Dataset: Buoyant weight data collected during acute and chronic stress exposures in outdoor mesocosms in Hawaii from January 2022 to March 2024

Overview

Seasonal mesocosm incubations of Montipora capitata and Pocillopora acuta were conducted over a two-year period (June 2022–March 2024) at the Hawai‘i Institute of Marine Biology (HIMB). Coral were collected from fringe reefs located around the institute. Experiments were carried out during winter (January 10–March 17) and summer (June 10–August 17) in outdoor flow-through mesocosms at the Hawaii Institute of Marine Biology in Kane`ohe, Hawai`i. Due to facility constraints, all treatments could not run simultaneously. Therefore, each seasonal experiment consisted of two consecutive 30-day exposures, each using newly collected coral colonies. All procedures were standardized across replicates, except for adjustments in total alkalinity (TA) and pH required to create the treatments.

 

Coral Collection and Pre-Experiment Holding

Coral colonies of Montipora capitata and Pocillopora acuta were collected by hand while snorkeling from shallow (1 m) fringing reef habitat surrounding HIMB in Kāne‘ohe Bay, Hawai‘i. Colonies were transported immediately to the mesocosm facility and held for one week in flow-through tanks that were identical in design to the experimental mesocosms.

One day prior to placement in experimental mesocosms, all colonies were stained for eight hours with alizarin red to mark the initial skeletal growth band (reference: Jokiel and Morrissey 1993).

At the start of each 30-day experiment, colonies were randomly assigned to mesocosms, buoyant weighed following Jokiel (1978), and placed into a standardized grid layout. Each mesocosm contained 20 colonies of each species arranged in alternating species order to avoid species-level clustering effects.

 

Mesocosm Facility Design

The flow-through mesocosm facility (Jokiel et al. 2014) consisted of twelve 450 L fiberglass tanks operating on continuous seawater flow. Tanks were arranged in four rows of three mesocosms, with each row supplied by a dedicated header tank. Water was pumped at ~3 m depth from the adjacent bay, supplying unfiltered seawater that preserved natural diel and seasonal variability.

Each header tank fed seawater into a 100 L mixing reservoir positioned above the system to facilitate chemical manipulation before seawater entered the mesocosms. Mesocosm residence time was greater than one hour. Each mesocosm contained two submersible circulation pumps and an airstone that ran continuously to maintain water movement and oxygenation.

 

Treatment Conditions and Carbonate Chemistry Manipulation

Four carbonate chemistry conditions were generated across consecutive 30-day exposures by altering total alkalinity (TA), dissolved inorganic carbon (DIC), and pH. Target conditions included:

1. Ambient control

2. Low pH (ΔpH ~ -0.3 from ambient)

3. High or low TA (±100 µmol kg-1 from ambient)

4. Combined low pH and TA alteration

TA adjustments were made using a peristaltic pump delivering either 1.0 M HCl (for low TA) at ~3 mL min-1 or 1.0 M Na2CO3 (for high TA) at ~2 mL min-1 directly into each row’s mixing tank.
pH reductions were achieved by bubbling pure CO2 or a CO2-air mixture directly into designated mesocosms using a Maxi-Jet 1600 pump-driven venturi injector.

Because each row was supplied by a single header tank, only one direction of TA manipulation (increase or decrease) could be applied per row in a given 30-day experiment. The subsequent 30-day experiment performed the opposite TA manipulation, ensuring all conditions were tested each season.

 

Seawater Sampling and Measurements

Daily Parameters

Temperature, salinity, dissolved oxygen, and pHNBS were measured daily at mid-day in all mesocosms and header tanks using a YSI multimeter (YSI ProDSS or YSI 556 MPS).

pH electrodes were calibrated daily using NIST-traceable pH 4, 7, and 10 buffers and corrected to the total scale using Tris buffer from A. Dickson (Scripps Institution of Oceanography). Dissolved oxygen calibrations followed manufacturer water-saturated air calibration protocols.

 

Total Alkalinity Sampling and Storage

Discrete seawater samples (100 mL) for total alkalinity (TA) were collected twice weekly.

Sampling procedures followed best practices described in Dickson et al. (2007):

Rinsed sample bottles three times with sample water.

Collected seawater in acid-cleaned 100 mL borosilicate bottles.

No headspace was left to prevent gas exchange.

Samples were analyzed within 12 hours of collection and stored at ambient temperature in the dark until analysis.

 

TA Determination

TA was determined using open-cell potentiometric titration on a Metrohm 877 Titrino Plus equipped with a Metrohm 9101 Herisau glass pH electrode.

All titrations were standardized using certified reference materials (CRM, batch number provided by Dickson Laboratory). Electrode slope, offset, and drift were checked daily.

Carbonate system variables (DIC, pCO2, HCO3-, CO32-, and Omega aragonite) were calculated using the R package seacarb.

 

Calcification Measurements

Calcification rates were quantified using buoyant weighing following Jokiel (1978). Colonies were weighed on day 0 and day 30. Calcification rate (G) was calculated as:

G (g CaCO3 d-1) = 1.54 × (Wf – Wi) / d

where 1.54 g cm-3 is the density of aragonite, Wf and Wi are final and initial buoyant weights, and d is the exposure duration in days.

Three colonies per species per mesocosm were randomly selected for further biometric analysis. Three branch tips per colony were sampled immediately before and after the exposure period. Collected fragments were frozen at -20 C until post-processing.

Mortality was low and did not differ across treatments; dead colonies were excluded from analysis. All corals were returned to the reef at the conclusion of experiments.

 

Post-Hoc Processing of Coral Samples

At the end of each experimental season, all frozen branch tips were transported to Texas A&M University–Corpus Christi for tissue, symbiont, pigment, and skeletal analysis.

Tissue Removal and Homogenization

Branch tips were thawed and sprayed with phosphate-buffered saline (PBS) using a Paasche airbrush.

Tissue slurry (25 mL) was sonicated for 20 seconds (Qsonica ultrasonic processor).

Samples were vortexed and centrifuged (VWR International centrifuge) to separate host tissue and symbionts.

 

Host Tissue Analyses

Aliquots of the host fraction were used to quantify protein concentration using spectrophotometry (SpectraMax M3). PBS was used as a blank.

 

Symbiont Analyses

Symbiont pellets were resuspended and counted using a Bright-Line hemocytometer (Hausser Scientific) under 10× magnification on a Leica ICC50W microscope.

 

Pigment Analyses

Chlorophyll a and c were extracted using 100 percent acetone. Absorbance was measured on a SpectraMax M3 spectrophotometer, with acetone blanks. Calculations followed Parsons et al. (1984).

 

Skeletal Preparation and Biometric Measurements

After tissue removal:

Skeletons were soaked in 10 percent bleach to remove remaining tissue.

Samples were dried for 4 hours at 60 C (DX302C drying oven).

Dry weights were recorded using a VWR-4002B2 analytical balance.

Skeletal volume was measured by water displacement in a 100 mL graduated cylinder.

Three-dimensional skeletal scans were generated using an Einscan-SE scanner and processed in MeshLab to estimate total surface area.