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Nonnative Oysters in the Chesapeake Bay 4 Oyster Biology GENERAL BIOLOGY OF OYSTERS Oysters are members of the family Ostreacea, class Bivalvia, in the phylum Mollusca. Under the current systematic schema, most commercially important species are classified in three major genera: Ostrea, Saccostrea, and Crassostrea and a number of minor genera (Carriker and Gaffney, 1996). Adults are intertidal and subtidal bottom dwellers found worldwide. Most oyster species form the basis of local fisheries or aquaculture operations. Oysters differ from other bivalves in having a highly irregular shell form. The shape of the shell is typically dictated by environmental constraints, and they are capable of growing over or around adjacent objects, including other oysters. Oysters are plankton feeders; they use their gills to filter microalgae and probably bacteria. During feeding, they relax their single adductor muscle, allowing the two valves of the shell to open slightly. In an action called “pumping,” specialized cilia on the gill draw water into the shell cavity (Newell and Langdon, 1996). Other gill cilia trap particles and funnel them toward the palps—large liplike structures, also covered with cilia that surround the mouth and on which particles are sorted. Some particles, such as microalgae, are sent into the mouth; others, such as sediment, are usually rejected and deposited as “pseudofeces” just outside the shell. Filtration rates are a function of several environmental factors, including temperature, salinity, and suspended particulate concentration. Rates increase with size, although per unit weight, small oysters filter more water than do large individuals
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Nonnative Oysters in the Chesapeake Bay (Shumway, 1996). Powell et al. (1992) reviewed the literature on filtration rates for numerous marine bivalves and found that the relationship between filtration rate and size was similar for all species examined, including several oysters. Oysters do not regulate their body temperature or the salinity of their body fluids; thus, their metabolic activity is closely tied to the temperature of their surroundings, and the salt content of their blood is the same as that of the ambient water (Shumway, 1996). The ability of oysters to tolerate different environments is species specific. For instance, the European oyster, Ostrea edulis, grows in relatively cool, clear, water of high salinity (Yonge and Thompson, 1976). Crassostrea species, in contrast, are more typically inhabitants of estuaries in which they tolerate wide fluctuations in temperature, salinity, and turbidity. The oyster’s energetic investment in reproduction is prodigious, with individual females capable of producing many millions of eggs. Oysters typically become reproductively mature as males and may become female in subsequent seasons. Reproductive activity is seasonal and in temperate regions is generally dictated by temperature. Spawning occurs predominantly during the warm season, although other factors, such as phytoplankton blooms, may also play a role. Members of the genus Crassostrea shed their gametes directly into the water where fertilization occurs, and larval life is spent entirely in the water column. In contrast, fertilization and partial larval development in Ostrea take place in the interior of the oyster’s shell. Females release eggs within the shell cavity, and fertilization occurs when sperm shed by nearby male oysters get drawn into the female cavity. The larvae develop partially among the female’s gill filaments, which turn dark and become gritty as the larvae produce shells and become pigmented. The female’s unpleasant appearance and texture at this time are the principal reason that eating Ostrea species is avoided in the summer (months without “R,” or May through August) when reproduction occurs. The larvae of Ostrea species are eventually expelled from the female’s shell cavity and complete their development in the water column. Oysters that are brooders produce smaller numbers of offspring than nonbrooders. The waterborne larval stage of oysters allows them to disperse from the immediate site of the parental stock, enhances genetic mixing, and allows the colonization of new locations. The larvae are both dispersed and concentrated by water currents and wind. At the end of the larval life, usually 2 to 3 weeks, the oysters “set.” Unlike clams, which can settle into mud and can shift around as adults, oyster larvae cement themselves to a clean, hard substrate and lose their mobility (Yonge and Thompson, 1976). The substrate may be another oyster, a piece of shell, a pebble, a tree root, or any other solid, clean surface. The concentrating effect of wind and
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Nonnative Oysters in the Chesapeake Bay currents, and the fact that larvae prefer to settle where there are other oysters, results in large assemblages on suitable substrates. The mangrove oyster, C. rhizophorae, for instance, congregates on the roots of mangrove trees in shallow water. Species that inhabit deeper water tend to form aggregates known as “reefs” or “beds.” The Eastern oyster, C. virginica, is particularly well known for the large, three-dimensional reefs that it builds as successive generations of oysters settle on each other. DISEASES OF OYSTERS Because the terminology is often confused or confusing, this discussion of oyster diseases begins with definitions of key terms. Disease may be caused by an infectious agent or by other factors such as poor diet, exposure to a harmful substance, or a genetic defect. Infection and disease are not synonymous. Infection refers to the establishment of a foreign organism (infectious agent or parasite) in the tissues of another organism, called the host. Disease indicates damage to a body part, organ, or system such that the affected organism no longer functions normally. Infection does not necessarily lead to disease. Many infectious agents cause localized tissue damage but relatively little overall harm to their hosts. Infectious agents capable of causing disease are termed pathogens. Some pathogens are so virulent that they cause disease and mortality in susceptible hosts regardless of the physiological state of the host. Examples include Haplosporidium nelsoni and Perkinsus marinus (the disease agents), which cause MSX and Dermo diseases, respectively, in the Eastern oyster, C. virginica (the host). Other pathogens are described as opportunistic. Host organisms that are otherwise “healthy” can prevent infection by, or control proliferation of, opportunistic pathogens through structural (e.g., shell or epithelial barriers) or biological (physiological activity or the internal defense system) mechanisms. Opportunistic pathogens, however, may proliferate and cause disease if the host is compromised in some manner so that it can no longer effectively defend itself or if the number of opportunistic pathogens in the environment is large enough to overwhelm host defenses. Examples are the various bacterial and fungal species that infect and cause mortalities of cultured molluscan larvae and juveniles (Elston, 1984). Similarly, the herpes viruses associated with mortalities of larval and juvenile stages of a number of molluscan species in commercial hatcheries and nurseries are thought to be promoted by culture conditions, especially high temperature and high density (Farley et al., 1972; LeDeuff et al., 1996; Arzul et al., 2001). For disease to occur, a potential pathogen must find a susceptible host in a favorable environment. A parasite may infect one species without causing apparent harm but can cause catastrophic disease outbreaks when
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Nonnative Oysters in the Chesapeake Bay it infects another species. The “other species” in such a case can be a resident host infected by an introduced pathogen or an introduced host infected by a resident pathogen. Pathogens may also be present in an environment that inhibits their proliferation. Under these conditions they remain undetectable, either by causing no observable effect (such as death of the host) or because they are too few to be found by standard diagnostic assays. Climate warming, for instance, is hypothesized to have favored outbreaks of Dermo disease from existing undetected foci of P. marinusinfected oysters in the northeastern United States and thus resulted in the apparent range extension of P. marinus (Ford, 1996; Cook et al., 1998). While all commercial molluscan species examined so far are infected by some parasites, oysters have more reported lethal diseases than any other commercial species (Bower et al., 1994; Ford, 2001; see Table 4.1). As a matter of fact, the molluscan diseases listed as “of concern” by the Office International des Epizooties, an international veterinary body concerned with animal health, are primarily those affecting oysters of various species and are all caused by water-borne protozoan parasites that invade through the gut or external epithelium and proliferate inside the soft tissues, killing the oyster when the parasite burden becomes high. Transmission of some parasites, such as P. marinus and Bonamia ostreae (cause of the disease bonamiasis in Ostrea edulis), is directly from oyster to oyster. The mode of transmission, and indeed the complete life cycle of others, such as H. nelsoni and Marteilia refringens (cause of the disease marteiliosis in O. edulis), remains unknown, although a recent study provides evidence of the involvement of a copepod in the life cycle of M. refringens (Audemard et al., 2002). Oyster “mass” mortalities have been recorded at least since the early 1900s. Those not attributable to predation, siltation, or freshwater influxes were simply ascribed to unknown causes (Orton, 1924; Roughley, 1926; Sindermann and Rosenfield, 1968), although one such case was later ascribed to a pathogen (Farley et al., 1988). Another early disease outbreak, which killed large numbers of C. virginica in Prince Edward Island, Canada, in 1913 to 1915, has been attributed to an infectious agent (Needler and Logie, 1947). The disease agent is still present but has yet to be identified. Not until the discovery of P. marinus and H. nelsoni in the late 1940s and late 1950s, respectively, were specific infectious agents clearly identified as the cause of any bivalve mortality. Shortly thereafter, pathogens were associated with catastrophic mortalities of two oyster species (C. angulata and O. edulis) in France. A virus identified in the gills of C. angulata was thought to be the cause of at least some of the mortalities that wiped out commercial production of this species in France in the 1970s (Comps, 1988). The loss of C. angulata prompted the importation of
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Nonnative Oysters in the Chesapeake Bay TABLE 4.1 Important or Common Parasites and Diseases of Oysters Disease/Condition Causative agent Host(s) Region affected Comments Herpes virus disease outbreaks Herpes virus Numerous bivalve species including oysters Worldwide Typically found associated with mortalities of larvae and juveniles in commercial culture; has been found in adults and in wild larvae and juveniles, but without observed mortality. Juvenile Oyster Disease (JOD) Probably bacterial Crassostrea virginica juveniles grown in culture Northeastern United States Causative agent unknown, but transmissible; probably has bacterial cause, but may also involve other factors. Caused mortalities from New York to Maine during the 1990s. Problem subsided in most regions in late 1990s. Summer Mortality Vibrio splendidus bacterium (and various other factors) C. gigas France Associated with mortality of juveniles, but adults also suffer. Probably has various causes. Nocardiosis Nodardia crassostreae C. gigas West Coast of Unitred States Associated with summer mortalities. Maladie du pied Ostracoblabe implexa (fungus) O. edulis, C. gigas, Saccostrea cucullata Europe, Canada, India Fungus grows in shells causing “wart-like” protuberances on inner shell. May weaken oyster and diminish marketability.
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Nonnative Oysters in the Chesapeake Bay European oyster haplosporidiosis H. armoricanum (protozoan) O. edulis and O. angasi Northern Europe Very low prevalence and no significant impact on population. Marteiliosis (Aber Disease) M. refringens (protozoan) O. edulis O. angasi, Tiostrea chilensis (=T. lutaria) Western Europe In Europe, causes epizootic mortalities in O. edulis; other species have proved susceptible when challenged experimentally, but are not known to be affected in their native ranged. QX Disease M. sydneyi (protozoan) Saccostrea glomerata (=S. commercialis) Australia Causes epizootic mortalities. Dermo Disease P. narinus (protozoan) C. virginica, C. gigas, C. ariakensis East and Gulf Coast of United States Causes epizootic mortalities in C. virginica. C. gigas and C. ariakensis become infected, but do not develop lethal infections. Bonamiosis B. ostrea (protozoan) O. edulis (European oyster) and other species Ostrea: O. angasi, O. denselamellosa, O. puelchana, Ostreola conchaphila (= O. lurida), and Tiostrea chilensis (= T. lutaria) Western Europe; Maine, United States; Northwestern United States In Europe, causes epizootic mortalities in; O. edulis; other species have proved susceptible when challenged experimentally, but are not known to be affected in their native ranges. Not known to caused mortalities in O. edulis in the United States. C. ariakensis(?)
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Nonnative Oysters in the Chesapeake Bay Disease/Condition Causative agent Host(s) Region affected Comments Australian Winter Disease Mikrocytos roughleyi (protozoan) S. glomerata Australia Mortalities caused by M. roughleyi apparently first reported in 1926. Denman Island Disease Mikrocytos mackini (protozoan) C. gigas British Colombia, Canada Can be controlled through appropriate aquaculture practices. MSX Disease Haplosporidium nelsoni (protozoan) C. virginica, C. gigas East Coast of the United States (C. virginica) Pacific Asia and United States (C. gigas) Causes epizootic mortalities in C. virginica. C. gigas becomes infected but no mortalities reported. Also found in C. gigas in California. SSO Disease H. costale (protozoan) C. virginica East Coast of the United States Restricted to higher salinity locations compared to H. nelsoni, Malpeque Disease Unknown C. virginica Atlantic Canada First outbreak in 1915-16; oysters in affected areas appear to have developed resistance. Hemic neoplasia (uncontrolled proliferation of blood cells) Etiology unknown, but reported to have genetic or environmental links Many species of oysters and other marine bivalves Widespread Contagious; may be associated with mortality.
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Nonnative Oysters in the Chesapeake Bay Summer mortality Various C. gigas West Coast of United States Japan Probably a multifactorial cause. Infection by Rickettsiales- or Chlamydiales-like organisms Intracellular bacteria-like All species of oysters examined and most other marine bivalves also Global Has been associated with mortality of marine bivalves, but not necessarily as the causative agent—more probably opportunist. Found more frequently in dense associations of bivalves (e.g., nurseries or culture parks). Other parasites Trematodespestodes, nematodes, ciliates, and flagellates All species of oysters examined and most other marine bivalves also Global Not deleterious. SOURCE: Susan Ford, Haskin Shellfish Research Laboratory, Rutger University, Port Norris, New Jersey.
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Nonnative Oysters in the Chesapeake Bay C. gigas as a replacement species (Grizel and Héral, 1991). The C. angulata mortalities were followed by disease outbreaks in the European oyster, O. edulis, caused by two newly discovered protozoan pathogens, M. refringens and B. ostreae. The resulting mortalities caused a precipitous decline in O. edulis production in France in the 1970s, and accelerated the use of C. gigas, which is not susceptible to the diseases caused by these two pathogens or the viral “gill disease.” Infections of marine molluscs by other agents, including parasitic worms, protozoans, Rickettsiales- and Chlamydiales-like organisms (RLOs and CLOs), bacteria, and viruses are not uncommon (Bower et al., 1994), and “new” cases continue to be described as more and more host species are grown in culture and examined by an ever-increasing number of scientists. Some are found when mortalities, often of cultured molluscs, are investigated. Culture conditions, in which the molluscs are grown at high density and often using poor animal husbandry practices, favor the proliferation and transmission of opportunistic pathogens, which can then cause or exacerbate disease and mortality in the cultured organisms (Meyers, 1979; Elston, 1984; Bower, 1987; Bricelj et al., 1992; Lacoste et al., 2001). Various bacterial species and the herpes virus are examples of pathogens most commonly associated with disease outbreaks in hatcheries and nurseries (Hine et al., 1998; Renault et al., 2000, 2001; Arzul et al., 2001). Others are encountered during routine surveys or health examinations required for the shipment of molluscs across governmental boundaries. Most occur at low prevalence and intensity and appear to cause no harm to the host. For instance, certain microorganisms, such as the intracellular bacterialike RLOs and CLOs, have been found in all bivalves examined so far, typically without evidence of being harmful. They have often been associated with mortality (Gulka and Chang, 1984; Le Gall et al., 1988; Norton et al., 1993; Villalba et al., 1999; Moore et al., 2000), although it is probable that they are opportunistic pathogens rather than the original cause of death. Although not exhaustive, Table 4.1 lists the most important or common diseases and parasites reported for oysters, along with the host species and regions where they are found. CRASSOSTREA VIRGINICA Life History C. virginica, the Eastern oyster, inhabits estuarine waters from the Canadian maritime provinces to the Gulf of Mexico, with reports of the species as far south as Brazil and Argentina (Carriker and Gaffney, 1996). Adults are intertidal and subtidal dwellers, typically found in assem-
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Nonnative Oysters in the Chesapeake Bay blages called reefs, bars, or beds that range in size from a few acres to hundreds. The general morphological, physiological, and life history characteristics of oysters described earlier apply equally to C. virginica. This section provides more detailed characteristics of C. virginica. Reproduction of C. virginica is seasonal and largely influenced by temperature. Gametogenesis begins in the spring and spawning occurs from late May to late September in the mid-Atlantic, with the season contracted or extended to the north and south, respectively (Shumway, 1996; Thompson et al., 1996). C. virginica are either male or female (the reported incidence of hermaphroditism is <0.5%) but may change sex over the winter when they are reproductively inactive. Small oysters (10 to 20 mm) sometimes develop gametes, almost always sperm. Under favorable growth conditions in the mid-Atlantic, this may occur during the late summer after setting, although it is uncertain whether such individuals actually spawn or produce embryos because they do not ripen until after the normal spawning period. In the southeastern United States and the Gulf of Mexico, sexual maturity is typically reached about 3 months after setting, and the prolonged reproductive period in this region increases the probability that these juveniles do participate in the overall reproductive effort of the population. Males are more sensitive to spawning stimuli, such as temperature and food, than females and tend to spawn first. The presence of sperm in the water stimulates females to release eggs, which are then fertilized externally. Gametes deteriorate within a few hours of spawning and can be rapidly diluted by water currents; thus, the proximity of oysters to one another increases the chances of synchronous spawning and successful fertilization. In the first 24 hours, oyster larvae develop a large ciliated structure, the velum, which acts as both a swimming and food-gathering organ. Initially, the shell is secreted as a single event at about 24 to 48 hours; thereafter, growth occurs through accretion to both thicken and extend the shell. The larval stage lasts for about 2 to 3 weeks, depending on food availability and temperature. Larvae appear to migrate vertically, particularly at later stages, tending to concentrate near the bottom during the outgoing tide and rising in the water column during the incoming tide, thus increasing their chance of being retained in the estuary (Kennedy, 1996; Shumway, 1996). Larval mortality rates are estimated to be close to 99%. As is the case with all oyster species, C. virginica larvae must eventually find a clean, solid surface on which to cement themselves. Oyster shells meet those criteria if they are not covered with silt or heavily fouled by other epifaunal organisms (although the larvae settle on any type of hard substrate, such as pilings, rocks, and ship bottoms). The suitability of oyster shell for setting, the concentrating mechanisms of wind and
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Nonnative Oysters in the Chesapeake Bay water currents, and the gregarious nature of setting all lead to the formation and persistence of oyster reefs. Without continuous setting and growth of juveniles on the reef, it will usually become covered with silt. Certain sites within estuaries are known to reliably obtain good “sets” of young oysters. These are locations where clean shell is often spread by management agencies or industry members to “catch” the set, after which it is typically moved to grow-out areas. Despite the knowledge of where the larvae consistently settle, the parental stock for these sets is rarely, if ever, known. The potential obviously exists for oysters to be carried long distances during their larval life, both within and between estuaries; the current state of knowledge is insufficient to predict where larvae originating from oysters in a particular area will be transported or to estimate the likelihood that larvae from one estuary will be carried, along the coast, to another estuary. Oyster larvae are common in summer in water samples collected in East Coast estuaries (Kennedy, 1996); however, investigators sampling nearshore waters off New Jersey for surf clam larvae report seeing only one or two Crassostrea sp.-like larvae during several years of sampling in the 1970s and 1990s (M. Tarnowski, Maryland Department of Natural Resources, Annapolis, personal communication, 2003; J. Grassle, Rutgers University, Port Norris, personal communication, 2003). Further, there is little evidence for a broodstock size/recruitment relationship for oysters, and very large sets can occur even when the stock of oysters is very low, as occurred in Delaware Bay several years after the MSX disease epizootic and heavy oyster drill predation had severely reduced the Chesapeake Bay population (Fegley et al., 1994). For the Gulf of Mexico, Livingston et al. (1999) reported rapid repopulation of Apalachicola Bay after the oyster population was decimated by two hurricanes in 1985. A widespread heavy set occurred in Chesapeake Bay in 1997 even though the oyster population was severely depleted. Unfortunately, most of the oysters suffered disease-caused mortalities before they reached marketable size. From a few hundred microns in size at the time of setting, C. virginica grow to sizes exceeding 150 mm. They are typically marketed in the United States when they reach about 75 mm (about 3 inches). Growth rates vary with temperature, food, turbidity, and salinity. In the mid-Atlantic, market-sized C. virginica are at least 2 years old but more typically are harvested when 3 to 4 years old. To the south, C. virginica may grow to marketable size in 12 to 15 months. Growth rates of oysters held in floating aquaculture trays are typically much greater than those of oysters on the bottom. The average life span is about 6 to 8 years; the maximum is probably about 25 years. Oysters provide food for numerous predatory species, including flatworms, crabs, oyster drills, starfish, and certain finfish. Mortality, mostly due to predation, is high on newly set oysters (spat), typically exceeding 40% in the first week (Haskin and Tweed, 1976; Newell et al., 2000) and
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Nonnative Oysters in the Chesapeake Bay CRASSOSTREA ARIAKENSIS Life History C. ariakensis is reported to be found along the entire Chinese coastline (Tschang and Tse-kong, 1956), southern Japan (Rao, 1987), Taiwan, the Philippines, Thailand, Vietnam, and northern Boreno and Malaysia (Zhou and Allen, 2003). There are also reports of C. ariakensis on the northwest coast of India and Pakistan. However, the distributional range outside China and southern Japan has not been genetically confirmed (Allen et al., 2002), and there is considerable taxonomic confusion about the species (Coan et al., 2000; Carriker and Gaffney 1996; Zhou and Allen, 2003). Sometimes C. ariakensis has been identified as Ostrea rivularis, C. rivularis, C. discoidea (e.g., Awati and Rai, 1931; Harry, 1985; Rao, 1987) or C. paulucciae (Carriker and Gaffney, 1996). As a result, biological information on C. ariakensis in its native distribution range is somewhat difficult to unravel because C. ariakensis may have been misidentified. C. ariakensis was inadvertently introduced to Oregon with shipments of C. gigas and C. sikamea spat from Japan in the 1970s (Breese and Malouf, 1977). Although C. ariakensis seed has been repeatedly outplanted on intertidal mudflats or suspended from floating rafts at several sites from Washington to central California (Breese and Malouf, 1977; Langdon and Robinson, 1996), there are no reports of established wild populations existing on the U.S. West Coast (Coan et al., 2000; J. T. Carlton, Williams College-Mystic Seaport Program, personal communication, 2003; R. Malouf, Oregon State University, personal communication, 2003). Apparently, seawater temperatures are not warm enough for the species to reproduce and maintain self-sustaining populations at these sites. C. ariakensis has limited aquaculture use in Washington and Oregon because of difficulties in obtaining sufficient quantities of seed for large-scale production (Langdon and Robinson, 1996). Field and laboratory experiments in Virginia (e.g., Calvo et al., 2001) and North Carolina have used the U.S. West Coast strain originally imported from Japan. In addition, scientists at VIMS have imported strains collected from the Yellow River estuary (northern China ariakensis) and from the Guangxi province near Beihai (southern China ariakensis) for use in comparative biological studies (S. K. Allen, Virginia Institute of Marine Science, personal communication, 2003). Laboratory work has also been conducted in France with the strain that was accidentally imported from Japan to the U.S. West Coast. No wild populations currently exist in France. While C. ariakensis has been extensively cultured throughout southern China and Japan for over 300 years (Cai et al., 1979), there is relatively little information on the ecology or biology of natural populations of the species in its native distributional range. In China the common
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Nonnative Oysters in the Chesapeake Bay name is Jinjiang-muli (meaning “close to river [Jinjiang] oysters [muli]”: Zhou and Allen, 2003). Indeed, Cai et al. (1992) reported that the oyster is cultured in estuarine mid- to shallow subtidal regions of Zhanjiang Bay where water temperatures range from 14 to 31.8oC and salinity varies from 7.5 to 30.2‰. Others report that the species is normally found in areas where the salinity varies between 9 and 30‰ (Cahn, 1950) and that it can tolerate salinities less than 10‰ for short periods of time (Amemiya, 1928). Guo et al. (1999) reported that the species can tolerate a wide salinity range, though settlement is most pronounced in low-salinity estuaries and river beds. Extensive Chinese aquaculture of C. ariakensis is typically found in intertidal and shallow subtidal estuarine habitats, where concrete stakes are transported to lower-salinity waters for spat collection and then moved to more saline areas for growout to marketable sizes (Guo et al., 1999). Similar to C. virginica, spawning activity in C. ariakensis is seasonal, and the reproductive cycle is influenced by temperature and regional environmental conditions. For China, Cai et al. (1992) reported that the reproductive season was between April and June and that larval settlement was most pronounced on the shady sides of hard surfaces. Moazzam and Rizvi (1983) reported that larval settlement in saltwater creeks in Pakistan was most pronounced from September through October, though recruitment was also observed during July to August. In a saltwater creek near Karachi, Pakistan, Ahmed et al. (1987) reported that larval settlement occurred from April to October, with highest spatfall recorded in April through July. In the Zhujiang River estuary the reproductive season is June to September and spawning is mainly in June to July; a second spawning may occur if environmental conditions are appropriate (Zhou and Allen, 2003). Asif (1979) reported that gonads generally were first present in individuals 2 to 3 months of age (0.4 to 0.6 cm in shell length) and that protandric hermaphrodites were found. Langdon and Robinson (1996) report that mature oysters collected from Yaquina Bay, Oregon, became available for hatchery spawning in early to mid-May, and female oysters had some percentage of mature eggs until December. From August to November, females were found with more that 50% mature eggs. There was a higher degree of annual variability in the gametogenic cycle of male oysters, though the highest percentages (>50%) of mature individuals were recorded from June to February. For Dabob Bay, Washington, Perdue and Erickson (1984) reported that gonadal proliferation began in late April to early June and the greatest proliferation occurred in mid-June to October. The authors reported that C. ariakensis failed to spawn, however, due to low summer water temperatures. Luckenbach (personal communication, Virginia Institute of Marine Science, Gloucester Point, 2003) has conducted prelimi-
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Nonnative Oysters in the Chesapeake Bay nary laboratory studies which suggest that C. ariakensis is capable of reproducing throughout the same salinity ranges as C. virginica (i.e., 5 TO 35‰). Modeling of larval salinity and temperature tolerances will be needed to predict the dispersal of C. ariakensis both within and beyond the Chesapeake Bay as has been done for other species (Lough, 1975; Goulletquer et al., 1994b). One concern regarding the potential introduction of C. ariakensis to areas with native populations of C. virginica is that both species may spawn at the same time and place, raising the possibility of interspecific hybridization. Laboratory experiments conducted by Allen et al. (1993) demonstrate that, although fertilization occurs in interspecific crosses, no viable progeny are produced. The larvae do not grow and begin to die a week after fertilization. Coan et al. (2000) reported that C. ariakensis can reach shell heights up to 20 cm, while Carriker and Gaffney (1996) noted that the oyster can be 20 to 24 cm high. Oysters growth relatively quickly in southern Chinese coastal waters and are typically harvested, when raised in extensive aquaculture conditions, within 2 to 3 years (10- to 15-cm shell length) after larval settlement (Guo et al., 1999). In U.S. waters, growth rates appears to vary with environmental conditions. For example, Langdon and Robinson (1996) reported no differences in the growth rate of C. ariakensis and C. gigas at sites in Puget Sound, Washington, and Coos Bay, Oregon, when planted on intertidal mudflats or when suspended from floating rafts in Yaquina Bay, Oregon. Both species attained shell heights of about 10 cm in roughly 2 years. C. gigas did grow faster than C. ariakensis when placed in mesh bags supported by intertidal trestle in Tomales Bay, California, (central California C. ariakensis grew to only about 6 cm in about 1.5 years, while C. gigas attained an average shell height of 10 cm). Calvo et al. (2001) grew triploid C. ariakensis in mesh bags in floating trays at six sites in the Virginia portion of Chesapeake Bay; two sites were located in low-salinity (<15‰), two in medium-salinity (15-25‰), and two in high-salinity (>25‰) waters. At deployment, oysters averaged 6.4 cm in shell height (2 years old), and growth was followed for about 9 months. It was found that shell growth was significantly slower (2.6 mm/month) in the low-salinity regime when compared to medium- (4.9 mm/month) and high-salinity waters (6.2 mm/month). A limited number of studies have examined the predators and competitors of C. ariakensis in its natural distribution range. Several species of carnivorous snails and crabs are reported to feed on adult oysters (Zhang and Lou, 1956), while sea urchins and sea stars consume oyster spat (Zhou and Allen, 2003). Barnacles have been noted to compete for food and space with settling oyster larvae (Cai et al., 1992). Calvo et al. (2001) reported that mortalities of oysters suspended in mesh bags above
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Nonnative Oysters in the Chesapeake Bay the bottom at several places in the lower Chesapeake Bay varied from 13 to 16% over a 9-month period, though the agent of mortality was not reported. The oysters were contained in mesh bags placed in suspended trays where benthic predators (e.g., crabs, snails) had reduced access to them. Moreover, the oyster leech (flatworm), Stylochus ellipticus, considered a significant predator on C. virginica, may affect C. ariakensis populations (Daniel et al., 1983; MacKenzie, 1997; Sagasti et al., 2001). Because C. ariakensis is similar in size and shell characteristics, it is likely that the same suite of benthic predators that prey on C. virginica in Chesapeake Bay would also forage on C. araikensis. Luckenbach (personal communication, Virginia Institute of Marine Science, Gloucester Point, 2003) thinks that the shell of C. ariakensis may not as be strong as C. virginica, which may suggest that the effective size refuge from crab predators is larger for C. ariakensis than for Eastern oyster. Investigations of the larval biology of C. ariakensis are restricted to laboratory studies, most of which were conducted in highly artificial conditions to assess optimal nutritional and environmental conditions for potential aquaculture purposes. Breese and Malouf (1977) reported that maximum larval growth was obtained at 28oC at both 20‰ and 30‰ salinity. The authors found that larval settlement was highest at 28oC and 20‰, with some settlement occurring at 28oC and 15‰ and 26oC and 20‰. Other temperature-salinity combinations yielded poor larval growth and no settlement. No larvae survived at 32oC. Langdon and Robinson (1996) found that larval settlement at salinities of 15 and 20‰ was greater than 25 and 30‰ and that no settlement occurred at 35‰. A similar pattern was found when examining larval growth rate. The authors found that growth of settled oyster spat over a 2-week period (at 20‰ salinity and on a diet of the diatom Chaetoceros calcitrans) was generally fastest at 25o to 30oC, though growth occurred at temperatures ranging from 20o to 35oC. Salinity and temperature conditions were not reported, but Cahn (1950) found that newly settled C. ariakensis could be found as early as June in Japan. Preliminary laboratory studies by Luckenbach (personal communication, Virginia Institute of Marine Science, Gloucester Point, 2003) revealed that swimming larvae of C. ariakensis tended to concentrate nearer the bottoms of culturing tubes while those of C. virginica were more commonly found nearer the surface of the containers. No field observations are available on C. ariakensis larval behavior and mortality rates. Like other oyster species, larvae of C. ariakensis must settle on solid surfaces. As mentioned earlier, suitable substrates for successful oyster settlement are those not heavily fouled by sediment or organisms (e.g., Osman et al., 1989). While Ahmed et al. (1987) reported that spatfall was primarily in the lower intertidal zone in Pakistan, a much
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Nonnative Oysters in the Chesapeake Bay broader settlement range (low tide to 10-m depth) has been reported in China (Nie, 1991). Larval settlement, 12 to 18 days after spawning (Zhou and Allen, 2003), varies primarily with temperature and other environmental conditions. In southern China, spatfall has been recorded from June to August (Cai and Li, 1990), while in Pakistan larval settlement has been observed from May to December (Moazzam and Rizvi, 1983; Hasan, 1960). There has been some uncertainty about the reef-building characteristics of C. ariakensis in different areas depending on substrate type. Yao (1988) reported finding an oyster reef primarily composed of fossil shells of the species in the tidal zone of Fujian, China. X. Guo (Graduate Institute of Environmental and Occupational Health, Medical College, National Cheng Kung University, personal communication, 2003) noted, “It is common knowledge among oyster (workers) in China that C. ariakensis is a reef builder. Areas [with] C. ariakensis reefs are north Shandong, Guangzi and Gunangdong [provinces].” There is a brief reference to reef building in the Bohai Sea cited in Wang et al. (1993, referred to by Guo). Lastly, there are some newspaper accounts of using historical reefs found in coastal low-salinity waters of the Bohai Sea in northern China to make concrete and apparently only C. ariakensis is found in this region, according to X. Guo. In Japan, however, the oyster apparently is restricted to lower intertidal muddy bottom habitats (Amemiya, 1928; Hirase, 1930). There are several reports that in India and Pakistan the oyster can be found on both hard substrates and muddy creek bottoms (Mahadevan, 1987; Patel and Jetani, 1991; Ahmed et al., 1987). Oyster filtration rates are typically related to environmental conditions (e.g., temperature, salinity, hypoxia) as well as the quantity and quality of suspended particulate materials (e.g., Higgins, 1980; Shumway et al., 1985; Riisgard, 1988). Zhang et al. (1959) examined the feeding biology of adult C. ariakensis relative to a number of environmental parameters, and concluded that feeding rate was highest when temperature and salinity were between 10 to 12oC and 15 to 30‰, respectively. While the oyster is known to be able to tolerate reduced salinity conditions, feeding rate was significantly reduced at less than 5‰ salinity. Zhang et al. (1959) also noted that feeding rate was not influenced by high levels of suspended material. In preliminary laboratory experiments conducted by R. Newell (Horn Point Laboratory, University of Maryland Center for Environmental Science, personal communication, 2002), size-specific filtration rates of C. ariakensis appeared similar to those of C. virginica. These results are consistent with Powell et al.’s (1992) conclusion that size-specific filtration rates are quite similar for most marine bivalve species.
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Nonnative Oysters in the Chesapeake Bay Disease and Disease Resistance Observed Diseases and Parasites in Native Range There is little information on disease outbreaks in C. ariakensis over its native range. Most C. ariakensis production appears to occur in China, which also encompasses a large portion of this species’ range. A recent publication describes recurrent large-scale mortalities (80 to 90% annually) recorded since 1992 in Guangdong province (Wu and Pan, 2000). The mortalities occur from February to May in 2- to 7-year-old oysters. The authors found heavy concentrations of what they considered to be a Rickettsiales-like organism in moribund oysters collected during one mortality episode. Rickettsiales-like organisms are common and ubiquitous in bivalves worldwide and are generally considered benign. In a few instances they have been associated with mass mortalities of other bivalves (Gulka and Chang, 1984; Gardner et al., 1995; Villalba et al., 1999), although it is not certain in any case whether these organisms are the original cause of the deaths or merely contributed to the deaths of otherwise stressed hosts. Further, the bodies identified by Wu and Pan as rickettsia have morphological and staining characteristics atypical of rickettsia and may have been misidentified. Although Wu and Pan do not mention it, toxic algal blooms are thought by others to cause the occasional large-scale mortalities of C. ariakensis in China (Zhang and Lou, 1956; Zhang et al., 1995). In a survey of molluscan aquaculture in China, Guo et al. (1999) noted reports of mass mortalities in scallops, abalone, and certain clams but none for oysters of any species. As part of the International Council for the Exploration of the Seas (ICES) protocol being employed in tests of C. ariakensis in Chesapeake Bay, the VIMS has examined tissue sections of 242 individual C. ariakensis from three separate stocks originating from the U.S. West Coast and north and south China. Cells characterized as “unusual” were found in three oysters from the northern China site. One of the oysters contained spores and plasmodia consistent in morphology with members of the Haplosporidia, the phylum containing H. nelsoni; cells in the other two oysters were not identifiable. Neither the haplosporidian infection nor the other cells were abundant in the affected oysters. VIMS is currently engaged in further testing of C. ariakensis. Samples were obtained from five sites along the Chinese coast in autumn 2002, and another set from the same sites will be collected in spring 2003, with further sampling planned if mortality is observed. Each sample includes 30 small and 30 market-sized cultured oysters and if present, 30 marketsized wild oysters, for a potential total of 900 individuals. The oysters are being examined by tissue slide histopathology for potential pathogens and pathological conditions, and, by polymerase chain reaction (PCR)
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Nonnative Oysters in the Chesapeake Bay using DNA primers specific for Perkinsus spp. and herpes virus. Using PCR, VIMS researchers have detected herpes virus in adult C. ariakensis collected in Japan (Itoki River, Ariaki Bay) but not in two spawns of C. ariakensis made in 2002 at the VIMS hatchery (K. Reece, Virginia Institute of Marine Science, personal communication, 2002). It should be noted that herpes virus is already present in the East Coast oyster population, as reported by Farley et al. (1972) who found it in C. virginica subjected to elevated temperatures in a power plant discharge pipe in Maine. French researchers at the Institut français de recherche pour l’exploitation de la mer La Tremblade Laboratory examined tissue sections of 15 female C. ariakensis from Hong Kong and found parasites in the egg cells of 7 individuals. The parasite could not be identified with the material available but resembles a Marteilioides-type parasite described as occurring in the egg cells of C. gigas in Japan (Comps et al., 1986). An “Exotic” Disease As a possible replacement for C. gigas in France, in the event the species began to have serious mortalities, French scientists imported C. ariakensis (from the Haskin Shellfish Research Laboratory in New Jersey), which they held in a “quarantine” system that treated outgoing, but not incoming, water. A sample was inspected histologically before the importation and found to have no detectable parasites, but after 7 months in the “quarantine” system, the oysters experienced high mortality. Tissues of moribund oysters were examined and found to be heavily parasitized by an organism that resembled, in all morphological and pathological details, B. ostrea, which infects Ostrea edulis, causing the disease “bonamiasis” in the same region (Cochennec et al., 1998). The authors “strongly suspected” that the C. ariakensis became infected through exposure to incoming untreated “waters…endemic for bonamiasis.” They pointed out that the parasite is the first report of a bonamia parasite in a species of Crassostrea; all other findings have been in Ostrea spp. Resistance to MSX and Dermo Diseases In several tests conducted since 1998, C. ariakensis has demonstrated rapid growth, high survival, and low infection levels after exposure to H. nelsoni and P. marinus in a number of locations in the Virginia portion of Chesapeake Bay and the coastal bays of Virginia’s Eastern Shore. The oyster has not been tested in Maryland, but the Virginia sites represented a variety of environments that spanned high-, medium-, and low-salinity regimes. The first trial, conducted by VIMS, extended over 18 months in 1998 and 1999. Mortality of C. ariakensis was about 15% compared to 80 to
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Nonnative Oysters in the Chesapeake Bay 100% of similar-sized lower Chesapeake Bay wildstock C. virginica (Calvo et al., 2001). Infection prevalence of H. nelsoni was low during the test, with a maximum in C. virginica of only 25%. No H. nelsoni was detected in any C. ariakensis. In contrast, both species developed high prevalences of P. marinus, with peaks of 68 to 84% in C. ariakensis and 100% in C. virginica. Despite the high P. marinus prevalences in C. ariakensis, all infections were categorized as “light” (unlikely to be lethal), whereas a substantial proportion of the infections in C. virginica became advanced and ultimately lethal. Subsequent trials of C. ariakensis, initiated in 2000 and 2001, were conducted by industry and did not include a disease diagnosis component; however, survival and growth of the species were reported to be extremely good. Within a year of the 2000 deployment, the C. ariakensis were of market size with little or no mortality, whereas half of the C. virginica, against which they were compared, were dead (Thompson, 2001). One concern about the 1998 to 1999 trial (Calvo et al., 2001) was that the C. virginica had been exposed to infection during the year before the trials began, which might have explained their higher infection levels compared to C. ariakensis. Infection prevalence and intensity, especially of P. marinus, typically increase with exposure in C. virginica. Although C. ariakensis did become infected with P. marinus, there was no consistent trend toward increased prevalence or intensity in this species over the 18-month trial, which included two summer infection periods. To investigate the possibility that longer exposure might increase parasite burdens in C. ariakensis, VIMS is now examining residual oysters from the 2000 and 2001 industry trials (E. Burreson, Virginia Institute of Marine Science, personal communication, 2002). Samples collected in June, September, and October 2002, after 2 to 3 years of exposure, showed 20 to 80% prevalence of P. marinus in the C. virginica, depending on test site, but no more than 20% (all very light infections comprising just a few parasites) in the C. ariakensis. No H. nelsoni have been detected in any C. ariakensis examined. A few instances of hemocytic infiltration into the digestive epithelium and lumena have been found, but no recognizable parasites (E. Burreson, Virginia Institute of Marine Science, personal communication, 2002). All evidence to date, then, indicates that even though C. ariakensis can become infected with P. marinus (and perhaps also with H. nelsoni, although no infections have yet been detected), infections do not develop to the point where they inhibit growth, fattening, and survival. If C. ariakensis actually does become infected with H. nelsoni, infections remain so light and localized that they have so far been undetectable by standard diagnostic methods. This result is similar to that demonstrated for C. gigas in earlier trials (Calvo et al., 1999). One difference, however, is that C.
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Nonnative Oysters in the Chesapeake Bay ariakensis performed well in low salinity, whereas C. gigas grew poorly and had high mortalities. Further, C. ariakensis, under the conditions of the test, did not develop the heavy mud blisters caused by the polychaete Polydora websteri that formed in C. gigas tested in Chesapeake Bay and were also reported in earlier trials in Louisiana (Kavanagh, 1940). More recently however, some 2-year-old C. ariakensis from the industry trials, which had been deployed at a low-salinity site, were observed to have a heavy infestation of Polydora blisters (S. Ford, Haskin Shellfish Research Laboratory, Rutgers University, personal communication, 2002). Infestation by Polydora is probably a site-dependent phenomenon, as the prevalence and intensity of blisters in the 2 to 3-year-old oysters from four sites being monitored by VIMS have not been noted as being heavier than those of C. virginica at the same locations (E. Burreson, Virginia Institute of Marine Science, personal communication, 2002). SUMMARY Oysters, like other bivalve filter feeders, feed on microalgae and are capable of removing large quantities of particulate matter from the water; they have a larval stage that allows them to be dispersed from the immediate site of the parental stock; and they require a clean, solid substrate on which the larvae can settle and cement themselves. C. virginica spawns during the summer, producing a larval stage that lasts for 2 to 3 weeks, during which time mortality is typically about 99%. Between settlement in mid-summer and winter, mortality of the “spat” due to predators may be nearly as high. Adults can tolerate a wide range of environments: subtidal and intertidal; salinities from <5 to 40 ppt; temperatures from freezing to 36oC, with short-term exposure to 50oC. They display genetically conserved, geographically distinct growth, reproduction, and disease susceptibility traits. Two important ecological functions have been attributed to oysters: habitat and resource provision and filtration of suspended particles from the water. The first function results from habitat created by oyster beds or reefs that provides refuge for the next generations of oysters and structure, refuge, and food resources for other benthic organisms and fish. The second ecological function is the removal of organic material, primarily phytoplankton, from the water and its transformation into inorganic nutrients, thus helping to reduce turbidity. Although these functions are often attributed to oysters, it is unclear how much impact oyster reef habitat and oyster filtering capacity have on the Chesapeake Bay ecosys-
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Nonnative Oysters in the Chesapeake Bay tem, especially given other stresses from sedimentation and nutrient overenrichment. Most of the arguments that the loss of oysters from the Chesapeake Bay has changed it from a “benthic”- to a “pelagic”- dominated system are based on very limited or anecdotal evidence. Oysters, like other organisms, become infected by some parasites that cause disease (damage severe enough to cause dysfunction of an organism) and by other parasites that cause little overall harm. The parasites responsible for MSX and Dermo diseases are so virulent that they cause disease in otherwise healthy oysters. Other parasites, including many bacteria and at least one virus (herpes), are “opportunists” that proliferate and cause disease in oysters that are stressed by other factors. Opportunistic parasites are most common in hatcheries and nurseries, where larval and juvenile bivalves are grown at high densities and elevated temperatures. Over the past two decades, above-average temperatures and repeated droughts have favored the parasites that cause MSX and Dermo. Not only have these conditions caused heavy oyster mortalities on existing reefs, they have impeded restoration efforts by causing mortalities on newly created reefs. Resistance to disease has been documented in wild populations of C. virginica after heavy selective mortality. Selective breeding has also produced C. virginica strains that are highly resistant to MSX disease and moderately resistant to Dermo disease; however, when tested under the same conditions, C. ariakensis shows a significantly higher degree of resistance (i.e., lower infection prevalence and intensity). The best C. virginica strains have shown 50 to 75% lower mortality than local (unselected) stocks, whereas C. ariakensis has shown 80 to 85% lower mortality. C. ariakensis is reported from Japan and Korea, south along the coasts of China, Malaysia, the Philippines, and westward to India and Pakistan, although genetic confirmation is lacking for most of this region and the species may have been misidentified in some cases. Relatively little biological and ecological data are available on C. ariakensis in its native range; however, available information indicates that C. ariakensis grows under environmental conditions very similar to those of C. virginica and has the same size-specific filtration rates. It is found intertidally and subtidally on reefs as well as on muddy bottom and is cultured in areas where the temperature ranges from 14o to 32oC and salinities from 7 to 30 ppt. The salinity range under which it reproduces is also similar to that for C. virginica, although it may require higher temperatures to induce spawning. Stocks imported to the West Coast of the United
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Nonnative Oysters in the Chesapeake Bay States in the 1970s have not naturalized, presumably because the temperature is too low for reproductive success. C. ariakensis grows faster than C. virginica but has a thinner shell, which may make it more susceptible to predators such as crabs; otherwise, it should be subject to the same predation pressures as C. virginica. Little information is presently available on parasites or diseases of C. ariakensis in its native range. Histological examinations of C. ariakensis at various institutes have found a few protozoan parasites in groups known to cause mortality in oysters, including a haplosporidian (MSX-like) and a Bonamia parasite, which was associated with mortality of C. ariakensis and was probably the same parasite that caused widespread mortalities of the European oyster. Other parasites, including a putative Rickettsiales-like organism and herpes virus, have been found worldwide in numerous marine bivalves.
Representative terms from entire chapter: