oysters (Crassostrea gigas), blue mussels (Mytilus edulis, Mytilus

galloprovincialis), and cockles (Cerastoderma edulis). Possibly also Crassostrea

virginica (see note below)



 Morbidity/mortality rates: Very variable, prevalence often very high (100%).

Mortality appears to occur when the parasite sporulates, which in Spain occurs

throughout the year.



 Transmission: Unknown, but direct horizontal transmission does not occur

(Figueras & Montes 1988, Berthe et al. 1998). Similarly, Marteilia sydneyi

appears to require an intermediate host (Roubal et al. 1989). A benthic deposit-

feeding invertebrate, such as an annelid, is thought to be involved. Transmission

occurs in summer (July-August in Europe).



4. Host impact:



 Tissue tropism: The epithelial cells of the stomach, digestive ducts and digestive

tubules.



 Brief description of major pathological and biological effects: Parasitism

inhibits digestion, starving the host. There is glycogen loss, discolouration of the

digestive gland, cessation of growth and reproduction (Villalba et al. 1993a),

tissue necrosis and mortalities. The main impact is on the condition and meat

weight of infected oysters, with weights reduced by ~40% (Morel and Tigé 1974).





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

53





Infection begins in the stomach and spreads to the digestive ducts and tubules

(Villalba et al. 1993b).



5. Diagnostics and disease control



 Key diagnostic features: Discolouration and translucency of the digestive gland.

Characteristic basophilic stages in digestive duct epithelial cells, and acidophilic

stages in digestive tubules.



 Overview of diagnostic methods, including sensitivity and specificity: Except

in light infections, the parasite is relatively easy to detect by routine

histopathology.



 Disease management activities in major producing countries: Controls on

stock movements. However, once a culture site becomes contaminated, the

parasite may persist for long periods in the intermediate host, making cleaning of

farm sites impossible.



Note: American oysters (Crassostrea virginica) became infected with a Marteilia sp.

resembling M. refringens and Marteilia maurini, while being held for assessment of

aquaculture potential in France (Renault et al 1995)



References



Berthe, F.C.J., Pernas, M., Zerabib, M., Haffner, P., Thébault, A. & Figueras, A.J.

(1998). Experimental transmission of Marteilia refringens with special consideration of

the life cycle. Dis. Aquat. Org. 34: 135-144.



Figueras, A.J. & Montes, J. (1988). Aber disease of edible oysters caused by Marteilia

refringens. Am. Fish. Soc. Spec. Publ. 18: 38-46.



Morel, M. & Tigé, G. (1974). Maladie de la glande digestive de l’Huître plate.

Science et Pêche. Bull. Inst. Pêches marit. 241: 33-36.



Renault, T., Cochennec, N. & Chollet, B. (1995). Marteiliosis in American oysters

Crassostrea virginica reared in France. Dis. Aquat. Org. 23:161-164.



Roubal, F.R., Masel, J. & Lester, R.J.G. (1989). Studies on Marteilia sydneyi, agent of

QX disease in the Sydney rock oyster, Saccostrea commercialis, with implications for its

life cycle. Aust. J. Mar. Freshwater Res. 40: 155-167.



Villalba, A., Mourelle, S.G., Carballal, M.J. & Lopez, M.C. (1993a). Effects of infection

by the protistan parasite Marteilia refringens on the reproduction of cultured mussels

Mytilus galloprovincialis in Galicia (NW Spain). Dis. Aquat. Org. 17: 205-213.



Villalba, A., Mourelle, S.G., Lopez, M.C., Carballal, M.J. & Azevedo, C. (1993b).

Marteiliasis affecting cultured mussels Mytilus galloprovincialis of Galicia (NW Spain).

I. Etiology, phases of infection, and temporal and spatial variability in prevalence. Dis.

Aquat. Org. 16: 61-72.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

54







5.2.2 MARTEILIOSIS OF CALICO SCALLOPS



1. Name and taxonomy of disease agent: Marteiliosis of calico scallops (Argopecten

gibbus) caused by Marteilia sp. (Paramyxea).



2. Agent stability and inactivation data: Unknown.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Only reported from the coast of

Florida.



 Host range: The calico scallop (Argopecten gibbus)



 Morbidity/mortality rates: : 100%. The fishery, which produced 11-40 million

pounds of adductor muscle meat annually before December 1988 was destroyed

within 1 month over 2,500 square miles of sea floor and, as of spring 1992, had not

recovered to harvestable quantities (Moyer et al. 1993). Not reported since the

original epizootic.



 Transmission: Presumed to be indirect and involve another host, as in other

Marteilia spp.



4. Host impact:



 Tissue tropism: Epithelium of the digestive tubules.



 Brief description of major pathological and biological effects: Infection of the

digestive tubules, inhibits digestion leading to starvation and death.



5. Diagnostics and disease control



 Key diagnostic features: Characteristic stages in the digestive epithelial cells.



 Overview of diagnostic methods, including sensitivity and specificity: Readily

detected by histopathology.



 Disease management activities in major producing countries: None.



Reference



Moyer, M.A., Blake, N.J. & Arnold, W.S. (1993). An ascetosporan disease causing mass

mortality in the Atlantic calico scallop Argopecten gibbus (Linnaeus, 1758). J. Shellfish

Res. 12: 305-310.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

55





5.2.3 MARTEILIA SP. (PARAMYXEA)



1. Name and taxonomy of disease agent: Marteilia sp. (Paramyxea).



2. Agent stability and inactivation data: Not reported.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Fiji



 Host range: Giant clams (Tridacna maxima)



 Morbidity/mortality rates: Not reported.



 Transmission: Probably horizontal via an alternative host.



4. Host impact:



 Tissue tropism: Ciliated columnar epithelium of the kidney.



 Brief description of major pathological and biological effects: Lesions appear

as chalk-white foci in the dark red-brown kidney. Histologically, kidney lesions

appear as numerous cyst-like structures lined with ciliated columnar epithelium.

Within the cysts are groups of protistan cells, ~2 m in diameter, with punctate

nuclei. They are enclosed within a larger cellular unit, ~4 m in diameter, that

stains grey with H & E, but is more eosinophilic further from the epithelium. They

contain irregularly-shaped refringent bodies (Norton et al. 1993).



5. Diagnostics and disease control



 Key diagnostic features: Macroscopic white cysts in the epithelium of the dark

red-brown kidney, containing organisms with a cell-within-cell configuration.



 Overview of diagnostic methods, including sensitivity and specificity:

Histology is the only approach used to date.



 Disease management activities in major producing countries: No known

methods of prevention or control.



Reference



Norton, J.H., Perkins, F.O. & Ledua, E. (1993). Marteilia-like infection in a giant

clam, Tridacna maxima in Fiji. J. Invertebr. Pathol. 61: 328-330.





5.2.4 BONAMIOSIS



1. Name and taxonomy of disease agent: Bonamiosis, caused by Bonamia ostreae

(Alveolata, Eukaryota). Although some authors suggest that the Bonamia sp. of







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

56





Australia and New Zealand may be conspecific with B. ostreae, sequencing of the

18S region of rDNA shows these are two separate species (Dr Rob Adlard,

Queensland Museum: pers. comm.). These sequences also suggest that, although B.

ostreae ultrastructurally resembles a haplosporidian, it is more closely related to the

dinoflagellates (Carnegie et al. 1997).



2. Agent stability and inactivation data: The antimicrobial peptide magainin 1 is

effective against B. ostreae in vitro (Morvan et al. 1994).



3. Epidemiological features of the disease:



 Geographic range and features of distribution: The Netherlands, southern

England, Atlantic and Mediterranean France, Spain, Portugal, Italy, Maine and

California, U.S.A., Argentina, and possibly Chile.



 Host range: The published literature suggests that the genus Ostrea (including

Tiostrea) is susceptible to infection, and a recent paper reports a Bonamia-like

parasite from Crassostrea rivularis introduced into France from the U.S.A.

(Cochennec et al. 1998).



 Morbidity/mortality rates: Very high for at least 6 years after uninfected beds are

exposed to infected oysters (Bower et al. 1994).



 Transmission: Direct horizontal transmission of vegetative stages, spores not

reported.



4. Host impact:



 Tissue tropism: B. ostreae is phagocytosed by haemocytes in which it grows and

divides, until the haemocyte lyses. Also infects gill epithelial cells (Montes et al.

1994).



 Brief description of major pathological and biological effects: The infective

dose is about 200 particles. Once inside the oyster the parasite is recognised as

foreign and is phagocytosed by haemocytes. Once inside the haemocytes it grows

and divides until the haemocyte lyses. The released parasites are then

phagocytosed, and go through the same process. In order to contain the infection,

increasing numbers of haemocytes are produced, at the expense of gametogenesis,

which ceases. Finally the infection overwhelms the oyster, which probably dies of

exhaustion and tissue damage due to the release of hydrolytic enzymes from lysed

haemocytes and the cells of surrounding tissues.



5. Diagnostics and disease control



 Key diagnostic features: A few oysters may have yellow discolouration, and/or

perforated ulcers on the gill and mantle. However, these signs can not be regarded as

pathognomonic. Histopathology will detect light to heavy infections, and

examination of heart imprints can be used to detect moderate to heavy infections.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

57





 Overview of diagnostic methods, including sensitivity and specificity:

Monoclonal antibody techniques have been developed (Boulo et al. 1989, Rogier et

al. 1991, Cochennec et al. 1992). PCR has more recently been used to amplify B.

ostreae DNA, from which a DNA probe has been developed by IFREMER. This

probe labels B. ostreae and Bonamia sp. from New Zealand, and weakly labels M.

roughleyi.



 Disease management activities in major producing countries: None except

controls on movement. Attempts are underway to insert genes for magainins

(antibiotic peptides) from toads (Xenopus) into host oysters (Ostrea edulis).



References



Boulo, V., Mialhe, E., Rogier, H., Paolucci, F & Grizel, H. (1989). Immunodiagnosis of

Bonamia ostreae (Ascetospora) infection of Ostrea edulis L. and subcellular

identification of epitopes by monoclonal antibodies. J. Fish Dis. 12: 257-262.



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994). Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4: 1-199.



Carnegie, R.B., Distel, D.L. & Barber, B.J. (1997). Amplification and sequencing of the

Bonamia ostreae 18S rDNA gene: phylogenetic considerations and applications. J.

Shellfish Res. 16: 328.



Cochennec, N., Hervio, D., Panatier, B., Boulo, V., Mialhe, E., Rogier, H., Grizel, H. &

Paolucci, F. (1992). A direct monoclonal antibody sandwich immunoassay for detection

of Bonamia ostreae (Ascetospora) in hemolymph samples of the flat oyster Ostrea

edulis (Mollusca: Bivalvia). Dis. Aquat. Org. 12: 129-134.



Cochennec, N., Renault, T., Boudry, P., Chollet, B & Gerard, A. (1998). Bonamia-like

parasite found in the Suminoe oyster Crassostrea rivularis reared in France. Dis. Aquat.

Org. 34: 193-197.



Montes, J., Anadon, R. & Azevedo, C. (1994). A possible life cycle for Bonamia ostreae

on the basis of electron microscopy. J. Invertebr. Pathol. 63: 1-6.



Morvan, A., Bachère, E., Pinto Da Silva, P., Pimenta, P. & Mialhe, E. (1994). In vitro

activity of the antimicrobial peptide magainin 1 against Bonamia ostreae, the

intrahemocytic parasite of the flat oyster Ostrea edulis. Mol. Mar. Biol. Biotechnol. 3:

327-333.



