To determine the position of P. damicornis larvae in the seawater in response to high pCO₂, larvae were transferred from collection containers to UV-transparent acrylic tubes (UV-T, ACRYLITE Colorless 0070 GT, Evonik Industries, New Jersey, USA) and incubated in situ at two depths, which centered the tubes vertically at ~0.3 m (July 2016) or ~3.3 m (August 2016). The tubes had a wall thickness of 6 mm, were 68-cm long with an inner diameter of 4.5 cm, and were sealed at either end with UV-T acrylic caps. A 6-mm hole was drilled in the top cap so the tubes could be filled with seawater; the hole was later covered with vinyl electrical tape. While the objective was to completely fill the tubes with seawater, in most cases small bubbles were introduced during the filling process and combined when the tubes were positioned vertically to create an air gap of ≤ 1-cm height at the top of the tube. The acrylic in these tubes transmitted ~92% of ultraviolet radiation (UV-R) (280–400 nm) (J. Bergman, unpublished data), ensuring larvae were exposed to ecologically relevant light conditions for shallow seawater (Gleason and Hofmann, 2011). Previous studies of the vertical movement of P. damicornis larvae in acrylic tubes (Harii et al., 2002), as well as preliminary observations of larval behavior used in the present study, show that larvae of P. damicornis exposed to ambient seawater generally aggregate either in the top or bottom 0–2 cm of tubes regardless of light conditions (i.e., light vs. dark). Therefore, the vertically-oriented tubes were marked into two sections for the purpose of scoring the larvae by position: an upper section (21-cm length) and lower section (47-cm length), since we reasoned it would be unlikely that the larvae would accumulate in the center of the tubes. Of the larvae found in the lower section, 83–88% were located in the bottom 23 cm of the tube, which suggested that the greater size of the lower scoring section in the tubes did not upwardly bias estimates of larvae scored as moving downward.

Eight tubes were used for each experiment, during which four were filled with ambient (~400 μatm pCO₂) seawater, and four with treatment (~1,000 μatm pCO₂) seawater that simulated the elevated pCO₂ conditions predicted to occur by 2100 under a pessimistic scenario of human activity [RCP6.0 (Moss et al., 2010; Intergovernmental Panel on Climate Change, 2014)]. Once tubes were filled with seawater, 50 larvae were haphazardly selected from the larval stock obtained by pooling larvae released that morning from maternal colonies, and added using a Pasteur pipette through the 6-mm hole in the cap of the tube. The hole was sealed when the tubes were stocked with larvae. Each tube was carefully inverted to gently mix the 50 larvae, so each trial began with the larvae scattered along the length of the tube. Although the pelagic larval duration (PLD) of P. damicornis can be >100 d (Richmond, 1987), their larvae commonly settle ≤ 24 h following release (Richmond, 1987; Isomura and Nishihira, 2001; Harii et al., 2002; but see Cumbo et al., 2012), and therefore, the present incubations were designed to last 12 h starting at 08:00 h.

After the tubes were stocked with larvae on the shore, they were transported by a snorkeler to an adjacent fringing reef and suspended vertically, with the midpoint of the tubes at a median depth (MD) of either ~0.3 m (July) or 3.3 m (August) (hereafter referred to as 0.3 and 3.3-m MD) (Figure 1). The median depth was recorded at the middle of the vertically oriented tubes, thus the depth range for the 0.3 m MD tubes extended from the surface to 0.7 m, and from 3 to 3.7 m for the 3.3 m MD tubes. To maintain this configuration, the tubes were attached to a weight at 5-m depth, and a float was used to keep them suspended vertically. While it is not possible to exclude an effect of time (July vs. August) in confounding the contrast of depth, experimental conditions were kept virtually identical between months to reduce temporal bias. Tubes were stocked with larvae shortly after sunrise, and were installed on the reef within 30 min of filling at ~07:30 h. Thereafter, the vertical position of the larvae was scored every 4 h starting at 08:00 h (initial) and finishing at ~19:30–20:00 h (sunset); the tubes were removed from the water at 08:00 h the following day and the larvae processed for lipid content (described below). At each census, the position of the larvae in the tubes was scored as the number of larvae in either the top or bottom sections of the tube. Tubes were not evaluated at night because of logistical constraints associated with nighttime snorkeling. The percentage of the total number of larvae present in the tubes at the time of each sampling was calculated for the top section of the tubes and used as the response variable to evaluate the effects of the treatments (depth and pCO₂).

High pCO₂ seawater was prepared in a single 10 L plastic reservoir by bubbling air or pure CO₂ gas into seawater using a mass-flow controller (HORIBASTEC, SEC-E40, Japan) that supplied gas at 16 mL min⁻¹. Adjustments of ± 0.1 mL min⁻¹ in the flow rate of CO₂ gas were made manually as necessary each morning leading up to the experiment to maintain the desired pCO₂, as evaluated from daily measurements of seawater pH and total alkalinity (A_T) used to calculate seawater pCO₂ using CO2SYS software (Lewis and Wallace, 1998). Seawater pH and temperature in the reservoir were measured daily between 09:00 and 11:00 h using a handheld meter (Multi 3410, WTW, Germany) fitted with a combination probe that recorded pH (±0.001 pH unit) and temperature (±0.1°C) (SenTix 940, WTW, Germany). The probe was calibrated daily prior to use with three NBS buffers (Nacalai Tesque, Japan). The salinity of the seawater used to fill the larval incubation tubes was measured using a conductivity meter (TetraCon 325, WTW, Germany), and A_T was determined using open-cell titrations conducted with an autoburette titrator (Kimoto, ATT-05, Japan). Titrations of certified reference materials (batch 155) provided by A.G. Dickson (Scripps Institute of Oceanography) yielded A_T values that differed on average ≤ 1.3 μmol kg⁻¹ from the certified value (SE = 9.2 μmol kg⁻¹, n = 6). Seawater pCO₂, HCO3-, CO32-, and Ω_arag were calculated from pH, temperature, A_T, and salinity using CO2SYS (Lewis and Wallace, 1998) with dissociation constants K1 and K2 from Mehrbach et al. (1973).

To test for changes in total lipid content in the larvae following incubations under ambient and high pCO₂, the initial lipid content of larvae was determined in three samples of 50 larvae, freshly released from the maternal colonies at 06:00 h and pooled among them. Each batch of larvae was placed in 1.5-mL vials for processing. The final lipid content of the larvae (i.e., 24 h after the start of incubations) was determined by processing batches of larvae in the same way following 24 h in ambient or high pCO₂ in either shallow (July) or deep (August) locations. Tubes incubated on the reef were returned to the lab 24 h after the start of each experiment (i.e., 08:00 h the following day), and all living larvae were collected by rinsing the contents of each tube into a collection container. Retrieved larvae were counted, and a single batch of 50 larvae from each tube at both depths was placed in a 1.5 mL vial for measurement of lipid content.

After transferring the batches of larvae to 1.5-mL vials, excess seawater was removed from the vials using a Pasteur pipette, and the larvae were frozen at −80°C for lipid analysis within 2–4 weeks. Following the protocol of Harii et al. (2010), modified by lengthening the extraction time in an ultrasonic bath to 15 min, lipids were extracted with 5 mL dichloromethane-methanol (6:4) at room temperature. Each batch of 50 larvae was extracted three times, with the solvent and eluted lipid pooled among extracts. Extracted lipids were concentrated in a rotary evaporator for 15–20 min, the residue was dissolved in dichloromethane 6 times, and then centrifuged to separate lipids from any remaining water. The solvent containing dissolved lipids was transferred to a new 50 mL flask after each separation. After the replicate dissolutions in dichloromethane, the samples were partially evaporated once again for ~5 min, and passed through a ~1 cm column of Na₂SO₄ crystals and plastic wool to remove any remaining water. Samples drained by gravity from the column into 4-mL pre-combusted glass vials (400°C for 4 h), in which they were then dried under nitrogen gas and weighed using a microbalance (±1 μg, MT5, Mettler-Toledo, USA). All solvents used were American Chemical Society (ACS) Guaranteed Reagent (GR) grade. Larval lipid content was expressed as μg larva⁻¹. The extraction efficiency of lipids was evaluated by comparing the lipid content of control larvae with the known lipid content of P. damicornis (Rivest and Hofmann, 2015).

Statistical analyses were conducted using SYSTAT Version 11 software (Systat Software, San Jose, CA). A two-factor Repeated Measures (RM) ANOVA was used to compare the effects of depth and pCO₂ on larval position in the tubes, with time of day as the RM factor, and arcsine-transformed values of the percentage of larvae found in the top of each tube as the dependent variable. Post-hoc analyses were conducted as paired t-tests for tubes at each depth and treatment combination, testing for differences between sequential time points using an α = 0.05/3 for three contrasts within each group. Differences in lipid content of larvae were evaluated using a two-sample t-test to compare the effect of pCO₂ on total lipid between depths, and a Kruskal-Wallis non-parametric test (due to violations of normality in the data) to compare lipid content after pCO₂ incubations regardless of month. Assumptions of normality and homogeneity of variance for the RM-ANOVA were assessed through graphical analyses of the residuals.

The pCO₂ in the tubes did not differ between the start and end of the 24 h incubations at both 0.3-m MD (t = 2.550, df = 5, p = 0.238) and 3.3-m MD (t = 0.007, df = 6, p = 0.995) (Table 1). Pooling pCO₂ treatment levels between depths, and including both initial and final (24 h later) measurements of seawater chemistry, mean treatment levels were 1,060 ± 31 μatm pCO₂ and 356 ± 13 μatm pCO₂ (n = 15, ± SE). Larvae incubated in high pCO₂ retained their shape and pale brown color at both depths, and visually were indistinguishable from larvae incubated under control conditions. One treatment incubation tube at 0.3-m MD broke prior to the larval census at 16:00 h, and exclusion of this tube from the statistical analysis provided too few observations to test all interactions in the model. To address this limitation, observations on larval distribution at 16:00 and 20:00 h for the damaged tube were replaced with an imputed mean value calculated from the remaining 3 tubes in the same treatment group, and the degrees of freedom were reduced by one for each replacement (Quinn and Keough, 2002; Zar, 2010).

The percentage of larvae found in the top section of each tube differed between pCO₂ treatments in a pattern that varied among times and depths [i.e., there was a significant interaction among depth, time, and treatment, F_{(3, 34)} = 3.762, p = 0.019, Table 2] (Figure 2). Post-hoc analyses were equivocal because of the small sample sizes, but sequential contrasts of times (three comparisons) suggest the interactive effect was caused by differences between 16:00 vs. 20:00 h in the high pCO₂ tubes at 0.3-m MD and ambient pCO₂ tubes at 3.3-m MD, between 08:00 vs. 12:00 h in the ambient pCO₂ tubes at 0.3-m MD, and between 08:00 vs. 12:00 h and 12:00 vs. 16:00 h in the treatment tubes at 3.3-m MD (all p < 0.05). At 0.3-m MD, the percentage of larvae in the top of the tubes in high pCO₂ described an inverse parabola as a function of time, increasing from ~42 to 64% between 8:00 and 16:00 h, but decreasing to 49% at 20:00 h (n = 43–50 larvae tube⁻¹). Larval behavior in high pCO₂ contrasted with that observed at ambient pCO₂, where at 0.3-m MD the percentage of larvae in the upper section of the tubes increased throughout the day, starting at 28% at 08:00 h and ending at 84% at 20:00 h. At 3.3-m MD in high pCO₂, 25–50% of the larvae were in the upper section of the tubes regardless of time. Larval behavior was similar at high and ambient pCO₂ at 3.3-m MD, where in ambient pCO₂ the percentage of larvae in the upper section of the tubes was 28 ± 4% between 8:00 and 20:00 h (mean ± SE, n = 16 based on 200 larvae at each time), although it decreased to 18% (n = 200 larvae) at 16:00 h. Overall, 66 ± 6% and 37 ± 4% of the larvae were found in the upper section of the tubes at 0.3-m MD and 3.3-m MD in high pCO₂, respectively (mean ± SE, n = 16), and in all cases the remaining larvae were swimming and behaving normally in the bottom sections of the tubes after 12 h.

The larvae used for lipid analysis were similar to one another in color and size (~1-mm long), and were visible in situ when freshly released from the maternal colonies, and following 24 h in the acrylic tubes. The analytical procedure for the lipid analyses required a minimum of 50 larvae for each determination, and this restriction excluded one tube from each pCO₂ treatment (high and ambient) in both July and August, when 10–15% of the larvae settled on the walls of the tube after 24 h. Mean dry lipid content at 0.3-m MD was 12.19 ± 0.23 μg lipid larvae⁻¹ after 24 h in ambient pCO₂, but this declined ~23% to 9.38 ± 0.79 μg lipid larvae⁻¹ after 24 h in elevated pCO₂ (both ± SE, n = 3). Similarly, at 3.3-m MD, mean dry lipid content was 11.96 ± 0.69 μg lipid larvae⁻¹ after 24 h in ambient pCO₂, declining ~25% to 8.99 ± 1.24 μg lipid larvae⁻¹ after 24 h in elevated pCO₂ (both ± SE, n = 3). Lipid content of freshly released larvae did not differ between July and August (t = 0.092, df = 4, p = 0.928), and averaged 11.98 ± 0.27 μg larvae⁻¹ (± SE, n = 6). As lipid content did not differ between months for larvae incubated in ambient pCO₂ (t = 0.248, df = 3, p = 0.820) or high pCO₂ (t = 0.267, df = 3, p = 0.803), the results of the lipid analyses following incubations were pooled between months and depths to test for an overall effect of high pCO₂. Lipid content of the larvae differed in high pCO₂ from ambient pCO₂ and initial release (Kruskal Wallis test, H = 9.710, n₁ = n₂ = n₃ = 6, p = 0.008) (Figure 3).

The second goal of the present study was to test the hypothesis that high pCO₂ affected the lipid content of P. damicornis larvae after 24 h exposure to high pCO₂ in situ. Exploration of this goal was motivated by the inference that a change in lipid content in response to high pCO₂ would affect larval buoyancy (sensu Harii et al., 2007). A leading determinant of buoyancy in invertebrate larvae is lipid content (Chia et al., 1984), and variation in lipid content can modulate their buoyancy, as in larvae of the abalone Haliotis spp. (Moran and Manahan, 2003) and a diversity of zooplankton (Lee et al., 2006). It is commonly accepted that brooded coral larvae are positively buoyant immediately following release (Oliver and Willis, 1987; Wood et al., 2016), and therefore quickly ascend to the surface of the seawater (Harii et al., 2002), where they are subject to rapid advection in wind-driven surface currents (Wolanski and Kingsford, 2014).

In the present study, the lipid content of P. damicornis larvae was measured at the start of the incubations and 24 h later. We infer that changes in lipid content occurred linearly over this period, which is consistent with trends observed in the larvae of other corals (Figueiredo et al., 2012) and invertebrates [e.g., abalone (Moran and Manahan, 2003) and sea urchins Sewell, 2005]. A linear decline in the lipid content of coral larvae could be a result of several non-exclusive mechanisms, including the catabolism of fatty acids through aerobic respiration (Harii et al., 2010), the incorporation of lipids into membrane phospholipids (Lee et al., 2006), or the loss of lipids in fat-rich mucus secretions (Gaither and Rowan, 2010; Ricardo et al., 2016). Distinguishing among these possibilities will require physiological analyses of different mechanisms by which lipid reserves are depleted by catabolism, and resolution of the lipids to their component classes (e.g., triacylglycerols and wax esters). The decline in lipid content of P. damicornis larvae that we recorded was detected based on initial and final sampling after 24 h in the treatment. Intermediate sampling that might have more completely resolved the rate of lipid decline was precluded by the challenges of working offshore at night. Nevertheless, the results show that larval lipid content declined after exposure to high pCO₂, but not after exposure to ambient pCO₂.

To our knowledge, the buoyancy of brooded coral larvae has been measured in only one study (Szmant and Meadows, 2006), likely because of the technical difficulty of separating buoyancy from swimming as mechanisms mediating larval motility in this taxon. These difficulties arise from the small size of most coral larvae, the difficulties of quantitatively sampling them in the plankton, and the challenges of quantifying the effects of ciliary swimming. However, larvae freshly released from P. damicornis in Mo'orea, French Polynesia, sank when they were anesthetized with eugenol, thus suggesting they were negatively buoyant (L. Bramanti, personal communication to JB), as has been described for the aposymbiotic larvae of the gorgonian Corallium rubrum (Martínez-Quintana et al., 2014). When interpreting larval movement in the present experiment, we therefore assume that larvae of P. damicornis are negatively buoyant.

Most brooded larvae of tropical scleractinians contain endosymbiotic Symbiodinium, which typically are vertically transmitted from the parent colony (Harrison and Wallace, 1990), and can translocate as much as 27% of their fixed carbon to the cnidarian tissue of the planula (Richmond, 1982). However, in addition to obtaining energy resources from their Symbiodinium, it is likely that brooded coral larvae also metabolize maternally-derived lipids to meet their energy needs (Harii et al., 2010). The use of lipid reserves for this purpose could be particularly important when the photosynthetic capacity of larval Symbiodinium is impaired by light or thermal stress (Haryanti et al., 2015), or in the context of our findings, by increased pCO₂, as has been seen in adult corals (Anthony et al., 2008). Understanding of the mechanisms of lipid catabolism is likely to be enhanced by more information on the lipid classes involved in the inferred catabolism. The lipids in P. damicornis larvae are composed of wax esters, triglycerides, and phospholipids, and as wax esters are typically the most abundant class (Harii et al., 2007; Figueiredo et al., 2012), it is reasonable to infer that lipids in the present larvae were also composed of mainly wax esters. The concentration of wax esters in the larvae of P. damicornis is particularly important to understanding their behavior, because wax esters have a lower density than triglycerides and phospholipids (Gurr et al., 2002; Lee et al., 2006). Wax esters are also positively buoyant in seawater (Arai et al., 1993), so they could modulate the buoyancy of larvae in which they occur.

Evidence for lipid catabolism as a mechanism to reduce the wax ester content of coral larvae comes from P. damicornis at Heron Island, Australia, where ≥85% of larval metabolic demand was derived from lipids under low light (~100 μmol m⁻² s⁻¹) conditions after 31 days in culture (Harii et al., 2010). Calculations corroborate this finding, because a larval respiration rate in P. damicornis of 0.15 nmol O₂ larvae⁻¹ min⁻¹ (Cumbo et al., 2013) would consume 2.42 μg lipid larvae⁻¹ day⁻¹, assuming lipid was the respiratory substrate and that it required 2 mL O₂ mg⁻¹ to be consumed in aerobic respiration (Schmidt-Nielsen, 1997). Over 24 h in the present experiment, larvae consumed ~2.8 μg lipid larvae⁻¹ in high pCO₂, so while the composition of lipids in P. damicornis larvae was not measured in the present analysis, it is feasible that the decline in lipid content included effects caused by catabolism of wax esters. A reduction of wax esters might be sufficient to reduce the buoyancy of the present larvae incubated at high pCO₂ and 0.3-m MD, therefore favoring downward movement in the tubes.

Elucidating the causes of the vertical movement recorded in the present analysis is challenging due to the likelihood that movement was caused by a combination of buoyancy and ciliary swimming. The inability to distinguish larval movement attributed to ciliary swimming from that attributed to passive buoyancy suggests that at least a portion of the movement recorded in the present study was a result of swimming. Utilization of ciliary swimming might explain the interactive effect of pCO₂ and depth on the position of larvae within the tubes, because the constancy of lipid content between depths at high pCO₂ suggests that changes in buoyancy were independent of depth in high pCO₂. According to this hypothesis, larvae might have swum more actively in high pCO₂. The downwards shift in vertical position of P. damicornis larvae in 3.3-m MD tubes (vs. 0.3-m MD) at high pCO₂ may reflect active swimming, as has been recorded for the larvae of many coral species (Vermeij et al., 2010; Martínez-Quintana et al., 2014), including P. damicornis (Te, 1992; Kuffner and Paul, 2004).

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.