Vibrio spp. (Larval and Juvenile Vibriosis) of Oysters
Category 4 (Negligible Regulatory Significance in Canada)
Common, generally accepted names of the organism or disease agent
- Bacillary necrosis, Larval necrosis, Acute pallial infection, Vibriosis.
- Juvenile vibriosis, Chronic extrapallial abscesses of juvenile oysters.
- Juvenile summer mortality.
Scientific name or taxonomic affiliation
- Vibrio tubiashii, Vibrio anguillarum, Vibrio splendidus, Vibrio kanaloae, Vibrio ordali, Vibrio alginolyticus, Vibrio aestuarianus, Vibrio crassostreae, Vibrio gigantis and Vibrio spp. Various species of bacteria in the genera of Pseudomonas and Aeromonas may also be involved in the disease. Some of these bacteria (Vibrio aestuarianus, Vibrio crassostreae, Vibrio gigantis and members of the V. splendidus group) have been associated with disease and mortalities among spat and adult Crassostrea gigas in France. (Faury et al. 2004, Le Roux et al. 2005, Garnier et al. 2007). Another species of unknown pathogenic status, Vibrio lentus was isolated from oysters on the Mediterranean coast of Spain (Macián et al. 2001).
- Vibrio alginolyticus, Vibrio spp. and unidentified rod-shaped bacteria observed in extrapallial abscesses.
- Vibrio splendidus (various strains).
- In all marine waters where bivalve hatchery and nursery culture is practiced. It is generally a problem only in the warmest months of the year. Vibrio splendidus bivar II was identified as the cause of bacillary necrosis leading to mass mortalities in C. gigas hatcheries in Japan (Sugumar et al. 1998). Vibrio tubiashii was originally isolated from the northeast Atlantic coast of the United States and southeast coast of England and re-emerged as a severe pathogen in shellfish hatcheries on the west coast of North America during 2006 and 2007 (Elston et al. 2008).
- Oyster seed production facilities in Maine, California and Washington, USA.
- Bay of Morlaix in North Brittany and other locations along the Atlantic coast of France (Lacoste et al. 2001, Le Roux et al. 2002).
- Crassostrea virginica, Crassostrea gigas, Crassostrea sikamea, Ostrea edulis, Ostrea conchaphila and other cultured bivalve larvae including clams, and scallops. However, some species of bivalves may be more resistant to the pathogenic effects of these bacteria than other species (Elston et al. 2008).
- Crassostrea virginica, Ostrea edulis, Crassostrea gigas, Crassostrea sikamea and other cultured juvenile molluscs including clams, and abalone.
- Crassostrea gigas spat in the field and in hatcheries.
Impact on the host
Vibriosis is the most commonly encountered disease associated with intensive bivalve culture in hatcheries and nurseries. Infections are initiated by the attachment of bacteria to the external shell surface along the peripheral valvular margin. Attached bacteria form colonies that grow and contact the mantle resulting in necrosis of mantle epithelium and penetration of the bacteria into all soft tissues via the coelomic cavity. During the process, the branchial epithelium may also be infected (Elston 1999). Birkbeck et al. (1987) reported that V. anguillarum (strain 5679) had a particularly high affinity for gill tissues of O. edulis and C. gigas. Systemic infection of the soft-tissues of the larvae and juveniles (spat or seed), result in tissue necrosis (due to production of exotoxin by the bacteria) and death. Two of the low-molecular-weight toxins produced by pathogenic Vibrio spp. include a proteinase (molecular weight =40,000) that degrades connective tissue and at least one ciliostatic toxin (molecular weight = 500-1,000) (Nottage et al. 1989). The signs of infection include the sudden onset, with affected larvae exhibiting reduced feeding rate, and erratic swimming behaviour probably due to the toxin-induced velar damage caused by the bacteria.
Oyster larvae seem to be incapable of repairing damage to the mantle during the early stages of infection. The capacity for mantle repair seems to increase with the increasing size of the juvenile oysters. Larger spat oysters are capable of sequestering the invading bacteria between successive layers of conchiolin deposition on the inner surface of the shell. This chronic extrapallial abscess disease of juvenile oysters is differentiated from acute pallial infections (that result in rapid overwhelming bacterial infections) by the presence of contained abscesses in the extrapallial space in an infection that does not exceed the capacity of the mantle for repair. Although the bacteria that cause chronic abscess disease can be sequestered by new shell deposits and the infection resolved, the condition can cause mortalities (when the mantle is breached leading to overwhelming bacteremia) and significant loss of growth in intensively cultured juvenile oysters (Elston 1999, Elston et al. 1999).
Experimental results of Lambert and Nicolas (1998) and Garnier et al. (2007) confirmed that different species and different isolates of the same species of Vibrio vary in their pathogenicity for bivalves. Gay et al. (2004a and b) found that injection of pooled strains into C. gigas spat (4 to 6 cm in shell length) increased virulence and suggested that Vibrio spp. strains may have additive/synergistic action leading to higher mortality rates. Some bacteria (e.g., Vibrio aestuarianus strain 01/32) produce extracellular products (ECPs) which display immunosuppressant activities on haemocyte functions and may play an important role in the pathogenicity of the bacteria (Labreuche et al. 2006). Vibrio tubiashii can produce a toxigenic non-invasive disease in bivalve larvae as well as causing lesions from a classical bacterial invasion of the bivalve larval tissues (Elston et al. 2008). In general, adult bivalves do not suffer high mortalities when experimentally challenged with larval pathogens (Paillard et al. 2004).
Although Vibrio spp. and many other species of bacteria occur in shellfish hatcheries, many (most) are not pathogenic and some may have potential for probiotic applications (Schulze et al. 2006). Adult oysters are known to harbour Vibrio spp. that can cause disease in humans (e.g., Vibrio parahaemolyticus and Vibrio vulnificus) with no apparent adverse effects on the oysters (Genthner et al. 1999, La Peyre et al. 1999, Volety et al. 1999). Adult C. gigas inoculated with bacteria (heat-killed strains of Micrococcus luteus, Vibrio splendidus and Vibrio anguillarum) produced a large and rapid elevation in Interleukin-17 homologue (CgIL-17) cDNA transcript abundance in haemocytes, suggesting that this is a very early response gene to pathogens that may be responsible for the stimulation of other immune genes in the oyster (Roberts et al. 2008).
Note: Definitive diagnosis of the disease as vibriosis or one resulting from other bacteria requires identification of the specific species or strain involved by appropriate biochemical, immunodiagnostic, or molecular methods. However, consistent isolation of numerically dominant bacteria (Gram negative rods) from tissues with characteristic lesions provides a strong presumptive diagnosis.
Wet mount: Extensive bacterial colonies only visible during the late stages of infection. During larvae development, early stages of vibriosis are evident as velar damage characterized by loss of normal texture, deciliation of velar epithelium, and detachment of some of the cells. The digestive glands may be lighter in colour because of reduced feeding and visceral atrophy may be evident. However, these early signs of disease only provide tentative diagnosis because toxins other than those of bacterial origin (e.g., biotoxins or anthropogenic toxins) or other etiologies may cause similar conditions. Histological observation of velar lesions and detachment of velar retractor muscles supports the diagnosis. As noted above, a confirmatory diagnosis with attribution to a particular bacterium requires identification of the specific species involved. Once the larvae are inactive, it is typical to observes swarms of bacteria around the damaged velum, within the shell cavity and apparently in the larval tissues. The extent of mortality in juvenile oysters can be estimated by enumerating empty shells and living oysters with a stereomicroscope.
Histology: Prior to examination, oysters should be fixed whole in Davidson’s fixative containing acetic acid with further decalcification of the shell as needed. For adequate tissue structure preservation, oysters less than 5 mm in shell height should be embedded in a plastic histological medium while larger oysters can be prepared for histological examination using routine paraffin embedding techniques. Indications of tissue necrosis and the presence of rod shaped bacteria (usually slightly curved and Gram negative) attached to or within the tissues of larvae. Usually associated with damage to the velum and detachment of the velar retractor muscles. In juvenile oysters, the bacteria initially attached to externally oriented periostracum and subsequently invading the tissues through the valve closure and along the internal shell surface. Contact necrosis and sloughing of mantle epithelium results and bacteria can invade the coelomic cavity. Moribund larval and juvenile oysters can also be colonized by various marine organisms, especially various species of ciliates. In chronic extrapallial abscess disease, abscesses containing host cells and bacteria occur in the extrapallial space with no signs of other infectious agents.
Culture: Isolation and culture (TCBS bacterial culture agar) colonies of Vibrio from the tissues of diseased oysters. For larvae and small juveniles (seed), the bacterial isolations can be prepared from homogenized whole larvae or seed because of their small size. Isolates from diseased oysters often have a mixture of bacteria from several taxa normally found in seawater. The bacterial composition of the moribund oysters can be highly variable especially if antibiotic treatment was employed. However, plating a dilution series will help identify the numerically dominant species. Two procedures (using either 15 ml conical tubes or tissue culture plates) were developed to rapidly screening bacterial strains for pathogenicity to C. gigas larvae (Estes et al. 2004). However, pathogenicity dose curves generated in the laboratory may not represent the same dose disease relationship in large-scale bivalve hatchery production. Usually, higher concentrations of pathogenic bacteria are required to cause outbreaks in production facilities in contrast to the lower concentrations that caused morbidity and mortality in the laboratory (Elston et al. 2008).
Immunological Assay: Immunofluorescent labeled antibodies specific for Vibrio alginolyticus and Vibrio spp. labeled antisera have been used to identify vibrio in commercial hatchery epizootics (Elston et al. 1981).
Molecular characteristics: Genomic analysis using polymerase chain reaction – restriction fragment length polymorphism (PCR-RFLP) and the nucleotide sequences of various genes (e.g., 16S ribosomal DNA (16S rDNA), small subunit ribosomal DNA (SSU rDNA), rpoA, rpoD, recA, pyrH and gyrase B subunit (gyrB) genes) are being employed to enable the differentiation between pathogenic and non-pathogenic strains and species of some Vibrio spp. (Le Roux et al. 2002, Le Roux et al. 2004, Paillard et al. 2004, Thompson et al. 2005, Garnier et al. 2007). Vibrio tubiashii has specific genes that code for a protease and haemolysin and pathogenic isolates secrete these peptides (Elston et al. 2008).
Bioassay: The pathogenicity of bacteral isolates was assessed by inoculating 24-hr suspensions of the test organism into 400-ml cultures of 2 to 7 day-old bivalve larvae (Tubiash et al. 1965). Vibrio splendidus was detected as the cause of summer mortality in C. gigas in France by focusing on juvenile oysters presenting reduced stress response capacities determined by circulating noradrenaline measurements, a characteristic of juvenile oysters entering an early phase of the disease (Lacoste et al. 2001).
Methods of control
Vibrio bacteria are ubiquitous, hence eradication of the aetiologic agent is impossible. Vibriosis appears to be directly related to poor husbandry and/or poor water quality such as the presence of pesticides, heavy metals, toxic phytoplankton (or their metabolites), or petroleum toxicants in the water supply. Sources of infection are broodstock, algal cultures and incoming seawater. For example, Elston et al. (2008) identified that the losses of larval and juvenile bivalves in shellfish hatcheries were linked to V. tubiashii blooms in the coastal environment which were associated with the mixing of unusually warm surface seawater and intermittent up-welling of cooler nutrient- and Vibrio spp. enriched seawater. To initiate methods of control, determine the source of infection by culturing bacteria from various candidates. Vibrio spp. can be harboured on system surfaces and their growth can be augmented by dissolved organic substrates generated by algal cultures, external algal blooms, or the metabolites of the cultured bivalves. Elston et al. (2008) identified several areas of persistent contamination with algal culture contamination being the most insidious because several algal species, often used in production hatcheries, can coexist in the presence of bacteria pathogenic to bivalves without apparent effects on algal growth rate or maximum density. Prevention and control strategies must include routine sanitation of system surfaces, water filtration, broodstock sanitation and maintenance of low dissolved organic levels. Temperature control may also play an important role in managing some of the bacterial diseases of oyster larvae and juveniles (seed). Elston et al. (2008) presented various techniques for the management and prevention of serious bacterial contamination in shellfish hatcheries.
Batches containing infected larvae should be destroyed in an approved manner; disinfect all containers and equipment in contact with the infected stock. In a nursery, disease can be avoided or mortalities reduced by disinfecting the external surfaces of oyster seed with freshwater rinses and/or sodium hypochlorite baths (concentrations ranging from 10 to 25 ppm for a duration of 3 to 30 minutes depending on size of seed (lower concentrations and shorter duration for smaller seed) and intensity of infection and with repetition of treatment as needed). Antimicrobial agents to reduce bacterial populations in and around the bivalve molluscs for the control and treatment of disease in mollusc hatcheries have been assessed (Tubiash et al. 1965, Le Pennec and Prieur 1977). However, the use of inhibitory compounds may lead to the rapid development of resistant pathogen populations, the elimination of beneficial organisms and the emergence of other microbial pathogens of the bivalves. Research is underway to select and test strains of bacteria for use as probiotics (Elston et al. 2000)
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Bower, S.M. (2009): Synopsis of Infectious Diseases and Parasites of Commercially Exploited Shellfish: Vibrio spp. (Larval and Juvenile Vibriosis) of Oysters.
Date last revised: December 2009
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