Haplosporidium costale (SSO) of Oysters

Category

Category 2 (In Canada and of Regional Concern)

Common, generally accepted names of the organism or disease agent

Seaside disease, Seaside organism, SSO, High salinity disease.

Scientific name or taxonomic affiliation

Haplosporidium costale, (=Minchinia costalis) currently within the family Haplosporidiidae, order Haplosporida, class Haplosporea of the phylum Haplosporidia (Burreson and Ford 2004). However, the taxonomy of haplosporidians needs a thorough revision (Hine et al. 2009).

Geographic distribution

  1. Long Island Sound, New York to Cape Charles, Virginia, USA, in high salinity waters (more than 25 parts per thousand (ppt)). Haplosporidians morphologically identical to H. costale were detected in C. viriginca from Tomales Bay, California, USA that had been transplanted from the vicinity of New Haven, Connecticut up to 18 months earlier (Katkansky and Warner 1970). In 2002-2003, plasmodia, but not spores, of H. costale were detected in low prevalence and intensity of infection in C. virginica from several locations in the Southern Gulf of St. Lawrence, Atlantic coast of Nova Scotia and Bras d'Or Lakes in Cape Breton, Nova Scotia, Canada.
  2. Reported in a low prevalance of Crassostrea gigas from the Bohai Sea and Yellow Sea coastal areas of China by Wang et al. (2010). Infections were detected and identified using molecular tools which included in situ hybridization of tissue sections, specific polymerase chain reaction (PCR) on genomic DNA extracted from the digestive gland tissue followed by DNA sequencing of the PCR products (Wang et al. 2010).

Host species

  1. Crassostrea virginica.
  2. Crassostrea gigas.

Impact on the host

  1. Can cause a sharp (4-6 weeks) seasonal mortality in May and June in Virginia and Maryland which coincides with sporulation of H. costale in C. virginica (Couch and Rosenfield 1968, Andrews and Castagna 1978, Andrews 1984). Prevalence of H. costale and, thus, oyster mortality is usually less than 20%, but can reach 40% in some years along the Virginia coast. Oyster mortality from SSO disease seems to be much lower in the northern parts of its distribution (Burreson and Stokes 2006). Generally, infections of H. costale are acquired in early summer but do not become patent until the following March. Plasmodia multiply rapidly with synchronous sporulation in late May/early June causing host death (Burreson and Stokes 2006). In Long Island Sound, rare cases (0.08%) of sporulation were detected in October to December (Sunila et al. 2002). The detection and confirmation of plasmodia of H. costale using DNA probes in October in Virginia and Long Island Sound (Stokes and Burreson 2001, Sunila et al. 2002) was unprecedented and challenged historical criteria for the seasonality and epizootiology of this parasite. It was unclear whether this apparent change in seasonality was real or simply the result of improved diagnostic sensitivity (Burreson and Ford 2004). Haplosporidium costale is restricted to high salinity coastal bays and it does not occur where salinity is less than about 25 ppt. Also, the life cycle of H. costale is unknown (Burreson and Stokes 2006). In situ hybridization assays indicate that co-infections with Haplosporidium nelsoni (MSX) occur.
  2. Effects of H. costale on C. gigas have not been reported.

Diagnostic techniques

Gross observations: There are no reliable gross clinical signs of infection. Usually SSO disease is acute and mortality so rapid that infected oysters can appear healthy (Burreson and Stokes 2006).

Histology: Histological examination using light microscopy of paraffin embedded tissue sections (5 to 6 µm thick) coloured with hematoxylin and eosin (H&E) stain is the standard diagnostic technique for H. costale (Burreson and Stokes 2006). Multinucleate plasmodia (typically less than 10 µm in diameter containing nuclei about 1.6 µm in diameter) and uninucleate spores containing bright red sporoplasm (when stained by a modified Ziehl-Neelsen carbol fuschin techniqu)e occur extracellularly in the connective tissue . Spores of H. costale produced a positive acid fast stain (AFS) reaction when histological sections were stained with acid-fast stain (Graczyk et al. 1998). Synchronous sporulation of the parasite occurs throughout the connective tissue of the digestive gland, mantle and gonads, but not in the epithelia of the digestive tubules. Plasmodia are only easily detectable in March through June and infection is often associated with intense haemocyte infiltration, but phagocytosis of plasmodia is rare (Burreson and Stokes 2006). Spores (about 3 to 4 µm in diameter) are usually found in moribund oysters (gapers). All stages of H. costale are approximately half as large as Haplosporidium nelsoni (MSX) and spores do not occur in the digestive tubule epithelia as for H. nelsoni. However, in the absence of spores, distinguishing between H. costale and H. nelsoni is nearly impossible using traditional histological examination. Haplosporidium costale cannot readily be detected between July and March.