Rogier, H., Hervio, D., Boulo, V., Clavies, C., Hervaud, E., Bachère, E., Mialhe, E.,

Grizel, H., Pau, B. & Paolucci, F. (1991). Monoclonal antibodies against Bonamia

ostreae (Protozoa: Ascetospora), an intrahaemocytic parasite of flat oyster Ostrea edulis

(Mollusca: Bivalvia). Dis. Aquat. Org. 11: 135-142.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

58





5.2.5 DENMAN ISLAND DISEASE



1. Name and taxonomy of disease agent: Denman Island disease, caused by

Mikrocytos mackini. Although currently congeneric with Mikrocytos roughleyi, these

two organisms are not closely related at the molecular level or ultrastructurally. In

particular, M. roughleyi has a mitochondrion, but M. mackini does not. This suggests

that M. mackini is a more primitive protist.



2. Agent stability and inactivation data: Limited by temperature, with water

temperature requirement of 12C, but subclinical infections may persist for 3 months at

15C (Bower: pers. comm.).



3. Epidemiological features of the disease:



 Geographic range and features of distribution: The coast of British Columbia,

Canada.



 Host range: Natural infections: Pacific oysters (Crassostrea gigas). Experimental

infections: Ostrea edulis, Ostreola conchaphila, Crassostrea virginica (Hervio et al.

1993, 1995a, Bower et al. 1994).



 Morbidity/mortality rates: About 30% in some years, on some beaches (Hervio

et al. 1995a). Mortalities occur predominantly in April and May after a 3-4 month

period when temperatures are less than 10°C (Bower et al. 1994). Severe

infections are restricted to older oysters (>2 years old).



 Transmission: Directly transmitted between oysters during North American

spring (March – June), but the disease does not occur until the following spring

(Hervio et al. 1995a).



4. Host impact:



 Tissue tropism: Vesicular connective tissue cells, but may also be observed in

infiltrating haemocytes and muscle cells (Bower et al. 1994).



 Brief description of major pathological and biological effects: Focal

intracellular infection of vesicular connective tissue cells causes haemocyte

infiltration and tissue necrosis. The lesions are macroscopic and resemble

abcesses.



5. Diagnostics and disease control



 Key diagnostic features: Green pustules (5 mm diameter) within the body wall

or on the surfaces of the mantle, palps, often with a brown scar on the shell

adjacent to the lesion on the mantle surface. The adductor muscle may also be

infected (Hervio et al. 1996).



 Overview of diagnostic methods, including sensitivity and specificity: The

parasite is small (2-3 m in diameter) and very difficult to detect histologically,







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

59





even in heavy infections. The parasite is more readily visualised in tissue imprints

examined at x 1,000 magnification (Hervio et al. 1996). Specific molecular probes

are being developed (Hervio et al. 1995b).



 Disease management activities in major producing countries: Controls on

movements. The effect of the disease on infected populations can be reduced to a

manageable level by harvesting and moving large oysters to higher tide levels (1-2

m) prior to March and not planting oysters at lower tide levels before June (Bower

et al. 1994).



References



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994). Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4: 1-199.



Hervio, D., Bower, S.M. & Meyer, G.R. (1993). Detection, isolation, and host specificity

of Mikrocytos mackini, the cause of Denman Island disease in Pacific oysters

Crassostrea gigas. J. Shellfish Res. 12: 136.



Hervio, D., Bower, S.M. & Meyer, G.R. (1995a). Life cycle, distribution and lack of

host specificity of Mikrocytos mackini, the cause of Denman Island disease in Pacific

oysters, Crassostrea gigas. J. Shellfish Res. 14: 228.



Hervio, D., Bower, S.M. & Meyer, G.R. (1996). Detection, isolation, and experimental

transmission of Mikrocytos mackini, a microcell parasite of Pacific oysters Crassostrea

gigas (Thunberg). J. Invertebr. Pathol. 67: 72-79.



Hervio, D., Meyer, G.R., Bower, S,M & Adlard, R.D. (1995b). Development of specific

molecular probes for serological and PCR assays for the identification and diagnosis of

Mikrocytos mackini, the cause of Denman Island disease in the Pacific oyster,

Crassostrea gigas. J. Shellfish Res. 14: 268.





5.2.6 HAPLOSPORIDIOSIS (MSX)



1. Name and taxonomy of disease agent: MSX, haplosporidiosis, caused by

Haplosporidium nelsoni (Haplosporidiidae, Haplosporida, Alveolata, Eukaryota).



2. Agent stability and inactivation data: Holding in vivo for up to 2 weeks in 10 ‰

salinity seawater at 20°C kills the parasite but not the host (Ford 1985). H. nelsoni

does not cause disease at below 15 ‰ salinity (Bower et al. 1994).



3. Epidemiological features of the disease:



 Geographic range and features of distribution: East coast of the U.S.A. from

Maine to Florida. A Haplosporidium sp., similar in size and pathology, occurs in

Pacific oysters (Crassostrea gigas) in California and in Matsushima Bay, Japan,

from which the Californian stocks were derived (Friedman et al. 1991, Friedman

1996). A sensitive and specific probe for H. nelsoni (Stokes & Burreson 1995)

labels the Californian and Japanese Haplosporidium, suggesting that it is also H.





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

60





nelsoni, and that H. nelsoni was originally introduced into California in Japanese

Pacific oysters. The sudden appearance of H. nelsoni among Crassostrea virginica

in Delaware Bay in 1957 may have been due to introduction with movement of

Pacific oysters from California onto the east coast. Another Haplosporidium sp.,

similar in size and pathology, occurs in Pacific oysters (Crassostrea gigas) in

France. It is also labelled by the probe, and is also considered to be H. nelsoni.



 Host range: Crassostrea virginica (U.S.A.), Crassostrea gigas (Japan, U.S.A.,

France).



 Morbidity/mortality rates: 90-95% (Haskin and Andrews 1988). Survivors are

seriously affected (Ford and Figueras 1988).



 Transmission: Unknown. H. nelsoni cannot be transmitted directly oyster to oyster

using plasmodia, either by injection or co-habitation, but there have been fewer

attempts to effect transmission using spores (Haskin and Andrews 1988, Ford 1992).

The evidence to date suggests involvement by an unknown intermediate host.

Surveys of likely intermediate hosts are being carried out using PCR technology

(Stokes et al. 1997).



4. Host impact:



 Tissue tropism: Gill epithelium, systemic in connective tissues, digestive

epithelium.



 Brief description of major pathological and biological effects: Initial invasion

occurs in the gill epithelium. The parasite then penetrates the basement membrane

and becomes systemic in the connective tissue. Finally it invades the epithelium of

the digestive ducts and tubules, where it sporulates within the epithelial cells. It is

extracellular in the connective tissue, and evades or repels haemocytes.

Gametogenesis is inhibited when parasite levels are high (Ford & Figueras 1988).

Mortality is often associated with sporulation, which is most often observed in

juvenile oysters.



5. Diagnostics and disease control



 Key diagnostic features: The genus Haplosporidium has characteristic spores. In

all species, except H. nelsoni, sporulation occurs in the connective tissue, below

the basement membrane of the digestive ducts and tubules. Only H. nelsoni

sporulates within the epithelium of the digestive tract, and not in the connective

tissues.



 Overview of diagnostic methods, including sensitivity and specificity:

Histology is more reliable than haemolymph analysis (Burreson et al. 1988), but is

only reliable in moderate to heavy infections, and is less reliable than PCR in light

infections (Stokes et al. 1996). Ziehl-Neelsen carbol fuchsin stain enhances

detection of the parasite. A DNA probe specific to H. nelsoni has been developed.

This probe does not label other parasites and pathogens including Haplosporidium

spp. (Haplosporidium louisiana, Haplosporidium costale, Haplosporidium

teredinis) (Stokes & Burreson 1995, Stokes et al. 1995) and the two known





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

61





Haplosporidia from Australian bivalves. The PCR primers were able to detect the

H. nelsoni SSU rDNA from 50 ng of infected oyster genomic DNA or from 10 fg

of cloned H. nelsoni SSU rDNA. In dot blot hybridizations the probe detected 100

pg of cloned H. nelsoni rDNA and the presence of 1 g of genomic DNA from an

infected oyster.



 Disease management activities in major producing countries: Controls on

movements of all aquatic organisms from infected areas (some may be infected

intermediate hosts) (Ford 1992). Oysters are grown in cold low salinity waters for

as long as possible, and warm high salinity environments are avoided. Some

oyster stocks are more resistant to infection than others, and are selected for

culture.



References



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994). Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4: 1-199.



Burreson, E.M., Robinson, M.E. & Villalba, A. (1988). A comparison of paraffin

histology and hemolymph analysis for the diagnosis of Haplosporidium nelsoni (MSX)

in Crassostrea virginica (Gmelin). J. Shellfish Res. 7: 19-23.



Ford, S.E. (1985). Effects of salinity on survival of the MSX parasite Haplosporidium

nelsoni (Haskin, Stauber, and Mackin) in oysters. J. Shellfish Res. 5: 85-90



Ford, S.E. (1992). Avoiding the transmission of disease in commercial culture of

molluscs, with special reference to Perkinsus marinus (Dermo) and Haplosporidium

nelsoni (MSX). J. Shellfish Res. 11: 539-546.



Ford, S.E. & Figueras, A.J. (1988). Effects of sublethal infection by the parasite

Haplosporidium nelsoni (MSX) on gametogenesis, spawning, and sex ratios of oysters

in Delaware Bay. Dis Aquat. Org. 4: 121-133



Friedman, C.S. (1996). Haplosporidian infections of the Pacific oyster, Crassostrea

gigas (Thunberg), in California and Japan. J. Shellfish Res. 15: 597-600.



Friedman, C.S., Cloney, D.F., Manzer, D., Hedrick, R.P. (1991). Haplosporidiosis of the

Pacific oyster, Crassostrea gigas. J. Invertebr. Pathol. 58: 367-372.



Haskin, H.H. & Andrews, J.D. (1988). Uncertainties and speculations about the life

cycle of the eastern oyster pathogen Haplosporidium nelsoni (MSX). Am. Fish. Soc.

Spec. Publ. 18: 5-22.



Stokes, N.A. & Burreson, E.M. (1995). A sensitive and specific DNA probe for the

oyster pathogen Haplosporidium nelsoni. J. Eukaryot. Microbiol. 42: 350-357.



Stokes, N.A., Flores, B.S., Burreson, E.M., Alcox, K.A., Guo Ximing & Ford, S.E.

(1997). Life cycle studies of Haplosporidium nelsoni (MSX) using PCR technology. J.

Shellfish Res. 16: 336.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

62





Stokes, N.A., Walker, J.G. & Burreson, EM. (1996). Comparison of Haplosporidium

nelsoni diagnostic techniques: Polymerase chain reaction outperforms histology. J.

Shellfish Res. 15: 498.





5.2.7 SEASIDE ORGANISM (SSO)



1. Name and taxonomy of disease agent: Seaside organism (SSO), haplosporidiosis,

caused by Haplosporidium costale (Haplosporidiidae, Haplosporida, Alveolata,

Eukaryota).



2. Agent stability and inactivation data: Restricted to high salinity (>25 ‰) waters.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Virginia and Maryland north to

Delaware Bay, and in Washington State, U.S.A., in high salinity waters.



 Host range: Crassostrea virginica. A similar organism has been reported from

Crassostrea gigas in France (Comps & Pichot 1991).



 Morbidity/mortality rates: Mortalities occur in May-June, but the size of these

mortalities cannot be accurately assessed, as two other pathogens, H. nelsoni and

Perkinsus marinus, co-exist with H. costale, and contribute to mortalities

(Andrews 1984a). In general, H. costale is considered less pathogenic than the

other two organisms. About 30-40% of oysters are infected, with mortalities

estimated at about 80-90% of prevalence, or about 25-35% of oysters (Andrews

1984b).



 Transmission: Unknown, but likely to be horizontal and indirect, involving an

unidentified intermediate host.



4. Host impact:



 Tissue tropism: Extracellular in connective tissue.



 Brief description of major pathological and biological effects: H. costale has a

well-defined life cycle with sporulation and mortality in May-June and infectivity

in June-July (Andrews 1984b). The parasite enters the oyster through the gills, and

proliferates in the connective tissue. Sporulation occurs throughout the connective

tissue, but not in the epithelia.



5. Diagnostics and disease control



 Key diagnostic features: In the areas of the eastern U.S.A. where H. nelsoni and

H. costale are enzootic, the pathogens can be distinguished by differences in spore

size, and site of sporulation. The mortality pattern also distinguishes the species,

except in May-June, as H. nelsoni sporulates and causes mortalities throughout the

year, whereas H. costale only sporulates and causes mortalities in May-June.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

63





 Overview of diagnostic methods, including sensitivity and specificity:

Histopathology is the main diagnostic tool, and the parasite is easily detected

using a Ziehl-Neelsen carbol fuchsin technique. Molecular tools are currently

being developed.



 Disease management activities in major producing countries: No known

control methods, but losses can be minimised by harvesting oysters at 18-24

months of age, and culturing them in low salinity (100 developing spores) bulges into the lumen of the digestive

gland tubule.



5. Diagnostics and disease control



 Key diagnostic features: Tumefactions and associated pathology, discoloured

digestive gland, dimensions of developmental stages and the one or two surface

filaments wrapped around the spore body.



 Overview of diagnostic methods, including sensitivity and specificity: In wet

mounts, the spores have one or two surface filaments wrapped around the spore

body.



 Disease management activities in major producing countries: None.



Note: A species of Minchinia has been recently reported from Mytilus

galloprovincialis in the Mediterranean (Comps & Tigé 1997). It was encountered

during routine monitoring of mussels under culture, when it was noticed that the

mussel was thin and whitish. Other than an electron microscopical description of the

parasite, no other information is available at present. A haplosporidan has also been

reported from blue mussels (Mytilus edulis) on the east coast of the U.S.A., but lack

of spores did not permit identification (Figueras et al. 1991). Another unidentified

haplosporidan lacking spores has been reported from bay scallops (Argopecten

irradians) in China (Chu et al. 1996).



References



Chu, F-L., Burreson, E.M., Zhang, F. & Chew, K.K. (1996). An unidentified

haplosporidian parasite of bay scallop Argopecten irradians cultured in Shandong and

Liaoning Provinces of China. Dis. Aquat. Org. 25: 155-158.



Comps, M. & Tigé, G. (1997). Fine structure of Minchinia sp., a haplosporidan

infecting the mussel Mytilus galloprovincialis L. Systematic Parasitol. 38: 45-50.



Figueras, A.J., Jardon, C.F. & Caldas, J.R. (1991). Diseases and parasites of mussels

(Mytilus edulis Linnaeus, 1758) from two sites on the east coast of the United States.

J. Shellfish Res. 10: 89-94.



Taylor, R.L. (1966). Haplosporidium tumefacientis sp. n., the etiologic agent of a

disease of the California sea mussel, Mytilus californianus Conrad. J. Invertebr.

Pathol. 8: 109-121.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

65





5.2.9 CARPET CLAM HAPLOSPORIDIOSIS



1. Name and taxonomy of disease agent: Carpet clam haplosporidiosis, caused by

Haplosporidium (=Minchinia) tapetis (Haplosporidiidae, Haplosporida, Alveolata,

Eukaryota).



2. Agent stability and inactivation data: None reported.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: France, Spain, Portugal.



 Host range: Carpet clams, Tapes (=Ruditapes) decussatus, and Manila clams

Tapes (=Ruditapes) philippinarum.



 Morbidity/mortality rates: Prevalence usually low (~4%), no mortalities

reported.



 Transmission: As for Haplosporidium spp. and other Minchinia, it is likely that

transmission is horizontal via an alternative host.



4. Host impact:



 Tissue tropism: The epithelia of the digestive tract, with sporulation occurring in

the interstitial connective tissue in the digestive gland and gills (Chagot et al.

1987, Figueras et al. 1992).



 Brief description of major pathological and biological effects: Histologically,

multinucleate plasmodia, 5-16 m in diameter, with 3-16 nuclei and prominent

nucleoli, occur in the epithelia of the digestive tract. The ovoid operculate spores

are 5-6 m long and 4-6 m wide.



5. Diagnostics and disease control



 Key diagnostic features: The dimensions of developmental stages are similar to

those of other Haplosporidium spp., but the presence of plasmodia in the digestive

epithelia of venerid clams (Tapes, Ruditapes, Venerupis) is only known to occur

in this Haplosporidium species.



 Overview of diagnostic methods, including sensitivity and specificity:

Histology is the only diagnostic approach at present.



 Disease management activities in major producing countries: None.



References



Chagot, D., Bachère, E., Ruano, F., Comps, M. & Grizel, H. (1987). Ultrastructural

study of sporulated instars of a haplosporidian parasitizing the clam Ruditapes

decussatus. Aquaculture 67: 262-263







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

66







Figueras, A., Robledo, J.A. & Novoa, B. (1992). Occurrence of haplosporidian and

Perkinsus-like infections in carpet-shell clams, Ruditapes decussatus (Linnaeus, 1758),

of the Ria de Vigo (Galicia, NW Spain). J. Shellfish Res. 11: 377-382.





5.2.10 EUROPEAN OYSTER MINCHINIASIS



1. Name and taxonomy of disease agent: European oyster minchiniasis, caused by

Minchinia armoricana (Haplosporidiidae, Haplosporida, Alveolata, Eukaryota).



2. Agent stability and inactivation data: Not reported.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: France and the Netherlands.



 Host range: Ostrea edulis, and Ostrea angasi imported into France for evaluation

as an aquaculture species (Bachère et al. 1987).



 Morbidity/mortality rates: Prevalence is low (1%), but the disease is fatal to

infected oysters.



 Transmission: Unknown, however Minchinia spp. are closely related to

Haplosporidium spp., and therefore it is likely that M. armoricana requires an

alternative host.



4. Host impact:



 Tissue tropism: Systemic in the connective tissue, in which M. armoricana

sporulates.



 Brief description of major pathological and biological effects: Infected oysters

are thin and watery in appearance, with a characteristic brownish colour due to the

mass of spores in the digestive gland. Histologically, eosinophilic plasmodia, (17-

25 m in diameter) spread throughout the connective tissue, develop to sporonts

(30-45 m) and sporocysts (35-50 m) containing 100-150 spores (5.0-5.5 x 4.0-

4.5 m) (Cahour et al. 1980).



5. Diagnostics and disease control



 Key diagnostic features: The watery appearance cannot be regarded as a key

diagnostic feature as oysters that have recently spawned, or which have other

haplosporidan infections, appear watery. The brownish colouration, dimensions of

developmental stages, and the two long (70-100 m) projections of the outer spore

wall, are the most reliable features.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

67





 Overview of diagnostic methods, including sensitivity and specificity: Wet

mounts are useful as the spores can be seen to have two long (70-100 m)

projections of the outer spore wall. Histology is also important in diagnosis.



 Disease management activities in major producing countries: None.



Note: Another Minchinia sp. has been reported from razor clams (Siliqua patula) on

the west coast of the U.S.A. (Moore & Elston 1991).



References



Bachère, E., Chagot, D., Tigé, G. & Grizel, H. (1987). Study of a haplosporidian

(Ascetospora), parasitizing the Australian flat oyster Ostrea angasi. Aquaculture 67:

266-268



Cahour, A., Poder, M. & Balouet, G. (1980). Présence de Minchinia armoricana

(Haplosporea, Haplosporida) chez Ostrea edulis d'origine francaise. C.R. Soc. Biol. 174:

359-368.



Moore, J.D. & Elston, R.A. (1991). Spore ultrastructure of a haplosporidan parasite of

razor clams, Siliqua patula. J. Shellfish Res. 10: 239.





5.2.11 PERKINSOSIS (DERMO)



1. Name and taxonomy of disease agent: Perkinsosis, “dermo”, caused by Perkinsus

marinus (Perkinsidae, Dinoflagellata, Alveolata, Eukaryota).



2. Agent stability and inactivation data: Low temperatures and low salinities inhibit

growth and development of disease. Both prevalence and intensity of infection

increase with increasing temperature and salinity (Burreson & Calvo 1996, Chu & La

Peyre 1993, Chu et al. 1993, 1996, Chu & Volety 1997). In vitro, P. marinus can be

controlled by a wide range of chemicals (Krantz 1994). Although lasolocid, malachite

green, cyclohexamide, monensin and sulphadimethoxine kill the parasite in vitro, only

cyclohexamide effectively kills the parasite in vivo. Infection persists, however, in

oysters treated with cyclohexamide at 10 mg/l after 30 days (Calvo & Burreson 1994).

The antibiotic bacitracin reduces levels of infection in vivo (Faisal et al. 1997). An

analogue of the peptide antibiotic polyphemusin caused complete inhibition of P.

marinus at 12 m g/ml and partial inhibition at 8 m g.ml (Pierce et al. 1997). In vitro

cultured parasites are killed by 300 ppm Cl2 within 0.5 h (Bushek et al. 1996).



3. Epidemiological features of the disease:



 Geographic range and features of distribution: The east coast of Northern and

Central America, from Maine to Venezuela.



 Host range: Epizootics occur in Crassostrea virginica, but P. marinus

experimentally infects clams (Mercenaria, mercenaria, Mya arenaria) and Pacific

oysters (Crassostrea gigas) (Barber & Mann 1994). C. virginica can become

infected via contact with snails (Boonea impressa), flat oysters (Ostrea lurida)





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

68





and a small fish (Gobiosoma bosci). The parasite occurs in a wide variety of

scavengers, and Perkinsus-like organisms occur in many bivalve groups (Ford

1992).



 Morbidity/mortality rates: These vary greatly, and are determined by the

genotypes of the pathogen and host (Bushek & Allen 1996), temperature and

salinity (Burreson & Calvo 1996). At high temperatures and salinities mortalities

are often >90% (Chu & La Peyre 1993).



 Transmission: Horizontal and direct, although gastropods may act as vectors

(Ford 1992). Although all three stages of the life-cycle (zoospores, meronts,

prezoosporangia) may effect transmission, the meront is the primary agent of

transmission in natural stocks (Chu 1996).



4. Host impact:



 Tissue tropism: P. marinus enters oysters via the external epithelia and may be

observed in the gut lumen, but not gut epithelia (Dungan et al. 1996). After entry

the organism becomes systemic, but is most common in the digestive gland and

least common in the haemolymph (Oliver et al. 1996).



 Brief description of major pathological and biological effects: After invasion

through surface epithelia, the pathogen goes through a cycle in which meronts

develop into zoospores by schizogony in presporangia, developing to sporangia.

The zoospore is released and has a flagellum, but it appears the meront is more

effective in establishing infections (Chu 1996). The minimum infectious dose

under experimental condition is 102 meronts (Chu & Volety 1997). After entry the

pathogen secretes several substances, with hydrolytic enzymes and serine

proteases thought to cause the widespread tissue damage observed (La Peyre et al.

1996). Haemocytes are unable to destroy phagocytosed parasites (Anderson

1996), possibly due to suppression of superoxide anion production (Anderson

1999). As infection progresses, oyster growth is slowed (Barber & Mann 1994,

Paynter 1996), and reproduction reduced (Kennedy et al. 1995). Death is probably

due to massive tissue damage.



5. Diagnostics and disease control



 Key diagnostic features: There are no key diagnostic features. The gross signs of

weakness and emaciation are typical of many oyster diseases. In moderate to

heavy infections, the parasite is readily recognizable in stained sections.



 Overview of diagnostic methods, including sensitivity and specificity:

Histology can be used for moderate to heavy infections, but is unreliable in light

infections. A more reliable method is incubation of pieces of tissue in Ray’s Fluid

Thioglycollate Medium (RFTM) (Bushek et al. 1994). However, RFTM

incubation takes at least 3 days, and therefore molecular tools have been

developed. A semiquantitative PCR assay has been developed that can detect 10

pg of total P. marinus DNA per 1 m g of oyster haemocyte DNA with ethidium

bromide staining of agarose gels; 100 fg total P. marinus DNA with Southern blot

autoradiography; and 10 fg of total P. marinus DNA with dot-blot hybridizations





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

69





(Marsh et al. 1995). A PCR-based assay was specific for P. marinus and did not

react with P. olseni, P. atlanticus and other Perkinsus spp.. It detected 1 P.

marinus in 30 mg of oyster tissue (Robledo et al. 1998). Comparison of PCR with

RFTM gave prevalences of 92-100% for PCR, but only 70-83% for RFTM, using

the same oysters for both assays (Robledo et al. 1998). A microplate ELISA using

polyclonal antibodies detected as little as 1.5 ng of P. marinus protein in

microplate wells also containing 100 m g of normal oyster protein (Dungan &

Hamilton 1997). Sensitive immunoassays have also been developed for the

detection of P. marinus in environmental samples (Dungan 1997, Yarnall et al.

1997).



 Disease management activities in major producing countries: Controls on

movement of oysters from contaminated areas are only partially effective, as the

parasite occurs in many other aquatic animals and appears to be spreading

naturally with increase in water temperature (Ford 1992, 1996). Oysters should be

moved to low salinity areas as temperatures rise in spring and early summer.



References



Anderson, R.S. (1996). Interactions of Perkinsus marinus with humoral factors and

hemocytes of Crassostrea virginica. J. Shellfish Res. 15: 127-134.



Anderson, R.S. (1999). Perkinsus marinus secretory products modulate superoxide

anion production by oyster (Crassostrea virginica) haemocytes. Fish Shellfish Immunol.

9: 51-60.



Barber, B.J., Mann, R. (1994). Growth and mortality of eastern oysters, Crassostrea

virginica (Gmelin, 1791), and Pacific oysters, Crassostrea gigas (Thunberg, 1793) under

challenge from the parasite, Perkinsus marinus. J. Shellfish Res. 13: 109-114.



Burreson, E.M. & Calvo, L.M. (1996). Epizootiology of Perkinsus marinus disease of

oysters in Chesapeake Bay, with emphasis on data since 1985. J. Shellfish Res. 15: 17-

34.



Bushek, D. & Allen, S.K. (1996). Host-parasite interactions among broadly distributed

populations of the eastern oyster Crassostrea virginica and the protozoan Perkinsus

marinus. Mar. Ecol. Prog. Ser. 139: 127-141.



Bushek, D., Ford, S.E. & Allen, S.K. (1994). Evaluation of methods using Ray’s fluid

thioglycollate medium for diagnosis of Perkinsus marinus infections in the eastern

oyster, Crassostrea virginica. Ann. Rev. Fish Dis. 4: 201-217.



Bushek, D., Holley, R. & Kelly, M. (1997). Chlorine tolerance of Perkinsus marinus. J.

Shellfish Res. 16: 260.



Calvo, L.M. & Burreson, E.M. (1994). In vitro and in vivo effects of eight

chemotherapeutants on the oyster parasite Perkinsus marinus (Mackin, Owen, and

Collier). J. Shellfish Res. 13: 101-107.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

70





Chu, F-L. E. (1996). Laboratory investigations of susceptibility, infectivity, and

transmission of Perkinsus marinus in oysters. J. Shellfish Res. 15: 57-66



Chu, F-L.E. & La Peyre, J.F. (1993). Perkinsus marinus susceptibility and defense-

related activities in eastern oysters Crassostrea virginica: temperature effects. Dis.

Aquat. Org. 16: 223-234.



Chu, F-L.E., La Peyre, J.F., Burreson, C.S. (1993). Perkinsus marinus infection and

potential defense-related activities in eastern oysters, Crassostrea virginica: salinity

effects. J. Invertebr. Pathol. 62: 226-232



Chu, F-L. E., Volety, A.K. (1997). Disease processes of the parasite Perkinsus marinus

in eastern oyster Crassostrea virginica: minimum dose for infection initiation, and

interaction of temperature, salinity and infective cell dose. Dis. Aquat. Org. 28: 61-68.



Chu F-LE, Volety AK, Constatin G (1996) A comparison of Crassostrea gigas and

Crassostrea virginica: Effects of temperature and salinity on susceptibility to the

protozoan parasite, Perkinsus marinus. J. Shellfish. Res 15: 375-380



Dungan, C. F. (1997). Perkinsus marinus: immunoassay detection in oyster tissues and

environmental samples and in vitro experimental systems. J. Shellfish Res. 16: 263.



Dungan, C.F. & Hamilton, R.M. (1997). Microplate ELISA assay for detection of

Perkinsus marinus in oyster tissues. J. Shellfish Res. 16: 330-331.



Dungan, C.F., Hamilton, R.M., Burreson, E.M. & Ragone-Calvo, L.M. (1996).

Identification of Perkinsus marinus portals of entry by histochemical immunoassays of

challenged oysters. J. Shellfish Res. 15: 500.



Faisal, M., La Peyre, J.F. & Kaattari, S.L. (1997). A promising chemotherapy for

Perkinsus marinus-infected oysters. J. Shellfish Res. 16: 263-264.



Ford, S.E. (1992). Avoiding the transmission of disease in commercial culture of

molluscs, with special reference to Perkinsus marinus (Dermo) and Haplosporidium

nelsoni (MSX). J. Shellfish Res. 11: 539-546.



Ford, S.E. (1996). Range extension by the oyster parasite Perkinsus marinus into the

northeastern United States: Response to climate change? J. Shellfish Res. 15: 45-56.



Kennedy, V.S., Newell, R.I.E., Krantz, G.E. & Otto, S. (1995). Reproductive capacity of

the eastern oyster Crassostrea virginica infected with the parasite Perkinsus marinus.

Dis. Aquat. Org. 23: 135-144.



Krantz, G.E. (1994). Chemical inhibition of Perkinsus marinus in two in vitro culture

systems. J. Shellfish Res. 13: 131-136.



La Peyre, J.F., Garreis, K.A., Yarnall, H.A. & Faisal, M. (1996). Emerging evidence of

extracellular proteases as important virulence factors of Perkinsus marinus. J. Shellfish

Res. 15: 501.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

71





Marsh, A.G., Gauthier, J.D. & Vasta, G.R. (1995). A semiquantitative PCR assay for

assessing Perkinsus marinus infections in the eastern oyster, Crassostrea virginica. J.

Parasitol. 81: 577-583.



Oliver, L.M., Fisher, W.S., Burreson, E.M., Ragone-Calvo, L.M., Ford, S.E. & Gandy, J.

(1996). Perkinsus marinus tissue distribution and seasonal variation in oysters

(Crassostrea virginica) from Florida, Virginia and New York. J. Shellfish Res. 15: 497.



Paynter, K.T. (1996). The effects of Perkinsus marinus infection on physiological

processes in the eastern oyster, Crassostrea virginica. J. Shellfish Res. 15: 119-125.



Pierce, J.C., Maloy, W.L., Salvador, L. & Dungan, C.F. (1997). Recombinant expression

of the antimicrobial peptide polyphemusin and its activity against the protozoan oyster

pathogen Perkinsus marinus. Mol. Mar. Biol. Biotechnol. 6: 248-259.



Robledo, J.A.F., Gauthier, J.D., Coss, C.A., Wright, A.C. & Vasta, G.R. (1998). Species

specificity and sensitivity of a PCR-based assay for Perkinsus marinus in the eastern

oyster, Crassostrea virginica: a comparison with the fluid thioglycollate assay. J.

Parasitol. 84: 1237-1244.



Yarnall, H.A, Stokes, N.A. & Burreson, E. M. (1997). Development of a PCR assay for

the quantitation of Perkinsus marinus. J. Shellfish Res. 16: 342-343.





5.2.12 PERKINSOSIS OF SCALLOPS (SPX)



1. Name and taxonomy of disease agent: SPX, perkinsosis of scallops, caused by

Perkinsus qugwadi (Perkinsidae, Dinoflagellata, Alveolata, Eukaryota).



2. Agent stability and inactivation data: P. qugwadi is pathogenic at low

temperatures (8-15°C) (Blackbourn et al. 1998).



3. Epidemiological features of the disease:



 Geographic range and features of distribution: British Columbia, Canada

(Bower et al. 1998), possibly Japan and Russia (see note).



 Host range: Japanese scallops (Patinopecten yessoensis), but the aetiological

agent is thought to be enzootic in western Canada, and therefore may exist at low

levels in other hosts. Native scallops (Chlamys rubida, Chlamys hastata) are

resistant to infection (Bower et al. in press). Japanese scallops show variable

susceptibility to infection, with those surviving epizootics having significant

resistance to infection (Bower et al. in press).



 Morbidity/mortality rates: 60% -100% (Bower et al. 1997, 1998).



 Transmission: Unknown but presumed to be horizontal and direct. As in P.

marinus, a vector may sometimes be involved.



4. Host impact:





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

72







 Tissue tropism: Connective tissue throughout the host (Bower et al. 1998,

Blackbourn et al. 1998).



 Brief description of major pathological and biological effects: The route of

invasion is unknown, but P. qugwadi proliferates systemically in connective

tissue, causing extensive tissue damage. Focal lesions appear as creamy white

pustules 5 mm in diameter on all organs, but particularly on the gonad, digestive

gland and mantle. Zoospores are only found in the interstitial spaces (Blackbourn

et al. 1998). The degree of cell-mediated response by the host often does not

reflect the degree of infection.



5. Diagnostics and disease control



 Key diagnostic features: Creamy white pustules 5 mm in diameter on organ

surfaces.



 Overview of diagnostic methods, including sensitivity and specificity:

Currently only histology can be used to detect infection. The pathogen cannot be

cultured by the RTFM method used to detect P. marinus (Blackbourn et al. 1998).



 Disease management activities in major producing countries: Controls on

movements and eradication of infected stocks.



Note: It is unclear whether P. qugwadi is the same Perkinsus sp. reported from

Patinopecten yessoensis in Japan and Russia (Kurochkin et al. 1986).



References



Blackbourn, J., Bower, S.M. & Meyer, G.R. (1998). Perkinsus qugwadi sp. nov.

(incertae sedis), a pathogenic protozoan parasite of Japanese scallops, Patinopecten

yessoensis, cultured in British Columbia, Canada. Can. J. Zool. 76: 942-953.



Bower, S.M., Blackbourn, J. & Meyer, G.R. (1997). A new and unusual species of

Perkinsus pathogenic to cultured Japanese scallops, Patinopecten yessoensis, in British

Columbia, Canada. J. Shellfish Res. 16: 333.



Bower, S.M., Blackbourn, J. & Meyer, G.R. (1998). Distribution, prevalence, and

pathogenicity of the protozoan Perkinsus qugwadi in Japanese scallops, Patinopecten

yessoensis, cultured in British Columbia, Canada. Can. J. Zool. 76: 954-959.



