Printer Friendly

Histopathology of the digestive tissues and whole-body anaerobic bacteria counts of the eastern oyster, Crassostrea virginica, after experimental exposure to anoxia.

ABSTRACT The Eastern oyster, Crassostrea virginica, is a sensitive bio-indicator of environmental changes ranging from tidal flux to heavy-metal pollution. Extreme fluctuations in dissolved oxygen levels are well documented in Mobile Bay, AL. Extended periods of low dissolved oxygen occurring during the summer months have been shown to cause oyster mortality. The current study examined the effects of anoxia on tubule morphology of the digestive gland in the Eastern oyster as well as the quantity and quality of whole-body anaerobic bacteria counts. Oysters were exposed to anoxic conditions (<0.1 mg/L [O.sub.2]) at 28[degrees]C in a laboratory setting and sampled at 0 h (preexposure), 24 h, 48 h, and 60 h, and after a 4-wk recovery interval. The whole-body anaerobic bacteria count for test oysters (Mean = 1.11 x [10.sup.6] cfu/mL) from the 60-h interval was significantly higher than counts for the preexposure oysters, 24-h, and 48-h experimental intervals. The most common bacteria isolated were Clostridium sp. Histological examination of oysters exposed to anoxic conditions indicated stress in the digestive gland. Changes in shape and size of digestive tubule epithelial cells occurred, as well as sloughing of secretory absorptive cells into the digestive tubule lumen. Necrosis and inflammation composed of aggregates of hemocytes along with bacterial infiltration into digestive tissue was noted in oysters exposed to anoxic conditions. Digestive tubule lumen ratios of oysters sampled alter a 60-h exposure to anoxia were increased significantly in comparison with preexposure oysters. Digestive tubule lumen ratios returned to preexposure morphology after a 4-wk week recovery period. Oysters have the ability to regenerate digestive gland tissue if environmental conditions return to normal. These findings indicate that anoxia exposure at summer temperatures contributes to digestive gland atrophy and necrosis in C. virginica. The combination of digestive gland atrophy, necrosis, and bacterial infiltration into digestive tissues suggests that oysters may be succumbing to infection during periods of anoxic stress.

KEY WORDS: Crassostrea virginica, oyster, anoxia, histology, digestive tubule, bacteria


Hypoxic events in Mobile Bay, AL, are documented as far back as 1876 (May 1973). Suspected causes of these events range from salinity stratification to eutrophication of the bay system (Turner et al. 1987, Schroeder & Wiseman 1988, Diaz & Rosenberg 1995, Breitburg 2002, Kemp et al. 2009). Regardless of the predisposing cause, sessile benthic fauna are negatively affected by hypoxia. Occurring during summer months when metabolic rates are at their highest, periods of low dissolved oxygen within the bay have been proved to cause significant mortality of benthos, including reef-dwelling oysters (Baker & Mann 1992, Lenihan & Peterson 1998, Saoud et al. 2000a, Saoud et al. 2000b).

The Eastern oyster, Crassostrea virginica (Gmelin, 1791) is a widespread and abundant zoobenthic inhabitant of Mobile Bay. The oyster is the focus of an economically valuable commercial seafood industry and also serves an array of important ecological functions. Like many filter-feeding organisms, it is considered a sensitive bio-indicator of environmental stress. Morphologic changes in the digestive tissue of oysters can be indicative of environmental stressors such as altered tidal flows, decreased nutrient levels, water pollution, heavy-metal pollution and possibly anoxia (Shaw & Battle 1957, Morton 1977, Couch 1984, Winstead 1995, Carricker & Gaffney 1996). During periods of environmental stress, normal cuboidal epithelial cells that line the digestive tubule transform into simple, low squamous epithelial cells (Couch 1985, Weis et al. 1995). Along with epithelial cell atrophy, a contemporaneous change in lumen shape and size can be observed. Rounded dilated lumens with increased tubule lumen ratios have been documented in nutritionally deprived oysters as well as oysters exposed to environmental pollutants (Winstead 1995, Weis et al. 1995). Cell death, necrosis, and sloughing of secretory absorptive cells into the digestive tubule lumen occur with prolonged stressful environmental conditions (Couch 1985).

An acute inflammatory response along with edema in the digestive tissues may also occur during a stressful event (Pauley & Sparks 1965, Couch 1985). The hemocyte is the primary defense mechanism in the oyster. Thus, infiltration of hemocytes into oyster tissues can be a sign of physiological distress or disease. Infectious and noninfectious disease can cause an inflammatory response in bivalves (Tubiash et al 1973, Sparks 1976, Pipe & Coles 1995). Environmental stress can decrease the oyster's immune response, thus increasing susceptibility to disease-causing organisms (Chu & Hale 1994, Pipe & Coles 1995, Macey et al. 2008).

If environmental conditions return to tolerable levels before oysters succumb to degeneration or infection, they may be able to regenerate digestive tubule epithelial cells. A positive relationship between tubule reepithelialization of affected tissue and the return to acceptable environmental conditions has been documented by Eble and Scro (1996) and Bielefeld (1991), as well as Mix and Sparks (1971). Mix and Sparks (1971) described the regeneration of digestive tubule tissue in Crassostrea gigas approximately 50-60 days alter the cessation of ionizing radiation.

Although there has been significant documentation describing bivalve digestive tubule response to nutritional deprivation, environmental contaminants, and heavy-metal pollution, there have been no reports regarding the morphological effects of anoxia on oysters. The objective of this study was to examine the changes in digestive gland morphology in response to anoxic conditions at summer temperatures with special consideration for changes in whole-body anaerobic bacteria counts, dominant bacterial flora, and the ability to recover after an extended exposure.



Five hundred hatchery-produced oysters (C. virginica) were collected in May 2006 from suspended culture racks located in Bon Secour Bay, AL, and transported to the Auburn University Shellfish Laboratory on Dauphin Island, AL. The oysters were 1 y of age with a mean shell size of 48.07 [+ or -] 5.57 mm. Oysters were cleaned of any fouling organisms and stocked into a 2.4 x 0.6 x 0.25-m tank for a 7-day acclimation period. The tank was supplied with flow-through seawater filtered to 35 [micro]m and delivered at a rate of 3.0 L/h/oyster. Oysters were allowed to feed on naturally occurring phytoplankton and were not supplemented with feed during the acclimation period. A chiller unit (1 HP; Aqualogic, Inc., San Diego, CA) maintained a temperature of 28[degrees]C. A Hydrolab MiniSonde 4 (Hach Co., Loveland, CO) was placed in the tank to monitor temperature, salinity, and dissolved oxygen.

Experimental Conditions

After the 7-day acclimation period, oysters were transferred to aquaria. One aquarium had normoxic conditions (n = 110) whereas another aquarium was filled with anoxic seawater (n = 256). Aquaria (113 L) were filled with natural seawater filtered to 1 [micro]m and sterilized with UV light. Anoxic conditions were established in sealed aquaria by bubbling nitrogen through air stones for 24 h to deoxygenate the water before the addition of oysters. Oxygen concentration for the anoxic aquaria was maintained below 0.1 mg/L at all times by bubbling air supplied by a piston compressor (Sweetwater model SL22; Aquatic Ecosystems Inc., Apopka, FL). Temperature was maintained at 28 [+ or -] 1[degrees]C in both normoxic and anoxic aquaria with a 500-W Finnex Titanium Heating Tube (Finnex International Corp., Countryside, IL). Temperature, salinity, pH, and dissolved oxygen readings were taken every 4-6 h in both normoxic and anoxic aquaria using an YSI Model 85 Handheld Oxygen, Conductivity, Salinity and Temperature System (YSI Inc., Yellow Springs, OH) coupled with an Accumet AR 20 pH/conductivity meter (Fisher Scientific, Pittsburgh, PA). Because anoxic conditions can interrupt feeding in oysters, neither normoxic nor anoxic aquaria were ted during the experimental period. Thus, the normoxic (control) treatment should account for any effects of starvation throughout the course of the experimental period. Oysters were checked for mortality and transferred to new aquaria with their respective normoxic or anoxic conditions (established as described earlier) every 24 h to ensure adequate water quality. Dead oysters were counted, documented, and discarded. At the completion of the experimental trials, oysters from both normoxic and anoxic aquaria were removed from their respective tanks and put into a recovery tank (2.4 x 0.6 x 0.25 m) with flow-through seawater filtered to 35 [micro]m and maintained at 25 [+ or -] 1[degrees]C by a 1 HP Aqualogic, Inc., chiller unit. Oysters were allowed to feed on naturally occurring phytoplankton and were not supplemented with feed during recovery. During the recovery period, a Hydrolab MiniSonde (Hach, Edmonton Alberta) monitored temperature, salinity, and dissolved oxygen at 30-min intervals.

Oyster Sampling

Prior to placing oysters in the experimental conditions, 10 oysters were sampled (preexposure, 0 h) from the total population for histological examination to determine the initial condition of the digestive tissue. Another group of 10 oysters was sampled (preexposure, 0 h) for determination of initial whole-body anaerobic bacteria counts.

After initial sampling, the remaining 366 oysters were assigned randomly to either a normoxic (n = 110) or anoxic (n = 256) treatment. Oysters in the anoxia treatments were sampled at 24 h, 48 h, and 60 h intervals for histological examination of digestive tissues and whole-body bacteriology. Intervals were determined by a preliminary ranging experiment that showed sampling intervals of 24 h, 48 h, and 60 h provided a sufficient number of stressed oysters to sample while maintaining adequate survival in the anoxic conditions.

