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Low CO2 results in a rearrangement of carbon metabolism to support C4 photosynthetic carbon assimilation in Thalassiosira pseudonana Adam B. Kustka1, Allen J. Milligan2, Haiyan Zheng3, Ashley M. New1, Colin Gates1, Kay D. Bidle4 and John R. Reinfelder5 1

Earth and Environmental Sciences, Rutgers University, 101 Warren Street, Newark, NJ 07102, USA; 2Department of Botany and Plant Pathology, Oregon State University, 2082 Cordley

Hall, Corvallis, OR 97331, USA; 3Biological Mass Spectrometry Facility, Rutgers University, 174 Frelinghuysen Road, Piscataway, NJ 08854, USA; 4Institute of Marine and Coastal Sciences, Rutgers University, 71 Dudley Road, New Brunswick, NJ 08901, USA; 5Department of Environmental Sciences, Rutgers University, 14 College Farm Road, New Brunswick, NJ 08901, USA

Summary Author for correspondence: Adam B. Kustka Tel: +1 973 353 5509 Email: [email protected] Received: 28 November 2013 Accepted: 28 May 2014

New Phytologist (2014) doi: 10.1111/nph.12926

Key words: C4 metabolism, fatty acid metabolism, glycine decarboxylase, marine diatoms, quantitative proteomics, pentose phosphate pathway, pyruvate phosphate dikinase (PPDK), pyruvate carboxylase.


The mechanisms of carbon concentration in marine diatoms are controversial. At low CO2, decreases in O2 evolution after inhibition of phosphoenolpyruvate carboxylases (PEPCs), and increases in PEPC transcript abundances, have been interpreted as evidence for a C4 mechanism in Thalassiosira pseudonana, but the ascertainment of which proteins are responsible for the subsequent decarboxylation and PEP regeneration steps has been elusive.
We evaluated the responses of T. pseudonana to steady-state differences in CO2 availability, as well as to transient shifts to low CO2, by integrated measurements of photosynthetic parameters, transcript abundances and quantitative proteomics.
On shifts to low CO2, two PEPC transcript abundances increased and then declined on timescales consistent with recoveries of Fv/Fm, non-photochemical quenching (NPQ) and maximum chlorophyll a-specific carbon fixation (Pmax), but transcripts for archetypical decarboxylation enzymes phosphoenolpyruvate carboxykinase (PEPCK) and malic enzyme (ME) did not change. Of 3688 protein abundances measured, 39 were up-regulated under low CO2, including both PEPCs and pyruvate carboxylase (PYC), whereas ME abundance did not change and PEPCK abundance declined.
We propose a closed-loop biochemical model, whereby T. pseudonana produces and subsequently decarboxylates a C4 acid via PEPC2 and PYC, respectively, regenerates phosphoenolpyruvate (PEP) from pyruvate in a pyruvate phosphate dikinase-independent (but glycine decarboxylase (GDC)-dependent) manner, and recuperates photorespiratory CO2 as oxaloacetate (OAA).

Introduction Ambient CO2 levels in seawater are vastly under-saturating for carboxylase activity by ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO). This leads to the possibility of significant oxygenase activity, which can compete with carboxylation (Cooper et al., 1969; Badger et al., 1998), and results in photorespiration at the expense of carbon fixation (Riebesell et al., 1993; Ogren, 1994). To maintain high rates of photosynthesis with these constraints, marine diatoms have evolved a CO2-concentrating mechanism, or CCM (Rotatore et al., 1995; Fielding et al., 1998; Burkhardt et al., 2001; Colman et al., 2002; Tortell et al., 2002), the underlying biochemistry of which remains uncertain (Reinfelder et al., 2004; Roberts et al., 2007). CO2 concentrations can be elevated in the proximity of RuBisCO through means commonly referred to as biophysical or biochemical. The biophysical pathway relies on an increased intracellular Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

bicarbonate concentration (through active transport), followed by conversion to CO2 (by carbonic anhydrase) in the vicinity of RuBisCO, and is well documented in cyanobacteria and green algae (Amoroso et al., 1998; Badger et al., 2006). Alternatively, CO2 can be concentrated biochemically through single-cell C4 photosynthesis (Reinfelder et al., 2000; Roberts et al., 2007; McGinn & Morel, 2008), although it is important to recognize that a biochemical CCM does not preclude the utility of pumping bicarbonate into the cell. In the most general terms, C4 photosynthesis is characterized by the fixation of bicarbonate and phosphoenolpyruvate (PEP) by PEP carboxylase (PEPC) to form a C4 compound in one ‘compartment’, followed by transport of the C4 compound or its derivative to other compartments for subsequent decarboxylation. These steps serve to fix CO2 and subsequently decarboxylate a C4 acid in close proximity to RuBisCO, and are segregated to avoid futile cycling between C3 and C4 compounds. This New Phytologist (2014) 1 www.newphytologist.com

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segregation was first (Hatch & Slack, 1966) and is most often (Sage, 2004) described among different types of cells in multicellular, vascular plant systems, but is also achieved by organelles within individual cells of multicellular plants (Sage, 2002; Voznesenskaya et al., 2002; Edwards et al., 2004; Bowes, 2011). The C4 pathway originated with the appropriation and regulatory modification of existing carbon metabolism enzymes, often followed by the selection for kinetic properties more suitable for C4 metabolism (Svensson et al., 2003; and references therein). Because there have been multiple and evolutionarily independent origins for C4 photosynthesis (Sage, 2004; Christin et al., 2008), some degree of flexibility, in which enzymes have been co-opted for C4 metabolism, could be expected. In virtually every known example of C4-assisted photosynthesis, the initial step involves the carboxylation of PEP to oxaloacetate (OAA) by PEPC (Fig. 1). A notable exception occurs in the marine macroalga Udotea flabellum, where the C4 pathway starts with the carboxylation of PEP by phosphoenolpyruvate carboxykinase (PEPCK; Reiskind & Bowes, 1991). In all known systems that begin with PEPC-mediated carboxylation, the three known decarboxylation pathways involve either one of two isoforms of malic enzyme (NAD-ME or NADP-ME) or PEPCK. In pathways mediated by a malic enzyme (ME) isoform, OAA is rapidly converted to malate through malate dehydrogenase (MDH), transported to a separate compartment and decarboxylated by ME to form pyruvate (PYR) and CO2. The cycling of PYR back to PEP is

Fig. 1 Two abridged models of single-cell C4 photosynthesis. Solid arrow indicates the first step of C4 photosynthesis mediated by phosphoenolpyruvate carboxylase (PEPC). The first step (indicated by solid lines) of almost all known C4 pathways begins with the carboxylation of phosphoenolpyruvate (PEP) via PEPC to form oxaloacetate (OAA). Although there are some variations on the identity of the C4 acid (OAA, malate or aspartate) transported to the organelle (most often chloroplast), the second step is characterized by one of two archetypical pathways that involve decarboxylation by malic enzyme (ME; dashed lines) or phosphoenolpyruvate carboxykinase (PEPCK; dotted lines). In the ME pathway, OAA is converted to malate by malate dehydrogenase (MDH), and PEP is regenerated from pyruvate (PYR) by pyruvate phosphate dikinase (PPDK). Carbonic anhydrase (CA) catalyzes the interconversion between carbon dioxide and bicarbonate. New Phytologist (2014) www.newphytologist.com

New Phytologist accomplished by pyruvate phosphate dikinase (PPDK) and other enzymes. In a PEPCK-mediated pathway, OAA is decarboxylated to PEP and CO2. Recent research on single-cell C4 photosynthesis in marine diatoms has yielded disparate results. On the one hand, some biochemical and molecular data are suggestive of C4-assisted photosynthesis in T. pseudonana. For example, O2 evolution is dramatically reduced by the PEPC inhibitor, 3,3-dichloro-2-dihydroxyphosphinoylmethylpropenoate (DCDP), for both T. weissflogii and T. pseudonana grown at CO2 concentrations that are subsaturating with respect to RuBisCO carboxylase (Reinfelder, 2011), and can be rescued by the addition of HCO3 (McGinn & Morel, 2008). In addition, McGinn & Morel (2008) observed c. three-fold greater PEPC transcript abundances for T. pseudonana under low CO2. However, although Roberts et al. (2007) showed that malate is among the first products of carbon fixation in T. weissflogii (supporting the earlier findings of C4 photosynthesis in this species; Reinfelder et al., 2000), similar external CO2 conditions resulted in minimal malate labeling in T. pseudonana. In addition, in contrast with McGinn & Morel (2008), Roberts and colleagues found similar PEPC transcript abundances for T. pseudonana grown under low and high CO2 (380 and 100 ll l1, respectively). These findings led Roberts and co-workers to conclude that T. pseudonana does not exhibit C4 photosynthesis. Carbon-concentrating mechanisms help to minimize photorespiratory losses in vascular plants. In diatoms, low CO2 induces canonical biomarkers of photorespiration, but this is not manifested in decreases in growth rate as might be expected, raising questions about both the roles and biochemistry of photorespiration in diatoms. All the required genes for photorespiration, apart from glycerate kinase, are readily identified in both T. pseudonana and P. tricornutum genomes (Kroth et al., 2008). In diel experiments with T. pseudonana, Granum et al. (2009) observed light-dependent increases in glycine decarboxylase (GDC) transcript abundances under low (100 ll l1) but not ambient (360 ll l1) CO2. These transcripts, as well as relative cellular glycolate concentrations, also increased with increasing light intensity at ambient CO2 (Parker et al., 2004; Parker & Armbrust, 2005). Despite these indicators of photorespiration, manipulations of CO2/O2 stoichiometry that should lead to elevated rates of photorespiration do not lead to discernible changes in growth rate (Beardall, 1989). These apparent discrepancies between markers of the photorespiratory pathway and the negligible photorespiratory losses themselves suggest the adoption of some mechanism to recuperate the terminal products of photorespiration, CO2 and NH4+, or that these marker proteins may have biochemical functions other than photorespiration at low CO2. It is important to recognize that these enzymes are essential at both low and ambient CO2 in other phototrophs (Nakamura et al., 2005; Eisenhut et al., 2008; Zelitch et al., 2009). We hypothesized that T. pseudonana, like T. weissflogii, exhibits C4-assisted photosynthesis in response to low CO2. We also hypothesized that C4 metabolism might not entirely eliminate oxygenase activity, and explored whether compensatory mechanisms might be at play. These hypotheses were tested by Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

New Phytologist evaluating the transcriptional, photo-physiological and proteomic responses of these diatoms under conditions of carefully controlled CO2 availability. From these data, we develop a new model for carbon fixation and other physiological responses under low-CO2 conditions.

Materials and Methods Culture conditions and photo-physiology on CO2 shift Thalassiosira pseudonana (Cleve 1873), strain CCMP1335, was grown in medium prepared from synthetic ocean water according to the Aquil recipe (Sunda et al., 2005) and maintained at 18°C and 200 lmol photons m2 s1. Macronutrients were added at 100 lmol l1 NO3, 100 lmol l1 Si(OH)4 and 10 lmol l1 PO43. All medium preparation and sample handling were carried out in a class 100 laminar flow hood. All plastic ware was acid cleaned by soaking in 10% trace metal grade HCl for 5–10 d and rinsed with 18.2 MΩ cm deionized water. Media and culture flasks were microwave sterilized according to Keller (1988) in polycarbonate bottles. Cell density was determined using a Coulter counter (Beckman-Coulter, Fullerton, CA, USA), and growth rates were computed from linear regressions of ln(cell density) vs time. Cells were acclimated to a constant CO2 (1780 ll l1 or 61 lM aqueous CO2) and then shifted to 210 ll l1 (7.1 lM) CO2 within c. 15 s. Our pre-shift conditions were designed to provide a saturating supply of CO2, at about twice the half-saturation concentration for diatom RuBisCO (c. 30 lM; Badger et al., 1998), to greatly reduce the demand for a CCM relative to that for ambient CO2 levels. This was performed by maintaining cultures in house-built pH stats set at pH 7.61 (National Bureau of Standards scale) for > 10 generations. At the shift, 1 M NaOH was added whilst mixing the culture bottle to achieve a pH of 8.48. Before and following the shift, the pH was continuously monitored. CO2 was calculated using these pH values and the TCO2 (total CO2) content of Aquil (2.38 mM) using CO2SYS (http://cdiac.ornl.gov/oceans/co2rprt. html). As we control pCO2 by varying the pH at constant dissolved inorganic carbon concentrations, these experiments resemble ocean acidification experiments, except that here the concentration of aqueous CO2 is decreased, as occurs during bloom progression. The pH stats were assembled as follows. A gel-filled combination pH electrode (9106BNWP; Thermo, Waltham, MA, USA) was mounted through the wall of a 1-l polycarbonate bottle using a bulkhead mount. The electrode potential was monitored using a pH relay (pH200 controller; Eutech Inst., Singapore). When the pH increased above a set threshold, the relay switched on a peristaltic pump (Master Flex C/L; Cole Parmer, Vernon Hills, IL, USA) and an aquarium air pump to deliver weak acid (0.03 M trace metal grade HCl) whilst mixing the culture. Before and following a shift to low-CO2 conditions, samples were collected for the determination of chlorophyll a (Chl a)concentration, cell concentration, growth rate and photosynthesis–irradiance (PE) relationships. Samples (20–30 ml) for Chl a concentration were collected on glass fiber filters (GFF Whatman, GE Healthcare, Piscataway, NJ, USA). Chl was extracted Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

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overnight with 90% acetone and determined spectrophotometrically following the methods of Jeffrey & Humphrey (1975). Cell concentrations and growth rate were determined as above. The PE relationships for samples were determined from 10 min incubations with 14C bicarbonate in a photosynthetron (Lewis & Smith, 1983). Ten irradiances (0–1750 lmol photons m2 s1) were used during each incubation, with the resulting data fitted to the equation of Platt & Gallegos (1980) using non-linear regression: PB ¼ Ps ð1  e a Þe b ;

Eqn 1

where PB is the photosynthetic rate normalized to Chl a and Ps is the maximum photosynthetic rate in the absence of photoinhibition. For the exponents of Eqn 1, a = aE/Ps and b = bE/Ps, where a is the initial slope of the P vs E curve, b is the descending slope at high light and E is the scalar irradiance. There was no evidence of photoinhibition in our P vs E data, and so b was set to zero for fitting purposes. Variable fluorescence (Fv/Fm = (Fm  F0)/Fm) was measured using a fast-repetition-rate fluorometer (FRRf; Kolber et al., 1998). Samples were removed from the culture vessel and immediately measured (no dark acclimation period) for F0 (Chl fluorescence yield when the photosystem II, PSII, reaction centers are mostly oxidized) and Fm (Chl fluorescence yield when all functional PSII reaction centers are reduced). We purposely did not provide an opportunity for acclimation to capture the state of variable fluorescence in the culture vessel. The capacity to induce non-photochemical quenching (NPQ) was determined in a temperature-controlled (18°C) pulse amplitude-modulated (PAM) fluorometer (DUALPAM-100; Heinz Walz GmbH, Effeltrich, Germany) fitted with a photomultiplier detector (DUAL-DPM). NPQ was determined using a PAM protocol entailing an initial low-light (16 lmol photons m2 s1) acclimation period of 15 min, followed by a dark period of 5 min. It has been shown that diatoms require a low-light exposure to allow NPQ mechanisms to fully revert to a non-quenching state (Grouneva et al., 2008; Miloslavina et al., 2009; Milligan et al., 2012). The relaxation period was followed by an actinic light exposure (300 lmol photons m2 s1) for 10 min to induce NPQ, and finally a second dark period of 21 min. The maximum fluorescence (Fm) in the NPQ relaxed state was determined before actinic light exposure and during actinic light exposure (Fm0 ). NPQ was calculated as (Govindjee, 2004): NPQ ¼ ðFm =Fm0 Þ  1

Eqn 2

Quantitative PCR analysis of transcript abundances on CO2 shift Cultures were maintained under steady-state high CO2 at 350 lmol photons m2 s1 and subject to CO2 shifts as in the photo-physiology experiments, except that Aquil (Sunda et al., 2005) was assembled using coastal seawater rather than synthetic ocean water. In this case, the high- and low-CO2 pH values of New Phytologist (2014) www.newphytologist.com

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7.61 and 8.48 correspond to CO2 values of 184 and 1500 ll l1 (54 and 6.6 lM) based on a coastal seawater total alkalinity of 2.04 mM (18°C, salinity = 25; Lee et al., 2006). Gene transcript abundances for seven potentially important enzymes in C4 metabolism were quantified using quantitative reverse transcription-polymerase chain reaction (Q-RT-PCR hereafter) over an eight-point time course corresponding to 30, 20, 10, + 10, + 20, + 30, + 60 and + 90 min relative to the time of shift to low CO2, using actin as the housekeeping gene. RNA extraction, complementary DNA (cDNA) synthesis and QRT-PCR procedures generally followed those of Kustka et al. (2007) with some modifications. RNA was extracted using a modification of the Trizol method; cells were scraped from a 47mm polycarbonate filter into 60–65°C Trizol, passed through a 25-gauge needle four to six times to disperse clumps, frozen in liquid nitrogen and stored at 80°C for < 30 d. On thawing, solids were cleared via centrifugation and the supernatant was transferred to a new vial. Chloroform extraction was performed, and the aqueous phase was removed, mixed with 1 : 1 v/v 67% ethanol and loaded onto an ‘Absolutely RNA’ spinkit (StratageneAgilent, La Jolla, CA, USA). To remove genomic DNA, two sequential 30-min reactions were performed with Turbo-DNAse (Ambion-Life Technologies, Grand Island, NY, USA). Q-RTPCRs were performed with a Stratagene Mx3000P thermal cycler using Power SYBR mastermix. Equal amounts of template (either 10 or 25 ng of RNA equivalent) were used in triplicate 20-ll reactions. The absence of interfering genomic DNA contamination was validated with PCRs containing ‘no reverse transcription’ RNA as template, using identical conditions. Copy numbers of genes of interest (GOI), relative to that of actin, were estimated as 2[1(CtGOI – Ctactin)] [shorthand as 2(DCt)] according to Pfaffl (2001). This quantity was determined from replicate analyses for each observation, within each experiment. Melt curves, inspected with each quantitative PCR analysis for each reaction well, indicated single peaks with melting temperatures consistent with the absence of primer dimers. This finding was additionally confirmed by gel electrophoresis for each primer from one treatment once from each of two biological replicates. Standard templates were generated by gel purification of PCR products using gene-specific primers and quantified with a NanoDrop 1000 (Wilmington, DE, USA) spectrophotometer. For each primer pair, real-time PCR efficiency was calculated by plotting Ct vs log(template copy number) from the analysis of a four-point serial dilution spanning a range of 256-fold template concentration. The slope was used to calculate the efficiency, Efficiency = 10(1/slope). The average efficiency for all quantitative PCR primers in this study was 1.90  0.023 (SD), close to the theoretical value of 2. The primers used in this study are given in Table 1. Validation of housekeeping genes and limits on quantification The suitability of a housekeeper depends on the lack of a discernible trend in cycle threshold value along gradients of treatments (or time in this case). The analysis of residual Ct values vs time New Phytologist (2014) www.newphytologist.com

