Haplosporidium nelsoni (MSX) of Oysters

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• Category
• Common, generally accepted names of the organism or disease agent
• Scientific name or taxonomic affiliation
• Geographic distribution
• Host species
• Impact on the host
• Diagnostic techniques
• Methods of control
• References
• Citation information

Category

Category 2 (In Canada and of Regional Concern)

Common, generally accepted names of the organism or disease agent

MSX (Multinucleate Sphere Unknown 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). However, the taxonomy of haplosporidians needs a thorough revision (Hartikainen et al. 2003, Hine et al. 2009).

Geographic distribution

1. 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). In the autumn of 2002, an epizootic outbreak of H. nelsoni 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. Ulrich et al. (2007a) reported H. nelsoni from the Gulf of Mexico and Caribbean Sea from sites ranging from Florida to Venezuela in oysters with no associated pathology. Reported infections were detected only by polymerase chain reaction (PCR) amplification of the ribosomal rRNA gene complex (Ulrich et al. 2007a). However, Burreson (2008) indicated that although this report was interesting and worthy of additional research, the assay used by Ulrich et al. (2007a) was not validated for the Gulf of Mexico and infections were not confirmed by histology. Thus, infections by H. nelsoni in C. virginica in the Gulf of Mexico and in other species of oysters in the Caribbean Sea needs to be confirmed (Burreson 2008). A widespread survey of oysters (210 from 40 sites examined using PCR, 180 of these using tissue-section histology) from around the Gulf of Mexico and Puerto Rico did not detect H. nelsoni (Ford et al. 2011). The disease is restricted to salinities over 15 parts per thousand (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.
2. Reported from Crassostrea gigas in California and Washington, C. gigas cultured in France, Japan, a low prevalence observed in Korea, a relatively low prevalence reported on the coast of the East China Sea, the Bohai Sea and the north coast of the Yellow Sea, China and from C. gigas in Ireland (Chun 1972; Kern 1976a; Kang 1980; Renault et al. 2000; Kamaishi and Yoshinaga 2002; Miossec et al. 2009; Burreson 2005; Wang et al. 2010a, b; Lynch and Culloty 2011; Lynch et al. 2013). 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). More 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. Also, haplosporidian plasmodia and spores that resemble H. nelsoni were detected in one of 2184 C. gigas sampled from the Galicia coast of Spain between 2004 and 2009 (Iglesias et al. 2012). 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

1. Crassostrea virginica (similar haplosporidians found in other bivalve species world wide). Ulrich et al. (2007a) reported H. nelsoni in Crassostrea rhizophorae from Puerto Rico and in C. rhizophorae or Crassostrea gasar from Venezuela, in oysters with no associated pathology. However, these host species and locations need to be ratified because the PCR assay used by Ulrich et al. (2007a) was not validated for these species of oysters and infections were not confirmed by histology (Burreson 2008). Subsequent examination of C. rhizophorae from Puerto Rico did not detect H. nelsoni (Ford et al. 2011).
2. Crassostrea gigas and possibly the Olympia oysters, Ostrea conchaphila (=lurida). Lynch et al. (2013) detected plasmodia of H. nelsoni in the heart tissue of a single Ostrea edulis among the 1310 cultured O. edulis (2-4 years of age) sampled from various locations on the coast of Ireland.

Impact on the host

a) Mortalities can reach 90% to 95% of the C. virginica 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 (Kern 1976b, Andrews and Wood 1967, Ford and Haskin 1982, Burreson 2000, Powell et al. 2008). In adjacent areas where no unusual mortalities were detected, the meat condition index of oysters was found to decline significantly (Abbe and Sanders 1988). Mortalities may commence early in the spring (infected animals unable to recover from the metabolic demands of over-wintering) with maximum mortality in late July and early August and infection of new oyster hosts occurs primarily in that season (Couch and Rosenfield 1968, Burreson 2005, Burreson and Stokes 2006). The widespread distribution and high virulence of H. nelsoni have greatly reduced traditional on-bottom culture and have limited the development of off-bottom oyster aquaculture in high salinity areas (Burreson 2000). Although the annual distribution and prevalence of H. nelsoni vary with salinity, its general prevalence and intensity have not decreased since the original epizootics (Burreson 2000). However, after nearly 50 years of infection pressure, there is some evidence that native oysters in Delaware Bay and lower Chesapeake Bay are developing a natural resistance to H. nelsoni (Burreson 2005). In some areas of Delaware's Inland Bays where oysters have historically been under the greatest threat from H. nelsoni, Perkinsus marinus may pose a greater threat to the recovery of oyster aquaculture (Ulrich et al. 2007b).

Disease activity increases during drought periods in association with high salinities (Andrews 1968). In mesohaline areas (about 15-25 ppt), infective pressure remained high and relatively stable despite scarcity of oysters (Andrews and Frierman 1974). 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). In South Carolina, the highest prevalence (28 to 42%) and intensity of H. nelsoni occurred during the fall with no apparent associated mortalities (Bobo et al. 1996). Infected oysters held in a ‘disease-free' area for up to 4 years displayed the same seasonal cycle with chronic infections becoming localized, relapsing into general infections, and then becoming localized again in a sequence that was probably controlled by temperature. However, the persistent disease and resulting poor condition of the oysters suggested that chronically infected individuals would have lowered resistance to additional stress (Ford 1985a). Nevertheless, C. virginica in some areas have developed resistance to mortality, which involves an ability to restrict infections and tolerate parasitism for prolonged periods (Ford and Figueras 1988a). Crassostrea virginica in Deleware Bay have become very resistant to H. nelsoni infection development and rarely exhibit histologically detectable lesions (Ford et al. 2009). Also, resistant populations of C. virginica were believed to make a substantial contribution to oyster reproduction in some areas of Chesapeake Bay (Carnegie and Burreson 2011a, b, c).

The complete life cycle of H. nelsoni is unknown (Haskin and Andrews 1988, Burreson and Ford 2004, Burreson 2005). Sporulation of H. nelsoni is sporadic in adult C. virginica but prevalent in juvenile oysters (Barber et al. 1991c, Burreson 1994, Barber and Ford 1995). 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. Spores are not directly infective to oysters and 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 (Powell et al. 1999, Burreson and Ford 2004, Burreson 2005, Burreson and Stokes 2006). Sunila et al. (2000a) were able to infect hatchery-raised, MSX-free juvenile C. virginica (57% prevalence 11 weeks after the beginning of the exposure and a mortality of 80% after 16 weeks) placed in up-weller tanks supplied with water filtered through a screen with 1 mm² openings. The water originated from an area overlaying naturally infected oysters beds (MSX prevalence 17 to 57%) thereby demonstrating transmission of H. nelsoni via water-borne infectious agents capable of passing through a 1 mm filter (Sunila et al. 2000a). 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 (Burreson et al. 2000, Burreson 2005). Temperature and salinity are important in regulating H. nelsoni infections in C. virginica (Burreson 2005, Burreson and Stokes 2006). Both parasite and oyster are inactive at temperatures below 5 °C, the parasite proliferates between 5-20 °C and infections are acquired and oysters killed above 20 °C. A salinity of 15 ppt is required for infection and 20 ppt is required for rapid parasite proliferation. Salinities of 10 ppt or below results in expulsion of the parasite from the oyster within 10 days at temperatures above 20 °C (Burreson 2005). 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 ppt), 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. Outbreaks of H. nelsoni in the NE United States were 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).

Laboratory experiments and field observations indicated that H. nelsoni does not thrive at reduced salinities (Sprague et al. 1969, Andrews 1983). Reduced prevalence of H. nelsoni in C. virginica held at a low salinity is probably due to a physiological inability of the parasite to tolerate reduced salinity rather than to enhance effectiveness of host defense mechanisms (Ford and Haskin 1988a). Haemolymph enzyme (aspartate aminotransferase, alanine aminotransferase, and phosphohexose isomerase) activities increase significantly during the gill lesion stage of MSX disease but were not significantly elevated during the general infection stage of the disease (Douglass and Haskin 1976). However, the reverse situation was reported for total haemolymph protein levels. When infections were confined to the gill, as was the case with mortality-resistant oysters, there was no depression of protein levels but concentrations fell sharply and in approximate proportion to disease intensity in oysters with systemic H. nelsoni infections. Oysters with heavy parasitism had about one-third the protein levels of uninfected individuals (Ford 1986a). The clearance rate (difference in particle concentration entering and leaving an experimental chamber containing the test oysters) and condition index of C. virginica with systemic infections of H. nelsoni were significantly reduced compared to oysters without the parasite (Newell 1985). Also, the mean condition index for C. virginica with both the shell-boring polychaete Polydora sp. and H. nelsoni was 13% lower than that for oysters with neither; oysters with only one of these two invaders had intermediate values (Wargo and Ford 1993). In addition to a lower condition index, C. virginica parasitized by H. nelsoni had less glycogen (% dry weight) and a reduced relative reproductive effort compared to non-infected oysters (Barber et al. 1988a). Other tissue biochemical components (including lipid and ash content) also generally declined with increased H. nelsoni infection intensity and duration (Barber et al. 1988b and c). Gametogenesis was significantly depressed in male and female C. virginica with systemic infections of H. nelsoni but not in oysters with infections that were confined to the gills (Ford and Figueras 1988a and b). If temperature-associated infection remission subsequently occurred, many infected oysters recovered, developed mature gonads, and spawned before new or recurrent infections proliferated in the fall (Ford and Figueras 1988a). Agglutinins tested in the serum of C. virginica played no role in the defense against H. nelsoni (Chintala et al. 1994). However, experimental repeated haemolymph sampling of experimental C. virginica was associated with increased parasitism by H. nelsoni (Ford 1986b).

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, Burreson 2005) and prevalence of infection is typically low (less than 5%) (Kamaishi and Yoshinaga 2002, Wang et al. 2010a).

Diagnostic techniques

The detection and confirmation of infections with Haplosporidium nelsoni should encompass a range of techniques including heart smear screening (histocytology), histology, standard polymerase chain reaction (PCR), direct sequencing of PCR products and in situ hybridisation (ISH) with specific DNA probes designed to detect H. nelsoni in order to substantiate the diagnosis (Lynch et al. 2013).

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 and infected oysters can appear healthy (Burreson and Stokes 2006).

Squash Preparations

In digestive gland tissue squashes of oysters (usually juveniles) with spore development, the operculate spores measure 7.5 × 5.4 μm (unfixed).

Wet Mounts

A process known as "panning" was used to obtain plasmodia of H. nelsoni from oyster haemolymph. Panning consists of layering haemolymph in a petri dish, allowing haemocytes to settle and adhere, washing with isosmotic sea water, then removing supernatant fluid containing parasites that do not attach (Ford et al. 1990b). This process was used by Ford et al. (1993) to investigate the in vitro interactions between bivalve haemocytes and H. nelsoni. Haemocytes from C. virginica and C. gigas failed to phagocytise living plasmodia whereas haemocytes from the ribbed mussel, Geukensia demissa, a species that co-occurs with C. virginica in some habitats but is not known to become infected by H. nelsoni, rapidly phagocytised the plasmodia (Ford et al. 1993).

Histocytology: Oyster blood cell suspensions can be screened for plasmodial stages because the plasmodial stages spread throughout the tissues via the haemolymph. Moderate to heavy infections were diagnosed with 93% to 100% accuracy using fresh haemolymph preparations (Kanaley and Ford 1988). A durable record can be prepared by allowing the haemocytes to settle onto glass slides followed by drying the slides and staining them with Wright, Wright-Giemsa or equivalent stain (e.g. Hemacolor, Merck; Diff Quick, Baxter).

Histology

Histological examination using light microscopy of paraffin embedded tissue sections (5 to 6 µm thick) and coloured with hematoxylin and eosin (H&E) stain is the standard diagnostic technique for H. nelsoni (Burreson 2005, Burreson and Stokes 2006). Multinucleate plasmodia (4-100 μm in diameter depending on the number and size of the nuclei) in which nuclei have a peripheral nucleolus occur extracellularly throughout the connective tissue and epithelium (Perkins 1968, Burreson 2005, Burreson and Stokes 2006). In C. virginica, plasmodia are detectable from mid-May to October. Plasmodial infections are apparently initiated in the gill (Sunila et al. 2000a). Plasmodia multiply along the basal lamina of the epithelium and eventually break through into the circulatory system. Infection is often associated with haemocytosis. In older infections in C. gigas, an intense infiltration of haemocytes surrounds the plasmodial foci and necrotic areas of host tissue. The production of spores (sporulation) is rare in adult oysters but has been observed as high as 40 % in young spat of C. virginica (Barber et al. 1991c, Burreson 1994, Burreson 2005). Sporogonic stages (sporocysts 20-50 μm in diameter) and acid fast spores (4-6 μm × 5-8 μm with an overhanging cap at one end) are restricted to the epithelium of the digestive gland in C. virginica (Couch et al. 1966, Farley 1967, Graczyk et al. 1998) 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. 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.

Electron microscopy

Nuclei within the plasmodia are spherical (1.5-3 μm in diameter) with a peripheral endosome or are elongated (up to 7.5 μm long). During sporulation, plasmodia develop into sporocysts with spore walls forming around each nucleus (Perkins 1968). The fine structure of MSX plasmodia in the gills was extremely diverse with the cytoplasm ranging from electron-light to dense depending on the number of free ribosomes present (Scro and Ford 1990). The cytoplasm of all plasmodia contained round dense membrane-limited bodies called haplosporosomes (about 0.18 μm in diameter) that often contained an inner electron-lucent configuration. Electron-light plasmodia usually had fewer haplosporosomes (Scro and Ford 1990). Another structure found in the cytoplasm and called the concentric body (about 0.3 μm in diameter) was round and had a dense core that was surrounded by two electron-dense concentric rings. The ring closest to the core appeared to be connected to the core by fine radiating fibers arranged in a spoke-like fashion (Scro and Ford 1990).

Immunological Assays

Immunogold silver staining (application of primary antibody to haplosporidian spores produced in rabbits followed by addition of affinity purified goat anti-rabbit IgG coated on 5-nm colloidal gold particles and enhanced by precipitation of metallic silver to produce a positive reaction which appeared as a dark brown to black signal at the site of each antigen-antibody complex under light microscopy) of histological sections was used to differentiate between the spores of a haplosporidan infecting shipworms (Teredo navalis) and H. nelsoni (McGovern and Burreson 1987).

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, 2001; Stokes et al. 1995; Day et al. 2000; Renault et al. 2000; Burreson 2000; Burreson and Stokes 2006). Diagnosis using polymerase chain reaction (PCR) was found to be more sensitive than histology and allowed infections to be detected much earlier in the natural infection cycle (Burreson 2000). Ko et al. (1999) developed a rapid method, utilizing both polymerase chain reaction (PCR based on specific ribosomal RNA genes) and enzyme-linked immunosorbent assay (ELISA) for detecting H. nelsoni in oysters. Positive signals were observed in infected oysters using this PCR-ELISA technique and this method may be applicable to early detection of infection (Ko et al. 1999). Wang et al. (2010b) reported that reamplification of the unpurified PCR product, obtained with the MSX-A' and MSX-B primers described by Renault et al. (2000), when used as the template in a second PCR reaction increased the sensitivity of the PCR assay. 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). However, Ford et al. (2009) suggested using suspension-feeding bivalves as particle collectors in conjunction with PCR detection to investigate the environmental distribution of H. nelsoni. Results from this approach indicated that H. nelsoni persists throughout Delaware Bay, although it is rarely detected using standard histology in the disease resistant C. virginica populations (Ford et al. 2009).

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 (ISH) 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, Burreson and Stokes 2006). However, adequate molecular information is needed for the development of rigorous molecular diagnostic assays, these assays must be thoroughly tested for sensitivity and specificity, and ultimately the protocols need to be validated against established diagnostic procedures such as histology, which allows visualization of the parasite in host tissues, prior to full implementation (Reece and Burreson 2004, Burreson 2008). For example, the extreme sensitivity of the PCR technique may allow amplification of DNA from non-viable or non-pathogenic organisms. Therefore, a PCR-positive result does not necessarily mean that viable, pathogenic organisms are present or that mortality will eventually occur. Nonetheless, molecular diagnostic techniques hold great promise for early diagnosis of disease outbreaks in aquaculture and natural populations (Burreson 2000).

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). Anecdotal evidence indicates that H. nelsoni was introduced into the marine Bras d'Or Lakes in Nova Scotia, Canada in ballast water (Burreson 2005). Eradication is not possible and there is no known chemical treatment for H. nelsoni infections (Burreson 2005). Early diagnosis of H. nelsoni infection in C. virginica is an essential management tool (Ford and Haskin 1988b). The earlier infections can be detected, the more time is available for oyster growers to decide whether to harvest to limit mortality, or whether to move oysters to low salinity to eliminate the parasite (Ford 1985b). Infections of H. nelsoni acquired in late summer or fall remain nonlethal until spring; early detection would also allow harvest to avoid losses from these late season infections (Burreson 2000).

Source, history, size and age of oysters, and timing of exposure are important variables that affect MSX disease activity in C. virginica (Andrews and Frierman 1974). Salinity and temperature are important determinants of the severity of MSX disease (Andrews and Frierman 1974, Haskin and Ford 1982, Andrews 1983, Ford 1985b, Haskin and Ford 1989). 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 10 °C (Ford and Haskin 1988a, b). For example, Barber et al. (1991b) “purged” experimental oysters of MSX by suspended them for several weeks in an area where salinity was less than 5 ppt. Water depth was also found to affect progression of disease caused by H. nelsoni in C. virginica. Oysters at residence depth less than 45 cm compared to those at depths greater than 90 cm had decreased susceptibility to the parasite (Volety et al. 2000). Thus, Volety et al. (2000) suggested that piling of shells and construction of artificial reefs was a better strategy for rejuvenating oyster stocks than spreading thin layers of oyster shells on the bottom in estuaries and coastal areas.

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 (Kern 1976b). The development of resistant strains of C. virginica, which may become available for future culture in endemic areas, was undertaken (Haskin and Ford 1979; Ford and Haskin 1987, 1988b; Ford et al. 1990a; Matthiessen et al. 1990; Vrijenhoek et al. 1990; Sunila et al. 1999a). Barber et al. (1991a) suggested that resistance in the selected strain may be the result of a physiological response that inhibits the development of H. nelsoni and a basic metabolic adjustment to parasitism. 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 were enzootic (lower York River, Virginia, USA) (Ragone Calvo et al. 2003). Carnegie and Burreson (2011a) determined that populations of resistant oysters in disease-enzootic waters of Chesapeake Bay made a substantial contribution to reproduction and suggested that reefs in mesohaline-polyhaline salinities, and not only those in disease-free lower mesohaline areas, should be the focus of conservation and restoration efforts.

Efforts to mitigate the impact of H. nelsoni involved 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 (Matthiessen and Davis 1992, Dégremont et al. 2012). Note: both selected disease resistant oysters and triploids may be carriers of infection. Use of brood stock that are free of infection is an important management step. However, specific-pathogen-free (SPF) C. virginica will not have improved chances of survival when deployed into enzootic areas experiencing high salinity (e.g., during multiyear drought conditions) but are likely to survive and grow to market size if subject to reduced salinities for a relatively short duration such as just one year of drought conditions (Albright et al. 2007). 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 μm and then ultraviolet light (30,000 μW 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).

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Citation Information

Bower, S.M. (2014): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Haplosporidium nelsoni (MSX) of Oysters.

Date last revised: December 2014

Comments to Susan Bower