Marteilia sydneyi of Oysters
Category 3 (Host Not in Canada)
Common, generally accepted names of the organism or disease agent
QX (Queensland unknown) disease.
Scientific name or taxonomic affiliation
Marteilia sydneyi in the phylum Paramyxea as substantiated by Berthe et al. (2000, 2004). The "haplosporidian" reported in Sydney rock oysters from Moreton Bay, Queensland, Australia by Wolf (1972) was identified as M. sydneyi by Perkins and Wolf (1976).
In coastal estuaries of southern Queensland and New South Wales, Australia (Adlard and Ernst 1995, Kleeman et al. 2004, Adlard and Wesche 2005). Infection was detected in 1 of 117 Saccostrea glomerata from Dampier Archipelago, Western Australia (Hine and Thorne 2000) and in 1 of 411 oysters from the same area in 1995 (Jones and Creeper 2006). A Marteilia-like parasite was reported in 2 of 29 Saccostrea forskali in Thailand (Taveekijakarn et al. 2002, 2008).
Saccostrea (=Crassostrea) glomerata (=commercialis) and possibly also Striostrea mytiloides (=Saccostrea =Crassostrea echinata) and Saccostrea forskali. Similar protistans were reported from Ostrea angasi and clams including the giant clam Tridacna maxima.
Impact on the host
Typically, infected oysters are in poor condition with their gonad completely resorbed. Apparently, M. sydneyi may kill up to 80% of infected oysters with up to 100% prevalence (Hine 1996). Nell (2002) indicated that wherever M. sydneyi has been detected, the oyster industry has severely declined. Although the decline was slow (taking 30 years) in some areas of northern New South Wales, in the Georges River, Sydney, the industry totally collapsed in 2001 within seven years of M. sydneyi first being detected at that area. The disease is also a threat to other S. glomerata production areas in Australia (Bezemer et al. 2006). Massive invasion of the digestive gland epithelial cells by M. sydneyi leads to complete disorganization of the infected tissue. Death results from starvation and may occur in less than 60 days after initial infection. Peters and Raftos (2003) indicated that phenoloxidase activity was significantly suppressed in oysters associated with QX disease and that inhibition of the prophenoloxidase cascade may facilitate lethal infection by M. sydneyi. However, the presence of M. sydneyi in oysters does not always result in the development of QX disease which is more closely associated with the sporulation of the parasite (Simonian et al. 2009a). Haemocytes of S. glomerata can recognise and phagocytose M. sydneyi, where granules fused with phagosome membranes, the pH of phagosomes decreased, phenoloxidase deposition occurred, and ingested and melanised M. sydneyi were detected in vivo among haemocytes from infected oysters (Kuchel et al. 2010). Butt and Raftos (2008) indicated that phenoloxidase activity led to the complete melanisation of phagosomes and suggested that phagocytosis and cellular melanisation are critical defensive responses of S. glomerata infected with M. sydneyi. Also, Butt and Raftos (2007) suggested that the presence of some transient environmental stressor may affect phenoloxidase activity during a key period of infection and thus increase the susceptibility of oysters to disease. Green and Barnes (2010) employing DNA analysis techniques found that the microbiota of the digestive gland of S. glomerata is changed by infection with M. sydneyi, becoming dominated by a Rickettsiales-like prokaryote, and generally less diverse.
Oysters are reported to be susceptible to M. sydneyi during the summer months. Lester (1986), Anderson et al. (1994), Wesche (1995) and Adlard (1996) conducted transplant experiments and determined that oysters may be exposed to infection over a very short interval (possibly only two weeks per year). Once infection has been acquired, warm temperatures favour parasite development and host mortalities are greatest at the end of summer. At lower temperatures, retardation of host mortality and suppression of parasite development occurs. In some cases, infected oysters may survive the summer months and over winter with the disease, however subsequent elevated temperatures usually induce mortality. At times, M. sydneyi has caused over 90% mortality on oyster farms in northern New South Wales and southern Queensland. There was no apparent correlation with epizootics (outbreaks) of M. sydneyi and fluctuations in pH, salinity and temperature (Anderson et al. 1994, Wesche 1995). Adlard and Wesche (2005) determined that the presence of M. sydneyi in estuaries where disease has not been recorded has emphasised the likely role played by the dynamics of the parasite’s lifecycle together with host immune defence and likely environmental factors as regulators of disease outbreaks in S. glomerata. No mortalities were associated with the detection of a M. sydneyi-like parasite in Saccostrea forskali from two sites at the upper part of the Gulf of Thailand (Taveekijakarn et al. 2008).