As a control for starvation effects, 10 oysters were sampled randomly from the normoxic treatment for histological examination of digestive tissues and whole-body bacteriology at 60 h. To assess the ability of oysters to recover from anoxic conditions and possible starvation effects, 20 oysters from both the normoxic and the anoxic treatments were sampled after a 4-wk recovery period for histological examination of digestive tissue (n = 10) and whole-body anaerobic bacteria counts (n = 10).


Oysters sampled for histological examination from preexposure (0 h), the normoxic treatments (60 h), and the anoxia treatments (24 h, 48 h, and 60 h) were measured, shucked, and wet meats weighed to the nearest 0.1 g. Whole oyster bodies were preserved in 2 parts Davidson's fixative:1 part ambient seawater at 25 psu for 24 h. After 24 h, the samples were transferred to 70% isopropyl alcohol (Howard & Smith 1983). Samples were transported to the Auburn University Department of Fisheries Southeastern Cooperative Fish Disease Laboratory in Auburn, AL, for histological processing.

A transverse section approximately 0.3 cm of each oyster located at the junction of the labial palps and the gills (Howard & Smith 1983) was processed, embedded in paraffin blocks, and sectioned. Three 5-[micro]m sections were mounted on each of 2 slides. Three sections on one slide were stained with Harris hematoxylin cosin (H&E) stain. The 3 sections on the other slide were stained with Giemsa stain. Tissue stained with Harris H&E was examined for changes in digestive gland structural morphology. Digestive tubule measurements from both normoxic and anoxic treatment oysters were also taken from H&E-stained slides. Tissue stained with Giemsa was examined for the presence of foreign organisms.

Digestive tubule lumen ratios were evaluated for normoxic (60 h) and anoxic treatment (24 h, 48 h, and 60 h) oysters sampled at each interval. Digestive tubules were measured and tubule lumen ratios calculated using techniques described by Winstead (1995). Each oyster section was divided into 4 quadrants. Three randomly selected digestive tubules were measured in each quadrant. Two sets of measurements were taken from each digestive tubule. The first set measured the perpendicular external diameters ([A.sub.1], [A.sub.2]) of the digestive tubule lumen: the second set measured the perpendicular internal diameters ([B.sub.1], [B.sub.2]) of the digestive tubule lumen (Fig. 1). Digestive tubule lumen ratios were calculated according to Winstead, 1995):

Tubule Ratio [B.sub.1] + [B.sub.2] / [A.sub.1] + [A.sub.2] (1)


Oysters sampled for bacteriological examination from preexposure (0 h), the normoxic treatments (60 h) and the anoxia treatments (24 h, 48 h, and 60 h) were individually scrubbed, measured, and then wiped with 70% isopropyl alcohol to sterilize the area. Oysters were shucked using aseptic technique into a sterile, tared homogenizer cup. The shucking knife was then cleaned with 70% isopropyl and set aside for the next sample. The wet meat weight was recorded to the nearest 0.01 g, then 10 mL sterile 25-psu seawater was added to the sample. Parafilm was secured to the top of the homogenizer cup and the sample was homogenized for approximately 30 sec using a commercial blender on high speed. A sterile, disposable 1-mL pipette was used to extract 1 mL homogenized sample, and the sample was then placed into a labeled, sterilized 15-mL centrifuge tube with 9 mL sterile, 25-psu seawater. The tube was capped, labeled with the sample number, and placed under a sterile UV hood for plating. The remainder of the homogenate was discarded. The homogenizer cup was then sterilized for the next sample. Plating of samples began after all oysters were homogenized and aliquoted to their respective tubes. Each interval had a specific plating dilution based on expected number of bacteria. The dilution for pretreatment oysters, 24 h, control, and recovery intervals was [10.sup.-4]. The dilution fox 48 h and 60 h was [10.sup.-5]. Expected bacterial numbers were determined by prior sampling of oysters in a preliminary experiment. Marine Agar and Brain Heart Infusion Agar with 1% NaCl were both chosen for plating bacteria because of their nonselective properties. Bacterial colonies grown on Brain Heart Infusion Agar were identified with the Rapid ANA II system (Innovative Diagnostic Systems, Inc., Atlanta, GA).


Each oyster dilution was plated by placing 0.1 mL on each of 4 Marine Agar plates and 2 Brain Heart Infusion with 1% NaCl plates and spread with a sterilized glass rod (Buck & Cleverdon 1960). Three Marine Agar plates and 1 Brain Heart Infusion with 1% NaCl plate were incubated in an anaerobic environment using BD Bio-Bag Environmental Chambers Type A (Becton, Dickinson and Co., Franklin Lakes, NJ). Anaerobic chambers were used to isolate facultative anaerobic bacteria for total body counts, isolation, and identification. The remaining Marine Agar and Brain Heart Infusion with 1% NaCl plates were incubated simultaneously in aerobic conditions. A simultaneous incubation was used to determine whether there was a difference between bacterial counts in an aerobic environment versus an anaerobic environment. During each plating session, 1 Marine Agar plate and 1 Brain Heart infusion with 1% NaCl plate were left open in the hood and incubated to account for any potential contamination.

Colony forming units (cfu) were counted and recorded for each plate after 24-48 h (as recommended by anaerobic chamber type A directions) of incubation in an oven at 31[degrees]C. Individual colonies were chosen from the anaerobic incubated plates for further isolation and identification based on morphological properties of the colony.

Only the most abundant colonies were used for identification. The Rapid ANA II system (Innovative Diagnostic Systems, Inc., Norcross, GA) was used to identify the bacteria along with a Gram stain, catalase test, oxidase test, and indole test.

Statistical Analysis

Statistical analyses were performed using Statistical Analysis System (SAS) version 9.1 (SAS Institute, Cary, NC). The mean digestive tubule lumen ratios were calculated at each sampling interval for oysters from the preexposure samples (0 h), the normoxic treatment group (60 h), and the anoxic (24 h, 48 h, and 60 h) treatment group. Bacteriological counts were averaged for each sampling interval and treatment group. Bacteriological samples were [log.sub.10] transformed prior to analysis. Data were analyzed using 1-way ANOVA with Tukey's "Studentized" range (HSD) test for determination of differences among groups with a significance level of P < 0.05.


Oysters in the anoxia treatment group experienced 9.0% mortality through the 60-h exposure. Mortality increased with the length of the exposure to anoxia, with mortalities of 0.39% after 24 h, 2.7% after 48 h, and 5.9% after 60 h. In contrast, there were no oyster mortalities in the normoxic treatment group throughout the experiment.

Microscopic examination of digestive tissue in preexposure (0 h) oyster samples revealed the digestive tubules to be in an absorptive phase or a holding phase according to the standards set by Morton (1977). The digestive tubules in preexposure (0 h) oysters consisted of simple columnar epithelium composed of dark-stained basophilic cells admixed with lightly stained secretory absorptive cells (Fig. 2). These tubules were surrounded by a thin eosinophilic rim of basement membrane. Connective tissue also known as Leydig tissue surrounded the digestive gland and was comprised of collagenous fibers. All Leydig tissue examined during the preexposure (0 h) interval was within normal limits (Fig. 2A).


After a 24-h exposure to anoxia, digestive tubule basophilic cells transformed from tall columnar cells into squamous cells (Fig. 2B). Sloughing of the secretory absorptive cells into the lumens of the digestive tubules and a notable deterioration of the tubule epithelium was also evident after 24 h of anoxia exposure (Fig. 2B). This degeneration continued through to the end of the anoxia exposure (60 h), leading to pyknotic nuclei, karyorrhexis, karyolysis, and increased eosinophilia of the cytoplasm, with loss of distinct cell borders consistent with coagulative necrosis. Edema of the Leydig tissue was apparent after 24 h of anoxia exposure and continued through to the 60-h sampling interval. Multifocal areas of hemocytic infiltration of the digestive gland, including the Leydig tissue and digestive tubules, occurred concurrently with sloughing of basophi[ic cells into the tubule lumen after 48 h of anoxia exposure and extended through to the termination of the anoxia exposure (60 h: Fig. 2C, D).

After a 4-wk recovery period, the digestive tubules of oysters from the anoxia treatment group had regenerated to the preexposure condition, with small lumens and tall columnar epithelial cells (Fig. 2F). No hemocytic infiltration associated with the digestive tubule or Leydig tissue was seen after the 4-wk recovery period.

Microscopic examination of oysters from the normoxic treatment group after 60 h revealed an increase in the size of the tubule lumens as well as a flattening of the basophilic cells. There was no hemocytic infiltration, necrosis, sloughing of basophilic cells into tubule lumens, or edema apparent in digestive gland tissue of the normoxic treatment group oysters at any time. Digestive tubules in oysters from the normoxic treatment group also returned to their preexposure (0 h) morphological condition after a 4-wk recovery period (Fig. 2G).

Mean digestive tubule ratios increased significantly from 0.252 [+ or -] 0.02 (mean [+ or -] SE) in preexposure (0 h) oyster samples to 0.635 [+ or -] 0.02 in oysters from the anoxic treatment group at the 24-h interval. Ratios for the anoxic treatment group remained high at the 48-h and 60-h sampling intervals--0.616 [+ or -] 0.02 and 0.615 [+ or -] 0.02, respectively (Fig. 3). Oysters from the normoxic treatment group had a mean digestive tubule ratio of 0.468 [+ or -] 0.02 at the 60-h interval, which was significantly lower than ratios for oysters in the anoxic treatment group at the 60-h interval (0.615 [+ or -] 0.02). Digestive tubule ratios for the 60-h normoxic treatment group were significantly higher than ratios in the preexposure samples (0 h), which were 0.252 [+ or -] 0.02.(Fig. 3). Digestive tubule ratios in oysters assessed after the recovery period from the anoxic treatment group (0.284 [+ or -] 0.02) were not significantly different from the preexposure (0 h) group (0.215 [+ or -] 0.02) or oysters from the normoxic treatment group (0.361 [+ or -] 0.02) after the recovery period (Fig. 3).

Giemsa stain of digestive tubule tissue from oysters in the anoxic treatment group from the 24-h, 48-h, and 60-h anoxia exposure intervals showed bacterial infiltration of digestive tubule hunens and the surrounding Leydig tissue (Fig. 4B). At the 24-h interval, 3 of the 10 oysters sampled had 4-8-[micro]m rod-shaped bacteria. At both the 48-h and 60-h interval, 2 of the 10 oysters were positive for 4-8-[micro]m rod-shaped bacteria. There was no evidence of bacterial infection in preexposure (0-h) oyster samples (Fig. 4A), normoxic treatment group oyster samples (60 h), or in oysters sampled after the 4-wk recovery period in either treatment group.

The mean [log.sub.10] transformed facultative anaerobic bacterial count for test oysters in the anoxic treatment group from the 60-h interval (5.87 [+ or -] 0.08 cfu mL) was significantly higher than counts for oysters from pre-exposure samples (5.05 [+ or -] 0.07 cfu/ mL) anoxic treatment group oysters from the 24-h (4.94 [+ or -] 0.11 cfu/mL) and 48-h (5.60 [+ or -] 0.09 cfu/mL) intervals, and normoxic treatment group oysters from the 60-h interval (5.69 [+ or -] 0.06 cfu/ mL; Fig. 5). After the 4-wk recovery period, both normoxic and anoxic treatment group oysters had significantly lower mean [log.sub.10] transformed facultative anaerobic bacterial counts (4.44 [+ or -] 0.07 cfu/mL and 4.58 [+ or -] 0.07 cfu/mL, respectively) than oysters from preexposure samples, oysters from all anoxic treatment group exposure intervals (24 h, 48 h, and 60 h), and oysters from the normoxic treatment group (60 h: Fig. 5). A total of 43 isolates were taken for identification. Three species of anaerobic bacteria accounted for 74% of the bacteria isolated throughout the experiment--Clostridium sp. (53%), Propionibacterium sp. (12%), and Actinomyces sp. (9%), with the remaining 26% being unidentifiable Gram-negative rods.




The results of this experiment indicate that anoxia is a probable cause of digestive tubule atrophy beyond the expected starvation response. Although oysters in the normoxic treatment group exhibited only digestive tubule atrophy, the oysters subjected to anoxic conditions exhibited significantly higher levels of atrophy with a concurrent inflammatory response, necrosis, and bacterial infiltration within the digestive tissues.

Although stressors such as starvation and pollution are well documented to induce morphologic changes in digestive tubules, there is little information describing the effects of anoxia on the oyster digestive gland. Digestive tubules are comprised of flagellated basophilic cells, nonflagellated basophilic cells, and digestive cells (Weinstein 1995). There are 5 main stages of normal digestive tubule conformation: (1) the holding stage, (2) the absorptive stage, (3) the second absorptive stage, (4) the disintegration stage, and (5) the reconstitution stage (Morton 1977). In the current study, digestive tubules in oysters before they were exposed to anoxic conditions were determined to be in a holding or absorptive stage. All oysters exposed to anoxic conditions including the 24-h, 48-h, and 60-h intervals were determined to be in a disintegration stage or exhibiting necrosis. After 24 h of exposure to anoxia, histological review of oyster tissue showed sloughing of secretory absorptive cells into tubule lumens as well as flattening of tubule epithelial cells. Deterioration of the digestive tubules continued through 48 h and 60 h of anoxia exposure, with atrophy of the tubule epithelium and sloughing of secretory absorptive cells into a large, dilated digestive tubule lumen. Similar results have been described for bivalves exposed to periods of starvation and pollution stress (Couch 1984, Bielefeld 1991, Winstead 1995).

Oysters from the normoxic treatment group were found to be in disintegration or a holding phase during the experiment, with no evidence of inflammation and necrosis, unlike that documented in the anoxia treatment group oysters. The observed atrophy of the digestive tubules in the control oysters is likely a starvation response concurrent with increased temperature. The significant difference in the levels of deterioration between normoxic and anoxic treatment group oysters suggests that anoxia promotes degradation, inflammation, and necrosis in the digestive tubules, and compounds the effects of nutrient deficiency.


Digestive tubule ratios were significantly higher in oysters from all the anoxic treatment intervals compared with preexposure and normoxic treatment oysters. The increase in the mean digestive tubule ratios in oysters from the anoxic treatments agree with the morphological changes witnessed microscopically. A significant increase in digestive tubule ratios in oysters from the normoxic treatment group at the 60-h interval compared with preexposure oysters indicated a possible starvation response associated with the experimental protocol. However, oysters exposed to the 60-h anoxia treatment had digestive tubule lumen ratios that were 286% larger than the preexposure digestive tubule lumen ratios. In comparison, oysters in the control 60-h treatment group that were exposed to normoxic conditions had digestive tubule lumen ratios that were only 217% larger than preexposure oysters (0 h). The additional 69% increase in digestive tubule lumen ratios for oysters exposed to anoxia can be attributed to the additional stress of the anoxia treatment. These results are similar in magnitude to the stress response seen in salinity and starvation studies performed by Winstead (1995), but strikingly different in terms of the speed of the response. Winstead (1995) subjected oysters to starvation at 26[degrees]C and, by day 15, digestive tubule ratios were 0.665 (SE [+ or -] 0.009). Oysters in this study had a similar ratio of 0.635 (SE [+ or -] 0.024) as early as 24 h of anoxia exposure at 28[degrees]C. The accelerated increase in tubule lumen ratios can also be attributed to anoxia exposure.

An inflammatory response was observed after oysters were exposed to anoxia, as indicated by infiltration of hemocytes into digestive tissues. Although hemocyte infiltration into gonadal tissues is a natural process alter spawning for the oyster, infiltration of these cells into the digestive gland is not a normal occurrence in healthy oysters. Inflammation and increased susceptibility to pathogens brought on by stressful environmental conditions has been documented in bivalves (Pipe & Coles 1995, Fisher et al. 1999, Chu et al. 2002, Macey et al. 2008). Pipe and Coles (1995) observed a decreased immune response with a concurrent increase in mortality when exposing marine mussels to doses of cadmium and copper after challenging them with Vibrio tubiashi. Both Fisher et al. (1999) and Chu et al. (2002) found that exposure to environmental pollution increases the intensity of preexisting infections of Perkinsus marinus as well as susceptibility to new infection by suppression of the host immune response. Most recently, Macey et al. (2008) demonstrated that C. virginica exposed to hypercapnic hypoxic conditions experienced a decrease in their ability to deactivate, degrade, and eliminate pathogens such as Vibrio campbellii.

Although normoxic treatment group oysters showed no hemocytic or bacterial infiltration into the digestive tissue, anoxic treatment group oysters had signs of congestion along with infiltration of hemocytes into Leydig tissue, as well as digestive tubule epithelium. After 24 h of exposure to anoxia, oyster hemocytes as well as rod-shaped bacteria were seen in the digestive tissues. The presence of hemocytes and bacteria was noted in the anoxic treatment group through the 60-h interval in association with necrosis of the tubule epithelium. Elston et al. (1987) described a similar reaction to clusters of pathogenic bacteria as an extensive amoebocytic inflammatory response resulting from Gram-positive bacterial infiltration into tissues of Pacific oysters C. gigas. Edema and leukocytic infiltration along with congestion of smaller blood vessels with hemocytes have been attributed to invasion of foreign matter into oyster tissues. Pauley and Sparks (1965, 1966) noted this inflammatory response when oysters where injected with turpentine and talc.

Oysters can survive exposure to anoxia and have the ability to recover to a normal morphological state if normoxic conditions return. In this study, oysters exposed to anoxic conditions regenerated digestive tubules to the holding phase with normal cuboidal epithelial digestive cells within a 4-wk recovery period. Similar tubule reepithelialization in C. gigas was seen after a 90-day recovery period from ionizing radiation treatments (Mix & Sparks 1971). A significant decrease in tubule ratios at the 4-wk recovery interval also provides substantial evidence that there is regeneration of the digestive tubules to the normal holding stage morphology. Starvation studies by Winstead (1995) showed digestive tubule ratios dropped significantly within 2 days after feeding was initiated and were similar to control oysters 6 days after reinitiating feed.

Bacteria proliferate under high temperatures as well as in anaerobic conditions. C. gigas and C. virginica carry a natural population of both aerobic and anaerobic bacteria. These include Pseudomonas. Vibrio and Flavobacterium as the major aerobic flora (Colwell & Liston 1960). Both aforementioned aerobic bacteria have proteolytic activity and are linked with postmortem spoilage of oysters. An oyster's natural anaerobic bacterial flora may include several species, but the major constituent is Clostridium spp. (Hariharan et al. 1995).

Clostridium species are known to cause enteric infections in both humans, and domestic and wild animals. As part of the normal flora of C. virginica, Clostridium perfringens and Clostridium difficile should not be overlooked as a pathogen. Both species can produce exotoxins that can have cytotoxic effects (Songer 1996). It is unclear at this point the role that Clostridium spp. or other facultative anaerobic bacteria may be playing in the necrosis and subsequent death of an oyster under anaerobic conditions. However, this study clearly documented an increase in numbers of facultative anaerobic bacteria as well as infiltration of rod-shaped bacteria concurrent with inflammation and necrosis within the digestive gland tissues of oysters exposed to anoxic conditions. The use of enzyme linked immunosorbent assay, PCR, or electrochemoluminescence assays may be useful to identify whether exotoxins are present within the oysters during anoxic events and to determine the levels at which they are present.

In conclusion, this study brings to light the physiological stress that C. virginica endures during an extended period of low oxygen at summer temperatures. Digestive tubule atrophy and inflammation with significantly increased anaerobic bacterial counts were the most prominent changes in oysters exposed to anoxic conditions. It is probable that the stress of being in an anoxic environment with elevated water temperatures typical of summer conditions in the Gulf of Mexico decreases oysters' defense mechanisms, in turn making them more susceptible to pathogens and subsequent mortality.


Baker, S. M. & R. Mann. 1992. Effects of hypoxia and anoxia on larval settlement, juvenile growth, and juvenile survival of the oyster Crassostrea virginica. Biol. Bull. 182:265-269.

Bielefeld, U. 1991. Histological observation of gonads and digestive gland in starving Dreissena polvmorpha (Bivalvia). Malacologia 33:31-42.

Breitburg, D. 2002. Effects of hypoxia, and the balance between hypoxia and enrichment on coastal fishes and fisheries. Estuaries Coasts 25:767 781.

Buck, J. D. & R. C. Cleverdon. 1960. The spread plate as a method for the enumeration of marine bacteria. Limnol. Oceanogr. 5:78-80.

Carricker, M. R. & P. M. Gaffney. 1996. A catalogue of selected species of living oysters. In: V. S. Kennedy, R. I. E. Newell & A. F. Eble, editors. The Eastern oyster Crassostrea virginica. MD: Maryland Sea Grant College, College Park. pp. 75-168.

Chu, F.- L. E., V. K. Aswani, R. C. Hale & Y. Huang. 2002. Cellular responses and disease expression in oysters (Crassostrea virginica) exposed to suspended field-contaminated sediments. Mar. Environ. Res. 53:17-35.

Chu, F.- L. E. & R. C. Hale. 1994. Relationship between pollution and susceptibility to infectious disease in the Eastern oyster, Crassostrea virginica. Mar. Environ. Res. 38:243-256.

Colwell, R. R. & J. Liston. 1960. Microbiology of shellfish: bacteriological study of the natural flora of Pacific oysters (Crassostrea gigas). Appl. Environ. Microbiol. 8:104-109.

Couch, J. A. 1984. Atrophy of diverticular epithelium as an indicator of environmental irritants in the oyster, Crassostrea virginica. Mar. Environ. Res. 14:525-526.

Couch, J. A. 1985. Prospective study of infectious and noninfectious diseases in oysters and fishes in three Gulf of Mexico estuaries. Dis. Aquat. Organ. 1:59-82.

Diaz, R. J. & R. Rosenberg. 1995. Marine benthic hypoxia: a review of its ecological effects and the behavioural responses of benthic macrofauna. Oceanogr. Mar. Biol. Annu. Rev. 33:245-303.

Eble, A. F. & R. Scro. 1996. General anatomy. In: V. S. Kennedy, R. I. E. Newell & A. F. Eble, editors. The Eastern oyster Crassostrea virginica. MD: Maryland Sea Grant College, College Park. pp. 19-73.

Elston, R. A., J. H. Beattie, C. Friedman, R. Hedrick & M. L. Kent. 1987. Pathology and significance of fatal inflammatory bacteremia in the Pacific oyster, Crassostrea gigas Thtinberg. J. Fish Dis. 10:121-132.

Fisher, W. S., L. M. Oliver, W. W. Walker, C. S. Manning & T. F. Lytle. 1999. Decreased resistance of Eastern oysters (Crassostrea virginica) to a protozoan pathogen (Perkinsus marinus) after sublethal exposure to tributyltin oxide. Mar. Environ. Res. 47:185-201.

Hariharan, H., J. S. Giles, S. B. Heaney, G. Arsenault, N. McNair & D. J. Rainnie. 1995. Bacteriological studies on mussels and oysters from six river systems in Prince Edward Island, Canada. J. Shellfish Res. 12:527-532.

Kemp, W. M., J. M. Testa, D. J. Conley, D. Gilbert & J. D. Hagy. 2009. Temporal responses of coastal hypoxia to nutrient loading and physical controls. Biogeosciences 6:2985-3008.

Howard, D. W. & C. S. Smith. 1983. Histological techniques for marine bivalve mollusks. National Oceanic and Atmospheric Administration technical memorandum NMFS-F/NEC-25. Woods Hole, MA:

National Oceanic and Atmospheric Administration. 97 p. Lenihan, H. & C. H. Peterson. 1998. How habitat degradation through fishery disturbance enhances impacts of hypoxia on oyster reefs. Ecol. Appl. 8:128-140.

Macey, B. M., I. O. Achilihu, K. G. Burnett & L. E. Burnett. 2008. Effects of hypercapnic hypoxia on inactivation and elimination of Vibrio campbellii in the Eastern oyster, Crassostrea virginica. Appl. Environ. Microbiol. 74:6077-6084.

May, E. B. 1973. Extensive oxygen depletion in Mobile Bay, Alabama. Limnol.. Oceanogr. 18:353-366.

Mix, M. C. & A. K. Sparks. 1971. Repair of digestive tubule tissue of the Pacific oyster, Crassostrea gigas, damaged by ionizing radiation. J. Invest. Pathol. 17:172-177.

Morton, B. S. 1977. The tidal rhythm of feeding and digestion in the Pacific oyster, Crassostrea gigas (Thunberg). J. Exp. Mar. Biol. Ecol. 26:135-151.

Pauley, G. B. & A. K. Sparks. 1965. Preliminary observations on the acute inflammatory reaction in the pacific oyster, Crassostrea gigas (Thunberg). J. Invest. Pathol. 7:248-256.

Pauley, G. B. & A. K. Sparks. 1966. The acute inflammatory response in two different tissues of the Pacific oyster, Crassostrea gigas. J. Fish. Board Can. 23:1913-1921.

Pipe, R. K. & J. A. Colts. 1995. Environmental contaminants influencing immune function in marine bivalve mollusks. Fish Shellfish Immunol. 5:581-595.

Saoud, I. G., D. B. Rouse, R. K. Wallace, J. Howe & B. Page. 2000a. Oyster Crassostrea virginica spat settlement as it relates to the restoration of Fish River Reef in Mobile Bay, Alabama. J. World Aquacult. Soc. 31:640-650.

Saoud, I. G., D. B. Rouse, R. K. Wallace, J. E. Supan & S. Rikard. 2000b. An in situ study on the survival and growth of Crassostrea virginica juveniles in Bon Secour, Alabama. J. Shellfish Res. 19:809-814.

Schroeder, W. W. & W. J. Wiseman, Jr. 1988. The Mobile Bay estuary: stratification, oxygen depletion, and jubilees. In: B. Kjerfve. editor. Hydrodynamics of estuaries II: estuarine case studies. Boca Raton, FL: CRC Press. pp. 42-52.

Shaw, B. L. & H. I. Battle. 1957. The gross and microscopic anatomy of the digestive tract of the oyster Crassostrea virginica (Gmelin). Can. J. Zool. 35:325-337.

Songer, J. G. 1996. Clostridial enteric diseases of domestic animals. Clin. Microbiol. Rev. 9:216-234.

Sparks, A. K. 1976. Inflammation and wound repair in oysters. Mar. Fish. Rev. 38:2-4.

Tubiash, H. S., S. V. Otto & R. Hugh. 1973. Cardiac edema associated with Vibrio anguillarum in the American oyster. Proc. Natl. Shellfish. Assoc. 63:39-42.

Turner, R. E.. W. W. Schroeder, & W. J. Wiseman. 1987. The role of stratification in the deoxygenation of Mobile Bay and adjacent shelf bottom waters. Estuaries 10:13-19.

Weinstein, J. E. 1995. Fine structure of the digestive tubule of the Eastern oyster, Crassostrea virginica (Gmelin, 1791). J. Shellfish Res. 14:97-103.

Weis, P. J., S. Weis, J. Couch, C. Daniels & T. Chen. 1995. Pathological and genotoxicological observations in oysters (Crassostrea virginica) living on chromated copper arsenate (CCA)-treated wood. Mar. Environ. Res. 39:275-278.

Winstead, J. T. 1995. Digestive tubule atrophy in Eastern oyster, Crassostrea virginica (Gmelin, 1971) exposed to salinity and starvation stress. J. Shellfish Res. 14:105-111.


(1) Departnwnt of Fisheries and Allied Aquaeulturesm Auburn University, 203 Swingle Hall, Auburn, AL 36849; (2) Auburn University Shellfish Laboratory, 150 Agassiz Street, Dauphin Island, AL 36528; (3) Auburn University Marine Extension and Research Center, 4170 Commanders Drive, Mobile, AL 36615

* Corresponding author. E-mail:

DOI: 10.2983/035.030.0305
COPYRIGHT 2011 National Shellfisheries Association, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2011 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Author:Fogelson, Susan B.; Rikard, F. Scott; Brady, Yolanda; Wallace, Richard K.
Publication:Journal of Shellfish Research
Article Type:Report
Geographic Code:1USA
Date:Dec 1, 2011
Previous Article:Isolation and evaluation of new probiotic bacteria for use in shellfish hatcheries: II. effects of a Vibrio sp. probiotic candidate upon survival of...
Next Article:Oyster hemocyte mobilization and increased adhesion activity after [beta]-glucan administration.

Terms of use | Privacy policy | Copyright © 2018 Farlex, Inc. | Feedback | For webmasters