The effects of temperature and salinity on apoptosis of Crassostrea virginica hemocytes and Perkinsus marinus.

ABSTRACT: Oyster culture, which is a large part of the North American aquaculture industry, is greatly hindered by the parasite Perkinsus marinus. The different oyster defense mechanisms and how the parasite seems to evade them are not fully understood, and the role of apoptosis as an oyster defense mechanism remains unclear. Apoptosis of eastern oyster (Crassostrea virginica) hemocyte subpopulations (granulocytes and hyalinocytes) was quantified at the single cell level using flow cytometric Annexin-V and TUNEL assays. The influences of habitat salinity (540 mOsm [18.9 [per thousand]] and 910 mOsm [31.9 [per thousand]] for oysters and 488 mOsm [17.1 [per thousand]] and 810 mOsm [28.4 [per thousand]] for P. marinus) and temperature (14.5[degrees]C and 25[degrees]C for oysters, 14.4[degrees]C and 22.5[degrees]C for P. marinus) on apoptosis frequencies were evaluated experimentally on the host hemocytes and the parasite. Apoptosis frequency was higher in granulocytes than hyalinocytes, regardless of temperature or salinity. Salinity affected apoptosis of granulocytes and hyalinocytes, with more apoptosis at 910 mOsm than at 540 mOsm (at 14.5[degrees]C and 25[degrees]C). Parasite apoptosis frequency was significantly higher on culture at the low salinity (488 mOsm) at both low and high temperatures (14.5[degrees]C and 22.5[degrees]C). Temperature did not significantly affect either hemocyte or parasite apoptosis frequencies. Investigation of apoptosis as an oyster defense mechanism and how temperature and salinity affect C. virginica and P. marinus apoptosis may lead to a better understanding of host/parasite interactions and the mechanisms leading to disease susceptibility or resistance. Such information could lead to better management strategies that may result in reduced disease morbidity and mortality for oyster producers.

KEY WORDS: apoptosis, Crassostrea virginica, Perkinsus marinus, salinity, temperature

INTRODUCTION

Commercial oyster culture substantially contributes to the North American aquaculture industry but is thwarted by the protozoal parasite P. marinus, which causes extensive economic losses (Andrews 1988). Although oyster resistance to P. marinus has been linked to cellular and humoral defense mechanisms, there is debate as to the relative contributions of each to disease resistance. Molluscan innate immunity includes cellular functions such as phagocytosis (Goedken & De Guise 2004), respiratory burst (Anderson et al. 1992), natural killer cell-like activity (Morsey & De Guise 2003), and apoptosis (Sunila & LaBanca 2003) as well as humoral factors including lysozymes, aminopeptidases, phospholipase C, antimicrobial peptides, and lectins (Mohandas & Cheng 1985, Kamiya et al. 1989, Cheng et al. 1995). Early studies stated that the pathogenicity of P. marinus was presumably associated with ineffective killing and degradation of parasites within oyster hemocyte phagosomes (Cheng 1975, Cheng 1984). A later review suggested that cellular defense mechanisms do not play a central role in actively defending against P. marinus infection (Chu 1999). Although phagocytic activity, hemocyte numbers, and granulocyte percentages were measured; this study did not evaluate respiratory burst-associated reactive oxygen intermediate (ROI) generation or apoptosis. Another study that did measure ROI activity reported a positive correlation between the severity of P. marinus infection and ROI generation but concluded that ROI generation by itself was an ineffective defense mechanism (Anderson et al. 1993). Anderson later reported that ROI production did not increase following the avid hemocyte phagocytosis of P. marinus but did on phagocytosis of the insoluble yeast preparation zymosan, suggesting that P. marinus can suppress or scavenge ROIs following the unaffected phagocytosis (Anderson 1999). Furthermore, the ROI scavenging ability of P. marinus has recently been demonstrated with the isolation of superoxide dismutase activity from cell extracts of P. marinus (Ahmed et al. 2003).

Because its oyster host is a poikilothermic osmoconformer, varying environmental conditions including water temperature and salinity have complicated establishing the relative significance of humoral and cellular defense mechanisms in oyster resistance to P. marinus infections. In controlled studies, P. marinus decimated oysters at water temperatures above 25[degrees]C (Fisher et al. 1992), whereas it rarely caused advanced disease in waters below 18[degrees]C (Chu & La Peyre 1993). Perkinsus marinus infection prevalence and intensity in the field dramatically decreased during winter and early spring, when water temperatures are coldest (Ragone Calvo & Burreson 1994). Although cold water reduced C. virginica hemocyte phagocytosis and respiratory burst-associated activity (Foley & Cheng 1975), it was not lethal to P. marinus (Burreson & Ragone Calvo 1993). Taken together, these studies suggest that hemocyte phagocytosis and respiratory burst are not important mechanisms in oyster resistance to P. marinus (Alvarez & Friedl 1990).

Although plasma lysozyme activity has been negatively correlated with oyster habitat salinity, temperature and prevalence of P. marinus infections (Chu 1999), and oyster lysozyme has bactericidal activity (Cheng & Rodrick 1975), it is not known if enhanced lysozyme activity results in reduced infection intensities (Chu & La Peyre 1993). Variables such as reduced parasite metabolic activity and pathogenicity at low temperatures (Mackin 1951), or temperature-related effects on oyster defense mechanisms yet to be identified, confound a thorough understanding of the relationships between water temperature, oyster defense mechanisms, and P. marinus infections.

Apoptosis is a regulated, physiologic form of cell death used by protozoans as well as metazoans to eliminate unwanted cells (Cotran et al. 1999). It consists of a defined sequence of stereotypical morphologic changes including cell surface blebbing, cell shrinkage, nuclear chromatin condensation, and fragmentation followed by apoptotic body formation and phagocytosis of apoptotic material, all in the absence of significant inflammation (Kroemer et al. 1995, Schlegel et al. 1996). This contrasts with necrosis, which is characterized by cell swelling, loss of cell membrane integrity, and local inflammation (Cotran et al. 1999). Initiation of apoptosis results in de novo gene expression of caspases and other enzymes responsible for controlled cellular dissolution (Krammer 1999). This process results in the removal of senescent or damaged cells for maintenance of tissue integrity and early defensive elimination of cells infected by intracellular pathogens (Cotran et al. 1999). Studies of apoptosis and its crucial role as a defense mechanism have been undertaken in organisms ranging from nematodes to mammals. Host cells infected by protozoa, bacteria, or viruses may counteract these intracellular infections by initiating their own death (cell suicide) and their subsequent removal by phagocytic cells (Luder et al. 2001).

While most of the data on apoptosis is from mammals or annelids, there are rare studies documenting apoptosis in shellfish. Apoptosis has been documented in shellfish using in situ terminal deoxynucleotidyl transferase-mediated dUTP-biotin nick end-labeling (TUNEL), electron microscopy, and Annexin-V staining. TUNEL staining has documented reduced hemocyte apoptosis in C. virginica naturally infected with P. marinus (Sunila & LaBanca 2003). Electron microscopy has demonstrated apoptotic cell morphology in herpes-virus infected Crassostrea gigas and Ostrea edulis (Renault et al. 2000). Treatment of oyster hemocytes with RDG-containing peptides also resulted in cell morphology changes characteristic of apoptosis (Gielazyn et al. 2003, Terahara et al. 2003). Annexin-V apoptosis staining in Crassostrea gigas hemocytes was documented upon their in vitro incubation with the neuroendocrine analog isoproterenol (Lacoste et al. 2002). These studies on apoptosis in bivalves addressed either specific metabolic pathways, or examined "snapshots" of the advanced stages of disease processes. Reports concerning controlled experiments to assess the effects on apoptosis frequencies of environmental conditions within the range of those normally encountered, have not been published before. Such applied experiments are necessary to elucidate hemocyte cellular functions and changes in parasite physiology that may affect the host-parasite relationship. In this study apoptosis in hemocytes and parasite cell populations was quantified at different temperatures and salinities. Understanding the role of environmental conditions on hemocyte and parasite apoptosis may expand the understanding of the host-parasite interactions and possibly resistance strategies of C. virginica against P. marinus, which could in turn be used for objective selection of disease resistant stocks, or more effective disease management strategies.

MATERIALS AND METHODS

NaCl, Tris buffer, magnesium chloride, DNase 1, ribonuclease A, dithiothreitol (DDT), DME and Hams F-12 media, sodium bicarbonate, glucose, galactose, trehalose, yeast extract, penicillin G, streptomycin sulfate, phenol red, potassium iodide, bovine serum albumin, and sterile seawater were obtained from Sigma-Aldrich Chemicals (St. Louis, MO). Propidium iodide (PI) and 1-[micro]m yellow-green fluorescent latex beads were purchased from Molecular Probes (Eugene, OR). Phosphate buffered saline (PBS), RPMI-1640 medium, HEPES, and L-glutamine were obtained from Gibco (Grand Island, NY). Ethanol was obtained from Pharmco (Brookfield, CT) and paraformaldehyde was purchased from Ted Pella, Inc. (Redding, CA). Recombinant human Annexin-V-FITC conjugate and Annexin-V binding buffer were from Caltag Laboratories (Burlingame, CA). Apoptag Fluorescein Apoptosis Detection Kit was obtained from Intergen (Purchase, NY). Synthetic sea salt and water quality test kits were purchased from Aquarium Systems (Mentor, OH). Algae were purchased from Reed Mariculture, Inc. (San Jose, CA). Perkinsus marinus isolate ATCC50862 (Dungan & Hamilton 1995) was provided by Cooperative Oxford Laboratory collaborators (Oxford, MD). Six- to eight-centimeters shell-height oysters were obtained from Fishers Island Oyster Farm (Fishers Island, NY) and 5- to 6-cm shellheight oysters from Taylor Shellfish Farms (Shelton, WA), between March 2001 and August 2003. The oysters used in this study were free of infection by P. marinus as confirmed by testing at the Connecticut Bureau of Aquaculture (Fishers Island Oyster Farm) and the absence of this pathogen on the United States Pacific coast (Taylor Shellfish Farms).

TUNEL Assay

Two milliliters of hemolymph samples were collected from the adductor muscle sinuses of eight oysters as previously described (Chu 1988). Hemocytes (2 x [10.sup.6]) were fixed in a chilled 1% (w/v) paraformaldehyde solution isotonic with the water in which oysters were kept. Cells were permeablized for 12 h at -20[degrees]C in 70% ethanol, and washed five times in 20[degrees]C PBS. Samples were washed and resuspended in isotonic Apoptag equilibration buffer adjusted with NaCl to the ambient salinity of the tanks' artificial seawater. Cells were then incubated in the terminal deoxynucleotidyl transferase (TDT) enzyme solution supplied with the kit. After a 30-min, 37[degrees]C incubation, the cells were washed in Apoptag stop/wash solution. Cells were counterstained with a 7.5 mM PI/PBS solution containing 7.8 mg/L RNase A for 15 min prior to analysis with a Becton Dickinson FACScan flow cytometer equipped with an argon laser. Apoptosis-associated fluorescence (FITC) was measured at 530 nm (FL-l), and DNA-associated fluorescence (PI) was measured at 575 nm (FL-2). Two-color cell associated fluorescence was evaluated using Cell Quest software (Becton Dickenson Immunocytometry Systems, San Jose, CA). Negative controls included samples incubated with equilibration buffer in lieu of Apoptag TdT enzyme reagent. Apoptotic rates were calculated as the percentage of apoptotic hemocytes relative to all hemocytes detected in analyzed samples.

Annexin V Assay

One- to two-milliliters hemolymph samples were collected from the adductor muscle sinus of oysters as previously described (Chu 1988). Cells (1 x [10.sup.6]) were resuspended in 1 mL of Caltag binding buffer adjusted with NaC1 to be isotonic to the oysters' environments. Five [micro]l of Caltag Annexin V-FITC conjugate and 0.5 [micro]g PI were added to 1 x [10.sup.5] hemocytes. After a 15-min incubation at 20[degrees]C in the dark, the cell solutions were diluted 1:4 with binding buffer. Apoptosis-associated fluorescence (FITC) and necrosis-associated fluorescence (PI) were measured using a FACScan flow cytometer and CellQuest software as described. Negative control hemocyte suspensions were incubated with isotonic seawater instead of PI and/or Annexin-V. The same protocol was used for cultured P. marinus cells. As previously validated for use in oysters (Lacoste et al. 2002), apoptotic cells were defined as Annexin-V-positive staining and PI-negative staining cells.

Experimental Conditions

Between 22 and 24 oysters were maintained in each of four 20-gallon tanks with temperature/osmolality combinations of 25[degrees]C & 540 mOsm [18.9%[per thousand]], 25[degrees]C/910 mOsm [31.9[per thousand]], 14.5[degrees]C/540 mOsm, and 14.5[degrees]C/910 mOsm, for 3 wk prior to experimentation.

Perkinsus marinus parasites in log-phase growth were propagated in 10 mL of DME: Ham's F-12 media (Dungan & Hamilton 1995) with similar temperature/salinity combinations of 22.5[degrees]C/488 mOsm [17.1[per thousand]], 22.5[degrees]C/810 mOsm [24.8[per thousand]], 14.5[degrees]C/488 mOsm, and 14.5[degrees]C/810 mOsm, for 3 days prior to experimentation.

Statistical Analyses

The percentage of apoptotic cells was calculated for granulocytes, hyalinocytes, and parasites. Significance for oyster and parasite salinity and temperature trials was established with 2-way analysis of variance. P < 0.05 was used to assess statistical significance of treatment differences in all data analyses.

RESULTS

Flow cytometry was used to quantify apoptosis of hemocytes at the single cell level using double staining with fluorescein isothiocynate (FITC)-labeled reagents and PI. Morphologically unique hemocyte subpopulations including hyalinocytes and granulocytes were defined by their relative size (forward scatter) and complexity (side scatter) (Fig. 1). Granulocytes were larger and more complex than hyalinocytes, as previously described (Goedken & De Guise 2004). TUNEL assays and 2-color flow cytometry were used to distinguish apoptotic hemocytes from nonapoptotic cells and to correlate apoptosis frequencies with cell cycle stage as estimated by relative cellular DNA contents (Fig. 2). FITC-conjugated d-UTP was specifically ligated to bound apoptotic DNA fragments in nuclei of fixed and permeablized hemocytes when incubated with the enzyme TdT. Subsequent PI staining of total DNA permitted estimation of the percentage of apoptotic hemocytes in [G.sub.2]/S phases (tetraploid DNA content) and [G.sub.0]/[G.sub.1] phases (diploid DNA content), with quadrants set using negative controls. The upper quadrants in Figure 2 contain apoptotic hemocytes, whereas the lower quadrants contain nonapoptotic cells. The fight quadrants contain hemocytes that are replicating DNA in preparation for, or in the process of, mitosis, while the left quadrants contain resting, haploid hemocytes. Although 12.08% of the cells were in [G.sub.2]/S phases, the proportion of apoptotic cells in that group was approximately three times higher than that in resting ([G.sub.0]/[G.sub.1] hemocytes (3.06% of hemocytes in [G.sub.2]/S phases vs. 1.10% of hemocytes in [G.sub.0]/[G.sub.1]phases).

[FIGURES 1-2 OMITTED]

Two-color flow cytometry utilizing AnnexinV-FITC and PI was used to distinguish unfixed, apoptotic hemocytes from viable nonapoptotic and necrotic hemocytes at the subpopulation level, as shown for granulocytes (Fig. 3). Annexin-V-FITC allows for sensitive cytometric recognition of externalized phosphatidyl serine (PS), a membrane phospholipid restricted to the inner leaflet in normal cells, but expressed on external membrane surfaces of apoptotic cells. The exclusion of PI by both apoptotic and normal viable cells, in conjunction with the rapid, high-affinity binding of externalized PS by Annexin-V, allowed accurate differentiation of apoptotic cells from necrotic and viable, nonapoptotic cells. Using this technique, 78.2% of the cells excluded PI (lower quadrants), of which 4.5% were undergoing apoptosis (lower right quadrant) and 73.7% were viable (lower left quadrant); 21.8% of hemocytes lacked intact membranes (based on their inability to exclude PI) and were classified as necrotic (upper quadrants). Cells in the upper right quadrant were not considered apoptotic because cell membranes permeable to PI might also allow Annexin-V-FITC to penetrate normal cells to label PS on the inner membrane leaflet of nonapoptotic cells, hence the importance of double labeling.

[FIGURE 3 OMITTED]

To measure physiologic variations associated with environmental conditions, hemocyte apoptosis was measured in oysters kept at different temperatures and salinities. Using the Annexin-V assay, differences in apoptosis frequencies were detected between hemocyte cell types (hyalinocytes vs. granulocytes) and between hemocytes from oysters held at different salinities (540 and 910 mOsm) (Fig. 4). In all test conditions, hyalinocytes had significantly lower percentages of apoptosis than granulocytes. Hyalinocyte apoptosis frequency was on average 44.4% less than granulocyte apoptosis frequency for the 93 oysters examined in these experiments. At both 14.5[degrees]C and 25[degrees]C, apoptosis rates were significantly lower in hemocytes from oysters held at 540 mOsm than from those held at 910 mOsm. There was 29% to 36% and 30% to 33% less apoptosis in hemocytes from oysters held at 540 mOsm than from those held at 910 mOsm, among granulocytes and hyalinocytes, respectively. Temperature had no significant effect on apoptosis rates in either granulocytes or hyalinocytes from oysters held at 14.5[degrees]C and 25[degrees]C.

[FIGURE 4 OMITTED]

In addition to the effects of temperature and salinity on hemocyte apoptosis, the influence of these environmental variables on apoptosis of in vitro propagated P. marinus was measured after incubation at similar temperatures in media of similar salinities (Fig. 5). Parasites at 14.5[degrees]C and 22.5[degrees]C had significant 2- to 5-fold increases in apoptotic rates when propagated at 488 mOsm compared with those propagated at 810 mOsm. Similar to hemocytes, there were no significant differences in apoptosis rates between low (14.5[degrees]C) and high (22.5[degrees]C) incubation temperatures, among parasite cells propagated in same-osmolality media.

[FIGURE 5 OMITTED]

DISCUSSION

Although modulation of apoptosis is frequently associated with host cell infections by intracellular pathogens, few publications have addressed the possible role of apoptosis as a cellular defense mechanism in poikilothermic, osmoconforming mollusks, or their parasites. The temperature and salinity of the estuaries that oysters live in are dynamic, and knowing the effect of these fluctuations on apoptosis of the host and pathogen may elucidate the host/ pathogen interaction. The objective of this study is to adapt and use flow cytometric assays to quickly and accurately quantify individual hemocyte and parasite apoptosis, to determine the effects of environmental conditions.

TUNEL and Annexin-V assays, which have been consistently and reliably used to detect apoptotic cells in mammals, were adapted to allow quantification of this innate defense mechanism in oysters by adjusting reagent osmolalities to that of osmoconforming oysters. Annexin-V and TUNEL assays were used in conjunction with PI staining to detect oyster hemocyte apoptosis. The TUNEL assay (Gavrieli et al. 1992, Gorczyca et al. 1993) was used to detect apoptotic cells with characteristic mononucleosomal (180 base pair fragments) and oligonucleosomal (multiples of 180 base pairs) double stranded DNA fragments (Duke et al. 1983). The enzyme TdT annealed fluorescein-conjugated dUTP to the exposed 3'-OH terminal (Gold et al. 1993). The assay is very sensitive and allowed detection of apoptotic DNA fragments in samples with a percentage of apoptotic cells as low as 1.1%. TUNEL assay reliability as an indicator of apoptosis has been confirmed by the hallmark morphologic characteristics of DNA fragmentation within intact cytoplasmic membranes (Homburg et al. 1995, Vermes et al. 1995). Propidium iodide, a fluorescent nucleic acid-binding dye, was concurrently used to quantify nucleic acids by fluorescence intensity (Nicoletti et al. 1991) and to correlate cell cycle status with apoptosis levels in samples that were fixed and permeablized prior to incubation with reagents. Because RNA was eliminated with ribonuclease A, associated fluorescence was from DNA, allowing differentiation of hemocytes undergoing or preparing to undergo mitosis, from resting cells. The few cells to the right of the boxed area in Figure 2 likely represent hemocyte doublets not excluded by the forward scatter gate (Fig. 1). While it is possible that some of the events enumerated in the right quadrants also represent doublets, it is unlikely, since these cells had a similar forward and side scatter profile to those in the left quadrants upon back-gating (data not shown). During DNA replication, nonlethal genetic mutations are either repaired or the mutated cell is eliminated by way of apoptosis. Because mutations and associated apoptosis are more likely to occur during replication of genetic material and at associated checkpoints ([G.sub.2]/S phases), it is not surprising to detect apoptosis three times more frequently in dividing cells than in resting cells ([G.sub.0]/[G.sub.1]). We found 1.12% of hemocytes in circulation that were undergoing cell division. Because this proportion of dividing cells would be alarmingly high in mammals, which have well defined hemopoietic centers, the high proportion of circulating dividing cells along with the lack of defined hemopoietic tissue in oysters may suggest that hemopoiesis occurs in circulation. The weakness of the TUNEL assay is a potential lack of specificity associated with necrotic cells. In necrotic cells, there can be fragmentation of DNA by enzymes unrelated to apoptosis (Froelich-Ammon & Osheroff 1995), which may be labeled by conjugated d-UTP to appear apoptotic.

To address this concern, Annexin-V was used to distinguish apoptosis by relying on the high affinity and specificity of this ubiquitous intracellular protein for [Ca.sup.2+]-dependant binding to the acidic phospholipid phosphatidyl serine (PS) (Vance 2003). Phosphatidyl serine, which is found within prokaryote and eukaryote cells, is a structural component of the inner cell membrane with a role in signal transduction pathways. The exteriorization of PS is an early event in the apoptotic process, which modulates recognition and phagocytosis of apoptotic fragments in a relatively non inflammatory fashion (Vance 2003). Because the membranes of apoptotic cells remain intact throughout most of the process, exogenous, extracellular Annexin-V will only bind PS expressed on the outer cell membrane leaflet (Kroemer et al. 1995). Propidium iodide is excluded by the intact membranes of unfixed, viable cells; but easily stains necrotic cells with compromised cell membrane integrity (Ormerod et al. 1993). This allows accurate differentiation of necrotic cells from early stage apoptotic cells, insuring highly specific recognition of apoptotic cells.

The results of this study suggest that, on average, there is 10% apoptosis (ranging from 4.5% to 15.3%) in circulating hemocytes of oysters kept within a normal range of environmental salinities and temperatures, when measured with the Annexin-V assay. Apoptosis rates measured with TUNEL assays averaged 1% (0.4% to 1.4%) for similar environmental conditions. This is to be expected because TUNEL assays measure end-stage apoptosis events, and a large percentage of cells with these DNA changes will have already been cleared through phagocytosis by neighboring cells, leaving fewer apoptotic cells for detection (Stummvoll et al. 2000). The low levels of apoptosis (~10%) are in the range of base-line hemocyte apoptosis rates estimated from samples taken from the vasculature of the cardiac sinus of C. gigas (Lacoste et al. 2002). There were marked differences between results of our study and another in the same oyster species that documented apoptosis in approximately 50% of hemocytes (Sunila & LaBanca 2003). This difference can be explained by the different source-location for hemocytes examined in these studies. While our study quantified apoptosis in circulating hemocytes, Sunila and LaBanca (2003) examined cells in tissues, presumably after they were recruited from the circulation to sites of inflammation in response to pathogens and/or tissue damage from infections emanating from the gastro-intestinal tract (Mackin 1951, Perkins 1976). Alternatively, senescent hemocytes could be undergoing apoptosis during their transit in the connective tissue where apoptotic bodies are more easily phagocytized than in circulation.

Granulocytes showed more apoptosis than hyalinocytes, regardless of temperature or salinity. This may be due to the higher metabolic activity in granulocytes with higher phagocytic and respiratory burst activity (Goedken & De Guise 2004) and is in agreement with previous findings of increased granulocyte apoptosis in C. virginica mucosa and submucosa (Sunila & LaBanca 2003).

Several studies have attempted to assess the impact of environmental conditions on rates of infection and development of disease. Temperature, which had no effects on oyster hemocyte or P. marinus apoptosis, was previously reported to be the greatest factor influencing disease prevalence (Chu et al. 1994), suggesting that other factors involved in either oyster resistance to P. marinus or pathogenicity of P. marinus are not related to apoptosis. Field studies have reported severe P. marinus infections in high-salinity (20[per thousand] to 30[per thousand]) coastal waters (Bayou Rigaud) and milder infections in low salinity (12[per thousand]) estuarine waters (Bay Chene Fleur) (Ray 1954). Experimental studies later confirmed the effects of salinity on severity of infection (Gauthier & Fisher 1991, Ragone & Burreson 1993). Our study approximated these natural conditions (although not reaching salinities as low as 12[per thousand]) and revealed that there was more hemocyte apoptosis in oysters maintained at high salinity as compared with lower salinity. Similar experiments with cultured P. marinus indicated the opposite trend with more parasite apoptosis at low salinity. Another variable affecting infection is increased flushing and dilution of parasite numbers by freshwater from rivers into intertidal areas, a phenomenon that would also influence water salinity (Bushek et al. 2000). Although oyster and P. marinus interactions are complex and difficult to decipher, it seems that higher salinity increases hemocyte apoptosis but has the opposite effect on parasite apoptosis. Further research to clarify the role of apoptosis as a mechanism of oyster disease resistance and/or parasite pathogenicity will be necessary before our results on the influence of temperature and salinity on apoptosis can be interpreted in the context of infection and disease. Comprehensive and objective strategies to revive the oyster industry, to include initiatives such as the development of disease resistant strains of oysters or new management strategies relative to disease resistance, will not be optimized without a better understanding of the host/parasite or environment interactions.

ACKNOWLEDGMENTS

This study was funded in part by USDA/CSREES #96-34341-3478, USDA/CSREES #CONS00718, a Connecticut Sea Grant Seed Grant and a University of Connecticut Research Foundation Small Grant. This research was completed with the support of Milton Levin, Steve Daniels, and Rosalee Hamilton (Maryland DNR).

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MICHEAL GOEDKEN, (1) BRENDA MORSEY, (1) INKE SUNILA, (2) CHRISTOPHER DUNGAN (3) AND SYLVAIN DE GUISE (1) *

(1) Department of Pathobiology and Veterinary Sciences University of Connecticut, 61 N Eagleville Road, Storrs, Connecticut 06269; (2) State of Connecticut Department of Agriculture, Bureau of Aquaculture, PO Box 97, Milford, Connecticut 06430; (3) Maryland Department of Natural Resources, Cooperative Oxford Laboratory, 904 S. Morris Street, Oxford, Maryland 21654

* Corresponding author. E-mail: [e-mail]