The developmental stages of M. sydneyi in S. glomerata were identified and illustrated by Kleeman et al. (2002a). The initial infective stage of M. sydneyi enters S. glomerata through the palps and gills where extrasporogonic proliferation (i.e., proliferation that does not result in the formation of spores) occurs in the epithelium. A daughter cell within a vacuole in the cytoplasm of the uninucleate stem cell divides by binary fission to produce four daughter cells within the enlarged stem cell. Internal cleavage results in the formation of an uninucleate cell within each daughter cell. The stem cell degenerates to release the daughter cells, which become new stem cells. Following proliferation, stem cells each containing a daughter cell are liberated into surrounding connective tissue and haemolymph spaces to form a transient systemic stage. Following systemic dissemination, the parasites infiltrate the digestive gland, penetrate the basal membrane of the tubules and become established as nurse cells at the base of the epithelial cells in the digestive gland tubules. The nurse cell elongates and forms pseudopodial extensions that protrude along the basal membrane. Daughter cells formed within the nurse cells divide and are thus spread along the basal membrane to establish new infections between adjacent epithelial cells until all available sites within the digestive tubule are occupied. In mature infections, the nurse cells degrade and each daughter cell becomes the primary cell described by Perkins and Wolf (1976). The primary cell internally cleaves a secondary cell that divides to form a sporangiosorus (sporont primordia) containing 8 to 16 sporonts (sporangia) marking the initiation of sporulation. The sporulation process involves internal cleavage to produce two tricellular spores within each sporont within a sporangiosorus (Perkins and Wolf 1976). Mature spores are shed into the tubule lumen for evacuation from the oyster and infected oysters shed large numbers of spores before oyster death. The rest of the life cycle is unknown but Roubal et al. (1989) implicated filter-feeding or detritivorous invertebrates rather than scavenging invertebrates or fish in the life cycle. Individual rock oysters have been observed to eliminate low levels of M. sydneyi infection by shedding the parasite and undergoing full recovery (Roubal et al. 1989).
Gross Observations: The pale yellow-brown colour of the digestive gland contrasts with the deep green of healthy oysters. The body is greatly shrunken and tissues are translucent due to the complete resorption of the gonad.
Wet Mounts: Refractile bodies in sporonts in wet smears of digestive gland (hepatopancrease) provide a simple and rapid diagnosis when sporonts are present in infected oysters.
Figures 2 and 3. Unstained wet mount squashes of the digestive gland of Saccostrea glomerata infected with mature stages of Marteilia sydneyi
Tissue Imprints: Dried imprints of the digestive gland (hepatopancrease) on glass slides stained with Wright, Wright-Giemsa or equivalent stain (e.g. Hemacolor, Merck; Diff-QuiK, Baxter) enables the rapid detection of all developmental stages including presporulation stages not detectable in wet mount preparations within a few days of initial infection (Kleeman and Adlard 2000). Scan slides for the presence of M. sydneyi, at 200 and 400 magnification. The oil emersion objective (1000x magnification) may be required for definitive identification of early stages of infection in the digestive gland (i.e. daughter or primary cell stages). In all but very early stages of M. sydneyi incursion into the digestive gland, the infection was sufficiently homogeneous that sub-sampling of tissue for diagnosis did not have a significant negative impact on the ability of diagnostic tests to detect the disease (Adlard and Wesche 2005). A comparison between tissue imprints (cytology) and histology showed that cytology (96.88% infected) had a greater sensitivity than histology (86.11% infected) (Adlard and Wesche 2005)
Histology: Examine cross-sections of the digestive gland for the presence of Marteilia in the digestive gland epithelial cells. Marteilia sydneyi can be differentiated from Marteilia refringens by 1) the lack of striated inclusions in the sporont primordia, 2) the formation of eight to sixteen sporonts (sporangial primordia, sporangia) in each sporangiosorus (plasmodium), instead of eight, 3) the occurrence of two, rather than four, spores in each sporont (sporangium), and 4) the heavy layer of concentric membranes surrounding mature spores, in comparison to the lack of such a covering around M. refringens spores. Detection of infection has been facilitated by recent descriptions of the early developmental stages and site of infection (Kleeman et al. 2001, 2002a,c) and haemocyte infiltration throughout the tissue is common in pre-sporulating infections (Adlard and Wesche 2005). However, detection of the initial infections is facilitated by the application of DNA probes via in situ hybridisation (see Figs. 8 and 9, Fig. 10 and details pertaining to the DNA Probes below, Kleeman et al. 2002b, Adlard and Wesche 2005).
