Antimicrobial activity of copper and zinc accumulated in eastern oyster amebocytes.

ABSTRACT The distribution of eastern oysters Crassostrea virginica near terrestrial watersheds has led to a general impression that low or variable salinity is imperative for survival. However, freshwater runoff contains numerous mineral elements from geologic deposits that could play significant roles in oyster physiology. Two metals of terrestrial origin, copper and zinc, are accumulated to extremely high concentrations in eastern oysters, even in the absence of anthropogenic sources. As yet, there has been no defendable demonstration of a physiologic function for such high concentrations. Both copper and zinc, however, are accumulated almost exclusively in the amebocytes and calcareous shell of oysters, a unique distribution that implicates a role in the functions of amebocytes. Amebocytes are migratory, diapedetic cells generally recognized to provide nutriment and defense through phagocytosis, killing, and digestion of invading or ingested microorganisms. There is sufficient evidence in existing literature to suggest that copper and zinc directly contribute to these antimicrobial activities. This review presents historical and recent findings that demonstrate a strong affinity of oyster amebocytes for copper and zinc (even in low ambient concentrations), prolonged retention of the metals despite a potential route of elimination, and strong circumstantial evidence of antimicrobial activity by accumulated copper and zinc. It is proposed that oysters actively concentrate copper and zinc as antimicrobial agents to be used in intracellolar and extracellular killing (direct toxicity) as well as extracellular clot formation (precipitation of hemolymph). This potential, combined with evidence of amebocyte involvement in deposition of oyster shell, provides an alternative framework for understanding amebocyte functions, defense activities, and coastal distributions of oyster populations. It also affords some resolution to the apparent contradiction of eastern oysters thriving at seemingly polluted locations.

KEY WORDS: oysters, Crassostrea virginica, marine bivalves, copper, zinc, metals, contaminants, amebocytes, leucocytes, hemocytes, internal defense, immunomodulation, immunosuppression, metal toxicity, sentinel species

INTRODUCTION

Eastern oysters Crassostrea virginica are distributed across the North American coast of the Atlantic Ocean and Gulf of Mexico in bays, estuaries, and coastal zones, most often near sources of fresh water. The proximity to fresh water has often led to a conclusion that oysters require the low or variable salinity provided by freshwater inflow (Cake 1983, Soniat & Brody 1988, Berrigan et al. 1991). Intermittent salinity reductions can deter stenohaline predators such as oyster drills (Loosanoff 1955, Lunz 1955, Hopkins 1956, Wells 1961, Menzel et al. 1966, Berrigan et al. 1991) and can slow incidence of disease caused by the protozoans Perkinsus marinus and Haplosporidium nelsoni (Sprague et al. 1969, Ford 1985, Ford & Haskin 1988, Chu & Greene 1989, Chu et al. 1993, La Peyre et al. 2003). However, there is ample evidence of near-shore, intertidal oyster beds in high-salinity estuaries and coastal zones (Beaven 1955, Gunter & Geyer 1955, Lunz 1955, Nelson 1955, Copeland & Hoese 1966). It is possible then, that eastern oysters are influenced by other factors that accompany freshwater influx, such as terrestrial elements, rather than reduced salinity. Copper and zinc, like many elements in the marine environment, originate from natural terrestrial sources and are delivered to oyster beds in watershed runoff (Prytherch 1934, O'Connor 2002). Runoff with sufficient volume (i.e., streams and rivers) can generate flows that bring elements far and deep into the receiving water where subtidal oyster beds exist. Without such runoff, oysters may be confined to shallow intertidal areas adjacent to land where the elements originate. If this is correct, the availability of terrestrial elements may well be a principal determinant in the distribution of eastern oysters.

Copper and zinc are terrestrial elements of special interest because they accumulate to extraordinary concentrations in eastern oysters. They are accumulated against chemical gradients, even from low ambient concentrations, and are retained within the oyster longer than other metals, despite the apparent availability of an elimination mechanism. Moreover, zinc and copper are exclusively sequestered in oyster amebocytes (Fig. 1). This brings them into direct contact with a cell type credited with many indispensable responsibilities for oyster survival, including antimicrobial activities for defense and nutrition. These considerations implicate a physiologic reliance on copper and zinc that, if true, would support the concept that terrestrial elements are key factors in oyster success and distribution. Furthermore, it would resolve the apparent contradiction that eastern oysters often thrive at relatively polluted locations (e.g., Abbe & Sanders 1986). Fortunately, there is ample information in the literature to examine such a possibility. Because of the highly accumulated concentrations and unique association with amebocytes, uptake and disposition of copper and zinc in oysters has been widely studied.

[FIGURE 1 OMITTED]

Throughout the literature a perception exists that high concentrations of metals, copper in particular, are harmful to oysters. Yet evidence for such a perception is lacking, at least for adult eastern oysters. In fact, the highest known tissue concentrations of copper and zinc for any marine organism are found in tissues of apparently healthy eastern oyster adults. The perception of harm originates from the knowledge that such high concentrations would be quite toxic to most other biota. Yet, eastern oysters not only survive high ambient concentrations of copper and zinc, but avidly recruit the metals and accumulate them without ill effect. Several studies have shown that concentrations in oyster tissues are much higher than in the ambient environment, and that oysters assimilate and retain copper and zinc front environments with low ambient concentrations. The latter refutes any suspicion that eastern oysters passively, or inadvertently, absorb metals or that they accumulate metals only when exposed to polluted environments. If we accept that successful organisms do not expend energy without purpose, then active uptake and storage implies that oysters must accumulate copper and zinc for a physiologic purpose.

The literature is reviewed here to characterize evidence that supports a physiologic function for copper and zinc as antimicrobial agents in amebocytes. High concentrations are likely required to provide an expansive and flexible defensive and nutritional capability. More specifically, it is hypothesized that they serve as precipitating agents in extracellular clotting and as toxic antimicrobial agents for both extracellular wound protection and intracellular phagocytic activity. It is further speculated that copper, zinc, and possibly other terrestrially derived elements are essential precipitating agents for adult shell deposition (Fisher 2004). Dependence of these activities on copper may have co-evolved with the apparently exclusive requirement for this metal in eastern oyster setting and metamorphosis (Prytherch 1934). Because of these dependencies, availability of copper and other terrestrially derived metals have a major influence on the success and distribution of eastern oysters.

There are several topics relevant to this proposal and an abundance of evidence to consider. A historical perspective (Section I) summarizes early observations of copper and zinc accumulation in oysters, including emergence of a perception that high concentrations of copper are abnormal and toxic to oysters. Section II reviews the close association of copper and zinc with amebocytes, including their exclusive storage in amebocyte granules and posilive association with amebocyte numbers. Section III introduces evidence of avid and non-toxic uptake and retention of copper and zinc in granules of eastern oyster amebocytes. Field studies summarized in Section IV demonstrated greater amebocyte numbers and antimicrobial activity for oysters with high tissue concentrations of copper and zinc. In Section V, data are presented that support a role for copper and zinc in oyster defense activities. Finally, in Section VI, a series of umbrella hypotheses are presented that generate an alternative framework for interpreting high concentrations of copper and zinc, amebocyte activities in nutrition and defense, and the role of terrestrial metals in coastal distributions of eastern oyster populations.

EARLY STUDIES OF COPPER AND ZINC IN OYSTERS

Eastern oysters, once called American oysters, inhabit the coastline along the Atlantic and northern Gulf of Mexico in North America. Scientific names for this estuarine and near-coastal species have included Ostrea virginiana (or sometimes O. virginica), Gryphaea virginica, and the current Crassostrea virginica. Although a subsistence food for Native Americans and early colonials, eastern oysters did not represent a substantial commercial interest until the late 19th century (Hargis & Haven 1999). As popular consumption increased, food safety issues grew within the industry. Bacterial and chemical impurities were recognized to cause illness in consumers and, as today, cases of poisoning greatly influenced success in the market.

Green Oysters and Suspected Copper Toxicity

One such chemical impurity was copper. The occasional occurrence of a blue-green tint to oyster meats fueled fears of copper contamination from mines or copper-painted ship bottoms. Oyster tissues resembled the color of copper-salt residues found on cooking vessels (Fig. 2, left). Concerns over green oysters in North America were amplified by the lack of scientific agreement in Europe, where the European flat oyster Ostrea edulis had long been a culinary staple. Some asserted that high concentrations of copper were responsible for green O. edulis (e.g., Bizio 1845, Thorpe 1896), whereas others believed green color was simply a pigment acquired from ingested food (Gaillon 1824, Thiselton Dyer 1877). The latter group maintained that O. edulis raised in holding ponds (claires) near the Marennes region of France acquired a green tinge along the gill filaments (see Fig. 2, right) from a diatom Navicula ostrearia, which grew in the ponds. The green oysters from Marennes were a highly regarded cuisine.

[FIGURE 2 OMITTED]

It is peculiar that a green oyster controversy even existed because O'Shaughnessy (1866) had earlier resolved the conflicting evidence. He related the trial of a tradesman who, "actuated by the lawless desire of premature aggrandizement", was found guilty of selling copper-laden oysters that sickened consumers. The tradesman was unable to obtain green oysters from Marennes, so instead collected substitutes from the outfall of a copper mine in Cornwall, England. O'Shaughnessy (1866) concluded that both diatoms and copper could elicit greenness in oysters. Nonetheless, subsequent investigators in both Europe and America continued to dispute the issue. Some of the dissension is worth recounting because it has fashioned, in part, an existing predisposition that copper is a toxic element for oysters.

Sensitive to the controversy in Europe, John A. Ryder of the Smithsonian Institution reported to the United States Fish Commission that green color in American oysters was most likely a pigment from ingestion of sea cabbage (Ulva latissima) or a diatom similar to Navicula ostrearia (Ryder 1882). Much of Ryder's conviction however, appears to be based on a notion that oysters could not possibly survive the high concentrations of copper it would take to turn them green.

   "That it is not copper we may be equally certain, for any
   such quantity of a copper salt as would produce the green
   gills and patches on the mantle, such as are often observed,
   would without doubt be as fatally poisonous to the oyster as
   to a human being." (Ryder 1882, p. 410)

Green oysters were tested at two independent laboratories, both substantiating Ryder's claim; there was no evidence of copper in the green portions of tissue. Ryder saw that the green color was actually confined to independent cells that moved freely about the mantle and gills. By virtue of their amebiform ability to migrate, he proposed that these 'blood cells' could enter the digestive tract and absorb green pigment from intestinal juices. But, in spite of his findings, Ryder remained convinced that any oyster with green tissues must be diseased.

Most of Ryder's observations were independently confirmed in studies by Professor E. Ray Lankester (University College, London), who dismissed all reports of copper, at least in those green O. edulis that he examined from Europe (Lankester 1886). He strongly believed that green coloration, which he called "marennin", was imparted solely from ingestion of Navicula. Like Ryder, Lankester noted that the color was confined to certain "secretion cells" in the gill epithelium. He was puzzled by the ability of these specialized epithelial cells to independently migrate across gill surfaces. Neither Ryder (1882) nor Lankester (1886) were privy to the pioneering insights on invertebrate amoebocytes later summarized by Elie Metchnikoff (1891).

Dr. Thorne Thorpe (1896) determined that "green-bearded" or "green-finned" oysters from Marennes were so-colored from the diatom Navicula, and he found that copper quantities in these oysters were sometimes far less than found in "normal" white oysters. Nonetheless, he accepted O'Shaughnessy's contention that greenness could also originate from copper:

   "But there is no question that the greenness of certain oysters,
   especially those found in Falmouth and Truro waters,
   is due to copper. The color, both in character and distribution,
   is, however, quite different from that of the Marennes
   oyster. The green Cornish oyster is unsaleable in this country--at
   least for immediate consumption--as it leaves a distinct
   metallic taste in the mouth, similar, it is said, to that
   due to 'sucking a penny'." (Thorpe 1896, p. 107)

The color, metallic taste and copper content of green oysters gradually dissipated to "normal levels", Thorpe found, with their transfer to cleaner water. These normal concentrations were judged to be more appropriate for accepted physiologic functions, in particular as a component of the respiratory pigment hemocyanin, which at the time had been described in the blood of octopus and other molluscs (Thorpe 1896).

Professor W. A. Herdman (1896) believed the green tint he observed on experimental O. edulis and O. virginica was a sign of disease. He ran chemical tests on these oysters and, at first, concluded unequivocally that "copper had nothing to do with the disease" (p. 164). Only 1 year later, however, he reversed his stance to accept the views of O'Shaugnessy and Thorpe.

   "There are evidently several kinds of greenness in oysters,
   and whereas some may be due to normal and healthy processes,
   others must be regarded as abnormal or diseased
   conditions. It is the latter, in our experience, that contain
   the copper." (Boyee & Hcrdman 1897, p. 31)

Boyce and Herdman (1897) described experiments with O. virginica transplanted to Fleetwood, England. They found American oysters bad a different shade of green and much higher concentrations of copper than any of the English, Dutch, or Marennes oysters (O. edulis), except for occasional green specimens from Falmouth. They recounted several findings: (1) green oysters contained up to four times the copper of white oysters; (2) green tissues contained more copper than white tissues; (3) green coloration was strongly associated with the distribution of green amebocytes (described as ameboid, wandering leukocytes, Fig. 3); and (4) copper was exclusively associated with the green amebocytes. The blue-green tint of the American oysters, they surmised, was due to copper-laden amebocytes that migrated into the mantle cavity and onto the external surfaces of the tissues. They also found tinted amebocyte aggregates in the heart and sinus vessels (Fig. 4), as well as among the connective and epithelial tissues. These findings remain undisputed.

[FIGURES 3-4 OMITTED]

Boyce and Herdman (1897), like Ryder (1882), felt that copper-laden green oysters were diseased. They suggested that copper accumulation in amebocytes was

   "... due to a disturbed metabolism, whereby the normal
   copper of the haemocyanin, which is probably passing
   through the body in minute amounts, ceases to be removed,
   and so becomes stored up in certain cells." (Boyce & Herdman
   1897, p. 38)

They described a degenerative reaction in green oysters, a striking increase in amebocytes that collected in clumps and abnormally distended the blood vessels (see Fig. 4). Their evidence was eventually published in an often-cited treatise (Herdman and Boyce 1899) that clearly distinguished "diseased" American and Falmouth oysters, stained with high concentrations of copper, from the healthy Marennes and Roach River oysters tinted by pigments from diatoms. In their definitive work, Herdman and Boyce (1899) detailed the location and severity of both types of green discoloration with color drawings (see Figs. 2-4), quantified concentrations of copper and iron in oysters from several sources, and reported limited success in staining oysters green through exposure to high concentrations of soluble copper oxide and copper chloride.

In spite of a long history and plentiful evidence, controversy over the role of copper in green oysters persisted into the 20th century (Pease 1911, Nelson 1916 [cited from Hunter & Harrison 1928], Ranson 1924, Ranson 1925, Yonge 1928, Galtsoff & Whipple 1930, Takatsuki 1934). Some confusion may have been due to the fact that a green tint is most easily seen when amebocytes are present on the surface of the tissues, a condition that may vary for reasons unrelated to copper accumulation. For eastern oysters, the controversy was seemingly resolved when Galtsoff and Whipple (1930) found green oysters from New Haven Harbor (Long Island Sound) to contain copper at concentrations of 1,217-2,719 [micro]g [g.sup.-1] (dry weight) compared with only 82-138 [micro]g [g.sup.-1] in white oysters from Onset Bay (MA). There is now a general acceptance that copper can accumulate in amebocytes to concentrations high enough to impart green coloration. In tact, Galtsoff (1964) noted that,

   "In the case of pronounced green discoloration the presence
   of metallic copper may be demonstrated by inserting in the
   tissues a well-polished steel knife; the surface becomes copper
   plated in a short time. This simple method can be used
   profitably for a qualitative demonstration of the presence of
   copper." (Galtsoff 1964, p. 388)

There is no longer any doubt that copper at high concentrations in eastern oysters will create greenish coloration. Abbe and Sanders (1986) predicted that concentrations of copper must be more than 700 [micro]g [g.sup.-1] (dry) for oysters to turn green, and that all oysters would be green at 2,000 [micro]g [g.sup.-1]. Along with this acceptance is the unmistakable perception that such high accumulations are abnormal, are a direct consequence of environmental pollution, and are responsible for a toxic or diseased reaction in oysters.

One additional study on the association of copper with eastern oysters is particularly significant. In both field and laboratory studies, Prytherch (1934) found copper was required for setting and metamorphosis of eastern oyster larvae. In studies at Milford, Connecticut, he found larvae would not set if copper was excluded from the seawater. He further showed that by removing copper from the seawater immediately after setting, metamorphosis would cease until its re-addition. Prytherch (1934) cemented oyster larvae to glass dishes to microscopically examine and chronicle morphologic changes during metamorphosis. He observed that larval pigment spots (more often called 'eye spots') were composed of about 300 aggregated, immobile amebocytes. Addition of copper to the seawater initiated movement of the pigment spot amebocytes and their inaugural migration into the bloodstream. Although important to this review and pivotal to the argument for amebocyte participation in shell deposition (Fisher 2004), these findings have been generally overlooked in the literature and have never been validated.

High Zinc Accumulation and Radioactive [sup.65]Zn from Nuclear Reactors

The earliest reports of zinc accumulation by eastern oysters stemmed from broad-based efforts to determine whether all living organisms contained zinc and, if so, what physiologic purpose the element might serve. Two early contributions (Bradley 1904, Mendel & Bradley 1905) revealed that concentrations of zinc oxide in the hepatopancreas of Sycotypus canaliculatus, a large carnivorous gastropod, were well above 10%. They found only negative or "doubtful" concentrations for a variety of other New England marine crustaceans and molluscs, including O. virginiana. Mendel and Bradley (1905), based on existing knowledge that many invertebrates contained copper in the respiratory pigment, suggested zinc might bind to an analogous respiratory pigment. Although it was never characterized, they named this presumptive pigment 'hemosycotypin' (Mendel & Bradley 1906, Mendel & Bradley 1907). Zinc was found in the tissues of several marine organisms from the Tortugas by Phillips (1917; Table 1), who concluded it must be a natural constituent because the Tortugas were far removed from anthropogenic influences (although some samples were collected from the moat at Fort Jefferson). This approach, surveying fauna to generate hypotheses of potential function, reached a pinnacle in the work of Vinogradov (1953), who examined metal content in several marine phyla.

Many early observations on zinc in oysters were focused on the high concentrations that were accumulated. Hilmer and Wichmann (1919) found zinc and copper accumulated to high concentrations in Atlantic coast eastern oysters; they believed the elements were accumulated from the food chain because concentrations were so much greater than for seawater.

   "There appears, moreover, to be no direct, uniform ratio
   between the quantity of copper and zinc in oysters and the
   amount in the seawater in which they are found, although it
   is true, in general that oysters contain larger proportions of
   the metals when grown in seawater highly contaminated
   with metallic wastes from smelters and other factories."
   (Hilmer & Wichmann 1919, p. 217)

Although they noted a potential role for zinc in respiration, Hiltner and Wichmann (1919) attributed no biologic significance to these exceptionally high concentrations. In fact, they concluded that the concentrations seemed too great to even consider a functional role.

Bodansky (1920) examined 20 different marine species from the Gulf of Mexico for accumulation of zinc and found the concentrations in eastern oysters extraordinary. It was, he suggested,

   "... a striking phenomenon that the oyster living in a water,
   not contaminated by industrial wastes, should contain in its
   tissues more than 35,000 times as much zinc as is present in
   an equivalent weight of water." (Bodansky 1920, p. 403)

Bodansky (1920) also concluded that the source of zinc was probably oyster food, and he believed that zinc might play a role in respiration. Based on chemical analyses of dissected oyster tissues, he reported that zinc was distributed evenly in the digestive organ, mantle, and gills, yet was considerably lower in the adductor muscle.

Investigations into zinc accumulation by oysters gained significance in the mid-1950s as coastal zones and estuaries were exposed to increasing levels of industrial waste as well as radioactive contamination from nuclear reactors, fuel reprocessing plants, and fallout from intensive nuclear weapons testing. In a study to characterize the radioactive pollution from a nuclear power station in Essex, England, Preston (1968) examined [sup.137]Cs, [sup.60]Co, [sup.55]Fe, and [sup.65]Zn in the receiving estuary and found that [sup.65]Zn was the only nuclide appreciably assimilated by oysters. The amount accumulated was proportional to the exposure, as demonstrated by diminishing concentrations of [sup.65]Zn as oyster collections moved further from the source (Table 2). Because of this capacity to accumulate [sup.65]Zn and a link with human food consumption, oysters were considered an ideal indicator organism to assess food-borne risk (Marthy et al. 1959, Fitzgerald & Skauen 1963, Wolfe 1970a, Romeril 1971). In general, these reports showed that oysters near radioactive effluents accumulated [sup.65]Zn at levels too low to affect human health; reports describing effects on oysters have not been found.

Impetus for food safety led to advances in radioactive tracing technology. Methods were developed to track the differential uptake of [sup.65]Zn by aquatic species and its passage through biogeochemical cycles (Chipman et al. 1958, Watson et al. 1961, Alexander & Rowland 1966, Duke 1967). In one such study, Chipman et al. (1958) showed that clams, scallops, and particularly oysters (C. virginica) accumulated zinc thousands of times higher than seawater. Using radioactive [sup.65]Zn. they found uptake and exchange Of zinc between the seawater and tissues to be rapid and in continuous flux. They also noted a considerable amount of [sup.65]Zn in oyster shells, which they attributed to adsorption because of the dynamic uptake and exchange. Their studies included measurement of uptake by excised oyster gill sections, influences of chelating agents, and tissue distributions of injected [sup.65]Zn. They attempted to extract [sup.65]Zn from oyster tissues and found that about half the zinc was extractable by water alone.

Wolfe (1970a) described studies on C. virginica from North Carolina that used both stable and isotopic zinc. He found that soft tissues contained about six times the zinc concentration of shells, but because the shells were so heavy they contained nearly 45% of the total zinc in an oyster. Similar to Bodansky (1920), Wolfe found that tissues with large external surface areas (gills, mantle, labial palps, gonad, and digestive gland) had higher concentrations of zinc than other tissues (adductor muscle, pericardial sac). Romeril (1971) demonstrated that C. angulata (Portuguese oysters) accumulated [sup.65]Zn to higher concentrations than O. edulis, but found similar tissue distributions. This fact, he believed, indicated a common uptake mechanism among oyster species. Romeril (1971) found a high concentration of [sup.65]Zn with the shell, as had previous investigators (Chipman et al. 1958, Fitzgerald & Skauan 1963, Wolfe 1970a), but showed that co-incubation with iron or cobalt would reduce this [sup.65]Zn concentration, presumably due to binding-site competition. This finding supported his conclusion that zinc was adsorbed to the shell and in constant flux.

Few hypotheses on a potential role for such high concentrations of accumulated zinc emerged from these studies. It was clear, since the study of Bodansky (1920), that some zinc was bound to protein and might serve as enzyme co-factors; but calculations by Pequegnat et al. (1969) indicated that accumulation was much greater than any known or speculated requirements. Potential roles for zinc in enzyme activation have been investigated: Zinc-dependent enzymes in C. virginica include carbonic anhydrase, alkaline phosphatase and malic dehydrogenase (Wolfe 1970b), and in O. edulis they include carbonic anbydrase, alkaline phosphatase, carboxypeptidase A, and [alpha]-D-mannosidase (Coombs 1972). However, in both cases the investigators concluded that accumulated zinc was far in excess of enzymatic zinc requirements. This inconsistency between requirement and accumulated concentrations led Wolfe (1970a) to suggest.

   "The accumulation of zinc and other trace metals may be
   partly a coincidental result of poor discrimination by the
   biologic mechanism for calcium uptake and shell deposition.
   This possibility is consistent with the observed seasonality
   of trace element concentration ... where contents of
   manganese, iron, copper, and zinc were higher during
   warm months when shell deposition is greatest." (Wolfe
   1970a, p. 55)

Considering the great interest that many early investigators placed on a role for copper and zinc in oyster physiology, the lack of ensuing studies is remarkable. This dwindling attention may have risen from a sense that such extraordinarily high concentrations precluded any possible physiologic purpose.

METAL ACCUMULATION IN OYSTER AMEBOCYTES

Ostensibly, eastern oysters flourish in shallow coastal areas and estuaries because freshwater inflow creates reduced or variable salinities that thwart marine predators (e.g., oyster drills; Wells 1961, Menzel et al. 1966) and marine pathogens such as the protozoans Perkinsus marinus (Andrews & Ray 1988) and Haplosporidium nelsoni (Ford & Haskin 1988). Yet, freshwater inflow also transports terrestrial elements, including both natural and anthropogenic metals, from the watershed. Any dependence by oysters on terrestrial metals must have evolved from natural geologic sources because anthropogenic sources are relatively recent. Nonetheless, the two sources are sometimes confounded. Both metals and polycyclic aromatic hydrocarbons (PAH), even though they are naturally derived, are often associated with human activities because their levels are elevated in urban and industrial discharge (Hiltner & Wichmann 1919, Hunter & Harrison 1928, Chipman et al. 1958, Galtsoff 1964, Pringle et al. 1968, Roosenburg 1969, Boyden & Romeril 1974, O'Connor 2002). Elevated concentrations of metals and PAH, along with wholly anthropogenic polychlorinated biphenyls (PCB) and pesticides, are usually considered indicators of environmental pollution. Although there are natural sources for metals, a perception persists that certain metals, including copper and zinc, are environmental contaminants with a potential for toxic, detrimental effects on human health and the condition of flora and fauna.

Bivalves as Sentinels of Environmental Pollution

Pesticides, PCBs, PAHs, and metals are the most common chemicals monitored in the coastal systems (O'Connor 2002). These chemicals are generally present in heavily used coastal zones and, accordingly, have been the focus of monitoring programs Io determine status and trend of chemical pollution. One of the most comprehensive exposure monitoring programs in the United States, the National Marine Fisheries Service Status and Trends Program Mussel Watch Project, uses soft tissue concentrations in sentinel mussels and oysters to characterize local exposure conditions in relation to other locations (O'Connor & Ehler 1991, O'Connor 2002). Bivalves, presumably because of their filter-feeding behavior, are notorious for their ability to concentrate metals. This characteristic has provided an opportunity to use bivalves to monitor environmental chemicals that are otherwise below detection in the water column or that are highly variable over time.

Bivalve tissue concentrations used in the Mussel Watch Project and other environmental monitoring programs are better used to characterize long-term trends than to estimate actual exposure concentrations (Kopfler & Mayer 1973, Ikuta 1958a, Ikuta 1958b, Roesijadi 1996, O'Connor 2002). Individual bivalves, even when exposed to the same ambient concentrations of chemicals, can accumulate different tissue concentrations. This is shown by the substantial variability among oysters sampled at the same time and location. Some concentrate elements with such effectiveness that they have been termed "super-accumulators" (Lobel & Wright 1983, Wright et al. 1985). Variability in accumulation is often linked to size, age, filtration rate, and reproductive status, but can also vary with environmental factors that include salinity, food availability, and chemical speciation (Chipman et at. 1958, Boyden & Phillips 1981, Phelps & Hetzel 1987, Wright & Zamuda 1987, Bryan & Langston 1992, Roesijadi 1996). Such differences have led to the general conclusion that 15 to 25 oysters (Fig. 5) must be analyzed to provide a representative estimate of metal concentrations for a given site (Kopfler & Mayer 1969, Gordon et al. 1980, Boyden & Phillips 1981, Wright et at. 1985, Sanders et at. 1991, Reidel et al. 1995, Jiann & Presley 1997).

[FIGURE 5 OMITTED]

Variable accumulation has confounded interpretations of bivalve tissue analysis and has limited its effectiveness as an indicator of environmental exposure. As Roesijadi (1996) states,

   "A consensus on the best approaches for using the metal
   content of oysters to monitor environmental metal contamination
   has yet to emerge, although the use of oysters as
   sentinel organisms is a common activity." (Roesijadi 1996,
   p. 520)

To offset this variability, monitoring programs often monitor during the same season each year and use animals of similar size. Other approaches include the use of inbred organisms in transplant scenarios (Roesijadi 1996).

High Concentrations of Copper and Zinc are Accumulated in Oysters

Despite uncertainties in interpreting tissue concentrations, evidence supporting zinc as the most prominent element in eastern oyster tissues is overwhelming (Table 3). Zinc is usually followed in quantity by copper (Shuster & Pringle 1969, NOAA 1987) or iron (Lytle & Lytle 1990, Jiann & Presley 1997). Concentrations of zinc invariably comprise 80% or more of the total metal accumulated in eastern oysters, a proportion unique among marine invertebrates (Vinogradov 1953, Pequegnat et al. 1969) and even among filter-feeding bivalves (Pringle et al. 1968, Boyden & Romeril 1974, Boyden 1975). O'Connor (2002) reported that eastern oysters accumulated about 10 times the copper, silver, and zinc of mussels Mytilus edulis. This finding is upheld by relatively long-term data collected through the Mussel Watch Project (O'Connor 1996, Table 4), as well as direct comparisons of mussel and oyster collections from the same sites in Long Island Sound (O'Connor 1994) and Chesapeake Bay (Reidel et at. 1995).

Copper has also been found particularly high in oysters relative to other lamellibranchs (Vinogradov 1953, Brooks & Rumsby 1965).

   "Except for species of Ostrea, all Lamellibranchiata contain
   copper in quantities not higher than the average
   amount found in Gastropoda ... species of Ostrea have
   proved to he richest in copper. " (Vinogradov 1953, pp.
   349-350)

Copper, as described earlier, can be extremely high in oysters exhibiting a green tint. Hiltner and Wichmann (1919) found a mean of 403 [micro]g copper [g.sup.-1] (dry weight) for normal "white" oysters in the northeast United States (see Table 3), but noted that bluish oysters collected from Perth Amboy, NJ, contained 7,435 [micro]g [g.sup.-1]. Similarly, Galtsoff and Whipple (1930) found green oysters to contain 1,954 [micro]g copper [g.sup.-1], much higher than the mean 109 [micro]g [g.sup.-1] they recorded for white oysters.

Crassostrea virginica may accumulate more copper and zinc than C. gigas. Pringle et al. (1968) found from broad geographic surveys that eastern oysters along the Atlantic coast contained 5 to 10 times the zinc and copper as Pacific Coast C. gigus. Okazaki and Panietz (1981) found higher accumulations of zinc (but not copper) in C. virginica than C. gigas at a site in California. Boyden (1975) found O. edulis to accumulate more zinc and less copper than C. gigas sampled from the same site. Although there is some consistency among reports, inferences from these data must be tempered by recognition of the high variability noted above.

Copper and Zinc Are Accumulated in Granular Amebocytes

Orton (1923) was among the first to measure high concentrations of copper and zinc in oyster amebocytes. During the summer of 1920, O. edulis in the commercial oyster beds of the Thames Estuary suffered unusually high mortalities. Because mortality events were frequent during 1919 to 1921 in Europe (Italy, France, the Netherlands, Ireland, and in several regions of England), Orton postulated a relationship to wartime contamination, (i.e., hazardous munitions and material strategically dumped or lost from damaged and sunken ships). Accordingly, he examined O. edulis for concentrations of trinitrotoluene, nitrites, oil, and metals. Metals in Thames Estuary oysters were low, so metal toxicity was excluded. Yet, his investigations revealed that the amebocytes of oysters could contain high concentrations of copper (25,900 [micro]g [g.sup.-1] dry weight, using 5x wet weight values), zinc (40,650 [micro]g [g.sup.-1]) and tin (2,450 [micro]g [g.sup.-1]). These concentrations were many times higher than whole animals or sediments from the same location. Orton (1923) inferred from his findings that most, if not all metals in oysters were concentrated in amebocytes.

His investigation led to another intriguing conclusion. When he transplanted metal-laden oysters to environments with low metal concentrations, they eventually eliminated their metal burdens. Rather than steady-state equilibria, he suggested a novel route of elimination.

   "It seems possible that metals are excreted by the blood-cells
   leaving the body of the oyster and carrying the metals
   with them." (Orton, 1923, p. 17)

Orton's (1923) contributions to the field were thus 3-fold. His studies sustained earlier speculation that green oyster amebocytes had high concentrations of copper; he suggested that zinc, tin, and possibly other metals were retained in amebocytes; and he proposed a novel method for elimination of metals (i.e., exomigration of amebocytes).

The chemical nature of the green pigment in amebocytes was systematically investigated in 1927 when Paul S. Galtsoff and Samuel Lepkofsky applied "microchemical reactions" to paraffin-embedded sections of normal and green colored American oysters from New England (Galtsoff & Whipple 1930). Treatment with potassium ferrocyanide and hematoxylin confirmed fully the speculations of Herdman and Boyce (1899) that the intensity in green color was in proportion to the copper content in the oyster and that the copper was located almost exclusively in the green leukocytes (amebocytes).

Ultimately, a detailed characterization of both copper and zinc in oyster amebocytes was provided by Craig L. Ruddell (1971) in research published from his doctoral dissertation. He used traditional and novel histochemical staining techniques to locate both of the metals in membrane-bound granules (lysosomal derivatives) in amebocytes of Pacific oysters, C. gigas. Amebocytes laden with copper and zinc were found distributed throughout the hemolymph and tissues. Neither metal was found in any other cell types, including gill and mantle epithelial cells. It seems that some amebocytes (basophilic granular ameboeytes, BGA) contained prinmrily zinc, whereas others (acidophilic granular amebocytes, AGA) contained primarily copper (as presaged by Boyce & Herdman 1897, p. 33). Even so, Ruddell suggested that some amebocytes might store both elements. He did not report on other metals, (e.g., tin [Orton 1923], iron [Galtsoff 1938, Galtsoff 1953] and manganese [Galtsoff 1964]), that might also be retained and transported by amebocytes.

Positive Association of Metal Content with Amebocyte Number and Distribution

Ruddell later hypothesized that concentrations of copper and zinc in different oyster tissues would reflect the tissue distribution of amebocytes, lie examined C. gigas and C. virginica from both contaminated and reference sites and established that a positive correlation existed between the number of tissue amebocytes and tissue concentrations of copper and zinc (Ruddell & Rains 1975). Mantle tissues of oysters from element-rich sites had higher zinc and copper in their amebocytes than those from element-poor sites. In addition, zinc concentrations in the mantle were found linearly and positively associated with the number of BGAs counted in mantle histologic sections (Table 5). This relationship was reinforced by a positive association between BGAs and zinc concentrations derived from examination of mantle (high zinc) and digestive diverticulum (low zinc) tissues from the same oysters. Copper analyses yielded similar trends, but the linear association between amebocytes and copper concentrations was not as distinct as that of zinc. The fact that copper was associated primarily with AGAs rather than BGAs (Ruddell 1971) might have created this difference.

Overall, the results of Ruddell & Rains (1975) established a positive relationship for copper and zinc with amebocyte number and distribution. The metals occurred wherever metal-carrying amebocytes were stationed, sometimes in the adductor muscle and pericardial sac but more often in the gills, mantle, labial palps, gonad, and digestive gland. Ultimately, Ruddell and Rains (1975) used data from both C. gigas and C. virginica to estimate that nearly all of the zinc (and probably copper) in an oyster was retained within amebocytes.

Localization of Copper and Zinc in Amebocyte Granules

Robert S. Brown (1975), in his doctoral dissertation, further characterized the relationship of copper and zinc with circulating amebocytes of eastern oysters. Using C. virginica from Maryland, Brown applied histochemical techniques to show that copper and zinc were localized in membrane-bound intracellular granules. Virtually all intracellular granules stained positive with Mallory's hematoxylin for copper (Fig. 6) or with a fluorescent dye labeled specifically to bind with zinc (Fig. 7). Nongranular portions of cytoplasm and agranular amebocytes were not stained by either chemical. Transmission electron microscopy (unstained sections) showed that only the intracellular granules of amebocytes exhibited the electron-dense signature of metals (Fig. 8). High-density material was confined inside the granules, and was not bound by a separate membrane distinct from that of the granule (Fig. 9). Brown concluded that the cytoplasmic granules of amebocytes, the same granules described for C. gigas by Ruddell (1971), were storage sites for electron-dense metals.

[FIGURES 6-9 OMITTED]

Brown (1975) was also able to quantity copper and zinc within amebocytes using a relatively novel technique that coupled energy dispersive x-ray analysis to scanning electron microscopy (EDAX-SEM; Fig. 10). Nearly all of the 250 amebocytes probed with EDAX-SEM contained both copper and zinc, although a few cells contained only zinc. The metals were absent from nuclei and from agranular portions of the cytoplasm. Brown determined that C. virginica amebocytes could contain as much as 0.3% of their dry weight in copper and 9% of their dry weight in zinc (Table 6). Using EDAX-SEM data, he calculated the content of a single amebocyte to be 6.0 x [10.sup.-13] g Cu and 2.5 x [10.sup.-11] g Zn, and the content of a single amebocyte granule was 8.5 x [10.sup.-15] g Cu and 3.5 x 10 [10.sup.-13] g Zn. These calculations assumed 89% granular amebocytes in a sample and 71 granules per cell. Using the same technique, cell-free hemolymph was found to contain copper at an average 0.159 [micro]g m[L.sup.-1] and zinc at an average 8.372 [micro]g m[L.sup.-1].

[FIGURE 10 OMITTED]

These reports of metal accumulation in amebocytes of C. virginica led George et al. (1978) to examine "green sick" O. edulis collected from metal-polluted sites in Cornwall. England, as a representative model for studies of bivalve metal detoxification. Their approach included a technique similar to that used by Brown (1975). Using X-ray microprobe analysis (XRP-TEM, a dispersed energy detector coupled to transmission electron microscopy), they located and quantified electron-dense elements within cells. Comparisons of numerous tissues confirmed for O. edulis the earlier findings on C. gigas and C. virginica; copper and zinc were localized exclusively within amebocytes. Further confirming the work of Ruddell (1971), George and co-workers found two types of metal-containing amebocytes, BGAs that contained zinc in 1-p,m-diameter granules and AGAs that contained copper in 0.8-[micro]m-diameter granules. Analysis of hemolymph and cell-free plasma showed that 70% to 77% of the copper and zinc in the hemolymph was contained in the circulating amebocytes. This estimate, and others comparing hemolymph to cell-free plasma, may well have been influenced by loss of metals from the amebocytes during centrifugation and processing.

Members of the same research learn (Pirie et al. 1984) later showed that amebocytes from O. edulis, C. gigas, and O. angasi all had a capacity to accumulate copper and zinc. The latter two species were reported Io possess a single cell type that accumulated both copper and zinc, whereas O. edulis had copper-specific cells and zinc specific cells as well as nonspecific copper/zinc (mixed) cells. The metal-specific cells in O. edulis were found only from a high-metal environment and mixed cells were from less contaminated sites. The authors acknowledged a possibility that the amount or type of metal present in the environment might have influenced the proportion of different cell types in their studies.

Thomson et al. (1985) compared metals in different cells of C. gigas from Tasmania and Wales. Whole body metal analyses showed that oysters from Tasmania contained approximately twice the copper and zinc concentrations of the Welsh specimens, yet XRP-TEM analytical results showed that individual amebocytes from the two groups of animals contained about the same concentrations (see Table 6). This supported the proposition of Ruddell and Rains (1975) that the copper and zinc concentrations were linked to amebocyte number. Thomson et al. (1985) estimated that amebocytes contained -90% of the copper and zinc in gill and mantle tissues and calculated the concentrations in amebocyte granules to be 506 [micro]g copper [g.sup.-1] and 6,375 [micro]g zinc [g.sup.-1] (dry weight, converted from mM [kg.sup.1]), values much less than estimates for O. edulis (Orton 1923) and C. virginica (Brown 1975).

ACTIVE, CONTROLLED UPTAKE AND RETENTION OF COPPER AND ZINC

Copper and zinc accumulation in eastern oysters is so great that many have dismissed the possibility of a physiologic role (Hiltner & Wichmann 1919, Pequegnat et al. 1969, Wolfe 1970a). For copper, at least, high accumulation has even been related to toxicity or disease (Ryder 1882, Herdman 1896, Boyce & Herdman 1897. Korringa 1952). This has led to an overriding impression that copper and zinc are in excess and sequestered in oyster amebocytes solely to prevent toxicity.

   "Iron, copper, and zinc maybe be stored in the tissues and
   in some blood cells as excess materials which are slowly
   eliminated." (Galtsoff 1964, p, 390)

Simkiss & Mason (19831 introduced three theories to explain the magnitude of copper and zinc deposits in oyster amebocytes: (I) Metals are phagocytosed as foreign material by amebocytes but, being resistant to digestion, are accumulated in the cytoplasm; (2) amebocytes are a specific detoxification system that remove metal ions from the hemolymph to keep concentrations below toxic levels: (3) amebocytes accept metals from other cells (e.g., gill cells) Io transport them through the blood stream to other tissues (e.g., kidney) for storage and eventual excretion. Yet, to the authors, none of the these theories were totally satisfying:

   Many of the more easily detected accumulations of metals
   in molluscs are tissue specific and they are often associated
   with particular deposits of granules. These are almost universally
   interpreted as detoxification or excretory systems,
   but there is as yet virtually no acceptable evidence
   for such a conclusion." (Simkiss & Mason 1983, pp.
   154-155)

In fact, for eastern oysters, any perception that copper and zinc are sequestered simply for eventual elimination is confounded by an assortment of existing data. Eastern oysters do not seem harmed by high concentrations of copper or zinc in the ambient environment or by high concentrations accumulated in their tissues. Rather, they stein to accumulate copper and zinc only when they have the amebocyte capacity to safely do so. When amebocytes are available, oysters can accumulate copper and zinc from low ambient concentrations and retain them even though effective elimination mechanisms are available. These considerations, described below, not only refute a perception that copper and zinc are sequestered solely for detoxification and elimination, but sustain a contingency that they are stored for a physiologic purpose.

Oysters Concentrate Copper and Zinc front Low Ambient Water Concentrations

Comparisons of oyster soft tissue concentrations with ambient water concentrations of copper and zinc have clearly demonstrated preferential accumulation (Korringa 1952, Boyden & Romeril 1974. Boyden 1975, Simkiss et al. 1982, Roesijadi 1996). Hunter & Harrison (1928) found high concentrations of copper and zinc in eastern oysters had no direct relationship to body weight, other metals, or concentrations in the seawater. They concluded,

   "There is reason to believe that oysters will absorb from the
   water almost any substance which it contains." (Hunter &
   Harrison 1928, p. 9)

Moreover, it was apparent from even the earliest reports (e.g., Boyce & Herdman 18971 that these metals could be accumulated from metal poor environments. Measured tissue concentrations have been so high that active, selective uptake seemed inevitable.

   "The high proportions of zinc and copper in oysters from
   beds in the vicinity of industrial plants using these metals
   can be readily accounted for. The high zinc content of those
   from beds far removed from any known source of metallic
   contamination may be explained by the probability that oysters
   gradually remove traces of the metals from the water
   and store them in their tissues." (Hunter & Harrison 1928.
   p. 8),

There is evidence that zinc is more avidly concentrated at low ambient water concentrations than at high. Chipman et al. (1958) compared zinc concentrations of eastern oysters and seawater from several locations along the US Atlantic seaboard. Zinc content was higher in oysters from locations with high ambient zinc, but concentration factors were five times higher at the lowest ambient concentration (Table 7). Preston (19661 estimated a [10.sup.5] concentration factor for zinc. Copper is also concentrated from ambient waters (Ikuta 1958a, 1958b, Pringle et al. 1968, Shuster & Pringle 1969, Kopfler & Mayer 19731. Roesijadi (19961 concluded that uptake of anthropogenic copper and zinc is superimposed on an already strong natural proclivity. This capacity demonstrates the frailty of tissue analyses to detect pollution, and refutes any perception that metal uptake is passive and indiscriminate.

It is re-emphasized that the substantial concentrations of zinc and copper in eastern oysters are almost exclusively confined to the granules of amebocytes (Section II). Zinc and copper distribution in oysters is thus determined by the number and distribution of amebocytes in different tissues (Ruddell & Rains 1975) and their individual capacity to retain the elements in granules. Concentrations of copper and zinc are high in oysters, higher in granular amebocytes, and even higher in amebocyte granules, underscoring the conclusion of George et al. (,1978) that copper and zinc are actively recruited into granules against a strong chemical gradient. Even so, there has been no published investigation of the mechanism of uptake and no evidence of metabolic cost. Galtsoff and Whipple (1930) colnpared oxygen consumption rates of 'normal' eastern oysiers from Onset Bay (MA) with green eastern oysters from New Haven Harbor (CT). Results showed a slight increase in oxygen consumption by green oysters, but the significance was considered doubtful, based partly on the high variability attributed to muscular activity, gill ciliary activity, and season. Regardless, there seems little doubt that oysters must expend energy to assimilate and retain these metals, especially when they are scarce in the ambient environment.

Logarithmic Uptake of Copper and Zinc in Eastern Oysters

Metal content in tissues of bivalves and other organisms at any given time is the difference between two dynamic processes, influx (uptake rate) and efflux (release rote) (Rnesijadi 1996). Changes in influx and efflux rates can be estimated by transferring organisms from low to high (to estimate influx) or high to low (to estimate efflux) exposure concentrations. The number of studies that examine influx outweigh those that examine efflux, but both are equally important in the ultimate disposition of metals in oysters.

Jiann and Presley (1997) suggested that, in general metals accumulate in oysters because of rapid uptake and slow elimination. Whereas rapid uptake may be accurate for most metals, copper and zinc exhibit a unique pattern. Shuster and Pringle 11969) continuously exposed eastern oysters in the laboratory to two elevated concentrations each of Cd, Cr, Cu, or Zn and followed their accumulation in soft tissues over a 211-week period. Upon exposure, cadmium and chromium showed an immediate spike in tissue concentrations that eventually slowed, presumably because they approached a steady state between influx and efflux (Fig. 11). The rate of uptake for copper and zinc, however, did not spike appreciably upon exposure. Instead, a consistent, logarithmic increase in tissue residues was maintained, without decline, throughout the study. Shuster and Pringle 11969) assigned these unexpected results to high initial tissue concentrations. More likely, the two accumulation patterns distinguished two different mechanisms of uptake. Cadmium and chromimn exhibited a phased pattern typical of direct tissue absorption whereas the pattern for copper and zinc imply a controlled assimilation that might explain why oysters held in high ambient concentrations do not necessarily have high body burdens (Reidel et al. 1995).

[FIGURE 11 OMITTED]

Controlled uptake may be a consequence of the need to sequester copper and zinc in amebocyte granules. Only a limited number of granules exist in an amebocyte (~71 granules, as estimated by Feng et al. 1971), so there is a limited amount of each element that can be retained (amebocyte metal-carrying capacity). Additional uptake of copper and zinc from ambient sources would be limited until new cells become available. The logarithmic rate of accumulation observed by Shuster & Pringle (1969) seems consistent with this supposition: although cell proliferation has not been well characterized, new cells are probably produced at a logarithmic rate. The fact that uptake might be controlled by a physiologic condition (i.e., the availability of amebocytes with metal-carrying capacity), challenges any perception that copper and zinc are indiscriminately absorbed from metal-rich environments. Rather, it seems that they are actively incorporated when (and only when) amebocytes with metal-carrying capacity are available. Once this capacity is exceeded, further accumulation requires recruitment of new amebocytes, either through proliferation (i.e., hemopoiesis) or activation of existing amebocytes.

Slow Elimination of Copper and Zinc by Eastern Oysters

Just as transfer of oysters from a reference to contaminated site can be used to estimate influx, the converse can provide insight to efflux, or elimination of chemicals. By transferring eastern and Pacific oysters in California from a contaminated to a reference site, Okazaki and Panietz (1981) estimated efflux of various metals. They monitored the loss of metals over a 56-d period from the mantle, gill, digestive gland and kidney, and calculated biologic haft-lives ([B.sub.1/2]) for each tissue. Half-lives for all four metals tested (Ag, Cu, Hg, Zn) were particularly long for eastern oysters (Table 8). and generally 5x longer than those for C. gigas. Ruddell and Rains (1975) had earlier found higher concentrations of copper and zinc and higher numbers of basophils in C. virginica than in C. gigas from these same two sites. It seems that eastern oysters retain metals for a relatively long period, and significantly longer than Pacific oysters.

Other studies have reached similar conclusions. Zaroogian (1979) collected oysters from Long Island Sound (New York) and held them in seawater troughs for 56 weeks to examine depuration of copper and cadmium. Copper concentrations in the troughs ranged from 1-2 [micro]g [L.sup.-1]. He found no statistically significant decrease in copper during this time, and actually noted an increasing trend. Greig and Wenzloff (1978) transferred eastern oysters from a contaminated site (Housatonic River. CT, USA) to an uncontaminated site (Beaufort, NC, USA) and found silver, cadmium, and zinc did not substantially decrease during 40 weeks, and copper was marginally decreased on only one (27-wk) sampling date.

Slow efflux of copper and zinc from eastern oysters indicates that, because even though amebocytes are proficient at detoxifying copper and zinc (i.e., sequestering large amounts in membrane-bound granules), they are strikingly inefficient at eliminating them. This inability is not for lack of a physiologic mechanism. The most likely means for oysters In eliminate copper and zinc is amebocyte exomigration.

   "Amoebocytes seem to have an important phagocytic role
   during excretion in molluscs, particularly in eliminating
   insoluble particles from the circulation. The wandering
   amoebocytes phagocytose the particles and migrate to the
   gut, the pericardium, the excretory organs or the mantle
   cavity; they either move out of the body through these viscera
   or return after releasing the particles." (Narain 1973,
   p. 8)

This is, of course, the same elimination mechanism anticipated by Orton (1923), Yonge (1928) and others who observed exomigration of green, copper-bearing amebocytes in oyster psuedofeces. Since then, amebocyte exomigration has been described as a means for oysters to eliminate a variety of ingested materials. including india ink, carmine dye, neutral red, aniline oils, coal tar, iron, and fluorescent beads (Takatsuki 1934, Ranson 1936, Stauber 1950, Galtsoff 1953, Ruddell 1971). In one report, Galtsoff (1953) suggested that iron was actively ingested and then eliminated by amebocytes passing across the mucous cells of the gill and mantle. Ports (1967) considered the evidence for amebocytic elimination strong enough to permit an analogy of these cells with the reticuloendothelial system of vertebrates.

A description of amebocyte exomigration in C. virginica was provided by Stauber (1950), who injected india ink into oyster hearts and collected samples over a 42-day period for gross and microscopic examination.

   "The ink suspensions agglomerated readily and produced
   emboli which virtually occluded the arterial vessels of viscera,
   mantle and adductor muscle. Subsequent events, with
   considerable overlapping, were in sequence: (a) phagocytosis
   of the injected ink particles by mobile phagocytes, (b)
   distribution of the ink in the phagocytic amebocytes to all
   parts of the organism with concomitant resolution of the
   emboli and (c) eventual elimination of the ink from the
   organism by the migration of ink-laden phagocytes through
   the epithelial layers of the alimentary tract, digestive
   diverticula, palps, mantle, heart and pericardium into lumina
   from which they were voided." (Stauber 1950, pp. 239-240)

The presence of ink-darkened 'dejecta and rejecta' supported Stauber's conclusion that the bulk of the injected ink was eliminated by migration of amebocytes into lumina that opened to the outside of the oyster. A similar process had been described for O. edulis by Takatsuki (1934), who showed that carmine particles injected into the body of the oyster were ingested by amebocytes and then distributed into excretory tubules, pericardial epithelium, gonoducts, rectum, mantle cavity and blood vessels. The presence of carmine-containing amebocytes in the mantle cavity provided evidence that they traversed epithelial layers to be discharged in the pseudofeces.

A mechanism is thus available to eastern oysters to eliminate metals and other substances but, by all indications, is not used for copper and zinc. Both metals are retained in the oyster amebocytes at high concentrations for periods well beyond that needed for exomigration. In all likelihood, retention of copper and zinc exceeds many times over the life span of an individual amebocyte, implying that the elements are re-incorporated into new cells. This is probably achieved through phagocytosis of damaged and dead amebocytes (Scro & Ford 1990), and may include apoptosis, wherein damaged and senescent amebocytes pinch off cytoplasmic fragments that are ingested by younger amebocytes without an inflammatory response (Sanderson 1982, Sunila & LaBanca 2003). Metals sequestered in amebocyte granules could be retained through numerous amebocyte cell cycles.

No Evidence of Lethal Copper or Zinc Toxicity to Eastern Oyster Adults

The sensitivity of embryonic and larval stages of C. virginica to water-borne metal toxicity has been well documented. Results of 2-day embryo tests (Calabrese et al. 1973) showed copper to be 100% lethal to eastern oyster embryos at 130 [micro]g [L.sup.-1] water concentrations (L[C.sub.50] = 103 [micro]g [L.sup.-1]) and zinc to be 100% lethal at 500 [micro]g [L.sup.-1] (L[C.sub.50] - 310 [micro]g [L.sup.-1]). In a subsequent study, MacInnes (1980-81) demonstrated a synergistic toxicity for copper and zinc in eastern oyster embryos at concentrations of 8-16 [micro]g Cu [L.sup.-1] and 100-200 [micro]g Zn [L.sup.-1]. For larvae of C. virginica, Calabrese et al. (1977) estimated lethal copper toxicity for a 12-day exposure as L[C.sub.50] = 32.8 [micro]g [L.sup.-1], with growth of surviving larvae reduced by 30%. Prytherch (1934) noted that larval exposure to very high copper (>800 [micro]g [L.sup.-1]) would prove lethal even with very short exposure periods (minutes to hours).

In contrast, there is little evidence of water-borne toxicity to adult oysters. One early study attempted to characterize anticipated toxic effects:

   "In the first place, we tried the effect of pieces of copper;
   copper filings, and copper dust lying in the bottom of the
   aquarium; and similarly, of steel filings, old rusty nails, and
   other fragments of iron. We also kept oysters for some time
   in an old copper vessel, and along with copper pyrites and
   other ores of copper. None of these gave any definite result."
   (Herdman & Boyce 1899, p. 32)

In subsequent experiments however, they exposed eastern oysters to 50 grains of copper oxide in a gallon of water, which nominally resulted in some green coloration and some mortality within a few weeks. Other than this unsubstantial report, there have been no definitive demonstrations of lethality for copper or zinc exposures to adult eastern oysters. This is underscored by O'Connor (2002), who reviewed a large data set compiled by Jarinen and Ankley (1999) on chemical effects to survival, growth, and reproduction of aquatic organisms; there were no data relating lethal effects of copper or zinc to adult eastern oysters. This in triguing lack of evidence led Roesijadi (1996) to conclude that adult eastern oysters are tolerant of high copper and zinc concentrations in the water column and in their tissues.

There is some published evidence of "negative" toxicity test results. In two carefully controlled, continuous-exposure experiments (Shuster & Pringle 1969), there were no lethal consequences to eastern oysters exposed to copper and zinc, even though tissue concentrations were significantly elevated over the 20-week exposure (see Fig. 11). Cadmium (100 and 200 [micro]g [L.sup.-1]) caused significant mortalities, but not copper (25 and 50 [micro]g [L.sup.-1]), zinc (100 and 200 [micro]g [L.sup.-1]), or chromium (50 and 100 [micro]g [L.sup.-1]). Copper accumulated in the oyster tissues to over 5000 [micro]g [g.sup.-1] (dry wt) and zinc to over 15,000 [micro]g [g.sup.-1]. There are, furthermore, numerous examples of extremely high concentrations of copper and zinc accumulated in apparently healthy eastern oysters (see Table 3). Frazier (1975, 1976) found adult C. virginica survived elevated concentrations of copper in experimental studies, but the shells deposited during this time were thinner and weaker. Okazaki (1976) showed copper toxicity to adult C. gigas, but at very high concentrations (96-h median tolerance limit = 560 [micro]g [L.sup.-1]) so this species may also be relatively tolerant.

It is not clear how sequestration in granules prevents copper and zinc toxicity. The membranous lining of the granules may shield vulnerable tissues, or the metals may be complexed within the granules so they are unavailable to vulnerable tissues. As long as additional uptake requires generation of new amebocytes and their protective granules, elevated ambient water concentrations will not lead to oyster toxicity. However, this mechanism relies on the ability of amebocytes to capture incoming (ambient) copper and zinc before contact with vulnerable tissues. Most likely, this phase is achieved through binding of the metals to mucus in the mantle cavity, as proposed below.

Mucus Capture of Ambient Copper and Zinc

Any water-borne or food-borne element entering an oyster first encounters the copious mucus covering all external surfaces of the soft tissues (Fig. 12). Hillman (1968, 1969) investigated cellular and compositional differences of mucus in Mercenaria mercenaria and concluded that mucus was, "playing far more sophisticated roles in the life activity of the clam than previously suspected" (Hillman 1978, p. 21). These roles include organismal protection. lubrication, and food capture. Mucus of bivalves contains complex carbohydrate sulphates that operate as an ion-exchange mechanism to capture and retain particles and elements (Pringle et al. 1968), especially divalent cations such as copper and zinc (Korringa 1952). If toxic metals were bound to mucus, then they would no longer be available for direct absorption into vulnerable tissues. Instead, they would flow with the mucus sheets across tissue surfaces toward the mouth for elimination (psuedofeces) or ingestion (alimentary tract). This is a common path for selection of oyster food particles (Loosanoff 1949, Menzel 1955); eastern oysters are capable of preferentially ingesting organic particles while rejecting inorganic particles with the pseudofeces (Newell & Jordan 1983). If bound to mucus, water borne copper and zinc entering the mantle cavity could be eliminated without ever contacting vulnerable tissues.

[FIGURE 12 OMITTED]

Accumulation, however, could still be achieved through amebocyte phagocytosis of metal-laden mucus and metal-laden food particles trapped in the mucus. Through diapedesis, amebocytes cuter both the mantle cavity and the alimentary tract to capture mucus and food. The process of phagocytosis includes formation of phagosomes, cytoplasmic inclusions (Section V) that exhibit membrane structures similar to copper- and zinc-bearing granules. Metals might be retained in the phagosome after food particles and mucus are digested. If so, phagosomes may be the precursors to cytoplasmic granules. Other possibilities for uptake of metals exist. In particular, vertebrate phagocytes have displayed natural resistance-associated macrophage protein (nramp) activity that transports metal ions for defensive purposes (Alkinson & Barton 1998, Jabado et al. 2000). It is believed that nramp proteins remove divalent cations from phagosomes, thereby depleting phagocytosed microorganisms of essential elements. Accumulation or retention of" metals in oyster amebocyte granules might use a similar. albeit inverted, process.

A Presumptive Biologic Function for Copper and Zinc

Information has been presented to dispel any perception that copper and zinc are sequestered by oysters solely to avoid toxicity. Amebocytes acquire high concentrations of copper and zinc from low ambient concentrations and retain them for relatively long periods despite the potential for elimination by exomigration. Retention of copper and zinc may exceed the life span of an individual amebocyte. The lack of toxicity to eastern oyster adults. even at high water and high tissue concentrations, is most likely attributable to binding of the metals by mucus and sequestration in membrane-lined granules of amebocytes. These findings support speculation that eastern oysters store high amounts of copper and zinc for eventual use in critical physiologic functions. The following section provides evidence that antimicrobial activity is one of those functions.

HEIGHTENED AMEBOCYTE ACTIVITIES AT CONTAMINATED SITES

Any attempt to understand the effects of chemicals on bivalve defense capacity must confront the conceptual obstacles holed at the beginning of Section II; the types and concentrations of chemicals vary in the water column over time and each chemical exhibits a unique rate of assimilation and elimination. Furthermore, variability in the natural environment can influence amebocyte defense responses (Fisher 1988). Oysters are poikilothermic and osmoconforming, so the frequent fluctuations in temperature and salinity of estuarine and coastal waters are reflected in oyster tissues. Temperature and salinity are both known to affect defense-related activities of eastern oyster amebocytes (Fisher & Newell 1986, Fisher & Tamplin 1988), as is the seasonal reproductive cycle (Fisher et al. 1989, Fisher et al. 1996).

Bivalves do not produce antibodies for specific recognition of antigens, but rely on robust, relatively non-specific cellular and humoral mechanisms to heal wounds and ward off microbial invaders. Bivalve amebocytes, also called coelomocytes, hemocytes, leukocytes, phagocytes, and blood-cells, are considered key to internal defense because of their ability to phagocytose, encapsulate, and degrade foreign material, including parasites and pathogens (Takatsuki 1934, Wagge 1955, Bang 1973, Narain 1973, Cheng 1975, 1981, 1984, Fisher 1986, Feng 1988, Chu 1988, Chu 2000). Essential to these activities is the mobility of amebocytes and their capacity to migrate across epithelial barriers (diapedesis). Defense is not the only role of bivalve amebocytes; they are reported to participate in a variety of other important biologic functions, most notably food digestion, excretion, and shell deposition (Yonge 1928, Wagge 1955, Narain 1973, Cheng 1977, Feng et al. 1977). Each of these roles is undoubtedly affected by both natural and anthropogenic stimuli in the environment.

Field Studies Positively Link Chemical Contaminants and Amebocyte Activities

Recent investigations in Florida characterized the prevailing responses of oyster defenses, particularly amebocyte-based defenses, to a spectrum of pollution types and intensities under natural conditions. The approach was intended to provide a "realworld" characterization of multiple factors without regard to independent effects of individual chemicals. Eastern oysters were collected from polluted and unpolluted sites to compare tissue chemical residues and defense activities. Although immunosuppression is a common expectation, the studies revealed a higher level of amebocyte activities for oysters collected at the more polluted sites. In the first study (Fisher et al. 2000), oysters were collected from 16 heavily-, moderately-, and lightly-contaminated sites in Tampa Bay (Long et al. 1991, Long et al. 1994, McCain et al. 1996). Circulating amebocytes of oysters at the more contaminated sites exhibited higher numbers and higher locomotory activity (Table 9). Positive associations with these putative defense characteristics were strongest at sites where oysters had high concentrations of metals, particularly copper and zinc. and polycyclic aromatic hydrocarbons (PAH). Based on a preconception that metals were sequestered solely for detoxification and elimination, it was suggested that the presence of high ambient metal concentrations triggered an elevated amebocyte response to protect the organism. Enhancement of defense activities was believed incidental to detoxification (Fisher et al. 2000).

A subsequent survey (Oliver et al. 2001) explored the geographic extent of this association by examining oysters collected from 22 locations across 5 Florida bays (St. Andrew, Choctawhatchee, Pensacola, Tampa, and Biscayne Bays, Fig. 13). Chemical concentrations and defense factors varied across bays and among sites within a bay. Within-bay comparisons reiterated the finding that oysters inhabiting contaminated sites had higher numbers of circulating amebocytes, higher percentages of mobile amebocytes, and higher rates of locomotion. These characteristics were positively associated with site-averaged concentrations of metals, particularly copper, tin, and zinc (Table 10), PAHs, and even with certain polychlorinated biphenyls (PCB). Data analyzed across all five bays also revealed a strong and consistently positive association between tissue chemical concentrations and amebocyte measurements. These results, corroborating those of the first study, led to a projection that the prevailing effect of environmental mixtures of chemicals on defenses of Florida oysters was enhancement. rather than suppression, of amebocyte numbers and defense capabilities.

[FIGURE 13 OMITTED]

Technical Constraints and Resolutions

Understanding the prevailing effect of enhanced defenses at contaminated sites was limited by two technical constraints. First, measurements of different amebocyte activities in both studies were performed as indicators of defense capacity ("putative" defense characteristics), rather than direct measures. Although it is easily accepted that more, and more highly active, amebocytes should augment the defense response, clear evidence of elevated microbicidal activity would have been profoundly more convincing. Accordingly, a vertebrate procedure to estimate microbial killing by amebocytes in vitro was adapted (Volety et al. 1999). Vibrio parahaemolyticus, common bacteria in oysters and coastal environments, were incubated with constant numbers of circulating amebocytes. After a challenge period, numbers of surviving bacteria were estimated by a colorimetric assay based on their reduction of tetrazolium dye. This technique, applied in this and other studies (Genthner et al. 1999, Volety & Fisher 2000), provided a direct and more definitive assessment of eastern oyster defense capacity.

The second technical constraint involved the decision to analyze chemical concentrations from a single composite of 20 oysters collected from each site, an experimental compromise necessitated by the need for sufficient tissue to assay a broad array of chemicals. This decision negated the statistical correlation of specific analytes with specific amebocyte characteristics, which were measured independently for each oyster. Chemical concentrations in oysters at the same site are known to be highly variable (see Section 11), and a single composite value could not adequately reflect concentrations of an individual.

Deployment Studies in Pensacola Bay Area

To overcome these two technical constraints, a subsequent study measured metals from individual organisms and applied the amebocyte killing assay as a direct estimate of amebocyte defense activity (Oliver et al. 2003). Only two sites in Pensacola Bay were sampled, but they differed markedly in type and magnitude of chemicals present. Chemical analyses were confined to metals and butyltins from individual oysters at each site, which allowed statistical correlation with amebocyte measures from the same oyster. Additional chemicals were analyzed as previously, from a composite. Oysters from one site (Bayou Chico) bore significantly higher concentrations of Cu, Mn, Sn, Zn, butyltins, PAH, and PCB and displayed significantly higher circulating amebocyte numbers and bactericidal activity. These oysters exhibited significantly lower concentrations of Al, Cr, Fe, Ag, Cd, and Hg than oysters from the East Bay site. Statistically-significant correlations were found between defense measurements and specific analytes (Table 11). Corroborating the previous findings, circulating amebocyte numbers and bactericidal activity were positively correlated with Cu, Sn, Zn, total metals, tributyltin, and total PAH.

Although an association between chemicals and defense activities was reasonably established, it was still unknown whether elevated defenses were induced through short-term chemical exposure (acclimation), were selected through survival of defensively-active oysters at chronically contaminated sites (adaptation), or both. An additional study was performed to determine whether short-term exposures would alter amebocyte activity (Fisher et al, 2003). Hatchery-reared oysters were deployed for 12 weeks in the summer at 3 sites in Pensacola Bay, Florida. The sites (Bayou Chico, Bayou Texar, and Santa Rosa Sound, respectively) were heavily-, moderately- and lightly-influenced by anthropogenic discharges. Tissue concentrations of Cu and Zn (Fig. 14) as well as Cr, butyltins, and PAH increased dramatically at Bayou Chico during deployment. Concomitantly, amebocyte number and bactericidal activity were significantly elevated at Bayou Chico. These results, while not excluding adaptation, demonstrated that amebocyte enhancement could be an acclimation response. Positive associations of chemicals with amebocyte number and activity corroborated the earlier studies, but results from a 16-week spring deployment were equivocal, implying that the stimulus, response, or capacity to detect a response was seasonally-dependent or influenced by other environmental factors.

[FIGURE 14 OMITTED]

Results from these field studies consistently demonstrated that environmental contaminants stimulated, rather than suppressed, amebocyte defense activities. The results portrayed cumulative effects of multiple chemicals under natural environmental conditions, not the effects of individual chemicals that might independently suppress or stimulate amebocytes. The studies did not examine, despite the obvious implication, any mechanistic linkage to oyster disease resistance. However, they established a consistent and defensible relationship between amebocyte numbers and defense activities with tissue concentrations of copper and zinc. These were not the only two analytes associated with elevated defenses but, because of their dominating concentrations in oysters and their physical containment within amebocytes, a possible cause-effect relationship became worthy of consideration.

ANTIMICROBIAL FUNCTIONS OF COPPER AND ZINC

It was reported in Section III that accumulations of copper and zinc in oysters were so extraordinarily high that many investigators believed they exceeded useful amounts (Hiltner & Wichmann 1919, Bodansky 1920, Korringa 1952, Pequegnat et al. 1969, Wolfe 1970a, Simkiss & Mason 1983).

   "It seems probable that zinc, as well as cooper, can be
   absorbed and retained in the tissues of the oysters in quantities
   far in excess of functional requirements, especially in
   oysters grown in waters badly polluted with metallurgical
   and factory wastes." (Hiltner & Wichmann 1919, p. 221)

Functions that might be served (e.g., respiratory pigment) (Mendel & Bradley 1905, Prytherch 1934) or enzyme catalysts (Pequegnat et al. 1969), simply did not require the high concentrations accumulated in tissues. Ruddell and Rains (1975) hypothesized that the metals might be used in numerous oyster functions, including hydration of C[O.sub.2], maintenance of extracellular and intracellular pH, production or entrapment of tree radicals, regulation of redox potentials, or traps for small molecular weight compounds with affinities for zinc or copper. Yet, these roles would be difficult to perform for metals sequestered within amebocyte granules and, even if true, all these functions combined would probably not require the concentrations that can be accumulated. This review rekindles a premise of functional roles for high accumulations of copper and zinc. Field studies described in Section IV demonstrated greater amebocyte numbers, mobility and bactericidal activity in eastern oysters inhabiting sites contaminated with high levels of copper and zinc. In particular, elevation of bactericidal activity implicated a true physiologic function for copper and zinc in oyster defense. Such a role, in fact, had already been proposed, as described below.

Copper and Zinc in Wound Healing

Ruddell (1971) confirmed the observations of Pauley and Sparks (1965) and DesVoigne and Sparks (1968) that green coloration often accompanied the healing of wounds in C. gigas. This coloration Ruddell interpreted as an inflammatory response involving copper-laden amebocytes, an association that had been well established (Section I). In his experiments, Ruddell provoked an inflammatory response by wounding the mantle tissues or implanting excised oyster tissue into an intact mantle cavity. Tracking the tissue changes histologically, he found that copper did, indeed, appear at the site of the wound, first bound within the granules of acidophilic granular amebocytes (AGA), but later in extracellular spaces bound to surfaces of muscle, nerve and epithelial tissues near the wound. Copper was subsequently found in granules of basophilic granular amebocytes (BGA) at the wound site, but not in BGAs distant from the wound site. These observations he interpreted to mean that copper, which was accumulated primarily in AGAs, was released at the site of a wound, distributed to tissues near the wound area, and then recovered into granules of BGAs. From these observations, Ruddell (1971) proposed that copper served as an antimicrobial agent:

   "Although it is apparent that copper plays an essential role
   in the oyster inflammatory response, the function of the
   copper in the response is not known. One can presume that
   copper might function to ward off, destroy, or inhibit the
   growth of potential oyster pathogens." (Ruddell 1971, p.
   110)

Ruddell (1971) also hypothesized a similar role for zinc. His evidence indicated that BGAs recruited to the site of a wound underwent swelling and released zinc from their granules. He also noted the release of other constituents, in particular diazo-positive material that ]re believed was a phenolic substance.

Using both C. gigas and C. virginica, Ruddell and Rains (1975) formulated a functional role for high accumulations of copper and zinc. Although they understood that zinc- and copper-rich amebocytes could be used in diverse ways, they believed that a primary role was in response to trauma,

   "It would seem probable that oysters do use their basophils
   and hence, by extension, the large amounts of zinc and
   copper incorporated in the basophils. Therefore, although
   oysters may contain very large quantities of zinc und copper,
   these metals must not be regarded as being stored away
   in u physiologically inactive form." (Ruddell & Rains, p.
   590)

Their studies provided compelling rationale for extracellular killing by copper and zinc, serving in their well-known capacity as "antimetabolites for a diverse number of animal, plant and microbial forms." Direct evidence of microbial killing is lacking, however, even though concentrations of copper and zinc in amebocytes (see Table 6) seem more than adequate to effect toxic action.

Extracellular Clot Formation

Ruddell's demonstration of extracellular release of copper and zinc led to later research on the formation of extracellular clots. Most organisms have some mechanism to form clots in the hemolymph or blood; clotting is a sequence of complex chemical and physical reactions that results in conversion of fluid (hemolymph or blood plasma) into a coagulum. Clotting in invertebrates, as described by Gregoire (1970), involves the exudation or eruption of blood cells to release granular material into the surrounding fluid. Circular clouds of a granular consistency form around the blood cells, and they eventually develop into "islands of coagulation" formed by networks of granular fibrils. In most early studies, lamellibranch hemolymph was found to lack extracellular clotting. Takatsuki (1934) noted clumping of amebocytes withdrawn from O. edulis, but he did not observe any extracellular clot formation.

   "The amoebocytes of the oyster were allowed to stand after
   being drawn from the body: the mass of amoebocytes did
   not become jelly-like by coagulation as in the case of crustacean
   and vertebrate blood, and no fibrin appears in the
   blood-plasma which remains fluid on standing in the air
   and even after being heated." (Takatsuki 1934, p. 402)

Eastern oysters became the first, and may well be the only, bivalve known to exhibit such a response (Narain 1973). Frederick B. Bang (1961) described clotting in eastern oysters as an extracellular, finely-granular "gel" that occurred around aggregates (clumps) of amebocytes (Fig. 15). Clots were observable on glass slides under phase contrast microscopy within 30 min of hemolymph withdrawal, and they resembled clots formed naturally in the hemolymph vessels of traumatized oysters. He noted that these clots could trap and immobilize bacteria, presumably making the bacteria more susceptible to phagocytosis.

   "It seems to have a real role in the repair of traumatized
   tissue, for: 1) it was found already' developing in small
   cellular clots taken directly from the heart; 2) it occurred
   predominantly around clumps of cells; 3) bacteria were
   immobilized by its development ... and, 4) it was obtained
   both immediately after opening an oyster and from some
   preparations which had been on the half shell as long as 24
   h. Its possible relation to cycles of feeding by the, amebocytes
   is unknown." (Bang 1961, p. 61)

[FIGURE 15 OMITTED]

Copper and Zinc Initiate Extracellular Clot Formation

Bang (1961) felt that the extracellular gel in eastern oysters may have originated from extrusions of the cellular granules of the amebocytes (degranulation), a process previously described for arthropods (Gregoire 1970). It was not until a decade later that one of Bang's doctoral students, Robert S. Brown, followed up on this hypothesis. He noted that extracellular clots would sometimes form when whole hemolymph or plasma was added to glass slides treated with alconox detergent (R. S. Brown, personal communication). Subsequently, he found that other detergents and a variety of other substances, including bases, alcohols, copper, and zinc, would also cause extracellular clotting (Fig. 16). Recognizing that copper and zinc were used in biochemical separation methods to precipitate proteins, Brown combined the work of Bang (1961) and Ruddell (1971) to forge his dissertation hypothesis that copper and zinc released from amebocyte granules precipitated hemolymph plasma proteins to form extracellular clots (Brown 1975). To test his hypothesis, Brown investigated 3 basic suppositions; (1) that oyster amebocytes were the source of the clot-promoting substance. (2) that copper and zinc were released from amebocytes into the plasma, and (3) that copper and zinc produced clots in plasma by precipitating proteins.

[FIGURE 16 OMITTED]

Using a variety of microscopic and analytical tools, Brown (1975) was able to support each of the suppositions. He reiterated Bang's finding that extracellular clots were always associated with amebocyte aggregates (Fig. 17) and would not form in hemolymph if amebocytes were removed. Clots would form if amebocytes were added back to cell-free hemolymph, even if the amebocytes had been fixed with glutaraldehyde, or were frozen, boiled and extracted before reintroduction. He verified for C. virginica the earlier evidence (Ruddell 1971) that C. gigas amebocytes sequestered copper and zinc (see Figs. 6 and 7) and that they were retained in amebocyte granules (see Figs. 8 and 9). He used a combination of techniques to verify that high concentrations of copper and zinc were excluded from agranular regions of the amebocytes (See Fig. 10).

[FIGURE 17 OMITTED]

To provide some evidence that amebocytes could release copper and zinc from the granules, Brown (1975) presented electron microscopic evidence that independent amebocytes were highly granulated, whereas those amebocytes in aggregates were either completely degranulated, in the process of degranulation, or in some cases, undergoing cytolysis (Fig. 18). He believed degranulation provided a mechanism for release of copper and zinc into the hemolymph. He also revealed that extracellular clots had a relatively high staining affinity (Fig. 19) and high concentrations of copper (356 [micro]g [g.sup.-1], dry weight) and zinc (10,571 [micro]g [g.sup.-1]); these were substantial amounts, particularly in comparison to the minute quantities found in clot-free plasma (copper < 0.2 [micro]g [mL.sup.-1] and zinc < 10 [micro]g [mL.sup.-1]).

[FIGURES 18-19 OMITTED]

Finally, Brown (1975) demonstrated that addition of either Cuacetate or Zn-acetate to cell-free oyster plasma created precipitates that were indistinguishable from in vitro clotting (Fig. 20). Precipitation required a minimum of either 0.6 [micro]g Cu [mL.sup.-1] or 0.15 [micro]g Zn [mL.sup.-1] hemolymph and, as expected, precipitation dramatically reduced the concentration of plasma proteins. When plasma was first deproteinized with trichloroacetic acid, no amount of copper or zinc would cause precipitation. Similarly, if copper were added to zinc-treated plasma, no further precipitation resulted. Boiling of the plasma did not influence the ability of copper or zinc to effect precipitation, leading to Brown's conviction that extracellular clot formation was a non-enzymatic chemical action. He found that ethylenediaminetetra-acetic acid (EDTA) and hydrochloric acid (HCl), both known to disperse metal-protein precipitates, also dispersed extracellular clots.

[FIGURE 20 OMITTED]

In all, Brown's (1975) data indicated that copper and zinc from amebocytes could interact with proteins in the hemolymph via a non-enzymatic, inorganic reaction to form locally-restricted precipitations, or clots, consisting of calcium and magnesium hydroxide, phosphates and carbonates. There may be other constituents released simultaneously by the amebocytes, such as the phenols described by Ruddell (1971), that may also play a role in clot formation. Phenoloxidase is a constituent of the prophenoloxidase clotting system in crustaceans (Soderhall 1982, Soderhall et al. 1994) that is believed to provide melanin for encapsulation of invading microorganisms. Although not linked to a clotting mechanism, phenoloxidase activity has also been reported in bivalves (Coles & Pipe 1994, Asokan et al. 1997, Carballal et al. 1997, Deaton & Dankert 1998, Peters & Raftos 2002).

Brown (1975) did not analyze the form of copper and zinc in the granules of amebocytes or in the extracellular clots. The work of Coombs (1972) led George et al. (1978) to conclude that granules contained copper sulfide and zinc phosphate. Metal speciation may be important because it could control the pH of material released from the amebocyte granules. Either high or low pH could contribute to release of metals from the granules and protein precipitation in the hemolymph. Beaven and Paynter (1999) have shown that phagosomes (see below) become highly acidified after fusion with lysosomal granules.

Intracellular Microbicidal Potential of Copper and Zinc

Ruddell & Rains (1975) confined their speculations to the extracellular microbicidal capacity of copper and zinc in wound healing. However, the possibility that these metals participate in intracellular killing by amebocytes is equally compelling. Metchnikoff (1891) brought attention to evidence that vertebrate and invertebrate amebocytes possess phagocytic ability. A complex process, phagocytosis requires that amebocytes recognize, locate, ingest, and either transport or digest foreign particles. In eastern oysters, as in other bivalves, phagocytosis is considered critical to both nutrition and internal defense (Cheng 1977, Feng et al. 1977). To provide nutrient, amebocytes migrate into the digestive tract to engulf particles of food, digest them, and then migrate into the tissues and hemolymph to release nutrient (usually as glycogen) to metabolizing cells. Yonge (1937, 1946), among others, believed this to be the primary nutritional pathway for oysters. Using a similar process, amebocytes can capture unwanted particles and microorganisms (parasites and pathogens) in the mucus and hemolymph to prevent microbial growth in the soft tissues. If the phagocytosed material has no nutritive value, it can be eliminated through feces or psuedofeces by amebocyte exomigration (see Section II). It has been postulated that phagocytosis was originally a nutrient-acquiring process that eventually evolved into a defensive capability (Cheng 1975, Feng et al. 1977).

Professor Leslie A. Stauber (1950) initiated the first of many studies that characterized phagocytosis in marine bivalves when he traced the fate and disposition of injected india ink particles in C. virginica. Three investigators, Sung Y. Feng, Marennes R. Tripp and Thomas C. Cheng, followed Stauber's initiative with innumerable experiments and cogent descriptions of bivalve internal defense, often using eastern oysters as a model system (see Cheng 1983). Iterative updates on their progress were provided by Cheng (e.g., Cheng 1967. 1975, 1981, 1984, 1996). Combined, these studies have detailed the fates of a diverse array of foreign particles and microorganisms in oysters and characterized the process of phagocytosis (e.g., Stauber 1950, Stauber 1961, S. Y. Feng 1958, 1966, 1967, 1988, Tripp 1958. 1960, 1970, J. S. Feng 1966, Tripp & Kent 1967, Acton et al. 1969, Fries & Tripp 1970, Feng et al. 1971, 1977, Cheng & Cali 1974, Foley & Cheng 1975, 1977, Cheng & Rudo 1976, Renwrantz et al. 1979, Hinsch & Hunte 1990). Most living microorganisms are killed upon ingestion by phagocytic amebocytes, including viral particles (J. S. Feng 1966, Fries & Tripp 1970) bacteria, phytoplankton (e.g., Navicula ostrearia), and metazoans. But some microorganisms are able to survive and even multiply within the phagosomes. Of particular interest for eastern oysters is the apparent ability of Perkinsus marinus, a devastating oyster pathogen, to grow and multiply within host amebocytes (Mackin 1951).

Ingestion, or endocytosis, of microorganisms by eastern oyster amebocytes reportedly can occur in three ways; some microbes adhere to the amebocyte filopodia (Bang 1961), some are taken into funnel shaped psuedopods (Renwrantz et al. 1979), and some are enveloped in a vesicle on the amebocyte surface (Cheng 1975). Regardless of the path of entry, the targeted microorganism is captured and drawn into a cytoplasmic invagination (Fig. 21) that encircles it with concentric lamellae to form a "phagosome" (Cheng 1975). Digestion of the microbe begins in this primary phagosome but, at least for eastern oysters, can be transferred to, or evolve into a secondary phagosome which has an electron-dense outer surface comprised of a thick outer wall. Lysosomes, which are membrane lined organelles containing acid hydrolases, have been observed to fuse with secondary phagosomes (Cheng 1975), and may fuse with primary phagosomes as well (Cheng 1996). Fusion of lysosomes with phagosomes is believed to expose the microorganism to degrading lysosomal enzymes. The fused organelle is sometimes called a "phagolysosome". Fusion with a lysosome results in relatively rapid acidification of the phagosome (Beaven & Paynter 1999).

[FIGURE 21 OMITTED]

Both granular and agranular (hyaline) amebocytes have been found capable of phagocytosis, but granular forms are recognized to be much more active. Foley and Cheng (1975) found over 80% of the granular amebocytes in C. virginica to phagocytose Staphylococcus aureus and Escherichia coli compared with less than 20% of hyaline amebocytes. With the exception of Ruddell (1971), who may not have included degranulated cells (fibrocytes) in his estimates (see Cheng 1981), it is generally maintained that granular amebocytes are more abundant and more actively phagocytic in oysters than agranular amebocytes. This may be because granules play a critical role in the phagocytic process. In fact, granularity and phagocytic activity may be tightly linked; Beaven and Paynter (1999) discovered that the number of visible granules in eastern oyster amebocytes declined as phagocytosis proceeded. It could be inferred that highly granular amebocytes are avidly phagocytic, but lose their potential as more granules fuse with phagosomes.

The amebocyte granules that contain copper mad zinc are morphologically indistinguishable from lysosomes, and there is little reason to believe that they are not the same. Ruddell (1971) called the copper and zinc granules 'lysosomal derivatives", but did not elaborate on similarities or differences. From the results of Brown (1975), it seems that virtually every granule in the eastern oyster amebocyte contains some copper or zinc (see Figures 6-9). There has been some suggestion that granules and lysosomes in eastern oysters differ morphologically (Feng et al. 1971, Cheng 1975), but distinctions between the two are vague. Yoshino and Cheng (1976) demonstrated through cytochemistry and electron microscopy that the granules in amebocytes of Mercenaria mercenaria were lysosomes, serving as storage organelles for hydrolytic acids. Foley and Cheng (1977) showed that lysosomes can be stimulated to move to the surface of the amebocyte and expel their contents (Fig. 22), a process that resembles, or is identical to the degranulation process described by both Ruddell (1971) and Brown (1975) for metal-containing amebocyte granules. It can be postulated from these observations that the copper- and zinc-bearing granules in eastern oysters are either lysosomes or closely-related structures.

[FIGURE 22 OMITTED]

It is possible to hypothesize a role for copper and zinc in intracellular microbicidal activity. When phagosomes fuse with cytoplasmic granules, the captured microorganisms are exposed to very high, and likely toxic, concentrations of metals and hydrolytic enzymes. Unlike the case for extracellular killing, metals used in intracellular killing remain inside the cytoplasm. If the phagosome breaks down, as has been described for the release of glycogen granules in nutritive digestion, the metals could be recovered by newly-formed granules.

Extracellular Toxicity to Encapsulated Microorganisms

A similar extrapolation might he drawn for the role of copper and zinc in amebocyte action against larger microorganisms (i.e., parasites that have been encapsulated). Encapsulation is the simultaneous envelopment of relatively large foreign bodies by numerous amebocytes (Fig. 23). The foreign body is usually too large for a single amebocyte to ingest; nonetheless, encapsulation may be the consequence of independent attempts by many amebocytes to phagocytose the invader (Cheng & Rifkin 1970). The cells become flattened against the surface of the foreign body and ultimately cut it off from host tissues.

[FIGURE 23 OMITTED]

It is not unlikely, assuming that encapsulation is an attempt at phagocytosis, that encapsulating amebocytes degranulate while encompassing the intruder. As noted by Cheng (1981),

   "... external contact with certain foreign substances will
   result in hypersynthesis of intracellular lysosomal enzymes
   which are released from haemocytes into the serum where
   digestion of the foreign material, such as bacteria, is initiated.
   The release of enzymes is effected by what has been
   termed degranulation (Foley and Cheng, 1977), a process
   involving the migration of lysosomes to the surface of the
   cell where the enclosed enzymes are discharged." (Cheng
   1981. p. 286)

Cheng et al. (1975) demonstrated the extracellular release of lysozyme from lysosomes during phagocytosis by Mercenaria mercenaria amebocytes. Similarly, McDade and Tripp (1967) demonstrated the presence of lysozyme in hemolymph of eastern oysters. Extracellular release of copper and zinc at the site of a wound (Ruddell 1971) is presumably a consequence of amebocyte degranulation, as is copper-plating of a "well polished steel knife" (Galtsoff 1964, p. 388). If metals and hydrolytic enzymes are expelled onto the surface of the invader, then the process of encapsulation attains a dual objective of both isolating and debilitating the invader.

Rationale for High Accumulations of Copper and Zinc

Evidence has been presented that copper and zinc serve antimicrobial functions in eastern oysters, yet it is not immediately obvious why such high accumulations are needed. Brown (1975), for example, estimated that the amounts of copper and zinc contained in eastern oyster amebocytes was much greater than thresholds necessary to form extracellular clots. Confounding the issue is the fact that living, healthy oysters exhibit widely varying tissue concentrations. The best explanation may simply be that higher concentrations have the capacity to generate a larger clot, or marshal stronger microbicidal action against phytoplankton, bacteria, and viruses. This increased capacity provides a competitive advantage to oysters with higher accumulated concentrations. Perhaps equally important, high accumulations may afford oysters greater flexibility in realizing this advantage. Amebocytes are highly mobile and can move across epithelial barriers to virtually any compartment of the oyster (diapedesis). They may, however, be limited in their ability to reach a wound quickly (or an algal cell needed for nutrition) and their movement may leave other areas of the oyster unprotected (or unexploited), High accumulations of copper and zinc, combined with the requisite high numbers and high activity of amebocytes, provide the oyster a robust, distributed antimicrobial capacity that allows quick responses to frequent or repeated challenges and opportunities throughout the organism.

HYPOTHESES AND RAMIFICATIONS OF AN ALTERNATIVE FRAMEWORK

Numerous independent investigations on metals in eastern oysters have been reviewed to establish that copper and zinc, if not other terrestrial elements, are actively captured and retained for physiologic purposes. This contradicts a historical perception that high accumulations are merely part of a process to detoxify and eliminate the metals. Copper and zinc are physically associated with amebocytes, so any physiologic role is likely related to amebocyte functions. Evidence presented here links copper and zinc to antimicrobial functions of amebocytes in defense and nutrition. A separate report (Fisher 2004) presents evidence of a role for terrestrial metals in shell deposition, which is facilitated by amebocytes. The findings contradict a tendency to regard copper, zinc, and other metals as unnecessary, if not undesirable, oyster constituents. Introduced through this review is a corollary proposition that oyster success and distribution in near-coastal areas, even at seemingly polluted locations, stems from a dependency on terrestrial elements. These proposals arise from the cumulative interpretations of many scientific contributions, and generate an alternative framework for understanding issues that range from mechanisms of amebocyte function to coastal distribution of eastern oyster populations. There are numerous considerations that bear on the legitimacy of this alternative framework, some supported more than others by existing evidence, but all deserving at least some corroborative research. These considerations are listed below in a series of broad, umbrella hypotheses (H1-H6).

Eastern Oyster Adults Actively Accumulate Terrestrial Copper and Zinc

(H1) The most consistent information amassed over several years and numerous studies is that accumulation of copper and zinc is extremely high in eastern oysters, a status supported by their ability to concentrate the metals from low ambient concentrations and retain them longer than other metals despite availability of an efficient elimination mechanism. Long retention may mean that copper and zinc are transferred from old to young amebocytes by phagocytosis or apoptosis. Active assimilation and accumulation of the metals signifies a cost to oyster energetics. Accumulations of copper and zinc, rare in marine environments, represent quantities acquired from terrestrial watersheds. This factor may limit oyster distribution to estuarine and near-coastal locations.

Copper and Zinc are Sequestered in Membrane-lined Granules of Amebocytes

(H2) There can be little doubt that copper and zinc are retained primarily, if not exclusively, in oyster amebocytes. Metals have been reported in amebocytes from microscopic observations (e.g., green oysters), various histologic preparations, and electron microscopic imaging techniques. Storage in amebocytes provides a transport mechanism to quickly distribute the metals to virtually any site in the organism. Copper and zinc are sequestered in membrane-lined granules in the amebocyte cytoplasm at high concentrations. Sequestration shields vulnerable tissues from exposure and is the primary reason that eastern oysters can safely accumulate high tissue concentrations. Acquisition of metals by the granules probably occurs by phagocytosis of metal-laden mucus (see later) or food particles (e.g., phytoplankton and bacteria). The chemical speciation of metals in the granules allows their discharge and bioavailability upon degranulation or fusion with phagosomes.

Ambient Dissolved Copper and Zinc are Bound in Mantle-cavity Mucus

(H3) Oysters assimilate copper and zinc from ambient water or food into protective amebocyte granules without prolonged or excessive exposure to unprotected tissues. Chemical properties of mucus ensure strong binding of any cations entering the oyster mantle cavity and prevent their absorption into tissues. Binding with mucus also provides an efficient elimination mechanism for unwanted metals; mucus accumulates in the psuedofeces, which is passed out of the body. Uptake of metals, on the other hand, may be achieved in a controlled manner by amebocyte phagocytosis of mucus and particles bound in mucus. Phagocytic activity captures copper and zinc in membrane-lined phagosomes, vesicles formed by invaginations of the cell membrane. These vesicles may ultimately form metal-containing granules. Embryo and larval stages, without the copious mucus production of adults, are vulnerable to water-borne copper and zinc toxicity.

Copper and Zinc Accumulation Depends on Availability of Amebocytes

(H4) Amebocytes are highly phagocytic and can migrate throughout the alimentary tract and mantle cavity to acquire copper and zinc from food or mucus. Safe accumulation of the metals, however, depends on the availability of amebocytes with metal-carrying capacity. Amounts of metal that can be stored in an amebocyte are presumably constrained by its size, its capacity to form and retain granules, and the metal-carrying capacity of the granules. Whenever existing amebocytes are saturated, additional up take from the environment requires additional amebocytes. Uptake by eastern oysters is thus dependent on the rate of amebocyte proliferation or recruitment. Specific amebocyte cell types may be recruited based on the metal present (i.e., BGA for copper and AGA for zinc). Higher numbers of amebocytes increase the oysters capacity to accumulate metals.

Degranulation Releases Copper and Zinc from Amebocyte Granules

(H5) In a process diametric to endocytosis, lysosomal membranes fuse with the outer cell membrane to discharge lysozyme into extracellular spaces (degranulation). Lysosomal granules also fuse with cytoplasmic phagosomes to discharge lysozyme into phagolysosomes for killing or digesting trapped microorganisms. Amebocyte granules that contain high concentrations of copper and zinc are similar, if not identical, to lysosomal granules. Copper and zinc, which may occur in all granules, are discharged simultaneously with lysosomal enzymes into extracellular spaces or phagolysosomes.

Copper and Zinc Provide Extracellular and Intracellular Antimicrobial Activity

(H6) Antimicrobial activities of copper and zinc include clotting of hemolymph and both intracellular and extracellular killing of microorganisms. For intracellular killing, granules fuse with phagosomes to discharge hydrolytic enzymes and toxic concentrations of copper and zinc, which may act in concert. Toxic metals released intracellularly are excluded from vulnerable host tissues and simultaneously retained for future use. For extracellular killing, a burst of either metal during degranulation will affect invading microbes without severe damage to surrounding tissues. The high mobility and flexible morphology of amebocytes allows degranulation to occur with relative precision at wound sites, possibly even directed at individual microorganisms. Metals are recouped to protect nearby tissues from prolonged exposure and for re-use. Extracellular discharge of copper and zinc might also pro vide antimicrobial activity against encapsulated microorganisms. Copper and zinc released from granules into the hemolymph precipitate proteins and generate a clot. Clotting provides a granular net to capture and immobilize bacteria and stem the flow of plasma (hemostasis). Amebocyte clumping, or aggregation, is also a factor in hemostasis and occurs simultaneously with extracellular clot formation.

Potential Ramifications of an Alternative Framework

Anticipating that some of the considerations above are ultimately justified by direct evidence, it is worth considering potential ramifications. The most consequential aspect is the influence of copper and zinc on amebocyte antimicrobial activity. Antimicrobial activity of amebocytes in defense and nutritional functions has been accepted for eastern oysters, if not most bivalves, for many years (Tripp 1960, Stauber 1961, Feng 1967, 1988, Narain 1973, Anderson 1975, Cheng 1975, 1981, 1996, Fisher 1986). Some early investigators, particularly Yonge (1926, 1937, 1946), believed that amebocytes were largely responsible for digestion of foods entering the alimentary tract. Although this emphasis has been disputed, amebocyte participation in nutrition is widely accepted (Takatsuki 1934, Wagge 1955, Owen 1966, Purchon 1968, Narain 1973, Cheng 1975, 1977, Feng et al. 1977).

One of the greatest concerns for fisheries managers over the last 50 years has been diseases caused by 2 protozoans, Perkinsus marinus and Haplosporidium nelsoni. Host defenses against these two diseases are not well characterized, but it seems that both agents may be susceptible, at least under some conditions, to phagocytosis by oyster amebocytes (Burreson et al. 1988, Ford & Kanaley 1988, Sero & Ford 1990, Anderson 1996, Volety & Fisher 2000). Both diseases are severe at high salinity locations (Andrews 1988, Andrews & Ray 1988, Ford & Haskin 1988, Haskin & Andrews 1988, Ford 1996, Soniat 1996) and disease intensity can be reduced with low salinity (Sprague et al. 1969, Ford 1985, Ford & Haskin 1988, Chu & Greene 1989, Chu et al. 1993, La Peyre et al. 2003). Hence, the potential enhancement of defenses by copper, zinc, or other terrestrial elements provided by freshwater inflow could be relevant to control of disease.

Also of great concern to the oyster fishery is contamination of commercial oysters with human pathogens such as Vibrio parahaemolyticus, V. vulnificus, and V. cholerae (Kelly & Arcisz 1954, Blake et al. 1979, 1980, Pavia et al. 1987, Bernard 1989, Daniels et al. 1999). Elimination of bacteria from oyster tissues (depuration) is at least partly achieved through phagocytic action of amebocytes (Rodrick & Ulrich 1984). The ability of oysters to successfully eliminate specific microorganisms partly depends on the species; Vibrio vulnificus, for example, is much more persistent in oysters than fecal pollutants such as Escherichia coli and Salmonella (Perkins et al. 1980, Richards 1988, Jones et al. 1991, Tamplin & Capers 1992). Such persistence may depend on whether the bacteria have light or heavy mucopolysaccharide coats that mask the bacteria from amebocyte recognition (Harris-Young et al. 1993, 1995, Genthner et al. 1999, Volety et al. 2001). The significantly higher in vitro capacity of amebocytes of oysters from copper- and zinc-contaminated sites to kill V. parahaemolyticus (Fisher el al. 2003, Oliver et al. 2003) supports the seemingly ironic speculation that elevated accumulations of copper and zinc in oyster amebocytes will reduce human health risks.

Studies of chemical influences on oyster defense processes have often demonstrated variable results. Perplexing is the fact that eastern oysters are known to thrive at some highly-polluted sites (Abbe & Sanders 1986). A few studies have linked chemical exposure of eastern oysters with higher prevalence or intensity of infectious disease (Winstead & Couch 1988, Chu & Hale 1994, Anderson et al. 1996, 1998, Fisher et al. 1999), but have never linked vulnerability to any particular defense mechanism (Oliver & Fisher 1999). Some studies have examined effects of individual chemicals on defense mechanisms of amebocytes (Cheng & Sullivan 1984. Cheng 1988a, Cheng 1988b, Larson et al. 1989, Fisher et al. 1990, Alvarez et al. 1991, Alvarez & Friedl 1992, Anderson et al. 1992, 1994, 1997, 1998, Sami et al. 1992, 1993, Baier-Anderson & Anderson 1997) and found both suppression and stimulation of defense responses (Baier-Anderson & Anderson 2000). Several studies have exposed amebocytes to chemicals in vitro, providing direct contact that may never occur it, under natural conditions, the chemicals are first bound to mucus. Studies of potential immunomodulation are complicated by the fact that chemical mixtures vary from location-to-location. Because each different chemical can elicit a different type and degree of response, it has been almost impossible to anticipate which response would predominate. The field surveys described in Section IV demonstrated enhanced activity at sites with high copper and zinc concentrations, apparently overwhelming effects of any other chemicals. The results provide a novel focus, tissue concentrations of copper and zinc, for comparing defense activities in eastern oysters.

The close association of copper and zinc accumulations with hemocyte number leads to a speculation that their availability in the ambient environment may lead to proliferation of amebocytes. Whereas highly conjectural, the significance of this would be extraordinary. Metal stimulation of amebocyte proliferation might parallel vertebrate immune responses that rely on antigen recognition and binding for lymphocyte proliferation. There is a possibility that increased exposure to only one metal might differentially stimulate recruitment of different amebocyte types (AGA for copper and BGA for zinc), just as antibody-specific cells are generated by vertebrate systems. Vertebrate studies have demonstrated numerous adverse effects of copper deficiency, including reduced bactericidal activity of neutrophils, decreased macrophage activity, reduced proliferation and activity of B and T lymphocytes, and impaired cell-mediated immunity (see Kramer & Johnson 1992 for review). In fact, deficiencies in copper, zinc, and iron are all recognized to alter immunocompetence in humans and other vertebrates (Sherman 1992).

Currently, we know very little of amebocyte proliferation in bivalves (Narain 1973). Cuenot (1891) suggested that amebocytes might originate from special lymph glands at the base of the gills, and subsequent investigators have suggested alternate sites. Wagge (1955) and others believed that new cells were mitotically generated from existing amebocytes. Stauber (1950) disagreed because "of the many phagocytes examined, only one has been observed in mitosis" (p. 233). Ruddell (1971) also found only a few granular amebocytes undergoing mitosis in histologic sections. However, Feng et al. (1977), in their studies of C. gigas, encountered binucleated granular amebocytes and amebocytes with centrioles "fairly frequently" (p. 61). They felt that Ruddell's (1971) observations might have been peculiar to activities of wound repair. If true that metals are responsible for stimulating mitotic activity, then different amounts available to oysters might well account for discrepancies in the number of mitotic amebocytes observed in different studies.

There is an alternate possibility that "dormant" amebocytes are activated by the presence of ambient copper. Prytherch (1934) found copper activated the pigment spot cells in mature eastern oyster larvae. Quiescent, or at least immobile until addition of copper, the cells were found to disaggregate and migrate into the hemolymph to become the first circulating amebocytes. This "catalytic"" action of copper (Prytherch 1934, p. 83) concurrently initiated larval metamorphosis. In vitro studies have also shown that reconstitution of cation-depleted media with seawater will initiate disaggregation and migration of adult oyster amebocytes (Fisher & Newell 1986). Mobilization of amebocytes has been further implied by the consistent finding of higher numbers of highly-locomotory amebocytes from oysters collected at metal-polluted sites (Section IV). Greater locomotory activity may account for higher numbers of amebocytes observed in hemolymph circulation. A mechanism for amebocyte activation by copper has not been investigated, but catalytic action of copper on oxidation of glutathione has been suggested to increase cell respiration (Prytherch 1934).

The distribution of eastern oysters throughout the Gulf of Mexico and along the Atlantic coast is determined by their ability to survive and reproduce in different environmental conditions. Several have proposed that the distribution of eastern oysters is dependent on freshwater inflow because of the low and variable salinities that hinder parasite growth and high-salinity predators. Low-salinity protection from both parasites and predators has been applied in the development of a "Habitat Suitability Index" for oyster (Cake 1983, Soniat & Brody 1988). However, there is ample evidence thai oysters are also successful in high-salinity environments (Beaven 1955, Lunz 1955, J.R. Nelson 1955), so an alternative reason may exist. One rationale is the availability of elements, particularly copper and zinc, from land-based sources. Terrestrial elements are carried by freshwater runoff into bays and estuaries (Reidel et al. 1995, 1998), greater volumes of water carrying them further into the receiving waters. If this dependence is found credible, then the coastal distribution of eastern oysters may be driven by a need for terrestrial elements that play critical roles in oyster physiology, including defense and nutrition (this review), shell deposition (Fisher 2004) and larval setting and metamorphosis (Prytherch 1934).

The investigations reviewed here varied widely in scope, purpose, and content. Consequently, several caveats are required. First, portions of the information reviewed may not have been intended for the purposes to which they have been applied. Even so, the pivotal studies seem well within the perceived intentions of the investigators. Although information was considered from research on several oyster species (particularly C. virginica, C. gigas, and O. edulis), the proposed function in defense is intended only for C. virginica. Adequate evidence to compare metal accumulation, adult avoidance of toxicity, and retention of accumulated metals among different oyster species was lacking. Similarly, the review has emphasized, and is limited to, the fates and effects of copper and zinc. This focus stems from the high concentrations found in C. virginica and the nearly exclusive retention of these metals within the amebocytes. Other reviewers might have included tin (Ortron 1923), manganese or iron (Galtsoff 1953, 1964) for consideration. These elements may serve physiologic roles similar to copper and zinc, but this is neither proposed nor defended here. Finally, because issues were wide-ranging, review of material for any specific topic has not been comprehensive. Many studies, such as those showing sublethal effects of copper or zinc exposure were ant included (e.g., Cheng 1988a, 1988b, Larson et al. 1989, Anderson et al. 1994, Roesijadi 1994, Ringwood et al. 1998, Butler and Roesijadi 2001). Nor were studies depicting environmental effects on bioavailability of metals (e.g., Zamuda & Sunda 1982, Wright & Zamuda 1987, Knezovich 1994), even though this topic is clearly relevant to the subject. Despite the caveats and selective focus, the evidence presented here supports a compelling image of copper and zinc as elements highly valued, if not fundamental, for oyster defense and nutrition. Such a perspective, in spite of the obvious need for testing and validation, provokes a need to re-evaluate many biologic, environmental, and resource issues that may be affected.

TABLE 1.
Concentrations of copper and zinc oxide detected in various marine
species from the Dry Tortugas as reported by Phillips (1917). Values
were recalculated from analytical results obtained with 20 g dry
tissue. Copper was determined electrolytically in nitric acid solution.
Zinc was precipitated as a sulphide in acetic acid solution and,
weighed as ZnO. Other analytes measured by Phillips (1917) include
Fe, MnO, and Pb[O.sub.2]

                                         Copper      Zinc Oxide
                                        ([micro]g     ([micro]g
        Species             Tissue     [g.sup.-1])   [g.sup.-1])

Fascioluria gigantea      Liver           2,170          180
Fasciolaria gigantea      Liver           3,725          615
Cassis Sp.                Liver             350           95
Cassis sp.                Liver              95           70
Strombus bituberculatus   Whole              30           70
Strombus gigas            Liver              30           95
Strombus gigas            Liver              40          380
Strombus gigas            Whole             125          255
Strombus gigas            Whole              10          190
Fulgur perversus          Whole              35          385
Palinurus                 Blood             700          None
Palinurus                 Liver             900           200
Palinurus                 Liver           1,100           190
Limulus polyphemus        Assorted          170           655
Limulus polyphemus        Blood             850            70
Aplysia                   Whole              14            55
Aplysia                   Whole              15            80
Aplysia                   Liver             110           100
Holothuria bermudiana     Muscle          Trace           215
Holothuria bermudiana     Intestines        200            65
Ciona atra                Whole              15            30
Gray tunicate             Whole             200           160

TABLE 2.
The amount of [sup.65]Zn in O. edulis tissues decreased with distance
from the source, a cooling water outfall from a nuclear generator in
Essex, England, as reported by Preston (1966).

Distance from Outfall   [[sup.65]Zn]

   (Statute Miles)        (pCi/g)

        0.33                100
        1.0                  42
        2.0                  33
        2.75                 19
        5.75                  9.1
        6.75                  4.5

TABLE 3.
Examples of copper and zinc concentrations ([micro]g [g.sup.-1] dry
weight) reported from tissues of eastern oysters Crassostrea virginica.

  Mean         Mean
  [Cu]         [Zn]                         Source

  403 (a)    3,263 (a)   (b) cm of 8 sites, Connecticut; Hiltner &
                           Wichman (1919, Table 2)
7,435       14,515       "Blue" oysters, New Jersey; Hiltner & Wichman
                           (1919, Table 3).
  219 (a)                (b) cm of 5 oysters, Texas; Rose & Bodansky
                           (1920)
             1,299 (a)   (b) cm of 5 oysters, Texas; Bodansky (1920)
  150 (a)      850 (a)   One eastern oyster; Severy (1923)
  109                    (b) cm of 9 lots of "white" oysters: Galtsoff
                           & Whipple (1930)
1,954                    Mean of 6 "green" oysters; Galtsoff & Whipple
                                                                (1930)
  426 (a)                North Atlantic states, winter; Coulson et al.
                           (1932)
   85 (a)                South Atlantic states, winter; Coulson et al.
                           (1932)
  134 (a)                Gulf States, winter: Coulson et al. (1932)
             6,744 (a)   (b) cm from 9 sites; Chipman et al. (1958)
  458 (a)    7,140 (a)   Mean of 100 Atlantic Coast sites; Pringle et
                           al. (1968)
   95 (a)    1,150 (a)   (b) cm from 7 sites in southeast USA: Kopfler
                           & Mayer (1969)
   82 (a)    2,969 (a)   (b) cm from 8 sites, Mobile, Alabama; Kopfler
                           & Mayer (1973)
   60 (b)    6,132       (b) cm from 4 sites in Appalachicola Bay,
                           Florida; Magley (1978)
  598                    Mean of 1978 samples, power plant, deployed;
                           Abbe (1982)
  219 (b)    9,188       (b) cm of 1978 data, Chesapeake Bay; Wright
                           et al. (1985)
  323 (b)    6,701       (b) cm of 1979 data, Chesapeake Bay; Wright
                           et al. (1985)
1,480        5,215       Chesapeake Bay power plant, deployed; Abbe &
                           Sanders (1986)
  130 (a)    3,020 (a)   (b) cm of 5 sites, Mississippi Sound: Lytle &
                           Lytle (1990)
  124 (b)    1,950       (b) cm from 8 annual geometric means:
                           O'Connor (1996) (c)
  310                    (b) cm of 10 oysters, Chesapeake Bay: Abbe et
                           al. (2000)
  415        5,240       Mean of 16 sites, Tampa Bay, Florida; Fisher
                           et al. (2000)
  682        5,374       Mean of 22 sites, five Florida Bays; Oliver
                           al. (2001)

(a) Wet weight to dry weight conversion; x5 (by convention).

(b) Calculated mean (cm): average of multiple means from different
collections documented in the original study.

(c) O'Connor (1996) reported geometric means generated from samples
collected annually at 154 sites across the United States from 1986-1993
(8 years).

TABLE 4.
Annual (1986-1993) geometric mean concentrations ([micro]g [g.sup.-1]
dry wt) of selected metals in soft tissues of oysters C. virginica
(composite of 20 for each site) or mussels Mytilus edulis (composite
of 30 for each site) collected from 154 sites across the US, as
reported from the NOAA National Status and Trends Mussel Watch Project
(O'Connor 1996). Oysters generally accumulate >l0x the copper and zinc
(and silver,  O'Connor 2002) accumulated by mussels.

             1986       1987      1988      1989      1990

Copper
  Mussels      9.9       9.9       9.3      10.0       9.9
  Oysters    110       110       130       120       150
Lead
  Mussels      2.1       2.2       2.1       1.7       1.7
  Oysters       .42       .53       .49       .45       .55
Zinc
  Mussels    140       130       130       120       140
  Oysters   1800      1700      2100      2100      2300

            1991      1992       1993     Grand Mean
Copper
  Mussels      9.0       8.7       8.1       9.2
  Oysters    120       130       120       124 (~13x)
Lead
  Mussels      2.1       2.3       1.7       1.9
  Oysters       .60       .50       .59       .58 (~0.3x)
Zinc
  Mussels    130       130       130       131.3
  Oysters   1700      2000      1900      1950 (~15x)

TABLE 5.
Tissue concentrations (dry weight) of zinc and copper were
associated with the density (number per unit area) of basophilic
amebocytes observed in histological sections of Pacific (C. gigas)
and eastern (C. virginica) oysters by Ruddell Rains (1975), which
provides statistical information. Metal analyses were performed on
whole oysters, mantle, and digestive diverticulum (dig. div.) tissue.

                         Density                   [Zn]         [Cu]
                           of        Tissue      [micro]g     [micro]g
Sample     Species      Basophils   Analyzed    [g.sup.-1]   [g.sup.-1]

  1      C. gigas          44.5     Whole           336         149
                                    Mantle          328         129
                                    Dig. div.       214          61
  2      C. gigas          85.4     Whole           422          19
                                    Mantle        1,039          47
                                    Dig. div.       400          18
  3      C. gigas         189.2     Whole           429         482
                                    Mantle        1,508         900
  4      C. gigas         346.4     Whole         1,656         319
  5      C. virginica     213.7     Whole         1,335         290
  6      C. virginica     743.0     Whole         4,598         913
                                    Mantle        5,285         968

TABLE 6.
Concentrations ([micro]g [g.sup.-1] dry weight) of copper and zinc in
amebocytes reported for various oyster species. Ruddell and Rains
(1975) estimated zinc comprised 6% of the amebocyte cell weight in C.
virginica. Brown (1975) estimated zinc to be 9% and copper 0.3%
of the cell weight. For O. edulis, Orton (1923) estimated that zinc
comprised 4.1% and copper 2.6% of the amebocyte cell weight. Thompson
et al. (1985) estimated that mantle amebocytes retained 93% of the
copper and 85% of the zinc in the mantle, and that gill amebocytes
retained 90% of the copper and 90% of the zinc in gill tissues.

  Species             Source            Method     Cell Type

C. virginica   Ruddell Rains 1975     Histo, AAS   BGA
               Brown 1975             AAS
C. gigas       Pirie et al. 1984      XRP-TEM      'Mixed'
               Thompson et al. 1985   XRP-TEM
O. edulis      Orton 1923             ?
               George et al. 1978     AAS
               Pirie et al. 1984      XRP-TEM      'Cu' cell
                                                   'Zn' cell
                                                   'Mixed'
O. angasi      Pirie et al. 1984      XRP-TEM      'Mixed'

                                             Amebocyte
  Species        Method     Cell Type           [Cu]

C. virginica   Histo, AAS   BGA
               AAS                           448-2,784
C. gigas       XRP-TEM      'Mixed'         211 (a),(b)
               XRP-TEM                      211 (a),(b)
O. edulis      ?                             25,900 (b)
               AAS                      (a) 5,571-65,000 (b)
               XRP-TEM      'Cu' cell       900 (a),(b)
                            'Zn' cell        69 (a),(b)
                            'Mixed'         991 (a),(b)
O. angasi      XRP-TEM      'Mixed'         660 (a),(b)

                                            Amebocyte
  Species        Method     Cell Type          [Zn]

C. virginica   Histo, AAS   BGA           57,447-63,227
               AAS                        20,548-89,676
C. gigas       XRP-TEM      'Mixed'       1,156 (a),(b)
               XRP-TEM                    1,228 (a),(b)
O. edulis      ?                            40,650 (b)
               AAS                      49,500-125,000 (b)
               XRP-TEM      'Cu' cell       28 (a),(b)
                            'Zn' cell     3,079 (a),(b)
                            'Mixed'        703 (a),(b)
O. angasi      XRP-TEM      'Mixed'       2,469 (a),(b)

(a) X-ray probe data: mM/kg converted to [micro]g [g.sup.-1] (multiply
by 0.6 for Cu and 0.625 for Zn).

(b) Wet weight to dry weight conversion; x5 (by convention).

Histo, histochemical: BGA, basophilic granular amebocyte: AAS, atomic
absorption spectrophotometry: XRP, X-ray probe microanalysis.

TABLE 7.
Zinc concentration factors or C. virginica at various sites in the US
calculated from data of Chipman et al. (1958). Lower
concentrations of zinc in the seawater coincided with lower zinc
concentrations in the tissues, but higher concentration factors.
Although not calculated by Chipman et al. (1958), this association
was observed by both Preston (1966) and Wolfe (1970a). (Note:
oyster tissue data is normalized to wet weight).

                                        Oyster
                     Seawater          ([micro]g      Concentration
   Location       (ng [g.sup.-1])   [g.sup.-1] wet)      Factor

Chesapeake Bay         24.0              2933           x122,208
Milford, CT            18.8              3174           x168,830
James River, VA         7.9              1484           x187,848
Beaufort, NC            4.6              1171           x254,565
Brunswick, GA           1.1               313           x284,545
Pensacola, FL           0.8               600           x750,000

TABLE 8.
Biological half lives of selected metals as calculated
by Okazaki and Panietz (1981) from concentrations in
specific tissues of C. virginica and C. gigas removed
from a contaminated site (Redwood Creek) and deployed for
56 d at a reference site (Tamales Ray) in California.

                 Tissue Concentrations
                 ([micro]g [g.sup.-1]
                          dry)

                                           Biological
               Contaminated   Reference   Half-Life (d)

C. virginica
  Ag               143            9.5        149.1
  Cu             1,548          438          156.2
  Hg                 3.36         0.38       133.5
  Zn            14,365        5,490          183.9
C. gigas
  Ag               162            3.2         26.4
  Cu             1,504           93           32.9
  Hg                 4.26         0.24        23.3
  Zn             5,531          474           36.7

(a) Average of concentrations deteintined from mantle, gill,
digestive gland and kidney on day 0 from both sites
(25 oysters).

(b) Average determined from estimates of half-life in separate
tissues (mantle, gill, digestive gland and kidney) of oysters
transplanted from the contaminated to the reference site for
56 d (5 groups of 5 oysters).

TABLE 9.
Summary of results from oysters collected at 16 sites in
Tampa Bay, Florida (Fisher et al. 2000). Top: Number of
significant positive and negative associations (P < 0.05)
between amebocyte defense characteristics and various
chemicals analyzed from oyster tissues. Defense characteristics
include amebocyte density (number [mL.sup.-1] hemolymph),
percent locomotory amebocytes, and rate of amebocyte locomotion.
Associations were determined by comparison of pooled chemical
data from each site (20 oysters) with averages of 20 individuals
from the same site. Bottom: R-values for copper, tin, and
zinc, all exhibiting significant positive associations.

              Amebocyte    Percent      Rate of
Association    Density    Locomotory   Locomotion

Positive       24             26         10
Negative        2              1          2
Metals
  Cu            0.748         --           --
  Sn            0.622         --          0.565
  Zn            0.605         --          0.546

TABLE 10.
Summary of results from oysters collected at 22 sites across
five Florida bays (Oliver et al. 2001). Top: Number of
significant positive and negative associations (P < 0.115)
between amebocyte defense characteristics and chemicals
analyzed from oyster tissues. Defense characteristics include
amebocyte density (number [mL.sup.-1] hemolymph), percent
locomotory amebocytes, and rate of amebocyte locomotion.
Associations were determined by comparison of pooled chemical
data from each site (20 oysters) with averages of 20
individuals from the same site. Bottom: R-values for arsenic,
copper, tin and zinc, all exhibiting significant positive
associations.

              Amebocyte    Percent      Rate of
Association    Density    Locomotory   Locomotion

Positive        25          26            4
Negative         0           0            2
Metals
  As             --          0.538          --
  Cu             --          0.499        0.570
  Sn             0.665       0.464        0.415
  Zn             0.463         --           --

TABLE 11.
Correlation analysis between amebocyte characteristics
and tissue metal concentrations in individual oysters
analyzed by Oliver et al. (2003). Twenty oysters were
analyzed from each of two sites (n = 40) differing
dramatically in chemical contamination. Bacterial killing
index is the percent of bacteria (Vibrio parahemolyticus)
killed in vitro and amebocyte density is the number of
amebocytes [mL.sup.-1] hemolymph. Pearson's correlation
coefficients are reported as significant at P < 0.05 *,
significant at P < 0.01 **, or not significant (ns).
Positive correlation coefficients are shown in bold type.

               Bacterial     Amebocyte
Metal        Killing Index   Density

Ag           -0.689 **         -0.437 **
Al           -0.487 **           ns
Ba           -0.387 *          -0.323 *
Cd           -6.18 *           -0.381 *
Cr           -0.571 *8       -320 *
Cu           +0.672# **        +0.462# **
Fe           -0.543 **           ns
Hg           -0.702 **         -0.441 **
Mn           +0.364# *           ns
Pb             ns                ns
Sb           -0.468 **           ns
Sn           +0.702# **        +0.485# **
Zn           +0.738# **        +0.488# *
TBT            ns                ns
DBT          +0.682# **        +0.614# **
All Metals   +0.731# **        +0.490# **

TBT, tributyltin; DBT, Dibutyltin.

'All Metal' represents a composite analysis of
all metals measured.

ACKNOWLEDGMENTS

Florida field survey data were obtained through the combined efforts of my colleagues, Leah M. Oliver and James T. Winstead, to whom I am grateful for many collaborations over the last decade. Thanks are also due our collaborators Edward R. Long, Aswani K. Volety, Fred J. Genthner, and Becky L. Hemmer. I am indebted to Dr. Robert S. Brown, who graciously provided both insight and material from his unpublished dissertation at Johns Hopkins University. Library material was assembled with the assistance of Sonya Doten and Liz Pinnell, and numerous graphics rendered by Stephen Embry. I am indebted to P. Chapman, W. Davis, J. Fournie, L. DiMichael, D. Shepard and C. Walker for meaningful discussions. Valuable comments and recommendations from reviewers of early versions, including L. Oliver, S. Jordan, R. Anderson, G. Roesijadi, and M. Carriker are greatly appreciated. This was prepared in support of the Gulf Ecology Division mission in Altered Habitat, and is GED contribution #1196.

LITERATURE CITED

Abbe, G. R. 1982. Growth, mortality, and copper-nickel accumulation by oysters (Crassostrea virginica) at the Morgantown Stam Electric Station on the Potomac River, Maryland. J. Shellfish Res. 2:3-13.

Abbe, G. R., G. F. Reidel & J. G. Sanders. 2000. Factors that influence the accumulation of copper and cadmium by transplanted eastern oysters (Crassostrea virginica) in the Patuxent River, Maryland. Mar. Environ. Res. 49:377-396.

Abbe, G. R. & J. G. Sanders. 1986. Condenser replacement in a coastal power plant: copper uptake and incorporation in the American oyster. Crassostrea virginica. Mar. Environ. Res. 19:93-113.

Acton, R. T., E. E. Evans & J. C. Bennett. 1969. Immunological capabilities of the oyster Crassostrea virginica. Comp. Biochem. Physiol. 29: 149-160.

Alexander, G. V. & R. H. Rowland. 1966. Estimation of zinc-65 background levels for marine coastal waters. Nature 210:155-157.

Alvarez, M. R. & F. E. Friedl. 1992. Effect of a fungicide on in vitro hemocyte viability, phagocytosis and attachment in the American oyster Crassostrea virginica. Aquaculture 107:135-140.

Alvarez, M. R., F. E. Friedl & C. M. Hudson. 1991. Effect of a commercial fungicide on the viability and phagocytosis of hemocytes of the American oyster, Crassostrea virginica. J. Invertebtr. Pathol. 57:395-401.

Anderson, R. S. 1975. Phagocytosis by invertebrate cells in vitro: Biochemical events and other characteristics compared with vertebrate phagocytic systems. In: K. Maramorsch & R. E. Shope, editors. Invertebrate Immunity. New York: Academic Press. p. 365.

Anderson, R. S. 1996. Interactions of Perkinsus marinus with humoral factors and hemocytes of Crassostrea virginica. J. Shellfish Res. 15: 127-134.

Anderson. R. S., L. L. Brubacher, L. Ragone Calvo. E. M. Burreson & M. A. Unger. 1997. Effect of in vitro exposure to tributyltin on generation of oxygen metabolites by oyster hemocytes. Environ. Res. 74: 84-90.

Anderson, R. S., L. L. Brubacher, L. Ragone Calvo, M. A. Unger & E. M. Burreson. 1998. Effects of tribulyltin and hypoxia on the progression of Perkinsus marinus infections and host defense mechanisms in oyster. Crassostrea virginica (Gmelin). J. Fish Diseases 21:371-380.

Anderson, R. S., L. M. Mora & S. A. Thomson. 1994. Modulation of oyster (Crassostrea virginica) hemocyte immune function by copper, as measured by luminol-enhanced chemiluminescence. Comp. Biochem. Physiol. 108C:215-220.

Anderson, R. S., L. M. Oliver & D. Jacobs. 1992. Immunotoxicity of cadmium for the eastern oyster [Crassostrea virginica (Gmelin, 1791)]: effects on hemocyte chemiluminescence. J. Shellfish Res. 11:31-35.

Anderson, R. S., M. A. Unger & E. M. Burreson. 1996. Enhancement of Perkinsus marinus disease progression in TBT-exposed oysters (Crassostrea virginica). Mar. Environ. Res. 42:177-180.

Andrews, J. D. 1988. Epizootiology of the disease caused by the oyster pathogen Perkinsus marinus and its effects on the oyster industry. Amer. Fish. Soc. Spec. Publ. 18:47-63.

Andrews, J. D. & S. M. Ray. 1988. Management strategies to control the disease caused by Perkinsus marinus. Amer. Fish. Soc. Spec. Publ. 18:257-264.

Asokan, R., M. Arumugam & P. Mullainadhan. 1997. Activation of prophenoloxidase in the plasma and hemocytes of the marine mussel Perna viridis Linnaeus, Dev. Comp. Immunol. 21:1-12.

Atkinson, P. G. P. & C. H. Barton. 1998. Ectopic expression of Nrampl in COS-1 cells modulates iron accumulation. FEBS Letters 425:239-242.

Baier-Anderson, C. & R. S. Anderson. 1997. The effect of pentachlorophenol on pyridine nucleotide production in oyster hemocytes: NADPH and immunomodulation. J. Shellfish Res. 16:111-114.

Baier-Anderson, C. & R. S. Anderson. 2000. Immunotoxicity of environmental pollutants in marine invertebrates. In: M. Fingerman & R. Nagabhushanam, editors. Recent advances in marine biotechnology, vol. 5, Immunobiology and Pathology. Enfield, USA; Plymouth. UK: Science Publishers, Inc. pp. 189-225.

Bang, F. B. 1961. Reaction to injury in the oyster (Crassostrea virginica). Biol. Bull. 121:57-68.

Bang, F. B. 1973. A survey of phagocytosis as a protective mechanism against disease among invertebrates. In: W. Braun & J. Ungar, editors. "Non-Specific" Factors Influencing Host Resistance. Basel, Switzerland: S. Karger. pp. 2-10.

Beaven, A. E. & K. T. Paynter. 1999. Acidification of the phagosome in Crassostrea virginica hemocytes following engulfment of Zymosan. Biol. Bull. 196:26-33.

Beaven, G. F. 1955. Various aspects of oyster setting in Maryland. Proc. Nat. Shellfish. Assoc. (1954) 45:29-37.

Bernard, F. R. 1989. Uptake and elimination of coliform bacteria by tour marine bivalve mollusks. Can. J. Fish. Aquat. Sci. 46:1592-1599.

Berrigan, M., T. Candies, J. Cirino, R. Dugas, C. Dyer, J. Gray, T. Herrington, W. Keithly, R. Leard, J. R. Nelson & M. Van Hoose. 1991. The oyster fishery of the Gulf of Mexico, United States: a regional management plan. Ocean Springs. MA: Gulf States Marine Fisheries Commission, pp. 5-20.

Bizio. 1845. Transactions of the Institute of Venice. Cited from Lankester 1886.

Blake, P. A., M. H. Merson, R. E. Weaver, D. G. Hollis & P. C. Heublein. 1979. Disease caused by a marine vibrio. N. Engl. J. Med. 300:1-7.

Blake, P. A., R. E. Weaver & D. G. Hollis. 1980. Diseases of humans (other than cholera) caused by vibrios. Ann. Rev. Microbiol. 34:341-367.

Bodansky, M. 1920. Biochemical studies on marine organisms. II. The occurrence of zinc. J. Biol. Chem. 44(2):399-407.

Boyce, R. & W. A. Herdman. 1897. On a green leucocytosis in oysters associated with the presence of copper in the leucocytes. Proc. Royal Society; London 62:30-38.

Boyden, C. R. 1975. Distribution of some trace metals in Poole Harbor, Dorset. Mar. Pollut. Bull. 6:180-187.

Boyden, C. R. & D. J. H. Phillips. 1981. Seasonal variation and inherent variability of trace elements in oysters and their implications for indicator studies. Mar. Ecol. Progr. Ser. 5:29-40.

Boyden, C. R. & M. G. Romeril. 1974. A trace metal problem in pond oyster culture. Mar. Pollut. Bull. 5:74-78.

Bradley, H. C. 1904. The occurrence of zinc in certain invertebrates. Science xix(474): 196-197.

Brooks, R. R. & M. G. Rumsby. 1965. The biogeochemistry of trace element uptake by some New Zealand bivalves. Limnol: Oceanogr. 10:521-528.

Brown, R. S. 1975. Evidence for metal-protein complexes precipitating around blood cells as the mechanism of extracellular blood clotting in the American oyster Crassostrea virginica (Gmelin), Ph.D. dissertation, Baltimore, MD: Johns Hopkins University. 106 pp.

Bryan, G. W. & W. J. Langston. 1992. Bioavailability, accumulation and effects of heavy metals in sediments with special reference to United Kingdom estuaries: a review, Environ. Pollut. 76:89-131.

Burreson, E. M., M. E. Robinson & A. Villalba. 1988. A comparison of paraffin histology and hemolymph analysis for the diagnosis of Haplosporidium nelsoni (MSX) in Crassostrea virginica. (Gmelin). J. Shellfish Res. 7:19-23.

Butler, R. A. & G. Roesijadi. 2001. Disruption of metallothionein expression with antisense oligonucleotides abolishes protection against cadmium cytotoxicity in molluscan hemocytes. Toxicol. Sci. 59:101-107.

Cake, E. W., Jr. 1983. Habitat suitability index models: Gulf of Mexico American oyster. US Fish Wildlife Service, FWS/OBS-82/10.57. 37 pp.

Calabrese, A., R. S. Collier, D. A. Nelson & J. R. MacInnes. 1973. Toxicity of heavy metals to embryos of the American oyster Crassostrea virginica. Mar. Biol. 18:162-166.

Calabrese, A., J. R. MacInnes, D. A. Nelson & J. E. Miller. 1977. Survival and growth of bivalve larvae under heavy-metal stress. Mar. Biol. 41:179-184.

Carballal, M. J., C. Lopez, C. Azevedo & A. Villalba. 1997. Enzymes involved in defense functions of hemocytes of mussel Mytilus galloprovincialis. J. Invertebr. Pathol. 70:96-105.

Cheng, T. C. 1967. Marine molluscs as hosts for symbioses: with a review of known parasites of commercially important species. Adv. Mar. Biol. 5:14-24.

Cheng, T. C. 1975. Functional morphology and biochemistry of molluscan phagocytes. Ann. N.Y. Acad. Sci. 266:343-379.

Cheng, T. C. 1977. Biochemical and ultrastructural evidence for the double role of phagocytosis in molluscs: defense and nutrition. In: L. A. Bulla, Jr., & T. C. Cheng, editors. Comparative pathobiology, vol 3. Invertebrate immune responses. New York; and London: Plenum Press. pp. 21-30.

Cheng, T. C. 1981. Bivalves. In: N. A. Ratcliffe & A. F. Rowley, editors. Invertebrate blood cells. London: Academic Press, pp. 233-300.

Cheng, T. C. 1983. Internal defense mechanisms of molluscs against invading microorganisms: personal reminiscences. Trans. Amer. Microscop. Soc. 102:185-193.

Cheng, T. C. (editor) 1984. Comparative Pathobiology Vol 6: Invertebrate Blood; Cells and Serum Factors. New York (USA) and London (England): Plenum Press. 211 pp.

Cheng, T. C. 1988a. In vivo effects of heavy metals on cellular defense mechanisms of Crassostrea virginica: total and differential cell counts. J. Invertebr. Pathol. 51:207-214.

Cheng, T. C. 1988b. In vivo effects of heavy metals on cellular defense mechanisms of Crassostrea virginica: phagocytic and endocytotic indices. J. Invertebr. Pathol. 51:215-220.

Cheng, T. C. 1996. Hemocytes: forms and functions. In: V. S. Kennedy, R. I. E. Newell, & A. F. Eble, editors. The Eastern Oyster Crassostrea virginica. College Park, Maryland (USA): Maryland Sea Grant College program, University of Maryland System. pp. 299-333.

Cheng, T. C. & A. Cali. 1974. An electron microscope study of the fate of bacteria phagocytized by granulocytes of Crassostrea virginica. Contemp. Topics Immunobiol. 4:25-35.

Cheng, T. C. & E. Rifkin. 1970. Cellular reactions in marine molluscs in response to helminth parasitism. In: S. F. Snieszko, editor. A symposium on diseases of fishes and shellfishes, vol. 5. Washington, D. C. (USA): Amer. Fish. Soc. Symp. pp. 443-496.

Cheng, T. C. & B. M. Rudo. 1976. Distribution of glycogen resulting from degradation of [sup.14]C-labelled bacteria in the American oyster. Crassostrea virginica. J. Invertebr. Pathol. 27:259-262.

Cheng, T. C. & J. T. Sullivan. 1984. Effects of heavy metals on phagocytosis by molluscan hemocytes. Mar. Environ. Res. 14:305-315.

Cheng, T. C., G. E. Rodrick, D. A. Foley & S. A. Koehler. 1975. Release of lysozyme from hemolymph cells of Mercenaria mercenaria during phagocytosis. J. Invertebr. Pathol. 25:261-265.

Chipman, W. A., T. R. Rice & T. J. Price. 1958. Uptake and accumulation of radioactive zinc by marine plankton, fish, and shellfish. US Fish Wild Serv. Fish Bull. 58:278-292.

Chu, F.-L. E. 1988. Humoral defense factors in marine bivalves. Amer. Fish. Soc. Spec. Publ. 18:178-188.

Chu, F.-L. E. 2000. Defense mechanisms of marine bivalves. In: M. Fingerman & R. Nagabhushanam, editors. Recent advances in marine biotechnology, Vol. 5. Immunobiology and pathology. Enfield USA; Plymouth, UK: Science Publishers. pp. 1-42.

Chu, F.-L. E. & K. H. Greene. 1989. Effect of temperature and salinity on in vitro culture of the oyster pathogen, Perkinsus marinus (Apicomplexa: Perkinsea). J. Invertebr. Pathol. 53:260-268.

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

Chu, F.-L. E., J. F. La Peyre & C. S. Burreson. 1993. Perkinsus marinus infection and potential defense-related activities in eastern oysters, Crassostrea virginica: salinity effects. J. Invertebr. Pathol. 62:226232.

Coles, J. A. & R. A. Pipe, 1994, Phenoloxidase activity in the haemolymph and haemocytes of the marine mussel Mytilus edulis. Fish Shellfish Immunol. 4:337-352.

Coombs, T. L. 1972. The distribution of zinc in the oyster Ostrea edulis and its relation to enzymic activity and to other metals. Mar. Biol. 12:170-178.

Copeland, B. J. & H. D. Hoese. 1966. Growth and mortality of the American oyster, Crassostrea virginica, in high salinity shallow bays in central Texas. Publ. Inst. Mar. Sci. Univ. Texas 11:149-158.

Cuenot, L. 1891. Etudes sur le sang et les glandes lymphatiques dans la serie animale (2 Portie: Invertebres). Arch. Z. Exper. 9:19-54.

Daniels, N. A., B. Ray, A. Easton, N. Marano, E. Kahn, A. McShan, L. Del Rosario, T. Baldwin, M. Kingsley, N. Puhr, J. Wells, D. Cameron, P. Griffin, R. Tauxe & F. Angulo. 1999. Large multistate outbreak of oyster-associated Vibrio parahaemolyticus infections: emergence of virulent serotype highlights prevention quandary. J. Gen. Intern. Med. 14:23.

Deaton, L. E. & J. R. Dankert. 1998. Phenoloxidase activity in the hemolymph of the molluscs Geukensia demissa and Mercenaria mercenaria. J. Shellfish Res. 17:323.

DesVoigne, D. M. & A. K. Sparks. 1968. The process of wound healing in the Pacific oyster, Crassostrea gigas. J. Invertebr. Pathol. 12:53-65.

Duke, T. W. 1967. Possible routes of zinc 65 from an experimental estuarine environment to man. J. Water Poll. Cont. Fed. 39:536-542.

Feng, J. S. 1966. The fate of a virus, Staphylococcus aureus phage 80, injected into the oyster, Crassostrea virginica. J. Invertebr. Pathol. 8:496-504.

Feng, S. Y. 1958. Observations on the distribution and elimination of spores of Nematopsis ostrearum in oysters. Proc. Nat. Shellfish. Assoc. 48:162-173.

Feng, S. Y. 1966. Experimental bacterial infection in the oyster Crassostrea virginica. Invertebr. Pathol. 8:505-511.

Feng, S. Y. 1967. Responses of molluscs to foreign bodies, with special reference to the oyster. Fed. Proc. 26:1685-1692.

Feng, S. Y. 1988. Cellular defense mechanisms of oysters and mussels. Amer. Fish. Soc. Spec. Publ. 18:153-168.

Feng, S. Y., J. S. Feng, C. N. Burke & L. H. Khairallah. 1971. Light and electron microscopy of the leuococytes of Crassostrea virginica (Mollusca: Pelecypoda). Z. Zellforsch. 120:222-245.

Feng, S. Y., J. S. Feng & T. Yamasu. 1977. Roles of Mytilus coruscus and Crassostrea gigas blood cells in defense and nutrition. In: L. A. Bulla. Jr., & T. C. Cheng, editors. Comparative pathobiology, vol 3, Invertebrate Immune Responses. New York: and London: Plenum Press. pp. 31-67.

Fisher, W. S. 1986. Structure and functions of oyster hemocytes. In: M. Brehelin, editor. Immunity in invertebrates. Berlin: Springer-Verlag. pp. 25-35.

Fisher, W. S. 1988. Environmental influence on bivalve hemocyte function. Amer. Fish. Soc. Spec. Publ. 18:225-237.

Fisher, W. S. 2004. Relationship of amebocytes and terrestrial elements to adult shell deposition in eastern oysters. J. Shellfish Res. 23.

Fisher, W. S., M. M. Chintala & M. A. Moline. 1989. Annual variation of estuarine and oceanic oyster Crassostrea virginica Gmelin hemocyte capacity. J. Exp. Mar. Biol. Ecol. 127:105-120.

Fisher, W. S. & R. I. E. Newell. 1986. Salinity effects on the activity of granular hemocytes of American oysters, Crassostrea virginica. Biol. Bull. 170:122-134.

Fisher, W. S., L. M. Oliver & P. Edwards. 1996. Hematologic and serologic variability of eastern oysters from Apalachicola Bay. Florida. J. Shellfish Res. 15:555-564.

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

Fisher, W. S., L. M. Oliver, J. T. Winstead & E. R. Long. 2000. A survey of oysters Crassostrea virginica from Tampa Bay, Florida: Associations of internal defenses with contaminant burdens. Aquat. Toxicol. 51:115-138.

Fisher, W. S., L. M. Oliver, J. T. Winstead & A. K. Volety. 2003. Stimulation of defense factors in eastern oysters deployed to contaminated sites in Pensacola Bay. Florida. Aquat. Toxicol. 64:375-391.

Fisher, W. S. & M. L. Tamplin. 1988. Environmental influence on activity and foreign-particle binding by hemocytes of American oysters, Crassostrea virginica. Can. J. Fish. Aquat. Sci. 45:1309-1315.

Fisher, W. S., A. Wishkovsky & F.-L. E. Chu. 1990. Effects of tributyltin on defense related activities of oyster hemocytes. Arch. Environ. Contam. Toxicol. 19:354-360.

Fitzgerald, B. W. & D. M. Skauen. 1963. Zinc-65 in oysters in Fishers Island Sound and its estuaries. In: V. Schultze & A. W. Klement, editors. Radioecology. New York: Reinhold, pp. 159-162.

Foley, D. A. & T. C. Cheng. 1975. A quantitative study of phagocytosis by hemolymph cells of the pelecypods Crassostrea virginica and Mercenaria mercenaria. J. Invertebr. Pathol. 25:189-197.

Foley, D. A. & T. C. Cheng. 1977. Degranulation and other changes in molluscan granulocytes associated with phagocytosis. J. Invertebr. Pathol. 29:321-325.

Ford, S. E. 1985. Effects of salinity on survival of the MSX parasite Haplosporidium nelsoni (Haskin, Stauber, and Mackin) in oysters. J. Shellfish Res, 5:85-90.

Ford, S. E. 1996. Range extension by the oyster parasite Perkinsus marinus into the northeastern United States: response to climate change? J. Shellfish Res. 15:45-56.

Ford, S. E. & H. H. Haskin. 1988. Management strategies for MSX (Haplosporidium nelsoni) disease in eastern oysters. Amer. Fish. Soc. Spec. Publ. 18:249-256.

Ford, S. E. & S. A. Kanaley. 1988. An evaluation of hemolymph diagnosis for the detection of the oyster parasite Haplosporidium nelsoni (MSX). J. Shellfish Res. 7:11-18.

Frazier, J. M. 1975. The dynamics of metals in the American oyster. Crassostrea virginica. I. Seasonal effects. Chesapeake Sci. 16:162-171.

Frazier, J. M. 1976. The dynamics of metals in the American oyster. Crassostrea virginica. II. Environmental effects. Chesapeake Sci. 17: 188-197.

Fries, C. R. & M. R. Tripp. 1970. Uptake of viral particles by oyster leucocytes in vitro. J. Invertebr. Pathol. 15:136-137.

Gaillon, B. 1824. Sur la cause de la coloration des huitres et sur les animacules qui servent a leur nutrition. Memoires de la Societe Linneene du Calvados, pp. 135-158.

Galtsoff, P. S. 1938. Shellfish investigations. In: E. Higgins, compiler. Progress in biological inquiries 1937 Washington DC: US Government Printing Office, U. S. Department of Commerce, Bureau of Fisheries Administrative Report No. 30. pp. 42-52.

Galtsoff, P. S. 1953. Accumulation of manganese, iron, copper and zinc in the body of American oysters (Crassostrea) Ostrea virginica. Anat. Rec. 117:601-602.

Galtsoff, P. S. 1964. The American oyster Crassostrea virginica Gmelin. Fish. Bull. 64:1-480.

Galtsoff, P. S. & D. V. Whipple. 1930. Oxygen consumption of normal and green oysters. US Department of Commerce. Bull. Bureau Fisheries XLVI:488-508.

Genthner, F. J., A. K. Volety, L. M. Oliver & W. S. Fisher. 1999. Factors influencing in vitro killing of bacteria by hemocytes of the eastern oyster (Crassostrea virginica). Appl. Environ. Microbiol. 65:3015-3020.

George, S. G., B. J. S. Pirie, A. R. Cheyne, T. L. Coombs & P. T. Grant. 1978. Detoxication of metals by marine bivalves: an ultrastructural study of the compartmentation of copper and zinc in the oyster Ostrea edulis. Mar. Biol. 45: 147-156.

Gordon, M., G. A. Knauer & J. H. Martin. 1980. Mytilus californianus as a bioindicator of trace metal pollution: variability and statistical considerations. Mar. Pollut. Bull. 11:195-198.

Gregoire, C. 1970. Haemolymph coagulation in arthropods. Symp. Zool. Soc. Lond. 27:45-74.

Greig, R. A. & D. R. Wenzloff. 1978. Metal accumulation and depuration by the American oyster, Crassostrea virginica. Bull. Environm. Contam. Toxicol. 20:499-504.

Gunter, G. & R. A. Geyer. 1955. Studies of fouling organisms in the northwestern Gulf of Mexico. Publ. Inst. Mar. Sci. Univ. Texas 4(1): 39-67.

Hargis, W. J. & D. S. Haven. 1999. Chesapeake oyster reefs, their importance, destruction and guidelines for restoring them. In: M. W. Luckenbach, R. Mann & J. A. Wesson, editors. Oyster reef habitat restoration: a synopsis and synthesis of approaches. Gloucester Point, Virginia: Virginia Institute of Marine Science Press. pp. 329-358.

Harris-Young, L., M. L. Tamplin, W. S. Fisher & J. W. Mason. 1993. Effects of physicochemical factors and bacterial colony morphotype on association of Vibrio vulnificus with hemocytes of Crassostrea virginica. Appl. Environ. Microbiol. 59:1012-1017.

Harris-Young, L., M. L. Tamplin, J. W. Mason, H. C. Aldrich & J. K. Jackson. 1995. Viability of Vibrio vulnificus in association with hemocytes of the American oyster (Crassostrea virginica). Appl. Environ. Microbiol. 61:52-57.

Haskin, H. H. & J. D. Andrews. 1988. Uncertainties and speculations about the life cycle of the eastern oyster pathogen Haplosporidium nelsoni (MSX). Amer. Fish. Soc. Spec. Publ. 18:5-22.

Herdman, W. A. 1896. Investigations on oysters and disease. Report for 1895 on the Lancashire Sea-Fisheries Laboratory, Transactions. Liverpool Biol. Soc. 10:158-177.

Herdman, W. A. & R. Boyce. 1899. Oysters and disease, an account of certain observations upon the normal and pathological histology and bacteriology of the oyster and other shellfish. London: Lancashire Sea-Fisheries Memoir No. 1. 8 plates. 60 pp.

Hillman, R. E. 1968. Histochemistry of mucosubstances in the mantle of the clam, Mercenaria mercenaria. I: a glycosaminoglycan in the first marginal fold. Trans. Amer. Microsc. Soc. 87:361-367.

Hillman, R. E. 1969. Histochemistry of mucosubstances in the mantle of the clam. Mercenaria mercenaria. II. Mucosubstances in the second marginal fold. Trans. Amer. Microsc. Soc. 88:420-425.

Hillman, R. E. 1978. Invertebrate mucus: model systems for studying diseases in man. In: P. N. Kaul & C. J. Sindermann, editors. Drugs and food from the sea myth or reality? Norman, Oklahoma (USA): Univ. Oklahoma. pp. 17-25.

Hiltner, R. S. & H. J. Wichmann. 1919. Zinc in oysters. J. Biol. Chem. 38:205-221.

Hinsch, G. W. & M. Hunte. 1990. Ultrastructure of phagocytosis by hemocytes of the American oyster. In: F. O. Perkins & T. C. Cheng, editors. Pathology in marine science. San Diego, CA: Academic Press. pp. 479-488.

Hopkins, S. H. 1956. The boring sponges which attack South Carolina oysters, with notes on some associated organisms. Contrib. Bears Bluff Laboratories, no. 23. Wadmalaw Island, SC: Bears Bluff Laboratories. 30 pp.

Hunter, A. C. & C. W. Harrison. 1928. Bacteriology and chemistry of oysters, with special reference to regulatory control of production, handling, and shipment. Washington, D.C: US Department of Agriculture technical bulletin, No. 64. pp. 1-75.

Ikuta, K. 1958a. Studies on accumulation of heavy metals in aquatic organisms -II. On accumulation of copper and zinc in oysters, Bull. Jap. Soc. Sci. Fish. 34:112-116.

Ikuta, K. 1958b. Studies on accumulation of heavy metals in aquatic organisms -III. On accumulation of copper and zinc in the parts of oysters. Bull. Jap. Soc. Sci. Fish. 34:117-122.

Jabado, N., A. Jankowski, S. Dougaparsad, V. Picard, S. Grinstein & P. Gros. 2000. Natural resistance to intracellular infections: natural resistance-associated macrophage protein 1 (nramp1) functions as a pH-dependent manganese transporter at the phagosomal membrane. J. Exp. Med. 192:1237-1247.

Jarinen, A. W. & G. T. Ankley. 1999. Linkage of effects to tissue residues: development of a comprehensive database for aquatic organisms exposed to inorganic and organic chemicals. Pensacola FL: Society for Environmental Toxicology and Chemistry.

Jones, S. H., T. L. Howell & K. O'Neill. 1991. Differential elimination of indicator bacteria and pathogenic Vibrio sp. from eastern oysters (Crassostrea virginica Gmelin, 1791) in a commercial controlled purification facility in Maine. J. Shellfish Res. 10:105-112.

Jiann, K.-T. & B. J. Presley. 1997. Variations in trace metal concentrations in American oysters (Crassostrea virginica) collected from Galveston Bay, Texas. Estuaries 20:710-724.

Kelly, C. B. & W. Arcisz. 1954. Survival of enteric organisms in shellfish. U S Pub. Health Rep. 69:1205-1210.

Knezovich, J. P. 1994. Chemical and biological factors affecting bioavailability of contaminants in seawater. In: J. L. Hamelink, P. F. Landrum, H. L. Bergman, & W. H. Benson, editors. Bioavailability: physical, chemical, and biological interactions. Ann Arbor, Michigan: Lewis Publishers. pp. 23-30.

Kopfler, F. C. & J. Mayer. 1969. Studies on trace metals in shellfish. In: R. J. Hammerstrom & W. F. Hill, Jr., editors. Proceedings gulf and south Atlantic States Shellfish Sanitation Research Conference. Cincinnati. Ohio: US Dept. Health, Education, and Welfare, Public Health Service. pp. 67-80.

Kopfler, F. C. & J. Mayer. 1973. Concentrations of five trace metals in the waters and oysters (Crassostrea virginica) of Mobile Bay, Alabama. Proc. Nat. Shellfish. Assoc. 63:27-34.

Korringa, P. 1952. Recent advances in oyster biology. Qtr. Rev. Biol. 27(3):266-308.

Kramer, T. R. & W. T. Johnson. 1992. Copper and immunity. In: S. Cunningham-Rundles, editor. Nutrient modulation of the immune response. New York: Marcel Dekker Inc. pp. 239-254.

Lankester, E. R. 1886. On green oysters. Qtr. J. Microscop. Sci. pp. 71-94, plus 7 plates.

Larson, K. G., B. S. Roberson & F. M. Hetrick. 1989. Effect of environmental pollutants on the chemiluminescence of hemocytes from the American oyster Crassostrea virginica. Dis. Aquat. Org. 6:131-136.

La Peyre, M. K., A. D. Nickens, A. K. Volety, S. G. Tolley & J. F. La Peyre. 2003. Environmental significance of freshets in reducing Perkinsus marinus infections in eastern oysters (Crassostrea virginica): potential management implications. Mar. Ecol. Progr. Ser. 248:165176.

Lobel, P. B. & D. A. Wright. 1983. Frequency distribution of zinc in Mytilus edulis (L.). Estuaries 6:154-159.

Long, E. R., D. MacDonald & C. Cairncross. 1991. Status and trends in toxicants and the potential for their biological effects in Tampa Bay, Florida. NOAA Technical Memorandum NOS OMA 58, Seattle. WA. 77 pp.

Long, E. R., D. A. Wolfe, R. S. Carr, K. J. Scott, G. B. Thursby, H. L. Windom, R. Lee, F. D. Calder, G. M. Sloane. & T. Seal. 1994. Magnitude and extent of sediment toxicity in Tampa Bay, Florida. NOAA Technical Memorandum NOS ORCA 78. Silver Spring. MD: US National Oceanic and Atmospheric Administration. 138 pp.

Loosanoff, V. L. 1949. On the food selectivity of oysters. Science 110:122.

Loosanoff, V. L. 1955. How to increase production of seed oysters in Connecticut. Proc. Nat. Shellfish. Assoc. (1954) 45:19-22.

Lunz, G. R. 1955. The general pattern of oyster setting in South Carolina. Proc. Nat. Shellfish. Assoc. (1954) 45:47-51.

Lytle, T. F. & J. S. Lytle. 1990. Heavy metals in the eastern oyster, Crassostrea virginica, of the Mississippi Sound. Ball. Environ. Contam. Toxicol. 44:142-148.

Mackin, J. G. 1951. Histopathology of infection of Crassostrea virginica (Gmelin) by Dermocystidium marinum Mackin, Owen and Collier. Ball. Mar. Sci. Gulf Caribb. 1:72-87.

MacInnes, J. R. 1980-81. Response of embryos of the American oyster, Crassostrea virginica, to heavy metal mixtures. Mar. Environ. Res. pp. 217-227.

McCain, B. B., D. W. Brown, T. Hom, M. S. Myers, S. M. Pierce, T. K. Collier, J. E. Stein. S.-L. Chan & U. Varanasi. 1996. Chemical contaminant exposure and effects in four fish species from Tampa Bay. Florida. Estuaries 19:86-104.

McDade, J. E. & M. R. Tripp. 1967. Lysozyme in the hemolymph of the oyster, Crassostrea virginica. J. Invertebr, Pathol. 9:531-535.

Mendel, L. B. & H. C. Bradley. 1905. Experimental studies on the physiology of the molluscs second paper. Amer. J. Physiol. xiv:313-327.

Mendel, L. B. & H. C. Bradley. 1906-07. Experimental studies on the physiology of the molluscs. Amer. J. Physiol. xvii:167.

Menzel, R. W. 1955. Some phases of the biology of Ostrea equestris Say and a comparison with Crassostrea virginica (Gmelin). Publs. Inst. Mar. Sci. Univ. Texas 4:73-148.

Menzel, R. W., N. C. Hulings & R. R. Hathaway. 1966. Oyster abundance in Apalachicola Bay, Florida in relation to biotic associations influenced by salinity and other factors. Golf Research Reports 2:73-96.

Metchnikoff, E. 1891. Lectures on the Comparative Pathology of Inflammation. Delivered at the Pasteur Institute in 1891, translated by F. A. Starling & E. H. Starling and originally published by Kegan Paul, Trench, Trubner & Co., Ltd. in 1893. Republished in 1968 by New York: Dover Publications Inc. 224 pp.

Murthy, G. K., A. S. Goldin & J. E. Campbell. 1959. Zinc-65 in foods. Science 130(3384): 1255-1256.

Narain, A. S. 1973. The amoebocytes of lamellibranch molluscs, with special reference to the circulating amoebocytes. Malacol. Rev. 6: 1-12.

Nelson, J. 1916. Copper content of green oysters. Report, Department of Biology, New Jersey State Agricultural Experiment Station (for 1915), Trenton, NJ, pp. 246-249.

Nelson, J. R. 1955. Symposium on various aspects of oyster setting. Oyster setting. Proc. Nat. Shellfish. Soc. (1954) 45: 16-18.

Newell, R. I. E. & S. J. Jordan. 1983. Preferential ingestion of organic material by the American oyster, Crassostrea virginica. Mar. Ecol. Prog. Ser. 13:47-53.

National Oceanic and Atmospheric Administration 1987. National status and trends program for marine environmental quality. A summary of selected data on chemical contaminants in tissues collected during 1984, 1985, and 1986. Technical Memorandum NOS OMA 38. US Department of Commerce, National Oceanic and Atmospheric Administration, Office of Oceanography and Marine Assessment, Ocean Assessments Division, Rockville, MD, 23 pp plus appendices.

O'Connor, T. P. 1994. The National Oceanic and Atmospheric Administration national status and trends mussel watch program: national monitoring of chemical contamination in the coastal United States. In: C. R. Cothern & N. P. Ross, editors. Environmental statistics, assessment, and forecasting. Boca Raton, Florida: Lewis Publishers. pp. 331-339.

O'Connor, T. P. 1996. Trends in chemical concentrations in mussels and oysters collected along the US coast from 1986-1993. Mat. Environ. Res. 41:183-200.

O'Connor, T. P. 2002. National distribution of chemical concentrations in mussels and oysters in the USA. Mar. Environ. Res. 53:117-143.

O'Connor, T. P. & C. N. Ehler. 1991. Results from NOAA national status and trends program on distributions and effects of chemical contamination in the coastal and estuarine United States. Environ. Monit. Assess. 17:33-49.

Okazaki, R. K. 1976. Copper toxicity in the Pacific oyster Crassostrea gigas. Bull. Environ. Contam. Toxicol. 16:658-664.

Okazaki, R. K. & M. H. Panietz. 1981. Depuration of twelve metals in tissues of the oysters Crassostrea gigas and C. virginica. Mar. Biol. 63:113-120.

Oliver, L. M. & W. S. Fisher. 1999. Appraisal of prospective bivalve immunomarkers. Biomarkers 4(6):510-530.

Oliver, L. M., W. S. Fisher, J. T. Winstead, B. L. Hemmer & E. R. Long. 2001. Relationships between tissue contaminants and defense-related characteristics of oysters (Crassostrea virginica) from five Florida bays. Aquat. Toxicol. 55:203-222.

Oliver, L. M., W. S. Fisher. A. K. Volety & Z. Malaeb. 2003. Relationships between oyster (Crassostrea virginica) defense measurements and tissue contaminant concentrations. Aquat. Toxicol. 64:363-373.

Orton, J. H. 1923. Summary of an account of investigations into the cause or causes of the unusual mortality among oysters in English oyster beds during 1920 and 1921. J. Mar. Biol. Assoc. UK. 13:1-23.

O'Shaughnessy, A. W. E. 1866. On green oysters. Ann. Mag. Nat. Hist. Ser. 3, vol. xviii, contribution XXXI, pp. 221-228.

Owen, G. 1966. Digestion. In: K. M. Wilbur & C. M. Yonge, editors. Physiology of mollusca, vol. 2. New York: Academic Press, pp. 53-96.

Pauley, G. B. & A. K. Sparks. 1965. Preliminary observations on the acute inflammatory reaction in the Pacific oyster, Crassostrea gigas (Thunberg). J. Invertebr. Path 7:248-257.

Pavia, A. T., J. F. Campbell, P. A. Blake, J. D. Smith, T. W. McKinley & D. L. Martin. 1987. Cholera from raw oysters shipped interstate. J. Amer. Med. Assoc. 258:2374.

Pease, H. D. 1911. Problems in the sanitary handling of opened oysters. Fishing Gazette 28:865-867.

Pequegnat, J. E., S. W. Fowler & L. F. Small. 1969. Estimates of the zinc requirements of marine organisms. J. Fish. Res. Bd. Canada 26:145-150.

Perkins. F. O., D. S. Haven, R. Morales-Alamo & M. W. Rhodes. 1980. Uptake and elimination of bacteria in shellfish. J. Food Prot. 43:124-126.

Peters, R. & D. Raftos. 2002. The effects of Marteilia sydneyi on the host defense responses of the Sydney rock oyster, Saccostrea glomerata. J. Shellfish Res. 21:437.

Phelps, H. L. & E. W. Hetzel. 1987. Oyster size, age and copper and zinc accumulation. J. Shellfish Res. 6:67-70.

Phillips, A. H. 1917. Analytical search for metals in Tortugas marine organisms (IV). In: Papers from the Department of Marine Biology of the Carnegie Institution of Washington. DC, vol. 10. pp. 89-93.

Pirie, B. J. S., S. G. George, D. G. Lytton & J. D. Thomson. 1984. Metal-containing blood cells of oysters: ultrastructure, histochemistry and x-ray analysis. J. Mar. Biol. Assoc. U. K. 64:115-123.

Potts, W. T. W. 1967. Excretion in the molluscs. Biol. Rev 42:1-41.

Preston, A. 1966. Concentration of [sup.65]Zn in the flesh of oysters related to the discharge of cooling pond effluent from the C.E.G.B Nuclear Power Station at Bradwell-on-Sea, Essex. In: B. Aberg & F. P. Hungate. editors. Radioecological concentration processes. Oxford. England: Pergamon Press. pp. 995-1004.

Preston, A. 1968. The control of radioactive pollution in a North Sea oyster fishery. Helgol. Meeresunters. 17:269-279.

Pringle, B. H., D. E. Hissong, E. L. Katz & S. T. Mulawka. 1968. Trace metal accumulation by estuarine mollusks. Proc. Amer. Soc. Civil Eng. J. Sanitary Engineering Division 94:455-475.

Prytherch, H. F. 1934. The role of copper in the setting, metamorphosis, and distribution of the American oyster, Ostrea virginica. Ecol. Monogr. 4(1):47-107.

Purchon, R. D. 1968. The biology of the mollusca. London: Pergamon Press.

Ranson, G. 1924. Note preliminaire sur le verdissement des huitres. Bull. Soc. Zool. France 49:512.

Ranson, G. 1925. La verdissement des huitres. C. R. Soc. Biol. 93:1100.

Ranson, G. 1936. Sur quelques maladies des huitres. Rev. Path. Comp. et d'Hyg. Gen. No 1. 475:1-21.

Reidel, G. F., G. R. Abhe & J. G. Sanders. 1995. Silver and copper accumulation in two estuarine bivalves, the eastern oyster (Crassostrea virginica) and the hooked mussel (Ischadium recurvum), in the Patuxent River Estuary. Maryland. Estuaries 18:445-455.

Reidel, G. F., G. R. Abbe & J. G. Sanders. 1998. Temporal and spatial variations of trace metal concentrations in oysters from the Patuxent River, Maryland. Estuaries 21:423-434.

Renwrantz, L. R., T. P. Yoshino, T. C. Cheng & K. R. Auld. 1979. Size determination of hemocytes from the American oyster, Crassostrea virginica, and the description of a phagocytosis mechanism. Jahrb. Zool. Abt. Physiol. Zoomorph. 83:1-12.

Richards, G. P. 1988. Microbial purification of shellfish: a review of depuration and relaying. J. Food Prot. 51:218-251.

Ringwood, A. H., D. E. Conners & A. Di Novo. 1998. The effects of copper exposures on cellular responses in oysters. Mar. Environ. Res. 46:1-5.

Rodrick, G. E. & S. A. Ulrich. 1984. Microscopic studies on the hemocytes of bivalves and their phagocytic interaction with selected bacteria. Helgol. Meeresunters. 37:167-176.

Roesijadi, G. 1994. Behavior of metallothionein-bound metals in a natural population of an estuarine mollusc. Mar. Environ. Res. 38:147-168.

Roesijadi, G. 1996. Environmental factors: response to metals. In: V. S. Kennedy. R. I. E. Newell. & A. F. Eble, editors. The Eastern Oyster Crassostrea virginica. College Park, Maryland: Maryland Sea Grant College Program. University of Maryland system, pp. 515-537.

Romeril, M. G. 1971. The uptake and distribution of [sup.65]Zn in oysters. Mar. Biol. 9:347-354.

Roosenburg, W. H. 1969. Greening and copper accumulation in the American oyster. Crassostrea virginica, in the vicinity of a steam electric generating station. Chesapeake Sci. 10:241-252.

Ruddell, C. L. 1971. Elucidation of the nature and function of the granular oyster amebocytes through histochemical studies of normal and traumatized oyster tissues. Histochemie 26:98-112.

Ruddell, C. L. & D. W. Rains. 1975. The relationship between zinc, copper and the basophils of two crassostreid oysters, C. gigas and C. virginica. Camp. Biochem. Physiol. 51A:585-591.

Ryder, J. A. 1882. Notes on the breeding, food, and green color of the oyster. Bull. US Fish Comm. (also cited as Bull. Bur. Fish.),Vol. 1 for 1881. Washington, DC: Government Printing Office. pp. 403-419.

Sami, S., M. Faisal & R. J. Huggett. 1992. Alterations in cytometric characteristics of hemocytes from the American oyster Crassostrea virginica exposed to a polycyclic aromatic hydrocarbon (PAH) contaminated environment. Mar. Biol. 113:247-252.

Sami, S., M. Faisal & R. J. Huggett. 1993. Effects of laboratory exposure to sediments contaminated with polycyclic aromatic hydrocarbons on the hemocytes of the American oyster Crassostrea virginica. Mar. Environ. Res. 35:131-135.

Sanders, J. G., G. F. Riedel & G. R. Abbe. 1991. Factors controlling the spatial and temporal variability of trace metal concentrations in Crassostrea virginica (Gmelin). In: M. Elliot & J. P. Ducrotoy, editors. Estuaries and coasts: Spatial and temporal intercomparisons. Virborg, Denmark: Olson and Olson Publishers. pp. 335-339.

Sanderson, C. J. 1982. Morphological aspects of lymphocyte mediated cytotoxicity. In: W. R. Clark & P. Golstein, editors. Mechanisms of cell-mediated cytotoxicity. New York: Plenum. pp. 3-26.

Scro, R. A. & S. E. Ford. 1990. An electron microscope study of disease progression in the oyster, Crassostrea virginica, infected with the protozoan parasite, Haplosporidium nelsoni (MSX). In: F. O. Perkins & T. C. Cheng, editors. Pathology in marine science. New York: Academic Press Inc. pp. 229-254.

Severy, H. W. 1923. The occurrence of copper and zinc in certain marine animals. J. Biol. Chem. 55:79-92.

Sherman, A. R. 1992. Zinc, copper and iron nutriture and immunity. J. Nutrition 122:604-609.

Shuster, C. N., Jr. & B. H. Pringle. 1969. Trace metal accumulation by the American eastern oyster, Crassostrea virginica, Proc. Natl. Shellfish. Assoc. 59:91-103.

Simkiss, K. & A. Z. Mason. 1983. Metal ions: metabolic and toxic effects. In: The mollusca Vol. 2 environmental biochemistry and physiology. New York: Academic Press. pp. 101-163.

Simkiss, K., M. Taylor & A. Z. Mason. 1982. Metal detoxification and bioaccumulation in molluscs. Mar. Biol. Letters 3:187-201.

Soderhall, K. 1982. The prophenoloxidase activating system and melanisation: a recognition mechanism of arthropods? A review. Dev. Camp. Immunol. 6:601-611.

Soderhall, K., L. Cerenius & M. W. Johansson. 1994. The prophenoloxidase activating system and its role in invertebrate defense. Ann. N. Y. Acad. Sci. 712:155-161.

Soniat, T. M. 1996. Epizootiology of Perkinsus marinus disease of eastern oysters in the Gulf of Mexico. J. Shellfish Res. 15:35-43.

Soniat, T. M. & M. S. Brody. 1988. Field validation of a habitat suitability index model for the American Oyster. Estuaries 11:87-95.

Sprague, V. E., A. Dunnington & E. Drobeck. 1969. Decrease in incidence of Minchinia nelsoni in oysters accompanying reduction of salinity in the laboratory. Proc. Nat. Shellfish. Assoc. 59:23-26.

Stauber, L. A. 1950. The fate of India ink injected intracardially into the oyster Ostrea virginica Gmelin. Biol. Bull. 98:227-241.

Stauber, L. A. 1961. Immunity of invertebrates, with special reference to the oyster. Proc. Nat. Shellfish. Assoc. 50:7-20.

Sunila, I. & J. LaBanca. 2003. Apoptosis in the pathogenesis of infectious diseases of the eastern oyster Crassostrea virginica. Dis. Aquat. Org. 56:163-170.

Takatsuki, S. 1934. On the nature and functions of the amebocytes of Ostrea edulis. Qtr. J. Microsc. Sci. 76:379-431.

Tamplin, M. L. & G. Capers. 1992. Persistence of Vibrio vulnificus in tissues of Gulf Coast oysters, Crassostrea virginica, exposed to seawater disinfected with UV light. Appl. Environ. Microbiol. 58:15061510.

Thiselton Dyer, W. T. 1877. Greening of oysters. Nature. September 6, 1877, pp. 397.

Thomson, J. D., B. J. S. Pirie & S. G. George. 1985. Cellular metal distribution in the Pacific oyster, Crassostrea gigas (Thun.) determined by quantitative X-ray microprobe analysis. J. Exper. Mar. Biol. Ecol. 85:37-45.

Thorpe, T. E. 1896. Oyster culture in relation to disease, Nature 55:105-107.

Tripp, M. R. 1958. Disposal by the oyster of intracardially injected red blood cells of vertebrates. Proc. Nat. Shellfish. Assoc. 48:143-147.

Tripp, M. R. 1960. Mechanisms of removal of injected microorganisms from the American oyster, Crassostrea virginica (Gmelin). Biol. Bull. 119:273-282.

Tripp, M. R. 1970. Defense mechanisms of mollusks. J. Reticuloendoth. Soc. 7:173-182.

Tripp, M. R. & V. E. Kent. 1967. Studies on oyster cellular immunity. In Vitro 3:129-135.

Vinogradov, A. P. 1953. Chemical composition of marine organisms. Memoir Sears found. Mar. Res. Report II, pp. 339-358.

Volety, A. K. & W. S. Fisher. 2000. In vitro killing of Perkinsus marinus by hemocytes of oysters Crassostrea virginica. J. Shellfish Res. 19: 827-834.

Volety, A. K., S. A. McCarthy, B. D. Tall, S. K. Curtis, W. S. Fisher & F. J. Genthner. 2001. Responses of oyster (Crassostrea virginica) hemocytes to nonpathogenic and fecal isolates of Vibrio parahaemolyticus. Aquat. Microb. Ecol. 25:11-20.

Volety, A. K., L. M. Oliver, F. J. Genthner & W. S. Fisher. 1999. A rapid tetrazolium dye reduction assay to assess the bactericidal activity of oyster (Crassostrea virginica) hemocytes against Vibrio parahaemolyticus. Aquaculture 172:205-222.

Wagge, L. E. 1955. Amoebocytes. Internat. Rev. Cytol. 4:31-78.

Watson, D. G., J. J. Davis & W. C. Hanson. 1961. Zinc-65 in marine organisms along the Oregon and Washington coasts. Science 133: 1826-1828.

Wells, H. W. 1961. The fauna of oyster beds, with special reference to the salinity factor. Ecol. Monogr. 31:239-266.

Winstead, J. T. & J. A. Couch. 1988. Enhancement of protozoan pathogen Perkinsus marinus infections in American oysters Crassostrea virginica exposed to the chemical carcinogen n-nitrosodiethylamine (DENA). Dis. Aquat. Org. 5:205-213.

Wolfe, D. A. 1970a. Levels of stable Zn and [sup.65]Zn in Crassostrea virginica from North Carolina. J. Fish. Res. Bd. Canada 27:47-57.

Wolfe, D. A. 1970b. Zinc enzymes in Crassotrea virginca. J. Fish. Res. Bd. Canada 37:59-69.

Wright, D. A., J. A. Mihursky & H. L. Phelps. 1985. Trace metals in Chesapeake Bay oysters: intra-sample variability and its implications tar biomonitoring. Mar. Environ. Res. 16:181-197.

Wright, D. A. & C. D. Zamuda. 1987. Copper accumulation by two bivalve molluscs: salinity effect is independent of cupric ion activity.

Yonge, C. M. 1926, Structure and physiology of the organs of feeding and digestion in Ostrea edulis, J. Mar. Biol. Assoc. 14:295-386.

Yonge, C. M. 1928. The absorption of glucose by Ostrea edulis. J. Mar. Biol. Assoc. U. K. 15:643-653.

Yonge, C. M. 1937. Evolution and adaptation in the digestive system of the Metazoa. Biol. Rev. 12:87-115.

Yonge, C. M. 1946. Digestion of animals by lamellibranchs. Nature 157: 729.

Yoshino, T. P. & T. C. Cheng. 1976. Fine structural localization of acid phosphatase in granulocytes of the pelecypod Mercenaria mercenaria. Trans. Amer. Microscop. Soc. 95:215-220.

Zamuda, C. D. & W. G. Sunda. 1982. Bioavailability of dissolved copper to the American oyster, Crassostrea virginica. I. Importance of chemical speciation. Mar. Biol. 66:77-82.

Zaroogian, G. E. 1979. Studies on the depuration of cadmium and copper by the American oyster Crassostrea virginica. Bull. Environm. Contam. Toxicol. 23:117-122.

WILLIAM S. FISHER

United States Environmental Protection Agency, Office of Research and Development, National Health and Environmental Effects Research Laboratory, Gulf Ecology Division, 1 Sabine Island Drive, Gulf Breeze, FL 32561

E-mail: [e-mail]