Detailed methodology and results are described in the following publication:
Hettinger, A., E. Sanford, T.M. Hill, J.D. Hosfelt, A.D. Russell, and B. Gaylord. 2013. The influence of food supply on the response of Olympia oyster larvae to ocean acidification. Biogeosciences 10: 6629-6638. doi:10.5194/bg-10-6629-2013

Briefly (excerpted from above):
Adult Olympia oysters (n = 140) were collected from Tomales Bay, California (38 deg 06'58" N, 122 deg 51'16" W) in June 2011, transported to Bodega Marine Laboratory (BML) and distributed evenly among four 100 L culturing cylinders. A similar average size of adult oysters was maintained among cylinders. Every day, adults in each cylinder were fed microalgae (Isochrysis galbana) at a high concentration to encourage larval release. Seawater, filtered to 0.45 um and held at 18– 22 degrees C, was changed every other day. Adults released larvae in two of the four culturing cylinders after 72 h. Larvae were distributed by pipette to 4.5 L glass jars on day 1 of the experiment (n = 1000 larvae per jar). Each jar held 2 L of filtered seawater at the appropriate pCO2 level, resulting in initial rearing concentrations of 1 larva per 2 mL of seawater.

In subsequent experimental trials, the investigators employed a target elevated seawater pCO2 concentration of 1000 uatm (~7.76 pH). The target control seawater pCO2 concentration was 500 uatm (~8.05 pH). All seawater used during larval rearing was pre-adjusted, prior to addition of larvae, to the appropriate pCO2 concentrations in 20 L carboys by bubbling filtered seawater for 2–3 days with NIST-traceable CO2 air mixtures (carboy water). To minimize net CO2 exchange across the seawater surfaces in the jars and help maintain seawater pCO2 concentrations at target levels within the jars, the same CO2 gas mixtures used to maintain carboy water were pumped continuously into sealed air spaces above the free surfaces of the seawater in the culture jars (hereafter "headspaces"). Five replicate jars per pCO2 concentration shared a given headspace. Carboys and jars were held in seawater tables maintained at 20 degrees C (+/- 0.2 degrees C).

Every other day, 90% of the seawater in each jar (jar water) was removed using reverse-filtration through 125 um mesh and replaced with carboy water at the target pCO2 concentration. Immediately following each water change, microalgae (I. galbana) were added to each jar to generate final densities in each jar of: 100,000, 50,000, or 10,000 cells per mL (n = 2 pCO2 levels × 3 food levels × 5 replicate jars = 30 jars). The three food levels were the equivalent of initial daily algal cell to larva ratios of 50, 25, and 5 cells per larva per day.

Seawater chemistry:
Jar water and carboy water were sampled for total alkalinity (TA) and dissolved inorganic carbon (DIC) every other day when a water change was performed. Seawater pH and temperature were measured using a potentiometric pH/temperature meter (Accumet Excel XL60). Raw pH readings (mV) were calibrated using two seawater buffers. pH was monitored primarily as a real-time indicator of changes in the carbonate system. Salinity was determined using a YSI Professional Plus Multiparameter instrument with a conductivity probe (YSI, Yellow Springs, OH). TA was measured using automated Gran titration with duplicates (Metrohm 809), and standardized using certified reference material (CRM) from A. Dickson. The offset between measured and certified TA values for Dickson CRM (Batches 104, 107, 111) was −0.01 (+/- 2.81) meq per kg (n = 91). Other carbonate system parameters were calculated using the carbonate system analysis software CO2SYS (Lewis andWallace, 1998).

Sampling of larval growth:
On day 1, prior to their placement in the culture jars, larvae were collected haphazardly by pipette (n = 20), placed on a 125 um Nitex plankton filter, rinsed, and left to dry at room temperature for 24 h. Larvae were photographed individually under a dissecting microscope (Leica M125 with DC290 camera) for analysis using ImageJ software (ver. 1.37, National Institutes of Health) to determine the initial projected area of the shell. Larvae were sampled similarly on day 5, 9, and 11 post-larval release (n = 10 larvae per jar at each time point). Larval growth was estimated as the change in projected shell area since larval release (day 1).

Sampling of larval total dry weight:
Total dry weight (body plus shell, ug) of the oysters was also determined on days 5, 9, and 11 post larval release. Larvae were collected haphazardly by pipette from each jar (n = 25), placed on a 125 um Nitex plankton filter, and rinsed. Total dry weights of larvae sampled were determined by transferring larvae individually to aluminum vessels pre-ashed at 500 degrees C for 3 h, drying the larval sample at 50 degrees C for >24 h, and weighing on a microbalance (Sartorius Ultramicro, Goettingen, Germany). Larvae were dried in their vessels a second time, reweighed to verify their weights, and combusted at 460 degres C for 4 h in a muffle furnace (Thermo Scientific FB1415M) to remove all organic matter (see also Gaylord et al., 2011). Ash-free dry tissue weights were determined from the weight difference before and after combusting.

Sampling of percent metamorphosis:
Settlement of larvae and metamorphosis into benthic juveniles was assessed daily starting on day 11, two days after larval transfers were made into the substrate jars. When <5% of larvae remained swimming (used as the assay point for settlement) in each pCO2 treatment, the bases of each substrate jar were removed, and the proportion of metamorphosed individuals was determined by first subtracting dead and non-metamorphosed larvae (e.g., pediveligers) from the total initial number of larvae, then dividing by the total initial number of larvae (1000). Any metamorphosed individuals on the walls of the glass jars were also included in the proportion of metamorphosed individuals.

food_larval_mass.csv