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Archived - Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish

Haplosporidium nelsoni (MSX) of Oysters

Category | Common Name | Scientific Name | Distribution | Host Species
Impact on Host | Diagnostic Technique | Methods of Control | References | Citation


Category

Category 2 (In Canada and of Regional Concern)

Common, generally accepted names of the organism or disease agent

MSX (Multinucleate Sphere X) disease, Haplosporidiosis, Delaware Bay disease, Haplosporidiosis of Pacific oysters.

Scientific name or taxonomic affiliation

Haplosporidium (=Minchinia) nelsoni, within the family Haplosporidiidae, order Haplosporida, class Haplosporea of the phylum Haplosporidia (Perkins 1990, 2000; Siddall et al. 1995; Flores et al. 1996; Reece et al. 2004; Burreson and Ford 2004).

Geographic distribution

a) Florida, USA north to Nova Scotia, Canada. Enzootic areas occur in Delaware Bay with occasional epizootics in Chesapeake Bay; Long Island Sound, Connecticut; Bayville, New York; Cape Cod and Cotuit Bay, Massachusetts; and Piscataqua River Estuary in Maine/New Hampshire. The Maryland, USA, Department of Natural Resources, Fisheries Service monitors the prevalence and distribution of H. nelsoni in Chesapeake Bay (Tarnowiski 2003, 2005). A recent (autumn of 2002) epizootic has occurred in the Bras d'Or Lakes, part of Cape Breton, Nova Scotia, Canada (Stephenson et al. 2003). However, the parasite has not yet been detected in oyster stocks between the southern end of Maine, USA and Bras d'Or Lakes. The disease is restricted to salinities over 15 ppt (H. nelsoni cannot survive below 10 ppt), rapid and high mortalities occur at 18-20 ppt (parasite proliferation is greatest above 20 ppt). There is some evidence that water temperatures exceeding 20 C may cause the disease to disappear.

b) Reported from Crassostrea gigas in California and Washington, C. gigas cultured in France, and a low prevalence observed in Korea  (Chun 1972, Kern 1976, Kang 1980, Renault et al. 2000). In 1989-1993, up to 10% of the C. gigas seed from Japan (Matsushima and Watanoha bays) being screened for importation into California were infected (Friedman et al. 1991, Friedman 1996). Recently (June 2007), H. nelsoni was detected with no associated mortalities in a few C. gigas from a grow-out facility in British Columbia, Canada. Haplosporidian plasmodia have also been reported in Olympia oysters (Ostrea conchaphila) from Oregon, USA that were imported from California (Mix and Sprague 1974).

Host species

a) Crassostrea virginica (similar haplosporidians found in other bivalve species world wide).

b) Crassostrea gigas and possibly the Olympia oysters, Ostrea conchaphila (=lurida).

Impact on the host

a) Mortalities can reach 90% to 95% of the oysters in a cohort within 2 to 3 years of being out-planted. When the disease first appeared in the late 1950s and early 1960s, mortalities of adult C. virginica approached 100% of the standing stock during a 3 year period in the high salinity areas of Chesapeake and Delaware bays (Andrews and Wood 1967, Ford and Haskin 1982). Mortalities may commence early in the spring (infected animals unable to recover from the metabolic demands of over-wintering) and infection of new oyster hosts occurs primarily in that season (Couch and Rosenfield 1968). Disease activity increases during drought periods in association with high salinities (Andrews 1968). Mortalities from new or recurrent infections occur throughout the summer and peak in August-September and are not associated with oyster population densities (Andrews 1968, 1984). The complete life cycle of H. nelsoni is unknown (Haskin and Andrews 1988). Sporulation of H. nelsoni is sporadic in adult C. virginica but prevalent in juvenile oysters (Barber et al. 1991b, Burreson 1994). When present, it occurs in summer and causes a gradual disruption of digestive tubule epithelia. Sporulation in juvenile C. virginica was associated with mortality rates of at least 30% in infected spat. Transmission directly between oysters has not been accomplished; requirement for an intermediate host to complete the life cycle is suspected and supported by model simulations (Burreson and Ford 2004). DNA analysis indicates that H. nelsoni was introduced to the east coast of the United States from California or from Asia with documented plantings of C. gigas. Modeling studies by Hofmann et al. (2001) indicated that when cold winter temperatures (less than 3 C) are followed by a year of low salinity (less than 15 parts per thousand), the prevalence and intensity of MSX disease are greatly reduced and winter temperatures consistently lower than 3 C limit the long-term development of the disease. However, the disease will occur when average environmental conditions return. The recent H. nelsoni outbreaks in the NE United States are related to increased winter temperatures (Burreson and Ford 2004). Also, the timing of maximum food supply for the oyster and stage in the parasites life cycle within the oyster is important in determining whether or not the parasite undergoes sporulation or density-independent growth of the plasmodia (Hofmann et al. 2001).

Reduced prevalence of H. nelsoni in low salinity is probably due to a physiological inability of the parasite to tolerate reduced salinity rather than to enhanced effectiveness of host defense mechanisms (Ford and Haskin 1988b). Agglutinins tested in the serum of C. virginica played no role in the defense against H. nelsoni (Chintala et al. 1994).

b) Effects on C. gigas have not been described but some authors speculate that it may be pathogenic, especially for juvenile oysters. However, haplosporidiosis has not been associated with mortality of C. gigas (Elston 1999) and prevalence of infection is typically low (less than 4%) (Kamaishi and Yoshinaga 2002).

Diagnostic techniques

Gross observations: Infected juvenile oysters may have pale digestive glands, appear emaciated and show no new shell growth. In adult oysters, mantle recession has been reported from heavily infected C. virginica, accompanied by extensive fouling along the inside peripheral left valve. Raised yellow-brown conchiolin deposits on internal valve surfaces of chronically infected oysters have been reported but these are not consistent and frequently are not observed. Affected oysters are typically thin and watery with pale digestive diverticula. The clinical signs are not unique to Haplosporidian-caused diseases (Farley 1968). In acute infections, the disease may progress rapidly resulting in no clinical signs prior to death.

Squash Preparations: In digestive gland tissue squashes of oysters (usually juveniles) with spore development, the operculate spores measure 7.5 5.4 m (unfixed).

Histocytology: Examine blood cell suspensions for plasmodial stages. Allow haemocytes to settle onto glass slides and stain with Wright, Wright-Giemsa or equivalent stain (e.g. Hemacolor, Merck; Diff Quick, Baxter). Plasmodial stages spread throughout the tissues via the haemolymph.

Histology: Multinucleate plasmodia (4-50 m in diameter depending on the number and size of the nuclei) occur extracellularly throughout the connective tissue (Perkins 1968). In C. virginica, plasmodia are detectable from mid-May to October. Plasmodial infections are apparently initiated in the gill. Plasmodia multiply along the basal lamina of the epithelium and eventually break through into the circulatory system. Infection is often associated with haemocytosis. Sporogonic stages (sporocysts 20-50 m in diameter) and acid fast spores (4-6 m 5-8 m with an overhanging cap at one end) are restricted to the epithelium of the digestive gland in C. virginica but may also occur (infrequently) in other tissues of C. gigas. The sporocysts can rupture the digestive epithelial cells thereby releasing developing and mature spores into the lumen of the digestive gland. In older infections in C. gigas, an intense infiltration of haemocytes surrounds the plasmodial foci and necrotic areas of host tissue. Spores are rarely observed in adult oysters but are more prevalent in juveniles (Barber et al. 1991b, Burreson 1994). Unlike Haplosporidium costale (SSO), there is no spore formation in the connective tissue of C. virginica, sporulation is asynchronous and the spores of H. nelsoni are larger than those of H. costale. However, in the absence of spores, distinguishing between H. costale and H. nelsoni is nearly impossible using traditional histological examination.

Digestive Gland Figure 1. Histological section through the digestive gland of Crassostrea virginica from Virginia, USA, infected with Haplosporidian nelsoni. Multinucleate plasmodia (P) occur in the connective tissue and haemal spaces as well as within the digestive tubule epithelium while sporulation and release of mature spores (S) are limited to the digestive tubule epithelium. Haematoxylin and eosin stain.

Electron microscopy: Nuclei within the plasmodia are spherical (1.5-3 m in diameter) with a peripheral endosome or are elongated (up to 7.5 m long). During sporulation, plasmodia develop into sporocysts with spore walls forming around each nucleus (Perkins 1968).

DNA Probes: The small subunit ribosomal RNA gene was sequenced (Fong et al. 1993) and sensitive and specific molecular probes were identified (Stokes and Burreson 1995, Stokes et al. 1995, Day et al. 2000). Polymerase chain reaction (PCR) technology was used as a tool in an unsuccessful attempt to identify the life cycle of H. nelsoni (Burreson et al. 1996, Stokes et al. 1999). Multiplex PCR (simultaneous testing of two or more pathogens in a single test reaction) was developed for H. nelsoni, H. costale (SSO), and Perkinsus marinus (Penna et al. 1999, 2001; Russell et al. 2000, 2004). DNA sequence equivalency (tested by in situ hybridization and amplification of genomic DNA of parasitized C. gigas with primers specific for a 565 bp fragment of the small subunit rRNA of H. nelsoni from C. virginica) is conclusive evidence that the haplosporidian in C. gigas is H. nelsoni.

PCR and in situ hybridisation assays developed to specifically identify H. costale in conjunction with molecular tools previously developed for H. nelsoni have overcome the limitations of histological examination, which could not be used to differentiate between the plasmodial stages of these two parasites that overlap in geographic distribution (Stokes and Burreson 2001). However, adequate molecular information is needed for the development of rigorous molecular diagnostic assays and the protocols need to be validated against established diagnostic procedures such as histology prior to full implementation (Reece and Burreson 2004)

Methods of control

Evidence indicates that the spread of H. nelsoni to new areas was caused by the introduction of diseased oysters from areas where the parasite was established (Krantz et al. 1972). Thus, do not introduce oysters or other marine organisms (that may serve as an intermediate hosts) from MSX-endemic areas (historical or current). Eradication is not possible. Salinity and temperature are important determinants of the severity of MSX disease (Haskin and Ford 1982, Ford 1985). The effect on infected populations can be reduced by holding infected oysters in cold, low salinity waters (less than 15 ppt) for as long as possible and reducing the amount of grow-out time in higher salinity, warm water. Infections can be eliminated in two weeks by exposing oysters to mean salinities of 10 ppt or less and temperatures above 20 C. Results of epizootiological investigations by Farley (1975) indicated that C. virginica from an enzootic area were more resistant to disease than oysters introduced from another area. Endeavors to develop resistant strains of C. virginica, which may become available for future culture in endemic areas, are in progress (Haskin and Ford 1979, Ford and Haskin 1987,1988a; Ford et al. 1990; Matthiessen et al. 1990; Sunila et al. 1999a). Field transmission studies using specific DNA primers and PCR technology suggest that native oysters at some locations in Delaware Bay have become resistant to the development of H. nelsoni infection (Ford et al. 2000a). Also, dual resistance to H. nelsoni and Perkinsus marinus was achieved through four generations of artificial selection at a location where both diseases are enzootic (lower York River, Virginia, USA) (Ragone Calvo et al. 2003).

Efforts to mitigate the impact of H. nelsoni involve the evaluation of disease resistant non-native oysters (Burreson and Ford 2004). Triploid C. virginica also seem more resistant to the disease and mortalities caused by MSX. Note: both selected disease resistant oysters and triploids may be carriers of infection. Use of broodstock that are free of infection is an important management step. Ford et al. (2000b, 2001) demonstrated that juvenile oysters from nursery systems (seed) that use raw water pumped from an enzootic area are highly likely to be infected although infections may be very light in intensity and low in prevalence. Treating raw water, by filtration to 1 m and then ultraviolet light (30,000 W s-1 cm-2 UV irradiation), will help protect hatchery produced seed from infection (Ford et al. 2001).

A mathematical model that simulates the host-parasite-environment interactions was developed to provide a quantitative framework for guiding future laboratory and field studies as well as management efforts (Ford et al. 1999; Hofmann et al. 1999; Paraso et al. 1999; Powell et al. 1998, 1999). Attempts to manage the October 2002 MSX outbreak in Atlantic Canada included analysis of historic oceanographic data, targeted surveillance and consideration of industry activities. The information was used to develop a zonation approach to curtailing the spread of the disease (Stephenson and Petrie 2005).

 


References

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Citation Information

Bower, S.M. (2007): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Haplosporidium nelsoni (MSX) of Oysters.

Date last revised:  October 2007
Comments to Susan Bower