Juvenile Disease of Eastern 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 1 (Not Reported in Canada)

Common, generally accepted names of the organism or disease agent

Juvenile oyster disease, JOD, Roseovarius oyster disease, ROD.

Scientific name or taxonomic affiliation

JOD is used to describe a syndrome of juvenile oyster morbidity and mortality occurring in Crassostrea virginica from the northern Atlantic coast of the United States. Although initially attributed to a toxin, perhaps of bacterial or microalgal origin, or by an intracellular protistan parasite (Small 1997), Boettcher et al. (1999b, 2000) proposed that JOD was associated with a novel species of alpha-proteobacterium which was tentatively named Roseimarina crassostreae (Maloy and Boettcher 2003). The bacterium was subsequently identified as Roseovarius crassostreae and is believed to be a member of the Roseobacter clade of the marine α-Proteobacteria (Boettcher et al. 2005, Maloy et al. 2007a). The deposition of organic material on the inner shell surface resembles the lesion observed in brown ring disease of Manila clams which is the result of a bacterial infection by Vibrio tapetis (Elston 1999, Paillard et al. 2004).

Geographic distribution

North eastern United States from Maine to New York. A similar shell disease was reported by Renault et al. (2002) in C. virginica reared in two experimental localities on the Atlantic coast of France.

Host species

Crassostrea virginica. Other bivalves (Mercenaria mercenaria and Ostrea edulis) cultured in nearby waters did not seem to be affected (Lewis et al. 2006a).

Impact on the host

Sporadic high mortalities (40% to over 90%) typically occur during July and August (when water temperatures exceed and remained over 20°C) in juvenile oysters (15 to 25 mm in shell length) during their first growing season at sites where salinities are above 18 ppt (Boettcher et al. 2006). This disease was first recognized in the mid 1980s (Ford 2006). Oysters in diseased populations experienced a reduction in growth rate followed by morphological signs as described below. Bacteria and ciliates were found associated with tissues of some affected oysters. Mortalities were usually observed coincident with or shortly after the onset of signs and within a week, losses often exceeded 50%. Most losses occurred in oysters between 6 and 30 mm shell height with the smaller size cohorts having the greatest mortality. From experimental field studies, Barber et al. (1998) concluded that the occurrence of JOD was site specific (not dependent on source of seed), and that under challenge from JOD, selected oysters not only grew faster than wild (unselected) oysters, but exhibited a genetically based tolerance of this disease. JOD is prevalent on the Rhode Island coast but severe mortality events were triggered only when oysters less than 20 mm in shell length were exposed to warm water temperatures above 25°C (Markey et al. 2008). Laboratory studies indicate that JOD is infectious and readily transmissible 3 to 7 weeks after exposure to JOD-infected oysters with mortalities and conchiolin deposition enhanced by warm temperatures (22 to 26°C) and salinities of 18 to 30 ppt (Ford and Tripp 1996, Lewis and Farley 1997, Lewis et al. 1996). Although the aetiology of JOD has been associated with various organisms including: an intracellular protistan parasite, elevated levels of Vibrio spp. and a ciliate (Mesanophrus (=Paranophyrs) magna) (Small 1997, Ford 2006 and Horwitz et al. 2006, respectively), the application of Koch's postulates has confirmed the disease to be caused by Roseovarius crassostreae (Boettcher et al. 2006, Maloy et al. 2007a). Maloy et al. (2007a) proposed that the disease now be known as Roseovarius oyster disease (ROD). However, secondary infections, loss of haemolymph and generalized physiological dysfunction may also contribute to oyster death (Boettcher et al. 2006).