Bower, S.M., Blackbourn, J. & Meyer, G.R. & Nishimura, D.J.H. (1992). Diseases of

cultured Japanese scallops (Patinopecten yessoensis) in British Columbia, Canada.

Aquaculture 107: 201-210.



Bower, S.M., Blackbourn, J. & Meyer, G.R. & Welch, D.W. (in press). Effect of

Perkinsus qugwadi on various species and strains of scallops. Dis. Aquat. Org. (see

www.int-res.com/journals/dao/daoForthcoming.html).









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

73





Kurochkin, Y.V., Tsimbalyuk, E.M. & Rybakov, A.V. (1986). Parazitî bolyyezni

(Parasites and diseases). In: Primorskii grebeshok. (The yezo scallop or Japanese

common scallop Mizuhopecten yessoensis [Ja], in Russian). Institute of Marine

Biology, Far East Science Centre, Academy of the U.S.S.R., Vladivostok. pp. 174-

182.





5.2.13 CLAM PERKINSUS DISEASE



1. Name and taxonomy of disease agent: Clam Perkinsus disease, caused by

Perkinsus atlanticus (Perkinsidae, Dinoflagellata, Alveolata, Eukaryota). Perkinsus

atlanticus shows similarities to Perkinsus olseni (Hamaguchi et al. 1998), but

Perkinsus marinus also shows similarities to P. atlanticus (Robledo et al. 1997).



2. Agent stability and inactivation data: Not reported, but P. atlanticus is very

closely related to, if not conspecific with, Perkinsus olseni, for which the following

survival parameters are known. P. olseni survives at least one day at 0°C and 4°C, and

at least 197 days at -60°C. Free prezoosporangia are killed within 30 minutes in 6

ppm of chlorine, whilst those enclosed in tissue survive for >2 hours. Free

prezoosporangia do not survive for 6 hours in 7 ‰

seawater, and 50%.



 Transmission: Life cycle direct, apparently ovoviviparous (Fleming et al

1981).



4. Host impact:



 Tissue tropism: The gill is affected.



 Brief description of major pathological and biological effects: Large

numbers cause unsightly appearance of the gills. Histologic changes reported

as infiltration of hemocytes around the turbellarians and engorged adjacent

blood sinuses. No detectable effect on condition index.



5. Diagnostics and disease control



 Key diagnostic features: Small oval or pyriform turbellarians 2 mm long,

on gill surfaces and between gill lamellae. Uniform ciliation, scattered

vitellaria, posterior terminal oral-genital pore, muscular penis bulb (Cannon

1986; Goggin & Cannon 1990; Bower et al 1994).



 Overview of diagnostic methods, including sensitivity and specificity:

Light microscopy of fresh gills. Histology of sections through the gill and

palp area.



 Disease management activities in major producing countries: Maintain

oysters in 8 o/oo for 8 days at 20oC (Bataller & Boghen, 2000)



References



Bataller,E. & Boghen, A.D. (2000) Elimination of the gill worm Urastoma cyprinae

(Graff) from the eastern oyster Crassostrea virginica (Gamelin) using different

salinity-temperature combinations. Aquaculture 182:199-208.



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994) Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann.Rev.Fish Dis. 4:1-199.



Caceres-Martinez, J., Vasquez-Yeomans,R., Sluys, R. (1998) The turbellarian

Urastoma cyprinae from edible mussels Mytilus galloprovincialis and Mytilus

californianus in Baja California, NW Mexico. J.Invert.Pathol. 72:214-219









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

84





Cannon, L.R.G (1986) Turbellaria of the World – a guide to families and genera..

Queensland Museum, Brisbane, 132 pp.



Goggin, C.L. & Cannon, L.R.G. (1990). Occurrence of a turbellarian from Australian

tridacnid clams. J.Invert.Pathol. 56:135-138.



Fleming, L.C., Burt, M.D.B., & Bacon, G.B. (1981) On some commensal Turbellaria

of the Canadian east coast. Hydrobiologia 84:131-137.



Lauckner, G. (1983). Diseases of Mollusca:Bivalvia. In Diseases of Marine Animals,

O. Kinne ed., Biologische Anstalt Helgoland, Hamburg. Vol. 2.



Villalba, A., Mourelle, S.G., Carballal, M.J., Lopez, C. (1997) Symbionts and

diseases of farmed mussels Mytilus galloprovincialis throughout the culture process in

the Rias of Galicia (NW Spain). Dis.Aq.Org. 31:127-139.





7.2 HELMINTH DISEASES IN BIVALVES NOT KNOWN IN AUSTRALIA



7.2.1 TURBELLARIA, POLYCLADIDA



1. Name and taxonomy of disease agent: Turbellaria (Stylochus pilidium, Stylochus

ellipticus, Stylochus frontinalis, Pseudostylochus ostreophagus, Stylochus sp.)

(Turbellaria, Polycladida).



2. Agent stability and inactivation data: Highly susceptible to dessication..

Readily killed by drying or freshwater.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: S. pilidium Mediterranean,

S. ellipticus Eastern USA, S. frontinalis Florida, Pseudostylochus

ostreophagus Western USA, Stylochus sp. Korea.



 Host range: Predators of oysters and other marine bivalves.



 Morbidity/mortality rates: Over 50% of oyster spat (Lauckner, 1983)



 Transmission: Life cycle is direct.



4. Host impact:



 Tissue tropism: The mantle cavity.



 Brief description of major pathological and biological effects: Readily

attack and kill weakened oysters. Reported to be primary predators on oyster

spat and other juvenile bivalves (Lauckner, 1983; Jennings & Newman, 1996).

Agents enter the mantle cavity, evert the pharynx and lyse oyster tissue.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

85







5. Diagnostics and disease control:



 Key diagnostic features: Presence of lesions containing polyclads with

reticulate dark colour, tentacles forward. Pharynx ruffled, much folded,

central. Copulatory apparatus anterior to male pore, prostatic vesicles free,

gonopores separate in posterior half of body (Cannon 1986). Species can be

separated by colour pattern, size and male reproductive structures (Jennings &

Newman, 1996)



 Overview of diagnostic methods, including sensitivity and specificity:

Gross examination for adult worms which are 10mm or greater in length.



 Disease Management activities in major producing countries: Not known.



References:



Cannon, L.R.G. (1986) Turbellaria of the World. Qld Museum, Brisbane



Jennings, K.A. & Newman, L.J. (1996). Four new Stylochid flatworms

(Platyhelminthes:Polycladida) associated with commercial oysters from Moreton Bay,

southeast Queensland, Australia. Raffles Bull. Zool. 44:493-508.



Lauckner, G. (1983) Diseases of Mollusca:Bivalvia. In Diseases of Marine Animals,

O. Kinne ed., Biologische Anstalt Helgoland, Hamburg. Vol 2.





7.2.2 TURBELLARIA, RHABDOCOELA, GRAFFILLIDAE



1. Name and taxonomy of disease agent: Turbellaria (Paravortex spp and others)



2. Agent stability and inactivation data: Delicate organisms outside shellfish, well

protected in gut.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: North Sea, Mediterranean

(Paravortex scrobiculariae); North Sea (P. cardii); Chesapeake Bay, Gulf of

Mexico, Eastern Canada (P. gemmellipara); Northwest USA (Graffila

pugetensis) (Schell 1986).



 Host range: Mytilidae, Cardiidae, Myoida, Tellinidae, Ostreiidae, other

bivalves.



 Morbidity/mortality rates: None recorded. Prevalence 50-70% in some

areas.



 Transmission: Life cycle is direct.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

86







4. Host impact:



 Tissue tropism: Paravortex spp found in the gut (Lauckner 1983); G.

pugetensis in the pericardial cavity (Schell 1986).



 Brief description of major pathological and biological effects: None

described for Paravortex. G. pugetensis displaces and constricts heart of

Macoma (Tellinidae) (Schell 1989).



5. Diagnostics and disease control:



 Key diagnostic features: Mouth in anterior of body, testes paired, sacciform

or lobed. Male copulatory organ unarmed or with simple stylet. Germinal

region anterior, without vagina, gonopore in anterior half of body (Cannon

1986).



 Overview of diagnostic methods, including sensitivity and specificity:

Examination of squashed tissues under dissecting microscope best.

Occurrence of turbellarian detectable in appropriate histological section

(Bower et al 1994). Species identification generally requires examination of

live specimens.



 Disease management activities in major producing countries: None.



References



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994) Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann.Rev.Fish Dis. 4:1-199.



Lauckner, G. (1983) Diseases of Mollusca:Bivalvia. In Diseases of Marine Animals,

O. Kinne ed., Biologische Anstalt Helgoland, Hamburg. Vol 2.



Schell, S.C. (1989) Graffilid from pericardial cavity of Macoma. J.Parasitol. 75:428-

430.





7.2.3 DIGENEANS (GROUP 1) SPOROCYSTS AND REDIAE

- Mollusc as 1st intermediate host.



1. Name and taxonomy of the disease agent: Bucephalidae, Sanguinicolidae,

Monorchiidae, Fellodistomidae, Gymnophallidae



2. Agent stability and inactivation data: Stable within mollusc. Unlikely to

survive freezing.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Varies with species. NW

Europe,France, Germany, Japan, UK, Sweden, Italy, Eastern USA.





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

87







 Host range: Wide range of species affected. Bucephalidae - Mytilus spp,

Ostrea lutaria, Pinctada martensi . Monorchiidae - Cardium edule (Cercaria

cerastodermae). Fellodistomidae - . Tapes spp Gymnophallidae - Mya

arenaria (Bower et al 1994; Lauckner 1983)



 Morbidity/mortality rates: Some species are direct cause of morbidity and

mortality.



 Transmission: Complex life cycles that require a mollusc, a vertebrate and a

third host.



4. Host impact:



 Tissue tropism: Digestive gland and gonad.



 Brief description of major pathological and biological effects: Castrate and

sterilise infected molluscs. Parasites displace gonad and much of digestive

gland. Major physiological changes and depletion of body reserves eventually

debilitate host.



5. Diagnostics and disease control:



 Key diagnostic features: Gross detection usually possible after opening

shell. Confirm presence of sporocysts or rediae or cercariae using dissecting

microscope. Easily seen in histological sections. Often not possible to

identify to genus without further experimental work.



 Overview of diagnostic methods, including sensitivity and specificity:

Identification by microscopy.



 Disease management activities in major producing countries: Not known.





References



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994) Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann.Rev.Fish Dis. 4:1-199.



Lauckner, G. (1983) Diseases of Mollusca:Bivalvia. In Diseases of Marine Animals,

O. Kinne ed., Biologische Anstalt Helgoland, Hamburg. Vol 2.







7.2.4. DIGENEANS (GROUP 2) METACERCARIAE

- Molluscs as 2nd intermediate host.



1. Name and taxonomy of disease agent: Monorchiidae, Fellodistomidae,

Echinostomatidae, Psilostomatidae, Renicolidae, Gymnophallidae.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

88





2. Agent stability and inactivation data: Stable within mollusc. Unlikely to

survive freezing.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Varies with species. Baltic

states, UK, North Sea, Japan, Holland, NW Europe.



 Host range: Wide range of species affected. Monorchiidae – Tellina,

Macoma, Cardium spp. Echinostomatidae – Cardium edule, Mytilus edullis.

Renicolidae – European mussels and cockles. Gymnophallidae – Cardium

spp, Macoma.



 Morbidity/mortality rates: Direct cause of morbidity and mortality reported

for some species.



 Transmission: Complex life cycles that require a mollusc, a vertebrate and a

third host.



