Abstract

A common, but not universal, effect of ocean acidification on benthic foraminifera is a reduction in the growth rate. The miliolid Archaias angulatus is a high-Mg (>4 mole% MgCO3), symbiont-bearing, soritid benthic foraminifer that contributes to Caribbean reef carbonate sediments. A laboratory culture study assessed the effects of reduced pH on the growth of A. angulatus. We observed a statistically significant 50% reduction in the growth rate (p < 0.01), calculated from changes in maximum diameter, from 160 μm/28 days in the pH 8.0/pCO2air 480 ppm control group to 80 μm/28 days at a treatment level of pH 7.6/pCO2air 1328 ppm. Additionally, pseudopore area, δ18O values, and Mg/Ca ratio all increased, albeit slightly in the latter two variables. The reduction in growth rate indicates that under a high-CO2 setting, future A. angulatus populations will consist of smaller adults. A model using the results of this study estimates that at pH 7.6 A. angulatus carbonate production in the South Florida reef tract and Florida Bay decreases by 85%, from 0.27 Mt/yr to 0.04 Mt/yr, over an area of 9,000 km2.

INTRODUCTION

Archaias angulatus (Fichtel & Moll, 1798), a soritid symbiont-bearing benthic foraminifer (Hallock, 1999), is useful in determining past water depths, sedimentary facies analysis (Wilson, 2006), and reefal and environmental change (Cockey et al., 1996; Souder et al., 2010). This species (Fig. 1) is common in shallow waters throughout the West Indies and Caribbean, and is particularly dominant in the shallow waters of Florida Bay (Hallock & Peebles, 1993). As such, it is a major source of foraminiferal carbonate sediments around Caribbean reefs (Hallock et al., 1986) and the South Florida shelf, where it constitutes >20% of the foraminiferal population (Lidz & Rose, 1989). Like many other carbonate-secreting biotic elements, its response to ocean acidification (Caldeira & Wickett, 2003) is unknown (Kleypas et al., 2005).

Ocean acidification (Caldeira & Wickett, 2003) occurs when atmospheric carbon dioxide (CO2) is absorbed by seawater, increasing aqueous pCO2, altering oceanic carbonate chemistry, and reducing the pH (Doney et al., 2009). Atmospheric CO2 concentration has increased from a pre-industrial 280 ppm to 400 ppm today (Keeling et al., 2013). The Intergovernmental Panel on Climate Change (IPCC) RCP8.5 emission-driven simulation projects that by 2100 atmospheric CO2 may reach a concentration of 985 ± 97 ppm (Collins et al., 2013).

The surface ocean contains >40 times more CO2 than the atmosphere, and acts as both a buffer and conduit to deeper water for atmospheric carbon (Broecker et al., 1982; Watson & Orr, 2003). Atmospheric CO2 rapidly equilibrates with seawater in the form of H2CO3, and dissociates to form HCO3 and CO32−. This process in turn results in increased concentrations of dissolved inorganic carbon (CT) with no change in total alkalinity (AT; Butler, 1991). As the oceanic CO2 reservoir absorbs increased quantities of atmospheric CO2, there is a concomitant increase in H2CO3, decrease in pH, and a reduction of saturation state (Ω) with respect to carbonate minerals (Broecker et al., 1971; Feely et al., 2009). Recent measurements (2010, 2011) of Ω indicate that >20% of Arctic Ocean surface water is already undersaturated with respect to aragonite (Robbins et al., 2013).

Laboratory experiments have shown that some taxa [e.g., Marginopora vertebralis Quoy & Gaimard, Baculogypsina sphaerulata (Parker & Jones)] have reacted to increased pCO2 with increased rates of calcification (Fujita et al., 2011; Vogel & Uthicke, 2012) and photosynthesis (Uthicke & Fabricius, 2012). However, the majority of studies have shown that ocean acidification will have deleterious effects on planktic foraminifera (Gonzalez-Mora et al., 2008; de Moel et al., 2009; Moy et al., 2009; Manno et al., 2012) and benthic foraminifera (Kuroyanagi et al., 2009; Dias et al., 2010; Fujita et al., 2011; Vogel & Uthicke, 2012; McIntyre-Wressnig et al., 2013). These effects include reduced calcification (Kuroyanagi et al., 2009; Moy et al., 2009; Haynert et al., 2011) and decreased growth rates (Manno et al., 2012; Reymond et al., 2013), which may reduce the fitness of calcifying foraminifers and lead to ecological replacement by non-carbonate producers (Kuffner et al., 2008; Dias et al., 2010). Previous studies on the response of calcareous benthic foraminifers to ocean acidification have found that, as pH decreases, growth rates and test preservation are adversely affected, although this response is not uniform (Bernhard et al., 2009; Kuroyanagi et al., 2009; Dias et al., 2010; Dissard et al. 2010; Fujita et al., 2011; Haynert et al., 2011; Sinutok et al., 2011; Uthicke et al., 2012; Uthicke & Fabricius, 2012; Vogel & Uthicke, 2012; McIntyre-Wressnig et al., 2013; Reymond et al., 2013; summarized in Keul et al., 2013).

Physical and chemical methods of quantifying carbonate production (e.g., amount of organic and inorganic carbon fixed, shell weights, growth rates, ultrastructural changes) have been used to estimate the response of biocalcifiers to changing CO2 concentration (Langdon et al., 2000; Barker & Elderfield, 2002; Ries et al., 2009). The biocalcification rates of individual taxa can be used to model regional rates of carbonate sediment production (Murray, 1967; Muller, 1974; Hallock et al., 1986; Harney & Fletcher, 2003).

Because Archaias angulatus is an important source of carbonate sediments in the Florida Keys, we assessed its potential responses to ocean acidification in a laboratory study of the effects of reduced pH on morphology (i.e., test growth rate and pseudopore size) and test chemistry (i.e., stable isotope composition and Mg/Ca ratio). Further, the observed change in the growth rate was used to model the impact of increased CO2 on A. angulatus carbonate sediment production in the Florida Keys.

Archaias angulatus, A Soritid Benthic Foraminifer

Benthic foraminifers are a prolific source of carbonate sediments (Hallock et al., 1986; Debenay et al., 1999) that can be used to assess the ecological status of benthic environments and to elucidate the paleoecological history of marine environments (Hallock et al., 1986, 2003; Hallock & Glenn, 1986; Cockey et al., 1996; Hallock, 2000; Barker & Elderfield, 2002). Additionally, they are useful in resolving paleo-water depths and making facies interpretations of the geologic past (Wilson, 2006). Finally, they are useful indicators of reef health (Cockey et al., 1996; Hallock et al., 2006).

Archaias angulatus has a planispiral, involute, porcelaneous test that is imperforate, covered with pseudopores, and composed of high-magnesium (10–15%) calcite (Blackmon & Todd, 1959; Culver et al., 1982; Cottey & Hallock, 1988; Hallock & Peebles, 1993; Macintyre & Reid, 1998; Hallock, 2000; Wilson, 2006). Ranging from sea level to at least 30-m water depth, the species is moderately euryhaline and eurythermal, but cannot tolerate reduced oxygen levels. Reproduction, which kills the parent, occurs near 9 months, at which point the adult is ~2.2 mm in diameter, although older individuals over 3.6 mm diameter have been documented (Hallock et al., 1986).

Test alterations have been observed in both living and dead specimens. Reid & MacIntyre (1998) reported that recrystallization of the test to minimicrite (<1 μm) begins while the foraminifera is still living, possibly caused by increased internal pCO2 (Macintyre & Reid, 1995). Dissolution of Archaias angulatus has been noted in undersaturated microenvironments with a pore water pH of 7.3 near the sediment-water interface (Cottey & Hallock, 1988). Tests of A. angulatus frequently glauconitize postmortem, possibly as a result of bacterial alteration in anoxic conditions (Wilson, 2006). Archaias angulatus, which moves by means of its pseudopodia, inhabits a range of substrates, including rubble, algal turf, seagrass, and macroalgae (Hallock & Peebles, 1993); higher densities have been associated with mixed seagrass-macroalgal substrates (Hallock et al., 1986). Lévy (1991), who surveyed A. angulatus along a transect from the Everglades to the Florida Straits, found that it was one of the most abundant species of miliolid foraminifers in water shallower than 3 m. Minimum and maximum seasonal values of seawater parameters between April 2009 and February 2012 (Manzello et al., 2012) near the Keys Marine Laboratory in Layton, Florida (location where experimental specimens were gathered), are shown in Table 1.

Stable Isotope and Mg/Ca Chemistry of Archaias angulatus

Stable isotope values (VPDB scale) for A. angulatus have been reported between approximately −1.0‰ and +1.0‰ for δ18O and between +1.0‰ and +4.5‰ for δ13C. In both cases, samples gathered from the same locations tended to exhibit local variations of approximately 1.0‰ (Gross, 1964; Wefer, 1985; Brasier & Green, 1993). Equilibrium and kinetic isotope effects cause fractionation of oxygen isotopes between CO2(aq), HCO3, and CO32−, the anions involved in carbonate precipitation; CO32− is depleted 16‰ relative to HCO3 and HCO3 is depleted 24‰ relative to CO2(aq) (Spero et al., 1997; Zeebe, 1999). Carbon isotopes are subject to equilibrium fractionation but are also subject to a metabolic and abiotic kinetic fractionation (Spero et al., 1997). However, miliolids such as A. angulatus precipitate carbon directly from seawater (ter Kuile & Erez, 1987).

Studies thus far report different species have varied responses to changes in pH and pCO2. A study of oxygen stable isotopes in Amphistegina spp. documented an increase of δ18O as pH and [CO32−] decreased (Rollion-Bard et al., 2008). Similarly, an earlier study of the symbiotic planktic foraminifer Orbulina universa d’Orbigny by Spero et al. (1997) demonstrated that lower pH levels, causing a reduction in δ18O-depleted [CO32−] and increase in δ18O-enriched [HCO3], resulted in increased δ18O and δ13C values. Elderfield et al. (2006) presented the carbonate ion hypothesis, which proposed that lower [CO32−] results in lower Mg/Ca in benthic foraminifera. However, analysis of the low-Mg, symbiont-barren, rotalid Ammonia tepida (Cushman) grown at different pCO2 concentrations reported no relationship between [CO32−] and the incorporation of Mg into the test (Dissard et al., 2010). On the other hand, a similar study by Raitzsch et al. (2010) reported that test Mg content declines as the Ωcalcite increases. They found a similar relationship in the high-Mg symbiont-bearing rotalid Heterostegina depressa d’Orbigny, although the effect was less pronounced. Archaias angulatus tests are also formed from high-Mg calcite, recently reported at 10–15 mol% (Toler et al., 2001; Souder, 2009). Until now, the response of A. angulatus to decreased pH and accompanying decreases in [CO32−] has not been reported. Because Mg interferes with calcification, weakens the calcite structure, and makes calcite dissolve more easily, we expect organisms secreting high Mg-calcite tests to be at a disadvantage as increasing atmospheric CO2 decreases the seawater calcite saturation state (Ωc).

METHODS

Sampling and Specimen Preparation

Samples of Archaias angulatus were collected in March 2011 at the Keys Marine Laboratory’s (KML) Dolphin Cove (Layton, Florida; 24.8268°N, 80.8144°W) in water ~1.5-m deep. Seawater temperature (Tseawater = 22.5°C), salinity (S = 35.1), and pH (~8.2) were measured at KML with a calibrated Oakton water analyzer. A water sample was later analyzed for total alkalinity (AT = 2239 μmol kg−1SW) at the U.S. Geological Survey (USGS) Coastal and Marine Science Center in Saint Petersburg following the methods of Yao & Byrne (1998). Clumps of brown algae containing A. angulatus were gathered into seawater-filled 2-L transparent plastic zipper storage bags.

The bags were sealed and agitated vigorously to dislodge A. angulatus specimens from the algae, which were then discarded. Seawater containing the dislodged A. angulatus was filtered through a 63-μm sieve in the KML wet lab. The foraminiferal residue was stored in 500-mL plastic jars containing 1–2 cm of the sieved sediment and fresh local seawater. The jars were placed in a cooler to maintain a constant temperature of ~22.5°C and transported the same day to the University of South Florida (USF) College of Marine Science (CMS) Reef Indicators Lab in Saint Petersburg. Approximately 40 L of sediment from the KML lagoon were collected for use as substrate in the experimental tanks. Local filtered seawater (AT = 2257 μmol kg−1SW) from Clearwater Beach, Florida, was used in the tanks. Random jars of sediment and associated A. angulatus specimens were emptied into a 10-L aerated holding tank and placed in an environmental chamber (T = 25°C). Extra 500-mL plastic jars containing sediment and A. angulatus specimens were also placed in the environmental chamber. Water in the aerated holding tank was changed with fresh KML seawater each week. Containers of extra seawater were stored in a dark cabinet.

Preparation of A. angulatus specimens for the experiment was performed at CMS. A subsample of sediment was removed from the holding tank and placed in a large Petri dish. The sediment was examined with a binocular microscope; specimens of A. angulatus were gently removed with a fine brush and sequentially placed in seven seawater-filled Petri dishes until ~20 individuals were in each dish. Only individuals with a green tint, an indication of healthy living specimens containing algal symbionts (Lee, 2006), were selected. The contents of each dish were transferred to a microscope slide and digitally photographed. The contents on each slide were then transferred into a 20-μm mesh bag (25 × 25 mm) that contained two algal-encrusted segments of live Halimeda spp. for food. The mesh bags were heat-sealed and transported to the USGS in a jar of seawater, where each bag was randomly placed in one of six tanks used in the experiment (described below). A 10-cm length of monofilament line, attached to a stainless steel weight, was attached to each mesh bag to ensure continuous submersion. These bagged replicates were placed near the center of each tank but away from the aerator. When present, air bubbles were gently dislodged from the bags. Every seventh replicate was immersed in deionized (DI) water for 15 minutes, air-dried, and archived as a reference set. A total of 42 bags were sorted; 36 were placed in tanks (six bags/tank) and six used in the archival reference set. Of the 36 bags in the tanks, 29 were ultimately used in the subsequent analysis. The remaining seven bags had unequal numbers of foraminifers before and after the experiment, attributed to leaks in the mesh bag caused by seam failure.

Experimental Setup

The culturing apparatus consisted of six clear glass 38-L tanks filled with ~20 L of seawater, a 2–3-cm thick aragonite sediment substrate, a 72 W light fixture, a transparent plastic lid, an aerator, and an automated pH-controlled CO2(g)-injection apparatus (Fig. 2). Tank AT was maintained at a target value of 2325 μmol kg−1SW as detailed in Appendix 1. The tanks were numbered sequentially from 1–6. Tanks 1, 3, and 5 were the experimental treatment tanks, and 2, 4, and 6 were the control tanks. Details of tank chemistry treatment are provided in Table 2 and Figure 3. A supplemental air conditioning unit in the laboratory provided overpressure to reduce the ambient CO2 levels. Laboratory air was measured six times, and [CO2] ranged between 390–410 ppm. Laboratory temperature was maintained by central air conditioning at approximately 24°C.

The experiment was conducted in a six-week trial, consisting of three control tanks (pH 8.0) and three pH 7.6 treatment tanks. The pH controllers automatically injected CO2(g) into each treatment tank’s headspace when that tank’s pH rose above the target value. Each silicone CO2(g) injection line was mounted 2 cm above the water level at the end of the tank opposite the pH probes. Injected CO2(g) was mixed with the tank’s seawater by the submersible aerator. The controllers were programmed to begin CO2(g) injection when treatment tank total-scale pH (pHT) was >7.61. Injection ceased when pHT was <7.60. Injections of CO2(g) occurred approximately every two hours; each injection cycle generally lasted 15 minutes and delivered ~2 ml of CO2(g)/cycle to each treatment tank. Treatment tank headspace [CO2(air)] averaged 700 [standard deviation (σ) = 87] ppm before injection and 1200 (σ = 265) ppm after injection. Treatment tank pHT averaged 7.61 (σ = 0.05). Control tank headspace [CO2(air)] averaged 412 (σ = 3) ppm. Control tank pHT averaged 8.00 (σ = 0.05).

Mesh bags of A. angulatus were removed after ~14 days, 28 days, and 42 days of immersion. Upon removal from the tanks, mesh bags were immersed in tap water for five minutes, DI water for one minute, and then set aside to air dry. The specimens were later transferred to archival slides and digitally photographed in preparation for analysis. Detailed methods are described in Appendix 1.

Growth and Pseudopore-Size Analysis

Digital images of specimens were analyzed using ImageJ version 1.45s software (Rasband, 2007). The perimeter of each specimen image was outlined to calculate the maximum diameter across the perimeter, referred to as the Feret diameter (Francus, 2004). It was inferred that the difference between pre-experiment and post-experiment diameter measurements represented growth that occurred as newly formed terminal chambers during the experiment. This assumption was verified by visually comparing pre-and post-experiment images of the same individuals. Detailed measurements are located in Appendix 2.

Slides of A. angulatus were randomly selected for scanning electron microscope (SEM) analysis, and random specimens were mounted flat on aluminum SEM specimen mounts using conductive adhesive tabs. Specimens were imaged in variable-pressure mode at the USF Electron Microscopy Lab with a Hitachi S-3500N SEM. All pseudopores that were completely within three 50 × 50 μm regions located on the terminal chambers of each specimen were counted and measured using the software’s “analyze particles” tool.

Isotope Ratio Mass Spectroscopy (IRMS)

Water samples from the tanks and the KML lagoon were analyzed for δ18O of seawater and δ13C of DIC. These samples were collected for stable isotopic analysis by completely filling 125-mL serum bottles, fixing with 50 μL of saturated mercuric chloride, and sealing with greased Teflon crimp caps. Analyses were completed at the USF School of Geosciences Stable Isotope Laboratory using a Thermo-Finnigan Delta V 3-kiloelectron volt (keV) Isotope Ratio Mass Spectrometer coupled to a Finnigan GasBench II preparation device.

Archaias angulatus specimens that were approximately the same diameter and which were visually and measurably larger in post-experiment images compared with pre-experiment images were selected at random. Specimens were weighed, sonicated in 25 μL of DI water for two minutes, rinsed with 500 μL of DI water, immersed in a 1% (v/v) H2O2 alkali-buffered oxidizing solution (Barker et al., 2003), heated for eight minutes at 70°C, and then sonicated for an additional two minutes. The oxidizing solution was replaced and the specimen was again heated for eight minutes, sonicated for two minutes, then rinsed twice with 500 μL of DI water and dried at 60°C. Specimens were reweighed. Finally, terminal chambers were removed from the test for use as the analyte until the required analysis mass (200 μg) was reached. The remainder of the specimen was set aside.

Analyses of δ18O and δ13C of carbonate samples were completed by flushing 12-mL Labco exetainers, containing 200 μg of sample, with He gas, adding 2.5 mL of 103% phosphoric acid (H3PO4), and equilibrating for 24 hr. Analyses of δ18O of H2O were completed by equilibrating 200 μL of sample with ~12 mL headspace of an ~0.3% CO2-He mixture in Labco exetainers (equilibration method after Epstein & Mayeda, 1953). Analyses of δ13C of DIC were completed by injecting 1-mL aliquots of sample water into ~12-mL vials that were pre-flushed with He and pre-filled with 1 mL of 85% H3PO4 (Assayag et al., 2006). In all three types of analyses, the relevant isotopic composition of the headspace pCO2 was measured after equilibration in controlled temperature conditions. The standards used as a reference for the δ-scale are VSMOW for H2O and VPDB for DIC and carbonate. Analytical precision (2σ) on water standards used was not >0.1‰ for δ18O and δ13C, and for carbonate samples <0.3‰ for δ18O (with one exception of 0.64‰) and <0.13‰ for δ13C.

Inductively Coupled Plasma Optical Emission Spectrometry (ICP-OES)

Elemental Mg/Ca ratios were determined using a PerkinElmer 7300 Dual View Inductively Coupled Plasma Optical Emission Spectrometer (ICP-OES) at the St. Petersburg USGS Coastal and Marine Science Center. An internal gravimetrical standard solution (IGS) was measured for Mg/Ca before and after dissolving each foraminiferal sample to correct for instrumental drift and noise (Schrag, 1999). The average corrected IGS precision for Mg/Ca was 0.0436 mmol mol−1 (1σ, n = 159). A second standard, homogenized powder of Euronorm certified reference material (ECRM), analyzed for Mg/Ca to test for any potential matrix effects had an average corrected precision of 0.0638 mmol mol−1 (1σ, n =16).

Archaias angulatus specimens were selected for ICP-OES analysis using the same procedure described in the Isotope Ratio Mass Spectroscopy (IRMS) section above. They were prepared by removing the terminal chambers from the test for use as analyte until sufficient material was gathered (250-μg pretreatment). Sample preparation was then performed by sonication, oxidation, and leaching, following the methods of Barker et al. (2003).

Statistical Analyses

Mean values in the tables are shown with their standard error of mean (mean ± SEM). Results were considered significant at p < 0.05. The Anderson-Darling test was used to confirm that data were normally distributed. A series of 1-sample t-tests were used to evaluate the mean temperature, salinity, pH, saturation state (Ω), and alkalinity differences between treatment and control tanks. A 2-sample t-test was used to evaluate the differences in weekly growth rates of the treatment and control, pre- and post-oxidation weights of the treatment and control foraminifers, pseudopore size of new growth of specimens from the treatment and control, and pre- and post-treatment pseudopore size of individual specimens from the treatment and control. One-way analysis of variance (ANOVA) was used to test the effect of pH on the oxygen and carbon isotope values. A 2-sample t-test was used to evaluate the Mg/Ca levels of specimens from the control and treatment tanks. Least-squares regression was used to test the effect of foraminiferal diameter on weight. All data were analyzed using Minitab 15 statistical software.

Calculations

Archaias angulatus daily test growth was calculated by subtracting the mean diameter of specimens in each bag measured before the experiment from the mean size of specimens measured after the experiment and dividing by the number of days in culture. The weekly growth rate was calculated by multiplying the daily growth rate by seven. Test diameters of mature (i.e., 1-year old individuals) were calculated by multiplying the modeled weekly growth (μm) by 52. Because weights of A. angulatus (Fig. 4) are strongly related to diameter (p < 0.01, r2 = 0.83), the regression equation

 

Weight (mg)=-0.545+0.000745×diameter(μm)

was used to calculate the weight of mature individuals from their diameters.

The contribution of A. angulatus to carbonate sediment production at reduced pH levels was estimated by multiplying the calculated value of A. angulatus production at pH 8.2, 60 g/m2/yr (Hallock et al., 1986), by the ratio of modeled adult weights at pH 8.0 (control) and pH 7.6 (treatment). These numbers were in turn multiplied by the area of observed A. angulatus habitat (9,000 km2).

Because A. angulatus precipitates calcite from seawater-filled vesicles in isotopic equilibrium (i.e., with little or no vital effect), the δ18O value of the precipitated calcite is highly dependent on that of local [HCO3] and [CO32−], essentially precipitating as the weighted average of the two ions, each of which are in equilibrium with local seawater (Zeebe, 1999). A mass-balance calculation at pH 7.6 has the following relationship:

 

δ18O=0.038CO32-(18.4δ18O)+0.962HCO3-(34.3δ18O)=33.7(SMOW)=2.7(PDB)

and at pH 8.0:

 

δ18O=0.097CO32-(18.4δ18O)+0.903HCO3-(34.3δ18O)=32.8(SMOW)=1.9(PDB).

These equations predict that δ18O should be enriched by 0.8‰, if the pH is reduced by 0.4 at constant temperature.

Elderfield et al. (2006) reported a temperature-corrected relationship between Mg/Ca and [CO32−] for Cibicioides wuellerstorfi (Schwager), in which

 

(Mg/Ca)Δ[CO32-]=-0.15+0.0086Δ[CO32-].

Combining this equation (which was derived from C. wuellerstorfi) with the control and treatment [CO32−] of this experiment (Table 2) predicts that the control specimens should have a Mg/Ca value 0.7 mmol/mol greater than the treatment.

RESULTS

Water Chemistry

Tank chemistry (control and treatment) is shown in Figure 3. There were no significant differences in the temperatures (n = 54, p = 0.984), salinity (n = 54, p = 0.963), or alkalinity (n = 54, p = 0.996) of the tanks; pH (n = 27, p = 0.493) or Ω (n = 27, p = 0.992) of the control (pH = 8.0) tanks; and pH (n= 27, p = 0.997) or Ω (n = 27, p = 0.995) of the treatment (pH = 7.6) tanks. When alkalinity is constant, as it was in this study, there is a strong correlation (p < 0.0001, r2 = 0.98) between pH and both [CO32−] and Ω.

Growth Analysis

Growth analysis (i.e., increase in diameter) was performed on specimens (n = 585) from 29 bags that contained equal numbers of individuals before and after the experiment. The mean diameter and increase in diameter for the replicates (n = 29) of Archaias angulatus are shown in Table 3. The presence of grazing tracks on the Halimeda segments enclosed in each bag for food indicated that the specimens were feeding.

There was a significant effect of treatment pH on the mean increase in diameter of foraminifer replicates [t(22) = −3.13, p = 0.005]. Mean weekly test growth (Fig. 5) in the pH 8.0 control averaged 2.8±0.3%, while in the pH 7.6 treatment the growth rate was 1.4±0.2%. The monthly increase in diameter of the control specimens, based on weekly growth (Table 4), was approximately 160 μm/28 days. The pH 7.6 treatment specimens had a growth rate 50% that of the control at 80 μm/28 days. There was no effect of pH on the difference of the pre- and post-oxidation weights (Fig. 4; n = 17, p = 0.89).

Pseudopore Area

The size of individual pseudopores (n = 811) increased as the pH decreased (Fig. 6). There was a significant effect of treatment pH on the mean pseudopore area after six weeks of culture [t(803) = 6.46, p < 0.001]; pseudopores from the pH 7.6 treatment (25.7±0.5 μm2) were 18% larger than pH 8.0 control pseudopores (21.7±0.4 μm2). A comparison of pseudopore size from the same specimens before and after six weeks of treatment (Fig. 7; n = 120) indicates that newly grown pseudopores at pH 7.6 were 13% larger than pre-treatment pseudopores [t7.6(52) = 2.24, p = 0.03], but pseudopore areas from the pH 8.0 control tanks were not significantly different before and after treatment [t8.0(57) = −0.69, p = 0.49].

Geochemical Analyses

There was a marginally significant effect of treatment pH on δ18O (VPDB) values (Fig. 8; n = 41) of A. angulatus after six weeks [F(2,40) = 3.21, p = 0.051] and no significant effect of pH on δ13C (p = 0.28). Mean δ18O of tank seawater (VSMOW) was 1.6±0.2; δ13C values were −19.7±0.6. A water sample from the Dolphin Cove lagoon had a δ18O value of 1.99 and δ13C value of −20.5. The Schrag-corrected Mg/Ca ratios of A. angulatus specimens after six weeks of treatment at pH 7.6 treatment (Fig. 9; n = 34) differed significantly from the Mg/Ca ratios of the pH 8.0 control specimens [t(17) = 2.17, p < 0.05]; treatment specimens had Mg/Ca ratios (131.4±2.3 mmol/mol) ~6 mmol/mol greater than that of the control specimens (125.4±1.5 mmol/mol).

DISCUSSION

Growth Rates

This study experimentally investigated Archaias angulatus’s physiological (i.e., growth rate and pseuodopore size) and chemical (i.e., stable isotope composition and Mg/Ca ratio) responses to different seawater pH levels induced by the injection of CO2(g) into the culture media. The treatment level was set at pH 7.6, corresponding to atmospheric CO2 levels (RCP 8.5, 1,300 ppm) projected to occur in the next 150 years (Collins et al., 2013). At constant alkalinity, pH is a proxy for [CO32−] and, therefore, Omega;calcite (Riebesell et al., 2010). The pooled responses of the treatment (pH 7.6) and of the control (pH 8.0) identified overall trends, although the variable responses of replicate bags, attributed to microenvironmental variations, were consistent with the findings of other researchers (Fujita et al., 2011; Glas et al., 2012; Vogel & Uthicke, 2012).

The reduction of the growth rate at lower pH (Fig. 5) and, therefore, lower [CO32−] (Table 2) indicates that calcification in A. angulatus is dependent upon externally supplied CO32−. This reliance is a consequence of its characteristic miliolid calcification method, which uses a supersaturated calcitic solution, largely the result of photosynthetic pH increases and generated directly from seawater, to precipitate the high-Mg calcite rods used to construct the test, as described by Angell (1980), Lowenstam & Weiner (1983), ter Kuile & Erez (1988), Wetmore (1999), Erez (2003), Goldstein (2003), and de Nooijer et al. (2009).

Because A. angulatus is dependent on externally supplied carbonate, even a mild reduction in [CO32−] will reduce growth. The consistent diameter-to-test-weight ratio independent of pH (Fig. 4) indicates that A. angulatus does not sacrifice structural integrity for increased size. The recorded growth response can be compared to studies of other dinoflagellate-bearing soritids, including Amphisorus hemprichii Ehrenberg (Fujita et al., 2011), Marginopora kudakajimensis Gudmundsson (Kuroyanagi et al., 2009), and M. vertebralis (Vogel & Uthicke, 2012). Fujita et al. (2011) found that calcification of Am. hemprichii decreased at higher pCO2. However, unlike A. angulatus, the change in size was accompanied by a pronounced change in the weight-to-diameter ratio, which decreased with increasing pCO2 (i.e., the foraminifers grew nearly as large, but weighed less). The results for M. kudakajimensis are similar to those of Am. hemprichii, although the response was less pronounced at intermediate pH levels (i.e., 7.9). In contrast, M. vertebralis increased its growth rate at very high CO2 levels (i.e., 1169 ppm, 1662 ppm) while maintaining its weight-to-size ratio. Each of these miliolids has dinoflagellate symbionts, whereas A. angulatus hosts chlorophytes. Specifically, Am. hemprichii and M. kudakajimensis both host Symbiodinium sp. and M. vertebralis hosts Gymnodinium sp. (Lee & Anderson, 1991). Symbiodinium (better known as zooxanthellae) is associated exclusively with symbiosis, whereas the related genus Gymnodinium contains free-living forms, some of which occur in acidic freshwater (Graham & Wilcox, 2000). When combined, the results of these studies show a potential relationship between the specific response of different foraminifers to ocean acidification and the symbiont type they host.

Using size and growth data (Table 5) reported by Hallock et al. (1986), the average size of A. angulatus in the experimental bags indicated that individuals were, on average, <6 months old at the end of the experiment. A longer lifespan could offset the pH-associated reduction in growth rate. However, such an increase in lifespan is unlikely because monthly mortality rates are strongly correlated with size (Hallock et al., 1986). The high mortality rate of smaller specimens precludes the increased time of survival necessary to “catch up” to untreated growth. In short, this reduction in growth rate and adult size indicates that A. angulatus living in a high-CO2 world probably would not live sufficiently long to grow as large as they are now, and because they will produce fewer offspring, the population will shrink. Reduction in the size of biotic lineages during stressful periods, such as mass extinctions, has been documented in the fossil record (Fermont, 1982; Keller, 1988; Twitchett, 2007; Borths & Ausich, 2011). These size reductions of fauna are referred to as the Lilliput Effect (reviewed in Harries & Knorr, 2009).

Pseudopore Size

Increased or decreased light penetration and enhanced gas exchange have been invoked to explain the presence of pseudopores on porcelaneous foraminifera (Hottinger, 1986; Lee & Hallock, 1987). However, these explanations do not account for an increase in pseudopore size (Figs. 6 and 7). As the pH and [CO32−] decreased, existing pseudopores increased in size and newly formed pseudopores were larger. Because pseudopores in the pH 8.0 control treatment remained the same size, dissolution of the test near the pseudopores is the most likely cause of increased pseudopore size in the lower pH treatment. The solubility and susceptibility to dissolution of calcite increases with Mg content (Andersson et al., 2008); an increase of Mg in A. angulatus calcite was noted at lower pH (Fig. 9). This combination of decreased Ωcalcite, gas (i.e., CO2) exchange around pseudopores, and a higher Mg/Ca ratio in secreted foraminiferal calcite may have promoted enlargement of pseudopores by dissolution.

The stages of dissolution of the A. angulatus test were investigated by Cottey & Hallock (1988). Indicators of extreme dissolution (e.g., exposure of the inner wall, pseudopore inversion, loss of test wall) were not noted in the cultured specimens from this study, although coalescence of pseudopores and dissolution at the pseudopore perimeter and interior (Fig. 10) was sometimes present. The angular, stepped boundary between pseudopores and the adjacent lateral walls exposes a larger proportion of unbonded surface area to the cytoplasm than the flat surface of the lateral walls (Morse & Mackenzie, 1990), an area where dissolution is first expressed in the form of enlarged pseudopores. Additionally, Macintyre & Reid (1998) showed that A. angulatus tests can begin to micritize while still living; a textural change of the calcite that forms the bulk of the test walls from Mg-calcite rods (1–2-μm long × 0.1-μm wide) to finer and denser minimicrite (0.1-μm equant crystals) could cause a deflationary reduction in volume without visibly damaging the outermost layer of tiled calcite.

Carbonate Production

Langer (2008) estimated that large benthic foraminifera annually produce a minimum of 130 Mt of CaCO3, which is ~2.5% of the oceanic carbonate budget. In the absence of other changes, a reduction of the growth rates of individual foraminifers will necessarily reduce the rate of carbonate production (Hallock, 1981). In a comprehensive study of sediment production in Key Largo Sound, Hallock et al. (1986) estimated Archaias angulatus carbonate production on shallow carbonate shelves at 60 g/m2/yr, based on a winter abundance of 5 × 103 individuals/m2 and a summer abundance of 15 × 104 individuals/m2; surface water in this vicinity had an estimated spring and summer pH > 8.0, measured between 1997 and 2001 (Millero et al., 2001). Studies of the carbonate production of other soritids have reported similar results. Lutze & Wefer (1980), in a study of Cyclorbiculina compressa (d’Orbigny) (Archaiasinae) from Bermuda, estimated a carbonate production rate of 60 g/m2/yr at an adult abundance of 500 individuals/m2; Hohenegger (2006), in a study of west Pacific reef flats, reported a carbonate production rate of 43 g/m2/yr at an abundance of 1.6 × 104 for M. kudakajimaensis (Soritinae); Doo et al. (2012) reported a carbonate production rate of 72 g/m2/yr at an abundance of 7.7 × 103 individuals/m2 for M. vertebralis (Soritinae) in an intertidal pool in the southern Great Barrier Reef. Given the similarity of soritid carbonate production values reported (i.e., 59±6 g/m2/yr CaCO3) by these geographically and taxonomically diverse studies, the A. angulatus carbonate production value (60 g/m2/yr) reported by Hallock et al. (1986) was used as a baseline for this study’s model.

In South Florida, A. angulatus is common to abundant from Key Largo (25°5′N, 80°26′W) to Dry Tortugas (24°38′N, 82°55′W) and throughout Florida Bay (Bock, 1971). A study by Lidz & Rose (1989) found that A. angulatus constituted between 20%–80% of foraminifera at depths ≤9 m and with no notable variation in distribution at depths <15 m. In later transects of Florida Bay to the reefs immediately south of the keys near the southern edge of the Florida platform, A. angulatus constituted up to 50% of the foraminiferal assemblage in areas with constant-to-moderate salinity fluctuations (Lidz & Rose, 1989; Lévy, 1991). Two other chlorophyte-bearing miliolids, Laevipeneroplis proteus (d’Orbigny) and Cyclorbiculina compressa, were found in reef rubble at depths ≤30 m (the maximum study depth) with A. angulatus, although abundances varied by species (Baker et al., 2009). In the portion of Florida Bay near mainland Florida, which exhibited larger salinity fluctuations, a similar chlorophytic Archaiasinae, Androsina lucasi Lévy, constituted almost 100% of assemblages (Lévy, 1991).

The following calculations assume that, based on their similarities, An. lucasi will respond to reduced pH in a similar fashion as A. angulatus. The relationship of size and weight of mature specimens to seawater pH is shown in Table 6; smaller specimens will exert the same proportionate effects. In seawater with a pH of 7.6, A. angulatus carbonate production as measured by weight is decreased by 87% relative to Hallock et al.’s (1986) estimate. Discounting the effects of reduced fecundity and assuming a constant growth rate at pH 7.6, this leads to a potential reduction in carbonate sediment production from 60 g/m2/yr (pH 8.2, 1986 estimate) to 8 g/m2/yr (pH 7.6). Such a reduction will have a considerable impact on sediment accumulation rates in South Florida’s shallow-water systems. The area of Florida Bay and the Florida Keys shallower than 15 m (Fig. 11), which is the deepest A. angulatus zone in Lidz & Rose’s study (1989), is ~9,000 km2 (bathymetric data from Robbins et al., 2007). Combining these values (Table 7) and assuming A. angulatus occurs in 50% of the suitable habitats (Bock, 1971) results in an 85% reduction of A. angulatus (and An. lucasi) calcite production at pH 7.6, from 0.27 Mt/yr to 0.05 Mt/yr.

Geochemistry

The δ18O values of A. angulatus increase as the pH and [CO32−] decrease (Fig. 8). This is in agreement with previous work on the relationship between carbonate speciation and δ18O response (McCrea, 1950; Spero et al., 1997; Zeebe, 1999; McConnaughey, 2003). This experiment’s isotopic results (Fig. 8) showed a 0.23‰ enrichment of δ18O when pH was reduced by 0.4. Small variations in tank temperature, vital effects, and the potential inclusion of low δ18O pre-experiment calcite may account for the 0.57‰ deviation from the predicted difference, as calculated in the Methods section. The lack of a significant response of δ13C to reduced pH can similarly be attributed to a combination of dependence on equilibrium, metabolic, and abiotic kinetic fractionation mechanisms (Spero et al., 1997).

Recent work by Andersson et al. (2008) theorized that marine calcifiers containing significant amounts of high-Mg (>4 mole% MgCO3) calcite, which is more soluble than low-Mg calcite or aragonite, should respond more adversely (e.g., slower rates of calcification) to ocean acidification than calcifiers containing little or no Mg. Additionally, because high-Mg calcite and aragonite form the majority of tropical shallow-water sediments (Tucker & Wright, 1990), soluble deposits of high-Mg calcite may become the “first responders” (sensuMorse et al., 2006) to ocean acidification, acting as a CO2 sink by dissolving into Ca2+ and HCO3− before less soluble forms of calcite. For low- and high-Mg calcite taxa, it has been suggested that the Mg content of foraminiferal calcite increases with temperature (Nürnberg et al., 1996); this relationship allows the Mg/Ca ratio to be used in paleothermometry (Hastings et al., 1998; Elderfield & Ganssen, 2000; Lear et al., 2002) and, in conjunction with their δ18O values, to calculate Cenozoic global ice volumes (Billups & Schrag, 2002).

Toler et al. (2001) previously reported Mg/Ca values of 120±13 mmol/mol for A. angulatus, comparable to this experiment’s pH 8.1 archival reference set value of 127±1 mmol/mol (Fig. 9). The slight increase in Mg/Ca from 125±2 at pH 8.0 to 131±2 at pH 7.6 is consistent with previous work (Raitzsch et al., 2010) on the high-Mg rotalid Heterostegina depressa d’Orbigny, which reported a weak relationship between increasing Ω and decreasing Mg/Ca. The increase is the opposite of that predicted by the Elderfield et al. (2006) carbonate-ion hypothesis, which was developed using non-symbiont bearing foraminifera.

More importantly, an 85% reduction in the production of high-Mg calcite by A. angulatus will have a pronounced local effect because miliolids are an importance source of high-Mg micritic mud (Hallock et al., 1986; Langer et al., 1997). Reduced production of high-Mg mud may affect the composition of local sediments and their associated biota. Infaunal organisms may encounter changes in sediment texture as the proportion of low-Mg sediments increases; because low-Mg sediments are less susceptible to dissolution than their high-Mg counterparts (Morse et al., 2007), the mean grain size should increase. Such an increase will, in turn, further reduce the dissolution rate because larger particles have a lower surface-to-volume ratio and, therefore, dissolve at a slower rate than smaller particles (Walter & Morse, 1984).

CONCLUSIONS

At constant alkalinity, lower seawater pH and consequent reductions in [CO32−] result in decreased growth rates of Archaias angulatus. A 0.6 reduction of seawater pH from recent values of 8.2 to a high-CO2-world seawater pH of 7.6 reduces the growth rate of A. angulatus by 87% and results in an order-of-magnitude decrease in local carbonate sediment production from 60 g/m2/yr to 8 g/m2/yr. Consequently, ocean acidification may reduce the contribution of A. angulatus to carbonate sediment production in South Florida from 0.27 Mt/yr to 0.04 Mt/yr. Finally, a reduction in pH appears to increase both the δ18O value and the Mg/Ca ratio; these observed trends agree qualitatively with calculated and previously reported observations of similar taxa, although detailed studies lasting for an entire developmental cycle are needed to clarify and fully quantify the effect.

ACKNOWLEDGMENTS

This study was funded by the USGS Coastal and Marine Geology Program, the Geological Society of America, and the USF Department of Geology. Tony Greco, USF Department of Marine Science-Electron Microscopy Lab, provided pertinent advice regarding the use of the SEM. Jennifer Flannery, USGS, ran the Mg/Ca analyses. Dr. Zachary Atlas, Manager of the USF Center for Geochemical Analysis, and Jessica N. Wilson, USF Geology Stable Isotope Lab IRMS-MS operator, arranged access to and assistance in running the stable isotope analyses. Any use of trade, firm, or product names is for descriptive purposes only and does not imply endorsement by the U.S. Government. This manuscript was improved by comments from two anonymous reviewers. Co-author Dr. Pamela Hallock, Editor of the Journal of Foraminiferal Research, was not involved in editing this paper.

APPENDIX 1

Supplemental Methods

Six 38-L 51 × 27 × 52 cm glass tanks were prepared by washing with a phosphate-free detergent, triple-rinsing with DI water, rinsing the interior for 5 minutes with 1 M HCl, and again triple-rinsing with DI water. Carbonate sediment gathered at KML was sieved with a 2-mm sieve. The coarse fraction was retained and spread out to dry. The sediment was microclaved for 15 minutes and triple-rinsed with DI water to remove residual organic particles. A 2–3-cm layer of this coarse fraction was then evenly spread in each tank, to which ~20 L of seawater was added. A peristaltic pump was used to transfer seawater from the storage container through a 0.45-μm filter into the tank. Initial water level was marked on each tank. A submersible aerator (Hydor Ario-2) which drew air from the tank’s headspace through a silicone tube was installed near the center of each tank.

Clear, 1-cm thick acrylic glass lids were constructed for each tank. A circular 3-cm port was cut into the center of each lid and the edges were lined with rubber sealing tape. Cables and tubes were routed through the port, which was sealed with plastic film (Parafilm). The lids were not completely air tight. Fluorescent light fixtures were placed approximately 15 cm above the tank. The fixtures each contained a 32 W bulb (Florasun 5000K T-8) and a 40 W bulb (Floraglo 2800K T-8). Light intensity was measured periodically with a light meter (Fisher Scientific) and fixture position was adjusted to ensure uniform irradiance. The lights were controlled by a digital timer and were operational from 0700 to 1900 each day. Mean mid-day (1000–1400) light intensity was 1097 lx (σ = 199); there was no significant difference in the median light levels of individual tanks. These light levels were below the threshold for photoinhibition (Walker et al., 2011).

A separate CO2− injection apparatus was constructed for each of the three experimental treatment tanks. Calibrated temperature probes (0.05°C accuracy) and laboratory-grade sealed Ag/AgCl pH probes (0.01 accuracy) were mounted on the tank wall near the ends of these treatment tanks and connected to AquaController pH controllers. The pH probes were calibrated each week relative to a fresh aerated sample of KML water, the pHT of which was determined spectrophotometrically following the methods of Clayton & Byrne (1993).

Approximately 10% of the water (2 L) in each tank was discarded every one–two weeks to maintain Ca2+ concentrations; filtered seawater from a container that had been opened to laboratory air for at least 12 hours was then added to raise the water level back to the initial mark on the tank exterior. After each water change, 1 mL of commercial trace element solution (Seachem Reef Trace) and 0.5 ml of iron solution (1% Seachem Flourish Iron) was added to each tank to maintain trace element concentrations.

Following the guidelines presented by Riebesell et al. (2010), a target AT value of 2325 μmol kg−1SW was used. The CO2SYS program (Lewis et al., 1998) was used to calculate the values for the carbonate system using the observed salinity, temperature, and pHT in conjunction with the target AT (Table 2). Temperature, salinity, pHT, and AT of seawater in each tank was measured and adjusted one to two times weekly using the following methods.

First, temperature was measured with a digital thermometer (0.01°C accuracy) and salinity was measured with a conductivity meter (YSI conductivity cell, K= 1.0, 0.1 accuracy). Next, AT was measured using an open-cell titration with an Ocean Optics USB4000-UV-VIS fiber optic spectrophotometer and bromocresol purple indicator (Yao & Byrne, 1998). Calibration of AT was performed periodically using Dickson certified reference materials (Dickson et al., 2003). Next, the pHT of each tank was measured with an Ocean Optics USB4000-UV-VIS fiber optic spectrophotometer and thymol blue indicator following established methods (Clayton & Byrne, 1993; Dickson et al., 2007). When necessary, reagent-grade NaHCO3(s) was diluted with 50 ml of KML water and added to each tank to raise AT back to the target value. A second measurement of AT was made ten minutes after the end of the CO2-injection cycle following addition of NaHCO3, at which time additional dilutions or additions were made as necessary. Previous trials with the experiment’s tank set-up determined that the addition of 100 μg of NaHCO3 increased AT by ~80 μmol kg−1SW. Individual measurements of foraminifera are located in Appendix 2.

APPENDIX 2

Size Measurements

Table of Archaias angulatus measurements made pre- and post-experiment. Bag indicates the mesh bag identification number; Tank is the experimental tank number; Type indicates whether the bag was part of the treatment (pH = 7.6) or control (pH = 8.0) set; Duration indicates the number of days the bag was immersed; Weeks is the approximate number of weeks the bag was immersed; and Diameter is the Feret diameter (i.e., maximum diameter of the specimen) in μm. This table can be found on the Cushman Foundation website in the JFR Article Data Repository (http://www.cushmanfoundation.org/jfr/index.html) as item number JFR_DR2015002.