Early larval development of the Sydney rock oyster Saccostrea glomerata under near-future predictions of C[O.sup.2]-driven ocean acidification.

ABSTRACT Anthropogenic emissions of carbon dioxide (CO2) from fossil fuel combustion and deforestation are rapidly increasing the atmospheric concentration of C[O.sub.2] and reducing the pH of the oceans. This study shows that predicted near-future levels of ocean acidification have significant negative effects on early larval development of the Sydney rock oyster Saccostrea glomerata (Gould, 1850). C[O.sub.2] was added to seawater to produce pH levels set at 8.1 (control), 7.8, and 7.6 (actual pH values were 8.11, 7.81, and 7.64, respectively). These treatments represent present-day surface ocean pH, as well as upper (A pH [equivalent to] -0.3) and lower (A pH [equivalent to] -0.5) pH predictions for the surface oceans in 2100. With decreasing pH, survival of S. glomerata larvae decreased, and growth and development were retarded. Larval survival decreased by 43% at pH 7.8 and by 72% at pH 7.6. Antero-posterior measurement (APM) was reduced by 6.3% at pH 7.8 and 8.7% at pH 7.6, and dorso-ventral measurement (DVM) was reduced by 5.1% at pH 7.8 and 7.5% at pH 7.6. The percentage of empty shells remaining from dead larvae decreased by 16% at pH 7.8 and by 90% at pH 7.6 indicating that the majority of empty shells dissolved within 7 days at pH 7.6. Scanning electron microscope images of 8-day-old larvae show abnormalities on the shell surface at low pH suggesting (1) problems with shell deposition, (2) retarded periostracum formation, and/or (3) increased shell dissolution. Larval life-history stages are considered particularly susceptible to climate change, and this study shows that S. glomerata larvae are sensitive to a high-C[O.sub.2] world and are, specifically, negatively affected by exposure to pH conditions predicted for the world's oceans for the year 2100.

KEY WORDS: climate change, ocean acidification, carbon dioxide, oyster, survival, larval development

INTRODUCTION

Over the last 23 million years, atmospheric carbon dioxide (C[O.sub.2]) concentrations have remained stable between 140 to 340 ppm and surface ocean pH has remained constant between 8.0 to 8.3 pH units (Pearson & Palmer 2000). Anthropogenic C[O.sub.2] emitted from fossil fuel combustion and deforestation since the onset of the Industrial Revolution has increased atmospheric concentrations of C[O.sub.2] from 280 to 380 ppmv in the last 200 y (Tans NOAA/ESRL records). This rapid increase in C[O.sub.2] of 35% is an order of magnitude faster than any change in C[O.sub.2] concentration that has occurred for millions of years (Doney & Schimel 2007). The increase in this greenhouse gas is not only causing the planet to warm but is also causing the oceans to acidify. This acidification is the result of increased C[O.sub.2] dissolving in seawater decreasing carbonate ion (C[O.sub.3.sup.2-]) concentration and increasing bicarbonate ion (HCO3) concentration in a series of equilibrium reactions that release hydrogen ions ([H.sup.+]).

Over the past 200 y, the oceans have taken up ~40% of the anthropogenic C[O.sub.2] emissions from the atmosphere (Zeebe et al. 2008). This uptake of C[O.sub.2] has caused a 0.1 unit drop in the pH of the surface oceans (Caldeira & Wickett 2003) which is equivalent to a 30% increase in [H.sup.+] concentration (Blackford & Gilbert 2007). By the 1980s, ocean pH was falling at a rate of 0.015 pH units per decade (Haugan & Drange 1996) and present day average global surface ocean pH is 8.07 compared with the preindustrial value of 8.17 (Cao et al. 2007).

Models predict a further reduction in ocean pH by 0.3 to 0.5 units by 2100 (Haugan & Drange 1996, Caldeira & Wickett 2005, Orr et al. 2005), which is equivalent to at least a 100-150% increase in [H.sup.+] concentration (Orr et al. 2005). Some reports suggest that increasing global temperatures caused by climate change will partially alleviate the effects of ocean acidification; however, Cao et al. (2007) show that ocean acidification is largely independent of temperature change. The predicted changes in ocean pH are not only greater but far more rapid than any experienced in the last 24 million years (see Blackford & Gilbert 2007) and possibly the last 300 million years (Caldeira & Wickett 2003).

Ocean acidification is likely to affect marine animals either by direct acidification of their environment or through hypercapnia. The decreasing pH of the oceans will lower the saturation state of seawater with respect to calcium carbonate and therefore increase the solubility of calcareous structures. Consequently, calcifying species such as corals, foraminifera, coccolithophores, molluscs, brachiopods, and echinoderms are thought to be particularly at risk. Kleypas et al. (2006) suggest that calcification rates could decrease by 60% this century. Under experimental conditions of the predicted atmospheric pC[O.sub.2] (-740 ppmv) in 2100 from the IPCC IS92a scenario, calcification was reduced by 25% in the edible mussel Mytilus edulis and 10% in the Pacific oyster Crassostrea gigas (Gazeau et al. 2007).

Other negative effects of ocean acidification have been demonstrated in various species, including decreased growth in the sea urchins Hemicentrotus pulcherrimus and Echinometra mathaei and the gastropod Strombus luhuanus (Shirayama & Thornton 2005), and reduced fertilization success in the sea urchin Heliocidaris erythrogramma (Havenhand et al. 2008). Experiments investigating the effects of seawater acidification at low pH (under scenarios of leakage from geological C[O.sub.2] sequestration stores) have demonstrated increased mortality in the sea urchin Psammechinus miliaris (Miles et al. 2007) and complex indirect effects on the induced defenses in the gastropod Littorina littorea (Bibby et al. 2007).

Lowered seawater pH reduces calcification in coral reefs (Langdon et al. 2003) and the combination of stresses of climate change and ocean acidification is expected to reduce coral reef system diversity, with corals becoming increasingly rare (Hoegh-Guldberg et al. 2007). Naturally occurring gradients of reduced pH around C[O.sub.2] vents in the Mediterranean Sea, have demonstrated ecosystem shifts and changes in species composition with a dramatic decrease in calcifying fauna in low pH areas (Hall-Spencer et al. 2008).

Understanding the effects of acidification in commercially important species is of prime importance. The Sydney rock oyster Saccostrea glomerata (Gould, 1850) was chosen as a model species for this preliminary study. It is an important commercial species supporting an industry that currently generates $34.5 million (AUD) annually in sales (O'Sullivan et al. 2008). S. glomerata is a warm temperate bivalve mollusc endemic to Australia and New Zealand. Dove and Sammut (2007) showed that acidification of estuarine waters from terrestrial acid sulphate soil runoff increased mortality in S. glomerata. Small oysters showed the greatest mortality because their thinner shells were perforated by acid erosion.

Early life-history stages are generally believed to be vulnerable to stress and change (Peck 2005). Studies have already shown that the early developmental stages of the oyster Crassostrea gigas (Kurihara et al. 2007), copepod Calanus finmarchicus (Mayor et al. 2007) and brittlestar Ophiothrix fragilis (Dupont et al. 2008) are negatively affected by acidified conditions. We aimed to identify the effects of near-future C[O.sub.2]-driven ocean acidification scenarios on the larval development of S. glomerata, and determine if this life-history stage is particularly susceptible to the stress of altered pH.

[FIGURE 1 OMITTED]

MATERIALS AND METHODS

Experimental Set-up and pH Control

Experiments were conducted at the Aquarium Complex of the Marine and Aquaculture Research Facilities Unit (MARFU) at James Cook University, Australia. Seawater obtained from the Australian Institute for Marine Science (AIMS) was filtered to 1 [micro]m and sterilized by ultra-violet radiation. Seawater was held in a 3,000-L reservoir tank in the laboratory and allowed to equilibrate to ambient temperature (26[degrees]C). Seawater was pumped from the reservoir into three 500-L header tanks, which filled on-demand via float valves (Fig. 1).

The pH of seawater was controlled by manipulation of the partial pressure of C[O.sub.2] (pC[O.sub.2]) reproducing the means by which atmospheric C[O.sub.2] will reduce the pH of the oceans. Seawater pH was regulated by a computerized control system (AquaMedic AT-Control) that introduced gaseous C[O.sub.2] into the header tanks as required via a solenoid valve, pH probes were calibrated with NBS buffer solutions and the pH values reported in this study are on the NBS scale. Values of pH were nominally set at 8.1 (control), 7.8, and 7.6. These treatments represent present-day pH, upper year 2100 pH predictions (A pH [equivalent to] -0.3) and lower year 2100 pH predictions ([DELTA] pH [equivalent to] 0.5) of seawater, respectively. Seawater from the header tanks was gravity-fed (2.5 1.[h.sup.1]) to individual 60-L treatment tanks within 3,000-L water-bath tanks. The actual pH values measured in the treatment tanks were 8.11,7.81, and 7.64 (Table 1). Seawater samples were analyzed for total alkalinity by titration. Other parameters of the carbonate system were calculated using the program CO2SYS (Lewis & Wallace 1998).

Seawater temperature was maintained at 26 [degrees]C (to 2 significant figures) based on optimum temperatures reported for growth, development, and survival of embryos and larvae of S. glomerata (Dove & O'Connor 2007). Salinity was maintained at 32 and seawater within the treatment tanks was mixed by aeration through a fine glass tube. Larvae were retained in the treatment tank by a fine mesh, which covered the outflow pipe (Southgate & Beer 1997). Used seawater was not recirculated into the experimental system. Treatment tanks were covered with transparent lids to minimize loss of C[O.sub.2] from the seawater to the atmosphere and were held under a 13 hL:11 hD lighting regime.

Spawning Procedure and Larval Culture

Sydney rock oysters were obtained from a commercial oyster farm in central New South Wales, Australia and strip-spawned. Embryos were incubated in a 500-L tank filled with 1-[micro]m filtered seawater containing the antibiotic streptomycin sulphate at a concentration of 10 mg.[L.sup.-1]. At 24 h old, the larvae had reached D-veliger stage. Larvae were retained on a 25-[micro]m mesh screen, washed with 1-[micro]n filtered seawater and 90,000 larvae were transferred into each 60-L treatment tank creating a stocking density of 1.5 larvae.mL L in each. In the treatment tanks, larvae were pulse fed daily with live cultured microalgae composed of a 1:1 mixture of Pavlova salina and Isochrvsis (clone T-[so) fed at increasing concentrations of 5,000-20,000 cells.[mL.sup.-1] from days 1-8. Feeding rate was based on previously published data for the larvae of S. glomerata (Nell & O'Connor 1991) and sufficient concentrations of algal cells were fed to ensure food supply was not a limiting factor in survival, growth and development.

Processing of Larvae

Larvae were removed from the treatment tanks when they were 8 days old by passing the contents of each treatment tank through a 37-[micro]m mesh screen. Larvae were counted on a Sedgewick rafter counter using a light microscope. The anteroposterior measurement (APM) and dorso-ventral measurement (DVM) of 30 larvae per treatment were recorded. Further samples of larvae were frozen, freeze-dried, and then weighed or fixed in a 2% gluteraldehyde solution in seawater for 2 h. Fixed larvae were dehydrated through graded ethanol solutions, air-dried and transferred onto stainless steel stubs. The stubs and larvae were gold coated and imaged on a JEOL JSM-5410LV scanning electron microscope (SEM) using secondary electrons.

Empty Shell Counts

Shells of both living and dead calcareous animals experience dissolution in the marine environment (see Harper 2000), so it was expected that some empty, or 'dead', larval shells would dissolve at each pH. To determine differences in shell dissolution between treatments, the percentage of empty shells remaining from larvae that had died during the experiment in each treatment tank was calculated. The number of larvae that had died, and therefore the number of empty shells that should have been present, was determined by subtraction of counts of surviving larvae at the end of an experiment from original stocking numbers. The actual numbers of empty shells remaining were counted on a Sedgewick rafter counter using a light microscope. The percentage of empty shells remaining was then calculated by dividing the number of empty shells counted by the number of dead larvae per treatment tank (Eq. 1), thus correcting for differences in larval mortality between treatments.

Percent empty shells remaining

= empty shells counted/number of dead larvae x 100% (1)

Statistical Analysis

Data were tested for normality where appropriate using the Shapiro-Wilk normality test. Relevant parametric (e.g., ANOVA, Pearson correlation) or nonparametric (e.g., Kruskal Wallis) statistical analyses were performed using SPSS 16.0.

RESULTS

In the following sections, nominal pH values are used to refer to the treatment pH conditions (i.e., pH 8.1, 7.8, and 7.6).

For actual pH values measured in the treatments and further water chemistry data, see Table 1.

Larval Survival

One-day-old D-veliger larvae were introduced to the treatment tanks. By 8-days-old, the majority had developed into umbonate larvae. The experiment was thus run between these two key developmental stages. Percentage larval survival decreases with decreasing pH (Fig. 2A). There is a significant positive correlation between pit and mean larval survival (r = 1.000, P = 0.004, n : 3 for nominal pH: r = 1.000, P - 0.032, n = 3 for measured pH). Compared with the control treatment (pH 8.1), survival decreased by 43% at pH 7.8 and by 72% at pH 7.6.

Larval Growth and Morphology

Decreased pH had an effect on the morphology and growth of the larvae. There was a significant difference between the antero-posterior measurement (APM) of larvae at different pH ([chi square] = 13.697, df 2, P = 0.001) and between the dorsoventral measurement (DVM) of larvae at different pH ([chi square] 12.276, df= 2, P = 0.002). Larval APM was reduced by 6.3% at pH 7.8 and 8.7% at pH 7.6 (Fig. 2B). Larval DVM was reduced by 5.1% at pH 7.8 and 7.5% at pH 7.6 (Fig. 2C). It was expected that pH would affect larval dry mass (Fig. 2D). However, preliminary mass data were limited and statistical tests showed no significant difference in mass among pH treatments.

[FIGURE 2 OMITTED]

Empty Shells Recovered

There was a significant difference in the percentage of empty shells remaining between pH treatments (F = 50.129, df = 2,7, P < 0.001). At pH 7.8 and 7.6, the saturation state of seawater with respect to calcite and aragonite fell below [OMEGA] = 1 (Table 1). The percentage of empty shells remaining decreased with decreasing pH by 16% in pH 7.8 and 90% in pH 7.6 (Fig. 2E).

Shell Surface Structure

Shell surface characteristics were markedly affected by pH (Fig. 3). In normal larval shell development (control treatment pH 8.1) the shell is smooth and rounded with even growth (Fig. 3A). At pH 7.8 and pH 7.6, growth checks can be seen on the surface of the larval shell (marked by "X" arrows) and are likely to be the result of physiological stress after the transfer to low pH conditions (Figs. 3B and 3C). Larval shell deposition under low pH (indicated by "Y"), was no longer smooth, but appeared pitted and deformed particularly around the valve edge. These abnormalities in new shell growth indicate potential problems with shell deposition and/or periostracum formation. At low pH, areas in the middle of the shell surface, which were deposited before transfer to low pH conditions, showed surface pitting and were rough in appearance indicating dissolution of the larval shell after transfer to low pH conditions.

DISCUSSION

Increasing anthropogenic emissions of C[O.sub.2] have already caused a decrease in ocean pH of 0.1 units (Caldeira & Wickett 2003). Since the chemistry of seawater and the carbonate system is well known, we can be moderately confident about model predictions for future ocean pH scenarios of [DELTA] pH -0.3 to -0.5 units by 2100 (Haugan & Drange 1996, Caldeira & Wickett 2005, Orr et al. 2005) and -0.7 units by 2300 (Caldeira & Wickett 2003).

Experiments were run with S. glomerata from 1 to 8 days old between key developmental stages: D-veliger to umbonate larvae. Compared to the control treatment (pH 8.1), survival decreased by 43% in pH 7.8 (A pH 0.3 upper 2100 prediction) and 72% in pH 7.6 ([DELTA] pH -0.5 lower 2100 prediction). Although mortality during larval culture in the experiment was high across all treatments, commercial scale hatcheries also experience mass mortalities (i.e. >80%) of S. glomerata larvae. For example, half of all hatchery runs with this species over a five-year period failed within the first 8 days (Heasman et al. 2000, Dove & O'Connor 2007). Furthermore, strip spawning of oysters is generally acknowledged to result in fewer viable larvae compared with natural or induced spawning methods (Gosling 2003).

Larval growth (APM and DVM) was retarded with decreasing pH suggesting that larval development could take longer under ocean acidification, or that larvae will be smaller at each developmental stage. SEM images revealed growth abnormalities in the surface of the larval shell at low pH. Larval shells were pitted and deformed, particularly where shell was deposited during exposure to low pH. These shell surface abnormalities indicate larvae may have been experiencing: (1) problems with shell deposition, (2) retarded periostracum formation, and/or (3) increased shell dissolution, at low pH.

Initially oyster larvae deposit a predominantly amorphous calcium carbonate (ACC) shell, which later partially transforms into aragonite (Weiss et al. 2002). This differs from the adult oyster shell, which is composed predominantly of calcite (Stenzcl 1964). ACC is more soluble than aragonite (Breeevic & Nielsen 1989), which in turn is more soluble than calcite (Vermeij 1993). Therefore the larval shell, particularly during the ACC phase, is the most susceptible shelled life-history stage of the oyster to dissolution caused by ocean acidification.

[FIGURE 3 OMITTED]

Empty shell count data show that the percentage of empty shells, left by larvae, which had died during the experiment, decreased considerably at pH 7.6. These data suggest the empty shells dissolved rapidly (within 7 days) at pH 7.6. In this experiment, calcite and aragonite were undersaturated ([OMEGA] < 1) at pH 7.8 and 7.6 putting these minerals into a dissolution state. However, calculations of saturation state are based on nonbiogenic minerals in artificial seawater. Biogenic calcium carbonate shell contains an organic matrix and is often covered by a periostracum. Therefore the tipping point for dissolution of biological CaC[O.sub.3] may not occur at exactly [OMEGA] = 1 for pure calcite or aragonite. In the experiment, there seemed to be a threshold between pH 7.8 and 7.6 at which rapid shell dissolution occurred and the parsimonious explanation is that the saturation state of seawater with respect to the larval shell itself was [OMEGA] < 1, and markedly so in the pH 7.6 treatment.

Larval dry mass was lowest in the most acidic treatment. This probably indicates a marked fall in larval soft tissue mass at pH 7.6 because dry mass in this treatment was approximately half that of the higher pH treatments and shell pitting clearly did not produce that level of decrease in shell mass (Fig. 3): this suggests a marked effect on energy balance at low pH either through enhanced costs or reduced feeding abilities. Greatest larval mass was recorded at pH 7.8. This may be because the larvae that were able to survive in this treatment were the healthiest and thus heaviest; however, further studies are required to confirm this.

The results of this study demonstrate negative effects on larval survival and size and increased shell abnormalities of S. glomerata in a high-C[O.sub.2] environment. Similar effects on early life-history stages of other marine invertebrates have been demonstrated. At low pH (7.4), reduced larval development and shell mineralization was shown in the oyster Crassostrea gigas (Kurihara et al. 2007). Other negative effects caused by low pH include reduced hatching rate and nauplius development in copepods (Kurihara et al. 2004) and decreased fertilization and larval size in sea urchins (Kurihara & Shirayama 2004). Dupont et al. (2008) showed a reduction in larval survival and size, as well as increased larval abnormalities in the brittlestar Ophiothrix fragilis at pH 7.9 and 7.7. Havenhand et al. (2008) found decreased sperm speed and motility and reduced developmental success in embryos and larvae of the sea urchin Heliocidaris erythrogramma at pH 7.7. Negative effects of ocean acidification on the early developmental stages of any species are likely to threaten its long-term viability by a reduction in recruitment of viable juveniles to the population.

Early reports assessing the effects of CO, on marine organisms have considered results of experiments that used mineral acid addition to lower the pH of seawater (see Auerbach et al. 1997). However, reducing pH by C[O.sub.2] addition compared with acid addition can have greater negative effects on marine organisms for a given pH (Kikkawa et al. 2004, Kurihara & Shirayama 2004). In recent years, there has been a move to manipulate pH by the addition of C[O.sub.2] (e.g., Kurihara et al. 2004, Gazeau et al. 2007; Havenhand et al. 2008, Dupont et al. 2008), thus creating slightly more realistic conditions in terms of the overall carbonate chemistry of seawater. This experiment aimed to create realistic near-future conditions of ocean acidification scenarios predicted to occur before the end of the century by the addition of C[O.sub.2] to seawater.

The through-flow system of elevated C[O.sub.2] seawater was a major advantage in this study, avoiding the need to use small containers in closed systems and providing the ability to rear larvae in acidified seawater for an extended period during their development. Using a closed system introduces organisms to chemical stress from changes in water quality, by the depletion of major nutrients and ions (such as [Ca.sup.2+]) and by the build up of waste products. Closed systems also induce physical stress during water changes. Our experiments allowed the continuous change of well-mixed seawater, which removed the need to handle larvae until the experiment was finished. One-Bm filtered pristine seawater was used continuously, avoiding potential water quality issues with recirculated water from the experiment. An additional advantage included the use of header tanks, avoiding the potential creation of pH gradients and/or considerable pH fluctuations in the experimental tanks. Manipulation of pH within the large (500 L) header tanks meant any fluctuations in pH caused by the C[O.sub.2] solenoid valves switching on and off, were not passed onto the treatment tanks. Large treatment tanks (60 L) also minimized any variation in physical and chemical parameters throughout the experiment.

This study demonstrates that exposure to predicted upper and lower ocean pH levels for the end of the century has a dramatic negative effect on the survival, growth, and shell formation of the early larval stages of the Sydney rock oyster, S. glomerata. Targeting a potentially vulnerable stage in a species' life-history is useful, but it is also important to research the effects of ocean acidification on all developmental stages, because organisms may differ in their "'ocean acidification tolerance windows" during different stages in their life-history. Such ocean acidification experiments in the laboratory and field, in addition to long-term ocean monitoring programs, are urgently needed to generate data on which to base policies to mitigate the effects of climate change.

ACKNOWLEDGMENTS

The authors thank John Morrison, Simon Wever, Matthew Wassnig and Michael Milione for technical assistance with this study. This study was funded by AIMS@JCU. The first author was funded by a studentship from the Natural Environment Research Council (NER/S/A/2005/13476) and a Co-operative Award in Science and Engineering studentship from the British Antarctic Survey.

LITERATURE CITED

Auerbach, D. I., J. A. Caulfield, E. E. Adams & H. J. Herzog. 1997. Impacts of ocean C[O.sub.2] disposal on marine life: I. A toxicological assessment integrating constant-concentration laboratory assay data with variable-concentration field exposure. Environ. Model. Assess. 2:333-343.

Bibby, R., P. Cleall-Harding, S. Rundle, S. Widdicombe & J. Spicer. 2007. Ocean acidification disrupts induced defences in the intertidal gastropod Littorina littorea. Biol. Lett. 3:699-701.

Blackford, J. C. & F. J. Gilbert. 2007. pH variability and C[O.sub.2] induced acidification in the North Sea. J. Mar. Syst. 64:229-241.

Brecevic, L. & A. E. Nielsen. 1989. Solubility of amorphous calcium carbonate. J. Cryst. Growth 98:504-510.

Caldeira, K. & M. E. Wickett. 2003. Anthropogenic carbon and ocean pH. Nature 425:365.

Caldeira, K. & M. E. Wickett. 2005. Ocean model predictions of chemistry changes from carbon dioxide emissions to the atmosphere and ocean. J. Geophys. Res. 110:12.

Cao, L., K. Caldeira & A. K. Jain. 2007. Effects of carbon dioxide and climate change on ocean acidification and carbonate mineral saturation. Geophys. Res. Lett. 34:5.

Doney, S. C. & D. S. Schimel. 2007. Carbon and climate system coupling on timescales from the Precambrian to the Anthropocene. Annu. Rev. Environ. Resour. 32:31-66.

Dove, M. C. & W. A. O'Connor. 2007. Salinity and temperature tolerance of Sydney rock oysters Saecostrea glomerata during early ontogeny. J. Shellfish Res. 26:93-9947.

Dove, M. C. & J. Sammut. 2007. Histological and feeding response of Sydney rock oysters, Saccostrea glomerata, to acid sulfate soil outflows. J. Shellfish Res. 26:509-518.

Dupont, S., J. Havenhand, W. Thorndyke, L. Peck & M. Thorndyke. 2008. Near-future level of C[O.sub.2]-driven ocean acidification radically affects larval survival and development in the brittlestar Ophiothrix fragilis. Mar. Ecol. Prog. Ser. 373:285-294.

Gazeau, F., C. Quiblier, J. M. Jansen, J.-P. Gattuso, J. J. Middelburg & C. H. R. Heip. 2007. Impact of elevated C[O.sub.2] on shellfish calcification. Geophys. Res. Lett. 34: L07603. DOI: 10.1029 2006GL028554.

Gosling, E. 2003. Bivalve Molluscs: biology, ecology and culture. Oxford: Blackwell Science. 456 pp.

Hall-Spencer, J. M., R. Rodolfo-Metalpa, S. Martin, E. Ransolne, M. Fine, S. M. Turner, S. J. Rowley, D. Tedesco & M. C. Buia. 2008. Volcanic carbon dioxide vents show ecosystem effects of ocean acidification. Nature 454:96-99.

Harper, E. M. 2000. Are calcitic layers an effective adaptation against shell dissolution in the Bivalvia? J. Zool. 251:179-186.

Haugan, P. M. & H. Drange. 1996. Effects of C[O.sub.2] on the ocean environment. Energy Convers. Manage. 37:1019-1022.

Havenhand, J. N., F. R. Buttler, M. C. Thorndyke & J. E. Williamson. 2008. Near-future levels of ocean acidification reduce fertilization success in a sea urchin. Curr. Biol. 18:R651.

Heasman, M. P., L. Goard, J. Diemar & R. B. Callinan. 2000. Improved Early Survival of Molluscs: Sydney Rock Oyster (Saeeoslrea glomerata). Aquaculture CRC Project A.2.1, NSW Fisheries Final Report Series No. 29, Nelson Bay, Australia.

Hoegh-Guldberg, O., P. J. Mumby, A. J. Hooten. R. S. Steneck, P. Greenfield, E. Gomez, C. D. Harvell. P. F. Sale, A. J. Edwards, K. Caldeira, N. Knowlton, C. M. Eakin, R. Iglesias-Prieto. N.

Muthiga, R. H. Bradbury, A. Dubi & M. E. Hatziolos. 2007. Coral reefs under rapid climate change and ocean acidification. Science 318:1737-1742.

Kikkawa, T., J. Kita & A. Ishimatsu. 2004. Comparison of the lethal effect of C[O.sub.2] and acidification on red sea bream (Pagrus major) during the early developmental stages. Mar. Pollul. Bull. 48:108-110.

Kleypas, J. A., R. A. Feely, C. Fabry, C. Langdon, C. Sabine & L. L. Robbins. 2006. Impacts of ocean acidification on coral reefs and other marine calcifiers: a guide for future research. A report of a workshop held April 18 20, 2005, St. Petersburg, Florida, sponsored by NSF, NOAA, and the US Geological Survey. 88 pp.

Kurihara, H., S. Kato & A. Ishimatsu. 2007. Effects of increased seawater pC[O.sub.2] on early development of the oyster Crassostrea gigas. Aquat. Biol. 1:91-98.

Kurihara. H., S. Shimode & Y. Shirayama. 2004. Effects of raised C[O.sub.2] concentration on the egg production rate and early development of two marine copepods (Acartia steueri and Aeartia erythraea). Mar. Pollut. Bull. 49:721 727.

Kurihara. H. & Y. Shirayama. 2004. Effects of increased atmospheric C[O.sub.2], on sea urchin early development. Mar. Ecol. Prog. Ser. 274:161 169.

Langdon, C., W. S. Broecker, D. E. Hammond, E. Glenn, K. Fitzsimmons, S. G. Nelson, T. H. Peng, I. Hajdas & G. Bonani. 2003. Effect of elevated C[O.sub.2] on the community metabolism of an experimental coral reef. Global Biogeochem. Cycles 17:1011. DOI: 10.1029/2002GB001941, 2003.

Lewis, E. & D. W. R. Wallace. 1998. Program Developed for CO, System Calculations. ORNL/CDIAC-105. Carbon Dioxide Information Analysis Center, Oak Ridge National Laboratory, US Department of Energy, Oak Ridge, Tennessee.

Mayor, D. J., C. Matthews, K. Cook, A. F. Zuur & S. Hay. 2007. C[O.sub.2]-induced acidification affects hatching success in Calanus finmarchieus. Mar. Ecol. Prog. Set. 350:91-97.

Miles, H., S. Widdicombe, J. 1. Spicer & J. Hall-Spencer. 2007. Effects of anthropogenic seawater acidification on acid-base balance in the sea urchin Psammechinus miliaris. Mar. Pollut. Bull. 54:89-96.

Nell, J. A. & W. A. O'Connor. 1991. The evaluation of fresh algae and stored algal concentrates as a food source for Sydney rock oyster Saccostrea commereialis (Iredale & Roghley) larvae. Aquaculture 99:277-284.

O'Sullivan, D., J. Savage & A. Fay. 2008. Status of Australian aquaculture in 2005/2006. Austasia Aquaculture Trade Directory 2008. Turtle Press, Hobart, Tasmania. pp. 6-24.

Orr, J. C., V. J. Fabry, O. Aumont. L. Bopp, S. C. Doney, R. A. Feely, A. Gnanadesikan, N. Gruber, A. Ishida, F. Joos, R. M. Key, K. Lindsay, E. Maier-Reimer, R. Matear, P. Monfray, A. Mouchet, R. G. Najjar, G. K. Plattner, K. B. Rodgers, C. L. Sabine, J. L. Sarmiento, R. Schlitzer, R. D. Slater, I. J. Totterdell, M. F. Weirig, Y. Yamanaka & A. Yool. 2005. Anthropogenic ocean acidification over the twenty-first century and its impact on calcifying organisms. Nature 437:681-686.

Pearson, P. N. & M. R. Pahner. 2000. Atmospheric carbon dioxide concentrations over the past 60 million years. Nature 406:695-699.

Peck, L. S. 2005. Prospects for survival in the Southern Ocean: Vulnerability of benthic species to temperature change. Antarct. Sci. 17:497-507.

Shirayama, Y. & H. Thornton. 2005. Effect of increased atmospheric C[O.sub.2]: on shallow water marine benthos. J. Geophys. Res. 110:C09S09. DOI: 10.1029/2004JC002618.

Southgate, P. C. & A. C. Beer. 1997. Hatchery and early nursery culture of the blacklip pearl oyster Pmctada margartifera (L.). J. Shellfish Res. 16:561-568.

Stenzel, H. B. 1964. Oysters: Composition of the Larval Shell. Science 145:155-156.

Tans, P. NOAA/ESRL (www.esrl.noaa.gov/gmd/ccgg/trends).

Vermeij, G. J. 1993. A Natural History of Shells. Princeton University Press, Princeton, New Jersey. 207 pp.

Weiss, I. M., N. Tuross. L. Addadi & S. Weiner. 2002. Mollusc larval shell formation: Amorphous calcium carbonate is a precursor phase for aragonite. J. Exp. Zool. 293:478-491.

Zeebe, R. E., J. C. Zachos, K. Caldeira & T. Tyrrell. 2008. Oceans Carbon emissions and acidification. Seience 321:51-52.

SUE-ANN WATSON, (1) * PAUL C. SOUTHGATE, (2) PAUL A. TYLER (1) LLOYD S. PECK (3)

(1) National Oceanography Centre, Southampton, School of Ocean and Earth Science, University of Southampton, European Way, Southampton, SO14 3ZH, United Kingdom; (2) AIMS at JCU and School of Marine and Tropical Biology, James Cook University, Townsville, Qld, 4811, Australia; (3) British Antarctic Survey, Natural Environment Research Council, High Cross, Madingley Road, Cambridge, CB3 0ET, United Kingdom

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

TABLE 1.
Water chemistry data for treatment tanks (mean [+ or -] 1 S.D.).

Nominal pH                       8.1                      7.8

Measured [pH.sub.NBS]    8.11 ([+ or -] 0.02)     7.81 ([+ or -] 0.03)
[pH.sub.NBS] range
  (min.--max.)                8.10-8.14                7.78-7.84
Alkalinity (mg/L as
  CaC[O.sub.3])         101.1 ([+ or -] 1.18)    102.27 ([+ or -] 1.42)
Temperature
  ([degrees]C)           25.7 ([+ or -] 0.2)      25.6 ([+ or -] 0.02)
pCO2 ([micro]atm)        220.3 ([+ or -] 16.1)    508.8 ([+ or -] 45.7)
[OMEGA].sub.cal]         1.67 ([+ or -] 0.06)    0.93 ([+ or -] 0.05))
[OMEGA].sub.arag]        1.15 ([+ or -] 0.04)     0.64 ([+ or -] 0.03)

Nominal pH                        7.6

Measured [pH.sub.NBS]    7.64 ([+ or -] 0.04)
[pH.sub.NBS] range
  (min.--max.)                 7.59-7.67
Alkalinity (mg/L as
  CaC[O.sub.3])         101.23 ([+ or -] 0.91)
Temperature
  ([degrees]C)            25.6 ([+ or -] 0.1)
pCO2 ([micro]atm)        775.6 ([+ or -] 5.7)
[OMEGA].sub.cal]         0.65 ([+ or -] 0.06)
[OMEGA].sub.arag]        0.45 ([+ or -] 0.05)