showed no trend (Supporting Information Table S1), suggesting that this housekeeping gene is suitable for studies of transcript abundance with shifting CO2. The minimum level of differential gene expression that can be detected (below which apparent shifts in transcript abundance may be caused by some small variation in measured actin transcript) was calculated with a 5% type I error rate by analyzing a population of standard deviations in actin Ct from all pairwise combinations of pre-shift time points. This threshold equaled a standard deviation of 0.26 Ct, which translates into a 20% copy number difference in actin transcripts between any two identical samples. For transcripts which exhibited shifts above these copy number difference thresholds (PEPC1 and PEPC2), differences between pre- and post-shift transcript abundances relative to actin (TRA) were evaluated using a one-tailed t-test on pooled data from both biological replicates. Quantitative proteomic analyses at steady state Cultures were grown under steady-state conditions of low and high CO2 (210 and 1780 ll l1) in pH stats with Aquil medium, as described above, except that either low- or high-CO2 cells were grown with 15N- nitrate (> 98%) or nitrate with a natural isotopic composition (i.e. 99.6% 14N). Two sets of independent biological replicates from each condition were processed. For each condition, c. 900 ml were harvested at (4–5) 9 105 cells ml1 and flash frozen in liquid nitrogen. Protein samples were extracted in 4% sodium dodecylsulfate (SDS), 7.5% glycerol, in 0.1 M Na2CO3 and protease inhibitor (Sigma-Aldrich P-2714), and quantified before the addition of 0.1 mM dithiothreitol Table 1 Primers used in quantitative PCR. Each gene name is as given in the text, followed by the protein identification number (in parentheses) and genome locus (underscored) Gene name locus forward and reverse primers (50 –30 ) Phosphoenolpyruvate carboxylase (3830) chr_3:1899971–1903147 CAGCGTCTCATGGCAGTTAGAAATC CTCCGAGCTTCCCTCACTTTCAC Phosphoenolpyruvate carboxylase (5027) chr_5:200957–204623 GGCGACCCCTGAGTTGGAACTT CACGGAATAGCTCTCAAACTGTCAACAC Malate dehydrogenase; cytoplasmic (41425) chr_7:1677378–1678600 CATGGAGAAGCTGCAAGTCACTGA GCCAACGAAGCAGTGGATACAGTAG Malate dehydrogenase; mitochondrial (175) chr_1:985790–987134 ATGGCGTATGCTGGCTATGTATTTACAG GCAAAATCTCCTCTACACCTCCCTTAC Malic enzyme (5100) chr_5:366187–368316 GCGAAACACTGCGAAAGACCAATC ACGGGCTGCCACTAGCAAAGATAC Phosphoenolpyruvate carboxykinase (5186) chr_5:566357–568410 GGTGGGAGCTTCCTGCCTCTATT AATCGACATCTCCTTTCCCCTTGTAC Pyruvate phosphate dikinase (5500) chr_5:1377134–1380283 GAGGCTGAGCGCATTAAAGTTGAAC TGGCTTGCTGTCTTCGTCGTCTA Actin (25772) chr_22:804575–806436 ACTGGATTGGAGATGGATGG CAAAGCCGTAATCTCCTTCG Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

New Phytologist (DTT). Fifty micrograms of protein from each condition were mixed, yielding a 1 : 1 ratio of 15N- to 14N-labeled protein mixture. Subsequent processing was carried out at the Biological Mass Spectrometry Facility of the UMDNJ-Robert Wood Johnson Medical School, New Brunswick, NJ, USA. The protein mixture was digested by trypsin. Digested peptides were solubilized in buffer A (20 mM ammonium formate, pH10) and subjected to high-pH reverse-phase high-performance liquid chromatography (HPLC) (Gilson 306 pumps, 805 manometric module and UV/VIS 155 detector) equipped with a XbridgeTM C18 column (3.5 lm, 2.1 9 150 mm, Waters, Milford, MA, USA). The gradients used for separation of the peptides were 2% buffer B (20 mM ammonium formate, 90% acetonitrile, pH 10) for 2 min, then 2–45% B in 43 min, 45–100% B in 5 min; 1-min fractions were collected and vacuum dried before being combined or individually analyzed by nano-LC-MS/MS. Nano-LC-MS/MS was performed using an RSLC system interfaced with an LTQ Orbitrap Velos (ThermoFisher, Waltham, MA, USA). Samples were loaded onto a self-packed 100 lm 9 2 cm trap packed with Magic C18AQ, 5 lm 200 A (Bruker Auburn, Auburn, CA, USA), and washed with buffer A (0.2% formic acid) for 5 min with a flow rate of 10 ll min1. The trap was brought in-line with the home-built analytical column (Magic C18AQ, 3 lm 200 A, 75 lm 9 50 cm) and peptides were fractionated at 300 nl min1 with a multistepped gradient (4–15% buffer B (0.16% formic acid, 80% acetonitrile) in 25 min and 15–25% B in 65 min and 25–50% B in 55 min). The mass spectrometer acquisition cycled through one MS in Orbitrap (resolution 60 000), followed by 20 MS/MS (CID) in LTQ with dynamic exclusion (two repeat counts within 30 s and exclusion time of 60 s). The LC-MS/MS data were analyzed using Proteome Discoverer software v1.3 (ThermoFisher). The data were first searched against T. pseudonana (composed of sequences queried from Uniprot) using a Sequest search engine through a ‘light’ search assuming normal nitrogen isotope distribution and a ‘heavy’ search assuming all amino acids were labeled with 15N. For both ‘heavy’ and ‘light’ searches, carbamidoethyl on cysteine was used as a fixed modification. For the ‘light’ search, oxidation of methionine was included as a variable modification. For the ‘heavy’ search, N-terminal modification of 0.997 Da was included as a variable modification. The identified peptides were quantified with a custom-built precursor ion quantification method within the software. The peptide quantification results of each HPLC fraction were combined into one list (MUDPIT). Peptides were filtered to include only top ranked identification with confidence above medium. The heavy/light ratio of each peptide was normalized to the median ratios of all peptides. The heavy/light ratio of protein was calculated as the median value of all peptides belonging to this protein. The protein ratio was accepted if three or more peptides were quantified. Detection limits for the responsive proteome The minimum difference in 15N : 14N ratios for differential protein abundances was determined by evaluating the distribution of relative protein abundances for T. pseudonana grown under Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

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Fig. 2 Thalassiosira pseudonana photosynthetic parameters before and following a shift (at t = 0) from high to low CO2. (a) The initial slope of the photosynthesis–irradiance (PE) relationship (ab, circles) and chlorophyllspecific maximum carbon fixation rate (Pbmax, squares) for PE relationships determined in 10-min 14C-bicarbonate assimilation assays. (b) Light reaction parameters: variable to maximum fluorescence (Fv/Fm, circles) determined immediately (no dark acclimation period) and nonphotochemical quenching (NPQ, rectangles) determined on samples in which NPQ was permitted to relax. The widths of the symbols for each parameter are equal to the time required to make the measurement, except for Fv/Fm, which is instantaneous. Error bars,  SD.

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Fig. 3 Relative gene expression for phosphoenolpyruvate carboxylase 1 (a) and phosphoenolpyruvate carboxylase 2 (b) in Thalassiosira pseudonana. At time zero, cells were shifted from high to low CO2. Gene expression relative to that for actin was determined in two replicate analyses per experiment. Experiments one and two are indicated as filled and hollow symbols, respectively. For each of two replicate experiments, error bars  SD of replicate analyses. New Phytologist (2014) www.newphytologist.com

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identical conditions (CO2 at c. 1780 ll l1), except that the N source was provided as either 14N-NO3 or 15N-NO3. This serves to integrate the error caused by the variability in mass spectrometer ionization efficiencies as well as biological variability. The raw 15N : 14N peak area ratio for each protein was normalized to the global median 15N : 14N value and then log2 transformed. The standard deviation of these log2-transformed 15 N : 14N ratios was 0.167 (Table S2). Setting the type I error rate at  1% corresponded to a threshold log2-normalized 15 N : 14N ratio of  0.388. Proteins from experimental treatments (grown in 14N and 15N under low and high CO2, respectively) were considered to be differentially expressed when the log2-transformed value in both biological replicates was either < 0.388 (corresponding to a median-normalized 15N : 14N ratio of 0.7641 or less; up-regulated under low CO2) or > 0.388 (corresponding to a median normalized 15N : 14N ratio of 1.309; up-regulated under high CO2). We applied this criterion to the 15 N : 14N ratios for each protein from both biological replicates. Assuming that the error of the 15N : 14N ratio measured for any protein is random and represented by the standard deviation measured here, this criterion translates to a false positive rate of 0.01%. For proteins differentially expressed in both biological replicates, we report the average fold change (geometric mean) in protein abundance, with an exception for proteins that are extremely abundant under one condition relative to the other. Isotopic ratios are difficult to constrain for such instances (i.e. those with

Subcellular protein targeting Targeting was assessed using the predicted localization of gene/ Uniprot models (Table S3). In some cases, gene and Uniprot models were re-evaluated and models were extended upstream. We followed the general approaches outlined in Gruber et al. (2007), Kroth et al. (2008) and Bender et al. (2012). Specific criteria to assess targeting are provided in more detail in Table S3.

Results Photo-physiology and transcripts in response to low - CO2 shift Maximum Chl-specific carbon fixation rates (Pbmax) declined by 54% within the first 10 min after the shift to low CO2, whereas the initial slope of the irradiance vs fixation curve (ab) was unaffected (Fig. 2a). Before Pbmax recovery, the variable fluorescence

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ratios < 1 : 12 or > 12 : 1; Huttlin et al., 2007). For this reason, we caution on the over-interpretation of the quantitative nature of such extreme peptide ratios, and rather support the interpretation that such peptides have abundance ratios much less (or greater) than 1 : 1. In cases in which the apparent fold increase in protein abundance for one replicate was > 12, we indicate the fold change in protein abundance as at least as great as the lesser fold change measured.

Fig. 4 Relative gene expression for proteins potentially involved in either of the two archetypical C4 decarboxylation pathways in Thalassiosira pseudonana. At time zero, cells were shifted from high to low CO2. (a) Cytosolic malate dehydrogenase, (b) NADdependent malic enzyme, (c) pyruvate phosphate dikinase, (d) mitochondrial malate dehydrogenase, (e) phosphoenolpyruvate carboxykinase. Gene expression relative to that for actin was determined in two replicate analyses per experiment. Experiment one and two are indicated as filled and hollow symbols, respectively. For each of two replicate experiments, error bars  SD of replicate analyses. These values were all below the 20% transcript relative abundance threshold (Supporting Information Table S1). Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

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(Fv/Fm) dropped (11%) and the capacity to induce NPQ rose by 37% (Fig. 2b). These responses are consistent with a more reduced photosynthetic electron transport chain under acute CO2 limitation. Within 30–50 min after their initial deviation, NPQ and Fv/Fm both recovered to values obtained under steady-state high CO2. The culture pH remained at 8.48 throughout the recovery period (data not shown). When cultures were shifted from high to low CO2, mitochondrial targeted PEPC1 TRA increased by 5.7-fold within 30 min of the shift (Fig. 3a). By 90 min, PEPC1 TRA had declined to 2.2-fold of pre-shift values. Similarly, TRA of PEPC2 (targeted to the chloroplastic endoplasmic reticulum (CER) or periplastidic space (PPS)) increased by 6.5-fold within 30 min of the shift (Fig. 3b), and decreased to c. 2.3-fold of pre-shift values by 90 min. These increases were above the minimum threshold of 20% (see the Materials and Methods section and Table S1). Transcript abundances were significantly higher

for PEPC1 (P < 0.003) and PEPC2 (P < 0.005) after exposure to low CO2. Transcript abundances for other genes potentially involved in pathways for C4 acid transport and decarboxylation were also evaluated (Fig. 4). The average differences in TRA between pre- and post-shift conditions for two MDHs, NAD-ME, PPDK and PEPCK were below the 20% TRA threshold (Table S1). Quantitative proteomic analyses at steady state Under steady-state growth conditions, growth rates during these experiments did not differ at the two pH/CO2 levels investigated (2.06  0.014 d1 at pH 7.61 and 2.03  0.071 d1 at pH 8.48), which is consistent with other findings that ocean acidification (lower pH) does not have an obvious direct effect on T. pseudonana growth (Crawfurd et al., 2011). Using a nitrogen isotope labeling approach, we were able to quantify 3688 proteins

Table 2 Proteins up-regulated under low CO2 in both replicates Uniprot ID

Description

Replicate 1

Replicate 2

Average fold change

B8BTB4 B8C083 B8C250 B8LE18 B8LE19 B8C0D9 B8CG97 B8C4I8 B8C0D8 B8LE17 B8BSW5 B8C8T5 B8BQZ0 B8CDL6 B8BY90 B8BZ35 B8BSI9 B8CE42 B8BPY6 B8BVD1 B8C3W3 B5YMF5 B8C1R7 B8LDH1 B8BX31 B8C596 B8C017 B8C1C4 B8C5V3 B8BXJ5 B8BWX9 B8BZM5 B8BYW8 B8BYN5

Delta carbonic anhydrase Putative DNA-binding heat shock factor protein Carbonic anhydrase Mucin-like protein HNKH Bestrophin 2, similar to low-CO2 inducible membrane protein Cadmium carbonic anhydrase Putative glycoprotein Bestrophin 1; similar to low-CO2 inducible membrane protein HNKH WAX2-like protein Glutamine-dependent carbamoyl phosphate synthase-like protein Ornithine cyclodeaminase AMP yield acyl-CoA synthetase Formate–tetrahydrofolate ligase Alanine glyoxylate aminotransferase/serine pyruvate transaminase HNKH Pyruvate carboxylase HNKH Related to biotin carboxylase and acetyl-CoA carboxylase HNKH Acetyl-CoA carboxylase Phosphoenolpyruvate carboxylase Putative SAM-dependent methyltransferase Glycine decarboxylase p-protein HNKH Alanine aminotransferase Cold-shock DNA-binding domain-containing protein Biotin synthase L-Lactate dehydrogenase Microsomal omega-6 fatty acid desaturase BolA-like protein Phosphoenolpyruvate carboxylase HNKH

580.00 140.23 506.71 15.18 7.89 4.75 77.83 2.36 3.51 3.19 1.36 2.28 1.88 2.37 1.93 1.60 1.49 1.99 1.95 1.75 1.83 1.62 1.71 1.64 1.58 1.50 1.48 1.50 1.59 1.55 1.46 1.47 1.32 1.39

11.37 14.68 9.08 8.67 5.36 3.78 4.04 4.74 3.13 2.30 2.59 1.63 1.95 1.37 1.66 1.95 2.06 1.54 1.51 1.63 1.34 1.45 1.35 1.40 1.45 1.49 1.48 1.46 1.34 1.35 1.42 1.39 1.46 1.38

≥ 11 ≥ 15 ≥9 ≥9 6.50 4.24 ≥4 3.34 3.31 2.71 1.88 1.93 1.91 1.80 1.79 1.77 1.75 1.75 1.72 1.69 1.57 1.53 1.52 1.52 1.51 1.49 1.48 1.48 1.46 1.45 1.44 1.43 1.39 1.38

The fold increase in protein abundance within each biological replicate was calculated as the inverse of the normalized 15N : 14N ratio. Averages were calculated as the geometric mean from biological replicates. When apparent fold increases for one replicate were > 12, the average was substituted with a range equal to or greater than the lesser value (see text for more details). Protein descriptions listed as HNKH were hypothetical proteins with no known described homologs according to BLASTp search (National Center for Biotechnology Information, NCBI) with a cut-off of E > 1030. Up-regulated but incomplete protein models without plausible N termini (Supporting Information Table S3; B8C5S5, B8C235, B8C333, B8LDG9, B8CEV2) are not shown here. Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

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8 Research Table 3 Proteins down-regulated under low CO2 in both replicates Uniprot ID

Description

Replicate 1

Replicate 2

Average fold change

B8C274 B8LE04 B8BW41 B5YNT2 B8LEL7 B8BW74 B8BVA0 B8C8L8 B8BXS2 B8LD81 B8LBQ0 B8C5Q7 B5YNK8 B8BSB3 B8BRH9 B8BT47 B8C7H4 B8LE89

Phosphoenolpyruvate carboxykinase ISIP3-like protein Similar to cobalamin synthesis protein Glutathione reductase Nickel ABC transporter HNKH Transaldolase HNKH Fructose-1,6-bisphosphatase K exchanger-like protein Putative NAD-dependent epimerase/dehydratase HNKH HNKH Putative triose phosphate translocator HNKH Similar to Lipid A export msbA protein HNKH HNKH

321 5.04 1724 4.00 579 2.34 2.50 2.21 2.31 2.11 1.80 19 1.37 1.71 1.59 1.61 1.37 1.36

5.08 1.84 2.71 1.36 2.50 1.86 1.44 1.55 1.40 1.36 1.67 1.72 2.00 1.52 1.39 1.34 1.36 1.33

≥5 3.05 ≥ 2.71 2.33 ≥ 2.5 2.09 1.90 1.85 1.80 1.69 1.73 ≥ 1.72 1.66 1.61 1.49 1.47 1.36 1.34

The fold decrease in protein abundance within each biological replicate is the normalized 15N : 14N ratio. Averages and protein descriptions are given as in Table 2. Down-regulated but incomplete protein models without plausible N termini (Supporting Information Table S3; B8BWI1, B8C0W2, B8C443, B8C231) are not shown here.

from the T. pseudonana proteome from both replicate experiments (Table S4). Of these, 39 and 22 proteins were significantly up-regulated (Tables 2, S3) and down-regulated (Tables 3, S3) under low CO2, respectively. Targeting predictions and relative abundances for select proteins are shown in Tables S3 and S5, respectively. The most highly up-regulated proteins at low CO2 included three carbonic anhydrases, CA-4, CA-6 and CA-3 (Uniprot IDs B8BTB4, B8C250 and B8CG97, respectively), following the nomenclature of Tachibana et al. (2011). CA-4 and CA-3 may be localized to the cytosol, whereas CA-6 may be targeted to the cell surface or ER membrane. Two chloroplast-targeted proteins in the bestrophin family of anion channels (B8C0D8 and B8C0D9) were also present in higher abundance at low CO2. Among the candidate C4 metabolism proteins, both PEPC paralogs were more abundant under low-CO2 conditions. PEPC2 (B8C1R7) exhibited SignalP targeting scores suggestive of targeting to the CER or PPS, whereas PEPC1 (B8BYW8) seemed to be mitochondrial. ME (B8C1Z0) abundances did not change in response to CO2 (Table S5), whereas PEPCK (B8C274) abundances decreased by at least five-fold. Notably, PPDK (B8C332) was not detected in these proteomes. The abundance of a chloroplast-targeted pyruvate carboxylase (PYC; B8CE42) was 1.8-fold higher under low CO2. The expression patterns of other proteins include the 1.6-fold down-regulation of a putative triose phosphate translocator (B8BSB3), which belongs to a family of proteins capable of triose phosphate or 3-phosphoglycerate (3-PGA) transport (Fl€ ugge, 1999), and has ER targeting and possible anchoring to the ER membrane. In addition, cytoplasmic fructose 1,6-bisphosphatase (B8BXS2), transaldolase (B8BVA0) and epimerase (B8LBQ0), components of the non-oxidative portion of the pentose phosphate pathway, were down-regulated at low CO2 (Table 3). Core proteins in the oxidative phase of the pentose phosphate pathway were not differentially expressed, but New Phytologist (2014) www.newphytologist.com

glutathione reductase (B5YNT2), which participates in a side reaction consuming NADPH, was down-regulated by c. 2.3-fold under low CO2 (Fig. 5; Table 3). Some photorespiratory proteins, including GDC P protein (B8BX31) and alanine glyoxylate aminotransferase (AGAT; B8BZ35), were up-regulated (by 1.5and 1.8-fold, respectively) under low-CO2 conditions (Table 2). Proteins involved in ornithine and carbamoyl phosphate metabolism were up-regulated under steady-state low CO2. Ornithine cyclodeaminase (B8BQZ0) was 1.9-fold more abundant under low CO2, whereas glutamine-dependent carbamoyl phosphate synthetase (B8C8T5) was 1.9-fold more abundant. However, no proteins involved in the ornithine–urea cycle (OUC) were differentially expressed (Table S5).

Discussion These experiments were designed to examine the physiological responses to decreasing CO2 which are known to occur during blooms. Our pre-shift conditions provided a supply of CO2 at about twice the half-saturation concentration for diatom RuBisCO (c. 30 lM; Badger et al., 1998) to greatly reduce the demand for a CCM relative to that at ambient CO2. Following the shift to low CO2 (7.1 lM), carbon fixation (Pbmax) dropped immediately. The concomitant drop in variable fluorescence and increase in NPQ capacity are consistent with the limitation of photosynthetic electron transport by CO2 availability. The recovery of all photophysiological parameters was complete within 100 min. Both PEPC transcripts were up-regulated on shifting to low CO2 and, after 90 min, these transcripts were c. two-fold greater than pre-shift values (Fig. 3). The slightly greater response (c. three-fold) observed by McGinn & Morel (2008) may reflect the lower CO2 values (at pH 8.9 vs pH 8.48 here) in their experiments. In contrast, Roberts et al. (2007) did not observe Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

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Fig. 5 Effects of low CO2 on cytoplasmic pentose phosphate pathway relative protein abundances in Thalassiosira pseudonana (determined as described in the text). