The physiological and molecular responses of larvae from the reef-building coral Pocillopora damicornis exposed to near-future increases in temperature and pCO2

  ORIGINAL PAPER The physiological and molecular responses of larvaefrom the reef-building coral Pocillopora damicornis exposedto near-future increases in temperature and p CO 2 H. M. Putnam • A. B. Mayfield • T. Y. Fan • C. S. Chen • R. D. Gates Received: 24 August 2012/Accepted: 12 November 2012 Ó Springer-Verlag Berlin Heidelberg 2012 Abstract Given the threats of greenhouse gas emissionsand a changing climate to marine ecosystems, there is anurgent need to better understand the response of not onlyadult corals, which are particularly sensitive to environ-mental changes, but also their larvae, whose mechanismsof acclimation to both temperature increases and oceanacidification are not well understood. Brooded larvaefrom the reef coral Pocillopora damicornis collected fromNanwan Bay, Southern Taiwan, were exposed to ambientor elevated temperature (25 or 29 ° C) and p CO 2 (415 or635 l atm) in a factorial experiment for 9 days, and avariety of physiological and molecular parameters weremeasured. Respiration and rubisco protein expressiondecreased in larvae exposed to elevated temperature,while those incubated at high p CO 2 were larger in size.Collectively, these findings highlight the complex meta-bolic and molecular responses of this life history stageand the need to integrate our understanding across mul-tiple levels of biological organization. Our results alsosuggest that for this pocilloporid larval life stage, theimpacts of elevated temperature are likely a greater threatunder near-future predictions for climate change thanocean acidification. Introduction Future predictions of anthropogenic greenhouse gas emis-sions and climate change represent global environmentalfactors that result in detrimental effects on marine organ-isms. These include two major physical drivers in marineecosystems: (1) elevated seawater temperature and (2)increased atmospheric CO 2 (van Vuuren et al.2008,2011). Increased temperature has well-documented negativeimpacts on organismal physiology, specifically aboveorganisms’ thermal optima (Hofmann and Todgham2010).Ocean acidification (OA), the ‘‘other CO 2 problem’’(Doney et al.2009b), occurs when CO 2 is taken up by theoceans, which shifts the carbonate buffering systemincreasing hydrogen ion concentration and decreasing pH,an environmental setting that is also detrimental to manymarine organisms (reviewed by Doney et al.2009a, Dupontet al.2010a,b; Kroeker et al.2010). Communicated by H.-O. Po¨rtner. Electronic supplementary material The online version of thisarticle (doi:10.1007/s00227-012-2129-9) contains supplementarymaterial, which is available to authorized users.H. M. Putnam ( & ) Á R. D. GatesHawaii Institute of Marine Biology, University of Hawaii,PO Box 1346, Kaneohe, HI 96744, USAe-mail: hputnam@hawaii.eduA. B. Mayfield Á T. Y. FanNational Museum of Marine Biology and Aquarium,2 Houwan Rd., Checheng, Pingtung, Taiwan, ROCT. Y. FanInstitute of Marine Biodiversity and Evolution,National Dong-Hwa University, Hualien, Taiwan, ROCC. S. ChenGraduate Institute of Marine Biotechnology,National Dong-Hwa University, Hualien, Taiwan, ROCC. S. ChenTaiwan Coral Research Center (TCRC), National Museumof Marine Biology and Aquarium, Checheng, Pingtung 944,Taiwan, ROCC. S. ChenDepartment of Marine Biotechnology and Resources, NationalSun Yat-Sen University, Kaohsiung, Taiwan, ROC  123 Mar BiolDOI 10.1007/s00227-012-2129-9  Relative to preindustrial values, near-future temperaturepredictions include increases from 1.5 to 3.5 ° C (Meins-hausen et al.2011). Increased temperatures have negativeimpacts on marine organisms that include changes in thesuccess rates of fertilization and development (Negri et al.2007), shifts in ecological range (Parmesan2006), and mortality (Fitt et al.2001). Additionally, greenhouse gasemission scenarios of representative concentration path-ways (RCPs) predict that near-future (2075) atmosphericCO 2 will range from * 440 to 700 ppm (van Vuuren et al.2011) and reach 490, 650, 850, and [ 1,370 ppm by 2100,based on IMAGE, GCAM, AIM, and MESSAGE models,respectively (Moss et al.2010). Ocean acidification affectsmarine organism reproduction (reviewed by Byrne2011a,b), and while it can be beneficial in some systems ascharacterized by enhanced development and growth rates(Dupont et al.2010a,b,2012), it can also result in detri- mental effects such as malformation during development(Kurihara2008; Talmage and Gobler2010), declines in adult calcification and growth (Kroeker et al.2010), andloss of discriminatory capacity for environmental cues(Munday et al.2009). Together, elevated temperature andOA have the potential to greatly perturb the stability andnet accretion of marine ecosystems, particularly thosebased on calcifying organisms (Hoegh-Guldberg et al.2007; Fabry et al.2008; Hofmann et al.2010). Of specific interest in this context, is the response of reef-buildingcorals, the calcifying structural engineers of highly pro-ductive and diverse ecosystems. Corals are symbioticorganisms that contain single-celled dinoflagellates of thegenus Symbiodinium within their gastrodermal cells. Thesephotosynthetic symbionts produce and translocate themajority of their fixed carbon to the coral host (Muscatineet al.1984) and are responsible for the high productivityand high rates of calcification of coral reef ecosystems.These framework-building corals are especially sensitive toenvironmental changes, and the synergistic impacts of elevated temperature and OA have the potential to drivecoral reef ecosystems past functional thresholds towardalternate ecosystem states (Pandolfi et al.2005; Hoegh-Guldberg et al.2007; Veron2011). Reef-building corals are of particular concern due totheir sensitivity to elevated temperature, which can causebleaching, and in severe cases, bleaching-related mortality(Coles and Brown2003). Likewise, OA has been shown toresult in decreased coral calcification (Langdon andAtkinson2005). To date, the severe declines in the healthof coral reef ecosystems that have been predicted underclimate change scenarios (Hoegh-Guldberg et al.2007;Veron2011) have predominantly been based on studiesdocumenting adult coral responses (reviewed by Lesser2011; Erez et al.2011). However, factors such as inter- specific variability, location-specific responses, physicalsynergisms and antagonisms, the potential for adaptationand acclimatization (Chauvin et al.2011; Edmunds2011; Fabricius et al.2011; Pandolfi et al.2011), and the importance of reproduction and recruitment (Albright2011) have now been recognized as being critical consid-erations in determining the impacts of disturbance regimeson corals and coral reefs.The maintenance of coral reefs demands the continuoussupply of new propagules, recruitment into the population,and persistence of these juveniles within the community.Larval tolerance may present a bottleneck for this process(Byrne2011b), and as such, there is an urgent need tobetter understand the capacity of this early life history stageto respond to temperature and CO 2 regimes expected tocharacterize reefs over the next 50–100 years (Kurihara2008; Byrne2011a,b). Elevated temperature can affect the larval response by modulating settlement choice (Putnamet al.2008) and can also reduce photosynthetic rates(Edmunds et al.2001), settlement success (Randall andSzmant2009), and survivorship (Edmunds et al.2001; Bassim and Sammarco2003; Yakovleva et al.2009). Likewise, larvae of reef-building corals have been shownto exhibit shifts in fertilization success (Albright et al.2010), settlement (Albright and Langdon2011; Nakamura et al.2011b), metabolic demands (Albright and Langdon2011; Nakamura et al.2011b), and survival (Nakamura et al.2011b) when exposed to elevated CO 2 .Despite the importance of this early life history stage,few studies have tracked the response of coral larvae acrossmultiple biological scales. This reflects the fact that earlylife history stages of corals are only very intermittentlyavailable and difficult to work with due to size and otherfactors, so most studies focus either on whole-organismphysiological characteristics, such as respiration, survival,and settlement (e.g., Edmunds et al.2001; Anlauf et al.2011; Nakamura et al.2011b), or solely on molecular parameters, such as gene expression (e.g., Rodriguez-Lanetty et al.2009; Aranda et al.2011; Meyer et al.2011). Here, with the goal of attaining a more comprehensivemechanistic understanding of the phenotypic responses of coral larvae, we examined the effects of increased tem-perature and OA on several aspects of both whole-organ-ism physiology and the sub-cellular response (Fig.1) of brooded Pocillopora damicornis larvae.We selected three physiological response variables toassess larval performance under exposure to elevatedtemperature and p CO 2 (Fig.1). First, we examined thephotochemical efficiency of PSII ( F  V  /  F  M ), in which adecline indicates potential damage to, or photoinactivationof PSII, a documented precursor to the bleaching cascade(Jones et al.2000; Fitt et al.2001). Second, we assessed holobiont metabolism via larval dark respiration measure-ments, with the expectation that metabolic performance Mar Biol  123  would decline under thermal and hypercapnia stress (Byrne2011a; Po¨rtner2008, but see Stumpp et al.2011b). Thirdly, we measured larval size as an indicator of dispersalpotential (Isomura and Nishihira2001), and more gener-ally, fitness.In addition to the physiological variables, we measuredthe expression of two broad categories of genes and pro-teins hypothesized to be important in coral acclimation toaltered environments using real-time quantitative PCR(QPCR) and western blotting, respectively. The first groupincluded four photosynthesis-related genes (Fig.1); pho-tosystem I (  psI  subunit III), phosphoglycolate phosphatase(  pgpase ), ribulose-1,5-bisphosphate carboxylase/oxygen-ase (rusbisco, rbcL  ), and ascorbate peroxidase ( apx1 ).Additionally, the expression of the rubisco protein, RBCL,was measured with western blotting. We hypothesized thatthe expression of the first three genes and the RBCL pro-tein would decrease in larvae exposed to elevated tem-perature and CO 2 , indicating potential damage to thephotosynthetic machinery, and the latter gene, apx1 , wouldincrease to detoxify reactive oxygen species (ROS)generated by damage to the photosynthetic pathway (e.g.,Lesser1997; Venn et al.2008). The second group included the molecular chaperone heat shock protein 70 ( hsp70 )gene in both the host coral and Symbiodinium and theholobiont HSP70 protein. Expression of both hsp70 orthologs and HSP70 was expected to increase in larvaeexposed to elevated temperature and CO 2 , as environ-mental stress has previously been shown to necessitaterepair of damaged protein (Downs et al.2000). Materials and methods Manipulative experimentTo obtain brooded larvae for experimental use, adult P. damicornis colonies were collected from Nanwan Bay,Southern Taiwan (21 ° 56.179 0 N, 120 ° 44.851 0 E), on March12, 2010. The lunar cycle of larval release for brooding P. damicornis in Southern Taiwan is well documented (Fanet al.2002), and peak release follows the new moon (lunar Fig. 1 Generalized schematic summarizing physiological and molec-ular assays for a the whole organism, b cellular response, and c photosynthetic pathway. Numbers within the circles indicate thebiological scale of measurement: 1 physiology, 2 gene expression, 3 protein expressionMar Biol  123  days 6–10). Adult P. damicornis colonies were held inlarval collectors (described in Putnam et al.2008,2010) under ambient light ( * 200 l mol photons m - 2 s - 1 ) andtemperature ( * 25 ° C). Swimming larvae were collectedfrom the tank outlet in mesh beakers via flowing seawateras described in Putnam et al. (2010) near peak larvalrelease on lunar day 9. The larvae from multiple colonieswere pooled and randomly subdivided into groups of eightylarvae that were placed in plankton mesh cylinders( * 100 ml volume, 170 l m mesh). Cylinders containinglarvae were then suspended in each of sixteen 40-l tanksrepresenting four treatments (  N  = 4 tanks per treatment).Larvae were exposed for 9 days (March, 23–April 01,2010) to one of four treatments of temperature and CO 2 (twolevels each) in a factorial crossed design as follows: (1)25 ° C, ambient CO 2 (ATAC), (2) 25 ° C, high CO 2 (ATHC),(3) 29 ° C, ambient CO 2 (HTAC), and (4) 29 ° C, high CO 2 (HTHC). These treatments used spring-time ambient tem-perature and the partial pressure of CO 2 (  p CO 2 ) of NanwanBay,Taiwan,aswellasnear-futurepredictionsof  * 600 ppmCO 2 (RPC 8.5, rising emissions scenario for the years2050–2075;Riahietal.2011; vanVuurenetal.2011) during a bleaching stress event scenario of  * ambient ? 4 ° C(Donner2009; Meinshausen et al.2011). While larvae were not fed during the experiment, P. damicornis larvae arereleasedwithafullcomplementofphotosyntheticsymbionts( * 3,000–7,000 cells larva - 1 ; Putnam et al.2010) andthereforehavecapacitytosustainthemselvesautotrophically(Muscatine et al.1981). In addition, the incoming seawaterwas sand filtered, allowing for natural seawater bacteria andDOM to pass through into the experimental aquaria andcontribute to larval energetic demands.Tanks were illuminated with metal halide lamps (MH-150 W), and irradiance (PAR) measurements were takenmultiple times daily in each tank using a cosine-correctedLi-Cor sensor (192-SA, Li-Cor). Average daily light levelsdid not differ between tanks ( F  (15,111) = 1.0739, p = 0.39)and treatments ( F  (3,123) = 1.8452, p = 0.14), and theaverage irradiance was 176 ± 2 l mol photons m - 2 s - 1 ,(mean ± standard error [SE]). Within the experimentaltanks, temperature was controlled using submerged heaters(AZOO 300w, Taikong Corporation) and external chillers(Aquatech) connected to recirculating pumps for eachtreatment tank. CO 2 control was created by the addition of either ambient or high premixed CO 2 concentrations bub-bled into each tank using an automated feedback CO 2 con-trol system (Qubit Systems; see Edmunds2011for details).Treatment temperatures were assessed several timeseach day using a certified thermometer (15-077-8, accuracy0.05 ° C, resolution 0.001 ° C, Control Company). Samplingfor seawater chemistry was carried out as described below,including the use of the recommended best practices forOA research and reporting (Riebesell et al.2010), andcertified reference materials (CRMs) for total alkalinity(TA) and pH standards obtained from the laboratory of Andrew Dickson (UCSD). In short, tanks were monitoredfor temperature, salinity, TA (potentiometric titrations,Dickson et al.2007, SOP 3B), and pH [total scale, m-cresoldye method, Dickson et al.2007SOP 6B, with modifica-tions to a 1-cm path length (Fangue et al.2010)]. Fromthese measurements, the carbonate chemistry parameters of   p CO 2 ( l atm), HCO 3 - , CO 32 - , DIC ( l mol kg - 1 sw), and X a (aragonite saturation state) were calculated withCO2SYS (Pierrot et al.2006) using the dissociation con-stants for carbonic acid by Mehrbach et al. (1973) refit byDickson and Millero (1987).Physiological parametersFollowing 9 days of exposure to the treatments, the pres-ence or absence of larvae in each tank, in relation to initiallarval sample size, was used to quantify percent survivor-ship. In addition, groups of larvae were sampled fromreplicate tanks of each treatment for response measure-ments. First, one group of 13 larvae was used to assess thedark-adapted yield, or photochemical efficiency of PSII( F  V  /  F  M ), of the Symbiodinium within the coral larvae witha Diving-PAM fluorometer (Walz GmbH) as described inPutnam et al. (2008,2010). Second, larval size was assessed as planar surface area measured from photographsof 10 larvae per tank (Putnam et al.2010) using ImageJsoftware (NIH,http://rsb.info.nih.gov/nih-image/ ). Third,dark respiration was measured as oxygen consumption(nmol O 2 larva - 1 min - 1 ) using a fiber optic oxygen sensor(FOXY-R Ocean Optics) and Ocean Optics spectropho-tometer (USB-2000, Ocean Optics) as described inEdmunds et al. (2011). Briefly, six larvae were placed in2 ml seawater in sealed glass vials and held in the dark for * 2 h. Oxygen concentrations were measured in thetreatment seawater immediately prior to sealing the vialsand prior to any air contact after the dark incubations ateach of the treatment temperatures and CO 2 conditions(ambient = 25.35 ± 0.06 ° C, high = 29.17 ± 0.02 ° C).Prior to use, the probe was calibrated to 0 and 100 %saturation at each treatment temperature. To avoid mea-surement artifacts from oxygen-dependent effects, allmeasurements were completed at [ 80 % O 2 saturation. Alllarval respiration rates were corrected by subtracting theoxygen change in treatment water vials containing no lar-vae under the same conditions.Molecular assaysSamples were assayed with real-time quantitative PCR(QPCR) for mRNA expression of photosynthesis and stress Mar Biol  123  response genes, including photosystem I (  psI  subunit III),phosphoglycolate phosphatase (  pgpase ), ribulose-1,5-bis-phosphate carboxylase/oxygenase (rusbisco, rbcL  ), ascor-bate peroxidase ( apx1 ), and heat shock protein 70 ( hsp70 ,for both host and Symbiodinium ). In addition, rubisco(RBCL) and HSP70 protein expression were assayed fromthe same samples using SDS-PAGE and western blotting.One group of 13 larvae from each tank was collected formolecular analysis, placed in 500 l l TRIzol TM (Invitro-gen), and immediately frozen at - 80 ° C. Preliminarytitration of DNA, RNA, and protein concentration as afunction of the number of larvae revealed that extraction of groups of  C 10 larvae resulted in * 3–4 l g RNA and DNA,and * 140 l g of protein. These quantities were more thansufficient for the molecular analyses.Nucleic acid and protein extractionsRNA, DNA, and protein were extracted from a group of 13 larvae from each of 15 of the 16 treatment tanks; dueto a spill, there were insufficient larvae in one tank of theHTAC treatment to conduct these analyses. Larvae werepulverized with a micropestle in 500 l l TRIzol in amicrocentrifuge tube, and when completely homogenized,an additional 500 l l TRIzol was added. RNAs wereextracted as in Mayfield et al. (2009) except that afterprecipitation, RNA pellets were solubilized in LysisBuffer A of the GeneMark  Ò Plant RNA miniprep puri-fication kit (Hopegen Biotechnology). RNAs were re-purified according to the manufacturer’s instructions,including the 15-min on-column DNase digestion, andeluted in 30 l l DEPC-treated water. The quantity andquality of the RNA were assessed using a NanodropSpectrophotometer (Infinigen) and formaldehyde–agarose(0.8 % TBE) gels post-stained with ethidium bromide,respectively.DNAs from the same samples were extracted as inMayfield et al. (2010) with two modifications. First, theDNA pellets were dissolved in Buffer PCR-A of theAxyprep TM PCR clean-up kit after precipitation (AxygenBiosciences). Second, the DNAs were dried for an addi-tional 5 min at 65 ° C to evaporate residual ethanol, asrecommended by the manufacturer. DNAs were eluted into30 l l of the manufacturer’s elution buffer, and quantity andquality were assessed using the Nanodrop spectrophotom-eter and native agarose gels (0.8 % TBE) post-stained withethidium bromide, respectively. Proteins were extractedfrom the organic phase of the same samples as in Mayfieldet al. (2011) and quantified with the 2-D Quant Ò kit(Amersham Biosciences) according to the manufacturer’sinstructions. RNA/DNA and protein/DNA ratios werecalculated for each sample to estimate total gene andprotein expression, respectively.Reverse transcription and QPCRRNAs (200 ng) were reverse transcribed with the HighCapacity Ò cDNA synthesis kit (Applied Biosystems) sup-plemented with a 1 9 Solaris Ò RNA spike (Thermo-Sci-entific) following the respective manufacturer’s protocols.Prior to QPCR, cDNAs were diluted threefold in DEPC-treated water. Triplicate PCRs were conducted for eachsample and primer set, and a serial dilution of a randomcDNA sample was run on each 96-well plate to estimatethe PCR efficiency of each primer set [ * 98–102 % (datanot shown)]. QPCR was carried out using 1 9 EZ-TIME Ò SYBR Ò Green I mastermix with ROX passive referencedye (Yeastern Biotech. Co., LTD) and 2 l l of cDNA in20 l l reaction volumes. Target gene expression was con-ducted with the primer concentrations, annealing temper-atures, and cycle numbers presented in Table1, and eachthermocycling profile consisted of an initial 10-min incu-bation at 95 ° C followed by cycling at 95 ° C for 15 s andthen 60 s at the respective annealing temperature (Table1).A melt curve was conducted after each run to ensurespecificity of the respective primer sets for all genes.QPCR standardization/normalizationIn order to standardize all assays, equal amounts of DNA,RNA, and protein were loaded into PCRs, reverse tran-scription (RT) reactions, and SDS-PAGE gels, respec-tively. Therefore, the data from the DNA, RNA, andprotein parameters are presented on a relative expressionchange basis, and so are not influenced by any larval sizedifferences. To control for potential differences in RTefficiency between the samples, QPCR amplification of theexogenous Solaris spike was conducted using the kitprimers (200 nM), but not probes (see details of SYBR Ò Green I mastermix above), according to the manufacturer’srecommendations. A melt curve analysis ensured that theSolaris primers were specific to the spike amplicon. Targetgene expression was first normalized to recovery of theSolaris RNA spike, thereby ensuring expression patternswere not influenced by differential RT efficiency.A DNA-based normalization was used (sensu Mayfieldet al.2011) to standardize for potential differences inbiological composition among samples (i.e., the proportionof host and endosymbiont material in each sample). Hostand Symbiodinium hsp70 genome copies were eachamplified in triplicate using 20 ng DNA (10 ng l l - 1 ), 1 9 EZ-TIME SYBR Green mastermix, and 500 nM each pri-mer in 20 l l reaction volumes (Table1). Thermocyclingwas conducted at 95 ° C for 15 min for one cycle, followedby 35 cycles of 95 ° C for 15 s and 59 ° C/62.5 ° C for 60 sfor the host and Symbiodinium ortholog, respectively. Amelt curve analysis was conducted after every reaction. Mar Biol  123