Publication associated with this data set:
McLachlan RH, Price JT, Muñoz-Garcia A, Weisleder NL, Levas SJ, Jury CP, Toonen RJ, Grottoli AG. Physiological acclimatization in Hawaiian corals following a 22-month shift in baseline seawater temperature and pH. Scientific Reports (in press) – updated February 2022.

Locations
Corals were collected at the following locations:
Sampan, O'ahu, Hawai'i, USA 21.452394 N, -157.794870 W, depth 0.5–5m.
Hawai'i Institute of Marine Biology, O'ahu, Hawai'i, USA, 21.434167 N, -157.786335 W, depth 0.5–5m.
Waimānalo, O'ahu, Hawai'i, USA, 21.326287 N, -157.674599 W, depth 0.5–5m.
Hale'iwa, O'ahu, Hawai'i, USA, 21.592516 N, -158.110337 W, depth 0.5–5m.

Experiments were conducted at the Hawai'i Institute of Marine Biology, O'ahu, Hawai'i.

Coral species, sample collection, and acclimation
The corals Montipora capitata (branching and encrusting), Porites compressa (branching), and Porites lobata (massive) were collected at 2 ± 1m depth between 29 August and 11 November 2015 from four reef sites around the island of Oʻahu, Hawaiʻi: Moku o Loʻe [21.434167 N, -157.786335 W] and Sampan [21.452394 N, -157.794870 W] within Kāneʻohe Bay, Waimānalo [21.326287 N, -157.674599 W], and Haleʻiwa [21.592516 N, -158.110337 W]. P. lobata was not found at Moku o Loʻe and was not collected there. This broad spatial sampling of corals helped to ensure that a representative sample of the genetic variation present in these species from Oʻahu was included in the study. Six genets of each species were collected at each site using a hammer and chisel for a total of 66 genets (24 parent colonies for M. capitata, 24 parent colonies for P. compressa, and 18 parent colonies for P. lobata). Species-specific microsatellite markers (developed by Concepcion et al. 2010; Gorospe and Karl 2013) were used to genotype all corals and ensure that they were genetically distinct. After removal from the reef, genets were placed in individual plastic bags filled with seawater from the collection site, and transported back to the Hawaiʻi Institute of Marine Biology [21.434167 N, -157.786335 W]. Four clonal ramets were cut from each genet using a band saw, and each ramet was mounted on a labelled ceramic plug using cyanoacrylate gel. The 264 ramets (i.e., 66 genets x 4 ramets) were distributed among the experimental outdoor flow-through mesocosm tanks, and allowed to recover and acclimate to the mesocosm system under ambient reef-supplied flow-through seawater for at least 12 weeks until 31 January 2016. Shade cloth above the mesocosm tanks attenuated sunlight by 30% to provide irradiance like that at collection depth, with a maximum instantaneous irradiance of ~1730 µmol m-2 s-1-1 (Wall et al. 2019).

Mesocosm experiment
The experiment consisted of four treatments (n = 10 mesocosms per treatment) as follows: control (present-day temperature with present-day pCO2), ocean acidification (present-day temperature with +350 μatm pCO2), ocean warming (+2 ℃ with present-day pCO2), and combined future ocean conditions (+2 ℃ with +350 μatm pCO2). The temperature and pCO₂ levels are consistent with current commitments under the Paris Climate Agreement (Rogelj et al. 2016). The ramets of M. capitata, P. compressa, and P. lobata were distributed among the 40 outdoor flow-through mesocosm tanks (70 L, 50 x 50 x 30 cm) at HIMB such that one ramet per genet was present within each of the four treatment conditions. Starting on 1 February 2016, temperature and pCO₂ were adjusted gradually over 20 days to minimize the likelihood of shocking the mesocosm communities. As the incoming waters from Kāneʻohe Bay are naturally slightly warmer and more acidic than other nearby reefs (Jury et al. 2013; Jury and Toonen 2019; Price 2020; McLachlan et al. 2021), the seawater being delivered to the control and ocean acidification treatment mesocosm tanks was chilled by 0.5 °C after 10 and 20 days while the seawater delivered to the ocean warming and combined future ocean treatment mesocosms was warmed by 0.5 °C after 10 and 20 days, then maintained at these offsets for the remainder of the study. At the same time, pH of the seawater being delivered to the ocean acidification and combined future ocean treatments was decreased by 0.05 units while the pH of the seawater being delivered to the control and ocean warming treatment mesocosm tanks was increased by 0.05 units after 10 days and again after 20 days then maintained at the offsets for the remainder of the study. Corals were maintained under experimental conditions for 22 months from 20 February 2016 to 13 December 2017 for a total of 662 days. Salinity, temperature, pCO₂, and pH were measured at mid-day in each mesocosm once weekly. This is a long-term experiment as defined by McLachlan et al. (2020e) and Grottoli et al. (2021) and the longest dual stress (i.e., combined ocean warming and acidification) experiment on corals to date (McLachlan et al., in press).

The mesocosms were designed to mimic the natural reef environment as closely as possible. Each mesocosm contained fragments from the eight most dominant reef-building coral species in Hawaiʻi (Montipora capitata, Montipora flabellata, Montipora patula, Porites compressa, Porites lobata, Porites evermanni, Pocillopora meandrina, and Pocillopora acuta) (Franklin et al. 2013; Rodgers et al. 2015), a layer of sand and carbonate rubble, a juvenile Convict tang (Acanthurus triostegus), and a juvenile Threadfin butterflyfish (Chaetodon auriga). Both fishes are generalist grazers, where the convict tang feeds on benthic algae and the butterflyfish feeds on non-coral invertebrates. They were at representative fish biomass densities for Hawaiian reefs (Gorospe et al. 2018), and together provide the essential functional role of herbivory and predation within the mesocosm communities. The flow-through mesocosms received unfiltered seawater pumped directly from the neighboring reef within Kāneʻohe Bay. Seawater was initially pumped into one of eight header tanks (two per treatment) within which temperature and pH were manipulated and then subsequently directed into the mesocosms such that each header tank supplied five mesocosms. Coral fragments were not directly fed but had access to dissolved and particulate organic matter from the reef-derived seawater and from daily feeding of the fish who were supplied ~3 g wet weight of frozen adult mysid or Artemia brine shrimp under flow-through conditions, thereby provisioning the fish and mesocosm communities with allochthonous (i.e., non-local, imported) non-living zooplankton at a rate similar to that measured in nature (Hamner et al. 1988).

The coral fragments grew faster than expected during the experimental period and therefore space within each mesocosm became limited near the end of the experiment. In order to prolong the experiment, three of the eight coral species (Montipora flabellata, Pocillopora meandrina, and Porites evermanni) were transferred into a secondary mesocosm system at HIMB in the last month of this study on 18 November 2017, thereby increasing the available space to all remaining coral fragments. None of the M. capitata, P. compressa and P. lobata corals showed any obvious adverse reactions to having M. flabellata, P. meandrina, and P. evermanni removed from the primary mesocosm tanks.

Coral fragments were photographed for surface area and ramet whiteness analysis, and buoyant weighed on the weeks of 20 March 2016 and 27 November 2017 corresponding to one month after the target temperature and pH conditions were reached and the end of the experimental period, respectively. During the last 20 days of the experimental period (23 November–13 December 2017) the following live physiological measurements were conducted on all surviving coral ramets: photosynthesis, respiration, total organic carbon flux, and maximum Artemia feeding capacity. Then, all surviving coral fragments were sacrificed by freezing at -20 ℃. Samples were transported on dry ice to the Ohio State University (OH, USA) where they were stored at -80 ℃ awaiting further analyses of biomass, lipids, proteins, Symbiodiniaceae density, and surface area according to methods published in protocols.io (McLachlan et al. 2020d, 2020c, 2020a, 2020b; McLachlan and Grottoli 2021).

Laboratory analyses

Coral color and surface area
Photographs of corals were taken from six different angles next to a scale bar and white reference card. Using ImageJ software, coral whiteness was assessed via photographic image analysis using the greyscale model to quantify the bleaching appearance (Amid et al. 2018). The percent whiteness was used as a proxy for bleaching intensity of corals because it is known to be highly correlated with chlorophyll a and Symbiodiniaceae density (Chow et al. 2016; Amid et al. 2018). Coral surface area was estimated from photographs using the geometric method (Naumann et al. 2009) for which a detailed protocol is described in McLachlan and Grottoli (2021).

Calcification
Calcification rate was determined using the buoyant weight technique (Jokiel et al. 1978). Daily calcification rates were calculated as the difference between initial and final weights, divided by the respective number of days elapsed, and normalized to the initial weight of the skeleton.

Photosynthesis, respiration, and total organic carbon flux
Maximal photosynthesis and day and night respiration rates were measured via changes in dissolved oxygen for each individual coral ramet (Rodrigues and Grottoli 2007) at respective treatment seawater temperatures and pCO₂ levels, and normalized to ash-free dry weight (AFDW). Total respiration was calculated by summing day and night respiration rates multiplied by the respective number of hours per day (i.e., 11 daytime hours and 13 nighttime hours at the time of our measurements in Hawaiʻi). Photosynthesis and respiration rates were corrected for any change in seawater oxygen concentration due to microorganism respiration which occurred in a seawater blank control chamber. Due to the high growth rates exhibited by many of the corals during the experiment, they were far too large to fit into the original respirometry chambers by the end of the study. Therefore, a smaller sub-ramet was cut from each ramet of M. capitata and P. compressa using a band saw with a diamond-coated blade and each sub-ramet was mounted on a labelled ceramic plug using cyanoacrylate gel for respirometry. The P. lobata grew primarily horizontally rather than vertically and thus were not cut prior to live physiological analyses and instead new wider respirometry chambers were constructed to accommodate them.

Total organic carbon (TOC) water samples were collected following night respiration incubations because coral feeding is known to occur primarily after dusk, using methods adapted from Levas et al. (2015). Following night respiration incubations, the water level in respirometry bins was lowered to expose the top of coral incubation chambers. While wearing nitrile gloves, lids were removed, and corals returned to experimental tanks. A 30 ml water sample was removed from each chamber using a new disposable 10 ml pipet, filtered through a 55 µm Nitex mesh, and collected in a pre-cleaned 50 ml Nalgene bottle. All water samples were acidified within 10 minutes of collection using 1 ml of 1.2 M hydrochloric acid (ACS Reagent Grade). TOC concentrations were determined using high-temperature catalytic oxidation using a Shimadzu model TOC-L analyzer, were corrected for the volume of water in respirometry chambers, and divided by the incubation duration to obtain the flux values per hour. The TOC value of the non-coral containing control chambers were subtracted from coral chamber TOC values. Corrected TOC fluxes were normalized to AFDW.

Carbon budget
The carbon budget of each coral ramet was calculated to determine the proportionate contribution of photosynthesis and heterotrophy to total metabolic demand (i.e., respiration). Photosynthesis and total respiration rates were used to calculate the percent Contribution of Zooxanthellae (i.e., Symbiodiniaceae) to Animal Respiration (CZAR) (Muscatine et al. 1981), while total respiration and nighttime TOC flux rates were used to calculate the percent Contribution of Heterotrophy from TOC to Animal Respiration (CHARTOC) (Levas et al. 2015). Artemia feeding capacity was not used to calculate CHARzoop as the Artemia concentrations were not representative of reef zooplankton densities or mesocosm zooplankton densities. The Contribution of the Total acquired fixed carbon relative to Animal Respiration (CTAR) (Grottoli et al. 2014) was calculated as the sum of CZAR and CHARTOC. However, we acknowledge that this is likely an underestimate of CTAR as it does not account for heterotrophic carbon derived from zooplankton nor any potential gains or losses in CHARTOC that may have occurred during the day.

Maximum Artemia capture rate
The maximum Artemia capture rate of corals was assessed using methods adapted from (Ferrier-Pagès et al. 2010). Briefly, corals were placed upon a small plastic stand in individual 500 ml glass beakers filled with seawater from their respective treatments. Beakers were placed on top of a magnetic stir plate (200 RPM) within a 100 L water bath maintained at the desired experimental temperature and placed in front of a window. Conducting feeding measurements under natural moonlight has been observed to increase polyp expansion and feeding behavior (Grottoli pers. obs.). Corals were placed in the feeding beakers one hour before sunset to ensure polyp expansion. Approximately 30 min after sundown, a concentrated solution of 2-day old Artemia salina nauplii was added to each beaker at an average concentration of 3000–3500 Artemia L-1. This concentration is much higher than in situ zooplankton concentrations, but was chosen to assess the maximum zooplankton capture rate of corals, as feeding rate in known to increase with prey concentration (Palardy et al. 2005, 2006). Five 10 ml subsamples were removed from each beaker using a 10 ml glass pipette after 2 and 40 minutes and the number of Artemia in the pipette was immediately counted under a light microscope. The counted Artemia solution was returned to the beaker within 30 seconds of its initial removal. After the final count, corals were removed from beakers and returned to their experimental tanks. The maximum Artemia feeding rate was calculated as the difference between average initial and end concentrations of Artemia, divided by the volume of water in the feeding beaker and the duration of the feeding trial. Capture rates were corrected for any change in Artemia concentration which occurred in a control beaker without coral. Maximum Artemia capture rates were normalized to AFDW.

Biomass, lipid, protein, and Symbiodiniaceae density
Frozen coral fragments were ground into a homogenous paste using a chilled mortar and pestle and partitioned using methods described in McLachlan et al. (2020a). Between 0.5–1 g of ground material was partitioned for analyses of total biomass ash-free dry weight, total soluble lipid, total soluble protein (henceforth referred to as biomass, lipid, and protein, respectively), and Symbiodiniaceae density based on pre-determined needs for each analysis. Briefly, biomass was quantified by drying ground coral subsamples to a constant weight (60 ℃ for 24 hr) and burning it (450 ℃ for 6 hr) according to protocol methods detailed in McLachlan et al. (2020a). Lipids were extracted using 2:1 chloroform methanol using methods modified from Rodrigues and Grottoli (2007) and the protocol detailed in McLachlan et al. (2020c). Protein was quantified using the Bradford method (Bradford 1976) with protocol details in McLachlan et al. (2020d). Symbiodiniaceae density was quantified by counting the number of cells in four replicate (4 µL) subsamples using a Countess™ II FL Automated Cell Counter which is detailed in McLachlan et al. (2020b). Coral biomass was normalized to surface area (McLachlan and Grottoli 2021), and lipid, protein, and Symbiodiniaceae density were normalized to AFDW to facilitate comparison among species of varying morphologies with different surface-area-to-volume ratios (Edmunds and Gates 2002).