Printer Friendly

Depuration of bacterial populations in the Indian backwater oyster Crassostrea madrasensis (Preston, 1916): effects on surface and bottom held oysters.

ABSTRACT The efficiency of depuration of the Indian backwater oyster Crassostrea madrasensis (Preston, 1916) using the fill-draw method (static method) with high-loading density was evaluated in this study. Depuration experiments were conducted with cartridge-filtered and UV-treated seawater at a salinity of 30.3% [per thousand], a pH of 8.3, and a temperature of 29.5[degrees]C. The oysters located in trays on the surface and on the bottom were compared for microbial loads. Samples were taken at 0 h, 8 h, 16 h, 24 h, 36 h, and 48 h of depuration. The results showed that in winter monsoon-sampled nondepurated oysters, the most probable number of fecal coliforms and Escherichia coli were greater than the limits according to NSSP and European Union regulations. The surface held oysters took 24 h to reduce the coliforms and E. coli levels to below safe limits whereas for bottom held oysters it took 48 h. The species Salmonella was never detected in the oysters sampled, whereas Vibrio spp. were present in the nondepurated oysters and were eliminated completely after 8 h of depuration. Variation in depuration of total coliforms, fecal coliforms, E. coli, total plate count, and fecal streptococci in oysters were significant (P < 0.05) between surface and bottom oysters. The study results recommend a loading density of 2 oysters/L water stacked in 1 layer as the optimum loading density for commercial depuration completed within 24 h.

KEY WORDS: Crassostrea madrasensis, depuration, oysters, bacterial population, E. coli


Farmed bivalve production in the world has increased continuously and substantially during the past century, increasing from approximately 1 million t in 1950 to 13.9 million t in 2010. China leads in production, followed by Korea and Japan. Currently, India ranks 13th in oyster production and has shown a steady increase for the past 2 decades (FAO 2012). The annual production of farmed oysters was 1,500 t in 2009/2010 and it increased to 2,500 t in 2010/2011 and to 3,200 t in 2011/2012 (Mohamed & Kripa 2013). The oyster Crassostrea madrasensis (Preston, 1916) is the main farmed species, which occurs extensively in brackish waters of both the east and west coasts of India. This environment, however, is also congenial for the growth of coliforms and pathogenic bacteria that pose a problem for human health. Molluscan shellfish, particularly oysters, concentrate microorganisms in their tissue from surrounding water during their filter-feeding process and are recognized as a reservoir for various microbial pathogens (Panicker et al. 2004). Naturally occurring pathogenic microorganisms such as Vibrio spp. and fecal pathogens such as Salmonella spp. inhabit marine and estuarine environments (Lee et al. 2003). Infections arise as a result of the consumption of raw, undercooked, or improperly processed oysters, which frequently cause outbreaks of gastroenteritis and septicemia (Sobsey & Jaykus 1991, Lees 2000, Koopmans et al. 2002, Le Guyader et al. 2003).

Since the 1970s, the U.S. Food and Drug Administration (FDA) has required the shellfish industry to use fecal coliforms as indicators of contamination in harvesting waters and oysters (Brands et al. 2005). The FDA requires each U.S. state to test harvesting waters 6 times/y for the presence of fecal coliforms. If the fecal coliforms are detected above the most probable number (MPN) of 230/100 g of oyster or 230/100 ml water sample, then the waters are closed for harvesting (FDA 2009). According to European Union (EU) regulations, for molluscs, in addition to the absence of Vibrio spp. and Salmonella spp., a maximum tolerable cell concentration is required both for fecal coliforms (300 MPN/100 g meat) and Escherichia coli (230 MPN/100 g meat) (EC 2007, Maffei et al. 2009). Washing and surface disinfection of shellfish may be effective in reducing postharvest contamination, but most of the microorganism outbreaks associated with shellfish are from preharvest sources of contamination, when the microorganisms are bioaccumulated (Richards et al. 2010).

Depuration is a process during which shellfish are held in tanks with clean seawater and are allowed to resume their natural filter-feeding activity, purging themselves of contaminants (Blogoslawski & Stewart 1983, Jackson & Ogburn 1999). This process was developed initially in response to a number of outbreaks of shellfish-associated typhoid illness (Richards 1988, Jones et al. 1991). Although depuration is not yet an established practice in India, it is used in many other developed countries such as the United Kingdom, France, Italy, the United States, and others, depending on the microbial quality of shellfish-growing waters and load of microbes in tissues. In India, there are no reports of such a classification for oyster-growing areas and also there is very limited scientific information available regarding bivalve depuration. This is probably a result of the low commercial production of bivalves and poor demand in markets until recently. Moreover, a live oyster (Crassostrea madrasensis) supply to high-end restaurants has been initiated lately, and a new value chain has been created (Mohamed & Kripa 2013).

The technology of oyster depuration has been well studied in temperate waters (Kelly 1961, Love et al. 2010), and is reviewed by Oliveira et al. (2011). Most of the studies focused on testing depuration in different types of water (Vasconcelos & Lee 1972, Kasai et al. 2011, Ramos et al. 2012), and at different temperatures (Chae et al. 2009, Love et al. 2010, Lopez-Joven et al. 2011) salinities (Rowse & Fleet 1984, Power & Collins 1990, Love et al. 2010), and densities (Lee 2010). However, there is little information on tropical oysters (Pillai & Selvan 1987) and no information on the effect of high-density stocking on depuration of tropical bivalves.

The current study investigates the reduction over time of bacterial loads in the farm-grown oyster Crassostrea madrasensis, during depuration specifically for total coliforms, fecal coliforms, Escherichia coli, fecal streptococci, Vibrio spp., and Salmonella spp. The efficiency of depuration of oysters placed in trays on the surface and on the bottom of the depuration tanks was compared.


Sampling Site and Sample Collection

The edible oyster Crassostrea madrasensis was harvested from a commercial oyster farm in Ashtamudi Lake (Dalavapuram) Kollam, Kerala, a brackish water lake located on the southwest coast of India (8[degrees]56' 16.31" N, 76[degrees]33'18.13" E). For experiments, approximately 1,000 2-y-old oysters were collected during the winter monsoon season of November and December. Samples of similar size and weight were selected to minimize differences. Morphometric measurements were taken at random for 20 oysters. The average length, width, depth, total weight, and meat weight of oysters used in the study was 86.70 [+ or -] 25.71 mm, 57.21 [+ or -] 10.02 mm, 29.60 [+ or -] 6.67 mm, 99.2 [+ or -] 46.97 g, and 9.2 [+ or -] 1.14 g, respectively. After harvesting, an initial sample of oyster and surface water was taken from the oyster farm and transported aseptically to the laboratory in an insulated icebox. On arrival at the laboratory, oyster and water samples were processed for the detection of total coliforms, fecal coliforms, Escherichia coli, fecal streptococci, Salmonella spp., Vibrio spp., and total plate counts (TPC) (Lalitha & Surendran 2004). The remaining organisms were transported to the depuration unit for additional experiments. Hydrographic parameters such as salinity, temperature, and pH of the water at the farm site and in the hatchery were checked.

Depuration Experiments

The fill-and-draw (static method) depuration method was followed (Lee et al. 2008). Depuration was conducted in 250-L Fiberglas tanks (1.25 x 0.75 x 0.75 m, 200 L seawater). The seawater was settled and then cartridge-filtered serially (10 pm, 5 [micro]m, 2 [micro]m, 0.2 [micro]m R.A-Mumbai) and UV (MOC: SS-313, UV intensity 900 [micro]W/min/[cm.sup.2]). Before starting depuration, attached barnacles and other fouling organisms were removed from the oysters by careful scrubbing and water jetting. Since the seawater from the sampling site and that in the experimental tanks was similar in physical-chemical parameters, acclimation of oysters was not done. For the experiment, 400 oysters were loaded in 4 perforated, plastic rectangular trays (50 x 30 x 15 cm) at a rate of 100 oysters per tray. Approximately 3 layers of oysters were kept in a tray. The density in the tank was 2 oysters/L seawater. The oysters in trays were elevated (by 5 cm) from the bottom of the tank to restrict recontamination with feces and pseudofeces. It has been established that the density of animals in the depuration tank influences the depuration process (Lee 2010), and therefore an alternate depuration tank was also kept in the same condition with same number of oysters for depuration. For every sampling, 10 oysters were taken for microbial analysis from tank 1, and the same number of the oysters was replaced from the alternate tank to maintain the density of oysters in depuration tank 1. The seawater of the depuration tank was completely changed and replaced with fresh seawater at 8 h, 16 h, 24 h, 36 h, and 48 h, and the depuration tank and trays were also cleaned with seawater to remove all the waste expelled by the oysters. Before every water change, water samples were drawn from the depuration tank, and oyster samples were taken from the surface and bottom layer of the tray for microbial analysis. The experiment was closed after 48 h.

