Contrasting modes of inorganic carbon acquisition amongst Symbiodinium (Dinophyceae) phylotypes

Contrasting modes of inorganic carbon acquisition amongstSymbiodinium (Dinophyceae) phylotypes

Patrick Brading1, Mark E. Warner2, David J. Smith1 and David J. Suggett1

1Coral Reef Research Unit, Department of Biological Sciences, University of Essex, Wivenhoe Park, Colchester, CO4 3SQ, UK; 2College of Earth, Ocean, and Environment, University of

Delaware, 700 Pilottown Rd, Lewes, DE 19958, USA

Author for correspondence:Patrick BradingTel: +974 33678460Email: [email protected]

Received: 16 March 2013Accepted: 23 May 2013

New Phytologist (2013)doi: 10.1111/nph.12379

Key words: carbon acquisition, carbon-concentrating mechanism (CCM),dinoflagellate, microalgae, oceanacidification, Symbiodinium.

Summary

! Growing concerns over ocean acidification have highlighted the need to critically understandinorganic carbon acquisition and utilization in marine microalgae. Here, we contrast these char-acteristics for the first time between two genetically distinct dinoflagellate species of the genusSymbiodinium (phylotypes A13 and A20) that live in symbiosis with reef-forming corals.! Both phylotypes were grown in continuous cultures under identical environmental condi-tions. Rubisco was measured using quantitative Western blots, and radioisotopic 14C uptakewas used to characterize light- and total carbon dioxide (TCO2)-dependent carbon fixation,as well as inorganic carbon species preference and external carbonic anhydrase activity.! A13 and A20 exhibited similar rates of carbon fixation despite cellular concentrations ofRubisco being approximately four-fold greater in A13. The uptake of CO2 over HCO"

3 wasfound to support the majority of carbon fixation in both phylotypes. However, A20 was alsoable to indirectly utilize HCO"

3 by first converting it to CO2 via external carbonic anhydrase.! These results show that adaptive differences in inorganic carbon acquisition have evolvedwithin the Symbiodinium genus, which thus carries fundamental implications as to how thisfunctionally key genus will respond to ocean acidification, but could also represent a key traitfactor that influences their productivity when in hospite of their coral hosts.

Introduction

With growing concern regarding the effects of ocean acidification(OA) on the marine environment (Guinotte & Fabry, 2008;Doney et al., 2009), there has been a renewed interest in howmarine microalgae acquire and utilize inorganic carbon (iC) tosupport carbon (C) fixation within the Calvin–Benson cycle(CBC) (Rost et al., 2008; Tortell et al., 2010; Brading et al.,2011). OA alters the speciation of iC within seawater and this, inturn, may affect the productivity and growth of marine microal-gae (Fu et al., 2008; Brading et al., 2011). Understanding themechanisms of iC acquisition is especially relevant for algaebelonging to the dinoflagellate genus Symbiodinium, which arefound both free-living and in symbiosis with reef-building coralsand other cnidarians. This symbiosis is essential for the viabilityof many cnidaria, and is key to the persistence of coral reef eco-systems under present day and future climates (e.g. Iglesias-Prietoet al., 2004; Berkelmans & van Oppen, 2006). However, our cur-rent understanding of the iC environment within the host tissueis limited (Gattuso et al., 1999; Venn et al., 2009), despite thefact that it may have a significant impact on the productivity andgrowth of the in hospite Symbiodinium population (Wooldridge,2009, 2010). The Symbiodinium genus is taxonomically dividedinto numerous phylotypes, physiologically distinct types that aredistinguishable by genetic variability in the second internal

transcribed spacer region (ITS2) of their ribosomal DNA(LaJeunesse, 2001). Different phylotypes can exhibit a range ofresponses to both environmental acclimation (Hennige et al.,2009) and stress (e.g. Robison & Warner, 2006; Suggett et al.,2008; Ragni et al., 2010; Buxton et al., 2012), allowing theircoral hosts to occupy a wide range of niches (e.g. Iglesias-Prietoet al., 2004).

Unlike inorganic macronutrients, such as nitrogen and phos-phorus, iC within ocean surface waters is relatively abundant (andconstant) at c. 2.2 mM. However, most of this iC is in the formof bicarbonate ions (HCO"

3 ), whereas aqueous carbon dioxide(CO2(aq)), the actual iC species fixed within the CBC, contributes< 1%. Rubisco, the enzyme employed by marine microalgae tocatalyse iC fixation, has a relatively low CO2 affinity and specific-ity, particularly within an O2-rich environment. Consequently,this key enzyme is, in fact, considerably undersaturated withrespect to current seawater concentrations of CO2(aq). This isparticularly true of dinoflagellates, such as Symbiodinium, whichpossess a form II Rubisco (Morse et al., 1995; Whitney et al.,1995) that has a significantly lower specificity for CO2 than theform I Rubisco of other phytoplankton taxa (e.g. Tortell, 2000).

In order to overcome the inherent enzymatic limitations ofRubisco, nearly all marine microalgae have evolved strategies,termed ‘carbon-concentrating mechanisms’ (CCMs), to activelyelevate CO2 at the active site of Rubisco (Giordano et al., 2005;

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Reinfelder, 2011); thus, the majority of marine primaryproducers are not considered to be C limited under present dayconditions (Giordano et al., 2005). CCMs not only allow marinemicroalgae to efficiently utilize the small and variable pool ofCO2(aq) for C fixation, but also take advantage of the much largerHCO"

3 fraction. The functional role of CCMs has been exten-sively reviewed within the literature (see Giordano et al., 2005;Reinfelder, 2011), and primarily involves one or more of the fol-lowing processes: (1) the active transport of CO2(aq) into the cell;(2) the dehydration of HCO"

3 to CO2(aq) at the cell surface,which is then transported into the cell; and (3) the active trans-port of HCO"

3 directly into the cell, where it is then converted toCO2(aq). Interconversion between HCO"

3 and CO2(aq), bothoutside and inside the cell, is catalysed by external and internalcarbonic anhydrases (eCA and iCA, respectively) (Reinfelder,2011), and the dehydration of HCO"

3 to CO2(aq) at the cell’sboundary layer by eCA can be further aided by membrane-boundproton (H+) pumps that acidify the immediate external environ-ment (Bertucci et al., 2010).

A number of CCM components have been identified forSymbiodinium, although no direct comparison of iC acquisitionacross different phylotypes has been performed within a singlestudy. Both eCAs and iCAs have been shown to be expressed bySymbiodinium, as well as H+-ATPase (a proton pump) and Na+/HCO"

3 co-transporters (Yellowlees et al., 1993; Al-Moghrabiet al., 1996; Leggat et al., 1999; Bertucci et al., 2010). It is alsoapparent that the expression of these mechanisms differs depend-ing on the host type (e.g. giant clam vs coral) and whether theSymbiodinium is in hospite or free-living (Al-Moghrabi et al.,1996; Leggat et al., 1999; Bertucci et al., 2010). Although thesevarious studies have demonstrated that the environment can playan important role in the type of CCM expressed, few have unfor-tunately specified the phylotype studied, with Bertucci et al.(2010) being the only exception. This has made it impossible toinfer whether differences in CCM type previously described forSymbiodinium are solely driven by environment (i.e. acclimationto different iC conditions/availability) or also by adaptive differ-ences between phylotypes.

It is widely accepted that the changes in productivity andgrowth under OA conditions represent ‘downstream effects’ oniC acquisition and fixation (Rost et al., 2008; Hurd et al., 2009).Recently, we have demonstrated that the effect of OA onfree-living Symbiodinium is phylotype specific, indicating thatkey differences in iC acquisition may also exist between phylo-types of Symbiodinium (Brading et al., 2011). In particular,phylotypes A20 (previously termed A2 in Brading et al., 2011)and A13 exhibit contrasting responses on exposure to an increasein the partial pressure of CO2 (pCO2) from c. 390 to 800 ppmv.Specifically, an increase in productivity, with no change in growthrate, was observed for A20, whereas the opposite was observedfor A13. Based on these previous observations, we sought toexamine these alternative physiological responses to elevated iCavailability (productivity vs growth), and hypothesized that: (1)A20, but not A13, is carbon limited under present day conditionsbecause of an inefficient CCM; and thus (2) A13 and A20 expressdifferent CCM types when grown in the same iC environment.

In order to evaluate the uptake and utilization of iC, we firstdetermined the light-dependent rates of productivity using twoapproaches. Light-dependent rates of C fixation and photosyn-thetic electron transfer (PET) were measured so as to comparethe efficiency with which energy (and reductant), derived fromPET, is utilized for C fixation vs other essential metabolicprocesses, such as CCM/CA activity. We subsequently assessediC affinity and species preference by quantifying the Rubiscoconcentration, measuring the total carbon dioxide (TCO2)dependence of photosynthetic C fixation and performing short-term 14C disequilibrium, which also estimated eCA activity(Martin & Tortell, 2006; Rost et al., 2007).

Materials and Methods

Phylotype selection

For this study, we focused on two Symbiodinium isolates withinclade A: A13 and A20. Phylotype A20 (accession numberEU449053) has yet to be identified from an intact symbiosis andis putatively considered to be a free-living Symbiodinium (M. E.Warner, pers. obs.). The photophysiology of both isolates has beenstudied in relation to photoacclimation (Hennige et al., 2009) andresponse to OA (Brading et al., 2011). Both phylotypes exhibitmarked differences in their physiological responses to light andpCO2, and recent work has shown A13 to be much more suscepti-ble to photoinactivation as a result of thermal stress (McGinleyet al., 2012). These two phylotypes are therefore good candidatesto explore adaptive differences in iC acquisition between closelyrelated Symbiodinium species (see also Brading et al., 2011).

Experimental design

Continuous cultures (volume, 3 l) of Symbiodinium phylotypesA13 and A20 were grown at 26°C under a growth irradiance ofc. 350 lmol photons m"2 s"1 and a 14 h : 10 h light : dark cycle.Fresh medium of nutrient-replete ASP-8A artificial seawater(Provasoli et al., 1957) was continuously supplied to each culture,via a 0.2-lm pore-sized filter (Polycap 36AS; Whatman Filters,GE Healthcare Biosciences, Pittsburgh, PA, USA), at a ratematching the growth rates of each culture, thereby maintaining aconstant cell biomass. The medium was modified to exclude thetris(hydroxymethyl)aminomethane buffer and adjusted to a totalcarbon alkalinity (TAC) of c. 2300 lEq kg

"1 seawater (see Brad-ing et al., 2011). Cultures were subjected to gentle stirring tominimize the settling of cells and to ensure thorough mixing ofthe medium with the gas inflow. Both phylotypes were grown atpCO2 levels representative of ‘present day’ conditions, that is,380–390 ppmv (Table 1; Solomon et al., 2007). The growthpCO2 was maintained using the ‘pH-stat’ culturing methodsdescribed in Brading et al. (2011). Cultures were allowed toacclimate to their pCO2 growth regimes for a period of 21 d(equivalent to at least nine generations), and were only sampledonce steady-state growth was achieved. The cultures weresampled approximately an hour after ‘lights-on’, and allmeasurements were completed before midday. Where certain

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measurements could not be made on both cultures simulta-neously, the order in which they were sampled was reversedbetween replicate measurements (n = 4). Replicate measurementsfrom the same cultures were separated by a period of 1 wk(c. three complete cell generations).

Growth rate

Daily cell counts were obtained microscopically with a Neubauerhaemocytometer (Fisher Scientific, Loughborough, UK) to mon-itor the growth rates of the cultures. As a result of the nature ofcontinuous culturing, changes in cell number were further cor-rected for the dilution rate and volumes of sample removed fromthe culture between cell counts to provide a true measure ofgrowth rate (Eqn 1).

l #loge

Ct$Dt %Vt$DtCt %Vt

! "$ Vwaste

Vvessel

! "

DtEqn 1

(t, time (d); Dt, time interval between sampling; C, concentrationof cells (cells ml"1); V, volume of medium (ml) in the culture ves-sel at a specific time of sampling (subscript ‘t’ or ‘t+Dt’), the wasteoutflow generated by the continuous dilution of the culture dur-ing the time period between sampling (subscript ‘waste’) and theculture vessel when maximally filled (subscript ‘vessel’)). The first

term, loge# Ct$Dt %Vt$Dt

Ct %Vt

$, describes the change in cell number for

the total population during the sampling interval, and the second

term,#Vwaste

Vvessel

$, describes the dilution rate. The specific growth rate

(l, d"1) was calculated as the change in cell concentration perday (d"1). Following the daily cell count, the dilution rate foreach culture vessel was adjusted in order to maintain a constantcell density within each culture. Cell densities of A13 and A20were maintained at c. 85 000 and 60 000 cells ml"1, respectively.

Chlorophyll a (Chla)

Aliquots of c. 100 ml were taken from each culture and filteredthrough glass fibre filters (GF/F). Each filter was snap frozen inliquid nitrogen and stored at"80°C for later extraction and anal-ysis. Each filter was ground in 5 ml of 100% methanol and storedat "20°C for 24 h. The samples were then centrifuged at 3400 gfor 5 min. The absorption of the supernatants between 600 and760 nm was read using a spectrophotometer (U-3000; HitachiHigh Technologies, Krefeld, Germany) against a referencecuvette containing deionized H2O (dH2O). Absorbance spectrawere then corrected for a baseline shift by subtracting a

methanol-only ‘blank’ spectrum, also read against a dH2O refer-ence, and the slope in the spectrum between 750 and 760 nm.The Chla concentration was then quantified using the coeffi-cients and equations of Ritchie (2006), and finally normalized tothe cell density to provide units of picogram Chla per cell.

Particulate organic carbon (POC)

Aliquots of 100 ml were filtered onto pre-combusted GF/Ffilters, dried at room temperature and then stored at "80°C forsubsequent analysis for cellular POC content. Inorganic carbon-ates were removed from the filters by acidification with c. 15%hydrochloric acid. After acidification, each filter was dried at60°C for 48 h. Analysis was performed on the solid module of aTotal Organic Carbon Analyser (Shimadzu, Milton Keynes, UK)against a glucose calibration standard. Once each sample wassealed within the analyser, the carrier gas was allowed to circulatefor up to 2 min before the sample was pushed into the furnace tomaximize the signal-to-noise ratio; this period ensured that thecarrier gas circulating through the furnace and module was suffi-ciently purged of CO2 before the analysis of the filter. The POCconcentration was then normalized to the cell density to giveunits of pg C per cell.

Rubisco

Aliquots of 100 ml were filtered onto GF/F filters, immediatelysnap frozen in liquid nitrogen and stored at "80°C for later anal-ysis of Rubisco content following the protocol described byRyan-Keogh et al. (2012). Briefly, the protein was extracted fromthe samples using a freeze–thaw cycle in liquid nitrogen and incombination with sonication in order to fully rupture the cellmembranes. The total protein concentration was then measuredusing the Lowry protein assay, and the Rubisco concentrationwas subsequently determined by quantitative Western blottingusing the Tricine-sodium dodecylsulfate-polyacrylamide gel elec-trophoresis (Tricine-SDS-PAGE) method of Sch!agger (2006);antibodies and protein standards were used to quantify the geneRbcL (Agrisera AB, V!ann!as, Sweden), which codes for the largesubunit of Rubisco, according to Brown et al. (2008). SDS-PAGE and membrane transfer were performed on an InvitrogenLife Technologies PowerEase 500TM (Invitrogen, Paisley,Renfrewshire, UK). Imaging of the membrane was performed ona BioRad VersaDoc imaging system (Bio-Rad, Hemel Hemp-stead, UK); these images were subsequently processed usingQuantityOne software to quantify protein samples and standards,with all quantification performed within the unsaturated part of

Table 1 The mean (& SE) carbon chemistry of the media in which the two Symbiodinium phylotypes were grown (6 wk). All calculations were performedwith the CO2SYS program developed by Pierrot et al. (2006) with the constants taken fromMillero et al. (2006)

Phylotype TAC (lEq kg"1 SW) pCO2 (ppmv) HCO"3 (lM) CO2"

3 (lM) CO2 (lM)

A13 2315 (& 44) 383 (& 8) 1763 (& 35) 225 (& 4) 10.6 (& 0.2)A20 2402 (& 96) 398 (& 17) 1831 (& 76) 233 (& 10) 11.0 (& 0.5)

Statistical analysis (t-test) of each variable indicated no significant difference between the growth conditions of the two phylotypes. SW, seawater.


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the calibration curve. Quantified RbcL was normalized to Chla,which was measured on subsamples of the protein extract using acalibrated Turner Designs 10-AU fluorometer (Turner Designs,Sunnyvale, CA, USA), to give units of mmol RbcLmol"1 Chla.

Light-dependent electron transfer rates (ETRs)

A FASTtracka II fluorometer, attached to a FASTact laboratory sys-tem (Chelsea Instruments, London, UK), was used to yield pho-tosystem II electron transfer rates (ETRPSII) via a fluorescencelight response curve (see Supporting Information Methods S1).Values of ETRPSII (mol e" g"1 Chla h"1) for each light step werecalculated following Suggett et al. (2008) as:

ETRPSII #E %rPSII%nPSII%F 0

q=F0m

Fv=Fm% '6:023( 105)%'3600( 10"6)

Eqn 2

(E, light intensity (lmol photons m"2 s"1); rPSII, effectiveabsorption cross-section of PSII (nm2 quantum"1); nPSII, con-centration of PSII reaction centres (mol RCII g"1 Chla)). Fv/Fmand F 0

q=F0m are both dimensionless and were calculated as Fv/

Fm = (Fm – F0)/Fm and F 0q=F

0m # 'F 0

m " F 0=F 0m), respectfully.

The conversion factor 6.0239 105 transforms the units of rPSIIto m2 RCII"1, whereas 36009 10"6 transforms the units of E tomol photons m"2 h"1. rPSII is weighted to the spectral quality ofthe light used to induce the fluorescence transient, that is, blueLED array, and was therefore spectrally corrected to match thespectrum of the white LED of the actinic light source:

rPSII # rPSII %"agrowth

"aLEDEqn 3

Values of "agrowth

"aLED were calculated according to the methods describedby Brading et al. (2011), and were 0.374 and 0.420 for A13 andA20, respectively. Values of nPSII were not measured, but assumedaccording to previous values obtained for these strains (grown atsimilar light intensity) using the O2-flash yield approach (Hennigeet al., 2009): 2.319 10"6 mol RCII g"1 Chla and2.509 10"6 mol RCII g"1 Chla for A13 and A20, respectively.

In order to calculate the light-limited electron transferefficiency (initial slope, aETR) and the light-saturated ETR (maxi-mum rate of electron transfer, ETRmax) from each of the rapid-light response curves performed, a modified version of the modelby Platt et al. (1980) was fitted to the data using SigmaPlot 10.0curve-fitting software:

PB # PBS % 1" exp

"aB %PFDPBS

% &% exp

"bB %PFDPBS

!

Eqn 4

where ETRmax # PBS %

#a

a$b

$%# ba$b

$ba and b describes the reduc-

tion in photosynthetic rate at high irradiance (photoinhibition).The saturation irradiance for electron transfer, termed Ek

ETR, wascalculated as (ETRmax/a

ETR) with units of lmolphotons m"2 s"1.

iC uptake

Measurements of radioisotopic 14C uptake, specifically 14C-labelled sodium bicarbonate (NaH14CO3) with a specific activityof 40–60 mCi mmol"1 (Perkin Elmer, Norwalk, CT, USA),were applied to three methods, each assaying different aspects ofiC acquisition. In all cases, fixed 14C was measured on a Tri-Card2910 TR liquid scintillation analyser (Perkin Elmer). The disin-tegrations per minute (DPM) for each sample were averagedfrom a single 20-min count and the rate of C fixation(lmol C l"1 h"1) was calculated as follows:

lmol Cl"1h"1 # DPMsample "DPMT0

DPMTC% VolumeTC

Volumesample% *TCO2+

tEqn 5

where the superscripts ‘sample’, ‘T0’ and ‘TC’ refer to the sample,the initial measurement at the start of an incubation and the totalactivity count, respectively. [TCO2] is the concentration of iC(lM) within the sample and t is the incubation in hours.

Light-dependent C fixation The photosynthetic light responseof each culture was characterized by incubating discrete samples,spiked with 14C radioisotope, at a range of light intensities fromc. 10 to 1330 lmol photons m"2 s"1 (see Methods S2). Theamount of fixed 14C at each photon flux density (PFD) was thenmeasured according to standard liquid scintillation proceduresand normalized to cell number.

The modified model by Platt et al. (1980), described in Eqn 4,was used to calculate the light-limited photosynthetic efficiency(initial slope, a14C) and the light-saturated photosynthetic capac-ity (maximum rate of C fixation, Pmax) of gross C uptake fromeach light curve. Pmax (mol C l"1 h"1) and a14C ((mol C l"1 h"1)(lmol photons m"2 s"1)"1) were normalized to the Chla concen-tration of the sample. The saturation irradiance for C fixation,termed Ek

14C, was calculated as (Pmax/a14C) with units of

lmol photons m"2 s"1.

TCO2-dependent C fixation The photosynthetic TCO2

response was also characterized for each culture between zero and5000 lM TCO2 (see Methods S3). The TCO2-saturated rate ofphotosynthesis (maximum velocity of C fixation, Vmax) and theTCO2 concentration at which the rate of photosynthesis is halfthat of Vmax (half-saturation concentration, Km(TCO2)) were cal-culated by fitting a Michaelis–Menton model to the data, usingSigmaPlot 10.0 curve-fitting software, which describes the veloc-ity of C fixation (v) as a function of TCO2:

v # Vmax % *TCO2+Km $ *TCO2+

Eqn 6

Values calculated for Km(TCO2) can also be used to obtain therespective half-saturation concentrations of CO2(aq) for C fixationby determining the relative speciation of the iC pool at pH 8.0and salinity 35 at Km(TCO2). This provides an indication as towhat the intracellular CO2(aq) concentration would be in the



absence of active uptake (i.e. passive diffusion only) when C fixa-tion is not carbon limited, and allows for comparison with thetheoretical Km(CO2) of Symbiodinium Rubisco (see the Discussionsection).

iC species preference and eCA activity Isotope disequilibrium(Espie & Colman, 1986) was used to quantify the relativecontributions of CO2 and HCO"

3 to iC uptake, following themodification to the technique proposed by Martin & Tortell(2006) (see Methods S4). In order to assess the presence andactivity of eCA, duplicate samples, pretreated with the mem-brane-impermeable CA inhibitor acetazolamide (AZ), were alsorun according to this protocol. To determine the relative contri-butions of CO2 and HCO"

3 to carbon uptake, data from boththe control and +AZ treatments were analysed according to themodel and protocol described by Martin & Tortell (2006), fittedusing SigmaPlot 10.0 curve-fitting software:

DPMt #Vt '1" f ) a1t$

DSACO2

SADIC

! "( '1" e"a1t )

! "

a1

$Vt 'f ) a2t$

DSAHCO"3

SADIC

% &( '1" e"a2t )

% &

a2

Eqn 7

(Vt, total rate of C uptake; f, fraction of C uptake supported bydirect transport of HCO"

3 into the cell; a1 and a2, first-order rateconstants for CO2 and HCO"

3 hydration and dehydration,respectively, which are temperature, salinity and pH dependent).In the absence of enzymatic activity, a1 and a2 were determinedto be 0.0894 and 0.1135 s"1, respectively (using the equations ofEspie & Colman, 1986), with corrections for temperature andsalinity (Johnson, 1982). SADIC is the specific activity of all theiC species at equilibrium, and DSACO2 and DSAHCO3- are thedifferences between the initial (i.e. at pH 7.0) and equilibrium(i.e. at pH 8.5) values of the specific activities of CO2 andHCO"

3 . The values of DSACO2/SADIC and DSAHCO3-/SADIC

were calculated using the difference in pH between 14C spike andbuffered ASP-8A, as well as TCO2 of the buffered ASP-8A.

In the +AZ treatments, the rate constants a1 and a2 wereassumed to equal the uncatalysed value, as any potential eCAactivity is blocked by the presence of the inhibitor. Therefore, inthe +AZ treatments, the model described in Eqn 7 was fitted tothe data by allowing Vt and f to vary and restraining the otherparameters to the values described previously. For the controltreatments, the model was fitted by restraining f to the value cal-culated in the corresponding +AZ model fit and, instead,

allowing a1 to vary as the model parameter (with a2 expressedas a function of a1 rather than as an actual model parameter; seeEspie & Colman, 1986), but constrained to be equal to or greaterthan the uncatalysed value. The change in a1 between the controltreatment and uncatalysed value of the +AZ treatment can thenbe used to describe the eCA activity employing the followingequation (Rost et al., 2007):

eCAact # acontrol1 " a$AZ1

a$AZ1

Eqn 8

Statistical analysis

The growth rate, cellular Chla content, parameters relating tolight-dependent C fixation and iC affinity, and the iC speciespreference and eCA activity under ambient pCO2 (i.e. c.380 ppmv) were compared between the two phylotypes using aparametric-independent samples t-test or a non-parametricMann–Whitney test where appropriate. The threshold ofstatistical significance was set at P < 0.05. All statistical tests wereperformed using SPSS statistical analysis software (version 16.0;SPSS, Portsmouth, UK).

Results

Growth rate, Chla, POC and Rubisco

Consistent with previous work for these strains, the growth rateof phylotype A20 was c. 50% higher than that of A13 (Table 2).Both cellular Chla and POC contents were also higher in A20than in A13 (Table 2), an observation that is perhaps unsurpris-ing given that the cell size of A20 is considerably larger than thatof A13 (see Brading et al., 2011). Indeed, POC normalized to thecorresponding values of Chla yielded no significant difference forthe C : Chla ratio between the phylotypes, despite the c. 30%higher values for A13 (Table 2). Chla-normalized Rubisco con-centrations (mmol RbcLmol"1 Chla) were approximatelyfour-fold greater for A13 than for A20 (Table 2). Therefore, A13has a greater capacity than A20 for C fixation, with a larger poolof Rubisco available to catalyse this reaction.

Light-dependent photosynthesis Two approaches were used tomeasure light-dependent photosynthesis, Chla fluorescence(ETR) and 14C uptake (C fixation), so as to determine whetherenergy derived from PET is closely coupled to C fixation orwhether additional processes also draw heavily from PET. Both

Table 2 Mean (& SE) growth rate (l, d"1), cellular chlorophyll a (Chla) (pg Chla per cell) and particulate organic carbon (POC) (pg C per cell) content,g C : g Chla ratio and Chla-normalized Rubisco concentration (mmol RbcLmol"1 Chla) of Symbiodinium phylotypes A13 and A20

Phylotype l pg Chla per cell pg C per cell C : Chla mmol RbcLmol"1 Chla

A13 0.287 (& 0.029) 1.03 (& 0.14) 111 (& 12) 110 (& 10) 1.85 (& 0.35)A20 0.425 (& 0.024) 4.16 (& 0.38) 358 (& 31) 86 (& 3) 0.47 (& 0.03)Difference s s s ns s

‘s’ and ‘ns’ denote whether there was a significant or non-significant difference between the two phylotypes, respectively, when analysed using a t-test.




ETRmax (Fig. 1a) and aETR (Fig. 1b) were the same for thetwo phylotypes, with combined mean values of c.2.76 mol e" g"1 Chla h"1 and c. 0.008 (mol e" g"1 Chla h"1)(lmol photons m"2 s"1)"1, respectively. Values of Ek

ETR werealso the same for the two phylotypes with a combined mean of c.348 lmol photons m"2 s"1, that is, values similar to the growthirradiance (Fig. 1c). Corresponding light response parametersmeasured by C fixation, Pmax (Fig. 2a) and a14C (Fig. 2b), werealso the same for both phylotypes, with mean values of c.282 mmol C g"1 Chla h"1 and c. 1.65 (mmol C g"1 Chla h"1)(lmol photons m"2 s"1), respectively. However, the Ek

14C valueof A20 was c. 18% lower than that of A13 (Fig. 2c) and both werelower than the growth irradiance by a factor of c. 2.3 and 1.9,respectively, suggesting that C fixation was not light limited forboth phylotypes.

Comparisons of the light response parameters derived fromETR vs those from C fixation provide a photosynthetic ‘currencyexchange rate’ (sensu Suggett et al., 2009) for both light-limiting(aETR : a14C) and light-saturating (ETRmax : Pmax) conditions(Table 3), and provide an insight into the coupling between PETand C fixation. Interestingly, the ‘exchange rate’ for both light-limited (aETR : a14C) and light-saturated (ETRmax : Pmax) photo-synthesis was the same for the two phylotypes, with values of c. 4.9and 9.8 mol e"mol C"1, respectively (Table 3). Furthermore, thecomparison of Ek

ETR with Ek14C (Ek

ETR : Ek14C, dimensionless)

shows that C fixation for both phylotypes is saturated at approxi-mately two-fold lower irradiances than PET (Table 3). That thevalues for ETRmax : Pmax were approximately two-fold greaterthan those for aETR : a14C and Ek

ETR : Ek14C for both phylotypes

suggests that the degree of coupling between PET and C fixationis dependent on light intensity (i.e. limiting vs saturating), butindependent of phylotype here.

TCO2-dependent C fixation Based on the values of Ek14C

obtained from the light-dependent C fixation measurements, the

incubations for TCO2-dependent C fixation were performedunder light-saturating conditions for C fixation for bothSymbiodinium phylotypes. Values of the Chla-specific Vmax

(mmol C g"1 Chla h"1), that is, the carbon saturated (maximum)velocity of C fixation, did not differ between the two phylotypes(Table 4), and were also statistically similar to their correspond-ing Pmax values. This similarity between Vmax and Pmax suggeststhat the light-dependent C fixation incubations were performedat saturating iC concentrations for photosynthesis and that thegrowth of the two phylotypes was thus not carbon limited. Pairedsamples for Rubisco quantification and TCO2-dependent C fixa-tion were not performed in the present study. However, the meanVmax can still be normalized to the mean Rubisco concentrationby performing error propagation to account for this (Table 4).Rubisco-normalized Vmax (mol C mmol RbcL"1) was c. 3.3-foldgreater for phylotype A20 than for A13, demonstrating that car-bon-saturated C fixation is not related to the size of the Rubiscopool, which was larger in A13. However, carbon-limited Cfixation might be. Comparisons of Km(TCO2) between the twophylotypes showed that A13 had a higher affinity for iC, as theKm(TCO2) value obtained for this phylotype was c. 36% lowerthan the Km(TCO2) value of A20. Thus, C fixation in A13 is satu-rated at a lower TCO2 than in A20, perhaps as a consequence ofits larger pool of Rubisco.

(a) (b) (c)

Fig. 1 Mean (& SE) (a) ETRmax (photosyntheticcapacity, mol e" g"1 Chla h"1), (b) aETR (initialslope or photosynthetic efficiency, (mol e" g"1

Chla h"1)(lmol photonsm"2 s"1)"1) and (c)Ek

ETR (light saturation photon flux density(PFD), lmol photonsm"2 s"1) ofSymbiodinium phylotypes A13 andA20.

(a) (b) (c)Fig. 2 Mean (& SE) (a) Pmax (photosyntheticcapacity, mmol C g"1 Chla h"1), (b) a14C

(initial slope or photosynthetic efficiency,(mmol C g"1 Chla h"1)(lmol photonsm"2 s"1)) and (c) Ek

14C (light saturationphoton flux density (PFD),lmol photonsm"2 s"1) of Symbiodiniumphylotypes A13 and A20. An asterisk abovethe data indicates statistical differencesbetween the two phylotypes for thatvariable.

Table 3 Mean (& SE) aETR : a14C, ETRmax : Pmax (mol e"mol C"1) andEk

ETR : Ek14C (dimensionless) of Symbiodinium phylotypes A13 and A20

Phylotype aETR : a14C ETRmax : Pmax EkETR : Ek

14C

A13 5.34 (& 0.45) 9.73 (& 0.54) 1.86 (& 0.18)A20 4.54 (& 0.29) 9.88 (& 1.20) 2.22 (& 0.34)Difference ns ns ns

‘s’ and ‘ns’ denote whether there was a significant or non-significantdifference between the two phylotypes, respectively, when analysed usinga t-test.



iC species preference and eCA activity Examples of 14C incor-poration of both A13 and A20, in the presence and absence ofthe eCA inhibitor AZ, during short-term 14C disequilibriumincubations can be seen in Fig. 3. Fitting the disequilibriummodel described in Eqn 7 to the +AZ samples provided an esti-mation of the fraction of C fixation attributable to direct HCO"

3

transport only. Both phylotypes of Symbiodinium exhibited verylittle direct HCO"

3 transport, as indicated by the low f(HCO"3 )

values of 0–6% (Table 5), suggesting that the iC species trans-ported across the cell membrane is predominantly CO2(aq). Fur-thermore, fitting the disequilibrium model when assuming‘CO2-only’ uptake (i.e. constraining f equal to zero) also pro-vided a good fit from the +AZ-treated samples (data not shown),thus supporting the inference that CO2(aq) is the predominant iCspecies transported across the cell membrane by both phylotypes.

Comparison of the catalysed rates of CO2 hydration (a1control)

with the uncatalysed rate (a1+AZ, a theoretical 0.0894 s"1; Fig. 4)

indicated that phylotype A20 possesses some degree of eCA activ-ity. eCAact (dimensionless), a proxy of eCA activity, was 2.3 and0.09 for A20 and A13, respectively. A20 is therefore capable ofutilizing the HCO"

3 pool, indirectly, for C fixation, whereas A13largely relies on direct CO2 uptake.

Discussion

Productivity

Rates of C fixation were similar between the two phylotypes andagree with the rates reported previously for Symbiodinium (Lesser,1996). In addition, ETRs were also similar, such that when these

rates were compared with their respective C fixation rates, littledifference was observed in the coupling of PET and C fixationbetween the two phylotypes at both low and high light conditions(i.e. aETR : a14C and ETRmax : Pmax, respectively). These compari-sons of ETR with rates of C fixation provide an indication ofenergy investment within the cell. In the present study, the com-parison of light-limited ETR with C fixation (aETR : a14C) yieldedan average ratio of c. 4.9 mol e"mol C"1, which agrees well withthe theoretical minimum PET : C fixation ratio of 5 (Suggettet al., 2009). Under light-limiting conditions, therefore, PET inPSII is closely coupled to the C fixation occurring in the CBC forboth phylotypes. Under saturating light conditions, however,these two processes become decoupled (ETRmax : Pmax of c.9.8 mol e"mol C"1), with C fixation becoming light saturatedbefore PET, as indicated by the Ek

ETR : Ek14C ratio of 2.

A similar observation of C fixation saturating at lowerirradiances than PET (i.e. Ek

14C < EkETR) was also reported for

the cyanobacterial species Synechococcus, whereby ETR continuedto increase with irradiance after the saturation of C fixation (Bai-ley et al., 2008), and is also consistent with the notion, observedfor many taxa across pelagic systems, that the rate of electrontransfer from the primary quinone acceptor to plastoquinone(PQ) (1/sQB?PQ) typically exceeds the rate of whole-chain elec-tron transfer (1/sPSII) (Moore et al., 2006). Such a pattern indi-cates the occurrence of alternative electron sinks downstream ofPSII, but before their use in C fixation (Suggett et al., 2009).Therefore, our observations for the two phylotypes ofSymbiodinium studied here suggest the existence of alternativeelectron sinks under high irradiances that exceed what is neces-sary to support C fixation.

Table 4 Mean (& SE) VmaxChl (mmol C g"1 Chla h"1), Vmax

RbcL

(mol Cmmol"1 RbcL h"1), Km(TCO2) (lM TCO2), Km(CO2) (lMCO2(aq)) andKm(HCO3-) (lMHCO"

3 ) of Symbiodinium phylotypes A13 and A20

Phylotype VmaxChl Vmax

RbcL Km(TCO2) Km(CO2)

A13 259 (& 55) 125 (& 36) 572 (& 85) 4.7 (& 0.7)A20 216 (& 33) 414 (& 68) 889 (& 75) 7.3 (& 0.6)Difference ns – s s

‘s’ and ‘ns’ denote whether there was a significant or non-significantdifference between the two phylotypes, respectively, when analysed usinga t-test.

(a) (b)

Fig. 3 Example results for the 14Cdisequilibrium incubations performed onphylotypes A13 and A20, in the absence(control, closed circles) and presence (+AZ,open circles) of the carbonic anhydrase (CA)inhibitor acetazolamide (AZ). Values of f(HCO"

3 ) represent the proportion of HCO"3

uptake relative to C fixation in the +AZsamples.

Table 5 The mean (& SE) fraction of carbon (C) fixation supported bydirect HCO"

3 uptake, f(HCO"3 ), and a measure of external carbonic anhy-

drase (eCA) activity for Symbiodinium phylotypes A13 and A20

Phylotype f(HCO"3 ) eCA activity

A13 0.005 (& 0.005) 0.09 (& 0.09)A20 0.061 (& 0.061) 2.30 (& 0.33)Difference ns s

Both variables are dimensionless. ‘s’ and ‘ns’ denote whether there was asignificant or non-significant difference between the two phylotypes,respectively, when analysed using a Mann–Whitney test.




In Brading et al. (2011), we demonstrated that a significantamount of light-enhanced O2 consumption occurs in both A13and A20. The pathways responsible for this, such as photorespira-tion and Mehler-Ascorbate-Peroxidase (MAP) activity, act as elec-tron sinks because of their requirement for electrons to reduce O2,and thereby perform a photoprotective role under high-light stress(Niyogi, 2000; Ort & Baker, 2002; Mackey et al., 2008). MAPand other alternative oxygen-consuming pathways have beenobserved previously in Symbiodinium under high-light stress, andspecifically within clade A phylotypes (McCabe-Reynolds et al.,2008; Suggett et al., 2008), as well as in other dinoflagellate spe-cies, such as Prorocentrum minimum (Suggett et al., 2009). Cyclicelectron transfer (CET) in PSI may also contribute to the decou-pling of ETR from C fixation in clade A Symbiodinium phylo-types, acting as an alternative sink for electrons, as well as anadditional source of ATP to linear PET (McCabe-Reynolds et al.,2008). It is not possible in the present study to determine whichof these processes is responsible for the observed decouplingbetween PET and C fixation under high light. However, ourobservations provide the first quantitative means by which fluores-cence-based measures of Symbiodinium productivity could betterevaluate C fixation, and demonstrate that coupling between PETand C fixation is primarily driven by irradiance and is independentof phylotype.

Rubisco

To our knowledge, the quantification of form II Rubisco inSymbiodinium spp. has never been published previously. Form IIRubisco is notoriously unstable once extracted (Whitney et al.,1995), and is therefore difficult to accurately quantify. Themethod employed in the present study, adapted from Brownet al. (2008), provides the first ever quantification ofSymbiodinium Rubisco concentration. The concentration of formII Rubisco in phylotypes A13 and A20 (1.9 and 0.5 mmolRbcLmol"1 Chla, respectively) was of a similar order of magni-tude to, but lower than, the concentrations of form I Rubisco

(3.6–8.2 mmol RbcLmol"1 Chla) reported previously for a num-ber of cyanobacteria species, including Trichodesmium spp. andSynechococcus elongatus (Brown et al., 2008). It is unsurprisingthat the concentration of Rubisco is greater in species of cyano-bacteria, given that their form I Rubisco consists of eight largesubunits, whereas form II Rubisco of dinoflagellates can have asfew as two large subunits (Tabita et al., 2008).

Despite having an almost four-fold greater concentration ofRubisco than A20, this did not translate to a greater maximumcapacity for C fixation in A13. When expressed relative to Rubi-sco, values of Vmax for A13 were considerably lower than those forA20, suggesting that C fixation at saturating concentrations of iCis not proportional to the size of the Rubisco pool. The lower Ru-bisco-normalized Vmax of A13 may be a consequence of insuffi-cient CO2(aq) elevation, relative to O2, at the active site of Rubiscoor, alternatively, a lower Rubisco specificity (s). Taxonomic differ-ences in s are well established, and some degree of variability for swithin taxonomic groups has also been shown, although notwithin dinoflagellates (reviewed in Tortell, 2000).

Alternatively, it may be that, although A13 has the larger poolof Rubisco, not all of it is in its active form (MacIntyre et al.,1997). In higher plants and several species of phytoplankton,Rubisco activity is controlled by both the presence of Rubiscoactivase, a soluble protein that facilitates the carbamylation ofRubisco, and CA1P, an inhibitor synthesized in the dark andbroken down in the light (Portis, 2003). The role of Rubisco acti-vase is to balance C fixation with the supply of ATP and reduc-tant from the light reactions of the thylakoid membrane (Portis,2003). In the case of Symbiodinium phylotype A13, the largepool of Rubisco, although seemingly unnecessary, may be a strat-egy used by this isolate to optimize carboxylation, activating anddeactivating the enzyme in response to changing supplies of iC,ATP and reductant. However, the actual presence of a Rubiscoactivase in Symbiodinium is yet to be determined (Lilley et al.,2010), and so we can only speculate at this time.

Although no direct measure of Km(CO2) for SymbiodiniumRubisco has been published to date, Leggat et al. (2002) approxi-mated a value of 50–60 lM CO2 by assuming a s value similarto that for form II Rubisco of Amphidinium carterae (i.e. s = 37)(Whitney & Andrews, 1998) and similar kinetic properties to theform II Rubisco of the proteobacterium Rhodospirillum rubrum.In the present study, Km(CO2) for C fixation was c. 4.7 lM forA13 and c. 7.3 lM for A20. If both phylotypes relied solely onpassive diffusion alone, this concentration of CO2(aq) would beconsiderably lower than that required to saturate Rubisco andsupport maximal C fixation. Such a consideration provides evi-dence that active processes (i.e. CCMs) must be functioning inA13 and A20 to increase the internal concentration of the CO2

(aq) pool relative to the external concentration by either: (1)actively accumulating CO2(aq) within the cell; or (2) supplement-ing/replacing the uptake of CO2(aq) with HCO"

3 .

iC affinity and species preference

As already mentioned, evidence of an active CCM inSymbiodinium has been published previously, but the exact

Fig. 4 Mean (& SE) catalysed rates of CO2 hydration (a1control, s"1) of

Symbiodinium phylotypes A13 and A20. The dashed line marks thetheoretical pH- and temperature-dependent uncatalysed rate of CO2

hydration (a1+AZ, s"1). An asterisk above the data indicates statistical

differences between the two phylotypes.



nature of the mechanisms utilized sometimes appears to conflictbetween studies, and it is unclear whether these differences aretaxonomically vs environmentally driven (Al-Moghrabi et al.,1996; Goiran et al., 1996; Leggat et al., 1999). Symbiodiniumrecently isolated from corals has been shown to preferentiallytransport HCO"

3 directly into the cell (Al-Moghrabi et al., 1996;Goiran et al., 1996), whereas that recently isolated from giantclams exhibited a preference for direct CO2 uptake (Leggat et al.,1999). Interestingly, in the latter study, this preference changedto direct HCO"

3 uptake after 2 d of being maintained in culture(Leggat et al., 1999). Together, these studies show that theexpression of a CCM can be environmentally driven for somephylotypes of Symbiodinium, with some degree of plasticity torespond to changes in the iC environment. Our study confirmsthat CCMs operate in Symbiodinium, but further shows that thenature of Symbiodinium CCMs are algal specific and are, to someextent, taxonomically driven. A13 and A20 both have iC uptakemechanisms that differ from those described previously forcultured Symbiodinium, further highlighting the physiologicaldiversity within this genus that has been reported previously inrelation to other environmental factors, such as light and temper-ature (Robison & Warner, 2006; Suggett et al., 2008; Hennigeet al., 2009; Ragni et al., 2010).

Although a direct comparison of the expression of CCM-related proteins between the phylotypes was not made, the 14Cdisequilibrium technique does provide an indication as to whichiC species was used and whether uptake was direct or aided byeCA activity. In the presence of AZ, and therefore in the absenceof any potential eCA activity, neither phylotype appeared to becapable of performing the direct transport of HCO"

3 into thecell, but instead were entirely dependent on direct CO2(aq)

uptake. Such an observation suggests an absence of proteinscapable of transporting HCO"

3 , such as the Na+/HCO"3

co-transporter, which has been proposed previously for aSymbiodinium sp. isolated from Galaxea fascicularis (Al-Moghra-bi et al., 1996). The inability to transport HCO"

3 directly has alsobeen observed previously in the dinoflagellates A. carterae andHeterocapsa oceanica (Colman et al., 2002; Dason et al., 2004).However, from the data available here, it is not clear whetherA13 and A20 lack the genes necessary to build the proteins thatfacilitate direct HCO"

3 uptake, as in A. carterae (Colman et al.,2002), or just do not express them under the growth conditionsused for this study.

In the absence of AZ, the calculated rate constant of CO2

hydration in the A20 samples was significantly higher than thatof the theoretical uncatalysed rate of 0.0894 s"1, and suggeststhat eCA was present in A20. Using the equation for eCA activityproposed by Rost et al. (2007), A20 had an eCA activity of c. 2.3,whereas that of A13 was c. 0.09. A20 is therefore also able to uti-lize HCO"

3 by first converting it to CO2(aq) using eCA, whereasA13 lacks the ability to utilize this additional iC pool. Clearly,this has important implications regarding the effects of OA,which has a greater effect on the CO2(aq) fraction of the iC pool.Previous work on Symbiodinium A1 has shown that dehydrationof HCO"

3 is further aided by the action of membrane-bound H+-ATPase which acidifies the periplasmic space around the cell

(Bertucci et al., 2010). It is not possible from the present study toconclusively determine whether a similar mechanism occurs inA20, but the fact that little HCO"

3 uptake was observed wheneCA was inhibited suggests that A20 lacks any other mechanismto promote HCO"

3 dehydration. Phylotype A13, however, doesnot support any of its C fixation by HCO"

3 uptake, either bydirect transport or indirectly by first converting it to CO2(aq) witheCA. The apparent lack of eCA activity in phylotype A13 hasalso been observed in A. carterae and H. oceanica (Colman et al.,2002; Dason et al., 2004), and also agrees with the findingsof Leggat et al. (1999), whereby the addition of AZ to a cultureof Symbiodinium, isolated from Tridacna gigas, had no effect onthe active uptake of iC. Although the phylotype was not specifiedin this study, it was most likely an A3 isolate, following the workby LaJeunesse (2001), which found that all of the Symbiodiniumisolates taken from host species within the Tridacna genus fellinto this classification.

Carbon acquisition and OA

In Brading et al. (2011), we showed that A13 and A20 exhibitedincreased growth rates and productivity (as O2 evolution),respectively, in response to a doubling of growth pCO2 from380 to 780 ppm. We interpreted these phylotype-specificresponses as an indication of CCM down-regulation andreinvestment of cellular energy into growth for A13 and as Climitation in A20 (Brading et al., 2011). In the light of the find-ings in the present study, we hypothesize that A13 may down-regulate the ATP invested in the active CO2 uptake pathwayunder OA conditions, afforded by increased availability of CO2

(aq) to support greater passive uptake, and utilize this energy incell division instead.

Having established that A20 is saturated with respect to iCunder present day pCO2, an alternative interpretation for itsobserved increase in productivity under OA conditions, otherthan C limitation, must be sought. Specifically, A20 showed a60% increase in O2 evolution at 780 ppm. Assuming a constantphotosynthetic quotient (PQ; mol O2; mol CO2), this wouldinfer that C fixation also increased by 60%. However, thisassumption is not necessarily valid. An increase in O2 evolution,independent of C fixation, may indicate a reduction in O2-con-suming pathways, such as photorespiration (Geider & Osborne,1992). It is possible, with the dataset provided in Brading et al.(2011), to estimate PQ using the values of net O2 evolution andthe product of A20’s growth rate and cellular POC concentra-tion. This approach estimates PQ of A20 to be 0.4 at 380 ppmpCO2 and 0.6 at 780 ppm pCO2. Using the direct measurementsof C fixation reported in this study, we also arrive at a similar esti-mate of PQ for A20 under 380 ppm pCO2 of 0.5. Althoughlower than the theoretical minimum PQ of 1, such instanceshave been reported previously for marine phytoplankton (e.g.Carpenter & Roenneberg, 1995), and may be a consequence ofsignificant photorespiratory activity (Geider & Osborne, 1992).

If the Rubisco specificity of A20 is considerably low or theconcentration of CO2(aq) at the site of Rubisco is not sufficientlyelevated, oxygenase activity will be favoured. Elevated CO2(aq),




as a result of OA, may therefore suppress this oxygenase activityand consequently increase the PQ of A2 at elevated pCO2. Thelack of a significantly increased growth rate, despite the higherrates of productivity observed for A20 under OA conditions(Brading et al., 2011), may be a consequence of continued ener-getic investment in eCA to support indirect HCO"

3 uptake ratherthan growth. This suggests that the CCM of A20 has limitedplasticity to acclimate to changes in the iC environment, such asthe increase in pCO2 associated with OA. It would be necessaryto study the plasticity of Symbiodinium CCMs under differentenvironmental conditions and, in particular, elevated pCO2 inorder to prove this.

Conclusions

The present study has shown that, although A20 has a loweraffinity than A13 for iC, C fixation in both phylotypes was satu-rated with respect to iC under present day pCO2 conditions.Both phylotypes were capable of active CO2(aq) uptake, but A20was also able to utilize HCO"

3 via eCA. This demonstrates thatdifferent mechanisms of uptake exist for these two phylotypesand, based on the differential effect of OA on other phylotypes ofSymbiodinium (Brading et al., 2011), no doubt further strategiesexist within this genus. These differences are adaptively drivenand will probably determine a phylotype’s response to environ-mental shifts, such as OA, when free-living, but may also haveimplications for their productivity when in hospite. The pH andiC environment within the coral host is complex and its interac-tion with the zooxanthellae is still not fully understood (Vennet al., 2009; and see commentary by Brownlee, 2009).

These adaptations should be considered against the wider phy-logenetic diversity of Symbiodinium in order to understand theevolutionary significance of these different CCMs, as they maycontribute greatly to the shaping of the diversity of this genusunder future environmental conditions. Further study is requiredto determine whether additional differences in iC acquisitionexist in other clades of Symbiodinium, and whether the regulationof the CCM to changes in the iC availability and speciation dif-fers between phylotypes. This study is also the first time thatdirect comparisons between ETR and C fixation rates have beenperformed for Symbiodinium, and showed that significant decou-pling between these rates occurs under saturating irradiances (i.e.where productivity may become increasingly limited by iC avail-ability; Muscatine et al., 2005), highlighting the dependence onadditional electron sinks, such as oxygen-consuming pathways ofMAP and chlororespiration.

Acknowledgements

We thank Dr Phillip Davey and Tania Maynard-Cresswell, at theUniversity of Essex, for technical support during this study, and DrMark Moore and Dr Tom Bibby, at the University of Southamp-ton (UK), for performing the Rubisco analysis. This research wasmade possible through a Natural Environment Research Council(NERC) PhD studentship to P.B. Funding to M.E.W. wasprovided by the National Science Foundation (CRI-OA 104940).

References

Al-Moghrabi S, Goiran C, Allemand D, Speziale N, Jaubert J. 1996. Inorganiccarbon uptake for photosynthesis by the symbiotic coral–dinoflagellateassociation II. Mechanisms for bicarbonate uptake. Journal of ExperimentalMarine Biology and Ecology 199: 227–248.

Bailey S, Melis A, Mackey KRM, Cardol P, Finazzi G, van Dijken G, Berg GM,Arrigo K, Shrager J, Grossman A. 2008. Alternative photosynthetic electronflow to oxygen in marine Synechococcus. Biochimica et Biophysica Acta 1777:269–276.

Berkelmans R, van OppenM. 2006.The role of zooxanthellae in the thermaltolerance of corals: a ‘nugget of hope’ for coral reefs in an era of climate change.Proceedings of the Royal Society B: Biological Sciences 273: 2305–2312.

Bertucci A, Tambutt#e #E, Tambutt#e S, Allemand D, Zoccola D. 2010.Symbiosis-dependent gene expression in coral–dinoflagellate association:cloning and characterization of a P-type H+-ATPase gene. Proceedings of theRoyal Society B: Biological Sciences 277: 87–95.

Brading P, Warner ME, Davey P, Smith DJ, Achterberg EP, Suggett DJ. 2011.Differential effects of ocean acidification on growth and productivity andphotosynthesis among phylotypes of Symbiodinium (Dinophyceae). Limnologyand Oceanography 56: 927–938.

Brown CM, MacKinnon JD, Cockshutt AM, Villareal TA, Campbell DA. 2008.Flux capacities and acclimation costs in Trichodesmium from the Gulf ofMexico.Marine Biology 57: 413–422.

Brownlee C. 2009. pH regulation in symbiotic anemones and corals: a delicatebalancing act. Proceedings of the National Academy of Sciences, USA 106: 16541–16542.

Buxton L, Takahashi S, Hill R, Ralph PJ. 2012. Variability in the primary site ofphotosynthetic damage in Symbiodinium sp. (dinophyceae) exposed to thermalstress. Journal of Phycology 48: 117–126.

Carpenter EJ, Roenneberg T. 1995. The marine planktonic cyanobacteriaTrichodesmium spp.: photosynthetic rate measurements in the SW AtlanticOcean.Marine Ecology Progress Series 118: 267–273.

Colman B, Huertas IE, Bhatti S, Dason JS. 2002. The diversity of inorganiccarbon acquisition mechanisms in eukaryotic microalgae. Functional PlantBiology 29: 261–270.

Dason JS, Huertas IE, Colman B. 2004. Source of inorganic carbon forphotosynthesis in two marine dinoflagellates. Journal of Phycology 40:285–292.

Doney SC, Fabry VJ, Feely RA, Kleypas JA. 2009.Ocean acidification: the otherCO2 problem. Annual Review of Marine Science 1: 169–192.

Espie GS, Colman B. 1986. Inorganic carbon uptake during photosynthesis.Plant Physiology 80: 863–869.

Fu F-X, Zhang Y, Warner ME, Feng Y, Sun J, Hutchins DA. 2008. Acomparison of future increased CO2 and temperature effects on sympatricHeterosigma akashiwo and Prorocentrum minimum. Harmful algae 7: 76–90.

Gattuso J-P, Allemand D, Frankignoulle M. 1999. Photosynthesis andcalcification at cellular, organismal and community levels in coral reefs: areview on interactions and control by carbonate chemistry. American Zoologist39: 160–183.

Geider RJ, Osborne BA. 1992. Algal photosynthesis: the measurement of algal gasexchange. London, UK: Chapman and Hall.

Giordano M, Beardall J, Raven J. 2005. CO2 concentrating mechanisms in algae:mechanisms, environmental modulation, and evolution. Annual Review ofPlant Biology 56: 99–131.

Goiran C, Al-Moghrabi S, Allemand D, Jaubert J. 1996. Inorganic carbonuptake for photosynthesis by the symbiotic coral–dinoflagellate association II.Bicarbonate absorption by animal host. Journal of Experimental Marine Biologyand Ecology 199: 207–225.

Guinotte JM, Fabry VJ. 2008.Ocean acidification and its potential effects onmarine ecosystems. Annals NY Academy of Sciences 1134: 320–342.

Hennige S, Suggett DJ, Warner ME, McDougall K, Smith DJ. 2009.Photobiology of Symbiodinium revisited: bio-physical and bio-opticalsignatures. Coral Reefs 28: 179–195.

Hurd C, Hepburn C, Currie K, Raven J, Hunter K. 2009. Testing the effects ofocean acidification on algal metabolism: considerations for experimentaldesigns. Journal of Phycology 45: 1236–1251.



Iglesias-Prieto R, Beltr#an V, Lajeunesse T, Reyes-Bonilla H, Thom#e P. 2004.Different algal symbionts explain the vertical distribution of dominant reefcorals in the Eastern Pacific. Proceedings of the Royal Society B: Biological Sciences271: 1757–1763.

Johnson KS. 1982. Carbon dioxide hydration and dehydration kinetics inseawater. Limnology and Oceanography 27: 849–855.

LaJeunesse TC. 2001. Investigating the biodiversity, ecology, and phylogeny ofendosymbiotic dinoflagellates in the genus Symbiodinium using the ITS region:in search of a “species” level marker. Journal of Phycology 37: 866–880.

Leggat W, Badger M, Yellowlees D. 1999. Evidence for an inorganic carbon-concentrating mechanism in the symbiotic dinoflagellate Symbiodinium sp.Plant Physiology 121: 1247–1255.

Leggat W, Marendy EM, Baillie B, Whitney SM, Lugwig M, Badger MR,Yellowlees D. 2002. Dinoflagellate symbioses: strategies and adaptations forthe acquisition and fixation of inorganic carbon. Functional Plant Biology 29:309–322.

Lesser MP. 1996. Exposure of symbiotic dinoflagellates to elevated temperaturesand ultraviolet radiation causes oxidative stress and inhibits photosynthesis.Limnology and Oceanography 41: 271–283.

Lilley RM, Ralph PJ, Larkum AWD. 2010. The determination of activity of theenzyme Rubisco in cell extracts of the dinoflagellate alga Symbiodinium sp. bymanganese chemiluminescence and its response to short-term thermal stress ofthe alga. Plant, Cell & Environment 33: 995–1004.

MacIntyre HL, Sharkey TD, Geider RJ. 1997. Activation and deactivation ofribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) in three marinemicroalgae. Photosynthesis Research 51: 93–106.

Mackey KRM, Paytan A, Grossman AR, Bailey S. 2008. A photosyntheticstrategy for coping in a high-light, low-nutrient environment. Limnology andOceanography 53: 900–913.

Martin CL, Tortell PD. 2006. Bicarbonate transport and extracellular carbonicanhydrase activity in Bering Sea phytoplankton assemblages: results fromisotopic disequilibrium experiments. Limnology and Oceanography 51: 2111–2121.

McCabe-Reynolds J, Bruns BU, Fitt WK, Schmidt GW. 2008. Enhancedphotoprotection pathways in symbiotic dinoflagellates of shallow-water coralsand other cnidarians. Proceedings of the National Academy of Sciences, USA 105:13673–13678.

McGinley MP, Aschaffenburg MD, Pettay DT, Smith RT, LaJeunesse TC,Warner ME. 2012. Transcriptional response of two core photosystemgenes in Symbiodinium spp. exposed to thermal stress. PLoS ONE 7: e50439.

Millero F, Graham T, Huang F, Bustos-Serrano H, Pierrot D. 2006.Dissociation constants of carbonic acid in seawater as a function of salinity andtemperature.Marine Chemistry 100: 80–94.

Moore CM, Suggett DJ, Hickman AE, Kim YN, Tweddle JF, Sharples J, GeiderRJ, Holligan PM. 2006. Phytoplankton photoacclimation andphotoadaptation in response to environmental gradients in a shelf sea.Limnology and Oceanography 51: 936–949.

Morse D, Salois P, Markovic P, Hastings JW. 1995. A nuclear-encoded form IIRubisco in dinoflagellates. Science 268: 1622–1624.

Muscatine L, Goiran C, Land L, Jaubert J, Cuif JP, Allemand D. 2005. Stableisotopes (dC-13 and dN-15) of organic matrix from coral skeleton. Proceedingsof the National Academy of Sciences, USA 102: 1525–1530.

Niyogi KK. 2000. Safety valves for photosynthesis. Current Opinion in Biology 3:455–460.

Ort DR, Baker NR. 2002. A photoprotective role for O2 as an alternativeelectron sink in photosynthesis? Current Opinion in Plant Biology 5:193–198.

Pierrot D, Lewis E, Wallace D. 2006.MS Excel program developed for CO2

system calculations. ORNL/CDIAC-105. [WWW document] URL http://cdiac.ornl.gov/oceans/co2rprt.html [accessed on 1 February 2011].

Platt T, Gallegos C, Harrison W. 1980. Photoinhibition of photosynthesis innatural assemblages of marine phytoplankton. Journal of Marine Research 38:687–701.

Portis AR, Li C, Wang D, Salvucci ME. 2008. Regulation of Rubisco activase andits interaction with Rubisco. Journal of Experimental Botany 59: 1597–1604.

Provasoli L, McLaughlin J, Droop M. 1957. The development of artificial mediafor marine algae. Archives of Microbiology 25: 392–428.

Ragni M, Airs R, Hennige S, Suggett DJ, Warner ME, Geider RJ. 2010. PSIIphotoinhibition and photorepair in Symbiodinium (Pyrrhophyta) differsbetween thermally tolerant and sensitive phylotypes.Marine Ecology ProgressSeries 406: 57–70.

Reinfelder JR. 2011. Carbon concentrating mechanisms in eukaryotic marinephytoplankton. Annual Review of Marine Science 3: 291–315.

Ritchie R. 2006. Consistent sets of spectrophotometric chlorophyll equations foracetone, methanol and ethanol solvents. Photosynthesis Research 89: 27–41.

Robison J, Warner ME. 2006. Differential impacts of photoacclimation andthermal stress on the photobiology of four different phylotypes ofSymbiodinium (Pyrrhophyta). Journal of Phycology 42: 568–579.

Rost B, Kranz SA, Richter KU, Tortell PD. 2007. Isotope disequilibrium andmass spectrometric studies of inorganic carbon acquisition by phytoplankton.Limnology and Oceanography: Methods 5: 328–337.

Rost B, Zondervan I, Wolf-Gladrow D. 2008. Sensitivity of phytoplankton tofuture changes in ocean carbonate chemistry: current knowledge, contradictionsand research directions.Marine Ecology Progress Series 373: 227–237.

Ryan-Keogh TJ, Macey AI, Cockshutt AM, Moore CM, Bibby TC. 2012. Thecyanobacterial chlorophyll-binding-protein IsiA acts to increase the in vivoeffective absorption cross-section of PSI under iron limitation. Journal ofPhycology 48: 145–154.

Sch!agger H. 2006. Tricine-SDS-PAGE. Nature Protocols 1: 16–22.Solomon S, Qin D, Manning M, Chen Z, Marquis M, Averyt KB, Tignor M,Miller HL, eds. 2007. Climate change 2007: the physical science basis.Contribution of Working Group I to the Fourth Assessment Report of theIntergovernmental Panel on Climate Change. Cambridge University Press, UK.

Suggett DJ, MacInytre HL, Kana TM, Geider RJ. 2009. Comparing electrontransport with gas exchange: parameterising exchange rates between alternativephotosynthetic currencies for eukaryotic phytoplankton. Aquatic MicrobialEcology 56: 147–162.

Suggett DJ, Warner ME, Smith DJ, Davey P, Hennige S, Baker NR. 2008.Photosynthesis and production of hydrogen peroxide by Symbiodinium(Pyrrhophyta) phylotypes with different thermal tolerances. Journal of Phycology44: 948–956.

Tabita FR, Satagopan S, Hanson TE, Kreel NE, Scott SS. 2008. Distinct form I,II, III, and IV Rubisco proteins from the three kingdoms of life provide cluesabout Rubisco evolution and structure/function relationships. Journal ofExperimental Botany 59: 1515–1524.

Tortell P. 2000. Evolutionary and ecological perspectives on carbon acquisitionin phytoplankton. Limnology and Oceanography 45: 744–750.

Tortell PD, Trimborn S, Li Y, Rost B, Payne CD. 2010. Inorganic carbonutilization by Ross Sea phytoplankton across natural and experimental CO2

gradients. Journal of Phycology 46: 433–443.Venn AA, Tambutt#e E, Lotto S, Zoccola D, Allemand D, Tambutt#e S. 2009.Imaging intracellular pH in a reef coral and symbiotic anemone. Proceedings ofthe National Academy of Sciences, USA 106: 16574–16579.

Whitney S, Shaw D, Yellowlees D. 1995. Evidence that some dinoflagellatescontain a ribulose-1,5-bisphosphate carboxylase/oxygenase related to that ofthe a-proteobacteria. Proceedings of the Royal Society B: Biological Sciences 259:271–275.

Wooldridge SA. 2009. A new conceptual model for the warm-water breakdownof the coral-algae endosymbiosisMarine and Freshwater Research 60: 483–496.

Wooldridge SA. 2010. Is the coral-algae symbiosis really ‘mutually beneficial’ forthe partners? BioEssays 32: 615–625.

Whitney SM, Andrews TJ. 1998. The CO2/O2 specificity of single-subunitribulose-bisphosphate carboxylase from the dinoflagellate, Amphidiniumcarterae. Australian Journal of Plant Physiology 25: 131–138.

Yellowlees D, Dionisio-Sese ML, Masuda K, Maruyama T, Abe T, Baillie B,Tsuzuki M, Miyachi S. 1993. Role of carbonic anhydrase in the supply ofinorganic carbon to the giant clam–zooxanthellate symbiosis.Marine Biology115: 605–611.

Supporting Information

Additional supporting information may be found in the onlineversion of this article.




Methods S1 Determination of light-dependent electron transferrates (ETRs) using fast repetition rate fluorometry (FRRF).

Methods S2 Determination of light-dependent carbon fixationusing radioisotopic carbon-14.

Methods S3 Determination of total carbon dioxide (TCO2)-dependent carbon fixation using radioisotopic carbon-14.

Methods S4 Determination of inorganic carbon species prefer-ence and external carbonic anhydrase (eCA) activity using isotopedisequilibrium.

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Contrasting modes of inorganic carbon acquisition amongst Symbiodinium (Dinophyceae) phylotypes

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