Dataset: Oyster Carryover Effects - Juvenile Growth: Expt 2
Deployment: BML_Gaylord

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 2:
Reduced pH induced a negative effect on juvenile growth rate. This outcome (of experiment 1) could have arisen from one of two causes: direct effects of seawater acidification during the early juvenile phase or carry-over effects of larval experience. Experiment 2 (September–October 2009) was conducted to distinguish among these alternatives.

In the first phase of Experiment 2, larvae were pooled across cohorts (300 larvae/cohort; 900 larvae/replicate jar) and reared through settlement under either control or low pH conditions(pH 8.0 or 7.8). The pH levels were randomly assigned to four boxes (2 pH levels x 2 boxes x 3 replicate jars = 12 jars). In the second phase of the experiment, and within 24 h of larval settlement, one-half of the juveniles reared as larvae in control pH conditions were transferred to low pH conditions, and one-half were returned to control pH conditions. Similarly, one-half of the juveniles that had been reared as larvae in low pH conditions were transferred to control pH conditions, and one-half were returned to low pH conditions.

The second phase of the experiment used 2 pH levels x 4 larval-juvenile pH treatments x 6 replicate jars = 24 jars. The juveniles in the two destination pH levels were reared until 13 days after settlement, and randomly selected juveniles on wedges (N=17 per jar) were photographed 7 and 13 days after settlement to quantify shell growth rate since settlement. Larval settlement occurred 24 hours earlier (day 13) under low pH conditions; therefore all photographs taken of settled juveniles in control conditions were delayed by one day.