Electron microscopy: During sporulation, plasmodia develop into sporocysts with spore walls forming around the nuclei. Spores (2.6 µm by 3.1 µm) are operculate and have a cap with an overhanging lid (Perkins 1969).

Immunological Assay: Polyclonal antibodies produced in rabbits against spores of H. costale recognized spores in paraffin sections of oyster tissue but not plasmodial stages of the parasite (Burreson 1988).

DNA Probes: A species-specific DNA probe from the small subunit ribosomal DNA sequence has been developed for H. costale (Ko et al. 1995). More recently, polymerase chain reaction (PCR) primers that target a region of the Small Sub Unit rRNA different from that of Ko et al (1995) have been identified and tested (Stokes and Burreson 2001). Multiplex polymerase chain reaction (MPCR which simultaneous tests for two or more pathogens in a single test reaction) was developed for H. costale, H. nelsoni, and Perkinsus marinus (Penna et al. 1999, 2001; Russell et al. 2000, 2004). PCR and in situ hybridization 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, , Burreson and Stokes 2006).

Methods of control

No known control methods. Losses can be minimized by harvesting oysters at 18-24 months of age. Salinities greater than 20 parts per thousand seem to favor M. costale (Andrews and Castagna 1978). The disease can be impeded by transplanting infected oysters to lower salinity waters (less than 25 ppt) (Ford and Tripp 1996). Particle filtration (1 µm filters) and UV irradiation of water entering hatcheries or nurseries should eliminate infective stages, as they do for H. nelsoni (ICES 2004). The mode of transmission is unknown.

References

Andrews, J.D. 1982. Epizootiology of late summer and fall infections of oysters by Haplosporidium nelsoni, and comparison to the annual life cycle of Haplosporidium costalis, a typical haplosporidian. Journal of Shellfish Research 2: 15-23.

Andrews, J.D. 1984. Epizootiology of diseases of oysters (Crassostrea virginica), and parasites of associated organisms in eastern North America. Helgoländer Meeresuntersuchungen 27: 149-166.

Andrews, J.D. and M. Castagna. 1978. Epizootiology of Minchinia costalis in susceptible oysters in seaside bays of Virginia's eastern shore, 1959-1976. Journal of Invertebrate Pathology 32: 124-138.

Andrews, J.D., J.L. Wood and H.D. Hoese. 1962. Oyster mortality studies in Virginia: III. Epizootiology of a disease caused by Haplosporidium costale, Wood and Andrews. Journal of Insect Pathology 4(3): 327-343.

Burreson, E.M. 1988. Use of immunoassays in haplosporidan life cycle studies. American Fisheries Society Special Publication 18: 298-303.

Burreson, E.M. and S. Ford. 2004. A review of recent information on the Haplosporidia, with a special reference to Haplosporidium nelsoni (MSX disease). Aquatic Living Resources 17: 499-517.

Burreson, E.M. and N.A. Stokes. 2006. 5.2.2 Haplosporidiosis of oysters, In: Executive Committee (ed.) Fish Health Section Blue Book, 2014 Edition, Suggested Procedures for the Detection and Identification of Certain Finfish and Shellfish Pathogens. Section 1, Diagnostic Procedures for Finfish and Shellfish Pathogens. Chapter 5 Diseases of Molluscan Shellfish. American Fisheries Society’s Fish Health Section, (For online / open access format of this section of the Blue Book see: http://afs-fhs.org/perch/resources/14069252585.2.2haplosporid2014.pdf).

Couch, J.A. and A. Rosenfield. 1968. Epizootiology of Minchinia costalis and Minchinia nelsoni in oysters introduced into Chincoteague Bay, Virginica. Proceedings of the National Shellfisheries Association 58: 51-59.

Ford, S.E. and M.R. Tripp. 1996. Diseases and Defense Mechanisms. In: Kennedy, V.S., R.I.E. Newell, A.F. Eble (eds.) The Eastern Oyster Crassostrea virginica. Maryland Sea Grant College, College Park, Maryland. pp. 581-660.

Graczyk, T.K., C.A. Farley, R. Fayer, E.J. Lewis and J.M. Trout. 1998. Detection of Cryptosporidium oocysts and Giardia cysts in the tissues of eastern oysters (Crassostrea virginica) carrying principal oyster infectious diseases. Journal of Parasitology 84: 1039-1042.

Hine, P.M., R.B. Carnegie, E.M. Burreson and M.Y. Engelsma. 2009. Inter-relationships of haplosporidians deduced from ultrastructural studies. Diseases of Aquatic Organisms 83: 247-256

ICES. 2004. Trends in important diseases affecting fish and molluscs in the ICES area 1998-2002. International Council for the Exploration of the Sea, ICES Cooperative Research Report No. 265. Copenhagen, Denmark. 26 pp. (Prepared and edited by the Working Group on Pathology and Diseases of Marine Organisms. For electronic publication see: http://www.ices.dk/sites/pub/Publication%20Reports/Cooperative%20Research%20Report%20(CRR)/crr265/crr265.pdf).

Katkansky, S.C. and R.W. Warner. 1970. The occurrence of a haplosporidian in Tomales Bat, California. Journal of Invertebrate Pathology 16: 144

Ko, Y.T., S.E. Ford and D. Fong. 1995. Characterization of the small subunit ribosomal RNA gene of the oyster parasite Haplosporidium costale. Molecular Marine Biology and Biotechnology 4: 236-240.

Mortensen, S., I. Arzul, L. Miossec, C. Paillard, S. Feist, G. Stentiford, T. Renault, D. Saulnier and A. Gregory. 2007. Molluscs and crustaceans, 5.3.21 Haplosporidiosis due to Haplosporidium costale. In: Raynard, R., T. Wahli, I. Vatsos, S. Mortensen (eds.) Review of disease interactions and pathogen exchange between farmed and wild finfish and shellfish in Europe. VESO on behalf of DIPNET, Oslo. pp. 408-409.

Penna, S., R.A. French, J. Volk, J. Karolus, I. Sunila and R. Smolowitz. 1999. Diagnostic screening of oyster pathogens: preliminary field trials of multiplex PCR. Journal of Shellfish Research 18: 319-320. (Abstract).

Penna, M.-S., M. Khan and R.A. French. 2001. Development of a multiplex PCR for the detection of Haplosporidium nelsoni, Haplosporidium costale and Perkinsus marinus in the eastern oyster (Crassostrea virginica, Gmelin, 1971). Molecular and Cellular Probes 15: 385-390.

Perkins, F.O. 1969. Electron microscope studies of sporulation in the oyster pathogen, Minchinia costalis (Sporozoa: Haplosporida). The Journal of Parasitology 55: 897-920.

Russell, S., S. Penna and R. French. 2000. Comparative evaluation of the multiplex PCR with conventional detection methods for Haplosporidium nelsoni (MSX), Haplosporidium costale (SSO), and Perkinsus marinus (Dermo) in the eastern oyster, Crassostrea virginica. Journal of Shellfish Research 19: 580-581. (Abstract). This abstract was repeated verbatum in Journal of Shellfish Research 19: 648.

Russell, S., S. Frasca Jr, I. Sunila and R.A. French. 2004. Application of a multiplex PCR for the detection of protozoan pathogens of the eastern oyster Crassostrea virginica in field samples. Diseases of Aquatic Organisms 59: 85-91.

Stokes, N.A. and E.M. Burreson. 2001. Differential diagnosis of mixed Haplosporidium costale and Haplosporidium nelsoni infections in the eastern oyster, Crassostrea virginica, using DNA probes. Journal of Shellfish Research 20: 207-213.

Sunila, I., N.A. Stokes, R. Smolowitz, R.C. Karney and E.M. Burreson. 2002. Haplosporidium costale (seaside organism), a parasite of the eastern oyster, is present in Long Island Sound. Journal of Shellfish Research 21: 113-118.

Wang, Z., X. Lu, Y. Liang and C. Wang. 2010. Haplosporidium nelsoni and H. costale in the Pacific oyster Crassostrea gigas from China's coasts. Diseases of Aquatic Organisms 89: 223-228.

Wood, J.L. and J.D. Andrews. 1962. Haplosporidium costale (Sporozoa) associated with a disease of Virginia oysters. Science (Washington, DC) 136: 710-711.

Citation Information

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

Date last revised: December 2014
Comments to Susan Bower

Date modified: