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 how marine microalgae acquire and utilize inorganic carbon (iC) to support 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, in turn, may affect the productivity and growth of marine microalgae (Fu et al., 2008; Brading et al., 2011). Understanding the mechanisms of iC acquisition is especially relevant for algae belonging to the dinoflagellate genus Symbiodinium, which are found both free-living and in symbiosis with reef-building corals and other cnidarians. This symbiosis is essential for the viability of many cnidaria, and is key to the persistence of coral reef ecosystems under present day and future climates (e.g. Iglesias-Prieto et al., 2004; Berkelmans & van Oppen, 2006). However, our current understanding of the iC environment within the host tissue is limited (Gattuso et al., 1999; Venn et al., 2009), despite the fact that it may have a significant impact on the productivity and growth of the in hospite Symbiodinium population (Wooldridge, 2009, 2010). The Symbiodinium genus is taxonomically divided into numerous phylotypes, physiologically distinct types that are distinguishable by genetic variability in the second internal transcribed spacer region (ITS2) of their ribosomal DNA (LaJeunesse, 2001). Different phylotypes can exhibit a range of responses 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 their coral hosts to occupy a wide range of niches (e.g. Iglesias-Prieto et al., 2004).
Unlike inorganic macronutrients, such as nitrogen and phosphorus, iC within ocean surface waters is relatively abundant (and constant) at c. 2.2 mM. However, most of this iC is in the form of bicarbonate ions (), whereas aqueous carbon dioxide (CO2(aq)), the actual iC species fixed within the CBC, contributes < 1%. Rubisco, the enzyme employed by marine microalgae to catalyse iC fixation, has a relatively low CO2 affinity and specificity, particularly within an O2-rich environment. Consequently, this key enzyme is, in fact, considerably undersaturated with respect to current seawater concentrations of CO2(aq). This is particularly true of dinoflagellates, such as Symbiodinium, which possess a form II Rubisco (Morse et al., 1995; Whitney et al., 1995) that has a significantly lower specificity for CO2 than the form I Rubisco of other phytoplankton taxa (e.g. Tortell, 2000).
In order to overcome the inherent enzymatic limitations of Rubisco, nearly all marine microalgae have evolved strategies, termed ‘carbon-concentrating mechanisms’ (CCMs), to actively elevate CO2 at the active site of Rubisco (Giordano et al., 2005; Reinfelder, 2011); thus, the majority of marine primary producers are not considered to be C limited under present day conditions (Giordano et al., 2005). CCMs not only allow marine microalgae to efficiently utilize the small and variable pool of CO2(aq) for C fixation, but also take advantage of the much larger fraction. The functional role of CCMs has been extensively reviewed within the literature (see Giordano et al., 2005; Reinfelder, 2011), and primarily involves one or more of the following processes: (1) the active transport of CO2(aq) into the cell; (2) the dehydration of to CO2(aq) at the cell surface, which is then transported into the cell; and (3) the active transport of directly into the cell, where it is then converted to CO2(aq). Interconversion between and CO2(aq), both outside and inside the cell, is catalysed by external and internal carbonic anhydrases (eCA and iCA, respectively) (Reinfelder, 2011), and the dehydration of to CO2(aq) at the cell's boundary layer by eCA can be further aided by membrane-bound proton (H+) pumps that acidify the immediate external environment (Bertucci et al., 2010).
A number of CCM components have been identified for Symbiodinium, although no direct comparison of iC acquisition across different phylotypes has been performed within a single study. Both eCAs and iCAs have been shown to be expressed by Symbiodinium, as well as H+-ATPase (a proton pump) and Na+/ co-transporters (Yellowlees et al., 1993; Al-Moghrabi et al., 1996; Leggat et al., 1999; Bertucci et al., 2010). It is also apparent that the expression of these mechanisms differs depending on the host type (e.g. giant clam vs coral) and whether the Symbiodinium is in hospite or free-living (Al-Moghrabi et al., 1996; Leggat et al., 1999; Bertucci et al., 2010). Although these various studies have demonstrated that the environment can play an important role in the type of CCM expressed, few have unfortunately specified the phylotype studied, with Bertucci et al. (2010) being the only exception. This has made it impossible to infer whether differences in CCM type previously described for Symbiodinium are solely driven by environment (i.e. acclimation to different iC conditions/availability) or also by adaptive differences between phylotypes.
It is widely accepted that the changes in productivity and growth under OA conditions represent ‘downstream effects’ on iC acquisition and fixation (Rost et al., 2008; Hurd et al., 2009). Recently, we have demonstrated that the effect of OA on free-living Symbiodinium is phylotype specific, indicating that key differences in iC acquisition may also exist between phylotypes 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 increase in the partial pressure of CO2 (pCO2) from c. 390 to 800 ppmv. Specifically, an increase in productivity, with no change in growth rate, was observed for A20, whereas the opposite was observed for A13. Based on these previous observations, we sought to examine these alternative physiological responses to elevated iC availability (productivity vs growth), and hypothesized that: (1) A20, but not A13, is carbon limited under present day conditions because of an inefficient CCM; and thus (2) A13 and A20 express different CCM types when grown in the same iC environment.
In order to evaluate the uptake and utilization of iC, we first determined the light-dependent rates of productivity using two approaches. Light-dependent rates of C fixation and photosynthetic electron transfer (PET) were measured so as to compare the efficiency with which energy (and reductant), derived from PET, is utilized for C fixation vs other essential metabolic processes, such as CCM/CA activity. We subsequently assessed iC affinity and species preference by quantifying the Rubisco concentration, 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
For this study, we focused on two Symbiodinium isolates within clade A: A13 and A20. Phylotype A20 (accession number EU449053) has yet to be identified from an intact symbiosis and is putatively considered to be a free-living Symbiodinium (M. E. Warner, pers. obs.). The photophysiology of both isolates has been studied in relation to photoacclimation (Hennige et al., 2009) and response to OA (Brading et al., 2011). Both phylotypes exhibit marked differences in their physiological responses to light and pCO2, and recent work has shown A13 to be much more susceptible to photoinactivation as a result of thermal stress (McGinley et al., 2012). These two phylotypes are therefore good candidates to explore adaptive differences in iC acquisition between closely related Symbiodinium species (see also Brading et al., 2011).
Continuous cultures (volume, 3 l) of Symbiodinium phylotypes A13 and A20 were grown at 26°C under a growth irradiance of c. 350 μmol 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-μm pore-sized filter (Polycap 36AS; Whatman Filters, GE Healthcare Biosciences, Pittsburgh, PA, USA), at a rate matching the growth rates of each culture, thereby maintaining a constant cell biomass. The medium was modified to exclude the tris(hydroxymethyl)aminomethane buffer and adjusted to a total carbon alkalinity (TAC) of c. 2300 μEq kg−1 seawater (see Brading et al., 2011). Cultures were subjected to gentle stirring to minimize the settling of cells and to ensure thorough mixing of the medium with the gas inflow. Both phylotypes were grown at pCO2 levels representative of ‘present day’ conditions, that is, 380–390 ppmv (Table 1; Solomon et al., 2007). The growth pCO2 was maintained using the ‘pH-stat’ culturing methods described in Brading et al. (2011). Cultures were allowed to acclimate to their pCO2 growth regimes for a period of 21 d (equivalent to at least nine generations), and were only sampled once steady-state growth was achieved. The cultures were sampled approximately an hour after ‘lights-on’, and all measurements were completed before midday. Where certain measurements could not be made on both cultures simultaneously, the order in which they were sampled was reversed between replicate measurements (n = 4). Replicate measurements from the same cultures were separated by a period of 1 wk (c. three complete cell generations).
Table 1. The mean (± SE) carbon chemistry of the media in which the two Symbiodinium phylotypes were grown (6 wk). All calculations were performed with the CO2SYS program developed by Pierrot et al. (2006) with the constants taken from Millero et al. (2006)
|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)|
Daily cell counts were obtained microscopically with a Neubauer haemocytometer (Fisher Scientific, Loughborough, UK) to monitor the growth rates of the cultures. As a result of the nature of continuous culturing, changes in cell number were further corrected for the dilution rate and volumes of sample removed from the culture between cell counts to provide a true measure of growth rate (Eqn 1).
(t, time (d); Δt, time interval between sampling; C, concentration of cells (cells ml−1); V, volume of medium (ml) in the culture vessel at a specific time of sampling (subscript ‘t’ or ‘t+Δt’), the waste outflow generated by the continuous dilution of the culture during the time period between sampling (subscript ‘waste’) and the culture vessel when maximally filled (subscript ‘vessel’)). The first term, , describes the change in cell number for the total population during the sampling interval, and the second term, , describes the dilution rate. The specific growth rate (μ, d−1) was calculated as the change in cell concentration per day (d−1). Following the daily cell count, the dilution rate for each culture vessel was adjusted in order to maintain a constant cell density within each culture. Cell densities of A13 and A20 were 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 filtered through glass fibre filters (GF/F). Each filter was snap frozen in liquid nitrogen and stored at −80°C for later extraction and analysis. Each filter was ground in 5 ml of 100% methanol and stored at −20°C for 24 h. The samples were then centrifuged at 3400 g for 5 min. The absorption of the supernatants between 600 and 760 nm was read using a spectrophotometer (U-3000; Hitachi High Technologies, Krefeld, Germany) against a reference cuvette containing deionized H2O (dH2O). Absorbance spectra were then corrected for a baseline shift by subtracting a methanol-only ‘blank’ spectrum, also read against a dH2O reference, and the slope in the spectrum between 750 and 760 nm. The Chla concentration was then quantified using the coefficients and equations of Ritchie (2006), and finally normalized to the cell density to provide units of picogram Chla per cell.
Particulate organic carbon (POC)
Aliquots of 100 ml were filtered onto pre-combusted GF/F filters, dried at room temperature and then stored at −80°C for subsequent analysis for cellular POC content. Inorganic carbonates were removed from the filters by acidification with c. 15% hydrochloric acid. After acidification, each filter was dried at 60°C for 48 h. Analysis was performed on the solid module of a Total Organic Carbon Analyser (Shimadzu, Milton Keynes, UK) against a glucose calibration standard. Once each sample was sealed within the analyser, the carrier gas was allowed to circulate for up to 2 min before the sample was pushed into the furnace to maximize the signal-to-noise ratio; this period ensured that the carrier gas circulating through the furnace and module was sufficiently purged of CO2 before the analysis of the filter. The POC concentration was then normalized to the cell density to give units of pg C per cell.
Aliquots of 100 ml were filtered onto GF/F filters, immediately snap frozen in liquid nitrogen and stored at −80°C for later analysis of Rubisco content following the protocol described by Ryan-Keogh et al. (2012). Briefly, the protein was extracted from the samples using a freeze–thaw cycle in liquid nitrogen and in combination with sonication in order to fully rupture the cell membranes. The total protein concentration was then measured using the Lowry protein assay, and the Rubisco concentration was subsequently determined by quantitative Western blotting using the Tricine-sodium dodecylsulfate-polyacrylamide gel electrophoresis (Tricine-SDS-PAGE) method of Schägger (2006); antibodies and protein standards were used to quantify the gene RbcL (Agrisera AB, Vännäs, Sweden), which codes for the large subunit of Rubisco, according to Brown et al. (2008). SDS-PAGE and membrane transfer were performed on an Invitrogen Life Technologies PowerEase 500™ (Invitrogen, Paisley, Renfrewshire, UK). Imaging of the membrane was performed on a BioRad VersaDoc imaging system (Bio-Rad, Hemel Hempstead, UK); these images were subsequently processed using QuantityOne software to quantify protein samples and standards, with all quantification performed within the unsaturated part of the calibration curve. Quantified RbcL was normalized to Chla, which was measured on subsamples of the protein extract using a calibrated Turner Designs 10-AU fluorometer (Turner Designs, Sunnyvale, CA, USA), to give units of mmol RbcL mol−1 Chla.
Light-dependent electron transfer rates (ETRs)
A FASTtracka II fluorometer, attached to a FASTact laboratory system (Chelsea Instruments, London, UK), was used to yield photosystem II electron transfer rates (ETRPSII) via a fluorescence light response curve (see Supporting Information Methods S1). Values of ETRPSII (mol e− g−1 Chla h−1) for each light step were calculated following Suggett et al. (2008) as:
(E, light intensity (μmol photons m−2 s−1); σPSII, effective absorption cross-section of PSII (nm2 quantum−1); nPSII, concentration of PSII reaction centres (mol RCII g−1 Chla)). Fv/Fm and are both dimensionless and were calculated as Fv/Fm = (Fm – F0)/Fm and , respectfully. The conversion factor 6.023 × 105 transforms the units of σPSII to m2 RCII−1, whereas 3600 × 10−6 transforms the units of E to mol photons m−2 h−1. σPSII is weighted to the spectral quality of the light used to induce the fluorescence transient, that is, blue LED array, and was therefore spectrally corrected to match the spectrum of the white LED of the actinic light source:
Values of were calculated according to the methods described by Brading et al. (2011), and were 0.374 and 0.420 for A13 and A20, respectively. Values of nPSII were not measured, but assumed according to previous values obtained for these strains (grown at similar light intensity) using the O2-flash yield approach (Hennige et al., 2009): 2.31 × 10−6 mol RCII g−1 Chla and 2.50 × 10−6 mol RCII g−1 Chla for A13 and A20, respectively.
In order to calculate the light-limited electron transfer efficiency (initial slope, αETR) and the light-saturated ETR (maximum rate of electron transfer, ETRmax) from each of the rapid-light response curves performed, a modified version of the model by Platt et al. (1980) was fitted to the data using SigmaPlot 10.0 curve-fitting software:
where and β describes the reduction in photosynthetic rate at high irradiance (photoinhibition). The saturation irradiance for electron transfer, termed EkETR, was calculated as (ETRmax/αETR) with units of μmol photons m−2 s−1.
Measurements of radioisotopic 14C uptake, specifically 14C-labelled sodium bicarbonate (NaH14CO3) with a specific activity of 40–60 mCi mmol−1 (Perkin Elmer, Norwalk, CT, USA), were applied to three methods, each assaying different aspects of iC acquisition. In all cases, fixed 14C was measured on a Tri-Card 2910 TR liquid scintillation analyser (Perkin Elmer). The disintegrations per minute (DPM) for each sample were averaged from a single 20-min count and the rate of C fixation (μmol C l−1 h−1) was calculated as follows:
where the superscripts ‘sample’, ‘T0’ and ‘TC’ refer to the sample, the initial measurement at the start of an incubation and the total activity count, respectively. [TCO2] is the concentration of iC (μM) within the sample and t is the incubation in hours.
Light-dependent C fixation
The photosynthetic light response of each culture was characterized by incubating discrete samples, spiked with 14C radioisotope, at a range of light intensities from c. 10 to 1330 μmol photons m−2 s−1 (see Methods S2). The amount of fixed 14C at each photon flux density (PFD) was then measured according to standard liquid scintillation procedures and 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, α14C) and the light-saturated photosynthetic capacity (maximum rate of C fixation, Pmax) of gross C uptake from each light curve. Pmax (mol C l−1 h−1) and α14C ((mol C l−1 h−1)(μmol photons m−2 s−1)−1) were normalized to the Chla concentration of the sample. The saturation irradiance for C fixation, termed Ek14C, was calculated as (Pmax/α14C) with units of μmol photons m−2 s−1.
TCO2-dependent C fixation
The photosynthetic TCO2 response was also characterized for each culture between zero and 5000 μM TCO2 (see Methods S3). The TCO2-saturated rate of photosynthesis (maximum velocity of C fixation, Vmax) and the TCO2 concentration at which the rate of photosynthesis is half that of Vmax (half-saturation concentration, Km(TCO2)) were calculated by fitting a Michaelis–Menton model to the data, using SigmaPlot 10.0 curve-fitting software, which describes the velocity of C fixation (v) as a function of TCO2:
Values calculated for Km(TCO2) can also be used to obtain the respective half-saturation concentrations of CO2(aq) for C fixation by determining the relative speciation of the iC pool at pH 8.0 and salinity 35 at Km(TCO2). This provides an indication as to what the intracellular CO2(aq) concentration would be in the absence of active uptake (i.e. passive diffusion only) when C fixation is not carbon limited, and allows for comparison with the theoretical Km(CO2) of Symbiodinium Rubisco (see the 'Discussion' section).
iC species preference and eCA activity
Isotope disequilibrium (Espie & Colman, 1986) was used to quantify the relative contributions of CO2 and to iC uptake, following the modification to the technique proposed by Martin & Tortell (2006) (see Methods S4). In order to assess the presence and activity of eCA, duplicate samples, pretreated with the membrane-impermeable CA inhibitor acetazolamide (AZ), were also run according to this protocol. To determine the relative contributions of CO2 and to carbon uptake, data from both the control and +AZ treatments were analysed according to the model and protocol described by Martin & Tortell (2006), fitted using SigmaPlot 10.0 curve-fitting software:
(Vt, total rate of C uptake; f, fraction of C uptake supported by direct transport of into the cell; α1 and α2, first-order rate constants for CO2 and hydration and dehydration, respectively, which are temperature, salinity and pH dependent). In the absence of enzymatic activity, α1 and α2 were determined to be 0.0894 and 0.1135 s−1, respectively (using the equations of Espie & Colman, 1986), with corrections for temperature and salinity (Johnson, 1982). SADIC is the specific activity of all the iC species at equilibrium, and ΔSACO2 and ΔSAHCO3- are the differences between the initial (i.e. at pH 7.0) and equilibrium (i.e. at pH 8.5) values of the specific activities of CO2 and . The values of ΔSACO2/SADIC and ΔSAHCO3-/SADIC were calculated using the difference in pH between 14C spike and buffered ASP-8A, as well as TCO2 of the buffered ASP-8A.
In the +AZ treatments, the rate constants α1 and α2 were assumed to equal the uncatalysed value, as any potential eCA activity is blocked by the presence of the inhibitor. Therefore, in the +AZ treatments, the model described in Eqn 7 was fitted to the data by allowing Vt and f to vary and restraining the other parameters to the values described previously. For the control treatments, the model was fitted by restraining f to the value calculated in the corresponding +AZ model fit and, instead, allowing α1 to vary as the model parameter (with α2 expressed as a function of α1 rather than as an actual model parameter; see Espie & Colman, 1986), but constrained to be equal to or greater than the uncatalysed value. The change in α1 between the control treatment and uncatalysed value of the +AZ treatment can then be used to describe the eCA activity employing the following equation (Rost et al., 2007):
The growth rate, cellular Chla content, parameters relating to light-dependent C fixation and iC affinity, and the iC species preference and eCA activity under ambient pCO2 (i.e. c. 380 ppmv) were compared between the two phylotypes using a parametric-independent samples t-test or a non-parametric Mann–Whitney test where appropriate. The threshold of statistical significance was set at P < 0.05. All statistical tests were performed using SPSS statistical analysis software (version 16.0; SPSS, Portsmouth, UK).
Rates of C fixation were similar between the two phylotypes and agree 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, little difference was observed in the coupling of PET and C fixation between the two phylotypes at both low and high light conditions (i.e. αETR : α14C and ETRmax : Pmax, respectively). These comparisons of ETR with rates of C fixation provide an indication of energy investment within the cell. In the present study, the comparison of light-limited ETR with C fixation (αETR : α14C) yielded an average ratio of c. 4.9 mol e− mol C−1, which agrees well with the theoretical minimum PET : C fixation ratio of 5 (Suggett et al., 2009). Under light-limiting conditions, therefore, PET in PSII is closely coupled to the C fixation occurring in the CBC for both 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 saturated before PET, as indicated by the EkETR : Ek14C ratio of 2.
A similar observation of C fixation saturating at lower irradiances than PET (i.e. Ek14C < EkETR) was also reported for the cyanobacterial species Synechococcus, whereby ETR continued to increase with irradiance after the saturation of C fixation (Bailey et al., 2008), and is also consistent with the notion, observed for many taxa across pelagic systems, that the rate of electron transfer from the primary quinone acceptor to plastoquinone (PQ) (1/τQB→PQ) typically exceeds the rate of whole-chain electron transfer (1/τPSII) (Moore et al., 2006). Such a pattern indicates the occurrence of alternative electron sinks downstream of PSII, but before their use in C fixation (Suggett et al., 2009). Therefore, our observations for the two phylotypes of Symbiodinium studied here suggest the existence of alternative electron sinks under high irradiances that exceed what is necessary to support C fixation.
In Brading et al. (2011), we demonstrated that a significant amount of light-enhanced O2 consumption occurs in both A13 and A20. The pathways responsible for this, such as photorespiration and Mehler-Ascorbate-Peroxidase (MAP) activity, act as electron 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). MAP and other alternative oxygen-consuming pathways have been observed previously in Symbiodinium under high-light stress, and specifically within clade A phylotypes (McCabe-Reynolds et al., 2008; Suggett et al., 2008), as well as in other dinoflagellate species, such as Prorocentrum minimum (Suggett et al., 2009). Cyclic electron transfer (CET) in PSI may also contribute to the decoupling of ETR from C fixation in clade A Symbiodinium phylotypes, acting as an alternative sink for electrons, as well as an additional source of ATP to linear PET (McCabe-Reynolds et al., 2008). It is not possible in the present study to determine which of these processes is responsible for the observed decoupling between PET and C fixation under high light. However, our observations provide the first quantitative means by which fluorescence-based measures of Symbiodinium productivity could better evaluate C fixation, and demonstrate that coupling between PET and C fixation is primarily driven by irradiance and is independent of phylotype.
To our knowledge, the quantification of form II Rubisco in Symbiodinium spp. has never been published previously. Form II Rubisco is notoriously unstable once extracted (Whitney et al., 1995), and is therefore difficult to accurately quantify. The method employed in the present study, adapted from Brown et al. (2008), provides the first ever quantification of Symbiodinium Rubisco concentration. The concentration of form II Rubisco in phylotypes A13 and A20 (1.9 and 0.5 mmol RbcL mol−1 Chla, respectively) was of a similar order of magnitude to, but lower than, the concentrations of form I Rubisco (3.6–8.2 mmol RbcL mol−1 Chla) reported previously for a number of cyanobacteria species, including Trichodesmium spp. and Synechococcus elongatus (Brown et al., 2008). It is unsurprising that the concentration of Rubisco is greater in species of cyanobacteria, given that their form I Rubisco consists of eight large subunits, whereas form II Rubisco of dinoflagellates can have as few as two large subunits (Tabita et al., 2008).
Despite having an almost four-fold greater concentration of Rubisco than A20, this did not translate to a greater maximum capacity for C fixation in A13. When expressed relative to Rubisco, values of Vmax for A13 were considerably lower than those for A20, suggesting that C fixation at saturating concentrations of iC is not proportional to the size of the Rubisco pool. The lower Rubisco-normalized Vmax of A13 may be a consequence of insufficient CO2(aq) elevation, relative to O2, at the active site of Rubisco or, alternatively, a lower Rubisco specificity (τ). Taxonomic differences in τ are well established, and some degree of variability for τ within taxonomic groups has also been shown, although not within dinoflagellates (reviewed in Tortell, 2000).
Alternatively, it may be that, although A13 has the larger pool of 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 Rubisco activase, a soluble protein that facilitates the carbamylation of Rubisco, and CA1P, an inhibitor synthesized in the dark and broken down in the light (Portis, 2003). The role of Rubisco activase is to balance C fixation with the supply of ATP and reductant from the light reactions of the thylakoid membrane (Portis, 2003). In the case of Symbiodinium phylotype A13, the large pool of Rubisco, although seemingly unnecessary, may be a strategy used by this isolate to optimize carboxylation, activating and deactivating the enzyme in response to changing supplies of iC, ATP and reductant. However, the actual presence of a Rubisco activase 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 Symbiodinium Rubisco has been published to date, Leggat et al. (2002) approximated a value of 50–60 μM CO2 by assuming a τ value similar to that for form II Rubisco of Amphidinium carterae (i.e. τ = 37) (Whitney & Andrews, 1998) and similar kinetic properties to the form II Rubisco of the proteobacterium Rhodospirillum rubrum. In the present study, Km(CO2) for C fixation was c. 4.7 μM for A13 and c. 7.3 μM for A20. If both phylotypes relied solely on passive diffusion alone, this concentration of CO2(aq) would be considerably lower than that required to saturate Rubisco and support maximal C fixation. Such a consideration provides evidence that active processes (i.e. CCMs) must be functioning in A13 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) supplementing/replacing the uptake of CO2(aq) with .
iC affinity and species preference
As already mentioned, evidence of an active CCM in Symbiodinium has been published previously, but the exact nature of the mechanisms utilized sometimes appears to conflict between studies, and it is unclear whether these differences are taxonomically vs environmentally driven (Al-Moghrabi et al., 1996; Goiran et al., 1996; Leggat et al., 1999). Symbiodinium recently isolated from corals has been shown to preferentially transport directly into the cell (Al-Moghrabi et al., 1996; Goiran et al., 1996), whereas that recently isolated from giant clams exhibited a preference for direct CO2 uptake (Leggat et al., 1999). Interestingly, in the latter study, this preference changed to direct uptake after 2 d of being maintained in culture (Leggat et al., 1999). Together, these studies show that the expression of a CCM can be environmentally driven for some phylotypes of Symbiodinium, with some degree of plasticity to respond to changes in the iC environment. Our study confirms that CCMs operate in Symbiodinium, but further shows that the nature of Symbiodinium CCMs are algal specific and are, to some extent, taxonomically driven. A13 and A20 both have iC uptake mechanisms that differ from those described previously for cultured Symbiodinium, further highlighting the physiological diversity within this genus that has been reported previously in relation to other environmental factors, such as light and temperature (Robison & Warner, 2006; Suggett et al., 2008; Hennige et al., 2009; Ragni et al., 2010).
Although a direct comparison of the expression of CCM-related proteins between the phylotypes was not made, the 14C disequilibrium technique does provide an indication as to which iC species was used and whether uptake was direct or aided by eCA activity. In the presence of AZ, and therefore in the absence of any potential eCA activity, neither phylotype appeared to be capable of performing the direct transport of into the cell, but instead were entirely dependent on direct CO2(aq) uptake. Such an observation suggests an absence of proteins capable of transporting , such as the Na+/ co-transporter, which has been proposed previously for a Symbiodinium sp. isolated from Galaxea fascicularis (Al-Moghrabi et al., 1996). The inability to transport directly has also been observed previously in the dinoflagellates A. carterae and Heterocapsa oceanica (Colman et al., 2002; Dason et al., 2004). However, from the data available here, it is not clear whether A13 and A20 lack the genes necessary to build the proteins that facilitate direct uptake, as in A. carterae (Colman et al., 2002), or just do not express them under the growth conditions used for this study.
In the absence of AZ, the calculated rate constant of CO2 hydration in the A20 samples was significantly higher than that of the theoretical uncatalysed rate of 0.0894 s−1, and suggests that eCA was present in A20. Using the equation for eCA activity proposed 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 utilize by first converting it to CO2(aq) using eCA, whereas A13 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 dehydration of 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 to conclusively determine whether a similar mechanism occurs in A20, but the fact that little uptake was observed when eCA was inhibited suggests that A20 lacks any other mechanism to promote dehydration. Phylotype A13, however, does not support any of its C fixation by uptake, either by direct transport or indirectly by first converting it to CO2(aq) with eCA. The apparent lack of eCA activity in phylotype A13 has also been observed in A. carterae and H. oceanica (Colman et al., 2002; Dason et al., 2004), and also agrees with the findings of Leggat et al. (1999), whereby the addition of AZ to a culture of Symbiodinium, isolated from Tridacna gigas, had no effect on the active uptake of iC. Although the phylotype was not specified in this study, it was most likely an A3 isolate, following the work by LaJeunesse (2001), which found that all of the Symbiodinium isolates taken from host species within the Tridacna genus fell into this classification.
Carbon acquisition and OA
In Brading et al. (2011), we showed that A13 and A20 exhibited increased growth rates and productivity (as O2 evolution), respectively, in response to a doubling of growth pCO2 from 380 to 780 ppm. We interpreted these phylotype-specific responses as an indication of CCM down-regulation and reinvestment of cellular energy into growth for A13 and as C limitation in A20 (Brading et al., 2011). In the light of the findings in the present study, we hypothesize that A13 may down-regulate the ATP invested in the active CO2 uptake pathway under OA conditions, afforded by increased availability of CO2(aq) to support greater passive uptake, and utilize this energy in cell division instead.
Having established that A20 is saturated with respect to iC under present day pCO2, an alternative interpretation for its observed increase in productivity under OA conditions, other than C limitation, must be sought. Specifically, A20 showed a 60% increase in O2 evolution at 780 ppm. Assuming a constant photosynthetic quotient (PQ; mol O2; mol CO2), this would infer that C fixation also increased by 60%. However, this assumption is not necessarily valid. An increase in O2 evolution, independent of C fixation, may indicate a reduction in O2-consuming 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 and the product of A20's growth rate and cellular POC concentration. This approach estimates PQ of A20 to be 0.4 at 380 ppm pCO2 and 0.6 at 780 ppm pCO2. Using the direct measurements of C fixation reported in this study, we also arrive at a similar estimate of PQ for A20 under 380 ppm pCO2 of 0.5. Although lower than the theoretical minimum PQ of 1, such instances have been reported previously for marine phytoplankton (e.g. Carpenter & Roenneberg, 1995), and may be a consequence of significant photorespiratory activity (Geider & Osborne, 1992).
If the Rubisco specificity of A20 is considerably low or the concentration of CO2(aq) at the site of Rubisco is not sufficiently elevated, oxygenase activity will be favoured. Elevated CO2(aq), as a result of OA, may therefore suppress this oxygenase activity and consequently increase the PQ of A2 at elevated pCO2. The lack of a significantly increased growth rate, despite the higher rates of productivity observed for A20 under OA conditions (Brading et al., 2011), may be a consequence of continued energetic investment in eCA to support indirect uptake rather than growth. This suggests that the CCM of A20 has limited plasticity to acclimate to changes in the iC environment, such as the increase in pCO2 associated with OA. It would be necessary to study the plasticity of Symbiodinium CCMs under different environmental conditions and, in particular, elevated pCO2 in order to prove this.
The present study has shown that, although A20 has a lower affinity than A13 for iC, C fixation in both phylotypes was saturated with respect to iC under present day pCO2 conditions. Both phylotypes were capable of active CO2(aq) uptake, but A20 was also able to utilize via eCA. This demonstrates that different mechanisms of uptake exist for these two phylotypes and, based on the differential effect of OA on other phylotypes of Symbiodinium (Brading et al., 2011), no doubt further strategies exist within this genus. These differences are adaptively driven and will probably determine a phylotype's response to environmental shifts, such as OA, when free-living, but may also have implications for their productivity when in hospite. The pH and iC environment within the coral host is complex and its interaction with the zooxanthellae is still not fully understood (Venn et al., 2009; and see commentary by Brownlee, 2009).
These adaptations should be considered against the wider phylogenetic diversity of Symbiodinium in order to understand the evolutionary significance of these different CCMs, as they may contribute greatly to the shaping of the diversity of this genus under future environmental conditions. Further study is required to determine whether additional differences in iC acquisition exist in other clades of Symbiodinium, and whether the regulation of the CCM to changes in the iC availability and speciation differs between phylotypes. This study is also the first time that direct comparisons between ETR and C fixation rates have been performed for Symbiodinium, and showed that significant decoupling between these rates occurs under saturating irradiances (i.e. where productivity may become increasingly limited by iC availability; Muscatine et al., 2005), highlighting the dependence on additional electron sinks, such as oxygen-consuming pathways of MAP and chlororespiration.
We thank Dr Phillip Davey and Tania Maynard-Cresswell, at the University of Essex, for technical support during this study, and Dr Mark Moore and Dr Tom Bibby, at the University of Southampton (UK), for performing the Rubisco analysis. This research was made possible through a Natural Environment Research Council (NERC) PhD studentship to P.B. Funding to M.E.W. was provided by the National Science Foundation (CRI-OA 104940).