Seasonal comparison of physiological adaptation and growth of Suminoe (Crassostrea ariakensis) and eastern (Crassostrea virginica) oysters.
Crassostrea (Environmental aspects)
Oysters (Environmental aspects)
Kelly, Christopher J.
Laramore, Susan E.
Newell, Roger I.E.
|Publication:||Name: Journal of Shellfish Research Publisher: National Shellfisheries Association, Inc. Audience: Academic Format: Magazine/Journal Subject: Biological sciences; Zoology and wildlife conservation Copyright: COPYRIGHT 2011 National Shellfisheries Association, Inc. ISSN: 0730-8000|
|Issue:||Date: Dec, 2011 Source Volume: 30 Source Issue: 3|
|Topic:||Event Code: 310 Science & research Computer Subject: Company growth|
|Product:||Product Code: 0913050 Oysters NAICS Code: 114112 Shellfish Fishing SIC Code: 0913 Shellfish|
|Geographic:||Geographic Scope: United States Geographic Code: 1USA United States|
ABSTRACT Shell growth, survival, and physiology were compared
between diploid Suminoe (Crassostrea ariakensis) and eastern oysters
(Crassostrea virginica) under conditions simulating a U.S. subtropical
estuary. Two age groups (4 mo and 28 too) of both oyster species were
grown for 9 mo (December 2006 through August 2007) in quarantine
mesocosms (700 L) supplied with ambient flowing ([greater than or equal
to] 10 L/min) water (annual temperature range, 18.6-30.4[degrees]C;
salinity, 28-37.7). There was no difference in overall rates of shell
growth between the 2 oyster species over the 9-mo period. Specific
growth rates for C. ariakensis did not differ over time, but they did
for C. virginica. The growth rate of C. virginica was slowest in the
winter (8.9 x [10.sup.-4] [mm.sup.2]/day) and fastest in the spring
(43.5 x [10.sup.-4] [mm.sup.2]/day). Mortality of both species rose
abruptly in April 2007, and all (100%) remaining C. ariakensis were dead
by the end of the study. Although 28% of the remaining C. virginica died
in April 2007, there was little further mortality in this species before
the study was terminated in August 2007. Physiological responses of both
species of oysters were compared under seasonal temperate euhaline
quarantine conditions to understand better how temperature affects these
species without the confounding unexplained mortality encountered in the
subtropical mesocosms. The clearance rate of C. ariakensis (1.2 L/g/h)
was half that of C. virginica (2.2 L/g/h) during the summer
(25[degrees]C); however, respiration rates for C. ariakensis (2.6 ml
[O.sub.2]/g/h) and C. virginica (2.5 ml [O.sub.2] g/h) were similar. The
low clearance rate of C. ariakensis during the summer resulted in a
negative scope for growth (-36.2 J/g/h) during this season. During the
winter, C. ariakensis remained physiologically active when water
temperatures were as low as 2[degrees]C; C. virginica were quiescent
during this time. We conclude that the "Oregon" strain of C.
ariakensis tested will not thrive in the warm subtropical waters of the
U.S. southeastern coast; however, given its native range in Asia, we do
not discount the possibility of an adaptation to warmer temperatures
KEY WORDS: oyster Crassostrea ariakensis, Crassostrea virginica, growth rate, reproduction, physiology
The eastern oyster (Crassostrea virginica Gmelin 1791) is an ecologically and economically important native species along the Atlantic and Gulf coasts of the United States (Newell 1988, MacKenzie 1996, Eggleston 1999, Posey et al. 1999, Coen et al. 2007). In Chesapeake Bay, populations of C. virginica have been in decline since the late 19th century as a result of a combination of habitat degradation, overharvesting, and epizootics of the protistan diseases MSX (Haplosporidium nelsoni) and Dermo (Perkinsus marinus) (Rothschild et al. 1994, Ford & Tripp 1996). It was proposed by the Maryland Department of Natural Resources that the introduction of the nonnative Suminoe oyster (Crassostrea ariakensis Fujita 1913) into Chesapeake Bay would help alleviate many of the problems associated with the loss of native C. virginica populations (National Research Council 2004). Because of the controversial nature of performing such an irreversible introduction, considerable research, including the work described here, was commissioned to help provide the scientific information necessary to inform such a decision. This body of research was used to develop a formal Ecological Impact Statement (U.S. Army Corps of Engineers, Norfolk District 2009). Ultimately, the introduction of the nonnative C. ariakensis was not considered to be a suitable means of enhancing oyster stocks at this time. The current management emphasis has now shifted to conservation and restoration of C. virginica.
The initial rationale for the proposed introduction of C. ariakensis was based on this species resistance to MSX and Dermo epizootic diseases that are major impediments to the restoration of C. virginica populations (Calvo et al. 2001, Paynter et al. 2008). If C. ariakensis were to be introduced into Chesapeake Bay, either deliberately, or accidentally from research facilities holding diploid broodstock, it was considered likely (National Research Council 2004) that it would establish feral populations along the U.S. Atlantic coast, including areas where C. virginica populations remain relatively robust.
The native range of C. ariakensis stretches from latitude 41[degrees]N in Liaoning, China, to latitude 20[degrees]N in Vietnam (Guo et al. 2008). In the Americas, this is equivalent to the coastline between Connecticut and the Yucatan Peninsula in Mexico (Guo et al. 2008). Most research in relation to the proposed introduction of C. ariakensis into Chesapeake Bay has been performed on triploid individuals grown under temperate estuarine conditions (Calvo et al. 2001, Grabowski et al. 2004, Hudson et al. 2005, McLean & Abbe 2008, Paynter et al. 2008, Kingsley-Smith et al. 2009) or on diploid C. ariakensis being assessed for aquaculture potential on the U.S. Pacific northwest coast (Breese & Malouf 1977, Perdue & Erickson 1984, Langdon & Robinson 1996). Considering that the natural range of C. ariakensis extends into subtropical regions (Zhou & Allen 2003, Guo et al. 2008), it is necessary to broaden past studies to include subtropical U.S. coastal environments. One such subtropical estuary is the Indian River Lagoon (IRL) on the Atlantic coast of Florida, which is highly biologically diverse (Gilmore 1985, Duncan et al. 2004) because of its location near the boundary between the temperate and subtropical regions. Subtropical conditions within the IRL promote the enhanced growth of many species as a result of year-round warm water temperatures and high phytoplankton availability. Populations of C. virginica in the IRL are relatively robust (Grizzle 1990, Boudreaux et al. 2006) in comparison with those in Chesapeake Bay.
To investigate the potential of C. ariakensis to form feral populations in subtropical conditions, we examined their growth, mortality, and reproductive capability over a 9-mo period in quarantined mesocosms supplied with seawater from the IRL. We hypothesized that under subtropical conditions C. ariakensis would perform better than or equal to C. virginica. Our initial hypothesis was not supported: therefore, to understand better C. ariakensis physiology under warm water temperatures, we then compared seasonal scope for growth in both species of oyster under salinities and summer water temperatures similar to that of the IRL. We hypothesized that C. ariakensis would have less energy to allocate toward somatic growth and gamete production during the summer as a result of physiological stress by high water temperatures.
MATERIALS AND METHODS
Two age classes (4 mo and 28 too) of both oyster species were used in this study. All C. virginica and the 4-too-old C. ariakensis were obtained as larvae from the Virginia Institute of Marine Science--Eastern Shore Laboratory (VIMS-ESL). The 28-mo-old C. ariakensis were obtained as larvae in 2004 from Taylor United Shellfish hatchery in Quilcene, WA. Larvae were allowed to metamorphose on large pieces of oyster shell and were reared as described by Newell et al. (2007) in a quarantined ambient flow-through facility at Horn Point Laboratory (HPL) in Maryland until use. All C. ariahensis larvae were progeny of adults originating from the "Oregon" stock (Newell et al. 2007).
Three circular (diameter, 1.6 m: height, 0.6 m) 700-L mesocosms at HBO1 were supplied with ambient seawater from the IRL at a flow rate of 10 L/min, and were aerated to maintain high dissolved oxygen levels (Table 1) and to keep seston in suspension. Effluent water was chlorinated to 2 ppm or greater free chlorine using a free chlorine analyzer (Foxcroft FX-1000p, Foxcroft Equipment and Service, Glen More, PA) and controller (Foxcroft FX-8500) to prevent any gametes or larvae produced by the diploid oysters from entering the IRL.
In September 2006, oysters of each species from the 28-mo-old cohorts and the 4-mo-old C. virginica cohort were separated into groups of approximately 50 individuals. Each group was placed into plastic trays (53 cm long x 38 cm wide x 14 cm high) with holes (diameter, 2.5 cm) drilled into the bottom and sides of each tray to allow for better water circulation. A tray for each oyster species was placed into the 3 mesocosms and suspended 5 cm above the bottom of the tank to avoid oysters becoming buried by particulate waste on the tank bottom. Tanks were cleaned periodically to remove accumulated sediment and organic matter. In December 2006, the additional 4-mo-old C. ariakensis transferred from HPL were acclimated to ambient salinity and temperature conditions for 7 days. They were then distributed into plastic trays and to each of the 3 mesocosms as described.
Mesocosm water temperature, salinity, and dissolved oxygen were measured daily using a conductivity meter (Model 85, Yellow Springs Inc., Yellow Springs, OH). One water sample from each of the 3 mesocosm tanks, and concurrent triplicate samples from the seawater intake in the IRL, were collected once per month in July and August 2006 for chlorophyll a and seston analysis using the EPA 160.2 (USEPA 1999) method for seston and the EPA SM 10200H method for chlorophyll a (American Public Health Association 1998).
Growth and Mortality
Oyster shells (16 for each species) with attached live oysters (2-9 per shell) from the 4- and 28-too-old C. ariakensis and C. virginica cohorts were labeled with an aluminum tag. A total of 42 C. ariakensis and 56 C. virginica attached to these shells were used for the growth and mortality assessments. Each 4- and 28-mo-old C. virginica oyster and 28-mo-old C. ariakensis oyster on the tagged shells was digitally photographed in September 2006 at the start of the experiment to obtain an initial reference size. Each 4-too-old C. ariakensis oyster was photographed digitally in December 2006 to obtain an initial reference size. Each oyster was then photographed monthly from December 2006 until the study ended in August 2007. All digital photographs included a scale with l-mm increments within the field of view to provide calibration.
Absolute shell growth was calculated by measuring the surface area (in square millimeters) of oysters at each sampling date using Image J software (Rasband & Image 1997 2009). Using absolute measurements to estimate growth may lead to an overestimation in the growth rate of larger oysters versus smaller oysters; therefore, to standardize growth rate relative to an oyster's size, we calculated growth as a daily specific growth rate (SPG):
SPG = (ln[A.sub.z]-ln[A.sub.1]) / [t.sub.2] - [t.sub.1]
where A is the area (in square millimeters) of the oyster shell at the beginning and end of each sampling period, and [t.sub.2] - [t.sub.1] is the time (in days) between each sampling date. The percent cumulative mortality of each species was estimated by the number of missing and dead oysters at each sampling date as verified by digital image analysis.
Reproductive assessment of mesocosm oysters on untagged shells was performed monthly between February 2007 and August 2007. Tissue samples from 12 21 oysters of each species (split between age classes), were wet weighed, fixed, paraffin embedded, sectioned transversely, and stained with hematoxylin and eosin (Kennedy et al. 1995). In March and July, several oysters from each cohort with untagged shells were also examined histologically for any type of cellular abnormalities. Tissue samples of 20 C. virginica (shell area, 22.9 [+ or -] 5.4 [cm.sup.2]) from the nearby natural population in the IRL were collected in August 2007 to make additional comparisons with mesocosm C. virginica at that same sampling date.
Gonadal tissue in histological sections for each oyster was examined microscopically, and the number of oysters in which gametogenesis had been initiated was enumerated. Oysters in this category had distinct follicles, but the cells had not yet differentiated sufficiently to discern whether they would develop as eggs or sperm. The number of those individuals in which gametogenesis had proceeded to the point that distinct eggs and sperm were visible in the follicles was enumerated separately.
Physiological studies were performed under ambient seasonal temperate conditions at VIMS-ESL in July 2008, October 2008, January 2009, and April 2009. The C. ariakensis (shell height, 3.7-12 cm) used in this experiment were from the same cohorts used in the HBOI mesocosm study, but reared at HPL. The size of C. virginica reared in HPL mesocosms (shell height, 2.5-5.3 cm) was smaller than C. ariakensis (Newell et al. 2009), so they were not used in this study. Instead, we collected C. virginica (shell height, 5.2-13.9 cm) directly from natural oyster reefs in Choptank River, Maryland, to study similar-size individuals of both species.
All oysters were maintained in ambient mesohaline (10-12) flow-through conditions at HPL. One month prior to each seasonal physiological study, we transferred 16 oysters of each species (32 total oysters) to VIMS-ESL, where they were gradually acclimatized to ambient conditions by increasing salinity by 5 salinity units every 4-5 day until ambient salinity (~30) was reached. During acclimatization, oysters were maintained in static tanks at ambient water temperature and fed a maintenance diet of cultured microalgae I. galbana clone T. iso. Oysters of both species acclimatized to conditions at VIMSESL as indicated by the presence of biodeposits and growth of new shell prior to the start of each seasonal study, except for C. virginica in winter, when all individuals were in a quiescent condition.
Ambient flow-through water was pumped from the adjacent Machipongo River into 2 head tanks that supplied water via lengths of Tygon tubing (6 mm i.d.) to 18 rectangular plastic pans (36 cm long x 30 cm wide x 10 cm high). A PVC plug with a precisely drilled hole was inserted into each tube that allowed a flow rate of either 40 L/h (for oysters [greater than or equal to]-5 cm in shell height) or 20 L/h (for oysters <5 cm in shell height) to the bottom of each pan. Preliminary studies showed that these high flow rates ensured that oysters would not be able to reduce particle concentrations appreciably during the experiment. All pans drained at the water surface though a standpipe at the end farthest from the inflow tube. This setup ensured adequate water column mixing through each pan. Waste effluent was collected in a holding tank and was chlorinated to 2 ppm or greater free chlorine for 2 h. Six oysters from each species were randomly assigned to 12 separate pans with an appropriate flow rate for their shell size. Controls, which did not contain oysters, were assigned to 3 pans with the higher flow rate and 3 pans with the lower flow rate. Each run of the experiment (3 total for each season) lasted 36 h, during which hourly water samples for seston analysis were taken using an ISCO water sampler (model 3700 Sampler Controller). Each oyster was held in a shallow plastic container placed in each pan to retain biodeposits. Appropriate containers were also placed in the control pans of both flow rates. Oysters were briefly removed from the pans after 12 h to eliminate biodeposits produced from seston that had been filtered and ingested before the start of the experimental run.
Oysters were removed from the pans at the end of the experiment. The shallow containers from oyster and control pans were carefully removed, sealed, and held at 5[degrees]C for 12 h to allow suspended material to settle. Overlying water was aspirated off, and the container was filled with 200 mL DI water to wash out salt from the deposits before holding at 5[degrees]C for another 12 h to once again allow suspended material to settle. The majority of DI water was aspirated off and 2 1-mL aliquots of biodeposits from the containers were removed for absorption efficiency determination. Each aliquot was placed onto 2 preweighed Whatman GF/C filters that had first been washed and heat treated at 450[degrees]C for l h. The remainder of the biodeposit slurry was transferred into a preweighed aluminum pan, which were placed into a 90[degrees]C drying oven for 24 h, after which dry weights were taken. The filters were also dried at 90[degrees]C and weighed to determine total dry weight. The filters were then heat treated at 450[degrees]C for 6 h to determine the organic fraction of the biodeposits. Material from the control containers was treated identically to the material from the oyster containers and was collected to determine the amount of material that naturally settled into the experimental containers independently of oyster feeding activity. To correct for this extra material in the oyster containers, the amount of organic and inorganic material from the control containers was determined as described earlier, and then subtracted from the total material present in the oyster containers.
Known aliquots of water (300-500 mL) collected by the ISCO (Teledyne Isco, Inc. Lincoln, NE) sampler were filtered through GF/C filters and treated in the same manner as bio-deposits to estimate seston concentration. Clearance rate (measured in liters per gram per hour) was calculated as milligrams of inorganic matter egested both as feces and pseudofeces per hour divided by milligrams of inorganic matter available per liter of seawater (Hawkins et al. 1996). Absorption efficiency was calculated using the Conover ratio (Conover 1966, Bayne et al. 1985).
For nitrogen excretion assays, ambient river water was filtered (0.45-[micro]m pore; Millipore) and used to fill beakers (200 900 mL) into which individual oysters were submerged or assigned as controls. Beakers were covered with plastic food wrap and incubated at ambient seawater temperature in a water bath for 2 h. Oysters were then removed from the beakers, and two 10-mL aliquots water from each beaker was placed into labeled test tubes. The phenol-hypochlorite method (Solorzano 1969, Bayne et al. 1985) was used to determine ammonium concentration. Ammonium excretion rates (measured as micrograms N[H.sub.4]-N per gram per hour) were calculated as described by Bayne et al. (1985).
Rates of oxygen consumption were measured using the methods described by Bayne et al. (1985). Individual oysters were then placed into either a large (2.3 L) or small (0.3 L) glass respirometer chamber supplied with ambient flow-through river water. These chambers were maintained at ambient river temperature by submerging them in a water bath. After 1 h of acclimatization, the water flow was stopped, and the decline in oxygen concentration was measured with a calibrated oxygen electrode (model E5047-0, Radiometer-Copenhagen). Controls were run using the same methods described earlier, but without an oyster in the respirometer chamber. Control respiration rates were subtracted from the oyster runs to eliminate background respiration. The calculation of oxygen consumption rates required the volume of the oyster to be subtracted from the total volume of water in the respirometer chamber (Bayne et al. 1985): therefore, oyster volume was determined by the displacement of water within a graduated cylinder. Respiration rates (measured in milliliters 02 per grain per hour) were calculated as described by Bayne et al. (1985).
Dry tissue weight (dw) of all experimental oysters was obtained by removing oyster tissue from its shell, placing it in a preweighed pan, and drying it at 90[degrees]C for 24 h. Seasonal physiological rates of individual oysters were regressed against their dw for C. virginica and C. ariakensis separately. Analysis of covariance (ANCOVA) was then performed to determine a species-specific common slope. Using this result, slope intercepts were recalculated using the allometric equation Y = a[X.sup.b]. These intercepts represent the seasonal physiological rates of an oyster of 1 g dw. This weight was selected because it is close to the average weight of the oysters studied, and also this animal weight is commonly used in comparisons of physiological rate functions within and between species of bivalves (Bayne & Newell 1983). Atomic ratios of oxygen consumption to nitrogen excretion (O:N) were calculated as described by Bayne et al. (1985) from standardized 1 g dw seasonal rates for each oyster species.
The physiological rates described earlier were converted into energy equivalents (measured in Joules per gram per hour (Bayne et al. 1985)) using a value for percent organic matter (%POM) of 23.5/mg (Widdows et al. 1979). This value is representative of the energy value for food materials such as seston (Slobodkin & Richman 1961, Bayne et al. 1985). Scope for growth (P) is the energy available for allocation to germinal and somatic tissue production, and it was calculated using (Bayne et al. 1985).
P(J/h) - a - (R + U)
where A is energy absorbed from seston, R is energy respired, and U is energy excreted.
The distributions of the specific growth rate for C. virginica and C. ariakensis were not normal and could not be made to approximate normality through transformation. Therefore, a nonparametric Freidman's test (Zar 1999), which is similar to repeated-measures ANOVA, was used to test for differences in growth rate between and within oyster species. Post hoc nonparametric Wilcoxon's signed rank sum tests (Zar 1999) were used to determine significant monthly differences in growth rate within species.
Percent cumulative mortality was arcsine-transformed to achieve approximate normality. A repeated-measures ANOVA was performed on the transformed data, and post hoc least significant difference (LSD) multiple mean comparison tests were conducted to determine significant monthly differences between species.
ANCOVA was used to test for differences in the seasonal physiological rates within each species. Post hoc LSD multiple mean comparison tests were performed to determine which seasons were significantly different from each other.
Percent absorption efficiency was arcsine-transformed to approximate normality. ANOVA was performed to determine whether there was a seasonal difference in absorption efficiency within species, and post hoc LSD multiple mean comparison tests were performed to determine significant seasonal differences.
Mesocosm water temperature, salinity, and dissolved oxygen concentrations were similar to the annual cycle observed in the adjacent IRL. Salinity values during the experiment ranged from 28.0-37.7 and were highest during the spring and early summer seasons (Table 1). These salinities were well within the optimal range for C. virginica (Shumway 1996). Water temperatures during this study ranged from 18.6-30.4[degrees]C and were highest in the late spring and summer seasons (Table 1). The percent dissolved oxygen saturation in mesocosms ranged from 74.1-91.2% (Table 1) and were similar to dissolved oxygen saturations found on natural oyster assemblages in the IRL (Wilson et al. 2005).
Seston and chlorophyll a concentrations in mesocosms and the IRL were measured in the summer to compare food availability. Seston was higher in mesocosms (July, 9.5 [+ or -] 3.6 mg/L: August, 8.6 [+ or -] 2.7 mg/L) than in the IRL (July, 4.0 [+ or -] 1.6 mg/L; August, 5.4 [+ or -] 1.3 mg/L): however, chlorophyll a values were similar among mesocosms (July, 3.6 [+ or -] 0.9 [micro]g/L: August, 4.4 [+ or -] 0.6 [micro]g/L) and the IRL (July, 3.7 [+ or -] 0.4 [micro[g/L; August, 6.8 [+ or -] [micro]g/L). These values are consistent with seston and chlorophyll a concentrations found throughout the IRL (Christian & Sheng 2003).
Application of a Friedman's test showed that there was no significant difference in SPG between C. ariakensis and C. virginica ([F.sub.r(3)] = 3.1, P > 0.05) for the 4-mo-old cohorts from January 2006 through April 2007. The absence of the 4-too-old cohort of C. ariakensis in the September to December 2006 sampling period, and high mortality rates of C. ariakensis after April 2007, precluded these sampling periods from being used in the analysis. Differences in SPG between C. ariakensis and C. virginica from the 28-too-old cohorts were nonsignificant ([F.sub.r(3)] = 7.4, P[alpha] < 0.05 = 0.06) between September 2006 and March 2007. High mortality rates of C. ariakensis from the 28-mo-old cohort after March precluded later sampling periods from being used in the analysis.
A Friedman's test performed on monthly C. ariakensis growth data (Table 2) showed that there was no significant difference in SPG for the 4-month-old ([F.sub.r(6)] = 9.3, P > 0.05) and 28-mo-old ([F.sub.r(5)] = 1.8, P > 0.05) cohorts. The same test performed on monthly C. virginica growth data showed that there was a significant difference among months in SPG for the 4-month-old ([F.sub.r(7)] = 52.10, P < 0.0001) and 28-mo-old ([F.sub.r(7)] = 19.09, [P.sub.[alpha]<0.05] = 0.008) cohorts over the course of the experiment (Table 2).
The 4-mo-old cohort of C. virginica had the fastest SPG between September and December 2006. Post hoc comparisons using Wilcoxon's signed rank sum tests showed that SPG during this period was significantly greater than all other sampling periods (Table 2). An additional increase in SPG occurred between April and May, and May and June; however, post hoc comparisons showed that SPG during these months was only significantly greater than SPG recorded between December and February (Table 2).
The 28-mo-old cohort of C. virginica also had their fastest SPG between September and December 2006 (Table 2): however, post hoc comparisons using Wilcoxon's signed rank sum test showed that SPG during this time was not significantly different from the rest of the study period, with the exception of January (S = 24.5. P> 0.0098), April (S = 17.5. P = 0.0391), and August (S = -10.5. P = 0.0313).
Repeated-measures ANOVA showed there was a significant monthly interaction in the cumulative mortality for the 4-mo-old C. ariakensis and C. virginica cohort ([F.sub.14,28] = 27.64, [P.sub.[alpha]<0.05] < 0.0001) as well as the 28-mo-old cohort ([F.sub.16,32] = 8.66, [P.sub.[alpha]<0.05] < 0.0001). The 4-mo-old cohort of C. virginica sustained high mortality between April and May 2007, during which cumulative mortality doubled. After this period, and until experiment termination, there was little mortality recorded. The 4-mo-old cohort of C. ariakensis also had low cumulative mortality until between April 2007 and May 2007, when cumulative mortality increased 4-fold (Fig. 1A). Unlike C. virginica, C. ariakensis continued to experience heavy mortality, and all oysters were dead by August 2007 (Fig. 1A). Post hoc LSD tests showed that mortality of 4-mo-old C. virginica was significantly lower than C. ariakensis between the June sampling and the conclusion of the study in August 2007 (Fig. 1A).
[FIGURE 1 OMITTED]
The 28-mo-old cohort of C. virginica also exhibited high mortality between April and May 2007 when 25% died; and additional mortality between July 2007 and August 2007 brought cumulative mortality to 52% (Fig. 1B). The 28-mo-old cohort of C. ariakensis experienced high cumulative mortality from February 2007 until July 2007, at which point 100% of the oysters in the mesocosms were dead (Fig. 1B). Post hoc LSD tests showed that there was a significant difference between the cumulative mortality of C. virginica and C. ariakensis after March 2007.
Some individuals from the 4-mo-old cohorts of both species exhibited early-stage gametogenesis for all sampling times between December and July, although fewer C. ariakensis than C. virginica exhibited gametogenesis during the March and April samplings (Table 3). The 28-mo-old cohorts of C. virginica and C. ariakensis also exhibited high levels of early-stage gametogenesis. Gametogenesis in C. virginica from the 28-mo-old cohort proceeded to the point that eggs and sperm could be identified clearly in follicles of 30-60% of individuals from May through August 2007, but no developed gametes were visible from any C. ariakensis (Table 3). All C. virginica (n = 10) oysters sampled from the IRL in August 2007 had distinguishable male and female gametes, in contrast to the low to moderate percentage of those in the mesocosms. Both age classes from the 2 oyster species showed an abrupt decline in the number of individuals with evidence of gonadal development in April 2007, compared with March and May (Table 3). This sharp decline in reproductive activity coincided with the reduced growth and increased mortality of these oysters that occurred during this same period.
Seasonal water temperatures at VIMS-ESL ranged from 27[degrees]C in the summer to 5[degrees]C in the winter: salinities remained euhaline ~30; Table 4). Seston loads at VIMS-ESL were high (13.8-49.3 mg/L) for all seasons sampled (Table 4), and %POM ranged from 9.3-17.9%. Seston and %POM were similar among all seasons sampled, with the exception of summer (July), when the seston load of 49.3 mg/L was 3 times greater, but the 9.3% POM was only approximately half that of the other seasons (Table 4). The seawater intake pipes at VIMSESL are located in a muddy creek that is subject to high tidal currents that can resuspend bottom sediments, thereby creating high seston concentrations and lower %POM.
There was no interaction between dw and season for clearance rate of C. virginica (ANCOVA; [F.sub.2,37] = 0.01, P > 0.05) or C. ariakensis (ANCOVA; [F.sub.3,53] = 1.42, P > 0.05). This allowed us to calculate a common slope for C. ariakensis (h = 0.62) and C. virginica (b = 0.44) for the regression equation that we then used to recalculate the intercept for each season, which equates to the clearance rate for a standardized oyster of I g tissue dw for each species over the 4 seasons. There were no significant differences in the clearance rate of C. virginica (Table 5) between summer, autumn, and spring (ANCOVA; [F.sub.2,39] = 1.39, P > 0.05). Winter data were not included because these oysters were not observed feeding and did not produce biodeposits during this period.
There were significant differences in the seasonal clearance rate (Table 5) of C. ariakensis (ANCOVA: [F.sub.3,56] = 36.40, [P.sub.[alpha]<0.05] = 0.0001), with significantly reduced rates during spring compared with autumn (LSD; [t.sub.56] = -2.38, [P.sub.[alpha]<0.05] = 0.0210) and summer ([t.sub.56] = 3.52, [P.sub.[alpha]<0.05 = 0.0009). Interestingly, C. ariakensis fed and voided biodeposits during the winter when temperatures ranged from 2-5[degrees]C.
There was significant seasonal interaction in absorption efficiency for the two oyster species (ANOVA: [F.sub.3,95] = 49.04, [P.sub.[alpha]<0.05] < 0.0001). The absorption efficiency of C. virginica differed significantly among seasons (Table 6), with the highest efficiency occurring in spring (44.1%) and the lowest in winter (0%). The absorption efficiency of C. ariakensis also differed significantly among seasons (Table 6), with the highest efficiency occurring in winter (43.5%) and the lowest in summer (15.4%).
There was no interaction between dw and season for the respiration rate of either C. virginica (ANCOVA: [F.sub.3,39] = 1.94, P > 0.05) or C. ariakensis ([F.sub.3,43] = 1.35; P > 0.05). This allowed us to calculate a common slope for C. ariakensis (b = 0.59) and C. virginica (b = 1.07) for the regression equation that was then used to recalculate the intercept for each season (Table 5).
The respiration rate of C. virginica during the summer was significantly greater than in the autumn (LSD; [t.sub.42] = -4.15, [P.sub.[alpha]<0.05] = 0.0002), winter ([t.sub.42] = 5.53, [P.sub.[alpha]<0.05] < 0.0001), and spring ([t.sub.42] = 5.09, [P.sub.[alpha]0.05] < 0.0001; Table 5). The respiration rate of C. ariakensis was also significantly greater in the summer than in the autumn (LSD; [t.sub.46] = -7.72, [P.sub.[alpha]<0.05] < 0.0001), winter ([t.sub.46] = 10.90, [P.sub.[alpha]<0.05] < 0.000l), and spring ([t.sub.46] = 8.03, [P.sub.[alpha]<0 05] < 0.0001; Table 5).
There was no interaction between oyster dw and season for ammonium excretion for C. virginica (ANCOVA; [F.sub.3,42] = 0.41, P > 0.05) or C. ariakensis ([F.sub.3,48] = 0.76, P > 0.05). This allowed us to calculate a common slope for C. ariakensis (b = 0.44) and C. virginica (h = 0.57) for the regression equation that was then used to recalculate the intercept for each season (Table 5).
The ammonium excretion rate of C. virginica was significantly higher in the summer than in the autumn (LSD; [t.sub.45] = 6.44, [P.sub.[alpha]<0.05] < 0.0001), winter ([t.sub.45] = -6.58, [P.sub.[alpha]0.05] < 0.0001), and spring ([t.sub.45] = 4.63, [P.sub.[alpha]<0 05] < 0.0001; Table 5). The ammonium excretion rate of C. ariakensis differed significantly among all seasons sampled (Table 5), with the highest rates occurring in summer and the lowest occurring in winter.
Both oyster species exhibited a seasonal pattern in their scope for growth (Fig. 2). For a C. virginica, of I g tissue dw, the greatest amount of energy available for tissue growth (somatic and germinal) occurred in the spring (46.3 J/g/h), whereas in winter a negative scope for growth (-4.5 J/g/h) was calculated because C. virginica were not feeding. There was a negative scope for growth for 1 g tissue dw of C. ariakensis during the summer (-36.2 J/g/h) when this species had high metabolic activity, and in winter (-1.02 J/g/h) when this species was physiologically active but metabolic activity was low (Table 5); there was a positive scope for growth for the other seasons.
The O:N ratio for C. virginica was lowest during the summer (<50) and highest (>100) in the autumn and winter seasons (Fig. 3A). The O:N ratio for C. ariakensis remained relatively low (<70) throughout the year, with the highest ratios (>50) during the winter and spring seasons (Fig. 3B).
Rates of shell growth did not differ significantly between C. ariakensis and C. virginica maintained under subtropical conditions. Although no statistical differences in growth rate were detected between the 28-mo-old cohorts of each species, C. ariakensis grew at ~25% of the daily rate of C. virginica during the first 90 days (September to December) of the study. There was no significant difference in the monthly growth rate of C. ariakensis between age cohorts; however, the 4-mo-old cohort did exhibit a higher rate of growth from May until August. The lack of a detectable monthly significance in growth in this cohort may have been a result of the severe mortality that reduced the sample size. and hence the power of the statistical test. There was a significant difference in the monthly growth rate of C. virginica, when high rates of growth for the 4-mo-old cohort occurred in May and June 2007. The enhanced growth of 4-mo-old cohort oysters was not likely caused by changes in environmental factors such as increased temperature or increased food availability, because this growth spurt was not seen within the 28-mo-old cohort of either species. The increased growth during this period may be the result of 4-mo-old oysters allocating more energy toward somatic growth than gamete development, an ontogenic shift in energy allocation that is typically seen in a long-lived invertebrate such as oysters (Thompson et al. 1996).
[FIGURE 2 OMITTED]
Both C. ariakensis and C. virginica exhibited high levels of very early gametogenesis; however, in very few individuals of either species or age class did gametogenesis proceed to the point that there were clearly distinguishable eggs or sperm within the follicles. Both oyster species showed a sharp, unexplained decline in the number of individuals with evidence of even early gametogenesis in April 2007, compared with March and May. The only cohort to show any gamete differentiation by the end of the experiment in August was the 28-mo-old C. virginica cohort (60%), whereas 100% of similar-size C. virginica sampled from the IRL in the vicinity of the seawater intake in August 2007 had clearly distinguishable male and female gametes. This is in accordance with reports by Wilson et al. (2005) that oysters in south Florida waters have ripe gametes present from May to October. Furthermore, in mesocosms simulating the mesohaline region of Chesapeake Bay and containing individuals from the same 28-mo-old cohort oysters used in the current study, Newell et al. (2009) observed distinguishable gametes in 60% of C. virginica and 80% of C. ariakensis in June 2007, compared with 30% of C. virginica and 0% of C. ariakensis in the subtropical system described here.
[FIGURE 3 OMITTED]
Both age classes of C. ariakensis suffered greater mortality than C. virginica within our experimental mesocosms. The 28-mo-old cohort of C. ariakensis began to die in mid February, with 100% mortality of all individuals by mid June. The 4-mo-old cohort of C. ariakensis began to die in mid May and experienced total mortality by August. The 28-mo-old cohort of C. virginica experienced a die-off in mid April, when ~20% of the oysters died, and in July an additional ~20% died. However, ~60% of C. virginica individuals from this cohort were still alive at experiment termination in August 2007. The C. virginica from the 4-mo-old cohort also experienced high mortality ~25%) in April, but little mortality was noted through the remainder of the study.
Oyster mortality within the mesocosms did not appear to be associated with any known adverse environmental conditions. Salinities in our experimental mesocosms were always fully euhaline (average salinity, 34) which is optimal for both C. virginica (Shumway 1996) and C. ariakensis (Calvo et al. 2001, Grabowski et al. 2004, Kingsley-Smith et al. 2009). Other environmental conditions in our mesocosms such as temperature, dissolved oxygen, and seston concentration also were typical of the annual cycle found in the I RL (Christian & Sheng 2003, Wilson et al. 2005). There is evidence that the warm temperatures encountered during the summer in Chesapeake Bay, and year-round in the IRL may reduce the growth rate of C. ariakensis. Calvo et al. (2001) observed no growth of either triploid C. ariakensis or diploid C. virginica during the summer (22-29[degrees]C) at high-salinity sites in Chesapeake Bay. The lack of summer growth for C. virginica reported by Calvo et al. (2001) is likely attributable to an intense outbreak of P. marinus, which subsequently caused heavy C. virginica mortality. The lack of C. ariakensis summer growth could not be attributed to disease or any other stressors (Calvo et al. 2001). They noted that the majority of growth in this species occurred during the spring and fall periods, when water temperatures were appreciably cooler. Conversely, other studies of triploid C. ariakensis in Chesapeake Bay (Paynter et al. 2008, Kingsley-Smith et al. 2009) and North Carolina estuaries (Grabowski et al. 2004) reported growth of C. ariakensis during the warm summer months. Langdon and Robinson (1996) reported that although C. ariakensis spat grew best at salinities of 25 35 at temperatures of 20-25[degrees]C during the summer, they also continued to grow equally well during the winter at several sites on the coast of Oregon. In its natural range C. ariakensis seems to flourish in waters with a wide annual temperature range of 3-28[degrees]C (Kang et al. 2000, Harding & Mann 2006, Yoon et al. 2008). Most evidence seems to support that C. ariakensis should grow well in high-salinity warm waters such as the IRL; however, we found that diploid C. ariakensis grew poorly and suffered high mortality when maintained under such conditions during our study.
The mortality of both species of oysters was also not associated with infections of any of the 3 well-recognized oyster parasites. Histological analysis did not reveal the presence of H. nelsoni in either species of oyster, and P. marinus (although present in both species of oysters) was at low prevalence and intensities (Scarpa et al. 2009). Bonamia sp. was detected in February 2007: however, the intensity of infection was not high enough to cause mortality (Scarpa et al. 2009). Grabowski et al. (2004) found high mortality of small triploid C. ariakensis grown in subtidal estuaries of North Carolina during the summer. They suggest that high mortality may limit the growth advantage C. ariakensis seems to have over C. virginica in high-salinity environments (Calvo et al. 2001, Paynter et al. 2008, Kingsley-Smith et al. 2009). Prevalence of P. marinus reported by Grabowski et al. (2004) in both species of oyster was light (0-16.7%) and not hypothesized to be the cause of the observed mortality, they did not test for the presence of H. nelsoni or Bonamia sp. Subsequent field trials in high-salinity estuaries of North Carolina have shown that smaller (shell height, <50 mm) C. ariakensis are particularly sensitive to Bonamia sp. infection, with mortality reaching 100% when temperatures exceed 20[degrees]C during the summer and early fall (Carnegie et al. 2008). Audemard et al. (2008) confirmed in laboratory studies that high salinities (20-30) coupled with high temperatures (>20[degrees]C) resulted in high Bonamia sp.-induced mortality of C. ariakensis. Prevalence of Bonamia sp. (60-100%) and intensity of infection reported by Carnegie et al. (2008) and Audemard et al. (2008) were much higher than the prevalence (0-40%) and intensity of infection in C. ariakensis from our mesocosms (Scarpa et al. 2009).
Concurrent with the onset of high mortality, the physiological condition of both species declined. Oysters appeared emaciated and edematous, which may be an indication of lack of feeding. The digestive gland in oysters sampled as part of the routine histological samples in March 2007 and July 2007 were examined microscopically for evidence of feeding activity and nutrient assimilation. Both C. ariakensis and C. virginica showed equal evidence of feeding activity as indicated by the presence of ingested food in the gut of 75-80% of the individuals examined. However, by July, C. virginica had a higher evidence of feeding activity (88%) than C. ariakensis (44%). Recent digestion of particles was indicated by columnar and cuboidal digestive gland epithelia, which were at similar levels in both species of oyster in March (67-72%). By July, recent particle digestion was inversely related to observed food ingestion frequencies, as 100% of C. ariakensis exhibited particle digestion compared with only 70% of C. virginica. A comparison of mesocosm and C. virginica freshly collected from location of the seawater intake in the IRL in August 2007 showed no difference in the feeding activity of mesocosm (78%) and wild oysters (77%), although recent particle digestion was lower for mesocosm oysters (65%) than for wild oysters (80%). Overall, there was no evidence of consistent deficiencies in feeding activity that may explain our gross observations of edematous emaciation in both species of oyster. It is possible that, although we observed the ingestion and digestion of food particles, these particles were composed of phytoplankton species that could not support the oysters' nutritional requirements; however, this would not explain the observed rapid increase in mortality rates in C. ariakensis compared with C. virginica.
Taken together, these findings of reduced growth rate, increased mortality, and decline in the reproductive activity of C. ariakensis, and to a much lesser extent in C. virginica, between mid February and mid April indicate that oysters in our mesocosms were subjected to some unknown stress during this period. We postulate that C. ariakensis' may have experienced thermal stress in the prolonged period of more than 20[degrees]C water temperatures, which are characteristic of the subtropical IRL. Bonamia sp. was present in our mesocosms at this time; however, prevalence and intensity of infection were low (Scarpa et al. 2009), and do not fully explain the mortality and decreased reproductive activity observed during this period. It is possible that water pumped from the IRL contained a toxin that was inadvertently released during routine maintenance of a vessel in the channel. Alternatively, there may have been a bloom of a toxic species of phytoplankton that was neither manifest (e.g., as a fish kill) nor readily detected by histology or measurement of environmental parameters (e.g., dissolved oxygen).
Physiological studies allowed us to examine whether differences in physiological responses to temperature were responsible for the poor growth and survival of C. ariakensis compared with C. virginica of a similar range of dw's (Table 5). These studies were performed in Virginia on oysters maintained under similar environmental conditions to those in our Florida mesocosms, but without the confounding factor of Bonamia sp. presence. Summer water temperatures were similar between the two locations, but the Virginia study site was subjected to a wider range of water temperatures during the remaining seasons. The scope for growth for both species of oysters showed a distinct seasonality in the amount of energy available for somatic growth and gamete production.
During the summer, at temperatures more than 25[degrees]C, C. ariakensis had a negative scope for growth (-36.2 J/g/h), which was the result of a low clearance rate (1.16 L/g/h) without an equivalent decrease in the other physiological rates. This supported our hypothesis that the mortality we observed in C. ariakensis in the Florida mesocosms was the result of high water temperatures imposing an energetic stress. Zhang et al. (1959, cited in Zhou and Allen (2003)) reported that C. ariakensis (= Ostrea rivularis) had a low feeding incidence (0-70%) during the summer when water temperatures were high (22 30[degrees]C), and salinities were low and variable (2-26): clearance rates were not reported. Zhou and Allen (2003) suggest that the decrease in feeding incidence may be more closely related to salinity than to temperature. Kelly (2011) reported clearance rates of 1.10 L/g/h for C. ariakensis from the same stock as we studied during the summer (temperature, 25[degrees]C; salinity, ~ 10). This is not different from the clearance rate of 1.2 L/g/h we measured from the high-salinity location in Virginia, indicating that clearance rates in this species are apparently not affected by salinities in the range of 10 24. During the winter, C. ariakensis remained active, with individuals observed to be feeding, producing biodeposits, and putting on new shell growth even when water temperatures dropped to 2[degrees]C. Absorption efficiency was highest in the winter; therefore, even with a reduced clearance rate, C. ariakensis was still benefiting from its continual activity by assimilating a greater portion of the food they were ingesting, although the calculated scope for growth was negative (-1.02 J/g/h). From spring to summer, C. virginica had a positive scope for growth that was primarily influenced by a relatively high clearance rate during the summer, and low respiration rates during the remaining seasons. The summer clearance rate for C. virginica (2.22 L/g/h) was 50% higher than C. ariakensis (1.16 L/g/h) in our study, but lower than what has previously been reported in the literature (Loosanoff & Nomejko 1946, Jordan 1987, Newell & Langdon 1996). It is possible that the clearance rate of C. virginica was negatively impacted by the high amount of seston (49.3 mg/L) present in the experimental system during the summer. High particle concentrations (>25 mg/L) have been shown to decrease C. virginica clearance rates at temperatures greater than 20[degrees]C (Newell & Langdon 1996). No feeding activity or biodeposit production was observed for C. virginica during the winter. Continued respiration and ammonium excretion, albeit at low rates, resulted in a negative scope for growth for C. virginica during the winter.
The O:N ratio is a measure of the relative utilization of protein in energy metabolism (Corner & Cower 1968, Bayne et al. 1985). An O:N ratio less than 20 indicates stress in marine bivalves (Bayne et al. 1985, Huang & Newell 2002). The O:N ratio for both species of oysters measured in Virginia during the summer is low, but still greater than the level indicative of nutritive stress in either species. This relatively low summer ratio may be the result of an unobserved spawning event prior to or during the acclimation period. Postspawning oysters are generally in a poor condition because of the need to reorganize or regenerate tissue (Bayne et al. 1985). Because the O:N ratios of C. virginica and C. ariakensis were almost identical during the summer, it is unlikely that spawning-induced stress was the reason for differences in the scope for growth between the oyster species. The O:N ratio of C. ariakensis remained relatively low (<60) and consistent throughout the year, indicating a greater affinity for obtaining energy through protein degradation rather than lipid and/or carbohydrate catabolism, compared with C. virginica; the reasons for this remain unclear.
These physiological studies indicate that high water temperatures impose a stress on C. ariakensis that results in highly reduced feeding activity, and a concomitant reduced scope of growth. In temperate locations, such as the Chesapeake Bay, where high water temperature occurs only for two summer months, such stress may not be lethal. During the summer, oysters can use nutrients accumulated in cooler months when the oysters are actively feeding. In the subtropical IRL, where our mesocosm studies were performed, water temperatures were more than 20[degrees]C for 11 mo and more than 25[degrees]C for 6 mo (Table 1). The possibility that heat stress alone was responsible for the high mortalities suffered by C. ariakensis in our study appears to be at odds with the species' geographical distribution in its native habitat, where its range is reported to extend into subtropical regions in Asia (Zhou & Allen 2003, Guo et al. 2008).
There has been some confusion surrounding the identification of C. ariakensis in its native range (Zhou & Allen 2003). Zhang et al. (2005) compared genetic variation of C. ariakensis in U.S. hatchery stocks with wild Asian populations using polymerase chain reaction with restriction fragment length polymorphism to analyze the mitochondrial COI gene region and found genetic differentiation between "northern-type" and "southern-type" strains. Using similar genetic analysis, Guo et al. (2008) found that C. ariakensis was a dominant member of mixed assemblages of oysters at only five spatially isolated sites in China, and was present at low abundances throughout the rest of its range. Only one of those populations occurred in a subtropical climate; the remaining sites had annual temperature ranges similar to the temperate region of the east coast of the United States.
It is important to note that all C. ariakensis used in this and other studies of C. ariakensis in North America during the past decade descend from a small founder population consisting of 7 males and 9 females (U.S. Army Corps of Engineers, Norfolk District 2009). Although the exact details concerning the introduction of C. ariakensis to Oregon from Asia are unknown, they were first isolated in the late 1960s among Crassostrea gigas oysters being cultured in Yaquina Bay, Oregon, and then subsequently bred in Oregon (Breese & Malouf 1977, Malouf, Oregon Sea Grant, pers. comm.). Zhang et al. (2005) reported that 97% of C. ariakensis in U.S. hatchery stocks are most genetically similar to the "northern-type'" strain.
Given the highly restricted genetic makeup of the Oregon stock-introduced oysters, and the fact that they likely originated in the cooler temperate regions of Asia, it is perhaps not surprising that they exhibit low tolerance to subtropical warm waters. Potential future introduction of additional C. ariakensis from southern regions of Asia may result in a population of new oysters with a greater temperature tolerance. It is also possible that a greater temperature tolerance may evolve in the Oregon strain of C. ariakensis. The recent northward range extension of C. gigas in Europe has been attributed to increasing summer water temperatures sufficient to allow the species to reproduce in waters that were previously too cold (Wrange et al. 2010). However, it is also plausible that sufficient time has elapsed since C. gigas was introduced into Europe in the early 1970s for adaptations to have evolved that allow this species to inhabit cooler waters. There is evidence that such physiological adaptations to temperature exist in latitudinally separated and reproductively isolated populations of the blue mussel along the east coast of North America (Thompson & Newell 1985).
In summary, our results indicate that if C. ariakensis were to be either deliberately or accidentally introduced into the Chesapeake Bay, their expansion into U.S. subtropical regions may be limited. A depressed clearance rate resulting in a negative scope for growth under warm water conditions would result in reduced growth rates for C. ariakensis, which may make them less competitive against the native C. virginica. Eastern oysters in the subtropical regions of the United States are primarily intertidal (Grizzle 1990, Coen et al. 2007), which provides them with a refuge against predation (O'Beirn et al. 1996). The intertidal zone has been found to be inhospitable to C. ariakensis, with mortality rates often reaching 100% largely as a result of physiological stresses possibly caused by some combination of thermal intolerance and desiccation stress (Kingsley-Smith & Luckenbach 2008, Kingsley-Smith et al. 2009). Our observations of reduced clearance rates and a negative scope for growth for C. ariakensis in year-round subtropical waters would likely result in a growth rate much lower than native C. virginica. This would prevent juvenile C. ariakensis in the subtropical subtidal zone from rapidly reaching a size refuge against intense predation pressures, which (when coupled with their relatively fragile shell (Newell et al. 2007)) might serve to enhance predation rates on this species.
We thank Krystal Baird, Eman El-Wazzan, Patrick Monaghan, James Webb, and David Wood (HBOI-FAU) for assistance. Mark Luckenbach kindly allowed us to use facilities at VIMSESL, where Stephanie Bonniwell and Reid Bonniwell provided extensive logistical support. From Maryland DNR, Carol McCollough assessed feeding activity and pathologies histologically; and Judson Blazek, Stuart Lehmann, and Kristi Shaw processed histological samples. Christopher Dungan of Maryland DNR and Ryan Carnegie of VIMS kindly provided results from their Bonamia sp. analyses. Funding for this project was provided by NOAA project no. NA06NMF4570245. This is HBOI-FAU contribution number 1835.
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CHRISTOPHER J. KELLY, (1) SUSAN E. LARAMORE, (2) JOHN SCARPA (2) AND ROGER I. E. NEWELL (1)
(1) Horn Point Laboratory, University of Maryland Center for Environmental Science, 2020 Horn Point Road, Cambridge, MD 21613; (2) Harbor Branch Oceanographic Institute at Florida Atlantic University, Aquaculture and Stock Enhancement Program, 5600 North U.S. Hwy 1, Fort Pierce, FL 34946
* Corresponding author. E-mail: firstname.lastname@example.org
TABLE 1. Mean (n = 3) water temperature, salinity, dissolved oxygen concentration (DO), and percent dissolved oxygen saturation (%DO) of ambient flow-through water in Florida mesocosms from September 2006 through August 2007. Temperature Month ([degrees]C) Salinity DO (mg/L) %DO September 2006 28.9 31.0 5.4 82.9 October 2006 25.8 32.0 6.0 87.1 November 2006 21.3 32.9 6.7 91.2 December 2006 21.8 33.2 6.4 87.9 January 2007 21.0 33.0 6.3 84.3 February 2007 18.6 32.8 6.9 89.5 March 2007 22.0 35.0 6.1 85.8 April 2007 22.9 36.2 6.1 87.1 May 2007 25.6 37.7 5.5 82.9 June 2007 28.3 35.7 5.1 79.0 July 2007 30.1 31.6 4.8 75.1 August 2007 30.4 28.0 4.8 74.1 TABLE 2. Mean absolute size ([+ or -] SE) and mean specific growth rate (SPG; [+ or -] SE) of 28 and 4-mo-old Crassostrea ariakensis and Crassostrea virginica in Florida mesocosms between September 2006 and August 2007. Absolute size Species Age Month ([mm.sup.2]) C. virginica 4 mo September 97.1 [+ or -] 8.7 December 212.2 [+ or -] 16.8 January 230.8 [+ or -] 17.4 February 237.3 [+ or -] 18.2 March 258.8 [+ or -] 18.1 April 285.9 [+ or -] 19.6 May 322.2 [+ or -] 26.9 June 368.1 [+ or -] 34.6 July 376.5 [+ or -] 33.8 August 387.9 [+ or -] 42.3 C. ariakensis 4 mo September December 249.5 [+ or -] 29.4 January 261.9 [+ or -] 30.3 February 273.9 [+ or -] 30.9 March 278.5 [+ or -] 31.9 April 304.2 [+ or -] 34.6 May 336.8 [+ or -] 37.9 June 376.8 [+ or -] 39.1 July 405.8 [+ or -] 36.0 August C. virginica 28 mo September 732.2 [+ or -] 32.9 December 894.4 [+ or -] 56.4 January 903.5 [+ or -] 55.2 February 934.5 [+ or -] 53.8 March 971.8 [+ or -] 66.3 April 996.6 [+ or -] 68.2 May 1,073.3 [+ or -] 84.7 June 1,068.6 [+ or -] 100.7 July 1,117.2 [+ or -] 111.3 August 1,173.0 [+ or -] 108.5 C. ariakensis 28 mo September 2,205.9 [+ or -] 245.5 December 2,262.7 [+ or -] 240.9 January 2,329.1 [+ or -] 272.7 February 2,318.1 [+ or -] 257.6 March 2,199.7 [+ or -] 317.2 April 1,450.9 [+ or -] 287.6 May 1,329.7 [+ or -] 267.2 June 866.1 [+ or -] N/A July August SPG [ln increase shell area Species Age Month ([mm.sup.2]/day)] C. virginica 4 mo September December 137.6 x [10.sup.-4] [+ or -] 19.8 x [10.sup.-4] January 12.8 x [10.sup.-4] [+ or -] 4.7 x [10.sup.-4] February 8.9 x [10.sup.-4] [+ or -] 3.7 x [10.sup.-4] March 19.7 x [10.sup.-4] [+ or -] 5.0 x [10.sup.-4] April 16.4 x [10.sup.-4] [+ or -] 7.5 x [10.sup.-4] May 35.7 x [10.sup.-4] [+ or -] 11.2 x [10.sup.-4] June 43.5 x [10.sup.-4] [+ or -] 14.6 x [10.sup.-4] July 17.4 x [10.sup.-4] [+ or -] 9.3 x [10.sup.-4] August 10.9 x [10.sup.-4] [+ or -] 10.9 x [10.sup.-4] C. ariakensis 4 mo September December January 21.0 x [10.sup.-4] [+ or -] 7.0 x [10.sup.-4] February 17.4 x [10.sup.-4] [+ or -] 12.7 x [10.sup.-4] March 4.7 x [10.sup.-4] [+ or -] 2.4 x [10.sup.-4] April 21 x [10.sup.-4] [+ or -] 9.4 x [10.sup.-4] May 35.3 x [10.sup.-4] [+ or -] 15.9 x [10.sup.-4] June 28.9 x [10.sup.-4] [+ or -] 13.3 x [10.sup.-4] July 28.7 x [10.sup.-4] [+ or -] 12.3 x [10.sup.-4] August C. virginica 28 mo September December 35.6 x [10.sup.-4] [+ or -] 11.8 x [10.sup.-4] January 3.0 x [10.sup.-4] [+ or -] 2.1 x [10.sup.-4] February 11.0 x [10.sup.-4] [+ or -] 4.0 x [10.sup.-4] March 13.7 x [10.sup.-4] [+ or -] 6.3 x [10.sup.-4] April 6.1 x [10.sup.-4] [+ or -] 4.5 x [10.sup.-4] May 12.0 x [10.sup.-4] [+ or -] 6.1 x [10.sup.-4] June 5.2 x [10.sup.-4] [+ or -] 5.2 x [10.sup.-4] July 5.3 x [10.sup.-4] [+ or -] 4.9 x [10.sup.-4] August 0.0 [+ or -] 0.0 C. ariakensis 28 mo September December 8.2 x [10.sup.-4] [+ or -] 4.1 x [10.sup.-4] January 3.6 x [10.sup.-4] [+ or -] 1.6 x [10.sup.-4] February 3.9 x [10.sup.-4] [+ or -] 2.3 x [10.sup.-4] March 6.6 x [10.sup.-4] [+ or -] 4.0 x [10.sup.-4] April 7.3 x [10.sup.-4] [+ or -] 7.3 x [10.sup.-4] May 0.0 [+ or -] 0.0 June 0.0 [+ or -] 0.0 July August P [less than Species Age Month n or equal to] 0.05 C. virginica 4 mo September 52 December 42 a January 46 d February 52 cd March 51 cbd April 41 cbd May 29 cb June 25 b July 28 cbd August 23 cbd C. ariakensis 4 mo September December 26 January 26 ns February 26 ns March 26 ns April 25 ns May 16 ns June 12 ns July 4 ns August 0 C. virginica 28 mo September 14 December 14 a January 14 b February 14 ab March 13 ab April 12 b May 9 ab June 9 ab July 8 ab August 6 b C. ariakensis 28 mo September 16 December 16 ns January 15 ns February 15 ns March 10 ns April 4 ns May 3 ns June 1 ns July 0 August 0 Different letters denote significant differences in growth rate within species of the same age class at [alpha] = 0.05. N/A, not applicable; ns, no difference (Wilcoxon's signed rank sum pairwise comparisons). TABLE 3. Reproductive and gonadal index data for Crassostrea ariakensis and Crassostrea virginica reared in Florida mesocosms. % Gonadal % Species Age Month n Development Differentiated C. virginica 4 mo February 4 25 0 March 4 100 0 April 3 66 0 May 9 89 0 June 2 50 0 July 11 91 0 August 4 25 0 C. ariakensis 4 mo February 9 78 0 March 10 60 0 April 14 21 0 May 15 93 0 June 16 81 0 July 13 92 8 August 0 C. virginica 28 mo February 8 63 0 March 8 100 0 April 7 43 0 May 8 100 50 June 10 80 30 July 3 100 33 August 5 80 60 C. ariakensis 28 mo February 11 91 0 March 9 100 0 April 3 33 0 May 5 80 0 June 4 50 0 July 1 100 0 August 0 Percent gonadal development indicates number of oysters that showed evidence of gametogenesis but no identifiable gender. Percent differentiated indicates number of oysters that had clearly distinguishable eggs or sperm. TABLE 4. Mean (n = 3) temperature, salinity, total suspended solids (TSS [+ or -] SE), and percent particulate organic matter (POM [+ or -] SE) of ambient flow-through water at VIMS-ESL during a 7-day period when seasonal physiological measurements were performed. Temperature Season ([degrees]C) Salinity TSS (mg [L.sup.-1]) July 2008 27 29.8 49.3 [+ or -] 7.2 October 2008 14 29.4 13.8 [+ or -] 3.7 January 2009 5 32 17.9 [+ or -] 0.9 April 2009 14 30 18.7 [+ or -] 1.4 Season POM (%) July 2008 9.3 [+ or -] 1.3 October 2008 16 [+ or -] 3.9 January 2009 17.1 [+ or -] 1.0 April 2009 16.2 [+ or -] 1.1 TABLE 5. Seasonal clearance, respiration, and ammonium excretion rates (a) for Crassostrea ariakensis and Crassostrea virginica measured in ambient flow-through conditions in Virginia. Rate Species b Season n a Clearance C. virginica 0.44 * Jul. 2008 15 2.22 rate Oct. 2008 14 1.86 (L/g/h) Jan. 2009 16 0 Apr. 2009 14 1.50 C. ariakensis 0.62 Jul. 2008 14 1.16 Oct. 2008 16 1.54 Jan. 2009 15 0.18 Apr. 2009 16 0.67 Respiration C. virginica 1.07 Jul. 2008 16 2.52 (mL Oct. 2008 16 0.78 [O.sub.2]/ Jan. 2009 4 0.22 g/h) Apr. 2009 11 0.51 C. ariakensis 0.59 Jul. 2008 13 2.60 Oct. 2008 15 0.70 Jan. 2009 9 0.32 Apr. 2009 14 0.61 Ammonium C. virginica 0.57 Jul. 2008 16 77.30 excretion ([micro]g Oct. 2008 14 5.57 N[H.sub.4]- Jan. 2009 6 2.14 N/g/h) Apr. 2009 14 11.65 C. ariakensis 0.44 Jul. 2008 14 72.33 Oct. 2008 16 25.19 Jan. 2009 11 6.18 Apr. 2009 15 11.36 P [less than or Rate Species b Season equal to] 0.05 Clearance C. virginica 0.44 * Jul. 2008 ns rate Oct. 2008 ns (L/g/h) Jan. 2009 Apr. 2009 ns C. ariakensis 0.62 Jul. 2008 a Oct. 2008 a Jan. 2009 c Apr. 2009 b Respiration C. virginica 1.07 Jul. 2008 a (mL Oct. 2008 b [O.sub.2]/ Jan. 2009 c g/h) Apr. 2009 bc C. ariakensis 0.59 Jul. 2008 a Oct. 2008 b Jan. 2009 c Apr. 2009 b Ammonium C. virginica 0.57 Jul. 2008 a excretion ([micro]g Oct. 2008 bc N[H.sub.4]- Jan. 2009 c N/g/h) Apr. 2009 bc C. ariakensis 0.44 Jul. 2008 a Oct. 2008 b Jan. 2009 d Apr. 2009 c Rate Species b Season Tissue weight (g) Clearance C. virginica 0.44 * Jul. 2008 0.93 [+ or -] 0.69 rate Oct. 2008 0.97 [+ or -] 0.51 (L/g/h) Jan. 2009 0.94 [+ or -] 0.47 Apr. 2009 0.92 [+ or -] 0.30 C. ariakensis 0.62 Jul. 2008 1.06 [+ or -] 0.96 Oct. 2008 0.71 [+ or -] 0.48 Jan. 2009 1.08 [+ or -] 0.68 Apr. 2009 1.68 [+ or -] 0.97 Respiration C. virginica 1.07 Jul. 2008 0.88 [+ or -] 0.69 (mL Oct. 2008 1.01 [+ or -] 0.51 [O.sub.2]/ Jan. 2009 1.04 [+ or -] 0.58 g/h) Apr. 2009 0.95 [+ or -] 0.27 C. ariakensis 0.59 Jul. 2008 1.13 [+ or -] 0.96 Oct. 2008 0.73 [+ or -] 0.48 Jan. 2009 1.01 [+ or -] 0.70 Apr. 2009 1.82 [+ or -] 0.94 Ammonium C. virginica 0.57 Jul. 2008 0.88 [+ or -] 0.69 excretion ([micro]g Oct. 2008 1.04 [+ or -] 0.51 N[H.sub.4]- Jan. 2009 1.00 [+ or -] 0.47 N/g/h) Apr. 2009 0.96 [+ or -] 0.30 C. ariakensis 0.44 Jul. 2008 1.06 [+ or -] 0.96 Oct. 2008 0.71 [+ or -] 0.48 Jan. 2009 1.13 [+ or -] 0.69 Apr. 2009 1.74 [+ or -] 0.97 Rate Species b Season Range (g) Clearance C. virginica 0.44 * Jul. 2008 0.17-2.15 rate Oct. 2008 0.34-2.03 (L/g/h) Jan. 2009 0.23-1.50 Apr. 2009 0.38-1.41 C. ariakensis 0.62 Jul. 2008 0.17-3.37 Oct. 2008 0.19-1.56 Jan. 2009 0.22-2.16 Apr. 2009 0.63-4.26 Respiration C. virginica 1.07 Jul. 2008 0.17-2.15 (mL Oct. 2008 0.34-2.03 [O.sub.2]/ Jan. 2009 0.23-1.50 g/h) Apr. 2009 0.46-1.41 C. ariakensis 0.59 Jul. 2008 0.17-3.37 Oct. 2008 0.19-1.56 Jan. 2009 0.22-2.12 Apr. 2009 0.68-4.26 Ammonium C. virginica 0.57 Jul. 2008 0.17-2.15 excretion ([micro]g Oct. 2008 0.34-2.03 N[H.sub.4]- Jan. 2009 0.23-1.50 N/g/h) Apr. 2009 0.38-1.41 C. ariakensis 0.44 Jul. 2008 0.17-3.37 Oct. 2008 0.19-1.56 Jan. 2009 0.22-2.12 Apr. 2009 0.63-4.26 Rates are standardized to an oyster with a 1-g dry tissue weight by the allometric equation Y = [aX.sup.b] (see text for details). Common slope (b) for clearance rate of C. virginica (*) does not include winter (January) 2009, because these oysters did not feed for the duration of the experiment during this season. Different letters denote significant difference in corrected a values for each species among seasons. ns, no difference (LSD pairwise comparisons, P [less than or equal to] 0.05). Mean ([+ or -] SD) and range of oyster dry tissue weights are shown for each season. TABLE 6. Mean ([+ or -] SE) seasonal percent absorption efficiency (Ae) of Crassostrea ariakensis and Crassostrea virginica in ambient flow- through conditions at VIMS-ESL. Species Season n Ae (%) [+ or -] SE C. virginica July 2008 11 25.5 [+ or -] 3.3 October 2008 16 37.1 [+ or -] 2.8 January 2009 15 0 [+ or -] 0 April 2009 14 44.1 [+ or -] 4.6 C. ariakensis July 2008 7 15.4 [+ or -] 2.8 October 2008 14 29.4 [+ or -] 3.3 January 2009 10 43.5 [+ or -] 3.8 April 2009 16 36.5 [+ or -] 4.5 Back-Transformed Average (%) [+ or -] SE P [less than Species Season Ae +SE -SE or equal to] 0.05 C. virginica July 2008 24.5 3.8 3.6 b October 2008 36.5 3.5 3.4 a January 2009 0 0 0 c April 2009 43.5 3.8 3.8 a C. ariakensis July 2008 14.9 4.1 3.6 c October 2008 28.7 3.5 3.4 b January 2009 43.4 4.5 4.4 a April 2009 35.2 3.5 3.4 ab Different letters denote significance of back-transformed means ([+ or -] SE) for each species among seasons (LSD pairwise comparisons, P [less than or equal to] 0.05).
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