4. Host impact:



 Tissue tropism: Penetrate soft tissue and encyst in a variety of areas

including siphon, kidney, foot, palps, mantle.



 Brief description of major pathological and biological effects: Interference

with function of organs, reduce resistance of the host to thermal, osmotic and

starvation stress. Cause gaping, poor growth, general debilitation, reduced

survival, inhibition of shell growth and shell deformities.



5. Diagnostics and disease control:



 Key diagnostic features: Readily seen in squash preparations under

compound microscope, and in appropriate histological sections.

Metacercariae of G. tokiensis predominantly around the labial palps (Bower et

al 1994). Metacercariae of Meiogymnophallus minutus under the hinge.

Often not possible to identify to genus without further experimental work.



 Overview of diagnostic methods, including sensitivity and specificity:

Main method of diagnosis is microscopy.



 Disease management activities in major producing countries: Not known.



References



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994) Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann.Rev.Fish Dis. 4:1-199.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

89







7.2.5 CESTODA



1. Name and taxonomy of disease agent: Lecanicephalida, Tetraphyllidea,

Trypanorhyncha. Representatives of these orders are found as juveniles

(metacestodes) in many species of bivalves.



2. Agent stability and inactivation data: Stable at temperatures suitable for the

host. Not usually killed by a host response. Probably all unable to survive

freezing.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Varies with species.

Usually present in cysts in small numbers. Adults occur in elasmobranchs.



 Host range: Varies with species.



 Morbidity/mortality rates: Some reports of mortality, not substantiated.



 Transmission: Probably through ingestion of eggs. Adults develop in the

spiral valve of elasmobranchs.



4. Host impact: Probably small.



 Tissue tropism: Metacestodes of leucanicephalids and trypanorhynchs

usually encapsulated in the Leidig tissue; tetraphyllideans free in the

lumen of the intestine.



 Brief description of major pathological and biological effects: Chronic

inflammation around the parasites in tissue; tissue displacement; no known

changes in intestinal forms. No significant tissue damage to the

elasmobranch definitive host.



5. Diagnostics and disease control:



 Key diagnostic features: Detect in unstained fresh squashes of host

tissue under low power of microscope. Easily seen in H&E stained

histological sections. It is not possible to identify lecanicephalids or

tetraphyllideans to species from the stages found in the mollusc,

particularly as many species remain undescribed even as adults.

Tryphanorhynchs of described species can be identified after eversion of

the spiny tentacles.



 Overview of diagnostic methods, including sensitivity and specificity:

Main diagnostic method is microscopy. Probably moderate sensitivity and

low specificity.



 Disease management activities in major producing countries: None

known. Gut parasites may be amenable to treatment by host starvation or





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

90





by an anthelmintic treatment in the water. Tissue parasites difficult to

treat.





7.2.6 ANNELIDA



1. Name and taxonomy of disease agent: Polydora hoplura, P. variegata,

Boccardia spp., Sabellidae. The taxonomy and biology of polychaetes is

relatively poorly known.



2. Agent stability and inactivation data: Australian infections of Polydora

websteri and Boccardia sp. in Sydney rock oysters are controlled by drying out

the shells at each tide. Boccardia knoxi on abalone shells can also be controlled

through drying the shells. Control method for sabellid on abalone shells in

California – hot fresh water, wax shells.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: An unknown sabellid has

become a serious pest of Californian abalone, introduced from South Africa

with stock.



 Host range: Wide range of mollusc shells infected. Successful colonisation

may depend more on the environment than the host species.



 Morbidity/mortality rates: Not known.



 Transmission: Direct, via trochophore larvae.



4. Host impact:



 Tissue tropism: The shell.



 Brief description of major pathological and biological effects: Reduce

growth, weaken the molluscs and deform the shells.



5. Diagnostics and disease control:



 Overview of diagnostic methods, including sensitivity and specificity:

Readily detected with a dissecting microscope. Difficult to identify to species.



 Disease management activities in major producing countries: Inspection of

seed before sale, improved farm sanitation practices, use of screens on farm

discharges. Regular drying of shells. Possible introduction of toxins with the

particulate feed of the sabellids.



References



Handlinger,J. (2000) pers. com.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

91





McBride, S.C. (1998) Current status of abalone aquaculture in the Californias.

J.Shellfish.Res. 17:593-600



Montague, P. (1998) Aquaculture CRC Ltd. Annual Report for 1997-8. pp 17-18.



Shields, J.D., Buchal, M.A., Friedman, C.S. (1998) Microencapsulation as a potential

control technique against sabellid worms in abalone culture. J.Shellfish.Res 17:79-83.







8.0 PARASITIC CRUSTACEANS



8.1 PARASITIC CRUSTACEANS IN BIVALVES IN AUSTRALIA



The only crustaceans reported from bivalves in Australia have been the copepods

Pseudomyicola spinosus and Myicola spp. in blue mussels (Mytilus edulis) (Pregenzer

1983). P. spinosus was found to prefer warmer waters and mussels near muddy or

silty seabeds. Several of the other genera that infest the gills of bivalves (Modiolicola,

Ostrincola, Conchylirus, Myocheres, Paranthessius) are probably symbionts which

have not been reported to cause any pathology in their hosts (Bower et al. 1994).

However, there are some claims that Mytilicola intestinalis harms its host, and

therefore it will be included in this review.



Reference



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994). Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4: 1-199.



Pregenzer, C. (1983). Survey of metazoan symbionts of Mytilus edulis (Mollusca:

Pelecypoda) in Southern Australia. Aust. J. Mar. Freshwat. Res. 34: 387-396.





8.2 PARASITIC CRUSTACEANS IN BIVALVES NOT KNOWN IN

AUSTRALIA



8.2.1 RED WORM DISEASE



1. Name and taxonomy of disease agent: Red worm disease caused by Mytilicola

intestinalis (Myicolidae, Copepoda).



2. Agent stability and inactivation data: Developmental rate is faster at 18°C and

decreases at >18°C (Gee & Davey 1986)..



3. Epidemiological features of the disease:



 Geographic range and features of distribution: European waters from Denmark

to Italy, including the U.K. and Ireland.



 Host range: Infects oysters (Ostrea edulis), clams (Ruditapes decussatus,

Macoma balthica), cockles (Cerastoderma edule) and mussels (Mytilus edulis,





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

92





Mytilus galloprovincialis), and many other bivalves, but not reported from

scallops.



 Morbidity/mortality rates: Heavy infection may reduce growth and condition

(Bayne et al. 1978, Durfort et al. 1982) at high temperatures (22-23°C), but

mortalities have not been reported.



 Transmission: Planktonic stages taken in during feeding.



4. Host impact:



 Tissue tropism: Lives in the main gut.



 Brief description of major pathological and biological effects: Heavy

infestations may lead to replacement of the ciliated columnar cells of the gut with

non-ciliated cuboidal cells (Moore et al. 1978), the ovacytes may show

ultrastructural alterations (Durfort et al. 1982), and feeding rate, and therefore

growth and condition, may be affected (Bayne et al. 1978). However, when other

factors that may influence condition are taken into account (host length, gonad

development, seasonal cycles, environmental parameters), the impact of the

copepod is negligible (Gee et al. 1977).



5. Diagnostics and disease control



 Key diagnostic features: The red elongated copepods can easily be seen

macroscopically.



 Overview of diagnostic methods, including sensitivity and specificity: Gross

observation or histology. Sensitivity is high, but high specificity requires

examination by expert practitioners.



 Disease management activities in major producing countries: None, but M.

intestinalis can be controlled by treatment with Dichlorvos at 30 mg/l for 2 hours

(Blateau et al. 1992).



References



Bayne, B.L., Gee, J.M., Davey, J.T. & Scullard, C. (1978). Physiological responses of

Mytilus edulis L. to parasitic infestation by Mytilicola intestinalis. J. Cons. CIEM 38: 12-

17.



Blateau, D., Le Coguic, Y., Mialhe, E. & Grizel, H. (1992). Mussel (Mytilus edulis)

treatment against the red copepod Mytilicola intestinalis. Aquaculture 107: 165-169.



Durfort, M., Bargallo, B., Bozzo, M.G., Fontarnau, R. & Lopez-Camps, J. (1982).

Alteration of the oocytes of Mytilus edulis, L. (Mollusca, Bivalvia) due to infestation of

the mussel by Mytilicola intestinalis Steuer (Crustacea, Copepoda). Malacologia 22:55-

59.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

93





Gee, J.M. & Davey, J.T. (1986). Stages in the life history of Mytilicola intestinalis

Steuer, a copepod parasite of Mytilus edulis (L.), and the effect of temperature on their

rates of development. J. Cons. CIEM 42: 254-264.



Gee, J.M., Maddock, L. & Davey, J.T. (1977). The relationship between infestation by

Mytilicola intestinalis, Steuer (Copepoda, Cyclopoidea) and the condition index of

Mytilus edulis in southwest England. J. Cons. CIEM 37: 300-308.



Moore, M.N., Lowe, D.M. & Gee, J.M. (1978). Histopathological effects induced in

Mytilus edulis by Mytilicola intestinalis and the histochemistry of copepod intestinal

cells. J. Cons. CIEM 38: 6-11.





8.2.2 BROOD-POUCH COPEPOD



1. Name and taxonomy of disease agent: Brood-pouch copepod on scallop gills,

caused by Pectenophilus ornatus (Copepoda, Crustacea).



2. Agent stability and inactivation data: Not reported.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Japan.



 Host range: Scallops, Patinopecten yessoensis, Chlamys farreri.



 Morbidity/mortality rates: Infects 100% of Japan’s scallop populations,

reduces host condition (Nagasawa & Nagata 1992), but mortalities not reported.



 Transmission: Moves directly from host to host.



4. Host impact:



 Tissue tropism: Attaches orally to the gill arch.



 Brief description of major pathological and biological effects: The attachment

site consists of hypertrophied host tissue which is perfused with host blood on

which the copepod feeds.



5. Diagnostics and disease control



 Key diagnostic features: A bright yellow sac-like female, up to 8 mm in width,

covered up by the scallop’s ciliated gill epithelium. It resembles a barnacle (Elston

et al. 1985)



 Overview of diagnostic methods, including sensitivity and specificity: Can be

easily visualized because of colour and size. Specificity and sensitivity high.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

94





 Disease management activities in major producing countries: None, but

infection can be reduced or avoided by hanging scallops well off the bottom

sediments (Nagasawa et al. 1991).



References



Elston, R.A., Wilkinson, M.T. & Burge, R. (1985). A rhizocephalan-like parasite of a

bivalve mollusc, Patinopecten yessoensis. Aquaculture 49: 359-361.



Nagasawa, K. & Nagata, M. (1992). Effects of Pectenophilus ornatus (Copepoda) on

the biomass of cultured Japanese scallop Patinopecten yessoensis. J. Parasitol. 78:

552-554.



Nagasawa, K., Takahashi, K., Tanaka, S. & Nagata, M. (1991). Ecology of

Pectenophilus ornatus, a copepod parasite of the Japanese scallop Patinopecten

yessoensis. Bull. Plankton Soc. Japan 1991: 495-502.







B. SIGNIFICANT DISEASES OF UNKNOWN AETIOLOGY



1. Name and taxonomy of disease agent: Malpeque disease.



2. Agent stability and inactivation data: Low salinity retards the disease.



3. Epidemiological features of the disease:



 Geographic range and features of distribution: Atlantic Canada.



 Host range: Only reported from Crassostrea virginica.



 Morbidity/mortality rates: Mortality rates 90%, but morbidity rates not

reported.



 Transmission: Unknown, but described as “highly infectious” (Bower et al.

1994).



4. Host impact:



 Tissue tropism: The disease occurs throughout the oyster, and is marked by a

haemocytosis, with haemocyte infiltration into all tissues, particularly connective

tissue.



 Brief description of major pathological and biological effects: Grossly the

affected oysters show mantle regression, gaping, oedema, and abscesses in the

mantle. Yellow to green pustules occur on the inner surface of the shell.

Histologically there are accumulations of ceroid, especially within the digestive

gland, oedema of the mantle, retardation in gonad development, connective tissue

abscess-like lesions and focal infiltration. The haemocytes have enlarged nuclei







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

95





and reduced cytoplasm, giving the superficial appearance of haemocytic

neoplasia.



5. Diagnostics and disease control



 Key diagnostic features: The combination of clinical signs and histopathology

given above suggest the presence of Malpeque disease, but individually some of

the signs are non-specific (mantle regression, gaping). Haemocytoses are a non-

specific response to a variety of factors (infectious disease, environmental stress,

trauma). The abcess-like lesions are probably the most characteristic feature of

diagnostic value, but very similar lesions occur in Australian flat oysters (Ostrea

angasi) infected with Bonamia sp.



 Overview of diagnostic methods, including sensitivity and specificity:

Histology is still the primary method for diagnosis of Malpeque disease. Attempts

at culturing organisms on marine agars have resulted in isolation of a fungus-like

organism, Labyrinthomyxa (Li et al 1980), but this is now regarded as an

opportunistic saprophyte infecting moribund oysters. Overall specificity and

sensitivity of diagnostic methods low/moderate.



 Disease management activities in major producing countries: Cessation of

movement of oysters from the affected area.



Note: From recent examination of infected tissues by one of us (P.M. Hine), it appears

that some of the abnormal haemocytes contain an intracytoplasmic microcell-like

organism. Microcell-like organisms in other hosts include Mikrocytos mackini causing

Denman Island disease in Crassostrea gigas off the western coast of Canada, Mikrocytos

roughleyi causing winter mortality in Sydney rock oysters (Saccostrea commercialis)

from Australia, and Bonamia spp. in Ostrea, Tiostrea, and Crassostrea. The similarity of

the abcess-like lesions in O. angasi caused by Bonamia sp. to the abcess-like lesions of

Malpeque disease is striking. However, changes in the appearance of haemocytes toward

more blast-like cells, and aggressive infiltration into connective tissue, may indicate an

infectious neoplastic disease, and retroviral neoplasia has been reported from clams

(Medina et al. 1993).



References



Bower, S.M., McGladdery, S.E. & Price, I.M. (1994). Synopsis of infectious diseases

and parasites of commercially exploited shellfish. Ann. Rev. Fish Dis. 4: 1-199.



Li, M.F., Traxler, G.S., Clyburne, S. & Stewart, J.E. (1980). Malpeque disease: isolation

and morphology of a Labyrinthomyxa-like organism from diseased oysters. ICES

COUNCIL MEETING 1980. ICES-CM-1980/F:15. ICES COPENHAGEN. 9 pp.



Medina, D.J., Paquette, G.E., Sadasiv. E.C. & Chang, P.W. (1993). Isolation of

infectious particles having reverse transcriptase activity and producing hematopoietic

neoplasia in Mya arenaria. J. Shellfish Res. 12: 112-113.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

96





C. BIVALVE MOLLUSCS AS PASSIVE VECTORS OF NOXIOUS

ORGANISMS



By virtue of their filtration capacity and, in the case of bivalves traded in the shell, the

water contained between the valves, these molluscs are capable of passively

transferring/transmitting a range of noxious organisms. This propensity is well recorded

in human medicine for viruses and bacteria, but documentation is lacking for pathogens

of veterinary importance. The ability of bivalves to carry vegetative stages or cysts of

toxic microalgae is insignificant in comparison to ballast water. It is quite feasible that

bivalves grown in proximity to fish farms experiencing epizootics of viral or bacterial

diseases, could contain significant titres of pathogens. If these molluscs were used as

feed for ornamental or other fish, or for fish bait, then it would be possible for these

pathogens to be transmitted.





OVERVIEW AND GENERAL COMMENTS



The serious and potentially serious diseases of bivalves listed above are summarised in

Appendix 1. From this the impression may be gained that oysters pose more of a risk

than other bivalves. Certainly the most serious diseases known are the OIE listed

diseases, and all of these are diseases of oysters, with only Marteilia refringens and

Perkinsus olseni infecting other bivalves. However, the diseases of oysters must be put

in context. Oysters have been cultured and moved around longer than the other bivalve

groups, and therefore more diseases may be expected to have emerged. Also, since the

last century, oysters have become the food of the wealthy, and have been cultured

intensively in developed countries (Europe, North America, Australasia) with scientific

expertise on hand to investigate disease problems. It is likely that serious diseases will

emerge in other bivalve groups and species as intensive culture expands.



A second impression that may be gained is that global bivalve diseases are largely

centred around France, Iberia, and North America (Appendix 2). Again this reflects the

availability of expertise, and aquaculture of these species. France has carried out more

research (assessed by scientific publications, and the existence of dedicated laboratories)

on all groups, than any other European country, and Spain and Portugal have undertaken

more studies on blue mussels (Mytilus spp.) and carpet clams (Ruditapes spp.), reflecting

their predominance in the aquaculture of these species. Some parasites and diseases have

emerged following the intercontinental movement of live animals for aquaculture.

Bonamia ostreae emerged as a pathogen following the movement of Ostrea edulis from

California to France, Perkinsus atlanticus is probably a subspecies of Perkinsus olseni

that was moved from Southeast Asia in Manila clams (Ruditapes philippinarum) to

Europe where it has subsequently drifted genetically, and spread into the naïve and more

susceptible native clam, Ruditapes decussatus. Similarly, Haplosporidium nelsoni causes

little disease in its natural host, Pacific oysters (Crassostrea gigas), but following

movement from Japan to the U.S.A., it has spread into the naïve and more susceptible

native oyster, Crassostrea virginica, causing haplosporidiosis (MSX). The infection of

several species of oyster (Crassostrea rivularis, Ostrea angasi, Tiostrea chilensis) with

Bonamia ostreae, Marteilia refringens and Minchinia armoricana in France, has

occurred after those oyster species were introduced into France for assessment as

aquaculture species, the infections being acquired after introduction.







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

97





The risks of introducing pathogens differs with different groups and their different

biology (Appendix 3). The haplosporidians (Bonamia, Haplosporidium, Minchinia) are

a high risk group because, as can be seen from the examples of B. ostreae and H. nelsoni

in the preceding paragraph, they cause serious disease when naïve hosts are exposed to

them. In the case of H. nelsoni it appears that even the requirement for an alternative

host is not limiting. The two stage movement of H. nelsoni from to Japan to the U.S.

west coast, and from the west coast to the U.S. east coast, suggests that suitable

alternative hosts belong to a widespread genus or genera. Similarly, Marteilia refringens

and Marteilia sydneyi readily establish at the site of introduction after movement,

suggesting the alternative host(s) must be common and widespread species. Perkinsus

spp. must also be regarded as a high risk genus at present because P. olseni (and

therefore probably the other Perkinsus spp.) can withstand prolonged freezing,

transmission is direct, and the distinction of species is confused. Consequently the OIE

has set up a network of laboratories that will sequence specified areas of the Perkinsus

genome from isolates obtained from the range of this genus, to determine how many

Perkinsus species exist and how their spread can be minimised. Until this is done,

Australia should consider any Perkinsus from outside national boundaries as exotic.



While the pathogens listed above may pose a risk if imported in a viable state in non-

viable product, importation of bivalves (particularly as chilled meats, or in the half shell)

from countries outside of Europe, North America and Australasia may pose just as much

risk. The health status of bivalves in Africa, the former U.S.S.R., Asia, and Central and

South America is almost totally unknown. It would be naïve to think that these vast areas

have less disease, because less has been reported. In Asia, for example, much more is

known about the impact of disease on the, much more studied, prawn farming industry,

than on molluscs. The economic loss attributed to outbreaks of disease in developing

countries in the Asian region was estimated to be at least US$1.4 thousand million in

1990, and the economic loss from prawn farms in 1993 in China alone was around

US$1.0 thousand million. In Thailand, yellowhead virus (YHV) caused losses of

5,000 metric tons in 1992, while in 1996 white spot syndrome virus (WSSV) was

blamed for losses of 40% of total production (70,000 metric tons) valued at over

US$500 million. Recent estimates, based on farm surveys in 16 Asian countries,

suggest that disease and environment-related problems have been associated with

annual losses of more than US$ 3 thousand million per year to aquaculture production

in Asian countries.



Similar levels of disease may also affect molluscs, but few data are available on

molluscan culture in Asia. For example, in 1994 live pearl oysters (Pinctada fucata)

were moved from Hainan Dao off the coast of China, to Japan. It was known that large

scale mortalities were occurring on Hainan Dao at the time, and soon identical

mortalities began to occur in Japan. From 1996 to the present, the annual mortality has

been 400 million akoya oysters in the western regions of Japan, constituting 50% of

Japanese stocks. The annual loss of oysters alone has been calculated at US$300

million, with a further loss of >50 tonnes of pearls annually. This disaster has not yet

become well known, because past surpluses in pearl production are being placed on

the market to supply current demand.



The risks of an exotic pathogen being introduced into Australian waters from imported

non-viable bivalves would seem to be small. The two areas of enhanced risk are

importation of oysters in the half shell, and chilled meats. Oysters in the half shell are





C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

98





sold in the half shell, and after consumption the shells are discarded. If discarded near or

into seawater, any pests infesting the shell may become established in the shells of other

molluscs. Probably the greatest risk is posed by boring sponges, as they are non-host

specific, and appear to be able to survive in seawater from subpolar regions to the

tropics. Several species infest the shells of living pearl oysters, making the nacre

valueless as mother-of-pearl. Mudworms exotic to Australia may also be introduced by

this route. It should be noted that ballast water, and oysters or mussels encrusting the

hulls of ships, are also likely to be a route of introduction of shell pests. Encrusting

bivalves may also introduce pathogens. For example, Marteilioides chungmuensis is a

protistan that infects the ova of several oyster species (Saccostrea echinata, Saccostrea

commercialis, Crassostrea gigas), and it has had a major impact on the fecundity of C.

gigas in Korea and Japan. In countries from which this pathogen has been reported, it

initially occurs in ports. In Australia it occurs in S. echinata in Darwin Harbour, but not

in surrounding bays (eg Bynoe Harbour).



There are no data on survival of bivalve pathogens in chilled meats, and therefore the

risk has to be deduced from general principles. The relative risk of introduction of

pathogens in chilled meats depends on the temperature to which the meat is chilled, and

the length of time between chilling and consumption. Chilled meats are usually

consumed, but mussel meat may be used as bait or feed for aquarium or aquaculture

species, allowing viable organisms direct or indirect entry into coastal waters. (Some

fish may act as hosts, otherwise the agent would have to survive digestion). Research is

needed to assess the real risks from chilled and frozen meats.









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

99









Appendix 1. A summary of the serious, or potentially serious, diseases,

parasites and pests, in different host groups and different host species.



EDIBLE OYSTERS (Ostreidae)



Disease/parasite/pest Geographical distribution

Crassostrea virginica

Juvenile oyster disease (JOD) Northeastern U.S.A.

Sirolpidium East coast, U.S.A.



zoophthorum

Mikrocytos mackini, Denman British Columbia, Canada,

Island disease experimental infection

Haplosporidium East coast of the United States,

from Maine to Florida

nelsoni

MSX

Haplosporidium costale, SSO Virginia and Maryland, Delaware

Bay, Washington State, U.S.A.

Perkinsus marinus East coast of North and Central

Dermo America to Venezuela

Malpeque disease Atlantic Canada

Crassostrea gigas

Oyster velar virus disease (OVVD) Washington State, U.S.A.

Gill necrosis virus disease (GNV) France, Portugal, Spain, U.K.

Haemocytic infection virus disease France, Spain

Summer mortality, nocardiosis British Columbia to California

Ostracoblabe implexa Widespread



Marteilia refringens Atlantic Europe from southern

England to Portugal

Mikrocytos mackini, Denman British Columbia, Canada

Island disease

Haplosporidium East coast of the United States,

Japan, France

nelsoni

MSX

Perkinsus marinus East coast, U.S.A.

Dermo

Crassostrea rivularis

Bonamia ostreae France

Ostrea edulis

Ostracoblabe implexa Widespread



Marteilia refringens Atlantic Europe from southern

England to Portugal

Bonamia ostreae East and West coasts, U.S.A.,

Atlantic and Mediterranean Europe

Mikrocytos mackini, Denman British Columbia, Canada,

Island disease experimental infection

Minchinia armoricana France, the Netherlands









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

100





Ostrea angasi

Marteilia refringens France



Bonamia ostreae France



Minchinia armoricana France

Ostrea puelchana

Bonamia ostreae Argentina

Tiostrea chilensis

Marteilia refringens France



Bonamia ostreae Chile

Ostreola conchaphila

Mikrocytos mackini, Denman British Columbia, Canada,

Island disease experimental infection

Saccostrea cuccullata

Ostracoblabe implexa Widespread





MUSSELS



Disease/parasite/pest Geographical distribution

Mytilus edulis

Marteilia refringens Atlantic Europe from southern

England to Portugal

Mytilus galloprovincialis

Marteilia refringens Atlantic Europe from southern

England to Portugal

Mytilus californianus

Haplosporidium tumefacientis California, U.S.A.



CLAMS AND COCKLES



Disease/parasite/pest Geographical distribution

Siliqua patula

Nuclear inclusion X Oregon and Washington States,

U.S.A.

Ruditapes decussatus

Brown ring disease France, Spain, Italy, Ireland

Haplosporidium tapetis France, Spain, Portugal



Perkinsus atlanticus Portugal, Spain,

Mediterranean Sea

Ruditapes philippinarum

Brown ring disease France, Spain, Italy, Ireland

Haplosporidium tapetis France, Spain, Portugal



Perkinsus atlanticus Portugal, Spain,

Mediterranean Sea

Mercenaria mercenaria

Sirolpidium East coast, U.S.A.



zoophthorum

Quahaug parasite X (QPX) Gulf of St Lawrence

Perkinsus marinus East coast, U.S.A., experimental

Dermo infection







C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

101





Venerupis aurea

Perkinsus atlanticus Portugal, Spain,

Mediterranean Sea

Mya arenaria

Perkinsus marinus East coast, U.S.A., experimental

Dermo infection

Cerastoderma edule

Marteilia refringens Atlantic Europe from southern

England to Portugal



SCALLOPS



Disease/parasite/pest Geographical distribution

Placopecten

magellanicus

Rickettsiosis Northeast coast of the U.S.

Bacterial abscess disease (BAD) Northeastern U.S.A., and Atlantic

Canada

Pecten maximus

Rickettsiosis France, Scotland, Sweden

Argopecten irradians

Sirolpidium East coast, U.S.A.



zoophthorum

Unidentified organism, formerly Atlantic Canada, northeast U.S.A.

Perkinsus karlssoni

Argopecten gibbus

Marteilia sp. Florida

Patinopecten yessoensis

Perkinsus qugwadi British Columbia, Canada, possibly

Japan and Russia









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

102





Appendix 2. A summary of the serious, or potentially serious, diseases,

parasites and pests, in the countries from which Australia imports, or may

in future support, bivalve molluscs. * current imports.

ARGENTINA*



Host species Disease/parasite/pest Tissue

Ostrea puelchana Bonamia ostreae Systemic



CANADA* - East coast



Host species Disease/parasite/pest Tissue

Crassostrea virginica Malpeque disease Systemic

Ostrea edulis Ostracoblabe implexa Shell

Mercenaria mercenaria Quahaug parasite X Systemic

Argopecten irradians Formerly Perkinsus Gut epithelia, connective

karlssonii tissue

Placopecten magellanicus Bacterial abscess disease Adductor muscle



CANADA* - West coast



Host species Disease/parasite/pest Tissue

Crassostrea gigas Mikrocytos roughleyi Systemic

Crassostrea gigas Nocardiosis Mantle, gill, adductor muscle, heart

Crassostrea gigas Ostracoblabe implexa Shell

Patinopecten yessoensis Perkinsus qugwadi, SPX Systemic

Patinopecten yessoensis Scallop protozoan G Connective tissue



CHILE



Host species Disease/parasite/pest Tissue

Tiostrea chilensis Bonamia ostreae Systemic



FRANCE



Host species Disease/parasite/pest Tissue

Crassostrea gigas Haplosporidium nelsoni Systemic

Crassostrea gigas Marteilia refringens Digestive tract epithelia

Crassostrea gigas Gill necrosis virus Gills

Crassostrea gigas Haemocytic infection virus Haemocytes

Ostrea edulis Marteilia refringens Digestive tract epithelia

Ostrea edulis Bonamia ostreae Systemic

Ostrea edulis Minchinia armoricana Systemic, connective tissue

Mytilus galloprovincialis Marteilia refringens Digestive tract epithelia









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

103





Cerastoderma edule Marteilia refringens Digestive tract epithelia

Ruditapes decussatus Brown ring disease Surface, mantle

Ruditapes philippinarum Brown ring disease Surface, mantle

Ruditapes decussatus Haplosporidium tapetis Digestive tract epithelia,

connective tissue

Ruditapes philippinarum Haplosporidium tapetis Digestive tract epithelia,

connective tissue

Ruditapes decussatus Perkinsus atlanticus Systemic, connective tissue

Ruditapes philippinarum Perkinsus atlanticus Systemic, connective tissue

Pecten maximus Rickettsiosis Gill endothelial cells



IRELAND



Host species Disease/parasite/pest Tissue

Ostrea edulis Ostracoblabe implexa Shell

Ostrea edulis Bonamia ostreae Systemic

Ruditapes decussatus Brown ring disease Surface, mantle

Ruditapes philippinarum Brown ring disease Surface, mantle

Patinopecten yessoensis Scallop protozoan G Connective tissue



ITALY



Host species Disease/parasite/pest Tissue

Ostrea edulis Bonamia ostreae Systemic

Ruditapes decussatus Brown ring disease Surface, mantle

Ruditapes philippinarum Brown ring disease Surface, mantle



JAPAN*



Host species Disease/parasite/pest Tissue

Crassostrea gigas Haplosporidium nelsoni Systemic

Crassostrea gigas Nocardiosis Mantle, gill, adductor muscle, heart

Ruditapes philippinarum Perkinsus sp. Systemic, connective tissue

Chlamys farreri Pectenophilus ornatus Gill arch

Patinopecten yessoensis Perkinsus qugwadi, SPX Systemic

Patinopecten yessoensis Scallop protozoan G Connective tissue

Patinopecten yessoensis Pectenophilus ornatus Gill arch



NETHERLANDS



Host species Disease/parasite/pest Tissue

Ostrea edulis Bonamia ostreae Systemic

Ostrea edulis Minchinia armoricana Systemic, connective tissue









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

104





PORTUGAL



Host species Disease/parasite/pest Tissue

Ostrea edulis Bonamia ostreae Systemic

Crassostrea angulata Ostracoblabe implexa Shell

Crassostrea gigas Gill necrosis virus Gills

Crassostrea gigas Ostracoblabe implexa Shell

Crassostrea gigas Marteilia refringens Digestive tract epithelia

Mytilus galloprovincialis Marteilia refringens Digestive tract epithelia

Cerastoderma edule Marteilia refringens Digestive tract epithelia

Ruditapes decussatus Haplosporidium tapetis Digestive tract epithelia,

connective tissue

Ruditapes philippinarum Haplosporidium tapetis Digestive tract epithelia,

connective tissue

Ruditapes decussatus Perkinsus atlanticus Systemic, connective tissue

Ruditapes philippinarum Perkinsus atlanticus Systemic, connective tissue



SPAIN

Host species Disease/parasite/pest Tissue

Crassostrea gigas Marteilia refringens Digestive tract epithelia

Crassostrea gigas Gill necrosis virus Gills

Crassostrea gigas Haemocytic infection virus Haemocytes

Crassostrea gigas Extracellular rickettsiae Gill surfaces

Ostrea edulis Marteilia refringens Digestive tract epithelia

Ostrea edulis Bonamia ostreae Systemic

Mytilus galloprovincialis Marteilia refringens Digestive tract epithelia

Cerastoderma edule Marteilia refringens Digestive tract epithelia

Ruditapes decussatus Brown ring disease Surface, mantle

Ruditapes philippinarum Brown ring disease Surface, mantle

Ruditapes decussatus Haplosporidium tapetis Digestive tract epithelia,

connective tissue

Ruditapes philippinarum Haplosporidium tapetis Digestive tract epithelia,

connective tissue

Ruditapes decussatus Perkinsus atlanticus Systemic, connective tissue

Ruditapes philippinarum Perkinsus atlanticus Systemic, connective tissue





UNITED KINGDOM*

Host species Disease/parasite/pest Tissue

Crassostrea gigas Gill necrosis virus Gills

Crassostrea gigas Marteilia refringens Digestive tract epithelia

Ostrea edulis Bonamia ostreae Systemic

Ostrea edulis Marteilia refringens Digestive tract epithelia

Ostrea edulis Ostracoblabe implexa Shell

Mytilus edulis Marteilia refringens Digestive tract epithelia

Cerastoderma edule Marteilia refringens Digestive tract epithelia









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc

105







UNITED STATES OF AMERICA* – East coast



Host species Disease/parasite/pest Tissue

Crassostrea gigas Haplosporidium nelsoni Systemic

Crassostrea virginica Haplosporidium costale Systemic

Crassostrea virginica Haplosporidium nelsoni Systemic

Crassostrea virginica Juvenile oyster disease Mantle epithelial cells

Crassostrea virginica Perkinsus marinus Systemic

Crassostrea virginica Sirolpidium zoophthorum Systemic

Ostrea edulis Bonamia ostreae Systemic

Mercenaria mercenaria Quahaug parasite X Systemic

Mercenaria mercenaria Sirolpidium zoophthorum Systemic

Ruditapes decussatus Sirolpidium zoophthorum Systemic

Argopecten gibbus Marteilia sp . Digestive tract epithelia

Argopecten irradians Sirolpidium zoophthorum Systemic

Placopecten magellanicus Rickettsiosis Gills, epithelial surfaces

Placopecten magellanicus Bacterial abscess disease Adductor muscle



UNITED STATES OF AMERICA* – West coast



Host species Disease/parasite/pest Tissue

Ostrea edulis Bonamia ostreae Systemic

Crassostrea virginica Haplosporidium costale Systemic

Crassostrea gigas Haplosporidium nelsoni Systemic

Crassostrea gigas Nocardiosis Mantle, gill, adductor muscle, heart

Mytilus californianus Haplosporidium tumefacientis Digestive gland, kidney

Siliqua patula Nuclear inclusion X Branchial epithelial cells









C:\Docstoc\Working\pdf\51af78aa-2201-4f5c-a1ee-021886db926e.doc