Mass mortality is often observed in cultured oysters during the period following spawning in the summer season. To examine the underlying causes leading to this phenomenon, thermotolerance of the Pacific oyster Crassostrea gigas was assessed using pre- and postspawning oysters that were sequentially treated with sublethal (37°C) and lethal heat shocks (44°C). The effects were examined on a range of immune and metabolic parameters in addition to mortality rate. A preventative 37°C significantly reduced oyster mortality after exposure to a second heat shock of 44°C, but in postspawning oysters mortality remained at 80%, compared with < 10% in prespawning oysters. Levels of the 72 kDa and 69 kDa heat shock proteins were low in the gill tissue from postspawning oysters stimulated by heat shock, indicating spawning reduced heat shock protein synthesis. The postspawning oysters had depleted glycogen stores in the mantle tissue and reduced adenylate energy charge after heat shock, indicative of lower energy for metabolic activity. A cumulative effect of spawning and heat shock was observed on the immunocompetence of oysters, demonstrated by reduced hemocyte phagocytosis and hemolymph antimicrobial activity. These results support the hypothesis that the energy expended during reproduction compromises the thermotolerance and immune status of oysters, leaving them easily subject to mortality if heat stress occurs in postspawning stage. This study improves our understanding of oyster summer mortality and has implications for the long-term persistence of mollusks under the influence of global warming.
- induced thermotolerance
- heat shock proteins
- energy trade-offs
- metabolic activity
due to rising concerns associated with global climate change (25, 35), there has been regained interest in the physiological mechanisms that animals use to tolerate extreme heat and adapt to thermal changes in their natural environment (52). Sessile intertidal invertebrates are of particular interest, because they can be exposed to extreme environmental fluctuations, including temperature changes during cyclic periods of emersion and immersion (28, 53). As an intertidal bivalve, the Pacific oyster Crassostrea gigas is an important environmental bioindicator, in addition to being one of the principle aquaculture species of economic value. In recent decades, mass summer mortality in oyster aquaculture has become a widespread phenomenon (5, 8, 21, 32, 49). It has been estimated that up to 50% of the harvestable crop can be lost in a given year, and these losses can be even higher in some areas (8, 55). One hypothesis to explain the high rate of summer mortality is a negative synergistic interaction between high temperature and the energetic cost of spawning (4, 11, 47, 49). Therefore, the motivation behind this study is a desire to further understand the impact of spawning on molluskan thermal tolerance. Specifically, we aimed to gather evidence from physiology and immunology to identify discrepant responses to temperature elevation between prespawning and postspawning oysters, and to assess the biological reasons for summer mortality.
Temperature has long been recognized as a key environmental factor that influences all physiological processes in oysters (40). As proteins must be sufficiently labile to interact with their substrates, they are often highly susceptible to protein-denaturing stresses, such as elevated temperature (24, 41). Heat shock usually results in a concomitant reduction in the synthesis of most proteins (50, 61). However, under thermal stress, heat shock proteins (HSP) perform critical protein-stabilizing functions (36, 41, 62) and thus play a crucial role in the development of thermotolerance (12, 36). HSPs can also participate in the preservation of essential defense functions of cells and contribute to resistance to disease (61). Previous studies have demonstrated that oysters have some ability to naturally tolerate thermal shock (12, 24), as they exhibit adaptive plasticity in the expression of HSPs, a response known as induced thermotolerance (17). HSP70 is the primary family of HSPs that is responsive to thermal stress. The thermal limits of oysters have been shown to correlate with changes in isoform expression in the HSP70 family (12, 13). Although there are a few isoforms of HSP70 identified in oysters (e.g., 69, 72, and 77 kDa), one isoform (69 kDa) has been acknowledged as an effective indicator of oyster-induced thermotolerance (12, 24). Thermal adaptation in oysters is known to be dependent on hydrographic and biological factors, including ambient temperature and reproductive stress (12, 15, 24). However, the impact of spawning on the expression of HSP70 isoforms under temperature stress has not been specifically investigated to date.
With elevated temperatures, animals need to increase metabolism to ensure an adequate energetic supply for survival (45). Beyond the thermal optimum, the scope for aerobic respiration may disappear, and transition to anaerobic metabolism occurs due to insufficient oxygen supply (40, 51, 52). If this process is incomplete or impossible, thermal tolerance will be diminished and mortality can ensue. In summer, oyster spawning is triggered by elevated temperature and is concomitant with energy expenditure (5, 44). The conflict of energy demand between thermal adaptability and spawning activity naturally leads to the question of whether thermotolerance in oysters is suppressed by spawning activity. When exposed to stressful stimuli, animals typically divert bioenergetic resources away from nonessential functions (e.g., reproduction) and redirect resources to combat, adapt, and overcome the stressful stimuli (39). However, after spawning, there may not be sufficient energy stores to facilitate the usual coping response, including the synthesis of HSPs.
Glycogen reserves play a central role in supplying energy for gametogenesis and the mobilization of energetic reserves in oysters (4, 57, 60). After spawning, glycogen storage reaches a minimum (5, 49), and thus postspawning oysters may be more susceptible to thermal stress. Impairment of the adenosine nucleotide balance will also result in a reduction of available energy and, consequently, physiological stress (33). Adenylates (such as adenosine mono-, di-, and triphosphates, i.e., AMP, ADP, ATP) represent the pool of metabolic energy in tissues and the balance between these adenosine nucleotides regulates cellular metabolism (16, 33). The adenylate energy charge (AEC) is a ratio of ATP to the total adenylate nucleotide pool (2), used as an index of metabolic activity and an indicator of metabolic potential available to the cell (34, 59, 65). Determination of glycogen storage and AEC levels will help establish whether postspawning oysters are in a fragile metabolic condition when faced with heat stress.
Physiological stress in marine mollusks is also implicated in outbreaks of infectious diseases (9, 21, 54). The manifestation of disease involves an interaction between the pathogen, environment and the physiological status of the host (3, 30). The growth of many marine pathogens has been correlated to elevated temperatures (25, 54), including marine Vibrios, which are often associated with disease in both the larval and adult stages of bivalves (10, 21, 38). Elevated temperatures can also impact directly on the immune defense system of marine animals (10, 21, 27, 54). The immune system of mollusks comprises both cellular and humoral functional components (22, 30). Cellular immunity is implicated in the activity of hemocytes, with phagocytosis providing the primary line of cellular immunity (48, 63). The humoral components of mollusk immunity include lysozyme activity, the phenoloxidase system, and other antimicrobial proteins and peptides (1, 48, 56), which all require energy for production. These cell-mediated and humoral components of the immune system effectively interact, whereby hemocytes synthesize and release antimicrobial molecules into the hemolymph, some of which can then act as opsonins promoting hemocyte phagocytic activity (9). Elevated temperature has been shown to inhibit the activity of hemocytes, phenoloxidase, and lysozyme in oysters leading to increased susceptibility to disease (14, 20, 27, 30). However, it is still not clear whether the lower energy supplies available after spawning can further compromise immune function in oysters.
In South Australia, heat stress occurs during summer, where intertidal organisms can be exposed to air temperatures in excess of 45°C for prolonged periods (>24 h) during low dodge tides (6). Similar high temperature is also reported in another geographic location (8) and these extremes in temperature are likely to become more frequent as a result of global warming. Therefore, this study was designed to investigate the impact of spawning on thermotolerance in the Pacific oyster during summer season with extreme temperature occur. As induced thermotolerance involves complex and robust regulatory mechanisms (17), heat stress was evaluated by measuring oyster mortality, metabolism, and immunocompetence at the cellular level. The synergistic effect between heat stress and spawning was examined using pre- and postspawning oysters subject to sublethal and lethal heat shock. This should reflect the likely consequences of spawning under thermal stress, thus contributing to our understanding of summer mortality in oysters.
MATERIALS AND METHODS
Pacific oysters C. gigas (shell length, 9–10 cm) were obtained from the same stock in Ceduna, South Australia, and acclimatized for a week at 15°C at the Aquatic Science Centre with the support of the South Australia Research and Development Institute. Because the proposed heat shock on ripe oysters could result in spawning, the heat shock trials on prespawning oysters were conducted in November 2005 when their gonads were not fully developed. In late December, 3 days after the ripe animals were spawned through thermal stimulation at 28°C, heat shock trials on postspawning oysters were commenced. Both pre- and postspawning oysters were starved for 24 h at 15°C prior to administering the heat shock.
Heat shock treatment.
According to Shamseldin et al. (58) and Clegg et al. (12), 37°C and 44°C are appropriate for administering sublethal and lethal temperature shock, respectively, in Pacific oysters. Consequently, in this study oysters were transferred directly from 15°C to 37°C (±0.1°C) seawater for 1 h for a sublethal heat shock treatment. During heat shock, animals were laid in a single layer on a perforated disk and the seawater was continually aerated, while maintaining the desired heat shock temperature. The oysters were then transferred back to 15°C seawater. Five days after 37°C heat shock, the oysters were shocked again at 44°C for 1 h and then transferred back into 15°C seawater.
Oyster mortality was first tested using three replicate batches of pre- and postspawning oysters for each treatment group, with 40 individuals per replicate with the above heat shock protocol. The first treatment group received the 37°C sublethal heat shock only. The second group that initially received the 37°C heat shock was then treated with a 44°C lethal heat shock 5 days later. The third group received the 44°C lethal heat shock only. In the control groups, oysters were kept at 15°C and received no heat shock. The mortality data presented the proportion of oysters that died 5 days after each treatment.
For physiological and immunological analysis, another batch of oysters was used for both pre- and postspawning oyster in treatment and control groups. Three replicates were used per group and each replicate contained 60 oysters. Due to the intensive sampling regimen required, only one heat shock treatment was used in this component of the study to assess the cumulative effects from sublethal and lethal heat shock. The treatment received was 37°C and 44°C shocks, whereas the control group comprised oysters without either heat shock. Five oysters were sampled from each replicate control and treatment group on days 0, 1, 2, and 3 after the 37°C heat shock, and then again on days 6 and10after the 44°C heat shock. The oysters were shucked, mantle and gill tissue were dissected out separately, and the oysters were immediately submerged in liquid nitrogen (total sampling time, <30 s). Later these tissues were transferred to −80°C for freezer storage before subsequent analyses (within 1 mo). In the meantime, hemolymph was collected quickly from the oysters’ pericardial cavity with a 1-ml sterilized syringe. To minimize interanimal variability, each replicate sample consisted of five pooled animals (but less on day 10 in postspawning oysters due to high morality).
Gill tissues for Western blots were prepared according to Clegg et al. (12). Briefly, these samples were homogenized on ice in glass tissue grinders containing potassium gluconate buffer (5 mM MgSO4, 5 mM NaH2PO4, 40 mM HEPES, 70 mM gluconic acid, 150 mM sorbitol; pH 7.5) at a concentration of 100 mg of tissue per milliliter of buffer. Homogenates were prepared for electrophoresis by mixing 1:1 in a 2× SDS sample buffer (6% SDS, 40% sucrose, 20 mM Tris pH 6.8, 0.15% bromophenol blue) and boiling for 5 min. Each solubilized sample (10 μg protein) was resolved on a 12% SDS-PAGE. To normalize between blots, multiple aliquots of a single sample of heat-shocked oyster gills were made as a standard. Proteins were transferred onto polyvinylidene difluoride membrane (GE HealthCare) and probed by using a monoclonal rat anti-HSP70 family antibody (MA3–006; Affinity Bioreagents, Golden, CO), and a horseradish peroxidase-conjugated goat secondary antibody (Sigma, St. Louis, MO). The signal was detected by chemiluminescence by using SuperSignal West Pico Chemiluminescent Substrate reagents (Pierce) by exposing blots to the photographic Kodak Biomax maximum resolution film. Densitometric analysis of the films was performed by scanning on VerseDoc (model 4000) and the digital images were analyzed with Quantity-One to quantify band intensities of the HSP70 isoforms. Given the inherent variation between film developments, data were normalized to standard samples included in each experimental protocol.
Glycogen and AEC assay.
The analysis of glycogen and AEC was conducted in the mantle tissue extraction, which was achieved using perchloric acid. Briefly, the frozen mantles (−80°C) from five oysters were ground into a fine powder on dry ice, and 5 ml of 0.6 M ice-cold perchloric acid was then added to 1 g fine powder (n = 3). Samples were vortexed for 30 s and left on ice for 10 min, followed by centrifugation at 1,500 g for 10 min at 4°C. Then 1 ml of the supernatant was removed for glycogen analysis. Another 1 ml retained supernatant for AEC analysis was prepared according to the method of Shofer and Tjeerdema (59). All samples were stored at −80°C prior to analysis.
Glycogen analysis was done using an iodine glycogen method (37). This procedure involved adding 1.3 ml iodine solution (1.92 ml I2KI to 500 ml saturated CaCl2 solution) to 0.2 ml of sample solution in a microcuvette. The samples were incubated at 25°C room temperature for 20 min before analysis at 460 nm on a Unicam UV-Visible spectrometer. Purified oyster glycogen (MP Biomedicals) (0, 0.1, 0.4, and 0.7 mg/ml) was used to create a standard curve. The final glycogen concentration was expressed in milligrams per gram of wet tissue weight.
The AEC analyses were carried out by HPLC using a Waters 2695 separation module (with Waters 2487 Dual Lambda Absorbance Detector) equipped with a reverse-phase C18 column (25 × 4.6 mm, 4.6-μm particle size fitted with a C18 guard cartridge, 20 mm × 4.6 mm, 4.6-μm particle size). The mobile phase consisted of 200 mM phosphate buffer (adding 1-liter distilled H2O to 5.23 g of K2HPO4 and 2.72 g of KH2PO4; pH 7.0) and methanol (99.8%). Each sample (20 μl) was injected into the column and allowed to run for 15 min. After sample injection, the column was eluted with 95% phosphate buffer, followed by a 10-min linear ramp to 40%, which was held steady for an additional 1 min and then returned to the starting condition over 4 min with 1 ml/min flow rate. The detector wavelength was set to 260 nm for nucleotide detection. A 5-min delay was added between analyses to permit column equilibration. Nucleotides were identified by their retention times and quantified by peak areas relative to those of the external standards of AMP, ADP, and ATP (Sigma). The AEC was calculated as defined by Atkinsons (2) using the formula: AEC = (ATP + 0.5 × ADP)/(ATP + ADP + AMP).
Hemolymph from five individual Pacific oysters per replicate was stored in a tube on ice. A subsample (300 μl) was then mixed with an equal volume of filtered seawater (0.2 μm) in a flow cytometer tube (n = 3). To assess phagocytic activity of hemocytes, fluorescent beads (Fluoresbrite YG Microspheres, 1.75 μm; Polysciences), as 4 μl of a stock suspension of 2.5% solids in filtered seawater per milliliter, were also added to each tube. The samples were analyzed on a FACScan flow cytometer (488 nm laser; Becton Dickinson, San Jose, CA) after 60-min incubation at 20°C in darkness (23). Phagocytic activity was expressed as the percentage of cells that ingested at least three fluorescence beads.
The remaining hemolymph was frozen in liquid nitrogen until ready for use in the antimicrobial activity assay, adapted from Vakalia and Benkendorff (64). Cultures of the marine pathogen Vibrio harveyii (obtained from the Tasmania Department of Primary Industries and Fisheries and maintained at −80°C in 15% glycerol) were prepared by streaking onto nutrient agar and incubating overnight at 37°C. A single bacterial colony from the culture plates was inoculated into a McCartney bottle containing sterile nutrient broth (1 g NaCl, 2 g yeast extract, and 1 g peptone per 100 ml distilled H2O), followed by incubating overnight at 37°C on an orbital mixer shaker (Ratek) at 200 rpm. The cultures were diluted to OD600nm = 0.1 on a spectrophotometer (Metertech UV/VIS SP8001) and returned to exponential growth phase (OD600nm = 0.18–0.2) prior to use in antimicrobial assays. The defrosted hemolymph was centrifuged at 500 g for 5 min to pellet the hemocytes, and then 90 μl aliquots of plasma were pipetted into a 96-well plate in triplicate. Then 10 μl of V. harveyii in exponential growth culture was added into each well. Negative controls consisted of 90 μl hemolymph incubated with 10 μl nutrient broth, and positive controls comprised 10 μl of V. harveyii in 90 μl of nutrient broth. After a 30-min incubation, 20 μl of CellTitre 96 Aqueous One Solution (Promega) was added to each well, and then the plates were returned to the incubator (37°C) for 2 h or until development of the red formazan product in control wells. The absorbance was measured at 492 nm using a 96-well plate reader (Spectra Max 250) (64). The background absorbance from hemolymph broth controls was subtracted from the treatment wells, and then cell viability was calculated as a percentage of the absorbance in positive control cultures. Antimicrobial activity was expressed by the formula: 1 − cell viability.
Data analyses were performed on SPSS (v. 14.0) using repeated-measures ANOVA to test the effects of heat shock and spawning. Data were divided into two time periods, i.e., 37°C heat shock (days 0-3) and 44°C heat shock (days 6-10). The Bonferroni test was used for multiple comparisons of significant treatment effects. Since the 44°C was imposed on day 5, the effect of 44°C heat shock was assessed using a Student's t-test between days 3 and 6. Oyster mortality was analyzed using two-way ANOVA. Data were transformed by square root or logarithm to meet assumptions of normality and homogeneity of variances if necessary in the above ANOVA analyses. Values are given as means ± SE. A significance level of P = 0.05 was used for all tests.
During heat shock, a similar behavior in response to the heat shock was observed between pre- and postspawning oysters. Most oysters gaped for 1–2 h when they were moved back to the 15°C seawater after receiving a 1-h 37°C heat shock. In contrast, the shells remained open for at least 3 h upon receiving a 1-h 44°C heat shock.
During the experimental period, no mortality occurred in control groups. However, 100% mortality was detected after applying a single 44°C heat shock in both pre- and postspawning oysters. There was a significant interaction detected between spawning and the three heat shock treatments (P < 0.001, Fig. 1). The 37°C heat shock led to about 6.7% mortality in postspawning oysters, but did not cause mortality in prespawning oysters (Bonferroni test, P < 0.001, Fig. 1). The preventive 37°C treatment also reduced oyster mortality after exposure to 44°C heat shock, especially in prespawning oysters where mortality was about eight times lower than in postspawning oysters (Bonferroni test, P < 0.001, Fig. 1).
Determination of HSP isoforms.
There were two isoforms of HSP identified in this study; one with a molecular mass of 72 kDa (HSP72), and the other one with 69 kDa (HSP69) (Fig. 2). The HSP72 isoform was detected in each treatment, while the HSP69 was only expressed in heat-shocked oysters.
The protein expression of HSP72 was affected by spawning and heat shock (Fig. 3A, Table 1). ANOVA revealed that a significant interaction occurred between spawning and the sublethal (P < 0.001) and lethal (P = 0.003) heat shock treatments relative to controls (Table 1). In the absence of heat shock, there was no difference in HSP72 expression between pre- and postspawning oysters (Bonferroni test, P > 0.05, Fig. 3A). In postspawning oysters, the HSP72 expression was not influenced by heat shock (P > 0.05). However, prespawning oysters expressed significantly more HSP72 when heat shocked (Bonferroni test, P < 0.001, Fig. 3A). Repeated-measures analysis revealed a significant interaction between time and 37°C shock treatment (P = 0.007, Table 1). In prespawning oysters, heat shock significantly stimulated the expression of HSP72, which increased more than twofold after 37°C shock (P < 0.001) and continued to increase after 44°C shock (Fig. 3A). In postspawning oysters, HSP72 expression increased slightly immediately after a 37°C shock but then remained consistently low from days 1 to 10 (Fig. 3A).
The HSP69 expression was also found to be affected by spawning (Fig. 3B). After the sublethal heat shock, a significant interaction was detected between spawning and time (P = 0.018). There was no difference in HSP69 expression between pre- and postspawning oysters initially (P > 0.05, Fig. 3B). The amount of HSP69 detected in prespawning oysters rapidly increased to about threefold of the original levels 2 days after a 37°C shock, but then started to drop off again by day 3 (Fig. 3B). After 44°C shock, the levels of HSP69 again increased, remaining at a high level between days 6 and 10 (P > 0.05). By comparison, the levels of HSP69 were consistently low in postspawning oysters (P < 0.001, Table 1, Fig. 3B). Levels of HSP72 and HSP69 as revealed by t-tests did not significantly differ between days 3 and 6 for any of the treatment groups (t-test, P > 0.05, Table 2).
Determination of glycogen level in mantle tissue.
The mantle glycogen levels varied from <2 mg/g in postspawning oysters to over 8 mg/g in prespawning controls (Fig. 4A). After the initial 37°C shock, glycogen was significantly reduced by spawning (P < 0.001) and heat shock (P = 0.001, Fig. 4A). However, after the subsequent 44°C shock, ANOVA showed that the interaction between spawning, heat shock, and time was significant (P < 0.006, Table 1). From days 6 to 10, the glycogen level in prespawning oysters increased significantly in mantle tissue (Bonferroni test, P < 0.001, Fig. 4A). Glycogen levels remained consistently low in postspawning oysters, but in heat-treated prespawning oysters, the glycogen level significantly dropped by day 6 compared with day 3 (t-test, P = 0.029, Table 2, Fig. 4A).
Determination of AEC in mantle tissue.
Postspawning heat-shocked oysters had the lowest AEC values throughout the study after heat shock (Fig. 4B). After the 37°C shock, ANOVA revealed that AEC value in the mantle tissue was significantly lowered by both spawning and 37°C shock (P < 0.001, Table 1, Fig. 4B). After 44°C heat shock, the interaction effect of spawning, heat shock, and time was significant (P = 0.003, Table 1). On day 10, AEC values in pre- and postspawning oyster controls was similar and the heat-shocked prespawning oysters were also not significantly different to the control (Bonferroni test, P > 0.05). The AEC value in postspawning oysters dropped to a level between 0.3 and 0.5 from days 6 to 10, whereas it was maintained between 0.5 and 0.7 in prespawning oysters (Fig. 4B). The t-test showed that the AEC value was significantly diminished by day 6 after the cumulative 37°C and 44°C shock in both pre- and postspawning oysters (P < 0.05, Table 2) but remained constant between days 3 and 6 in control groups (P > 0.05).
Phagocytic activity of oyster hemocytes.
ANOVA revealed that hemocyte phagocytosis was significantly diminished by 37°C shock and spawning (P < 0.001, Fig. 5A, Table 1), but the impact of 37°C heat shock varied over time (P = 0.003, Table 1). In control groups the phagocytic activity remained relatively constant over the first 3 days, whereas after a 37°C shock the activity dropped significantly below control levels, reaching a minimum on day 2 but then increasing on day 3 (Bonferroni test, P ≤ 0.025, Fig. 5A). After exposure to the 44°C shock, the interaction between spawning and heat shock became significant (P = 0.009, Table 1). Heat shock treatment had a large impact on the phagocytic activity of postspawning oysters such that the hemocyte phagocytic activity was close to zero ondays 2, 6, and 10 (Fig. 5A). The impact of spawning was still significant in the control (Bonferroni test, P = 0.027). There was no treatment effect on phagocytosis between days 3 and 6 (t-test, P > 0.05, Table 2).
Antimicrobial activity in oyster hemolymph.
The hemolymph antimicrobial activity was significantly affected by an interaction between spawning and 37°C shock (P = 0.025, Table 1). In the controls, the antimicrobial activity was similar between pre- and postspawning oysters (Fig. 5B). However, after 37°C shock treatment, postspawning oysters had much lower antimicrobial activity than prespawning oysters (Bonferroni test, P = 0.002), whereas prespawning oysters were not influenced by 37°C shock (Bonferroni test, P = 0.551). After 44°C shock, the antimicrobial activity of the hemolymph was significantly reduced by both spawning and heat shock (P ≤ 0.001, Table 1). Relative to untreated prespawning oysters, the largest drop in antimicrobial activity was observed after 44°C shock in postspawning oysters, followed by postspawning controls and then heat-shocked prespawning oysters (Fig. 5B). Between days 3 and 6, there was a significant reduction in the antimicrobial activity of hemolymph for both pre- and postspawning oysters with 44°C heat shock (P = 0.038 and 0.002 respectively, Fig. 5B, Table 2).
Our study has revealed that postspawning oysters are significantly less tolerant to heat shock than prespawning oysters. The observed synergistic impacts of heat shock and spawning on induced thermotolerance, energy reserves, and immunocompetence provide a valid explanation for the phenomenon of summer mortality in cultured mollusks, such as the Pacific oyster. Although all oysters in this study were recorded to have some adaptability of induced thermotolerance via the expression of HSP69, the ability to resist sublethal and lethal temperature shock was reduced in postspawning oysters, as indicated by significantly higher mortality and a reduced expression of critical HSPs. This reduced thermotolerance in postspawning oysters coincides with greatly reduced energy reserves for protein synthesis. The low energy reserves also correspond with lowered immunocompetence in spawned oysters, thus increasing their vulnerability to heat stress.
Glycogen heat shock typically causes protein denaturation, thus inducing protective HSPs (15). In this study, two isoforms of HSP were identified in the gill tissue, as was previously observed by Cruz-Rodriguez and Chu (15). These are the cognate isoform HSP72 and inducible isoform HSP69. The level of cognate HSP70 isoforms indicates the capacity for protein synthesis, whereas the levels of inducible isoforms may be more accurate indices for heat stress adaptation (18, 19). We found that both HSP72 and HSP69 had heat shock-dependent stimulation in prespawning oysters, but not in postspawning oysters. The low level of HSP72 in heat-treated postspawning oysters indicates a reduced capacity for protein synthesis, as well as lowered ability to withstand heat shock. Furthermore, although HSP69 was detected in postspawning oysters, the level was much lower than that in prespawning oysters and did not increase in response to a lethal heat shock. In agreement with Clegg et al. (12), our study suggests that induced thermotolerance in oysters correlates with HSP69 expression but further suggests a role for the inducible expression of HSP72. As suggested by Kregel (36), morbidity and mortality from thermal shock are due to the dysfunction of some critical target tissue that is heat sensitive and vital to the animal. Impairment of HSP synthesis in the gill and the consequent protein denaturation is likely to lead to hyperthermic killing (41), and our study supports the idea that this is a critical factor in Pacific oysters. Ultimately, dysfunction in the gill, the most important respiratory tissue of most marine animals, will cause a mismatch between oxygen delivery and the ability to respond to heat shock, which finally leads to the collapse of physiological function (51, 52).
The cost of protein synthesis is 18–26% of the energy budget in most ectotherms (31), so the metabolic cost of heat damage to proteins could significantly affect an oyster's energy budget (29). The accumulation of glycogen in mantle tissues provides the main energy supply for the mobilization of energy reserves (5). However, glycogen stored in the mantle and the gonads also provides the energy required for reproduction. Our study indicates that the risk of mortality in Pacific oysters is concomitant with the postspawning period when glycogen levels reach their lowest levels. This is consistent with previous studies by Mori et al. (47) and Perdue et al. (49). We also found that the expenditure of glycogen reserves was coincident with heat shock treatment. Given that higher temperature causes insufficient oxygen levels in ectothemic animals (51, 52), more energy reserves are catabolized for anaerobic metabolism, thereby indicating that oyster thermal tolerance is partially dependent on the glycogen storage. However, despite greatly reduced glycogen reserves, postspawning oysters only suffered 6.7% mortality after sublethal heat shock, implying that postspawning oysters are still able to mobilize their residual glycogen reserves to withstand mild short-term heat shock. However, in the event of an extreme (lethal) heat shock, postspawning oysters exhibited higher mortality and appear unable to mobilize sufficient glycogen reserves to meet the energy costs required for withstanding heat shock.
The more stressed an animal becomes, the more energy it uses to counteract the stress, thus lowering its AEC value (65). In this study, spawning and heat shock were concomitant with decreasing AEC value. After heat shock, the AEC value in prespawning oysters was maintained within the range of 0.5 to 0.7, which is the suboptimal stress range according to Cattani et al. (7). This indicates that prespawning oysters can maintain normal energy metabolism at a cellular level regardless of induced thermotolerance. However, postspawning oysters had much lower AEC levels, especially after lethal heat shock. The low AEC values and high mortality recorded in postspawning oysters subject to lethal heat shock, are consistent with a study by Shofer and Tjeerdema (59) where an AEC value between 0.3 and 0.5 was suggested to be a critical range from which the capacity to recover from stress is impossible (59). This critical limit may be attributed to the transition to anaerobic metabolism when temperature is beyond thermotolerance, which leads to insufficient cellular energy supply (52). In this stressful situation, the physiological tradeoffs will divert energy reserves toward essential maintenance (40), although the residual energy reserves (glycogen) seem not to be adequate for energy modulation in postspawning oysters.
Our study has also demonstrated that both heat shock and spawning have significant impacts on cell-mediated immunity. It has been previously suggested that heat shock is life threatening to oysters due to lethal effects on hemocytes (20, 26). Here we have shown that exposure to sublethal heat shock, in both pre- and postspawning oysters significantly reduces hemocyte phagocytosis, and the effect is further augmented by a lethal heat shock. The impact of heat shock was greater in postspawning oysters, which had significantly lower phagocytosis rates than prespawning oysters and were approaching zero after a lethal heat shock. Without heat shock, the hemocyte phagocytic activity in postspawning oysters indicates that they can still maintain some effective cellular immunity despite lower activity compared with prespawning oysters. However, it has also shown that spawning can reduce the number of circulating hemocytes in oysters (11). The number of hemocytes and the resilience of their activity provide important indicators for the surveillance of immune health in oysters (61). This study demonstrates that the impact of spawning on phagocytosis can last more than 10 days, during which time the oysters will be extremely vulnerable to infectious disease.
Theoretically, the shifts in hemocyte activity should also lead to changes in antimicrobial activity in the hemolymph. It is not surprising then that a dramatic reduction in antimicrobial activity was seen in postspawning oysters after lethal heat shock. Prespawning oysters subject to lethal heat shock also showed a significant reduction in antimicrobial activity. After only the sublethal heat shock, however, the effects were subtle, and only the postspawning oysters suffered significantly reduced activity compared with untreated controls. Although antimicrobial agents are synthesized by the hemocytes, they are stored in the cells as reserves and only usually were released after stimulation by microbial infection or other similar challenges (1, 9). Consequently, hemolymph antimicrobial activity may counteract the shortcomings of cellular immunity in postspawning oysters, if there is only a mild temperature stress. However, the significant decrease in antimicrobial activity in both pre- and postspawning oysters after 44°C treatment further suggests that oysters are under immunosuppression during induced thermotolerance. As oysters are exposed to various stressful situations in the environment (13), any immunosuppression would compromise their defense system against opportunistic parasites and pathogens (25). Considering the fact that many marine pathogens appear to have increased growth rates at higher temperatures (21, 25, 54), the decline of antimicrobial activity due to spawning could make oysters much more susceptible to mortality caused by disease when heat shock occurs.
Overall, this study illustrates some of the underlying causes for summer mortality in Pacific oysters. Postspawning oysters are highly vulnerable to temperature stress and have a reduced ability to resist infectious disease. Initial results from a large-scale collaborative investigation into Pacific oyster summer mortality by the French Government (MOREST program) indicate that increased vulnerability can occur at temperatures as low as 19°C in their farming area, depending on genetic factors and the intensity and duration of spawning (46). Consequently, further research into the physiological and immunological responses of C. gigasfrom different genetic origins and various environmental conditions are warranted. Huvert et al. (32) suggested that the summer mortality in Pacific oysters resulted from immunological and/or energetic dysfunctions. Our study supports a combination of both of these factors, in addition to a reduced ability to synthesis-inducible HSPs. As prespawning oysters have more energy reserves (glycogen), they can promote sufficient HSP production to withstand heat shock, which was also indicated by the relative high-energy metabolic activity (AEC). On the other hand, the lack of energy reserves available to postspawning oysters led to a diminished capacity to respond to heat shock and therefore reduced thermotolerance. Since oyster hemocytes are also responsible for energy and nutrient transport (56), any physiological dysfunction also necessitates a downregulation of immune function. Hence, after spawning, the oysters need to recover and start accumulating energy for thermal adaptation. This recovery process is long (in excess of 10 days) and will depend on the food availability in the ocean (42). The recovery process can also be inhibited by encountering stress stimuli, because stress causes a redirection of bioenergetic resources toward adaptive physiological functions (43). Given the fragile metabolism and immune system during the recovery period after spawning, high summer temperatures make postspawning oysters more susceptible to elevated temperatures and opportunistic pathogens, inducing mass mortality. It is possible that these incidents of mass mortality in marine organisms including species such as abalone, mussels, and clams, which have great economic value, will increase globally coincident with unpredictable extremes in weather patterns. The Pacific oyster offers a good opportunity to monitor the impacts of global warming on ocean productivity, as an important worldwide aquaculture species whose reproduction is triggered by temperature increases.
This work was supported by International Postgraduate Research Scholarship, Flinders University Research Scholarship (to Y. Li), and the Marine Innovation South Australia Initiative (to X. Li).
We are grateful to Tong Chen from Biological Sciences School for Western blotting technical assistance, Dr. Daniel Jardine from Advanced Analytical Laboratory in Flinders University for HPLC technical assistance, Dr. Jeremy Carson from the Fish Health Unit, Department of Primary Industry and Fisheries, Tasmania, for kindly providing the strain of marine bacteria, and Gary Zippel from Zippel's Enterprise in Ceduna, South Australia for supplying Pacific oysters. Suggestions and comments of Jim Mitchell (Flinders University), Steven Clark (South Australia Research and Development Institute), and three anonymous reviewers are greatly appreciated.
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- Copyright © 2007 the American Physiological Society