Detailed methodology and results are described in following publication:
Hettinger, A., E. Sanford, T.M. Hill, A.D. Russell, K.N. Sato, J. Hoey, M. Forsch, H.N. Page, and B. Gaylord. 2012. Persistent carry-over effects of planktonic exposure to ocean acidification in the Olympia oyster. Ecology 93: 2758-2768. doi:10.1890/12-0567.1

Briefly (excerpted from above):
Oysters were reared at Bodega Marine Laboratory (BML) in two experiments. These data are from experiment 1. In both, seawater CO2 concentrations in the treatement cultrues were increased relative to present-day levles by 100 and 400 ppm. Thus, seawater CO2 concentrations employed in the experiments were: 700 (used as an operational control), 800, and 1100 ppm. Such CO2 levels correspond to pH values of approximately 8.0, 7.9, and 7.8 (NBS scale). These nominal pH levels were used subsequently to identify the treatments.

Oysters were reared from early larval life in 4.5-L glass culture jars held in seawater tables maintained at 20.0 (+/- 0.02) degrees C. All seawater used during rearing was filtered at 0.45 um and pre-adjusted to appropriate pH levels in 20 L carboys by bubbling for 2-3 days with NIST-traceable CO2 air mixtures (hereafter referred to as "carboy water"). Acrylic boxes mounted over each seawater table received the same mixed gases and provided a common head space for six jars, minimizing off-gassing during culturing. Levels of pH used for each box, and jar position within a box, were randomly assigned.

Larval culturing:
Adult Olympia oysters (4-7 cm in length) were collected from Tomales Bay and transported to BML. They were cleaned and distributed among multiple 100-L cylinders containing seawater filtered at 0.45 um and held at 18-22 degrees C. At least one female per cylinder released larvae within 48 hours post-collection, enabling acquisition of independent "larval cohorts". Following release, larvae were transferred into culture jars containing 2L of seawater filtered at 0.45 um (day 1 of the experiment). Every other day, 90% of the sewater in each jar was changed and replaced with carboy water, whose pH had stabilized to the appropriate level.

Water chemistry:
Samples of jar water and carboy water were collected every day. Seawater pH (NBS) and temperature were measured using a pH/temperature meter (Accumet Excel XL60; Thermo Fisher Scientific, Waltham, Massachusetts, USA), and salinity was determined using a YSI 6600V2 multiparameter instrument (YSI, Yellow Springs, Ohio, USA). Alkalinity was measured using automated Gran titration (Metrohm 809; Metrohm, Herisau, Switzerland), and standardized using certified reference material from A. Dickson at Scripps Institution of Oceanography. Other carbonate system parameters were calculated using the software, CO2SYS (Lewis and Wallace 1998).

Sampling of larvae and juveniles:
Oysters in the culture jars were sampled at key time points during each experiment to quantify shell size and growth rate (change in shell area per day). On day 1, 100 larvae per larval cohort were collected haphazardly by pipette, fixed in 95% ethanol, and individually photographed under a microscope (Leica DM1000 with DC290 camera; Leica Microsystems, Wetzlar, Germany) for analysis using ImageJ software (version 1.37) to determine the initial projected area of the shell (software available online). Larval shell growth rates at later time points were calculated similarly. Juvenile shell growth rate following settlement was determined by measuring the projected shell area of settled individuals from photographs, subtracting the area of the larval shell (which remains visually distinct), and dividing that value by the number of days postsettlement.

Experiment 1:
In the first experiment (July–September 2009), oyster larvae from four larval cohorts were reared in pH 8.0, 7.9, and 7.8 seawater through the duration of planktonic larval development, and for 52 days post-settlement. Each of three pH levels were replicated by two boxes, and all four larval cohorts were represented by one to two culture jars in each box (N = 36 jars total). Fifteen larvae were haphazardly sampled from each jar at day 9 and fixed in 95% ethanol for later photographing. Larvae were allowed to settle directly onto the bottoms of the jars. Larvae reached competency on approximately day 11, and by day 13, the majority of larvae had settled across all treatments. Seven days after settlement, the bottom of each jar was removed and juveniles attached to the jar bottoms were photographed (N = 20 per jar). The juveniles were then transferred to a flow-through seawater table maintained at 20 degrees C, where they received ambient food concentrations from Bodega Bay. After 45 days in this common garden environment (52 days after settlement), juveniles were again photographed to determine their size.

larval_growth_exp1.csv