This methodology describes this dataset and other datasets from this experiment. See "Related Datasets" section for data access and more details of each related dataset.

Physiological analyses

Coral survivorship was assessed visually daily, and only one coral fragment died during the experiment. Net calcification, surface area (a proxy for extension; Rathbone (2021)), volume, and dark-adapted photosynthetic efficiency (Fv/Fm) of coral fragments were measured six times during the experiment (~2 week intervals) via buoyant weight and photogrammetry using previously described methods (Davies, 1989; Brown et al. 2021; Ferrari et al., 2016) (Brown et al. 2022 Supp Methods, Figure S4, Figure S5). At the end of the experiment, metabolic rates (net photosynthesis, dark respiration and light-enhanced dark respiration) were assessed via changes in oxygen evolution using oxygen optodes connected to an OXY-10 (PreSens) optical analyzer (Brown et al. (2019) Supp Methods). Upon completion of these living analyses, half of the coral fragments were flash frozen in liquid nitrogen and stored at -80°C. Subsequent laboratory analyses were done on these 48 specimens. For these analyses, corals (n = 12) were water-piked on ice to remove coral tissue from the skeleton using 50 mL of 0.1 M phosphate buffered saline solution. The tissue slurry was centrifuged at 4°C once for 5 min at 2500 g to sufficiently separate host tissue and the intracellular endosymbiont cells. Host tissue was analyzed for host-soluble protein concentration and mycosporine-like amino acids (MAAs) concentrations spectrophotometrically (Whitaker and Granum, 1980). Endosymbiont densities were determined from cell counts of three aliquots using a hemocytometer (Brown et al., 2019). Host protein concentration and endosymbiont cell densities were standardized to surface area (cm2), which was determined using the single wax-dipping technique (Holmes, 2008), whereas MAAs were normalized to host protein content. Endosymbiont photopigments were extracted in 100% acetone for 24 hours and concentration of chlorophyll a was determined via absorbance at 630, 663, and 750 nm using the equations in (Jeffrey and Humphrey, 1975). Pigment concentrations were standardized to both surface area and endosymbiont densities. Wax-dipping was also used to determine calcium carbonate (CaCO3) bulk density, where the skeleton was sealed with a coat of wax, dry weighed, and then buoyant weighed (Tambutté et al., 2015). The difference between dry weight and buoyant weight was calculated to determine the bulk volume, which was subtracted from the dry weight to yield bulk density. The other half of the fragments were transported alive from Heron Island to the University of Queensland, Brisbane to assess intracellular acid-base status and acidification resilience following established methods (Innis et al., 2021). Briefly, cells were loaded with SNARF-1AM and imaged using a confocal microscope (Zeiss LSM 710) via excitation at 561 nm, with SNARF-1 fluorescence emission acquired in two channels (585 and 640 ± 10 nm) simultaneously (see full details in Supp Methods of Brown et al., 2022).

Physiological analyses

Net calcification of coral fragments was measured six times across the experiment (22 Jan, 30 Jan, 12 Feb, 26 Feb, 12 Mar and 19 Mar of 2021) using the buoyant weight technique (Rathbone et al., 2021; Camp et al., 2018). At the same time, surface area and volume were quantified via non-invasive three-dimensional photogrammetry, with fragment reconstructions created from a set of ≥50 photographs using the program Autodesk ReCap Photo (Camp et al., 2018; Brown et al., 2022 Figure S4). On the night prior to growth measurements, dark-adapted photochemical efficiency (Fv/Fm) was quantified using a Diving-PAM (Walz GmbH) approximately 1 hour after sunset. Measurements were made using the Diving-PAM 5-mm diameter fibre-optic probe at a standardized distance 5 mm above the coral tissue after F0 stabilized (n=3 per fragment). At the end of the experiment, metabolic rates were assessed via changes in oxygen evolution using oxygen optodes connected to an optical analyzer (OXY-10, PreSens) (Kenkel and Matz, 2016). Coral fragments were analyzed at the end of the experimental period between 08:00 and 18:00 within 140 cm3 clear acrylic chambers on top of a magnetic stirrer to allow for continuous mixing. Corals were dark-adapted for at least 30 min prior to each assay, which followed a light program of 20 min of darkness (0 µmol quanta m-2 s-1) to measure dark respiration, 25 min of midday light levels (~500 µmol quanta m-2 s-1) to determine maximum net photosynthesis and 15 min of darkness (0 µmol quanta m-2 s-1) to determine light enhanced dark respiration. Seawater conditions were replicated to those experienced in the tanks by: (1) using seawater collected from treatment tanks to provide initial pCO2 concentrations and (2) using a water bath to maintain respective treatment temperatures within the incubation chambers. Tank water was filtered using a 0.22µm filter and oxygen content of the seawater was lowered to 70% using N2 to avoid hyperoxia. Chambers were completely drained and cleaned with a soft sponge in between trials.

Assessment of coral intracellular acid–base homeostasis

The other half of the fragments were transported alive from Heron Island Research Station (HIRS) to the University of Queensland, Brisbane to assess intracellular acid-base status and acidification resilience following established methods. Corals were held in an indoor, closed system (186 L per treatment; n = 2 tanks per treatment) that replicated stable and variable pCO2 conditions similar to the controllers at HIRS via the addition of CO2 or CO2-free air using Apex controllers and probes (Neptune Systems). PAR followed a 12 hr:12 hr day:night cycle, with mean PAR ~125 µmol quanta m−2 s−1. Corals were held for a maximum of two weeks, and were selected for analysis randomly. Coral cells were isolated by submerging the fragment in a shallow dish of 50mL filtered seawater (FSW) and gently brushing with a soft toothbrush. Cells were filtered through at 100 µM cell strainer into a clean 50mL tube and centrifuged for 4 min at 350 x g to pellet the cells. The supernatant was decanted, and the pellet was resuspended in 2 mL FSW. Cells were loaded with SNARF- 1 AM by adding 1 µL SNARF-1 AM (20 mM) and 1 µL pluronic acid (20% w/v) to 2 mL cells for 30 min in darkness. Isolated cells were spun down using a benchtop microfuge for 5 sec, discarding the supernatant and resuspending in ambient filtered seawater (pHe 8.0) or in acidified seawater (pHe 7.4) in darkness to measure cellular response to acidosis. Cells were imaged using a confocal microscope (Zeiss LSM 710) via excitation at 561 nm, with SNARF- 1 AM fluorescence emission acquired in two channels (585 and 640 ± 10 nm) simultaneously. A total of 8–10 cells containing algal symbionts (symbiocytes) and 8–10 cells without symbionts (non-symbiocytes) were imaged for each coral fragment. To measure acid stress response, SNARF-1 AM loaded cells were imaged over time (0, 15, 30, 45, 60, 75 minutes after acid stress) in darkness. SNARF-1 AM fluorescence emission was quantified in ImageJ by drawing two regions of interest per cell within the coral cytoplasm. A region drawn in the surrounding medium was used to subtract background fluorescence, and for each region of interest, a fluorescence ratio was calculated and converted to pH using a calibration curve generated as previously described (Oliver and Palumbi, 2011). The rate of pHi recovery was calculated as the slope of pHi from 15 to 60 minutes post-acidification.

For more detailed information, please see: Brown et al. (2022).

ph_i.csv