Domain Editor-in-Chief: Donald R. Zak; School of Natural Resources & Environment, University of Michigan, Ann Arbor, Michigan, United States
Associate Editor: Jessica J. Hellmann; Department of Biological Sciences, Environmental Change Initiative, University of Notre Dame, Notre Dame, Indiana, United States


Introduction

The problem of disease in marine ecosystems has recently attracted considerable attention, especially as the exacerbating effects of human activities on these systems have become apparent (Burge et al., 2014; Harvell et al., 2002; Lafferty et al., 2004). Some of the most important impacts that humans have had on marine diseases are the introduction of disease agents into new environments, the movement of naïve hosts (intentional or not) for trade and commerce, and the modification of environments in a way that changes the interaction of pathogens with their hosts (Harvell et al., 1999; Martin et al., 2010). Among the best-studied marine diseases are those affecting benthic species, such as sea urchins, corals and molluscs (Baker et al., 2008; Feehan and Scheibling, 2014; Hofmann and Ford, 2012; Sokolow, 2009). These may also be valuable commercial species or key members of an ecosystem, providing habitat and shelter for other species or foraging on significant ecosystem components (Coen et al., 2007); thus the loss of such organisms through disease-caused mortality can have both economic and ecological consequences (Feehan and Scheibling, 2014; Jones et al., 2004; Lafferty et al., 2015; National Research Council, 2004; Powell et al., 2008).

Two parasite-caused diseases of the eastern oyster (Crassostrea virginica) are especially important because of their widespread and devastating economic effects (Burreson and Ragone Calvo, 1996; Bushek et al., 2012; Ford and Bushek, 2012; Soniat, 1996). Dermo disease was enzootic in the southeastern United States and the Gulf of Mexico when it was first recognized in the late 1940s. Its etiological agent, Perkinsus marinus, was likely moved into the northeastern United States with the importation of infected oysters and later became epizootic during a period of rapid warming in the early 1990s (Cook et al., 1998). MSX disease is caused by a second protozoan, Haplosporidium nelsoni, which is enzootic to Asia, where it infects the Pacific oyster, C. gigas, but at low prevalence and without reported population-level effects (Burreson and Ford, 2004). Although the method of introduction of H. nelsoni to the east coast of North America is uncertain, the native eastern oyster was highly susceptible and has experienced catastrophic mortalities from the Chesapeake Bay to Nova Scotia, Canada in the years since the pathogen was first discovered in Delaware Bay in 1957 (Burreson et al., 2000).

The response of oyster populations to these two introduced pathogens has been documented over decades. The resulting long-term data sets offer insight into host-parasite interactions during which the environment changed over both short and long term periods (Burreson and Ragone Calvo, 1996; Bushek et al., 2012; Ford and Bushek, 2012). One of the resulting observations has been that MSX disease prevalence has declined in some areas where native oysters have experienced heavy selective mortality (Carnegie and Burreson, 2011; Ford and Bushek, 2012). In comparison, Dermo disease prevalence in wild populations shows much less evidence of a declining trend, with variations in prevalence largely a function of short-term climate cycles (Burreson and Ragone Calvo, 1996; Bushek et al., 2012).

Because knowledge of oyster disease and its consequent mortality are critical for management of this important resource and for basic understanding of natural selection, a number of studies have been undertaken to evaluate whether the development of resistance is responsible for these observations. Transplant and common garden experiments are often used to study how a host or host population responds to a particular pathogen and to begin to elucidate any underlying genetic basis for resistance or susceptibility (Nuismer and Gandon, 2008). With few exceptions (Bushek and Allen, 1996) in the case of oysters, such studies have been conducted using oysters deployed in field experiments and exposed to natural infections (Andrews, 1954; Brown et al., 2005a, 2005b; Burreson, 1991; Carnegie and Burreson, 2011; Encomio et al., 2005; Ford and Bushek, 2012; Haskin and Ford, 1979; Ragone Calvo et al., 2003). This has been necessary because of the logistical constraints of maintaining and feeding sufficiently large numbers of suspension-feeding oysters in the laboratory and because the MSX disease agent, H. nelsoni, has never been successfully transmitted in the laboratory. These studies have been transplant experiments in which the response of adult oysters from areas that have experienced disease mortality has been compared, under the same exposure conditions, with that of stocks imported from regions unaffected by disease.

