To evaluate whether rates of sea urchin grazing on C. nereostratum have changed or will change with ocean warming and acidification, we cultured C. nereostratum and S. polyacanthus under experimental conditions mimicking past, present, and predicted future levels of ocean temperature and pCO2 in the region, then followed this three-month culturing period with a controlled feeding experiment conducted under the same conditions. Small C. nereostratum colonies (~4-5 cm diameter) and large S. polyacanthus (~45-60 mm test diameter) were live collected from Adak in 2015 and immediately transported to the Northeastern University Marine Science Center in Nahant, Massachusetts. There, all specimens were acclimated to laboratory conditions at 8.5 °C for two weeks, after which individual C. nereostratum colonies were attached to the underside of plastic petri dishes using cyanoacrylate glue and then allowed to acclimate for an additional two weeks before being moved to experimental aquaria. Conditions were then incrementally modified to achieve target temperature and pCO2 levels (see below) over a one-week period. After reaching target conditions, each 42-L aquarium was dosed with 213 mL of calcein fluorescent dye (Western Chemicals Inc.), which was recirculated in the aquaria for three days and then flushed from the system. Coralline algae incorporate the dye into their skeleton, thus creating a distinct line that can be viewed via fluorescent microscopy to demark the region of new growth within each individual.

We employed four pCO2 conditions and three temperatures that, while factorially crossed, spanned pre-industrial, present-day, and projected year 2100 conditions (assuming an IPCC "business as usual" carbon emissions scenario; Pachauri and Meyer 2014). More extreme temperature (12.5 degrees C) and pCO2 (2800 micro-atm) conditions were also employed in the broader experiment but were not included in our experimental feeding assay because they are not predicted to occur until year 2500, or later. For each treatment, we set temperature to average summertime conditions, the time when ~75% of C. nereostratum growth occurs (Adey et al. 2013).

All treatments (4 pCO2 concentrations x 3 temperatures, fully factorial) were housed on individual shelves and consisted of three 42-liter acrylic aquaria and one 65-liter sump (n = 3 tanks/treatment). The aquaria were connected to a sump via a common overflow and return line but were each independently and continuously replenished with new seawater-thereby establishing them as true experimental replicates. The sump contained a filter box with a nylon mesh particle filter and activated carbon, a protein skimmer (Eshopps PSK-75), and a return pump, all of which was connected to a water chiller (Coralife 1/4HP). Filtered natural seawater was added via Darhor manual flow controllers at a rate of 50 mL/min/tank, resulting in full replacement of treatment water every ~21 hours-sufficiently fast to prevent material depletion of the dissolved constituents of the seawater yet slow enough to allow the mixed gases being sparged into the experimental treatments to approach equilibrium with the seawater. Mixed gases were sparged into each tank with 91 cm long flexible bubblers at the rate of ~1 L/min via Darhor needle-valve gas flow controllers. Two 12,000K LED light arrays (Ecoxotic Panorama, Pro 24V) were mounted above each tank and set to an irradiance that mirrored average summer daylight irradiance at 10 m depth in the Aleutian Islands (~258 micro-E m-2 s-1; 12 hr light:12 hr dark cycle).

Over the course of the four-month experiment, we measured pH (Accumet AB15 pH meter with Accufet solid state probe), salinity (YSI3200 meter with K=10 conductivity electrode and temperature probe), and temperature (NIST traceable red spirit glass thermometer) in each tank every Monday, Wednesday, and Friday. The pCO2 of the gas mixtures was measured with a Qubit S151 infrared CO2 analyzer and calibrated with certified mixed CO2 from Airgas Incorporated. Every 10 days, we characterized the full carbonate system chemistry of the experimental treatments from measured total alkalinity, dissolved inorganic carbon, temperature, and salinity. For this, seawater samples were obtained in 250 mL borosilicate ground-glass-stoppered bottles and immediately poisoned with 100 micro-L of saturated HgCl2 solution to halt biological activity (Dickson et al. 2007). Total alkalinity was measured via closed-cell potentiometric Gran titration and dissolved inorganic carbon was measured with a UIC 5400 Coulometer on a VINDTA 3C (Marianda Incorporated) using Dickson certified seawater reference material. Seawater pCO2, pH, carbonate ion concentration ([CO32-]) bicarbonate ion concentration ([HCO3-]), aqueous CO2, and calcite saturation state were calculated with the program CO2SYS (Lewis and Wallace 1998), using Roy and colleague's (1993) values for the K1 and K2 carbonic acid constants, the Mucci (1983) value for the stoichiometric calcite solubility product, the seawater pH scale, and an atmospheric pressure of 1.015 atm.

At the beginning of the experiment we measured the buoyant weight of each specimen. We then scrubbed each specimen with a toothbrush and reweighed it every month and at the end of the experiment. With each weighing, we also photographed the specimen with a ruler and Reef Watch coral bleaching card in the field of view. We then measured the 2-d surface area (coralline algae) or test diameter (urchin) of the photographed specimens (Image J, NIH). Following a three-month incubation, a subset of the coralline algae was placed individually in cages and paired with a single urchin to quantify bioerosion rate under the different treatments, while the remaining algae were retained as experimental controls during the 20-day feeding experiment, and subsequently measured for skeletal density. Following the feeding experiment, all coralline algae were sectioned with a diamond lapidary saw (Inland Craft SwapTop 6.5" Diamond Trim Saw) and either frozen for genetic analysis or sectioned into 6 mm slices, rinsed in a series of two 90% Ethanol baths, and allowed to air dry for further examination of growth and skeletal density.

After three months of culturing C. nereostratum and large S. polyacanthus (mean test diameter ± SE = 53 ± 1 mm) under various temperature and pCO2 conditions (see above), we conducted a controlled feeding assay. Individual coralline algae were randomly paired with sea urchins from the same tank and placed into small cages (n = 5/tank, 15/treatment) and returned to aquaria, with the remaining coralline algae from each tank serving as controls. Buoyant weight measurements and photographs of each alga and urchin were obtained at the beginning of the experiment (as described above) and again every five days. We excluded replicates where: (i) the sea urchin appeared moribund or died during the assay; or (ii) negligible grazing occurred throughout the experiment, indicating severe stress to the animal. Buoyant weights were converted to dry weight (mg) using an empirically-derived conversion factor (linear regression: -0.117448 + 1.746361 * buoyant mass; adjusted R^2 = 0.9996; p < 0.001). To calculate the amount of C. nereostratum (mg CaCO3 dry weight) consumed by each urchin at each sampling interval while also accounting for the potential gain or loss of mass within the treatment alga, we used the equation [Ti x (Cf/Ci)] - Tf, where Ti and Tf is the initial and final mass (respectively) of an alga exposed to herbivory, and Ci and Cf  is the initial and final mass (respectively) of its paired control. For this exercise, we averaged correction factors (Cf/Ci) for all control algae in a given tank, and applied this average correction factor to each treatment alga in the same tank. We then computed grazing rate (amount consumed/day) for each sea urchin, as well as per capita grazing rates standardized by the surface area of C. nereostratum.