Interactive Effects of Ocean Acidification, Elevated Temperature, and

Article pubs.acs.org/est

Interactive Effects of Ocean Acidification, Elevated Temperature, and Reduced Salinity on Early-Life Stages of the Pacific Oyster Ginger W. K. Ko,† R. Dineshram,† Camilla Campanati,† Vera B. S. Chan,† Jon Havenhand,‡ and Vengatesen Thiyagarajan*,† †

The Swire Institute of Marine Sciences and School of Biological Sciences, The University of Hong Kong, Hong Kong Special Administrative Region, China ‡ Department of Biological & Environmental Sciences, Tjärnö, University of Gothenburg, SE-452 96 Strömstad, Sweden S Supporting Information *

ABSTRACT: Ocean acidification (OA) effects on larvae are partially attributed for the rapidly declining oyster production in the Pacific Northwest region of the United States. This OA effect is a serious concern in SE Asia, which produces >80% of the world’s oysters. Because climate-related stressors rarely act alone, we need to consider OA effects on oysters in combination with warming and reduced salinity. Here, the interactive effects of these three climate-related stressors on the larval growth of the Pacific oyster, Crassostrea gigas, were examined. Larvae were cultured in combinations of temperature (24 and 30 °C), pH (8.1 and 7.4), and salinity (15 psu and 25 psu) for 58 days to the early juvenile stage. Decreased pH (pH 7.4), elevated temperature (30 °C), and reduced salinity (15 psu) significantly delayed pre- and post-settlement growth. Elevated temperature lowered the larval lipid index, a proxy for physiological quality, and negated the negative effects of decreased pH on attachment and metamorphosis only in a salinity of 25 psu. The negative effects of multiple stressors on larval metamorphosis were not due to reduced size or depleted lipid reserves at the time of metamorphosis. Our results supported the hypothesis that the C. gigas larvae are vulnerable to the interactions of OA with reduced salinity and warming in Yellow Sea coastal waters now and in the future.

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INTRODUCTION

The coastal and surface oceans have been forecast to become increasingly unsuitable for calcifying marine invertebrates because of rising anthropogenic CO2 levels.9 Increasing pCO2 reduces seawater pH, carbonate ion concentration, and the saturation states of calcium carbonate minerals, which are collectively termed “ocean acidification”, hereafter referred to as “decreased pH”.10 Simultaneously, climate change induced heavy precipitation in some parts of the world is expected to reduce surface salinity.11 Both reduced salinity and warming of seawater are known to be the important physical stressors that may challenge the fitness of marine animals.9 The negative impact on the carbonate system by these environmental factors in conjunction with decreased pH has been seldom studied. Moreover, our understanding of the impact of climate change is still mainly derived from experiments that analyzed only single stressors12,13 and usually with short exposure periods.14−18 However, climate change stressors do not act in isolation and usually interact together to potentially cause detrimental effects, in particular on the susceptible early life stages of marine

Oysters are ecologically and economically important shellfish that have a complex life cycle, consisting of multiple larval stages adapted for rapid growth, feeding, and dispersal. After a period of pelagic life, the larvae enter a competent state for metamorphosis before transforming into benthic juveniles in just a few minutes.1,2 Metamorphosis is a crucial step in the life cycle of most marine benthic invertebrates, involving the recognition of environmental signals, settlement on a suitable substrate, and subsequent tissue remodeling.3 In addition to these complex and energetically expensive processes, the oyster larvae also actively alter their shell mineral composition from aragonite to calcite.4 The intricate processes involved in metamorphosis could be a bottleneck that determines the success of these marine invertebrates, particularly in relation to the impact of climate-related variables. How this crucial transitional stage in the life history of oysters is affected by external factors is still poorly understood.5 The majority of past studies have focused on the early larval forms or the adult stages. The inherent relationship between these life stages through settlement and metamorphosis means that we need to consider the bigger picture in relation to climate change effects.6−8 © 2014 American Chemical Society

Received: Revised: Accepted: Published: 10079

April 3, 2014 July 9, 2014 July 10, 2014 July 11, 2014 dx.doi.org/10.1021/es501611u | Environ. Sci. Technol. 2014, 48, 10079−10088

Environmental Science & Technology

Article

organisms.19,20 For example, decreased pH combined with suboptimal elevated temperature have been reported to have negative effects in larvae of C. gigas resulting in dramatically reduced fertilization success, slower larval development, and abnormal larval morphology.21 However, these effects may be population dependent.18,22 Oysters metamorphosed under OA conditions also showed carry-over effects; individuals grown and metamorphosed at pH 7.8 showed delayed and suppressed growth as juveniles.5 Whether combined climate stressors interact and exacerbate the effects of decreased pH in any particular ecosystem and population is still unknown.23 This is especially important in China’s Yellow Sea area, which is the center for C. gigas production, and China in general as it accounts for >80% of the world’s shellfish production.24 The success of shellfish aquaculture is greatly determined by the performance of early life stages, such as the embryo and early larval stages.25 However, the early larval performance as observed in a short experimental period may not be indicative of a similar performance in later larval stages. A longer duration of exposure to stressors may have a profound impact on the success of metamorphosis, leading to a greater vulnerability or resistance to stressors after metamorphosis.26,27 The performance of the competent larvae, the completion of metamorphosis, and the fitness of the subsequent juveniles, therefore, would provide a more ecologically realistic picture of the success of an organism. Information such as metamorphosis and juvenile fitness are also critical for estimating the productivity of oysters in hatchery and off-shore aquaculture farms.28 Given the many stressors associated with climate change, there is an urgent need to develop a better understanding of the effects of these stressors on the different larval stages, metamorphosis, and postmetamorphic fitness through long-term acclimation studies.23 Knowledge of the adaptive capacity of larvae to multiple stressors will not only enhance our ability to predict the consequences of warming, reduced salinity, and ocean acidification but also assist the development of protective measures to mitigate the impacts of climate on marine shellfish resources.8,29 In this study, we investigated the effects of decreased pH together with elevated temperature and reduced salinity of seawater on the C. gigas populations in the oyster culture zones of the Yellow Sea. We examined how decreased pH, elevated temperature, and reduced salinity affect pelagic larval growth rates and how shell size and energy reserves at the time of settlement affect the metamorphosis success and postsettlement growth. Our study provided a comprehensive and integrated view of the biological responses to future warming, acidification, and reduced salinity during the early ontogeny and the life stage transitions of the commercially important Pacific oyster, C. gigas, in the Yellow Sea.

conditions represented the general Pacific oyster larvae culture conditions in northern Chinese aquaculture areas and were subsequently used as the ambient (or control) conditions in this study. The eggs from C. gigas were fertilized by pooling gametes from multiple parents (8 males and 23 females) in order to minimize sperm-egg incompatibility and to reduce the influence of genetic effects from an individual.31 Gametes were separated by “strip-spawning” and suspended in 0.22 μm filtered seawater (FSW) in ambient conditions with a salinity of 25 psu and pH 8.1 at 24 °C. About five million eggs were fertilized by mixing with 20 mL of concentrated sperms.32 Embryos were isolated from the sperms by filtration in a 20 μm nylon mesh and were cultured in FSW in 50 L plastic tanks in the above ambient conditions.22 The majority of the embryos hatched into the trochophore larval stage under these optimal culture conditions. Early stage larvae with an overall homogeneous size (diameter, ∼50 μm) were separated from undeveloped embryos and smaller sized larvae by washing in a 35 μm nylon mesh for use in the following experiment. Experimental Design for Multiple Stressors. A three factor orthogonal experimental design (2 × 2 × 2) was used to examine the interactive effect of pH, salinity, and temperature on the development of pre- and post-settlement life stages of the C. gigas. There were two levels for pH (pH 8.1 and pH 7.4), two levels for salinity (25 psu and 15 psu), and two levels for temperature (24 and 30 °C). The eight treatments were as follows: control (Control, C: 24 °C, 25 psu, pH 8.1), three single stressor treatments (reduced salinity, S: 15 psu; decreased pH, P: pH 7.4; elevated temperature, T: 30 °C), three dual stressor treatments (SP: 15 psu, pH 7.4; TS: 30 °C, 15 psu; TP: 30 °C, pH 7.4), and one triple stressor treatment (TSP: 30 °C, 15 psu, pH 7.4). Each treatment included four replicate cultures. These conditions represent seawater chemistry conditions experienced by the C. gigas due to seasonal fluctuations in a typical aquaculture area in the Yellow Sea. The seasonal fluctuations in salinity, temperature, and algal production can cause significant fluctuations in the surface seawater pH in the Yellow Sea area ranging from pH 7.67 to 7.92 in winter months and on average pH 8.06 in summer months (June to August).33,34 Rising anthropogenic CO2 can partially account for the declining pH in the Yellow Sea.33 Similarly, surface seawater temperatures can range from 5 °C in winter to 25 °C in summer. The semienclosed Dayanowan Bay, which is located in the north of the Yellow Sea and serves as an intensive culture zone of Pacific oysters, experiences highly variable seawater salinity and pH levels on a daily and monthly basis due to tidal and terrestrial influences.35 The experimental treatment ranges also cover the expected average pH, temperature, and salinity in near-future climate scenarios due to rising anthropogenic CO2, global warming, or climate change driven heavy precipitation in the Yellow Sea coastal areas.34 The decreased pH treatment (pH 7.4) was maintained by continuously bubbling CO2 enriched air directly into the culture tanks.22 The concentration of CO2 in the air was regulated through dual variable-area flow controllers with a flowmeter (Cole-Parmer Inc., U.S.A.). The ambient FSW salinity was diluted from 30 psu using freshwater to 25 psu (ambient) and 15 psu (reduced salinity). The water temperature was maintained using a circulating water bath heated with temperature control. The pH and temperature of the cultures were monitored two or three times per day using a pH analyzer (Orion Star, Thermo Electron Co., U.S.A.) with an Orion

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MATERIALS AND METHODS Collection, Maintenance, and Fertilization of the Study Organism. Sexually matured Pacific oysters Crassostrea gigas (shell length, 10−15 cm) were collected during June 2012 from the oyster aquaculture areas of Tsingdao (China) in the Yellow Sea (36° 04′N, 120° 22′E). The collection site had an average summer climate with a mean summer water temperature of about 23 °C, a salinity of 25 to 30 psu, and pH 8.2.30 The animals were transported in dry conditions to the Swire Institute of Marine Science, Hong Kong, and were acclimatized in a running seawater aquarium at pH 8.1, salinity of 25 psu, and temperature of 24 °C for 2 days. The acclimatization 10080

dx.doi.org/10.1021/es501611u | Environ. Sci. Technol. 2014, 48, 10079−10088


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replicate. The pre-settlement growth rate for the early larval period (1−10 days), late larval period (10−15 days), and the overall larval period (1−15 days) was calculated.22 Pediveligers were sieved using a 280 μm mesh after 15 days for the elevated temperature treatments or 18 days for the ambient temperature treatments. Irrespective of treatments, the pediveligers had a shell length between 280 to 350 μm. Similarly sized pediveligers of a similar physiological age allowed us to examine the effects of multiple stressors in terms of the larval metamorphosis rate and the lipid index independently from the virtual age (no delayed or enhanced growth).41 The extent of metamorphosis, lipid index, and shell mechanical properties were measured in these pediveliger larvae. However, these were the fastest growing larvae from each treatment group, which may not fully represent the entire larval population. Larval Metamorphosis. A portion of randomly selected pediveligers was transferred into a 1 L plastic bioassay vessel containing about 750 mL of FSW with a plastic sheet covered with 7-day-old natural biofilm to induce larval settlement and metamorphosis.42 The percentage of settled and metamorphosed individuals on each biofilm surface and tank wall was counted after 48 h incubation. Four replicate tanks were used for each treatment, but only two replicate cultures were useable for TP and TSP treatments. The pH, salinity, and temperature of each bioassay vessels were preadjusted and maintained during the whole assay period according to the larval treatment conditions. Larval Lipid Index. To determine the larval physiological condition, a portion of pediveliger larvae from each treatment was fixed to assess the lipid index using Nile red staining (Sigma-Aldrich, U.S.) to measure the lipid energy reserves,43 which are visible as stained intracellular lipid droplets. Between five and 14 pediveliger larvae per replicate tank were stained with 1.25 μg/mL of Nile Red dissolved in acetone for 90 min.44,45 During the staining period, respective larval treatment conditions were maintained. After staining, larvae were rinsed with FSW and fixed in 95% formaldehyde. Lipid reserves in stained larvae were visualized using a fluorescent microscope (Nikon 80i, U.S.), and photographs were taken within 4 h before the fluorescence faded. The images were converted to gray scale, and the lipid area of 10−12 larvae per sample was measured using the ImageJ software (ImageJ 1.45s, NIH, U.S.). The lipid index was determined as the ratio of the lipid area to the total surface area of the larva. Post-settlement Growth Rate. The juvenile growth rate was determined using the samples from the previous settlement bioassay. Again, the same original treatment conditions were used to culture the juveniles. Randomly chosen post-settlement individuals attached on the plastic sheets in each treatment tank were marked with permanent ink to identify them for repeat measurements. Crowded or overlapping individuals were not used. The shell growth and survival rate of juveniles were monitored after 43 days post-settlement. The growth rate was monitored for one to eight juveniles per replicate tank. There were 4 replicates per treatment except for TP and TSP, which had only 2 replicate tanks. The area of each juvenile’s right valve (outer shell) was determined using the Leica Q Win V3 software. Juvenile growth rate was calculated as the difference (growth increment) between the initial and the final shell area divided by number of days exposed to treatment conditions. Statistical Analysis. The effects of the three treatments (pH, salinity, and temperature) and their interactions on larval

8102BN Ross combination electrode (Thermo20 Electron Co., U.S.A.). The salinity and total dissolved oxygen levels were measured periodically with a refractometer and portable dissolved oxygen meter (ORI-1212503Orion 3-Star, Thermo Fisher Scientific Inc., U.S.), respectively. The algal food concentration was determined using a hemocytometer. Total alkalinity (TA) was measured using an Alkalinity Titrator (ACA2, Apollo SciTech’s Inc., U.S.) using 50 mL water samples free of larvae, poisoned with 10 μL of 250 mM mercuric chloride. The TA measurement was standardized with a certified seawater reference material (Batch 106, A.G. Dickson, Scripps Institution of Oceanography, U.S.). The carbonate system parameters were calculated using the CO2SYS software program with equilibrium constants K1, K2, and KSO4.36 Pre- and Post-settlement Cultures. A batch of trochophore larvae collected from ambient conditions was raised to early juvenile stage (post-settlement stage) in the eight different experimental conditions (8 treatments × 4 replicates). The plastic culture tanks contained about 10 larvae mL−1 in 40 L of UV-treated FSW. The culture tanks for the ambient and elevated temperature treatments were positioned in separate water baths. The respective temperature tanks were randomly assigned to the pH and salinity treatments. All the tanks were covered with tightly fitting plastic lids. Depending on the treatment, ambient air or CO2 enriched air was bubbled gently through an air stone at the bottom of the tank. Besides maintaining the corresponding treatment pH, the bubbles helped to circulate the larvae and the algal food.22 During the first 7 days of culture, larvae were fed with live Isochrysis galbana (5 × 105 cells mL−1) once a day.37 After 7 days, both larvae and juveniles were fed with a mixture of live I. galbana and Chaetoceros gracilis (10 × 105 cells mL−1, 1:1 ratio) twice a day.38 Metamorphosis was induced on plastic plates coated with 7-day-old natural biofilm developed in an open water circulating system at ambient conditions. The biofilm coated plates were immersed in larval culture tanks for inducing metamorphosis. Metamorphosed juveniles that had settled on the plates were kept in 1 L tanks for post-settlement growth with the same original treatment conditions as the larvae cultures. Juveniles were transferred to new culture medium every 4 day. Samples were periodically taken from the larva and juvenile cultures for measurements. Pre-settlement Growth Rate. The effects of the different stressors on larval survival rate, larval growth rate, and relative proportions of the different developmental stages were determined on days 1, 10, and 15 post-fertilization. The samples were filtered using a 50 μm mesh and concentrated into 200 to 250 mL FSW seawater of respective treatment. A subsample (1 mL) was taken from each replicate of the thoroughly mixed concentrated larvae. Samples were immediately fixed in 10% buffered formalin. To calculate the survival rate, the number of empty shells without tissue inside (dead at the time of fixation) and shells with larval tissue inside (alive at the time of fixation) were counted under a compound microscope equipped with a digital camera (Leica DFC 280, Leica, Germany). Larvae were also classified according to the different developmental stages. The developmental stages were identified by the shell morphology and the presence of Dshaped larva, early umbo, late umbo, and pediveliger stages with eyespots.39 All larval shells were then photographed at ×63 magnification. Shell areas of the right valve from each picture were measured to determine the growth using the Leica QWin V3 software.40 About 15 to 61 larvae were measured for each 10081



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Table 1. Representative Seawater Physiochemical Conditions Used To Study the Effect of Decreased pH (P), Reduced Salinity (S), Elevated Temperature (T), and Their Interaction on Pre- and Post-settlement Growth of Crassostrea gigasa measured condition pre-settlement C S P SP T TS TP TSP post-settlement C S P SP T TS TP TSP

pHNBS

temp (°C)

calculated

salinity (psu)

TA (μmol kg−1)

pCO2 (μatm)

CO32−(μmol kg−1)

ΩCal

ΩAr

8.04 7.97 7.47 7.37 8.05 8.02 7.63 7.48

± ± ± ± ± ± ± ±

0.01 0.01 0.06 0.04 0.12 0.02 0.03 0.03

22.0 22.1 22.1 21.9 30.1 30.2 30.0 29.4

± ± ± ± ± ± ± ±

0.1 0.2 0.1 0.1 0.1 0.0 0.1 0.6

25.0 18.4 24.5 18.9 24.7 15.0 25.0 14.5

± ± ± ± ± ± ± ±

0.0 0.9 0.6 0.3 0.5 0.0 0.0 0.7

1830 1433 1837 1577 1827 1383 1848 1706

± ± ± ± ± ± ± ±

64 43 40 268 19 83 47 36

519 534 2166 2532 532 517 1566 2396

± ± ± ± ± ± ± ±

35 37 385 602 31 28 113 95

95 53 28 16 120 65 52 24

± ± ± ± ± ± ± ±

1.8 1.5 4.6 1.9 5.2 4.4 4.6 1.0

2.45 ± 0.05 1.43 ± 0.05 0.73 ± 0.12 0.43 ± 0.05 3.19 ± 0.13 1.90 ± 0.13 1.37 ± 0.12 0.71 ± 0.03

1.55 ± 0.03 0.88 ± 0.03 0.46 ± 0.07 0.26 ± 0.03 2.06 ± 0.09 1.16 ± 0.08 0.89 ± 0.08 0.43 ± 0.02

8.07 8.03 7.48 7.40 8.13 8.05 7.52 7.47

± ± ± ± ± ± ± ±

0.00 0.00 0.00 0.01 0.00 0.00 0.00 0.01

23.9 23.8 24.0 23.9 29.9 29.7 29.8 30.0

± ± ± ± ± ± ± ±

0.0 0.0 0.0 0.0 0.1 0.1 0.1 0.1

24.0 15.0 24.0 15.0 24.0 15.0 24.0 15.0

± ± ± ± ± ± ± ±

0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0

1608 1085 1584 1218 1664 1122 1865 1161

± ± ± ± ± ± ± ±

25 13 33 254 283 72 292 0.49

437 372 1848 1971 396 388 2024 1618

± ± ± ± ± ± ± ±

6 6 11 475 73 27 316 69

90 44 26 12 123 55 39 17

± ± ± ± ± ± ± ±

2.2 0.4 1.1 2.2 21.1 3.3 6.0 0.5

2.35 ± 0.06 1.23 ± 0.01 0.67 ± 0.03 0.34 ± 0.06 3.30 ± 0.87 1.60 ± 0.30 1.05 ± 0.34 0.49 ± 0.02

1.49 ± 0.04 0.74 ± 0.01 0.42 ± 0.02 0.21 ± 0.04 2.12 ± 0.36 0.97 ± 0.06 0.68 ± 0.10 0.30 ± 0.01

Measurements shown in this table (mean ± S.D, n = 4) were taken from larval culture tanks at the time of sample collection for larval shell size analysis on 15 day post-fertilization. The undersaturated states of calcite or aragonite are indicated in bold. Abbreviations: partial pressure of carbon dioxide (pCO2), carbonate ion concentration (CO32−), aragonite saturation state (ΩAr) and calcite saturation state (ΩCal), and total alkalinity (TA). The treatment abbreviations are explained in Figure 1. a

control (C) or salinity (S) treatments (Table S1). The pH was reduced to pH 7.37 in reduced salinity (SP) of 15 psu. All four decreased pH treatments (P, TP, SP, and TSP) led to aragonite undersaturation (ΩA < 1). Reduced salinity caused ΩA to decrease from 1.55 to 0.88 at 24 °C and from 2.06 to 1.16 at 30 °C (Table 1), but the pH or the pCO2 level was not altered in either temperature. Reduced salinity in combination with the decreased pH treatment reduced ΩA further to the lowest value of the eight treatments (0.26, Table 1). At elevated temperatures (30 °C), the pCO2, carbonate ion concentration, and the ΩA were all lower than at ambient temperatures (24 °C). The dissolved oxygen levels were similar in all eight treatments, ranging from 5.2 to 5.8 mg L−1. Pre-settlement Growth. Larval growth response to stressors differed between early (1 to 10 days) and late (10 to 15 days) growth periods (Figure 1A). Early larval growth was generally faster in elevated temperature (T, TS, TP, and TSP) compared to the ambient (C, S, P, and SP). Early growth rate was reduced in reduced salinity, but no significant interactions were seen among the three stressors (Figure 1A; Table S2). In contrast, late larval growth (10 to 15 days) was generally slower in decreased pH conditions (P, SP, and TSP), and this response was more apparent in combinations of stressors (TP, TS, and TSP) (Tables 2 and S2). The negative effect of decreased pH (C > P; S > SP) on the growth rate at 24 °C was offset by elevated temperature at 30 °C (T = TP), but this mediating effect failed in the presence of reduced salinity (TS > TSP) (Figure 1A). We did not observe any statistically significant three-way interaction effects on overall growth rate for 1 to 15 days (Table 2), but larval growth was slower in decreased pH (P and SP) and faster in elevated temperatures (Figures 1A and B). Interestingly, the reduction in growth rate we observed at reduced salinity at 30 °C (T > TS, TP > TSP) did not occur at 24 °C (C = S, P = SP). The combination of the three stressors (TSP) negated the increases in growth rate caused by elevated

growth rates, metamorphosis, lipid reserves, and juvenile growth rate were examined using three-way ANOVA. Data were first transformed to improve normality and equality of variance. Data that still did not conform with the assumption of homoscedasticity were ranked-transformed before performing three-way ANOVA. Due to high variability and unequal sample sizes within treatments, the statistical power of some of these tests was low.46 Under these circumstances, Student’s t-test was used a priori to compare treatment effects of interest against the control. Bonferroni correction, normally applied to adjust the P-values for multiple comparisons, can increase type II errors, and therefore this was not used.46 Nevertheless, we interpreted our results from these tests with caution based on the magnitude of the differences between the treatment and the control or between two treatments of interest (see below). Importantly in this regard, ocean acidification research has recently emphasized understanding the magnitude of a given effect, especially with an effect size of zero or “no response”, rather than reliance on null-hypothesis testing.47 Particularly for our data set, measurements of a standardized effect size with confidence intervals (CI) are more informative than traditional ANOVA.48 Therefore, we studied the effects of different treatments on larval response variables by plotting the mean log response ratio (LnRR = loge [response in treatment]/loge [response in control]) ± 95% CI.49 This measurement is statistically robust, easy to interpret, and visualizes clearly the treatment effect.25

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RESULTS Carbonate Chemistry. Throughout the experiments, the observed pH values did not deviate substantively from expected levels in any of the treatments. The replicate variability was low in all of the treatments (Figure S1 and Table 1). In the four treatments at ambient temperature (24 °C), the mean pH under decreased pH (P) was pH 7.47 which was lower than the 10082



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temperature, leading to growth rates that were similar to those observed in the control (Figure 1B). In parallel with the increasing growth rate, warmer temperatures also increased larval development rate. In the elevated temperature, between 7% and 40% of larvae reached competence (pediveliger) by day 15 compared to 90% of these larvae were in late umbo stage, Figure S2). The interaction of the three stressors (TSP) appeared to result in a wider spread of developmental stages within the population, which contained more early umbo (25%), considerably more late umbo (44%), and some pediveliger (∼7%) stages. This was in contrast to the other elevated temperature treatments, which had greater degree of later developmental stages (2%−7% early umbo, 55%−63% late umbo, and 30%−40% pediveliger). Although elevated temperature treatments slightly reduced the survival rate, the effect was not statistically significant (C: 99.80 ± 0.20% and TSP: 78.48 ± 13.81%) (Tables 2 and S3). Larval Metamorphosis. After 15 days, about 20% to 40% of the larvae in the elevated temperature treatment conditions (T, TS, TP, and TSP) had developed to the pediveliger stage with an eyespot and a foot, indicating that they were competent to attach and metamorphose into spat. However, at ambient (control) temperature (C, S, P, and SP), the majority of larvae reached the pediveliger stage only after 18 days of development. Nevertheless, pediveliger larvae used in the bioassay had similar shell sizes. Few pediveligers metamorphosed successfully in decreased pH (P, SP, TP, and TSP) (Table S4; Figures 2A and B). In contrast, larvae cultured in the warmer treatments had greatly increased metamorphosis success (T, TS > C) (Figure 2A). Although there was no significant effects of TSP in the three-way ANOVA due to the small number of replicates (n = 2) (Tables 2 and S4), only few larvae (33%) metamorphosed when exposed to three interacting stressors in TSP compared to the control (TSP: t(4) = 2.924, p < 0.05; Figure 2B). Larval Physiological Condition (Lipid Index). Larval samples were categorized into high, medium, and low lipid reserve groups based on the lipid index measurements (Figure 3A). Mean values of the lipid index varied in a complex manner

Figure 1. (A) Effects of decreased pH, reduced salinity, elevated temperature, and their combinations on larval growth rates of Crassostrea gigas during 1−10 days (white bars), 10−15 days (gray bars), and 1−15 days (black bars) post-fertilization. Each bar represents the mean ± SE of four replicates. (B) Mean logarithmic response ratios (95% CI) of the effects of three stressors (P, S, and T) and their combinations on the overall growth rate of C. gigas larvae (n = 4; during 1−15 days post-fertilization). C = control; T = elevated temperature; S = reduced salinity; P = decreased pH; TS = elevated temperature and reduced salinity; TP = elevated temperature and decreased pH; SP = reduced salinity and decreased pH; TSP = elevated temperature, reduced salinity, and decreased pH.

Table 2. Summary Results of 3-Way ANOVA Showing the F-Ratios for the Main and Interactive Effects of pH (8.1 and 7.4), Salinity (25 and 15 psu), and Temperature (24 and 30 °C) on Proportion of Different Developmental Stages; Pre-settlement Growth Rate; Larval Physiological Condition; and Post-settlement Growth Rate of the Crassostrea gigasa condition pre-settlement larval development day 15 developmental stage D-shaped early umbo late umbo pediveliger survival rate growth rate overall day 1 to 10 day 10 to 15 larval physiological condition lipid index post-settlement development larval metamorphosis juvenile growth rate a

pH (P)

salinity (S)

temp (T)

34.404 10.274 3.964 1.590 1.351

5.955 19.345 0.025 7.237 2.746

34.303 9.415 103.915 96.742 5.614

12.397 0.180 13.178

15.603 8.420 0.0294

30.954 31.447 1.847

0.557

1.458

4.681

27.045 20.627

0.294 4.046

1.027 9.699

P×S

P×T

S×T

S×P×T

34.303 5.641 0.006 2.827 0.602

5.955 20.475 0.101 5.152 2.747

5.955 7.687 6.095 1.809 0.629

0.455 1.766 5.645

10.030 0.127 8.201

0.803 3.163 5.555

0.146

0.824

2.069

0.014

0.0725 11.934

2.946 17.066

4.200 5.776

0.976 8.221

5.955 8.446 0.311 8.657 0.217 1.460 1.840 × 10 1.504

−3

Significant effects (p < 0.05) are indicated in bold. Detailed information of ANOVA results are provided in Supporting Information Tables 1−4. 10083



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Figure 2. (A) Effects of decreased pH, reduced salinity, elevated temperature, and their combinations on metamorphosis of Crassostrea gigas as a percentage. Each data point represents the mean ± SE of four replicate measurements, but TP and TSP treatments had only two replicates. (B) Mean logarithmic response ratios (95% CI) of the effects of three stressors (P, S, and T) and their combinations on the metamorphosis of C. gigas larvae. See Figure 1 for abbreviations.

Figure 3. (A) Florescence microscopy images of Crassostrea gigas pediveliger larvae stained with Nile red for 1.5 h to detect lipid content. Images show green areas containing lipid droplets under UVlight. Larvae were qualitatively grouped or classified into high, medium, or low lipid index groups according to stain intensity. Scale bar = 280 μm, magnification was the same for all images. (B) Effects of decreased pH, reduced salinity, elevated temperature, and their combinations on the mean lipid index of C. gigas larvae at the time of settlement and metamorphosis (15 days post-fertilization in elevated temperature treatments and 18 days post-fertilization in ambient temperature treatments). Black bars represent a high lipid index, gray bars represent a medium lipid index, and white bars represent a low lipid index. Each data point represents the mean ± SE of four replicate measurements, but TP and TSP treatments had only two replicates. (C) Mean logarithmic response ratios (95% CI) of the effects of three stressors (P, S, and T) and their combinations on the lipid index of C. gigas larvae. See Figure 1 for abbreviations.

(C=S=TS > P=SP=TSP > TP=T; Figure 3B). 95% confidence intervals around mean log response ratios (LnRR) indicated no marked effect of salinity (S) on the lipid index but clear negative effects in most decreased pH treatments (P, SP, TP, and TSP) (Figure 3C). Increased temperature dramatically reduced (>40%) the lipid index in larvae (T, TP, and TSP) (Tables 2 and S4). Noticeably, the lipid index in larvae cultured in the TSP group was slightly higher than in the T treatment group and was equivalent to the controls (Figure 3C). Post-settlement Growth. Juvenile shell growth rate was significantly and negatively influenced by the three stressors in combination (TSP treatments) (Tables 2 and S4; Figure 4A). Individuals in the TSP treatment appeared to experience negative growth due to dissolution at day 17 post-settlement. Juveniles typically grew faster at decreased pH (Table S4), leading to an almost 10 times faster growth rate (in P and SP) at ambient temperature. At elevated temperature and decreased pH, there was also an increase in growth rate but to a lesser degree. No increased growth occurred in the combined elevated temperature and reduced salinity treatment (TS) (Figure 4B).

stressors (TSP) delayed the larval growth rate at a crucial period just before settlement and metamorphosis (i.e., 10 to 15 days post-fertilization) and negated the positive effect of elevated temperature on the larval growth rate. These findings are interesting because there is a distinct lack of studies on the effects of TSP on bivalve species in the literature. Due to practical limitations, we did not examine the impact of TSP on very early life stages of fertilization and the embryo. Our results may have underestimated the impact of TSP on the entire life cycle of oysters, because the two highly susceptible early life stages show decreased developmental growth or enhanced mortality in decreased pH as reported in previous studies.50 Although the larvae were fed ad libitum in our study, food limitation in nature may exacerbate some of the observed effects.51 Similarly, pooling of gametes from several individuals

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DISCUSSION This multiple stressors study involving multiple end-points and life stages tested the hypothesis that combined elevated temperature, reduced salinity, and decreased pH (TSP) would reduce growth, metamorphosis, energy reserves at settlement, and also post-settlement growth rate. Our findings supported only the first hypothesis. Multiple interacting 10084



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effects of reduced salinity and decreased pH (Figure 1B). At elevated temperature, warming results in an increased carbonate saturation state (Table 1) that may have driven the required mechanisms for maintaining the cellular environment for somatic growth and calcification, and larvae generally tend to have an overall increased metabolic rate offsetting the negative effects of decreased pH.54,55 Similar benefits of ocean warming have been demonstrated in calcifiers coping with decreased pH, such as certain coral species.56 On the contrary, the TP combination can negatively affect larvae in several species of sea urchin57 and delay larval development in bivalves.21,44 Thus, the effects of TP interaction may be developmental stage, species, and treatment specific.58 Although elevated temperature appeared to reverse the effects of pH and salinity stressors, the ability of larvae to speed up their growth rate may only be possible at the expense of their stored energy. The food source was replenished once or twice a day, so the observed effect on developmental growth was not a result of food availability. The decreased pH level in combination with reduced salinity and elevated temperature may deplete carbonate ions in the seawater, resulting in an environment prone to dissolution of carbonate structures. Therefore, more energy would be required to overcome the osmotic stress, acid−base regulation, and an elevated internal pH to maintain homeostasis for shell building at the calcification site.59 The reduced salinity can cause further osmotic challenge and elevate basal energy demand in a decreased pH environment, leading to lower energy levels available for growth or shell formation in bivalves.59 In addition, many marine invertebrates expend energy by expressing heat shock proteins in response to heat stress. When all three stressors are acting together, oyster larvae may exhaust their energy reserves and have less available energy to allocate for shell growth. Interestingly, early larval forms were robust to climate stressors than those of late larval forms. One possible explanation could be that early larval forms (e.g., veliger) are less dependent on feeding and associated behavior for energy resources than those of late larval stages (e.g., pediveliger). Additionally, early stages are functionally less diverse than those of pediveliger; the latter has to find suitable habitat and prepared to metamorphose. Nevertheless, physiological mechanisms that underlie/mediate the observed interactive effect of TSP are far from being fully understood. Metamorphosis of Pediveliger. Generally, the metamorphosis success of the C. gigas was substantially reduced at decreased pH regardless of elevated temperature or reduced salinity (Figure 2B). pH appears to play a key role in the larval metamorphosis success of C. gigas. Although elevated temperature increased metamorphosis, it could not counteract the negative effect of decreased pH. Although the statistical analysis did not detect a significant effect on metamorphosis for the TSP treatment, the effect of TSP is biologically important because 0.05). Similarly, there was no relationship between larval lipid store and environmental pH levels in this study (r2 = 0.10, p > 0.05). Thus, the observed treatmentspecific effects on metamorphosis success (Figure 2A) cannot be fully explained by lipid or energy reserves in larvae at the time of settlement (Figure 3B). Similarly, this study did not detect any relationship between pediveliger larval shell sizes and percentage of metamorphosis. The faster growing larval population that reached pediveliger stage on day 15 (at 30 °C) or day 18 (at 24 °C) post-fertilization had similar shell sizes. Therefore, our results cannot be ascribed to bias or confounding effects of larval size and developmental stage at the time of settlement. Our results from the shell analysis or lipid index measurements cannot fully explain why C. gigas larval metamorphosis was reduced in decreased pH treatments. We suspect that a complex interaction of exogenous and endogenous factors disrupts the metamorphosis of C. gigas under decreased pH, especially with TSP. Post-settlement Growth. It is widely accepted that postsettlement juvenile growth in a wide range of marine invertebrates is not independent of the pre-settlement period, rather it is strongly dependent on pre-settlement growth history, especially the level of energy stored in larva at the time of settlement.2,26 A few studies have shown that larvae exposed to decreased pH or reduced salinity or elevated temperature conditions resulted in decreased post-settlement growth in corals and barnacles.60,61 Such carry-over effects of delayed larval growth and decreased metamorphosis under ocean acidification on post-settlement growth was recently reported in Olympia oyster (Ostrea lurida) juvenile.5 In contrast to the main consensus, the post-settlement growth rate of C. gigas was significantly increased in decreased pH (Figure 4). Similar results have been observed with juvenile mussels.62 However, juvenile growth rate was dramatically decreased with the three stressors in TSP (Table 2 and Figures 4A and B). In agreement with our findings, the calcification of the C. virginica declined markedly at decreased pH 7.4 only with elevated temperature or reduced salinity.63 In our study, the lower growth rates caused by TSP or enhanced growth rates at decreased pH cannot be solely explained by the level of the energy stores at the time of settlement. Similarly, differences in juvenile growth rates cannot be explained by differences in size at the time of settlement. Juvenile shell dissolution could have contributed to the observed negative effects in the TSP treatment.64,65 The treatment conditions used in this study covered present and near-future (for the year 2100) environmental variability in oyster culture zones in China. The decreased pH values used (pH 7.4) is not normally considered to be near-future average under ocean acidification conditions, but this extremely decreased pH scenario is locally and ecologically relevant in the oyster culture zones from where broodstocks were collected for this experiment. In our previous study, the C. gigas larvae

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ASSOCIATED CONTENT

S Supporting Information *

Three tables showing the statistical analysis information for the combined effect of three climate-related stressors on larval performance. Figure 1 shows the pH variability during experiment. Figure 2 shows the effect of three stressors on larval composition. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors would like to thank Ziniu Yu of the South China Sea Institute for Oceanology for providing the oysters used in our experiments. We would also like to thank Tan Shau Hwai Aileen, Richard Bellerby, Mary Sewell, Jason Hall-Spencer, Sam Dupont, Ishimatsu Atsushi, and other members of the ISOACC group for their valuable discussions during the course of this project. We thank 3 anonymous reviewers for constructive comments on the previous version of this manuscript. This study was funded by three GRF grants from the HKSAR-RGC (Grant Numbers: 780510M, 705511P, and 705512P).

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