Studies in Delaware Bay examined reasons for the decline in MSX disease in the Bay nearly 30 years after its appearance was first noted. A comparison of infection levels in native oysters from a single, high-salinity (20–23), site in the lower Bay, which historically experienced heavy infection pressure, with those of naïve oysters deployed at the same location showed that the naïve stock consistently developed high prevalence and heavy H. nelsoni infections, whereas the natives showed few if any infections (Ford and Bushek, 2012). Haplosporidium nelsoni infection pressure decreases in an upbay direction, in association with lowered salinity (Haskin and Ford, 1982), which raised the question of whether oysters in the upper Bay would respond similarly to lower Bay oysters. Thus, a subsequent experiment involved transplanting oysters from two locations in Delaware Bay, with average salinities of ∼15 and ∼10 and moderate to very low MSX-disease-caused mortality, into the high-infection site. Prevalence and intensity of H. nelsoni infections were compared over a single season (May to October) to the local lower Bay stock and to a naïve stock that had never experienced selective mortality due to the pathogen. Native Delaware Bay stocks developed far fewer and lighter H. nelsoni infections than did the naïve stock, suggesting that resistance to this pathogen was widespread in the bay, but results also showed a weak negative association of both mortality and infection levels with distance upbay into lower salinity waters (Ford and Bushek, 2012). In both studies, however, P. marinus infection prevalence was high in the native, as well as in the naïve, oysters. A similar transplant experiment by Carnegie and Burreson (2011) verified that the observed decline in MSX disease in oysters having experienced selective mortality in the lower Chesapeake Bay was due to the development of resistance. In the Chesapeake Bay, however, resistance was not as widespread as in Delaware Bay, with unselected populations persisting in low-salinity regions that experience incursions of the H. nelsoni only during droughts.

A complicating factor in these studies, as well as in most other such comparisons, has been the use of adult oysters, raising the possibility of differential selection prior to the test (Kawecki and Ebert, 2004). Thus, the present study was an attempt to overcome the deficiencies of previous experiments (e.g., (Andrews, 1954; Brown et al., 2005a, 2005b; Burreson, 1991; Carnegie and Burreson, 2011; Encomio et al., 2005; Ford and Bushek, 2012; Haskin and Ford, 1979; Ragone Calvo et al., 2003) by comparing hatchery-reared cohorts produced from broodstock collected in different regions of Delaware Bay with those from a non-enzootic area, thus eliminating the problem of possible pre-test differential selection posed by using adult oysters. All test groups would thus have experienced the same environment prior to exposure, in a true common garden experiment, but would represent potentially distinct genotypes from different regions in the Bay and its tributaries. We were particularly interested in those from the upper-most natural reefs in the estuary as well as in locations in tributary rivers, where infections are extremely rare and, if present, not lethal (Ford et al., 2012). The response to disease challenge of these groups would enable us to infer the degree of genetic connectivity, via larval transfer, within the oyster metapopulation of the Bay, including the outlying subpopulations, as simulated in a modeling study by Munroe et al. (2015)

Materials and methods

Adult oysters for broodstock were collected from three low-salinity sites in Delaware Bay, brought into the hatchery and spawned at the Haskin Shellfish Research Laboratory, Rutgers University, Cape Shore Facility in the summer of 2007 (Fig. 1, Table 1). One site was higher upbay and two were in tributary rivers, and all were in somewhat lower salinity than those described above and presumably where selection pressure was minimal or non-existent (Ford and Bushek, 2012). Newly settled oysters (spat) at the Cape Shore site, where selection is intense, were included as a fourth Delaware Bay group. Naïve oysters from the Damariscotta River, Maine were spawned along with the 2007 Delaware Bay groups. Although an outbreak of MSX disease occurred in Maine in 2010, histological examination of the Maine broodstock in 2007 revealed no detectable infections of H. nelsoni and P. marinus is rare or absent in the river; thus oysters from this location are extremely susceptible to both pathogens. We repeated the test with two additional Delaware Bay cohorts spawned in 2008 in order to extend our source broodstock farther upbay (Fig. 1, Table 1).

doi: 10.12952/journal.elementa.000119.f001.
Figure 1.  

Map of Delaware Bay.

Sampling (circles) and deployment (stars) locations. Note that wild-set oysters of the 2007 year class were collected from the Cape Shore site for testing with 2007 hatchery-reared cohorts at both deployment sites.

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Table 1.

Study period span and times sampled for Perkinsus marinusandHaplosporidium nelsoni detection

2007 Offspring Stock Code Salinity of Broodstock Locationa Cape May Cape Shore
3/08–10/09 3/08–10/09
Maine (naïve control) ME07 28–31 6 X 4 X
Cohansey River CR07 11 6 X 5 X
Round Island Bed RI07 10 6 X 6 X
Leipsic R LP07 13 6 X 6 X
Cape Shore CS07 20-23 6 X 6 X
2008 Offspring 5/09–10/10 5/09–10/10
Hope Creek HC08 8 6 X --
Shell Rock SH08 15-16 6 X --

Broodstock was conditioned at the Cape Shore Facility before spawning. Each hatchery cohort was produced in a factorial design that involved strip spawning of 8 to 15 females and 8 to 10 males. Equal numbers of eggs from each female were pooled and divided into aliquots according to the number of males. Each egg aliquot was fertilized by one male. Fertilized eggs were then pooled and reared until setting. This procedure, which produced about 100 families in each cohort, increases the effective population size and maximizes genetic diversity among the offspring. Newly set spat were kept in the hatchery for a week until they were ∼1 mm and then moved to a land-based nursery system. Hatchery water was filtered and UV-treated then supplemented with cultured phytoplankton for food whereas the nursery received untreated bay water filtered through a 150-µm bag that provided a natural diet from local waters. Spat were maintained in the nursery for approximately three to four months until they were placed into mesh bags and moved into a protected site for overwintering.

Cohorts in each year class were deployed in April of the year following spawning for exposure to natural infections at two intertidal locations: Cape May Harbor and Cape Shore (Fig. 1). Both are intertidal sites, although the stocks at Cape May were not always completely exposed at low tide; salinity in the Harbor is about 30–32; at the Cape Shore it is 20–23. Infection pressure from H. nelsoni is heavy at both locations, but P. marinus infections are usually rare and very slow to develop at the Cape May Harbor site. The difference in P. marinus levels is most likely due to the temperature differential between the sites: in the summer when the parasite is most active, water temperatures at the Cape Shore can be up to 10o C higher than at Cape May (e.g., 31o vs 21o C). Thus, despite not being able to experimentally infect with H. nelsoni, we could compare stocks affected primarily or only by H. nelsoni and by both H. nelsoni and P. marinus. The cohorts were monitored for infection prevalence and intensity of both pathogens, and for mortality over 18 months.

At deployment in April of 2008 and 2009, the oysters were approximately eight months old. Duplicate bags of 500 to 700 spat each were placed on racks at each site. During the study, the bags were opened and cleaned, and all live and dead oysters counted and recorded at seven to ten sampling dates. Dead oysters were removed and live ones returned. Samples for infection assays were typically collected six times during the monitoring period (Table 1), in months designed to capture previously documented summer and fall peaks, and early spring lows, in the infection cycles of both pathogens (Ford and Tripp, 1996).

At each sampling time, 20 individuals from each group (10 per bag) were measured (hinge to lip), assayed by standard tissue-section histology for prevalence and intensity of H. nelsoni infections and by Ray’s Fluid Thioglycollate Medium (RFTM) incubation for P. marinus infections (Howard et al., 2004). Infection intensities were scored according to Ford et al. (1999) for H. nelsoni and Ray (1954) for P. marinus. Advanced infections are considered to be those that are lethal, or are approaching lethal levels: stage 4 (of 4) for H. nelsoni and 3–5 (of 5) for P. marinus (Dungan and Bushek, 2015; Ford and Haskin, 1982)

Cumulative mortality (mean of the duplicate bags) was calculated from the date of deployment at the two test sites. Although differential overwinter mortality during the first year does occur, it is not consistently associated with H. nelsoni infections (Haskin and Ford, 1979). Some infections may be acquired in the late summer/fall when the oysters are only a few months old, but they typically do not become heavy enough to cause mortality until the following late spring (Andrews, 1966). Perkinsusmarinus infections are extremely rare in spat that are only a few months old. Our data reflected that finding to some extent: one of the highest overwinter mortalities (30% for the ME07s) corresponded with H. nelsoni prevalence of 50% (Fig. 2 A and C), although that value was measured in the March 2008, after the mortality. Another group, CR07 experienced only 11% overwinter loss even though the March 2008 sample had 25% prevalence. In the absence of pre-winter infections assays, the post winter assays are the closest we can come to inferring relative infection levels that might have affected overwinter mortalities. Using this measure, we found no evidence of a relationship between prevalence of H. nelsoni and overwinter mortality (N = 7, r2 = 0.0056). Post winter Perkinsus marinus levels were essentially zero. Thus, to avoid confusion between deaths that are clearly caused by disease agents and those associated with other (unknown) factors, mortality of experimental stocks is calculated beginning in the spring after their birth year and before they are exposed to a full infection period. We also found no relationship between the salinity of the broodstock location and initial (overwinter) mortality (N = 7, r2 = 0.0124)

doi: 10.12952/journal.elementa.000119.f002.
Figure 2.  

Seasonal patterns of total and advanced Haplosporidium nelsoni infections.

Haplosporidium nelsoni (MSX disease agent) prevalence in oysters sampled over an 18-month period at Cape May (A and B) and Cape Shore (C and D) sites for the 2007 cohorts sampled in 2008 and 2009. E&F: Total and advanced infection prevalence for the 2008 cohorts sampled in 2009 and 2010 at Cape May. Each point represents a 20-oyster sample; both total and advance prevalence data are relative to the entire sample. Legend in plot D refers to A-D; that in F, refers to E and F. See Table 1 for stock codes.

Results

Growth

Mean shell heights at deployment (March 2008) of all the 2007 cohorts were between 13 and 18 mm except for the Cape Shore set (CS07 – see Table 1 for stock codes). This group was larger, averaging 29 mm, because it was a natural set with greater access to food than the other groups, which were reared onshore with more limited food availability. In October 2009, survivors of the 2007 year class were generally 55 to 60 mm in shell height at both Cape May (CM) and Cape Shore (CS) sites, except for the ME07 and CR07 at CS, which were less than 50 mm (Table 2). Both 2008 groups were deployed only at CM and first measured in May of 2009. Therefore, the initial size measurements were larger than those of the previous year, at 28 to 29 mm, but by October 2010 when the study ended, marked differences were evident between the two groups: averages of 56 mm for HC08 and 64 mm for SR08 (Table 2).

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Table 2.

Mean (SE) shell height in millimeters of experimental oysters at the start and end of exposure period

Stock Codea At Deployment At Termination
Cape May Cape Shore
ME07 16.9 (0.6)b 60.4 (2.4) 41.4 (0.8)
CR07 13.5 (0.5) 55.0 (2.3) 46.3 (1.3)
RI07 17.4 (0.6) 57.3 (1.9) 57.4 (1.7)
LP07 17.7 (0.6) 60.6 (2.7) 60.0 (2.0)
CS07 29.4 (1.6) 59.8 (2.7) 57.7 (1.9)
HC08 29.0 (1.5) 56.1 (2.0) NSc
SR08 27.7 (1.0) 63.5 (2.3) NSc

Infections

Haplosporidium nelsoni

Examination of samples taken at the time of deployment (March 2008) showed that some oysters in all 2007 groups already had detectable infections, most likely acquired while they were in the land-based nursery system during the previous late summer (see Ford et al., 2001). Prevalence ranged from an average of 14% (0–5% advanced) in the four Delaware Bay groups to 50% (10% advanced) in the ME07s, and did not vary dramatically over the study period (Fig. 2). Both total and advanced infection levels in the ME07s remained higher by 15 to 50 percentage points compared to the Delaware Bay groups at both test sites (Fig. 2). This difference was clearly evident in the mean values for data averaged over the study, which showed that, with the exception of the CR07s, the average prevalence of advanced infections in the ME07s at both locations was approximately three times that of the average total prevalence of the Delaware Bay groups (Fig. 3A and B). The CR07s stood out among the Delaware Bay groups as having somewhat higher prevalence, especially at the Cape Shore.

doi: 10.12952/journal.elementa.000119.f003.
Figure 3.  

Mean total and advanced Haplosploridium nelsoni infection prevalence.

Mean total and advanced infection prevalence of Haplosploridium nelsoni (MSX disease agent) in oysters sampled over 18-month periods at Cape May (A) and Cape Shore (B) sites. The tops of bars represent the total prevalence; the solid portion represents prevalence of advanced infections. Each bar is the mean of six samples (120 individuals). See Table 1 for stock codes.

The two cohorts produced in 2008 also acquired H. nelsoni infections while they were in the nursery that summer. They were subsequently deployed at CM only. Fifty-five percent of the HC08 oysters and 30% of the SR08 group had advanced infections when first sampled in May 2009. Thereafter, until the end of the study in October 2010, infection levels were similar in both groups, varying between 5 and 40% total, and 0 and 5% advanced infections (Fig. 2E & F), with marginally higher overall values for the HC08 group (Fig. 3A).

Perkinsus marinus

At the time of deployment in March 2008, prevalence was mostly 0% in the 2007 groups. With the exception of the CS07 group, it remained <5% at the CM site until the very end of the study period in October 2009 when it rose rapidly, reaching 85% in the ME07, 75% in the CS07 and 20 to 25% in the rest (Fig. 4A); however, only the ME07s showed substantial advanced infections at 50% (Fig. 4B).

doi: 10.12952/journal.elementa.000119.f004.
Figure 4.  

Prevalence of Perkinsus marinus infections.

Prevalence of Perkinsus marinus (Dermo disease agent) infections in oysters sampled over an 18-month period at Cape May (A and B) and Cape Shore (C and D) sites. A and B: Total and C and D advanced infection prevalence for the 2007 cohorts sampled in 2008 and 2009; E&F: Total and advanced infection prevalence for the 2008 cohorts sampled in 2009 and 2010. Each point represents a 20-oyster sample; both total and advance prevalence data are relative to the entire sample. Legend in plot D refers to A-D; that in F, refers to E and F. See Table 1 for stock codes.

The picture was very different at CS, where P. marinus is enzootic and where all groups quickly became infected, with total prevalence reaching close to 100% by late summer 2008 (Fig. 4C). Infection intensity followed a similar seasonal pattern, but reached the advanced stage faster in the ME07s compared to the Delaware Bay stocks. By August 2008, 70% of the ME07s vs 5 to 44% of the Delaware Bay groups had advanced infections, and this differential was still evident, albeit to a lesser degree, two months later, in October (Fig. 4D). Following the typical seasonal pattern, P. marinus was cleared from oysters in all stocks except the ME07s in spring 2009, but prevalence rose again to nearly 100%, with the majority being advanced infections, by the end of the study (Fig. 4D). Means for prevalence data averaged over the study showed somewhat elevated values for the ME07s and the CS07, although all were low overall compared to those at the Cape Shore (Fig. 5A and B).

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Figure 5.  

Mean total and advanced Perkinsus marinus infection prevalence.

Mean total and advanced infection prevalence of Perkinsus marinus (Dermo disease agent) in oysters sampled over 18-month periods at Cape May (A) and Cape Shore (B) sites. The tops of bars represent the total prevalence; the solid portion represents prevalence of advanced infections. Each bar is the mean of four (ME07), five (CR07) or six (RI07, LP07 and CS07) samples (80 to 120 individuals). See Table 1 for stock codes.

Like the 2007 cohorts, the 2008 groups deployed at CM had almost no detectable infections at the first sampling in May 2009. In both 2009 and 2010, infections developed earlier in the HC08 group than in the SR08s (Fig. 4E and F). Mean prevalence, calculated across the study period, was 49% in the HC08s and 22% in the SR08s; mean advanced prevalence was 22% and 11%, respectively (Fig. 5A).

Mortality

Mortality of the naïve ME07 group reached nearly 90% at CM and 95% at CS between April and October 2008, the end of the first full summer exposure period (Fig. 6A & B). Both groups were terminated after one year because so few individuals remained: less than 8% at CM and 1%, at CS; however, sampling for infections continued at CM using oysters from the same cohort that were not part of the mortality assessment. The difference between the naïve groups and the four native Delaware Bay groups was especially striking at CM (Fig. 6A), where the disease agent was almost exclusively H. nelsoni. Although all had substantially less mortality than the naïve controls, one Delaware Bay group stood out from the others: offspring of the Cohansey River (CR07) parents had higher mortality after one year (42% vs 19–26% for the other groups), and this differential persisted until the end of the study when death in the CR07 oysters reached 58% vs 36–39% for the others (Fig. 6A).

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Figure 6.  

Mortality patterns.

Cumulative mortality of oyster stocks deployed at Cape May and Cape Shore sites during 18-month periods at Cape May (A and C) and Cape Shore (B). Each point is the mean of two replicate bags. Error bars are omitted for the sake of visual clarity, but differences were rarely greater than 5 percentage points.

At CS, where oysters were exposed to heavy infection pressure from both H. nelsoni and P. marinus, mortalities were not only higher overall, but there were smaller differences between the naïve and native groups than at CM. Nevertheless, the same rank pattern was evident among the Delaware Bay groups, with the CR07 cohort experiencing much heavier mortality than the rest during the first year. At the end of one year, cumulative mortality was 90% in this group compared to 43% and 46% for LP07 and RI07, respectively. The native oysters that had set at the Cape Shore in mid-summer 2007 (CS07) experienced higher losses (62%), likely because they had had longer exposure, especially to P. marinus, as also indicated by the infection data (Fig. 4A and B, 5A). The death rate in the other groups accelerated towards the end of the study period reaching a cumulative mortality total by October 2009 of between 74 and 88% (Fig. 6B).

Offspring of the two Delaware Bay groups, HC08 and SR08, produced in 2008 and tested at CM in 2009 and 2010 experienced similar mortality patterns but at distinctly different levels (Fig. 6C), with the differential established primarily in the first season. Between March and October 2009, mortality of the HC08 group, from the most upbay oyster bed, reached 65%. Offspring from the SR group from a lower-bay bed, was 36% during the same period. At the end of the second year, October 2010, mortality was 83% vs 50% in these two groups, respectively (Fig. 6C).

It is noteworthy that deaths during the first summer and fall of exposure were largely responsible for establishing the mortality differentials among groups and they were associated with concurrent elevated infection levels of one or both of the pathogens (Figs. 2 and 6). This is particularly true for the Maine group, which displayed high infection levels, especially of H. nelsoni, during the period of most rapid mortality. The two Delaware Bay stocks that appeared less resistance to disease compared to others (CR07 and RI08) also stood out as having higher infection levels during that initial exposure period.

Discussion

The overall goal of this study was to further understand how a metapopulation of oysters responded to epizootic mortalities caused by two pathogens, each of which was influenced by anthropogenic factors: climate, pathogen introduction or both. How these factors influence disease has been investigated and reviewed for numerous terrestrial and aquatic systems, including ours (Anderson et al., 2004; Hofmann et al., 2009; Lafferty, 2009; Lindgren et al., 2000; Soniat et al., 2009); however, we are unaware of any other system in which the responses of the host population has been so well documented as in Delaware Bay.

To accomplish the goal, our study had two objectives. The first was to use hatchery-reared stocks to validate prior observations that resistance to these pathogens had developed naturally in wild stocks from Delaware Bay, USA. The second was to delineate the spatial extent and degree to which resistance is present in different Delaware Bay subpopulations, especially those at the extreme, low-salinity edges of the population boundaries, which experience little or no selection pressure. Measuring disease impacts in populations where individuals cannot be followed over time is difficult. In such cases, mortality often is a better gauge of disease impact than are “snap shot” infection measures. We were able to combine cumulative mortality, a time-integrated measure, with infection assessment, confirming the presence and intensity of the disease agents at key points in the infection cycle, especially periods when mortality rates among the tested groups differed substantially.

The results confirmed those of previous studies (Ford and Bushek, 2012): the naïve stock from outside the estuary stood out as experiencing much more rapid, and overall greater cumulative, mortality over the study period than did any of the native groups, including those in the uppermost reefs and tributaries. In fact, mortality exceeding 90% during the first summer alone prohibited following the naïve stock in both study locations after the end of the first year. But the study also underscored the very different response of the oysters to the two pathogens. The greater mortality of the naïve stock was paralleled by higher infection prevalence and intensity, predominantly of the MSX disease agent, H. nelsoni. Despite the heavy infection pressure by this pathogen, the native oysters tested at the same time developed far fewer infections and almost none were advanced. In contrast, native oysters, although slower to develop advanced infections of the Dermo disease agent, P. marinus, eventually reached prevalence and mortality levels that almost matched those of the naïve stock.

Efforts to produce selectively bred oysters obviously employ hatchery-reared progeny in exposure trials (see review by Degremont et al., 2015), but studies of resistance in wild populations typically employ adults. In contrast, our study employed hatchery-reared and newly settled oysters in order to eliminate the possibility that results using adults might have been influenced by local selection prior to the test (Kawecki and Ebert, 2004). This common-garden approach also minimized physiological adaptations that may have been present in adults, reducing their ability to combat disease challenge and removing confounding effects associated with transplant experiments (Nuismer and Gandon, 2008). Although we cannot discount the possibility that genetic adaptation to local environments, especially low salinity, might stress oysters placed at high salinity and thus affect their ability to resist infection and disease, our experience with H. nelsoni and P. marinus over more than 50 years of testing various stocks at the Cape Shore site has never provided evidence that this is the case for these two pathogens. They are so virulent that they cause disease and mortality even in originally very healthy individuals – as illustrated by their impact on naïve wild populations wherever they have appeared along the northeast coast of North America (Burreson and Ford, 2004; Ford and Smolowitz, 2007; Ford and Tripp, 1996).

Results from our study were essentially the same as the previous ones using adults: the hatchery-reared native stocks, including those from the population margins, were much more resistant to MSX disease than was the naïve stock and there was far less difference between native and naïve groups in Dermo disease levels. By removing the confounding environmental effects, we confirmed the distribution of genetically based resistance throughout the Delaware Bay, USA. The apparent lowered resistance of the two most upbay/upriver stocks, Cohansey River (CR07) and Hope Creek (HC08) (Fig. 1), compared to groups from the rest of the Bay, was consistent with results of adult-only trials, which showed a similar association of lessened resistance for stocks in low-salinity regions (Carnegie and Burreson, 2011; Ford and Bushek, 2012).

By deploying experimental stocks at sites where infection pressure from the two pathogens differed, we were able to assess differences in resistance to the development of the two diseases while still using field deployments. Comparison of outcomes at the two locations clearly shows the relative lack of resistance to the development of Dermo disease compared to that for MSX disease. Oysters at both Cape May Harbor and Cape Shore acquired infections of the MSX-disease pathogen, Haplosporidium nelsoni at similar levels, with natives rarely showing advanced (i.e., lethal) infections in more than 5% of the samples compared to 20 to 50% for the naïve stock. At Cape May, however, the Dermo disease agent, Perkinsus marinus, infections were largely absent until the very end of the study, whereas they were acquired rapidly at the Cape Shore location and reached 80–100% prevalence almost immediately in both native and naïve groups. The major difference was that advanced infections developed to a somewhat lesser extent in the natives during the first summer, but by the end of the study, after two seasons of exposure, 75–85% of the natives had advanced P. marinus infections, the same as recorded in the naïve oysters after a single season.

At Cape May, where infection pressure came almost exclusively from the MSX pathogen, one-year mortality in the Delaware Bay stocks was 20–40% compared to more than 90% in the naïve group. At Cape Shore, where both disease agents were prevalent, the predominance of Dermo disease as a cause of mortality in the native oysters can be inferred: 37% to 64% of the native stocks and 95% of the naïve group died during the first year, but by the end of the 18-month study, 70% to more than 95% of the natives had died. These results are consistent with the results of annual surveys of Delaware Bay oyster populations, which show little evidence of a trend towards lower P. marinus infection prevalence or consequent mortality since the 1990 epizootic (Bushek et al., 2012). Nevertheless, a delay in the development of advanced infections is clearly of benefit to the oyster grower as it provides extra time for the crop to reach market size before the danger of mass mortality. In fact, this is in part, how resistance to MSX disease-caused mortality developed: selected animals were those able to delay development of lethal infections (Ford and Haskin, 1987).

Why is it that the oysters developed significant resistance to MSX disease-caused mortality so quickly and clearly, whereas resistance to Dermo disease has been modest at best, even after 25 years of continuous disease pressure over large portions of the Delaware Bay oyster beds? Several explanations have been considered. The first is that susceptible oysters become heavily infected with the MSX pathogen and a large proportion die before they can spawn (Haskin and Ford, 1979), whereas infections caused by the Dermo pathogen typically do not become debilitating until oysters are at the end of their second year of life, thus allowing susceptibles among them to spawn at least once before they die (Bushek et al., 2012; Dittman et al., 2001). Consequently, MSX disease exerts a much stronger selective force. This explanation, based on wild populations, is weakened, however, by data from selective breeding trials, which employ broodstock that have undergone mortality caused by Dermo disease. Results of studies to date are inconsistent, with some showing modest improvement, others showing none (Frank-Lawale et al., 2014; Guo et al., 2003; Ragone Calvo et al., 2003)

Another possibility is that balancing-selection occurs such that genotypes with resistance to Dermo disease are more susceptible to mortality caused by other factors and die before selection for resistance has a chance to act. Under this scenario, the resistant genotypes are less fit overall and are prevented from becoming dominant, if not fixed in the population. Finally, the genes that play a role resistance to Dermo disease may be more numerous than those involved in resistance to MSX disease. Using a gene-based model, Powell et al. (2011) showed that the time to development of resistance to a disease increases with the number of loci involved. With the same gene-based model, Munroe et al. (2015) concluded that the only way to adequately simulate the results of selective breeding for resistance to MSX disease (Haskin and Ford, 1979) was to assume asingle highly influential locus” even though other, less influential, loci could be involved.

Although marine epizootics can affect wide areas, refuges from infection do exist. They are likely to be associated with environmental conditions such as temperature (Perrigault et al., 2009; Scheibling and Hennigar, 1997) and salinity (Haskin and Ford, 1982), and may also be temporally or spatially transient (Ford et al., 2012), modified by short-term weather cycles and longer-term climate trends. In estuaries, variable freshwater input, wind direction and circulation patterns create conditions that favor or prevent parasites from moving into upbay and upriver sites and initiating infections (Wang et al., 2012). The results of the present study indicate that oysters in Delaware Bay “refuges”- in the higher reaches of the Bay and its tributary rivers - are measurably less resistant to the development of infections of both H. nelsoni and P. marinus and to resulting mortality than are oysters in those sections of the bay not protected from the parasites, as might be expected. Genetic analysis is consistent with this finding. Examination of oysters from 9 sites in Delaware Bay, including those from which broodstock was taken for the present study, showed that oysters from the main body of the bay are genetically homogeneous, but different from the peripheral areas (including the Cohansey River and Hope Creek sites) at microsatellite loci linked to disease resistance (Hofmann et al., 2009). Nevertheless, oysters in these outlying areas are still much more resistant to the MSX parasite, and measurably, although less so to the Dermo parasite, than naïve oysters from outside the estuary.

Ford et al. (2012) documented the existence of refuges from both MSX and Dermo diseases in the upper bay and tributaries of the Delaware estuary based on the absence of infective stages and/or the absence of detectable infections. Whether the genetic contribution to the overall metapopulation by oysters in these refuges could delay the development of resistance in the metapopulation, as discussed by Ford et al. (2012) and simulated by Munroe et al. (2015) is almost a moot point in the case of MSX disease, because the majority of the oyster population has become very resistant. The more interesting outcome of our study is the degree of resistance that these subpopulations have, in fact, achieved despite a lack of evidence for in situ disease or disease-caused mortality. What it does demonstrate is the high degree of genetic connectivity within the Delaware Bay oyster metapopulation, allowing the offspring of highly selected subpopulations in the lower estuary to extend upbay to the very edges of the zones inhabited by the oysters, even against a general trend of downbay larval transport (Wang et al., 2012). This contrasts with the findings of Carnegie and Burreson (2011), who reported that clear evidence of resistance to MSX disease in Chesapeake Bay oysters was limited to the higher salinity portions of the lower estuary. The difference is likely due to the fact that Chesapeake Bay is much larger and morphologically more complex than Delaware Bay, with many oyster habitats existing in the major river systems flowing into the Bay. Our results, however, support their recommendation that conservation and restoration efforts should include populations in the higher-salinity regions where selection can occur, rather than just in the disease-free lower salinity regions. These higher salinity regions often support fisheries where they are commonly thought to be larval sinks due to the general net downstream transport of larvae. However, results from Wang et al. (2012), Munroe et al. (2015) and this study indicate that enhancement of fishery populations where disease selection pressure is high may help provide resilience to the larger metapopulation by allowing resistance to spread as some larvae are inevitably transported upstream via tidal circulation.

In a review of the evolutionary impact of introduced pathogens on native populations, Strauss et al. (2006) remarked that “surprisingly little evidence has been collected to show the degree to which [introduced disease agents] are important selective agents in native populations.” These authors cited observational studies of plants and terrestrial vertebrates, but the long-term data on eastern oysters combined with experimental studies may provide some of the best-documented examples of how populations respond to introduced pathogens and how those pathogens influence the evolution of affected populations. Our results also provide evidence of the potential resilience of wild populations to major population declines caused by anthropogenically introduced disease agents, a finding that should be considered in planning and pursuing restoration activities. A case in point is the biologically counter-intuitive fisheries management strategy in which restoration and enhancement efforts reflectively avoid areas of disease, thereby ignoring the natural resilience inherent in the population and its potential for self-correction.

Data accessibility statement

Data sets upon which this article is based will be made available upon request made to the corresponding author.

Copyright

© 2016 Bushek and Ford. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Contributions

Contributed to conception and design: DB, SEF

Contributed to acquisition of data: DB, SEF

Contributed to analysis and interpretation of data: DB, SEF

Drafted and/or revised the article: DB, SEF

Approved the submitted version for publication: DB, SEF

Competing interests

The authors have no competing interests.

Funding information

The National Science Foundation, Ecology of Infectious Diseases Program, grant numbers NSF OCE 0622672 and 1216220 provided financial support and an NSF-REU supplement provided support for student participation. This work was supported in part by USDA NIFA Hatch project #1009201 through NJAES Hatch project NJ32114.