Abstract
Consumption of raw oysters is known to cause serious health conditions due to bioaccumulation of contaminants. As filter feeders, oysters ingest bacteria along with phytoplankton from their surrounding habitats. Ensuring seafood safety for human consumption is always a concern. Since oysters are consumed raw, disease causing organisms, environmental contaminants, toxins, chemicals, and even physical hazards such as soils and metals retained in the oysters can enter through feeding. The objective of this study was to determine the quality of oysters collected from Delaware Inland Bays (DIB) and compare them with market oysters. Environmental parameters were monitored from local waters of DIB classified as closed versus open for shellfish harvesting. Total aerobic bacteria and vibrio were higher in market oysters during the warmer months, with open water having the least microbial loads. There were no significant differences in total vibrio counts between the study sites (), but significant differences were recorded over time (). Water temperature and turbidity were directly proportional to total vibrio in oysters, and salinity was inversely related. Research findings in this study may help bring awareness of changes in bacterial loads due to seasonal changes and additional handling and storage.
1. Introduction
Nearly 20 million Americans consume raw oysters, and when eaten raw, they pose health risks due to the potential exposure to pathogenic bacteria such as Vibrio vulnificus and Vibrio parahaemolyticus [1, 2]. People with health conditions such as liver disease, iron overload disease (hemochromatosis), diabetes, cancer, and stomach disorders are at high risk for serious illness or death from V. vulnificus infections [1]. Recently, the Florida Department of Health issued a press release warning to residents with certain health conditions to avoid eating raw oysters and exposing open wounds to seawater and estuarine areas due to the potential exposure to V. vulnificus bacteria [3]. V. parahaemolyticus, a human pathogen, which causes gastrointestinal illnesses, is commonly found in shellfish from coastal bay systems of the United States and Canada [4]. Oysters are the most harvested shellfish in the world [5]; they are nonmotile marine organisms and obtain their food by filtering the water in which they live [6]. Because of filter feeding, they can accumulate pathogenic microorganisms and other chemical contaminants, which cannot always be removed by depuration [7, 8]. Most oysters are stored at refrigeration temperatures (3 to 5°C) and sold live or shucked without further processing.
Previous studies stated that the environment in which the oysters live affects the physical characteristics of oyster meat [9, 10]. Mackenzie et al. [11] indicated that there are consistently excess nutrients such as nitrogen, phosphorus, and low dissolved oxygen in the Delaware Inland Bays because of the intensive agricultural operation such as corn, soybean, and poultry production in Delaware. These bays are very shallow (i.e., 1 to 2.4 meters) and poorly flushed by tidal movement, and they are especially sensitive to environmental changes. Increases in pollutants, changes in salinity due to the frequency of changes in precipitation or drought events, climate change-related fluctuations in water temperature, episodic hypoxic and anoxic conditions, and frequent harmful algal blooms can all have detrimental effects on the health of native eastern oysters (Crassostrea virginica) and humans who consume these oysters. Consumption of uncooked oysters can cause serious health conditions and illnesses due to the bioaccumulation of contaminants [12]. For the last decade, bacterial monitoring in oysters coupled with ecological and environmental monitoring and oyster growth studies have been conducted to determine the relationships between total bacteria and total vibrio levels in relation to water quality parameters such as turbidity, dissolved oxygen, temperature, pH, and total suspended solids in the bays [13]. For the two years of the decadal study, physical and chemical quality attributes and microbial contaminants including total vibrio counts in oysters were analyzed to assess the overall quality of oysters from local waters and local seafood market.
Oysters are high in zinc and omega 3 fatty acids, low in fats, and an important source of protein. Previous studies suggest that relative proportions of proteins, carbohydrates, and lipids are approximately the same in all oyster species. Depending on washing and processing methods and their frequencies, oysters may lose some of their soluble salts. Yurimoto [14] investigated the types of tissues accumulating glycogen and seasonal changes in glycogen content and found lower glycogen contents in autumn for all three species, pen shell Atrina lischkeana, ark shell Scapharca kagoshimensis, and Manila clam Ruditapes philippinarum, while he found higher glycogen contents in spring. Summer glycogen content was lower in the ark shell but higher in the Manila clam. Yamamura and Watanabe [15], Akashige and Fushimi [16], and Shinomiya et al. [17] investigated glycogen contents in oysters and found it to be dependent on the physiological state of the organism. They concluded physiological condition in different bivalve species such as Pacific oysters (Crassostrea gigas) and pearl oysters (Pinctada fucata martensii) can be evaluated by using glycogen content. Ren et al. [18] reported that glycogen concentration in oysters decrease when the outside temperatures drop and as gonadal development increases. According to Galtsoff [19], chemical compositions of oysters change throughout the year because of seasonal, environmental, and anthropogenic changes. Salinity and temperatures of the water systems where the oysters live are the two major environmental factors that affect the chemical composition of oysters [19–21]. Thus, environmental conditions are very important in maintaining the oyster quality. Reports show that along with temperature, handling and storing practices are other important factors that determine the shelf life of shellfish [22]. In addition, temperature control helps regulate the rate of bacterial spoilage; higher temperatures decrease shelf life and promote bacterial growth and enzyme breakdown [23].
Our research objective was to determine the microbial loads and physical and chemical attributes of oysters collected from the Delaware Inland Bays and those of market oysters exposed to additional handling and storage. Eastern oyster quality was determined through physical (color, texture, pH, and moisture), chemical (protein, total lipids, fatty acids, and glycogen), and microbial (aerobic and psychrotropic bacteria and total and fecal coliform) analyses during the warmer months in this two-year study. Our hypotheses in this project are twofold: total bacteria and Vibrionaceae concentrations are strongly related to environmental quality and market oyster quality is poorer than the Delaware Inland Bays’ oysters.
2. Materials and Methods
2.1. Sample Collection and Preparation for Analyses
Eastern oysters (Crassostrea virginica) were obtained from three locations: closed canal (Rehoboth, Delaware), open water (Indian River Marina at Delaware Seashore State Park), and local seafood market (Smyrna, Delaware) on the day of analysis. At the closed canal site, there is a moratorium on oyster harvesting for human consumption, whereas the open water site is in an area open for fishing and seafood consumption. Although market oysters are local oysters harvested from the Delaware Bay and transported in bushels to the seafood market, they were not always sold to the consumers on the same day they were shipped to the market. The primary purpose of using market oysters was to investigate any negative impacts that may have occurred during handling and storage of live oysters, as market oysters are the most commonly available source for consumption by public. Oysters from the local seafood market were transported on ice at 0–4°C without having oysters directly in contact with ice to the aquatic sciences and seafood safety laboratories at Delaware State University. While oysters from the closed canal and open waters were transported in a temperature-controlled cooler with internal temperature of 4–7°C and analyzed immediately the same day, they were obtained for the microbial testing. Samples were processed the same day for the physical and chemical attributes and analyzed following microbial testing. Physical, chemical, and microbiological analyses were performed once a month in 2013 and twice a month in 2014 from June through November. Based on our previous research efforts [13], late May through early November were the months bacterial contaminants were detectable in shellfish. On each sampling day, eighteen oysters were processed from each location of which three oysters were cleaned, shucked, and used for color and texture analyses for each sampling location. Nine oysters were cleaned, shucked, and homogenized using a stomacher (Stomacher 400 Circulator, Seward, England). The homogenate, which consisted of oyster and its liquor, was used for pH, moisture, microbiological, and chemical analyses. The remaining six oysters were used for the chemical analyses.
2.2. Water Quality
During the sampling dates, physical water quality parameters were recorded for temperature, dissolved oxygen (DO), pH, and salinity using a 556 YSI Multiprobe (YSI Inc., Yellowstone, OH) at open water and closed canal study sites in the Delaware Inland Bays. Chemical water quality parameters were analyzed in triplicates according to the HACH methods for total nitrogen (Method 10071—Persulfate Digestion Method), total phosphorous (Method 8190—PhosVer 3 with Acid Persulfate Digestion Method), nitrate (Method 8171—Cadmium Reduction Method), and total suspended solids (TSS) (Method 8006—Suspended Solids Photometric Method). Alkalinity and turbidity were analyzed with the YSI 9500 photometer (YSI Inc., Yellowstone, OH).
2.3. Physical Analyses of Oysters
2.3.1. Color
Oyster tissue color was analyzed with a colorimeter (Miniscan XE Plus, Reston, VA) using the Hunter L, a, and b scale (with L representing the lightness on a scale of 0 (dark) to 100 (white), +a for redness, −a for greenness, +b for yellowness, and −b for blueness). Three oysters per treatment and three measurements were taken per oyster on the ventral body of the oyster tissue. The colorimeter was calibrated before each series of measurements using a white tile and black glass (X = 80.0, Y = 84.9, Z = 90.2) as per the manufacturer’s instruction.
2.3.2. Texture
The ventral body of the clean oyster tissue was analyzed using a TX. XT Plus texture analyzer (Scarsdale, NY). A 2 mm cylinder probe and a TA-90H plate (plate with a hole) were used. The probe penetrated the samples with a test speed of 1 mm/s and a distance of 10 mm. The peak force values were recorded as the force in grams, and mean values were used in data interpretation.
2.3.3. Oyster Homogenate pH
The pH of oyster homogenates was measured [6] using a pH meter (Thermo Fisher Scientific Inc., Bridgewater, NJ). The pH meter was calibrated prior to each sampling time for the three-scale pH calibration (pH: 4–7–10) using pH buffers (Fisher Scientific Co., Hanover Park, IL).
2.3.4. Moisture
Oyster samples were weighed and then dried in an oven (Fisher Scientific Co, Hanover Park, IL) at 100°C for 24 h. After drying, oyster samples were placed in a desiccator to cool overnight and then weighed again. The moisture percentage was calculated as a percentage of moisture in the sample (%W).
2.4. Microbiological Analyses of Oysters
2.4.1. Total Aerobic Bacterial and Psychrotropic Counts
Oyster homogenates were serially diluted with phosphate-buffered saline (PBS) in tenfold dilutions. Plate counts were determined according to the method of Cousin et al. [24]. Bacterial counts of oysters were determined by spread plating on Tryptic Soy Agar (TSA) plates. Aerobic plate counts (APCs) were determined by incubation at 35°C for 48 h. Psychrotropic plate counts (PPCs) were determined by incubating the plates at 7°C for 10 days. The results were recorded as log colony-forming units (CFU) per gram oyster tissue.
2.4.2. Total and Fecal Coliform Counts
Total and fecal coliforms were analyzed with the five-tube most probable number (MPN) method [25]. Dilutions of samples from aerobic plate count were inoculated into Lauryl tryptose broth (LTB) with incubation at 35°C for 24 h. Each LTB tube with gas production was inoculated into Brilliant Green Lactose Bile (BGLB) broth with incubation at 35°C for 48 h. Each BGLB tube with gas production confirmed the presence of coliforms in the samples. Those tubes were inoculated with Escherichia coli (EC) broth incubating at 44.5°C for 48 h. Each EC broth with gas production confirmed the presence of fecal coliforms in the samples. Fermentation tubes were placed in all tubes to confirm the presence of gas production. The results were recorded as log MPN/g.
2.4.3. Total Vibrio Counts
Total vibrio counts in oysters were determined by colony overlay procedure for peptidases (COPP) Assay method according to Richards et al. [26]. In this method, cellulose acetate (CA) membranes were used to identify the total vibrio colonies from the total bacteria in the Petri dish. For the COPP Assay (Figure 1), the plates with the CA membranes were incubated for 10 min at 37°C. After incubation, the membranes were carefully removed and placed colony side up on a new Petri dish. The membranes were then observed under an ultraviolet (UV) lamp at a long wavelength (364 nm); the fluorescent colonies were counted as they correspond to total vibrio. Colonies were calculated into log colony-forming units (CFU) to determine the total vibrio loads in oyster samples according to Richards et al. [26].
2.5. Chemical Analyses of Oysters
2.5.1. Salt Extractable Protein
Salt extractable protein (SEP) was analyzed using the Biuret method [27]. Tissue homogenate (2 g) with 39 ml of extraction buffer (4.2% lithium chloride (LiCl) in 0.02 M lithium carbonate (LiCO3)) was vortexed and centrifuged for 30 min at 13,000 rpm. The concentration of the protein in the supernatant was measured using a spectrophotometer (General purpose UV/Vis Spectrophotomer, DU 720, Beckman Coulter, Indianapolis, IN, USA) at 540 nm wavelength. Standard curves were generated using 0.5, 1, 2, 3, 4, and 5 ml portions of bovine serum albumin (BSA), and the solution was made up to 5 ml using distilled water and analyzed using the sample methods used for tissue samples. Protein was expressed as SEP % in 100 g of oyster tissue.
2.5.2. Glycogen Content
Oyster homogenate was frozen using liquid nitrogen and then ground into powder. A sample size of 10 mg was weighed from the ground sample. Glycogen concentration was analyzed by the phenol-sulfuric acid method [28]. One mL of 30% potassium hydroxide (KOH) was added to the samples and incubated at 100°C for 20 min and then cooled. 1.5 mL of ethanol was then added to prevent the precipitation of polysaccharides and incubated at 100°C for 15 min. Distilled water was added to bring the volume to 6.6 mL, and 2 mL aliquots with 80% phenol (100 µL) and sulfuric acid (5 mL) were placed in tubes. The samples were incubated at room temperature of 20–21°C for 30 min and then observed using a spectrophotometer at a wavelength of 485 nm (general purpose UV/Vis Spectrophotomer, DU 720, Beckman Coulter, Indianapolis, IN, USA). The glycogen stock was made with 1 : 1 ratio of glycogen powder and distilled water. A standard curve was generated by adding 0.4, 0.8, 1.2, 1.6, 2, and 4 mL portions of glycogen stock, and the final solution was made up to 20 mL with distilled water. Analysis was completed using the sample methods used for tissue sample. Glycogen concentration was recorded as mg glycogen/g of oyster tissue absorbance of the standard [29].
2.5.3. Total Lipid and Fatty Acids
Total lipids were extracted according to the method of Bligh and Dyer [30]. One gram of liquid nitrogen frozen and ground oyster homogenate was used. Three milliliters of 2 : 1 ratio of methanol : chloroform was added and mixed on a rotator (Large 3-D, Thermo Scientific) for 20 min. After adding 1 mL of chloroform and 1.8 mL of distilled water, the samples were placed on a shaker for 5 min. The samples were then centrifuged at 3,500 rpm for 1 min, and the lower layer (organic phase) was transferred into a new tube. Empty tubes were weighed before transferring the lower layer. Total lipids were determined by calculating the difference in weight of the tubes before and after drying under liquid nitrogen.
Transmethylation procedure adapted from Caroline Gott (unpublished data—University of Delaware, Newark, DE) was also used in this study. Five milliliters of potassium hydroxide (KOH) and one milliliter of internal standard (C17) was added, vortexed, and flushed with nitrogen gas. Samples were placed in a water bath at 37°C for one hour and vortexed every 10 min. Two milliliters of acetic acid was added and mixed; further 5 mL of hexane was added, and the top layer was removed. Samples were washed one more time with hexane, and then the top layer was removed and nitrogen flushed. Two milliliters of hexane was added, and samples were then placed in vials for the gas chromatographic analysis (GCA) (GC-2010 Plus, Shimadzu, Japan).
Fatty acid methyl esters (Supelco 37 Component FAME mix) were used as the standard to identify individual fatty acid peaks following GCA. GCA was carried out using a Shimadzu (GC-2010 Plus, Shimadzu, Japan) gas chromatograph with a flame ionization detector, equipped with an AOC-20i auto-injector (injection volume of 1 µl). A SP2560 fused silica column (100 m × 0.25 mm × 0.2 µm) was used for separation. Helium was the carrier gas with a pressure of 440.4 psi. The oven temperature was initially held at 140°C for 5 min, followed by an increase to 240°C at a rate of 4°C and finally held at that temperature for 15 min. Fatty acid profiles were recorded as percentages of individual fatty acids.
2.6. Statistical Analysis
Data obtained from the analyses were tested for skewness and kurtosis, once identified as nonnormal data, nonparametric testing was performed. The Kruskal–Wallis test allowed for identification of significant differences () between treatments while accommodating nonnormal data (IBM SPSS Statistics for Windows, Version 22.0. Armonk, NY). The Mann–Whitney U test, as a nonparametric test, was used in place of an unpaired t-test. Principal component analysis (PCA) was performed using PRIMER 6 (Primer-E Ltd, Plymouth, UK) software to identify the significant environmental variables affecting the oyster microbial quality. values < 0.05 were considered statistically significant.
3. Results and Discussion
3.1. Water Quality
Water quality results throughout the study period showed higher salinity and dissolved oxygen levels in the open water than the closed canal site. Figures 2(a) and 2(b) show the physical water quality parameters from open water and closed canal in both years. The percent dissolved oxygen (DO%) in the open water study site stayed within the base and peak limits, whereas the closed site illustrated a major dip in DO%. Although salinity and temperatures varied between the sites, there were no major differences in the pH of water samples between the two study sites and no significant differences in the pH among the oyster tissue samples. Table 1 illustrates the parameters for optimal water quality conditions for the survivability for marine life including oysters.
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Of all the parameters analyzed for nutrients, only total nitrogen (TN) and total phosphorous (TP) remained well below the maximum levels allowed for optimal oyster survivability (Table 2). The recorded TN was higher at the closed canal study site in July and October of 2014 and was high in the open study site in October 2014 while TP was high at both study sites throughout the study period. These higher concentrations of TN and TP can be attributed due to the environmental events (i.e., horseshoe crab die offs) (personal observation) and the proximity of the sites to farming (i.e., poultry farm activities, residential, and commercial areas for TN and TP).
3.2. Physical Analyses of Oysters
Physical analyses were performed in triplicates per sampling date from June through November. Table 3 shows the Hunter , , and values of the oyster samples. Market oyster samples were highly pale in color, with increased pH, and moisture contents. The market oyster samples had the highest and values in July, August, and September showing a lighter coloration than the oysters collected from open water and closed canal sites.
For color value, significant differences were observed between the treatments (), but there were no significant differences between the study months (). In addition, significant differences between oysters from open water versus market () and closed canal versus market oysters () were recorded, but there were no significant differences between the oysters from open water and closed canal ().
For color , there were no significant differences between the treatments () and the study months (). For color , there were significant differences between the treatments (), but there was no significant difference between the months (). The statistical values for the oysters from open water and market oysters were and the values for open water and closed canal were , whereas the values for closed canal and market oysters were . Although Ozbay et al. [35] studied fish, they explained the increase in expressible fluid (in this study, percent moisture) correlated positively with fading/lighter coloration () and negatively with redness (), resulting market oysters and B∗ values much higher and oyster quality lower. There are strong correlations between the expressible fluid/percent moisture, protein binding, and the Hunter , , and values of the seafood product [35]. A small variation in the oyster pH was recorded among the three samples in relation to water quality conditions (Figure 3(a)). The increase in texture (hardness) was recorded in August for oysters from the closed canal (Figure 3(b)). There were no differences in the texture of oysters collected from the Inland Bays (open water and closed canal) and the market samples (), but there was a significant difference in the texture of oysters between the study months (). Mudoh et al. [36] investigated the effects of storage temperatures (5, 10, and 20°C) on growth of vibrios as well as other microbial, sensory, and textural characteristics of postharvest shell stock eastern oysters (Crassostrea virginica) over a 10-day period. They found no consistent pattern for the pH of oyster homogenates with storage time and temperature while olfactory acceptance reduced with time and increase in storage temperatures. Loss of freshness as judged by appearance and odor was significant during 10-day storage (). They also found that toughness of oysters increased with storage time at 5 and 10°C from days 1 to 3 but was inconsistent after day 7. Their results indicated that the length of storage and temperature had a significant effect on olfactory acceptance of oysters but had an inconsistent effect on texture.
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Percent moisture content was high in warmer months for all oyster samples (Figure 3(c)) that is in accordance with the previous research efforts by Lampilla and McMillin [37]. Moisture content was slightly higher in the market samples than the oyster samples collected from the Delaware Inland Bays. According to the Kruskal–Wallis analysis, there was no significant difference between treatments (). There was, however, evidence of significant differences between the sampling months ().
3.3. Microbiological Analyses of Oysters
The total aerobic bacterial counts (ABCs) in the oyster samples are shown in Figure 4(a). Bacterial counts were high in the warmer months for oysters from all the study sites. The highest bacterial counts were obtained in the market oysters during June, July, August, and the middle of September. When considering the DIB oysters, closed canal samples had the highest bacterial counts in the colder months than the open water samples (end of September, October, and November). Total aerobic bacteria in oysters from the open water (harvestable waters) remained below 6 log CFU/g throughout the study period while closed canal (nonharvestable waters) and market samples reached above 6 log CFU/g. Oyster samples from the closed canal had higher bacteria levels than oysters from the open water. Oysters normally inhabit large numbers of bacteria from their natural environment even after bacteria concentration decreased with decrease temperature [38]. Givens et al. [39] quantified Vibrio vulnificus and Vibrio parahaemolyticus in fish, oyster, sediment, and water using culture-independent (quantitative PCR, qPCR) and culture-dependent (direct plating-colony hybridization, DP-CH) techniques during winter and spring and found higher concentrations of V. vulnificus and V. parahaemolyticus in fish intestine samples greater than oyster, sediment, and water samples. Chase et al. [40] recovered culturable V. vulnificus bacteria most frequently from oyster samples (70%), followed by vegetation and sediment (∼50%) and water (43%) and discussed sediment habitats may serve as seasonal reservoirs for V. vulnificus. As the closed canal has less free flowing water, we expected higher levels of total bacteria in oysters harvested from the closed site because water becomes stagnant frequently.
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Total psychrotropic bacterial counts (PBCs) increased as temperatures increased in warmer months, and oysters from seafood market had higher PBCs than oysters of closed canal and open water (Figure 4(b)). For PBCs, the Kruskall–Wallis test showed significant differences among the study sites and local seafood market () and the study months (). The Mann–Whitney U test also showed significant differences between oysters from the closed canal and local seafood market () and between oysters from the open water and local seafood market (). There were no significant differences between oysters from the closed canal and open water (). Psychrotropic bacteria generally cause winter outbreaks, but these illnesses are largely dependent on the temperature, type of pathogen, and the environment. Studies have reported that even though there are a large number of bacteria in the seawater, they may not be harmful to oysters [41].
Both fecal and total coliform bacterial counts in oysters were higher during the warmer months (Table 4). Based on the statistical analysis, there were no significant differences in total coliform and fecal coliform counts of oysters collected from open and closed waters and market oysters, respectively (), but there were significant differences in total coliform and fecal coliform counts over time (). Mignani et al. [42] evaluated the total and thermotolerant coliform densities in the oyster culture water of Cananeia, SP, Brazil, in relation to environmental variables and tidal variations. In their study, they reported that coliform density increased with increase in rainfall but decreased with increase in salinity. They also found higher average total coliforms during summer (279 MPN 100 per mL) and autumn (70 MPN 100 per mL), and the average increased during winter (209 MPN 100 per mL) and spring (210 MPN 100 per mL). They found the highest average thermotolerant coliforms in spring (206 MPN 100 per mL), followed by summer (79 MPN 100 per mL), autumn (43 MPN 100 per mL), and winter (50 MPN 100 per mL). However, they found no significant differences between seasons and coliform levels for total () and thermo-tolerant coliforms (). According to their assessment, simple diagnosis of environmental conditions of the farm is insufficient to assess water condition during harvesting; therefore, continuous monitoring is necessary to harvest product from healthy water and ensure safe consumption of seafood.
Total vibrio levels were high in warmer months with the highest count in June recorded for the market oysters (Figure 4(c)). Oyster samples from the open water had lower total vibrio levels than oysters from the closed canal. There were no significant differences in total vibrio counts among the treatments (), but there were significant differences over time (). Total vibrio of oysters from the local market seems to be consistently higher during the warmer months. Previous study by Fay et al. [13] provided the first baseline levels for total Vibrionaceae and the relationships between water quality, total bacteria, and total Vibrionaceae in the Delaware Inland Bays. They also reported that total bacteria and total Vibrionaceae levels were related to water temperature and salinity. Total Vibrionaceae levels were found to be related to total phosphorous and inversely related to dissolved oxygen and total nitrogen levels at the bay site. Mudoh et al. [36] found halophilic plate count (HPC: 4.5–9.4 log CFU/g) were highest on day 7 at 20°C, and Vibrio parahaemolyticus (Vp) counts increased over time from 3.5 to 7.5 log MPN/g by day 10 during a 10-day storage trial for the eastern oysters. Based on their results, they confirmed the length of storage, and temperature had a significant effect on bacterial counts, which aligns with our study results. In addition, DePaola et al. [43] reported that when larger numbers of other kinds of bacteria are present in oysters, they have the natural tendency to selectively remove vibrios.
3.4. Oyster Chemical Analyses
Figure 5(a) shows the percent salt extractable protein (SEP%) of the oysters from the open water, closed canal, and the seafood market for the months June to November. The concentration of protein in the oyster samples increased during the warmer months and decreased as temperatures reduced. The highest SEP% was recorded in oyster samples from the open water during August. There were no significant differences in the SEP% of oysters between the study sites (), but there were significant differences in SEP% of oysters analyzed between the months (). The SEP% was low in the market samples during 2013 and was low in the June and July months of 2014. Studies show that oysters and fish have similar compositions such as high protein content and low lipid levels. Reza et al. [44] reported that the protein content in fish decreased during chilled storage. Ozbay et al. [35] concluded that the binding of water to protein in steelhead fillets decreased at lower temperatures during the storage. These results are in agreement with our results, and the decrease in the protein content of market samples may be due to storage. It has been suggested that protein and fat content is reduced during storage because of the loss of proteins and fat due to leaching of water-soluble components from the muscle. There were no significant differences in percent total lipid content of oysters collected from the two study sites and local market () nor were any significant differences in lipid content of oysters analyzed over time () (Figure 5(b)). There were changes in saturated fatty acid (SFA), monounsaturated fatty acid (MUFA), and polyunsaturated fatty acid (PUFA) percentages in oysters from closed canal, open water, and market over time. Oyster samples from open water during the study period from June to November resulted in higher SFA percentages when compared to closed canal and market samples. SFA percentages in all oyster samples increased in the warmer months and declined in the cooler months while MUFA percentages increased in the cooler months and PUFA decreased. This is in agreement with the study by Martino and Maria da Cruz [45] that the sum of SFA in mangrove oysters was significantly lower () in winter than in the other seasons in their study.
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Per Martino and Maria da Cruz [45], monounsaturated fatty acids (MUFA), predominantly consisting of the oleic acid and palmitoleic acid in oysters, were the lowest fatty acid group found in the oysters, and they were significantly lower in spring than in other seasons, confirming the results we obtained in our study.
The MUFA percentages were high in market samples in the cooler months and were lower in the warmer months when compared to closed and open samples (Figures 6(a)–6(c)). Studies by Ren et al. in 2003 [18] reported that lipid content in oyster somatic tissue decreases with water temperatures, and these results are in partial agreement with our study results when considering MUFA levels in oysters. It has also been reported that the compositions of proteins and lipids in oysters change due to the change in their life cycle and their metabolism. It has also been reported that when glycogen reserves decrease, oysters use fat sources in their tissues in order to maintain their metabolic activity, thereby decreasing the lipid content.
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3.5. Correlation between Water and Oyster Quality
For total aerobic bacterial counts, there was a significant difference between the treatments according to the Kruskall–Wallis [K–W] test (). Additional tests of pairwise comparisons (Mann–Whitney U test [M–W]) showed that there were no significant differences in total aerobic bacterial counts of oysters from closed canal and open water sites (). There were significant differences between oysters from closed canal and local seafood market (). There was also a significant difference between oysters from open water and local seafood market (). The Kruskall–Wallis test also showed differences in aerobic bacterial counts between the study months (). Total aerobic bacterial counts were near log 10 CFU/g for the market oyster samples, while Hunter and values were highest for these oysters, confirming that market oysters are poor in quality than the oysters analyzed directly from the Inland bays.
Principal component analysis (PCA) was performed to identify the significant environmental variables that can affect oyster microbiological quality. The relationships between total aerobic bacteria in oysters and water quality (temperature, salinity, DO%, and pH) are shown in Figure 7; this plot explains for about 75% variation among the oyster samples that were analyzed. Temperature was recorded to be high at both open water and closed canal sites whereas dissolved oxygen and salinity were higher at the open water site. Bacterial counts were high in oyster samples from closed canal. This analysis did not include oysters used from seafood market, as there were no water quality measurements to report. PCA analysis shows that temperature (PC1 = 0.456) plays a major role in increasing the total bacterial counts in oysters.
Dissolved oxygen (PC1 = −0.519), salinity (PC1 = −0.440), and pH (PC1 = −0.473) are indirectly proportional to total bacterial counts in oyster samples. Our result is aligned with the study conducted by Chase et al. [40] that the salinity (ranging from 1 to 35 ppt) was negatively correlated with V. vulnificus levels in water and sediments in their study. Ozbay et al. [46] also found during the controlled laboratory experiment that oysters under low salinity treatments consistently yielded the highest total aerobic bacteria and Vibrionaceae levels. Their study confirms that high aerobic bacteria and Vibrionaceae levels are primarily salinity dependent.
Correlation between turbidity, nitrate, total aerobic bacteria, total vibrio, and total psychrotropic bacteria for closed canal, open water and market oysters are provided in Table 5. Total aerobic bacterial counts in relation to turbidity are shown in Figure 8(a). Although there is no significant correlation, there was a positive correlation, when the turbidity was high in the closed canal, the bacterial count in oysters increased. Similarly, correlation between oyster bacterial counts and turbidity of open water was not significant.
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(b)
Figure 8(b) shows the total aerobic bacterial counts in relation to nitrate. Again, no significant correlation was found between the total aerobic bacteria and nitrate for the closed canal site; however, correlation was strong between aerobic bacteria in oysters from open water and nitrate, R = 0.77. This is considered relatively significant for environmental scale research. This infers that as the nitrate levels increased in open waters the bacterial counts in oysters increased. Negative correlation was monitored between total aerobic bacteria and total vibrio counts in both closed canal and open water sites is negative, although this correlation was weak (Table 5). The correlation values for total aerobic bacterial counts and total vibrio counts in the market oysters is positive (0.52), which shows that when the total aerobic bacterial levels increased, the total vibrio levels also increased for market oysters and is significant (). Correlation between total aerobic and psychrotropic bacterial counts in oysters from the market samples is also positive (0.65) (Figures 9(a) and 9(b)), whereas no strong correlation was detected between total aerobic and total psychrotropic bacterial counts in oysters from closed canal and open water.
(a)
(b)
The glycogen content of oysters was low in warmer months and was high in the cooler months and lowest at the end of July and the highest at the end of September as shown in Figure 10. The correlation of water temperature and glycogen content in oysters from the closed canal and open water was not significant (Table 5). These results show that water temperature in both open water and closed canal sites did not have a significant effect on the oyster glycogen content; however, the trend shows that when temperature is high, glycogen content is low. Even though the graph shows that glycogen content increased in September as the temperature declined for closed canal and open water oysters and water samples, this is not statistically significant. Overall, there were no significant differences in glycogen contents between the study sites () or months (). Studies by Ren et al. [18] reported that when phytoplankton levels decreased, the glycogen content decreased, and low glycogen content was observed during the winter months in their study. This shows that the glycogen content of oysters is directly related to the availability of food and may not be solely dependent on the seawater temperatures. On the other hand, Martino and Maria da Cruz [45] studied mangrove oysters, Crassostrea rhizophorae, and found glycogen significantly () higher in oysters in spring (4.4%) and winter (4.2%) than in summer (2.7%) and autumn (2.9%).
As stated by Fish and Wildlife Service [47], moisture content changes even if the oysters are measured immediately after they are shucked or measured later. As they reported, the quality of oyster meat is related primarily to the amounts of protein and carbohydrates. Their ratios change with the season and reproductive cycle and so does the moisture content [45]. Fish and Wildlife Service [47] reported that the percentage of protein decreases to less than 40 percent of the dry weight in late April and May while carbohydrates increase to their maximum, of about 60 percent, during the same time. Protein content is less pronounced because of the increased glycogen storage in the oyster tissue. As a reserve material for oyster, glycogen is stored primarily in the oyster connective tissue of the mantle and labial palp, which is being used during egg and sperm formation; hence, the glycogen reserves are limited by the end of the reproductive cycle. Such glycogen storing is exemplified by oysters in New England waters where glycogen supply reaches its maximum peak during late autumn and early winter and remains high until gonadal development and spawning [47].
Salinity and dissolved oxygen were higher in the open water than the closed canal. The total nitrogen, nitrate, total phosphate, total suspended solids, and turbidity of closed canal and open water increased in warmer months, as nitrate levels increased in open waters, and the total bacterial count in oysters also increased. Total aerobic bacteria and vibrio loads were higher in local market samples when compared to oysters from the open water and closed canal. The market oyster samples are lighter in color and had higher bacterial counts than the open water and closed canal samples. Open water had lower levels of bacteria than the closed and market samples.
Although oysters were sold as fresh in local seafood markets, due to handling stress and transportation time, bacterial counts can increase. As temperature decreases, there is a reduction in bacterial growth and percent moisture in all oysters. In August, with increased temperatures, an increase in moisture and SEP% has been observed. A positive correlation between the moisture content and SEP% of oysters has also been recorded. There seems to be a strong correlation between glycogen, SEP%, and water quality parameters. As the temperature declined in September, percent moisture and SEP% declined, and when glycogen content was increased, alkalinity was slightly higher in colder months in the study sites. Glycogen content is used as a good condition index, and environmental stress causes glycogen content to decrease especially in warmer months [48, 49].
The results from this study are aligned with the previous research suggesting that water temperature is a significant driver for bacterial loads, percent moisture, and salt extractable protein levels [47]. Oysters collected from the open water and closed canal had higher quality than oysters purchased from the local seafood market; however, oysters in open water are in better quality than the oysters from closed canal. Although no specific vibrio species were monitored in this study, our results show that temperature and dissolved oxygen control the total vibrio levels and are aligned with the studies carried by Pfeffer et al. [50] and DePaola [51]. Temperature, salinity, pH, and high levels of dissolved organics are the primary environmental factors that affect bacterial outbreaks. The study by Givens et al. [39] identified more favorable and less favorable environmental conditions for V. parahaemolyticus at mean salinities 4.3 and 13 ppt, respectively. Ramos et al. [52] found a positive correlation between V. vulnificus counts and the seawater temperature and a negative correlation between the V. parahaemolyticus counts and salinity in their study. Although not statistically significant, the results in this research showed an inverse relationship between the total bacterial and vibrio counts in the closed canal and open water compared to the strong relationship detected by Fay et al. [13]. Total vibrio counts may not always be correlated with the total bacterial counts, and this seems to be dependent on the environmental conditions from which the oysters were sampled. Although there are differences in the quality of oysters between the closed canal and open water, market oysters had poorer quality during warmer months. Additional handling and storage of raw oysters in the market seem to have immediate effects on the microbial contaminant loads and physical parameters like color than the chemical parameters.
4. Conclusion
There is an increasing concern for the microbiological safety of oysters and a demand to retain their nutrient quality and visual appearance. Oysters are one of the most desired shellfish products and consumer’s demand sustainable harvest nationwide. In this study, various environmental parameters were measured to study their relationships to oyster quality and safety. Overall, microbial loads are much higher during the warmer months, and oysters from the local market consistently had higher bacterial levels than the natural oysters during warmer months. A positive correlation was observed between oyster bacterial counts and turbidity in the closed canal system. Higher nitrate levels increased bacterial counts in the open water, whereas there were no correlations between the nitrate and the bacterial loads in oysters from the closed canal system. Of various environmental parameters tested in this study, temperature and turbidity correlated with Vibrionaceae densities. Salinity is much less severe and inversely correlated to Vibrionaceae densities in our study. This study displayed relationships between environmental quality in which oysters live, and how additional handling and storage can increase the microbial loads in oysters after harvesting. When microbial loads and physical analyses display significant trends for differences for the oysters from the bays and oysters from the market, chemical parameters of oysters seem to be the last affected by the seasonal changes and also handling and storage in the market. We recommend microbial and physical analyses to be tested for immediate response when studying the impact of environment on oyster quality for consumption. Chemical changes may take longer, and further analyses can be done to understand overall health of the animals. This study will help to understand the trends in the quality of oysters collected from Delaware Inland Bays and the market, and the potential risk of their consumption during warmer months and their quality.
Additional Points
Practical Application. Research focus is to assess the general safety of consuming oysters from local seafood market versus oysters from local coastal waters either close or open for shellfish harvesting. This study displayed relationships between water quality conditions and quality of oysters and how additional handling and storage can increase the microbial loads and worsen physical appearance in oysters after harvesting.
Conflicts of Interest
The authors have no conflicts of interest to declare.
Authors’ Contributions
Dr. Gulnihal Ozbay directed and planned the research, trained the staff, interpreted the data and results, and prepared the manuscript. Dr. Ozbay and Dr. Chintapenta contributed equally to this research. Dr. Lathadevi Karuna Chintapenta assisted with staff training and was involved in research, interpretation of the data and results, and preparation and editing the manuscript. Ms. Talaysha Lingham was involved in the research, writing, and data analysis and interpretation of the results and edited the manuscript. Dr. Stephen Lumor assisted with the fatty acid analysis and staff training. Dr. Jung-lim Lee assisted with the microbial testing of oysters. Dr. Bettina Taylor provided her assessment on the experimental approach and data interpretation. Dr. Shobha Sriharan provided assistance with the water quality data and overall assessment on how environmental parameters are correlated to the total bacteria in the project. Dr. Samuel Besong assisted with the results and overall application of the manuscript.
Acknowledgments
This project was funded by the U.S. Department of Agriculture Evans-Allen Grant Program (no. DELXDSUGO2015), U.S. Department of Agriculture National Institute of Food and Agriculture (NIFA) Grant Program (no. 2013-38821-21246), and NSF EPSCoR (no. EPS-1301765). The authors would like to thank Dr. Haiqian Chen for his assistance with the protocol for the microbial testing of oysters, Ms. Caroline Gott at the University of Delaware for assistance with fatty acid analysis, Dr. Ozbay’s lab group including Eunice Handy, Peta-gay Jackson, Laurieann Phalen, and Katherine Ommanney at Delaware State University for their assistance and support for this research.