Figures 5 to 7. Extrasporogonic replication of Marteilia sydneyi in the gill and palp epithelium of Saccostrea glomerata following the initiation of infection from an unknown source. Haematoxylin and eosin stain.
Figures 8 and 9. Tissue sections through the digestive gland of Saccostrea glomerata demonstrating an early stage of disease development.
Figures 10 to 15. Presporulating stages of Marteilia sydneyi in the digestive gland tubules of Saccostrea glomerata. Haematoxylin and eosin stain except for Fig. 10 which was coloured by in situ hybridisation using the Smart 2 probe as described below.
Figures 16 and 17. Tissue section through the digestive gland of Saccostrea glomerata demonstrating an advanced stage of disease with sporulating stages of Marteilia sydneyi. Haematoxylin and eosin stain.
Electron Microscopy: Formalin fixed material processed for transmission electron microscopy can be used for species determination of M. sydneyi by visualizing the two tri-cellular spores in a sporont (Adlard and Wesche 2005).
Immunological Assay: An Indirect Fluorescent Antibody Test (IFAT) for detecting most stages of M. sydneyi has been developed by Roubal and Lester (1987). However, this assay did not react to the contents of the spore (Roubal et al. 1989). Anderson et al. (1994b) determined that the IFAT based on the polyclonal anti-QX antibody developed by Roubal et al. (1989) recognised sporulating stages of M. sydneyi but not those of related species. However, this polyclonal antibody failed to detect presumed presporulation stages of M. sydneyi in the connective tissue of recently infected oysters. Therefore, Anderson et al. (1994b) suggested that antigens of M. sydneyi were stage specific and indicated that a DNA probe rather than immunohistochemical tests may be more useful in detecting this parasite.
DNA Probes: Sequence data for the first internal transcribed spacer (ITS1) region of M. sydneyi rDNA described by Anderson et al. (1995) were used in the development of Polymerase Chain Reaction (PCR) and in situ hybridisation assays (Kleeman and Adlard 2000). PCR assays on known numbers of sporonts purified from the digestive gland of infected oysters indicated that DNA equivalent to 0.01 sporonts was detectable using agarose gel electrophoresis. However, concentrations of host DNA in excess of 50 ng per 20 µl reaction reduced the sensitivity of the test. A DIG-labelled DNA probe constructed from M. sydneyi unique primers could detect 10 pg of M. sydneyi PCR amplified DNA in dot-blot hybridisation. This probe hybridised to all developmental stages of M. sydneyi in paraffin sections of infected oyster digestive gland with no non-specific binding, however, inconsistent detection of mature sporont stages was observed (Kleeman and Adlard 2000). PCR protocols using this probe for the detection of M. sydneyi in oyster tissue were optimised and validated by Adlard and Worthington Wilmer in 2003 as reported by Adlard and Wesche (2005). PCR was found to be more sensitive in detecting infection than examining tissue imprints (98.99% infected versus 58.11% in a sample of 1839 oysters) because PCR is able to detect M. sydneyi infections at a much earlier stage of development long before pre-sporulating and sporulating stages are easily identifiable through microscopy in the oyster digestive gland (Adlard and Wesche 2005). However, according to Simonian et al. (2009a), sporulation is a more reliable marker of QX disease than PCR because sporulation corresponds with the disease state, rather than simply reflecting the presence of M. sydneyi. Simonian et al. (2009a) used proteome maps produced by two dimensional electrophoresis to identify significant differences in the expression of four proteins in oysters with sporulating M. sydneyi infections.
The "Smart 2 probe" designed by Le Roux et al. (1999) from within the 18S rRNA of M. refringens also detected M. sydneyi via in situ hybridisation, and, although not a species specific test, allowed more reliable detection of all life cycle stages (Kleeman et al. 2002b). The combined use of the M. sydneyi specific ITS1 probe and the Smart 2 probe in in situ hybridisation allowed elucidation of the occurrence of nurse cells of M. sydneyi between epithelial cells and adjacent to the basal membrane of the digestive gland tubules (Fig. 10). Nurse cells are usually not evident in normal histology stained with haematoxylin and eosin stain. The application of the in situ hybridisation assay to lightly infected oysters allowed the visualisation of early developmental stages of infection (Fig. 9) and determination of a hypothetical development cycle of M. sydneyi in S. glomerata (see Fig. 1 and Kleeman et al. 2002a).
The ITS1 PCR and in situ hybridisation assays proved specific to M. sydneyi when tested for their potential to cross-react with related species of Paramyxea, including Marteilia refringens in Ostrea edulis from France, Marteilioides chungmuensis in Crassostrea gigas from Japan, and Marteilioides sp. from Striostrea mytiloides (=Saccostrea echinata) the blacklip pearl oyster from Darwin Harbour, Australia (Kleeman et al. 2002b). Although the Smart 2 probe (Le Roux et al. 1999) cross-reacted with various species of Paramyxea (Kleeman et al. 2002b), this probe provided a stronger signal in the detection of sporont stages and was more reliable in the detection of mature spores of M. sydneyi than the ITS1 probe (compare Figs. 20 to 19 and Figs. 21 to 22). Thus, Kleeman et al. (2002b) indicated that the Smart 2 probe was preferred for use in the screening or surveillance of oyster populations and that the ITS1 probe should be used as one means of confirming the specific identity of the pathogen as M. sydneyi. Kleeman et al. (2002b) also determined that in the absence of comparative sequence data, primers designed within the internal transcribed spacer regions (ITS) are best suited for the production of species specific PCR tests.
Figures 18 to 20. Detection of Marteilia sydneyi in the digestive tubules of Saccostrea glomerata.
Figures 21 and 22. Detection of immature and mature sporont stages within sporangiosori of Marteilia sydneyi in tissue sections of Saccostrea glomerata by in situ hybridisation.
Kleeman et al.(2004) determined that single strand conformation polymorphism (SSCP)-based analysis of a region (195 bp) of the ITS1 of ribosomal DNA could be employed to detect the existence of at least two genetic variant of M. sydneyi. This molecular data supported the proposal of the origin and subsequent southward distribution of this parasite along the south eastern coast of Australia (Kleeman et al. 2004).
Methods of control
Disease control is attempted by altering culture techniques: oysters are not planted in areas of risk during the austral summer (January to March), young oysters are held in high salinity water where they grow more slowly but remain free of infection until after the risk of infection is past (late April), large oysters are harvested prior to the onset of the transmission period (before late December) and restocking for grow-out from QX free areas in autumn. In addition, determining the risk factors that result in QX disease outbreaks will allow farmers to position their farms in locations where QX disease outbreaks are less likely to occur (Green et al. 2011). Farm management practices of transferring oyster stocks between estuaries increases the risk of introducing the parasite to new areas. Legislation (disease zoning) in New South Wales has been in force to prevent the transfer of potentially infected stocks from QX-endemic areas to QX-free southern estuaries (Adlard and Ernst 1995). However, this control measure hindered the farming of S. glomerata which historically relied on transferring wild-caught spat between estuaries for on-growing to market size and did not prevent the occurrence of QX disease in new locations (Green et al. 2011). According to Green et al. (2011) the future of the S. glomerata industry relies in part on the successful commercialization of hatchery-produced QX-disease resistant S. glomerata.
To control the disease, a QX disease resistance-breeding program was established in 1996 by the Australian New South Wales Department of Primary Industries (Nell et al. 2000, Nell and Hand 2003). Initially, it was reported that phenoloxidase activity was enhanced in S. glomerata selectively bred for resistance to QX disease (Newton et al. 2004). Unfortunately, about 10 years (four generations) of selective breeding of these oysters inadvertently resulted in a stock with significantly less phenoloxidase POb a defensive enzyme associated with resistance to disease (Bezemer et al. 2006). Nevertheless, Nell and Perkins (2006) reported that progeny of third-generation selected S. glomerata breeding lines had evaluated resistance to disease caused by both M. sydneyi and Bonamia roughleyi compared to a non-selected control. But, selection for resistance to M. sydneyi did not appear to confer resistance to B. roughleyi and the converse also applied (Nell and Perkins 2006). However, the selection process that was employed may have allowed for the selection of broodstock that had been challenged by multiple diseases and thus acquired some resistance to some of them (Green et al. 2008). Simonian et al. (2009b) developed a proteomic approach using 2-dimensional electrophoresis and mass spectrometry to identify two potential markers of QX disease resistance that were in addition to phenoloxidase activity. Kuchel et al. (2010) suggested that resistance against QX disease may be associated with enhanced phagolysosomal activity in QX disease resistant oysters.