Microbiological A n a lysis

Enumeration of Coliforms and Escherichia coli

Bacterial contamination in oysters was determined before and during depuration at 8 h, 16 h, 24 h, 36 h, and 48 h. Oysters were opened aseptically in a laminar flow cabinet with flame blades. As per the method given by Almeida and Soares (2012), soft tissues and intervalve liquids (25 g) were weighed and placed in sterile bags containing enough saline solution to make a 1:10 dilution, and were then blended for 2 min in a stomacher. The concentration of total coliforms, fecal coliforms, and Escherichia coli was determined by the MPN procedure using 5 tubes per dilution of 10 ml, 1 ml, and 0.1 ml (Almeida & Soares 2012). The total coliforms, fecal coliforms, and E. coli were enumerated using Lauryl Tryptose Broth (Himedia, India). The presence of total coliforms was confirmed by gas production in brilliant green bile broth (Himedia) culture within 48 [+ or -] 3 h at 35 [+ or -] 0.5[degrees]C. Gas production and growth in E. coli broth (EC broth: Himedia) culture within 24 [+ or -] 2 h was considered positive for fecal coliforms; E. coli was confirmed positive in EC-MUG tubes and indole production in Tryptose broth (Lalitha & Surendran 2004, American Public Health Association 2012).

Enumeration of TPC and Fecal Streptococci, and Qualitative Analysis of Salmonella and Vibrio

Total plate count was determined by using plate count agar (Himedia). Sample dilution of [10.sup.-1], [10.sup.-2], and [10.sup.-3] with buffered peptone water were taken. After inoculation, the plates were incubated for 24 h at 36 [+ or -] 1[degrees]C. Colonies were counted and data reported as colony-forming units per gram (Sengor et al. 2004, Obadai et al. 2010). Fecal streptococci were determined by using KF Streptococcus agar (Himedia). The plates were incubated at 36 [+ or -] 1[degrees]C for 48 h, and dark-red colonies and colonies with red and pink centers were counted as fecal streptococci colonies. The results are expressed in number of colony-forming units per milliliter or grams (American Public Health Association 1970).

The presence of Salmonella spp. was determined by the U.S. FDA bacteriological analytical method using RV medium and TT broth (Himedia) (Andrews & Hammack 2001). Isolation of Vibrio spp. was performed using the FDA BAM method (Elliot et al. 2001). Enriched peptone water after incubation was streaked onto TCBS agar (Himedia) plates that were incubated for 18-24 h at 35-37[degrees]C. The typical green and yellow colonies present in the TCBS agar were confirmed as recommended by the FDA BAM method.

Statistical Analysis

All MPN values for total coliforms, fecal coliforms, Escherichia coli, TPC (in colony-forming units per gram), and fecal streptococci (in colony-forming units per gram) were converted to [log.sub.10] values before analysis with 2-way ANOVA using SPSS version 16.0. Because the interaction between type and time was significant, the interaction means were compared using Student's t-test. Differences between treatment means were considered significant at P < 0.05. Standard deviations of the 3 replicates of each sample were reported as bars in figures.


Environmental Parameters

The salinity, temperature, and pH of the farm site were 30.5% [per thousand], 29.5[degrees]C, and 8.3, respectively. In the hatchery, salinity, temperature, and pH were 32.5% [per thousand], 30.3[degrees]C, and 8.1, respectively.

Occurrence of Microorganisms in Oysters Collected from the Farm

In the oyster samples prior to depuration, the [log.sub.10] MPN of total coliforms and fecal coliforms was 4.16 MPN/100 g and 4.06 MPN/100 g, and Escherichia coli was 3.88 MPN/100 g. The [log.sub.10] TPC was 4.23 CFU/g and fecal streptococci was 1.46 CFU/g. Prior to the start of the depuration process, the oyster MPN value for total coliforms, fecal coliforms was more than 18,000 MPN/100 g, and E. coli was 9,100 MPN/100 g. According to the FDA (NSSP) regulations, the MPN value of Ashtamudi Lake oyster and oyster-growing water comes under restricted area when the waters are subject to limited amounts of pollution such that shellfish must be depurated or relayed prior to sale.

Changes in Coliform Levels in Surface and Bottom Oysters at Different Time Intervals

The total and fecal coliform levels in surface held oysters decreased from [log.sub.10] 4.16 MPN/100 g to [log.sub.10] 3.45 MPN/100 g within 8 h of depuration (Figs. 1A-B), but the bottom held oysters showed no reduction in the coliform counts until after 8 h of depuration. Similarly the Escherichia coli level in the surface oysters decreased from [log.sub.10] 3.88 MPN/100 g to [log.sub.10] 3.22 MPN/100 g (Fig. 1C, but there was no change in the bottom held oysters within 8 h of depuration. The surface held oysters had fecal coliform levels and E. coli counts less than the limit after 24 h of depuration; the bottom held oysters took 48 h of depuration to reduce the coliform levels to less than the limit. After 48 h of depuration, the coliform counts were less than 20 MPN/100 g in both the surface- and bottom held oysters. Results (Table 1) revealed that, in the case of oysters held at the surface, total coliforms, fecal coliforms, and E. coli levels reduced significantly (P < 0.05) after 8 h, 16 h, and 24 h of depuration, but additional reductions at 36 h and 48 h were not significant (P > 0.05). In the case of bottom held oysters, a significant (P < 0.05) reduction in total coliforms and fecal coliforms was observed only after 24 h of depuration, and an additional significant reduction was seen only at 48 h of depuration. In the case of E. coli levels of bottom held oysters, the counts were reduced significantly (P < 0.05) at 24 h and reduced even more at 48 h of deputation (Table 1).

Changes in TPC and Fecal Streptococci Counts of Oysters During Depuration

The TPC of nondepurated oysters was [log.sub.10] 4.23 CFU/g, and it was reduced to logic 3.87 CFU/g after 8 h of depuration in surface held oysters. After 8 h of depuration, TPC decreased rapidly, from [log.sub.10] 3.87 CFU/g to [log.sub.10] 3.22 CFU/g after 16 h of depuration. At the end of the depuration experiment (48 h of depuration), the TPC was logic 2.27 CFU/g. However, after 8 h of depuration, bottom held oysters showed a slower depuration rate. The TPC was reduced from logic 4.23 CFU/g to [log.sub.10] 3.16 CFU/g (Table 1, Fig. 1D) after 48 h of depuration. Analysis of variance showed a significant (P < 0.05) difference in TPC between the surface and bottom oysters, and there were no significant interaction differences (t-test, P = 0.426). The fecal streptococci reduced to [log.sub.10] 0.99 CFU/g in surface oysters and logic 1-36 CFU/g in bottom oysters from [log.sub.10]1.46 CFU/g at the end of 8 h of depuration. Furthermore, after 16 h of depuration, the fecal streptococci in surface oysters reduced to [log.sub.10] 0.81CFU/g; in the bottom oysters, it reduced to logic 1-30 CFU/g. No fecal streptococci was detected after 24 h of depuration in surface oysters, but in bottom oysters it was present ([log.sub.10] 0.36 CFU/g; Fig. IE) until the end of 48 h of depuration. Results showed that, in the case of oysters held at the surface, the fecal streptococci level reduced significantly (P < 0.05) at 24 h; there were no significant (P > 0.05) reductions at 36 h and 48 h. In the case of bottom held oysters, there was no significant (P > 0.05) reduction after 8 h, 16 h, 24 h, and 36 h of depuration; a significant (P < 0.05) reduction was seen only after 48 h of depuration (Table 1).

Counts of Vibrio and Salmonella

Species of Salmonella were not detected in analyzed oysters or in water samples. Species of Vibrio were found in the oyster as well as in the water sample before depuration, and were eliminated completely after 8 h of depuration.


The oyster samples used in this experiment were from a commercial oyster farming area. In oysters, the level of fecal bacteria (Escherichia coli and fecal coliforms) was greater than safe limits as stipulated by the NSSP and EU regulations. The oysters were sampled during the winter monsoon, when harvest from oyster beds is greatest, and the high level of fecal coliforms could be the result of feral wastes brought by river runoff. Similar results were reported for mussels along the west coast of India, where high levels of fecal coliforms were present only during the monsoon season in the Arabian Sea (Sasikumar & Krishnamoorthy 2010). They suggested depuration of mussels before consumption during the monsoon season.

It is considered that the period of depuration is 1 of the limiting factors that is effective in reducing bacterial loads (Son & Fleet 1980, Yang 2008, Lopez-Joven et al. 2011). Another important factor in the depuration process is the placement (surface or bottom) of the oysters within the depuration tank, for which there are no previous reports. In the current study, using a static method, within 24 h oysters placed in trays near the surface of the depuration tank reduced the level of fecal coliforms to less than 300 MPN/100 g and reduced the level of Escherichia coli to less than 230 MPN/100 g, which are the threshold levels of the NSSP and EC, respectively (EC 2007. FDA 2009). Vasconcelos and Lee (1972) showed that depuration with UV-treated seawater could reduce fecal coliforms and E. coli counts below the limit after 24 h of depuration in Pacific oysters. However, the current study shows that 24 h of depuration was insufficient for oysters placed in the bottom of the trays. It took 48 h to reduce the fecal coliforms to less than 300 MPN/100 g, and E. coli to less than 230 MPN/g. Oysters held at the surface showed 2-phase depuration dynamics, with faster depuration rates during the first 24 h and a slower phase during the last 24 h for fecal coliforms, E. coli, and TPC. The bottom held oysters also showed a 2-phase depuration rate except for E. coli, for which the curve appeared smooth from 0-48 h. Many researchers (Power & Collins 1990, Dore & Lees 1995, Mcghee et al. 2008, Love et al. 2010) who used flow-through systems have reported that a 2-phase depuration is a result of differential lysosomal digestion. There are very clear indications that fecal coliforms--more specifically, E. coli--are not retained selectively in the gut and are eliminated readily with oyster feces during depuration of temperate oyster species such as Crassostrea virginica, Crassostrea gigas, Crassostrea commercialis, and Ostrea edulis (Souness & Fleet 1979, Son & Fleet 1980, Dore & Lees 1995, Love et al. 2010, Kasai et al. 2011). The current study also shows that, in the case of the tropical oyster species Crassostrea madrasensis, E. coli are eliminated rapidly from the oyster gut.

The time required for purification of polluted oysters depends on the initial level of contamination (Son & Fleet 1980). In general, bacteria are reduced rapidly to undetectable levels within 48 h of depuration (Dore & Lees 1995). Son and Fleet (1980), who studied depuration of Crassostrea commercialis in Australia, reported that the reduction of Escherichia coli numbers from 100 cells/g to undetectable levels took place within 48 h. Similarly, Rowse and Fleet (1984) showed that Salmonella charity and E. coli were virtually undetectable after 38 h of depuration in C. commercialis. Correa et al. (2007) reported that the combination of UV irradiation and chlorine treatments for 12 h eliminated all bacteria in Crassostrea gigas in Brazil, and Kasai et al. (2011) reported that in the Japanese oyster C. gigas, naturally occurring E. coli counts were reduced significantly to less than the detection limit (30 E. coli MPN/100 g) after depuration with UV-treated seawater for 24 h. Similarly, the current study shows that surface held oysters required only 24 h for complete depuration. However, in the case of bottom held oysters, 48 h was needed to reduce fecal streptococci and E. coli to less than the threshold. Earlier studies have not examined the depuration time with respect to density of animals in the tank (Chae et al. 2009, Love et al. 2010, Kasai et al. 2011, Phuvasate et al. 2012). The siphoning activity and filtration capacity of oyster is size dependant and this, too, can affect the time taken for depuration (Jones et al. 1991, Oliveira et al. 2011). In the current study, 2-y-old oysters (length, 60-110 mm) were used, and these are the normal sizes at which oysters are harvested. Any change in size of oysters used for depuration can affect the results obtained in the current study.

The natural occurrence of Vibrio spp. in nondepurated oysters, although at low concentration, might be the result of prevailing water temperatures. Several studies have demonstrated a positive correlation between water temperature and occurrence of Vibrio spp. (Deepanjali et al. 2005, Chae et al. 2009, Ramos et al. 2012). The current study shows that the depuration of oysters not only reduces the levels of fecal coliforms and Escherichia coli, but also it eliminates human pathogens such as Vibrio spp. and Salmonella spp. to below detectable levels. Therefore, these fecal bacteria are good indicators of the bacteriological quality of depurated oysters. The current study shows that depuration reduced bacterial loads significantly in the oysters. Elimination of bacterial loads in the oysters was faster in surface held oysters compared with bottom held oysters. It was obvious that the bottom oysters did not function with the same filtration efficiency as the surface oysters.


The results obtained from the current experiment show that depuration of the oyster Crassostrea madrasensis can reduce the microbial load considerably, and it varies significantly between the oysters located in trays on the surface and on the bottom. The depuration rate was faster in the surface held oysters compared with the bottom held oysters. As per the current protocol, 24 h of depuration is needed for surface held oysters to reduce fecal coliform levels and Escherichia coli counts to 300 MPN/100 g and 230 MPN/100 g, which are the NSSP and EU standards, respectively. The bottom held oysters require 48 h of depuration to reduce fecal coliforms and E. coli counts to acceptable levels. We recommend a loading density of 2 oysters/L water stacked in 1 layer as the optimum loading density for commercial depuration.


This study was funded by National Agricultural Innovation Project (NAIP) of the World Bank (P. Code 2000035102), which we gratefully acknowledge. We are also grateful to the director of the Central Marine Fisheries Research Institute (CMFRI), Kochi, for facilities and encouragement. We thank Dr. T. V. Sathianandan for guidance on statistical analysis. We are also much indebted to 2 anonymous reviewers whose comments greatly improved the manuscript.


Almeida, C. & F. Soares. 2012. Microbiological monitoring of bivalves from Ria Formosa Lagoon (south coast of Portugal): a 20 years sanitary survey. Mar. Pollut. Bull. 64:252-262.

American Public Health Association. 1970. Recommended procedures for the examination of seawater and shellfish, 411' edition. Washington, DC: American Public Health Association. 874 pp.

American Public Health Association. 2012. Recommended procedures for the examination of seawater and shellfish, 21st edition. Washington, DC: American Public Health Association, Washington. 1496 pp.

Andrews, W. H. & T. S. Hammack. 2001. Salmonella. In: FDA bacteriological analytical manual. AOAC International.

Blogoslawski, W. J. & M. E. Stewart. 1983. Depuration and public health. J. World Maricult. Soc. 14:535-545.

Brands, A. D., A. E. Inman, C. P. Gerba, C. J. Mare, S. J. Billington, L. A. Saif. J. F. Levine & L. A. Joens. 2005. Prevalence of Salmonella spp. in oysters in the United States. Appl. Environ. Microbiol. 71:893-897.

Chae, M. J., D. Cheney & Y. C. Su. 2009. Temperature effects on the depuration of Vibrio parahaemolyticus from the American oyster (Crassostrea virginica). J. Food Sci. 74:62-66.

Correa, A. A., J. D. Albarnaz, V. Moresco, C. R. Poli, A. L. Teixeira, C. M. Simoes & C. R. Barardi. 2007. Depuration dynamics of oysters (Crassostrea gigas) artificially contaminated by Salmonella enterica serovar Typhimurium. Mar. Environ. Res. 63:479-489.

Deepanjali, A., H. Sanathkumaur, I. Karunasagar & I. Karunasagar. 2005. Seasonal variation in abundance of total and pathogenic Vibrio parahaemolyticus bacteria in oysters along the southwest coast of India. Appl. Environ. Microbiol. 70:3575-3580.

Dore, W. J. & D. N. Lees. 1995. Behaviour of Escherichia coli and male-specific bacteriophage in environmentally contaminated bivalve molluscs before and after depuration. Appl. Environ. Microbiol. 61:2830-2834.

EC. 2007. (Commission regulation) no. 1441/2007 of December 2007 amending regulation (EC) no. 2073/2005 on microbiological criteria for foodstuffs. Official Journal of the European Union. L322, 12-29.

Elliot, E. L., C. A. Kaysner, L. Jackson & M. L. Tamplin. 2001. Vibrio cholera, Vibrio parahaemolyticus, Vibrio vulnificus and other Vibrio spp. In: FDA bacteriological analytical manual. AOAC International.

FAO. 2012. FishStat: universal software for fishery statistical time series. Rome: FAO. 218 pp.

FDA. 2009. National Shellfish Sanitation Program guide for the control of molluscan shellfish 2007 revisions. Available at http://

Jackson, K. L. & D. M. Ogburn. 1999. Review of depuration and its role in shellfish quality assurance. FRDC project no. 96/335. NSW Fisheries final report series no. 13. ISSN 1440-3544. Pyrmont, Australia. 63 pp.

Jones, S. H., T. L. Howell &K. R. O'Neil. 1991. Differential elimination of indicator bacteria and pathogenic Vibrio spp. from eastern oysters (Crassostrea virginica Gmelin, 1971) in a commercial controlled purification facility in Maine. J. Shellfish Res. 10:105-112.

Kasai, H., K. Kawana, M. Labaiden, K. Namba & M. Yoshimu. 2011. Elimination of Escherichia coli from oysters using electrolysed seawater. Aquaculture 319:315-318.

Kelly, C. B. 1961. Disinfection of seawater by UV radiation. Am. J. Public Health 51:1670-1680.

Koopmans, M., C.- H. von Bonsdorff, J. Vinje, D. de Medici & S. Monroe. 2002. Food borne viruses. FEMS Microbiol. Rev. 26:187-205.

Lalitha, K. V. & P. K. Surendran. 2004. Bacterial microflora associated with farmed freshwater prawn Macrobrachium rosenbergii (de Man) and the aquaculture environment. Aqua. Res. 35:629-635.

Lee, R. 2010. CEFAS protocol for inspection and approval of purification (depuration) systems: Food Standard Agency, Weymouth, England and Wales. 41 pp.

Lee, R., A. Lovatelli & L. Ababouch. 2008. Bivalve depuration: fundamental and practical aspects. FAO Fisheries technical paper 511. Rome: FAO. 139 pp.

Lee, C. Y., G. Panicker & A. K. Bej. 2003. Detection of pathogenic bacteria in shellfish using multiplex PCR followed by CovaLink[TM] NH microwell plate sandwich hybridization. J. Microbiol. Methods 53:199-209.

Lees, D. 2000. Viruses and bivalve shellfish. Int. J. Food Microbiol. 59:81-116.

Le Guyader, F. S., F. H. Neill, E. Dubois, F. Bon, F. Loisy, E. Kohli, M. Pommepuy & R. L. Atmar. 2003. A semi-quantitative approach to estimate Norwalk-like virus contamination of oysters implicated in an outbreak. Int. J. Food Microbiol. 87:107-112.

Lopez-Joven, C., I. Ruiz-Zarzuela, I. de Blas, M. D. Furones & A. Roque. 2011. Persistence of sucrose fermenting and non-fermenting Vibrios in tissues of manila clam species, Ruditapes philippinarium, depurated in seawater at two different temperatures. Food Microbiol. 28:951-956.

Love, D. C., G. L. Lovelace & M. D. Sobsey. 2010. Removal of Escherichia coli, Enterococcus fecalis, coliphages MS2, poliovirus and hepatitis A virus from oysters (Crassostrea virginica) and hard shell clam (Mercinaria mercinaria) by depuration. Int. J. Food Microbiol. 143:211-217.

Maffei, M., P. Vernocchi, R. Lanciotti, M. E. Guerzoni, N. Belletti & F. Gardini. 2009. Depuration of stripped Venus clam (Chamelea gallina L.): effects on microorganisms, sand content and mortality. J. Food Sci. 74:1-7.

Mcghee. T. J. J. A. Morris, R. T. Noble & P. K. Flower. 2008. Comparative microbial dynamics in Crassostrea virginica (Gmelin, 1751) and Crassostrea ariakensis (Fujita, 1913). J. Shellfish Res. 27:559-565.

Mohamed, K. S. & V. Kripa. 2013. Oyster farming: new hope for increasing mariculture production in India. MPEDA Newsl. 22:55-57.

Obadai, E. A., H. D. Nyarko & S. L. Amponsah. 2010. Effect of depuration on microbial content of mangrove oyster (Crassostrea tulipa) from Benya lagoon, Ghana. Ethiopian J. Environ. Stud. Manage. 3:47-53.

Oliveira, J., A. Cunha, F. Castilho, J. L. Romalde& M. J. Pereira. 2011. Microbial contaminants and purification of bivalve shellfish: crucial aspects in monitoring and future perspectives: a mini-review. FoodContr. 22:805-816.

Panicker, G., D. R. Call, M. J. Krug & A. K. Bej. 2004. Detection of pathogenic Vibrio spp. in shellfish using multiplex PCR and DNA microarrays. Appl. Environ. Microbiol. 70:7436-7444.

Phuvasate, S., M. Chen & Y. Su. 2012. Reduction of Vibrio parahaemolyticus in Pacific oyster (Crassostrea gigas) by depuration at various temperatures. Food Microbiol. 31:51-56.

Pillai, K. V. & K. Selvan. 1987. Study on bacterial quality on edible oyster Crassostrea madrasensis and its purification. Bull. Cent. Mar. Fish. Res. Inst. 42:426-431.

Power, U. F. & J. K. Collins. 1990. Tissue distribution of a coliphages and Escherichia coli in mussels after contamination and depuration. Appl. Environ. Microbiol. 56:803-807.

Ramos, R. J., M. Miotto, F. J. Squella, A. Cirolini & J. F. Ferreira. 2012. Depuration of oysters (Crassostrea gigas) contaminated with Vibrio parahaemolyticus with UV light and chlorinated seawater. J. Food Prot. 75:1501-1506.

Richards, G. P. 1988. Microbial purification of shellfish: a review of depuration and relaying. J. Food Prot. 51:218-251.

Richards, G. P., C. Mcleod & S. Le Guyader. 2010. Processing strategies to inactivate enteric viruses in shellfish. Food Environ. Virol. 2:183-193.

Rowse, A. J. & G. H. Fleet. 1984. Effects of water temperature and salinity on elimination of Salmonella charity and Escherichia coli from Sydney rock oysters (Crassostrea commercialis). Appl. Environ. Microbiol. 48:1061-1063.

Sasikumar, G. & M. Krishnamoorthy. 2010. Faecal indicators and sanitary water quality of shellfish harvesting environment influences of seasonal monsoon and river runoff. Indian J. Geo. Mar. Sci. 39:434-444.

Sengor, G. F., H. Kalafatoglu & H. Gun. 2004. The determination of microbial flora, water activity and chemical analysis in smoked, canned mussels (Mytilus galloprovincialis, L.). Turk. J. Vet. Anim. Sci. 28:793-797.

Sobsey, M. & L. Jaykus. 1991. Human enteric virus and depuration of bivalve molluscs. In: W. S. Otwell, G. E. Rodrick & R. E. Martin, editors. Molluscan shellfish depuration. Boca Raton, FL: CRC Press, pp. 71-114.

Son, N. T. & G. H. Fleet. 1980. Behaviour of pathogenic bacteria in oyster Crassostrea commercialis, during depuration, re-laying and storage. Appl. Environ. Microbiol. 40:994-1002.

Souness, R. A. & G. H. Fleet. 1979. Depuration of the Sydney rock oyster, Crassostrea commercialis. Food Technol. Austr. 31:397-404.

Vasconcelos, G. J. & J. S. Lee. 1972. Microbial flora of pacific oysters (Crassostrea gigas) subjected to ultraviolet-irradiated seawater. Appl. Microbiol. 23:11-16.

Yang, Q. 2008. Refrigerated seawater depuration for reducing Vibrio parahaemolyticus contamination in raw oysters (Crassostrea gigas). MS thesis. Oregon State University. 96 pp.


Molluscan Fisheries Division, Central Marine Fisheries Research Institute, PB No. 1603, Kochi, Kerala State. 682018 India

* Corresponding author. E-mail:

DOI: 10.2983/035.033.0209


Levels of total coliforms (TC), fecal coliforms (FC),
Escherichia coli (E. coli), total plate count (TPC), and
fecal streptococci (FS) for surface-layer and bottom held
oysters at different depuration times.

Sample    Time          TC (MPN/g)               FC (MPN/g)

Surface   0       4.16 [+ or -] 0.17 (a)   4.06 [+ or -] 0.17 (a)
          8       3.45 [+ or -] 0.23 (b)   3.29 [+ or -] 0.04 (b)
          16      2.97 [+ or -] 0.30 (c)   2.85 [+ or -] 0.41 (c)
          24      2.18 [+ or -] 0.03 (d)   1.84 [+ or -] 0.10 (d)
          36      2.18 [+ or -] 0.03 (d)   1.81 [+ or -] 0.23 (d)
          48      1.88 [+ or -] 0.05 (d)   1.32 [+ or -] 0.26 (e)
Bottom    0       4.16 [+ or -] 0.17 (a)   4.06 [+ or -] 0.17 (a)
          8       4.16 [+ or -] 0.17 (a)   4.06 [+ or -] 0.17 (a)
          16      4.06 [+ or -] 0.17 (a)   3.88 [+ or -] 0.13 (a)
          24      3.59 [+ or -] 0.23 (b)   3.34 [+ or -] 0.03 (b)
          36      3.49 [+ or -] 0.20 (b)   3.34 [+ or -] 0.03 (b)
          48      2.05 [+ or -] 0.09 (c)   1.58 [+ or -] 0.03 (c)

Sample    Time       E. coli (MPN/g)          TPC (CFU/g)

Surface   0       3.88 [+ or -] 0.13 (a)   4.23 [+ or -] 0.12
          8       3.22 [+ or -] 0.02 (b)   3.87 [+ or -] 0.21
          16      2.25 [+ or -] 0.24 (c)   3.22 [+ or -] 0.63
          24      1.60 [+ or -] 0.20 (d)   3.00 [+ or -] 0.10
          36      1.30 [+ or -] 0.30 (d)   2.76 [+ or -] 0.11
          48      1.26 [+ or -] 0.17 (d)   2.27 [+ or -] 0.34
Bottom    0       3.88 [+ or -] 0.13 (a)   4.23 [+ or -] 0.12
          8       3.88 [+ or -] 0.13 (a)   4.12 [+ or -] 0.03
          16      3.34 [+ or -] 0.17 (b)   3.70 [+ or -] 0.62
          24      2.84 [+ or -] 0.25 (c)   3.47 [+ or -] 0.49
          36      2.71 [+ or -] 0.60 (c)   3.33 [+ or -] 0.56
          48      1.58 [+ or -] 0.03 (d)   3.16 [+ or -] 0.18

Sample    Time          FS (CFU/g)

Surface   0       1.46 [+ or -] 0.30 (a)
          8       0.99 [+ or -] 0.26 (b)
          16      0.81 [+ or -] 0.14 (b)
          24       0.001 [+ or -] 0 (c)
          36       0.001 [+ or -] 0 (c)
          48       0.001 [+ or -] 0 (c)
Bottom    0       1.46 [+ or -] 0.30 (a)
          8       1.36 [+ or -] 0.02 (a)
          16      1.30 [+ or -] 0.21 (a)
          24      1.07 [+ or -] 0.19 (a)
          36      0.93 [+ or -] 0.05 (a)
          48      0.36 [+ or -] 0.30 (b)

All values are [log.sub.10] mean [+ or -] SD. Nonidentical
superscript letters within columns indicate a significant
difference at P < 0.05. The TPC between the surface and
bottom oysters were not significantly different (t-test, P =
0.426). MPN, most probable number.
COPYRIGHT 2014 National Shellfisheries Association, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2014 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Author:Chinnadurai, Shunmugavel; Mohamed, Kolliyil Sunil; Venkatesan, Vellathi; Sharma, Jenni; Kripa, Vasan
Publication:Journal of Shellfish Research
Article Type:Report
Geographic Code:9INDI
Date:Aug 1, 2014
Previous Article:A robust, spatially explicit model for identifying oyster restoration sites: case studies on the Atlantic and Gulf Coasts.
Next Article:Wheat germ agglutinin-binding glycoprotein extract from shells of conspecifics induces settlement of larvae of the Pacific oyster Crassostrea gigas...

Terms of use | Privacy policy | Copyright © 2020 Farlex, Inc. | Feedback | For webmasters