Diagnostic techniques

Gross Observations

Mortalities associated with reduced tissue and shell growth, fragile and uneven shell margins, cupping of the left valve, reduced condition index, mantle retraction and abnormal proteinaceous brownish material (conchiolin, an organic shell matrix) deposits around the periphery of the mantle on the inner shell surface several millimeters from the shell's edge. The conchiolin deposit (a non-specific host response by oysters) often appears as a raised ridge around the retracted mantle edge but may cover the entire mantle surface and interfere with the attachment of the adductor muscle to the shell (Ford and Borrero 2001, Boettcher et al. 2006). Dying oysters may also display unequal shell growth with the left valve outgrowing the right valve, and tissue emaciation. Depending on the size of the oyster, not all clinical signs may be present. Oysters less than 15 mm in shell length often die before they mount the characteristic conchiolin response. Oysters 15 to 25 mm in shell length typically show all gross signs of JOD and the highest mortality rate occurs among oysters less than 25 mm in shell length. Juvenile oysters over 25 mm in shell length often show abnormal conchiolin deposition but growth affects (including uneven valve margins) and mortalities are less common. Oysters that survive JOD may develop external shell checks that result when normal growth resumes (Boettcher et al. 2006).

Wet Mounts

Oyster (host) cell debris and foreign material is usually sequestered beneath successive layers of inner shell deposition. Roseovarius crassostreae can be visualised on the inner edge of affected valves or within flakes of conchiolin after staining with a nucleic acid viability stain (BacLight Live/Dead stain, Molecular Probes, Eugene, OR) and viewed with fluorescence microscopy (Boardman et al. 2008).

Histology

Lesions (increased haemocyte accumulations in the sinusoidal tissues of the mantle underlying the shell epithelium associated with attenuation and cuboidal metaplasia and/or hyperplasia of shell epithelium) occur in the mantle before abnormal conchiolin is deposited (Boettcher et al. 2006). The main conchiolin deposit was of uniform consistency (4 to 6 µm thick). In extreme cases, the conchiolin covered the entire mantle surface and the soft tissue of the oyster became detached from the shell. Successive organic sheets interspersed with haemocytes and debris, including bacteria, were laid down on the mantle surface between the conchiolin deposit and the epithelium. Abscess-like lesions were sometimes present adjacent to the deposits. These lesions contained dense, spherical bodies (1 to 7 µm in diameter), haemocytes aggregations in the connective tissue under the mantle epithelium, diapedesis of haemocytes through the epithelium, presumptive autophagosomes (Perkins 1996) and epithelial-cell erosion. Also, protozoa and bacteria were common in the area where the epithelium was eroded (Ford and Borrero 2001). However, standard histopathologic examination using standard haematoxylin and eosin stains did not reveal the identity of the etiological agent which does not appear to invade the oyster soft tissues (Bricelj et al. 1992, Lewis et al. 1996, Perkins 1996, Ford and Borrero 2001, Ford 2006. Lewis et al. 2006a, Boettcher et al. 2006, Maloy et al. 2007a). Roseovarius crassostreae may be visualised within the lesions after the application of specific stains such as immunofluorescence or in situ hybridisation.

Electron Microscopy

Scanning electron microscopy of diseased individuals can reveal abundant colonization by R. crassostreae on and within layers of host-derived conchiolin on the inner valve surfaces of the shell. The bacteria are 1.1 to 4.8 µm in length and 0.6 to 1.2 µm in width and the majority have one or two flagella at lateral, polar or subpolar positions. Occasionally bacteria have a single polar fimbriae tuft and rarely, both a fimbriae tuft and a flagellum may occur at the same pole of a single cell (Boettcher et al. 2005). Most R. crassostreae are attached to oyster tissue at the cell poles, which is consistent with the ability of R. crassostreae to express polar fimbriae (Boardman et al. 2008).

DNA Probes

All isolates examined by Boettcher et al. (1999a, 2000) had identical 16S ribosomal DNA (rDNA) regardless of the year, location or severity of the epizootic. However, sight variations in genotypes were detected (Maloy and Boettcher 2003) and two genotypes were identified based on the internal transcribed spacer (ITS) between the 16S and 23S rDNA genes (Paillard et al. 2004). Species-specific primers that amplify the ITS region using the polymerase chain reaction (PCR) were described by Maloy et al. (2005). Confirmation of product (amplicon) identity was accomplished by restriction fragment length polymorphism (RFLP) via overnight digestion of the amplicon with the restriction enzyme Ava I and resolved by electrophoresis on a 1.5% agarose gel (Maloy et al. 2005, 2007a). The assay has the potential to detect as few as 10 cells of R. crassostreae per oyster when samples are taken from the inner valve surfaces of the animal. Inclusion of material from soft body surfaces is not necessary, and may reduce sensitivity approximately 10-fold (Maloy et al. 2005). Two genetic signatures were discerned in the ITS region by Ava I digestion and genetic sequencing localized the Ava I site to a 100 base-pair region of low sequence identity consisting of 11 distinct genotypes (Maloy et al. 2007b).

Culture

Wash diseased oysters in filter-sterilized (0.2 µm) 70% seawater (FSSW). Aseptically suspend material scraped from the affected inner area of the shell into FSSW and spread suspension onto the surface of a seawater-typtone (SWT) agar plate (prepared with 0.5% tryptone, 0.3% yeast extract, 0.3% glycerol, and 1.5% agar in 70% seawater). Incubate the plates at room temperature (22–27°C) for 5–7 days and identify putative R. crassostreae colonies by morphological and growth characteristics (Boettcher et al. 2005, Boardman et al. 2008). Young colonies appear uniformly round, non-mucoid and semi-translucent with a characteristic umbonate shape, a somewhat chalky consistency and leave a noticeable impression when removed from the surface of an agar plate. After about 5 days of growth typical colonies are usually beige to pinkish-beige and about 1.0 mm in diameter or some isolates may have smaller colonies (about 0.5 mm in diameter) with a greenish-yellow appearance. The bacteria are Gram-negative, aerobic, nonfermentative, oxidase and catalase-positive, strictly marine and rod or ovoid in appearance. They are actively motile by one or two flagella, but cells are also observed to produce tufts of polar fimbriae (Boettcher et al. 2005). Confirmation of colony identity can be conducted using species-specific primers to PCR-amplify the ITS spacer region between the 16S and 23S rDNA followed by restriction fragment length polymorphism (RFLP) analysis (Maloy et al. 2005).

Methods of control

Mortality was reduced or eliminated in oysters maintained in 25 µm filtered water diluted with high salinity well water. The reduction of stocking densities within growing trays, increased flow rate in up-wells and mesh size of 6 mm or greater in grow-out containers also lessened mortalities (Boettcher et al. 2006). Also, Barber et al. (1996) found that cumulative mortality of oysters was related to timing of deployment with cohorts placed in the Damariscotta River, Maine before June or after August having significantly lower mortalities (20%) then cohorts deployed between these dates (with 64-90% cumulative mortalities). Barber et al. (1996) suggested that potential management options to reduce JOD impact include either 1) early spawning and deployment in May, ensuring a mean shell height greater than 25 to 30 mm is reached prior to August, a strategy supported by Lewis et al. (2006b) or 2) late spawning and deployment after mid-August to avoid exposure to JOD. Selection of disease resistant stocks may also be a successful management strategy for avoiding the devastating effects of JOD (Farley et al. 1997, Barber et al. 1998, Lewis et al. 2006b, Rawson et al. 2008). Davis and Barber (1999) reported that selected sublines of C. virginica had reduced mortality presumably because of their faster growth and hardiness thus enabling the cohort to reach a refuge size rage sooner thereby reducing susceptibility to JOD. Boettcher et al. (2000) reported that oysters raised at a JOD enzootic site that were unaffected by JOD were stably and persistently colonized by Stappia stellulate -like bacterial strains which appeared to inhibit the growth the JOD bacterium in an in vitro assay. However, the potential protective abilities of these bacteria require further investigation.

References

Barber, B.J., R.B. Carnegie and C.V. Davis. 1996. Effect of timing of seed deployment on growth and mortality of oysters, Crassostrea virginica, affected by Juvenile Oyster Disease (JOD). Journal of the World Aquaculture Society 27: 443-448.

Barber, B.J., C.V. Davis and M.A. Crosby. 1998. Cultured oysters, Crassostrea virginica, genetically selected for fast growth in the Damariscotta River, Maine, are resistant to mortality caused by Juvenile Oyster Disease (JOD). Journal of Shellfish Research 17: 1171-1175.

Barber, B., C. Davis and R. Hawes. 1998. Genetic selection of oysters for fast growth and disease resistance in Maine. Abstracts of the First Annual Northeast Aquaculture Conference and Exposition, Rockport, Maine, USA, November 18-19, 1998. pp. 78-79 (Abstract).

Barber, B.J., C.V. Davis, R.B. Carnegie and K.J. Boettcher. 2000. Management of Juvenile Oyster Disease (JOD) in Maine. Journal of Shellfish Research 19: 641. (Abstract).

Boardman, C.L., A.P. Maloy and K.J. Boettcher. 2008. Localization of the bacterial agent of juvenile oyster disease (Roseovarius crassostreae) within affected eastern oysters (Crassostrea virginica). Journal of Invertebrate Pathology 97: 150–158.

Boettcher, K.J. and B.J. Barber. 1998. Use of antimicrobials to investigate the etiological agent of JOD in Crassostrea virginica. Journal of Shellfish Research 17: 348. (Abstract).

Boettcher, K.J., B.J. Barber and J.T. Singer. 1999a. Use of antibacterial agents to elucidate the etiology of juvenile oyster disease (JOD) in Crassostrea virginica and numerical dominance of an α-Proteobacterium in JOD-affected animals. Applied and Environmental Microbiology 65: 2534-2539.

Boettcher, K.J., J.T. Singer and B.J. Barber. 1999b. A novel species of alpha-proteobacterium is associated with signs of juvenile oyster disease (JOD) in Crassostrea virginica. Journal of Shellfish Research 18: 295-296. (Abstract).

Boettcher, K.J., B.J. Barber and J.T. Singer. 2000. Additional evidence that juvenile oyster disease is caused by a member of the Roseobacter group and colonization of nonaffected animals by Stappia stellulata -like strains. Applied and Environmental Microbiology 66: 3924-3930.

Boettcher, K.J., K.K. Geaghan, A.P. Maloy and B.J. Barber. 2005. Roseovarius crassostreae sp. nov., a member of the Roseobacter clade and the apparent cause of juvenile oyster disease (JOD) in cultured Eastern oysters. International Journal of Systematic and Evolutionary Microbiology 55: 1531–1537.

Boettcher, K., R. Smolowitz, E.J. Lewis, B. Allam, H. Dickerson, S. Ford, A. Huq, K. Reece, G. Rivara and C.M. Woodley. 2006. Juvenile oyster disease (JOD) in Crassostrea virginica : synthesis of knowledge and recommendations. Journal of Shellfish Research 25: 683-686.

Bricelj, V.M., S.E. Ford, F.J. Borrero, F.O. Perkins, G. Rivara, R.E. Hillman, R.A. Elston and J. Chang. 1992. Unexplained mortalities of hatchery-reared, juvenile oysters, Crassostrea virginica (Gmelin). Journal of Shellfish Research 11: 331-347.

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Maloy, A.P. and K.J. Boettcher. 2003. Roseimarina crassostrea (gen. nov., sp. nov.) associated with JOD-signs in the absence of significant mortalities, and first isolation from a New York epizootic. Journal of Shellfish Research 22: 343. (Abstract).

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Maloy, A.P., B.J. Barber and K.J. Boettcher. 2005. A PCR-based diagnostic assay for the detection of Roseovarius crassostreae in Crassostrea virginica affected by juvenile oyster disease (JOD). Diseases of Aquatic Organisms 67: 155–162.

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Citation Information

Bower, S.M. (2010): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Juvenile Disease of Eastern Oysters.

Date last revised: May 2010
Comments to Susan Bower