Wesche et al. (1999) determined that spores of M. sydneyi were relatively short-lived once isolated from oysters and that the majority die within 7 to 9 days (maximum longevity was 35 days at 15 °C and 34 ppt salinity). Spores did not survive more than 2 hours following ingestion by birds or fish but they remained viable for over 7 months at -20 and -70 °C. Exposure to chlorine (from granular calcium hypochlorite 650 g per kg) at 200 ppm killed 99.5% of the spores within 2 hours and all spores within 4 hours of exposure (Wesche et al. 1999).
Adlard, R.D. and I. Ernst. 1995. Extended range of the oyster pathogen Marteilia sydneyi. Bulletin of the European Association of Fish Pathologists 15: 119-121.
Adlard, R.D. and S.J. Wesche. 2005. Aquatic Animal Health Subprogram: Development of a disease zoning policy for Marteilia sydneyi to support sustainable production, health certification and trade in the Sydney rock oyster. Final Report 2001/214. Australian Fisheries Research and Development Corporation and Queensland Museum, Brisbane, Australia, pp. 1-46.
Anderson, I.G. 1990. Diseases in Australian invertebrate aquaculture. In: Proceedings and Abstracts, Fifth International Colloquium on Invertebrate Pathology and Microbial Control, Society for Invertebrate Pathology, 20-24 Aug. 1990, Adelaide, Australia, p. 38-48.
Anderson, T.J., S. Wesche and R.J.G. Lester. 1994a. Are outbreaks of Marteilia sydneyi in Sydney rock oysters, Saccostrea commercialis, triggered by a drop in environmental pH? Australian Journal of Marine and Freshwater Research 45: 1285-1287.
Anderson, T.J., T.F. McCaul, V. Boulo, J.A.F. Robledo and R.J.G. Lester. 1994b. Light and electron immunohistochemical assays on paramyxea parasites. Aquatic Living Resources 7: 47-52.
Anderson, T.J., R.D. Adlard and R.J.G. Lester. 1995. Molecular diagnosis of Marteilia sydneyi (Paramyxea) in Sydney rock oysters, Saccostrea commercialis (Angas). Journal of Fish Diseases 18: 507-510.
Berthe, F.C.J., F. Le Roux, E. Peyretaillade, P. Peyret, D. Rodriguez, M. Gouy and C.P. Vivarès. 2000. Phylogenetic analysis of the small subunit ribosomal RNA of Marteilia refringens validates the existence of Phylum Paramyxea (Desportes and Perkins, 1990). The Journal of Eukaryotic Microbiology 47: 288-293.
Berthe, F.C.J., F. Le Roux, R.D. Adlard and A. Figueras. 2004. Marteiliosis in molluscs: A review. Aquatic Living Resources 17: 433-448.
Bezemer, B., D. Butt, J. Nell, R. Adlard and D. Raftos. 2006. Breeding for QX disease resistance negatively selects one form of the defensive enzyme, phenoloxidase, in Sydney rock oysters. Fish and Shellfish Immunology 20: 627-636.
Butt, D. and D. Raftos. 2007. Immunosuppression in Sydney rock oysters (Saccostrea glomerata) and QX disease in the Hawkesbury River, Sydney. Marine and Freshwater Research 58: 213-221.
Butt, D. and D. Raftos. 2008. Phenoloxidase-associated cellular defence in the Sydney rock oyster, Saccostrea glomerata, provides resistance against QX disease infections. Developmental and Comparative Immunology 32: 299-306.
Green, T.J. and A.C. Barnes. 2010. Bacterial diversity of the digestive gland of Sydney rock oysters, Saccostrea glomerata infected with the paramyxean parasite, Marteilia sydneyi. Journal of Applied Microbiology 109: 613-622.
Green, T.J., B.J. Jones, R.D. Adlard and A.C. Barnes. 2008. Parasites, pathological conditions and mortality in QX-resistant and wild-caught Sydney rock oysters, Saccostrea glomerata. Aquaculture280: 35–38.
Green, T.J., D. Raftos, W. O'Connor, R.D. Adlard and A.C. Barnes. 2011. Disease prevention strategies for QX disease (Marteilia sydenyi) of Sydney rock oysters (Saccostrea glomerata). Journal of Shellfish Research 30: 47-53.
Hine, P.M. 1996. Southern hemisphere mollusc diseases and an overview of associated risk assessment problems. Revue Scientifique et Technique de l'Office International des Epizooties 15: 563-577.
Hine, P.M. and T. Thorne. 2000. A survey of some parasites and diseases of several species of bivalve mollusc in northern Western Australia. Diseases of Aquatic Organisms 40: 67-78.
Jones, J.B. and J. Creeper. 2006. Diseases of pearl oysters and other molluscs: a Western Australian perspective. Journal of Shellfish Research 25: 233-238.
Kleeman, S.N. and R.D. Adlard. 2000. Molecular detection of Marteilia sydneyi, pathogen of Sydney rock oysters. Diseases of Aquatic Organisms 40: 137-146.
Kleeman, S.N., R.D. Adlard and R.J.G. Lester. 2001. Detection of early infective stages of Marteilia sydneyi and their development through to sporogenesis. Book of Abstracts, European Association of Fish Pathologists, Tenth International Conference "Diseases of Fish and Shellfish". Trinity College Dublin, Ireland, 9 - 14 September 2001. pg. O-004.
Kleeman, S.N., R.D. Adlard and R.J.G. Lester. 2002a. Detection of the initial infective stages of the protozoan parasite Marteilia sydneyi in Saccostrea glomerata and their development through to sporogenesis. International Journal for Parasitology 32: 767-784.
Kleeman, S.N., F. Le Roux, F. Berthe and R.D. Adlard. 2002b. Specificity of PCR and in situ hybridization assays designed for detection of Marteilia sydneyi and M. refringens. Parasitology 125: 131-141.
Kleeman, S., R. Adlard and B. Lester. 2002c. Discovery of the early infective stage of the protozoan parasite Marteilia sydneyi in oysters and the implications for disease detection and control. Handbook and Abstracts, Fifth Symposium on Diseases in Asian Aquaculture, Queensland, Australia, 24-28 November 2002. Pg. 100.
Kleeman, S.N., R.D. Adlard, X. Zhu and R.B. Gasser. 2004. Mutation scanning analysis of Marteilia sydneyi populations from different geographical locations in eastern Australia. Molecular and Cellular Probes 18: 133-138.
Kuchel , R.P., S. Aladaileh, D. Birch, N. Vella and D.A. Raftos. 2010. Phagocytosis of the protozoan parasite, Marteilia sydneyi, by Sydney rock oyster (Saccostrea glomerata) hemocytes. Journal of Invertebrate Pathology 104: 97-104.
Le Roux, F., C. Audemard, A. Barnaud and F. Berthe. 1999. DNA Probes as potential tools for the detection of Marteilia refringens. Marine Biotechnology 1: 588-597.
Lester, R.J.G. 1986. Field and laboratory observations on the oyster parasite Marteilia sydneyi. In: M. Cremin, C. Dobson and D.E. Moorhouse (eds), 'Parasite Lives'. University of Queensland Press, pp. 33-40.
Mortensen, S., I. Arzul, L. Miossec, C. Paillard, S. Feist, G. Stentiford, T. Renault, D. Saulnier and A. Gregory. 2007. Molluscs and crustaceans. 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. Chapter 5.3.16, pp. 381-384. For electronic version see www.dipnet.info under "Documents" subgroup "Reports and project deliverables".
Nell, J. 2002. The Australian oyster industry. World Aquaculture 33: 8-10.
Nell, J.A. and R.E. Hand. 2003. Evaluation of the progeny of second-generation Sydney rock oyster Saccostrea glomerata (Gould, 1850) breeding lines for resistance to QX disease Marteilia sydneyi. Aquaculture 228: 27-35.
Nell, J.A. and B. Perkins. 2006. Evaluation of the progeny of third-generation Sydney rock oyster Saccostrea glomerata (Gould, 1850) breeding lines for resistance to QX disease Marteilia sydneyi and winter mortality Bonamia roughleyi. Aquaculture Research 37: 693-700.
