Bonamia ostreae of Oysters
Category 2 (In Canada and of Regional Concern)
Common, generally accepted names of the organism or disease agent
Microcell disease, Bonamiasis, Bonamiosis, Haemocyte disease of flat oyster, Haemocytic parasitosis.
Scientific name or taxonomic affiliation
Bonamia ostreae (Pichot et al. 1980). Results of initial ultrastructural studies suggesting that this protist was affiliated with the Haplosporidia despite the lack of a spore stage (Bonami et al. 1985, Brehélin et al. 1982). This taxonomic affiliation was subsequently confirmed by DNA analysis (Carnegie et al. 2000b, Reece et al. 2004, López-Flores et al. 2007). Related species include Bonamia exitiosa a pathogen of New Zealand dredge oysters Ostrea chilensis, Bonamia (=Mikrocytos) roughleyi a pathogen of Sydney rock oysters Saccostrea glomerata, Bonamia perspora a parasite of the crested or horse oyster Ostreola equestris and other unidentified Bonamia spp. from various species of oysters in distant locations.
Western Europe along the coast from Spain to Denmark, Ireland and Great Britain (excluding Scotland and Wales). Bonamis ostreae was reported for the first time from Morocco in 2005 (Culloty and Mulcahy 2007) and was detected in archived (in 1990) samples of Ostrea edulis from the Manfredonia Gulf (Adriatic Sea) of Italy and again detected along with Bonamia exitiosa in a few (3 of 750) O. edulis collected in 2007 (Narcisi et al. 2010). Earliest records were from the west (California and Washington) and east (Maine) coasts of the USA. In California during the mid 1960s, Katkansky and Manzer (1967) reported high mortalities and heavy infections of microcells in O. edulis that had originated from Milford, Connecticut a few years earlier. In both Washington and Maine, the prevalence of infection is usually low and heavy infections are rare. Current evidence suggests that B. ostreae was inadvertently introduced into Maine, Washington and Europe from California by the translocation of infected O. edulis in the late 1970s (Elston et al. 1986, Friedman and Perkins 1994, Cigarría and Elston 1997). In the fall of 2004, this parasite was detected for the first time in O. edulis farmed in British Columbia, Canada (Marty et al. 2006).
Ostrea edulis and also known to infect Ostrea angasi, Ostrea chilensis, (=Tiostrea chilensis, =Tiostrea lutaria, =Ostrea lutaria), Ostrea puelchana, Ostrea denselamellosaand Crassostrea ariakensis (=rivularis), and Crassostrea angulata (Grizel et al. 1982, Carnegie and Cochennec-Laureau 2004). The Pacific oyster, Crassostrea gigas (Renault et al. 1995, Cao et al. 2009, Lynch et al. 2010), mussels, Mytilus edulis and Mytilus galloprovincialis, and clams, Ruditapes decussatus and Venerupis (=Ruditapes) philippinarum could not be naturally nor experimentally infected and these bivalves did not appear to act as vectors nor intermediate hosts for the parasite (Culloty et al. 1999). However, C. gigas may act as a carrier or reservoir host ofB. ostreae as indicated by Lynch et al.. (2010) who reported detecting positive Polymerase chain reaction (PCR) signal and visualized a few B. ostreae-like cells in haemocytes and extracellularly in two C. gigas. Microcells in the vesicular connective tissue cells of Ostrea conchaphila (=Ostrea lurida) from Oregon, USA were speculated to be B. ostreae (Farley et al. 1988). However, Elston (1990) indicated that although experiments suggest that O. conchaphila may contract the disease, infection has not been positively demonstrated and Arzul et al. (2005a) could not infect O. conchaphila by cohabitation for 11 months with diseased O. edulis.
Impact on the host
Thebault et al. (2003) listed and applied 24 epidemiological and experimental criteria to demonstrate that B. ostreae was the infectious agent responsible for mass mortalities of O. edulis. Bonamia ostreae, in conjunction with earlier epizootics caused by Martelia refringens, caused a drastic drop in the French production of O. edulis from 20,000 t per year in the 1970's to 1,800 t in 1995 (Boudry et al. 1996; Arzul et al. 2005b, 2006). Bonamia ostreae has also had a significant negative impact on O. edulis production throughout its distribution range in Europe (Tigé et al. 1981, 1982; Grizel 1983, Culloty and Mulcahy 2007). Although many infected oysters appear normal, others may have yellow discolouration and/or extensive lesions (i.e. perforated ulcers) in the connective tissues of the gills, mantle and digestive gland. Pathology appears correlated to haemocyte destruction and diapedesis due to proliferation of B. ostreae (Balouet et al. 1983, Berthe 2004). The transmission of infection between oysters is direct with no requirement for an intermediate host (Tigé et al. 1982, Tigé and Grizel 1982, Poder et al. 1983, Hervio et al. 1995, Culloty et al. 1999) and this has been experimentally demonstrated in the field as well as by cohabitation and injection in the laboratory (Bachère et al. 1986, Lallias et al. 2008). However, the possible involvement of a carrier/reservoir host should not be ruled out (Lynch et al. 2006, 2010). When benthic macroinvertebrates and zooplankton from a B. ostreae-endemic area were screened for the presence of parasite DNA, using polymerase chain reaction (PCR), 8 benthic macroinvertebrates and 19 grouped zooplankton samples gave positive results, and in the laboratory the transmission of B. ostreae was effected to two naïve O. edulis cohabiting in the laboratory with the brittle star, Ophiothrix fragilis (Lynch et al. 2007). Although these positive results with alternate hosts could be indicative of parasitism, it is equally as plausible that the animals were only casually associated with B. ostreae or had consumed infected oysters (Culloty and Mulcahy 2008).
Infection was demonstrated to result in the increase in the number of tissue infiltrating haemocytes (granulocytic reaction) (Balouet and Poder 1985, Cochennec-Laureau et al. 2003). Although some flat oysters die with light infections, others succumb to much heavier infections. Heavily infected oysters tend to be in poorer condition than uninfected oysters. In one study, the presence of Bonamia was better related to size than to age of O. edulis and infection level was statistically independent of gonadal development stage (Cáceres-Martínez et al. 1995). In another study, prevalence was highest in the largest oysters in spring and declined disproportionately in autumn, possibly due to high mortality of large oysters before autumn, suggesting that prevalence depends on oyster age (Engelsma et al. 2010). However, Robert et al (1991) and Culloty and Mulchy (1996) found that two years appeared to be the critical age for disease development in O. edulis in the Bay of Arcachon, France and on the south coast of Ireland, respectively. Nevertheless, both 0+ and 1+ year-old O. edulis are susceptible to infection and can develop a high prevalence and intensity of infection over a six-month period with associated mortalities (Lynch et al. 2005a, b; Lallias et al. 2008). Arzul et al. (2011) demonstrated that larvae of O. edulis can be infected with B. ostreae in the epithelium surrounding their visceral cavities while being held within the pallial cavity of infected mother oysters. Larvae might thus contribute to the spread of the parasite during their planktonic life. Some studies reported a seasonal pattern of prevalence and mortality, with highest levels occurring in autumn-winter (Montes 1990, Van Banning 1991, Culloty and Mulcahy 1996). In France, transmission occurred throughout the year but rates of infection seemed to be less from July to November (15% prevalence compared to 50% prevalence during March to June) (Tigé and Grizel 1982). Also in marine Lake Grevelingen in the Netherlands, B. ostreae was detected in flat oysters throughout the year with a higher prevalence in spring than in autumn (Engelsma et al. 2010). Male and female oysters were equally affected (Culloty and Mulchy 1996).
In vitro tests were used to determine that haemocytes of C. gigas were able to bind more B. ostreae than were haemocytes of O. edulis (Fisher 1988). This difference in the ability of haemocytes to bind the parasite in conjunction with the apparent inability of O. edulis haemocytes to digest the parasites once they are ingested (Balouet et al. 1983, Chagot et al. 1989, Hervio et al. 1989, Chagot et al. 1992, Xue and Renault 2000) may be factors relevant to the differences in susceptibility to infection and disease development in the two species of oysters. Cochennec-Laureau et al. (2003) reported that the proportion of granulocytes (granulated haemocytes) in O. edulis decreased with infection possibly as a result of these cells being destroyed or degranulated by B. ostreae suggesting that hyalinocytes (agranular haemocytes) may be involved in parasite survival and/or development. Da Silva et al. (2008) also found a similar correlation and supported the hypothesis that a high percentage of granulocytes and low percentage of hyalinocytes in a stock of O. edulis would enhance oyster immune ability and, consequently, would contribute to lower susceptibility to disease and longer lifespan. Insignificant correlation was found between haemolymph protein concentration and lysozyme levels and infection of O. edulis by B. ostreae (Cronin et al. 2001). From the results of in vitro experiments, Morga et al. (2009) suggested that B. ostreae actively contributed to the modification of haemocyte activities (decrease in non specific esterase activities and reactive oxygen species production) in order to ensure its own intracellular survival.
Gross Observations: Bonamiosis is sometimes accompanied by yellow discoloration and extensive lesions on the gills and mantle of O. edulis infected with B. ostreae. However, most of the infected oysters appear normal (Arzul and Joly 2011).
Tissue Imprint: Make acetone- (or methanol-) fixed impression smears from gill or heart tissue (preferably the ventricle since the auricles contain an abundance of serous cells which make detection of the parasite difficult). Stain with Wright, Wright-Giemsa, May-Grunwald-Giemsa or equivalent stain (e.g., Hemacolor, Merck; Diff-QuiK, Baxter). Examine for 2-5 µm spherical or ovoid organisms with a central nucleus within or outside the haemocytes (Arzul and Joly 2011). (Note: the organisms are enlarged by this method compared to those in fresh or histological preparations.) This method will also detect B. ostreae in hearts of oysters frozen and stored at -20°C for at least four years and held at 4°C for several hours before testing. Zabaleta and Barber (1996) observed that results obtained from the examination of stained haemolymph smears (histocytology) and histological preparations of an infected O. edulis populations were the same but suggested that histology was preferred for detecting light infections. O'Neill et al. (1998) recommended that the ventricular heart smear technique be used in conjunction with either haemolymph smears or histology to increase the possibility of detecting light infections. Culloty et al. (2003) indicated that the stained heart smear technique is not reliable for detecting latent infections. Lynch et al. (2008) claimed that heart smear examination was the most sensitive individual technique compared to histology, a Polymerase chain reaction (PCR) test and an in situ hybridization (ISH) assay, but a greater sensitivity of detection was obtained when results of heart smear and PCR screening were combined.
Histocytology: Haemolymph is withdrawn from the adductor muscle into an anticoagulent using a syringe and needle (21 Gauge). The haemocytes are placed (by cytocentrifugation or cell adhesion) in a monolayer onto poly-L-lysine coated glass slides and stained and examined as for tissue imprints. Da Silva and Villalba (2004) found this technique to be more sensitive in detecting B. ostreae than tissue imprints and histology.
Histology: Examine haematoxylin and eosin stained tissue cross-sections for tiny protozoa (2-4 µm in diameter) within haemocytes. Bonamia ostreae is distributed systemically in advanced infections (Balouet and Poder 1985). In early infections, B. ostreae are often observed within haemocytes, associated with dense focal haemocyte infiltrations in the connective tissue of the gill and mantle, and in the vascular sinuses around the stomach and intestine (Bucke 1988). Bachère et al. (1982a) preferred stained imprints of gill tissue over histological examination of the digestive gland for the diagnosis of B. ostreae. Arzul and Joly (2011) indicated that histopathology appears more reliable than tissue imprints for the detection of the parasite in case of low level of infections. However, tissue imprints are more rapid and less expensive than histopathology. Van Banning (1990) proposed that B. ostreae was an ovarian tissue parasite for part of its life cycle.
Electron Microscopy: Uninucleate, diplocaryotic and plasmodial stages with 3 to 5 nuclei have been described and illustrated (Pichot et al. 1980, Comps et al. 1980, Brehélin et al. 1982, Bonami et al. 1985, Montes et al. 1994). Intracellular structures include mitochondria, haplosporosomes, Golgi apparatus and persistent intranuclear microtubules. Two forms of B. ostreae were described: dense forms, 2-3 µm in diameter with cytoplasm rich in ribosomes, haplosporosomes and one or two mitochondria; and clear forms, 2-4 µm in diameter with a large nucleolus in the nucleus (Grizel 1987, Bucke 1988). Transmission electron microscopy is not recommended as a diagnostic technique because it is time consuming and not practical for routine application but is recommended when Bonamia like parasites are described in a new host species (Arzul and Jolu 2011). Although Hine et al. (2001) presented ultrastructural differences between B. ostreae and B. exitiosa, Narcisi et al. (2010) found that the ultrastructural characteristics of B. exitiosa occurring in Italy were so variable that they cannot be used to definitively identify a Bonamia species. Montes et al. (1994) observed B. ostreae within branchial epithelial cells of O. edulis. Immunological Assay: An immunofluorescent technique based on monoclonal antibodies was developed by Mialhe et al. (1988b). However, this technique gave unclear results when tested extensively on oysters from Maine, USA (Zabaleta and Barber 1996). Although direct monoclonal antibody sandwich immunoassay for the detection of B. ostreaee in haemolymph samples of O. edulis was developed (Cochennec et al. 1992) and marketed commercially for a few years in the mid 1990s, it is no longer available on the market.
DNA Probes: Segments of the ribosomal RNA locus (including parts of the small subunit (SSU rDNA or 18S rDNA) and internal transcribed spacers (ITS1)) and two actin genes have been sequenced by polymerase chain reaction (PCR) and molecular cloning (López-Flores et al. 2007). A PCR reaction specific for a rDNA amplicon (528 base pairs (bp) spanning 341 bp of 18S rDNA and 187 bp of ITS1) with a gene sequence resembling that belonging to members of the Phylum Haplosporidia was identified and found to detect the parasite in naturally infected O. edulis in Maine, USA (Carnegie et al. 2000a, b). This amplicon has also been developed into an in situ hybridization (ISH) assay (Carnegie et al. 1999, 2001, 2003). The PCR assay proved to be more sensitive and less ambiguous than standard cytological (tissue imprint) techniques (Carnegie et al. 2000b, Carnegie and Cochennec-Laureau 2004, Lynch et al. 2005b) and histology (Balseiro et al. 2006). Another DNA probe, that amplifies a 300 base pair product, was identified from the same area of the genome by Cochennec et al. (2000). In addition to detecting B. ostreae, these probes also detected Bonamia exitiosa and Haplosporidium nelsoni but B. ostreae can be differentiated from the other Haplosporidia by the application of restriction fragment length polymorphism (RFLP) analysis (Hine et al. 2001, a standard operating procedure for this technique is presented at http://www.eurl-mollusc.eu/SOPs). Marty et al. (2006) developed a real-time TaqMan® PCR assay that amplified a 68-bp target DNA fragment of the 18S rDNA and was designed not to amplify DNA of other Haplosporidia. This assay proved to have greater diagnostic sensitivity than histopathology even when used to analyse paraffin sections (Marty et al. 2006). Corbeil et al. (2006) also developed a real-time TaqMan® PCR assay for the detection of Bonamia spp. (but not Haplosporidium nelsoni nor Haplosporidium costale) that was comparable to conventional PCR in sensitivity but produced more rapid results with a low risk of sample cross-contamination and can be optimised to determine the intensity of infection. The real-time PCR assay developed by Robert et al. (2009) did not cross-react with closely related parasites, including Bonamia exitiosa, was at least 10-fold more sensitive than conventional PCR (performed according to Cochennec et al. (2000)) and was quantitative. If PCR was used to detect infection beyond the know geographic and host range of B. ostreae, visualization of the parasite and/or sequencing the product is required for diagnosis and to confirm that the DNA detected by PCR is that of B. ostreae (Culloty and Mulcahy 2007, Narcisi et al. 2010). In general, DNA based diagnosis tools need validation, specificity definition and further development prior to full implementation (Renault 2008). Nevertheless, ISH with a digoxigenin (DIG) labelled probe has been employed to locate light infections of B. ostreae within histological sections of the gills and epithelium of the digestive tract suggesting that these tissues may be the sites of first infection. Also, fluorescent ISH using a cocktail of 3 fluorescein labeled probes did not cross-react with H. nelsoni (Carnegie et al. 2003).
Culture: Limited multiplication of B. ostreae from explants of gills from heavily infected oysters was achieved after 3 days in vitro at 20°C (Comps 1983). Protocols for the preparation of purified B. ostreae cell suspensions from infected oysters have been described using a discontinuous density gradient of Percoll (Bachère et al. 1982b, Bachère et al. 1986) and a discontinuous sucrose gradient (Mialhe et al. 1988) The purified cells from both techniques retained infectivity and ultrastructural morphology and have been used in cytochemistry assays of the parasite (Hervio et al. 1991). Purified isolates have also been used to determine that in vitro, B. ostreae had a significantly lower survival at 25°C compared to 4°C and 15°C (especially after 48 hours of incubation), and high salinities (greater than or equal to 35 grams per litre salt in seabed borewater supplemented with natural salt) favoured parasite survival (Arzul et al. 2009).
Methods of Control
Ensure that no flat oysters from the United States of America or Europe are introduced into areas where bonamiasis is not known to occur. Pathogen transfers via movements of aquatic organisms appear to be a major cause of epizootics (Renault 2008). Some oysters from endemic areas may be asymptomatic and show no sign of Bonamia using routine detection techniques. Because larvae of O. edulis can be infected with B. ostreae while being held within the pallial cavity of infected mother oysters, the transfer of larvae for aquaculture purpose should be controlled especially when they are exported from areas where B. ostreae is present (Arzul et al. 2011). If infected animals are introduced into a naïve population, high mortalities can be expected for at least 6 years (van Banning 1985, 1991). To date, there are no known eradication procedures. Despite early attempts to eradicate B. ostreae from the Netherlands (Van Banning 1988), this parasite is now endemic to O. edulis in marine Lake Grevelingen, the Netherlands (Engelsma et al. 2010).
Mortalities due to bonamiasis can be reduced using suspension culture, reduced handling stress and lower stocking densities (Tigé et al. 1984). In Galicia, Spain, raft cultured oysters suspended at 1-2 meters depth had lower prevalence of infection and fewer mortalities then cohorts held at 8-9 meters depth suggesting that proximity to the sea floor may be a factor in transmission (Lama and Montes 1993). Subtidal growing areas also appear to be less severely affected than intertidal areas. Oyster seed from natural settlement should be avoided because these oysters tend to be significantly more parasitized than seed produced by hatcheries (Conchas et al. 2003). Montes et al. (2003) observed that O. edulis could be successfully cultured in areas of Galicia, Spain, contaminated with B. ostreae if they were promptly marketed after about 15 to 18 months of culture. Also, Arzul et al. (2006) indicated that bonamiosis kills oysters older than two years of age but O. edulis can reproduce after year one. Thus, oyster stocks that are regularly harvested for further growth or marketing results in the elimination of highly infected oysters. Le Bec et al. (1991) suggested that culturing O. edulis with C. gigas, which are not naturally susceptible to infection, may help to reduce infection in O. edulis. However, in one study, the growth of O. edulis was reduced when they were cultured with C. gigas (Robert et al. 1991). Also, B. ostreae may weaken the competitive ability of O. edulis relative to the introduced Pacific oyster C. gigas, particularly in years with high water temperatures (Engelsma et al. 2010). Despite management practices of reducing stocking densities under suspension culture or selling oysters at a lower weight before significant B. ostreae-induced mortalities occur, the production of O. edulis in Europe has remained low due to bonamiosis (Lallias et al. 2008).
Experimental infection by inoculation of B. ostreae into O. edulisfrom three separate populations in France found no significant difference in susceptibility between the populations (Bachère and Grizel 1983). However, field studies to investigate the potential disease resistance in a number of O. edulis populations from various locations in Europe indicated that some stocks performed significantly better (determined by prevalence and intensity of infection measurements and cumulative mortality) in some trials than others (Culloty et al. 2004). In Quiberon Bay, France where commercial production of O. edulis depends on the transfer of oysters from other regions of Brittany prior to marketing, despite the risks related to transfers of live molluscs, and where B. ostreae has been detected since 1980, the prevalence of B. ostreae is usually lower than 15% with less severe outbreaks than in the past suggesting that the oysters have developed a relative natural tolerance to the parasite (Arzul et al. 2005b). Also, detection frequencies recorded in the two main grow-out areas of France (Quiberon and Cancale bays) were not significantly correlated suggesting that environmental parameters and aquacultural practices have more impact on the evolution of the disease than initial parasite burden (Arzul et al. 2006). Montes et al. (1996) also reported that in Galicia, Spain, the prevalence of infection in experimentally exposed oysters varied significantly with location.
The breeding of bonamiosis-resistant flat oysters is reported to have some success (Martin et al. 1993; Boudry et al. 1996; Baud et al. 1997; Naciri-Graven et al. 1998, 1999; Culloty et al. 2001; Lallias et al. 2008). However, there is evidence from DNA microsatellite loci analysis that a population bottleneck has occurred during the selection process in some stocks of bonamiosis-resistant O. edulis. The small effective number of breeders are expected to lead to increasing inbreeding and have important consequences for the future management of at least three selected bonamiosis-resistant populations (Launey et al. 2001). Quantitative trait loci (QTL) analyses using a two-stage testing strategy and interval mapping methods were used to detect resistance to B. ostreae in a family of O. edulis derived from a cross between a wild oyster and an individual from a family selected for resistance to bonamiosis (Lallias et al. 2009). Utilizing a proteomic approach, Cao et al. (2009) envisaged the application of two-dimensional electrophoresis to the analysis of haemolymph proteins to understand the interaction between oysters and B. ostreae and to find the bases of tolerance/resistance to bonamiosis. Morga et al. (2010) studied the haemocyte response of O. edulis to B. ostreae at the transcriptome levels based on the use of real time PCR assays and suggested using a combination of glyceraldehyde 3-phosphate-dehydrogenase (GAPDH) and elongation factor 1 alpha (EF1-α) as reference genes (for which they characterized the complete open reading frame (ORF)) when examining expression levels of housekeeping genes in haemocytes of O. edulis. Morga et al. (2011) also used suppression subtractive hybridisation (SSH) to identify five oyster genes (omega glutathione Stransferase (OGST), superoxide dismutase (SOD), tissue inhibitor of metalloproteinase (TIMP), galectin, interferon regulatory factor (IRF-like) and filamin genes) with increased expression in haemocytes infected with B. ostreae. The expressed sequence tags (ESTs) of interest including genes involved in cytoskeleton, respiratory chain, detoxification membrane receptors, and immune system.
A controversial approach to developing bonamiosis-resistant was suggested by Morvan et al. (1994) and Morvan et al. (1997) who determined that B. ostreae and not O. edulis were sensitive to the antimicrobial peptides magainin 1 (originally extracted from the skin of the frog Xenopus laevis) and tachyplesin 1 (extracted from haemocytes of the Japanese horseshoe crab Tachypleus tridentatus), which may provide effective gene sequences that could possibly be used to genetically transform molluscs.
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Bower, S.M. (2011): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Bonamia ostreae of Oysters
Date last revised: April 2011
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