Nell, J.A., I.R. Smith and C.C. McPhee. 2000. The Sydney rock oyster Saccostrea glomerata (Gould 1850) breeding program: progress and goals. Aquaculture Research 31: 45-49.
Newton, K., R. Peters and D. Raftos. 2004. Phenoloxidase and QX disease resistance in Sydney rock oysters (Saccostrea glomerata). Developmental and Comparative Immunology 28: 565-569.
Perkins, F.O. and P.H. Wolf. 1976. Fine structure of Marteilia sydneyi sp. n. - haplosporidan pathogen of Australian oysters. The Journal of Parasitology 62: 528-538.
Peters, R. and D.A. Raftos. 2003. The role of phenoloxidase suppression in QX disease outbreaks among Sydney rock oysters (Saccostrea glomerata). Aquaculture 223: 29-39.
Potter, M.A. 1983. Growth rates of cultivated Sydney rock oysters, Saccostrea (Crassostrea) commercialis, in two estuaries in subtropical southern Queensland. Queensland Journal of Agriculture and Animal Science 40: 137-140.
Roubal, F.R., J. Masel and R.J.G. Lester. 1989. Studies on Marteilia sydneyi, agent of QX disease in the Sydney rock oyster, Saccostrea commercialis, with implications for its life cycle. Australian Journal of Marine and Freshwater Research 40: 155-167.
Simonian, M., S.V. Nair, W.A. O'Connor and D.A. Raftos. 2009a. Protein markers of Marteilia sydneyi infection in Sydney rock oysters, Saccostrea glomerata. Journal of Fish Diseases 32: 367-375.
Simonian, M., S.V. Nair, J.A. Nell and D.A. Raftos. 2009b. Proteomic clues to the identification of QX disease-resistance biomarkers in selectively bred Sydney rock oysters, Saccostrea glomerata. Journal of Proteomics 73: 209-217.
Taveekijakarn, P., G. Nash, T. Somsiri and S. Putinaowarat. 2002. Marteilia-like species: first report in Thailand. The Aquatic Animal Health Research Institute Newsletter 11: 1-2.
Taveekijakarn, P., T. Somsiri, S. Puttinaowarat, S. Tundavanitj, S. Chinabut and G. Nash. 2008. Parasitic fauna of rock oyster (Saccostrea forskali) cultured in Thailand. In: Bondad-Reantaso, M.G., Mohan, C.V., Crumlish, M. and Subasinghe, R.P. (eds.) Diseases in Asian Aquaculture VI. Fish Health Section, Asian Fisheries Society, Manila, Philippines, pp. 335-342. For electronic version see http://www.fhs-afs.net/publications.htm, proceedings of The Sixth Symposium on Diseases in Asian Aquaculture held in Colombo, Sri Lanka in 2005.
Wesche, S.J. 1995. Outbreaks of Marteilia sydneyi in Sydney rock oysters and their relationship with environmental pH. Bulletin of the European Association of Fish Pathologists 15: 23-27.
Wesche, S.J., R.D. Adlard and R.J.G. Lester. 1999. Survival of spores of the oyster pathogen Marteilia sydneyi (Protozoa, Paramyxea) as assessed using fluorogenic dyes. Diseases of Aquatic Organisms 36: 221-226.
Wisely, B., J.E. Holliday and B.L. Reid. 1979. Experimental deepwater culture of the Sydney rock oyster (Crassostrea commercialis = Saccostrea cucullata). 1. Growth of vertical clumps of oysters ("ren"). Aquaculture 16: 127-140.
Wolf, P.H. 1972. Occurrence of a haplosporidan in Sydney rock oysters Crassostrea commercialis from Moreton Bay, Queensland, Australia. Journal of Invertebrate Pathology 19: 416-417.
Wolf, P.H. 1977. Diseases and parasites in Australian commercial shellfish. Haliotis 8: 75-83.
Wolf, P.H. 1979. Life cycle and ecology of Marteilia sydneyi in the Australian oyster, Crassostrea commercialis. Marine Fisheries Review 41 (1-2): 70-72.
Bower, S.M., Kleeman, S.N. (2011): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Marteilia sydneyi of Oysters.
Date last revised: October 2011
Comments to Susan Bower
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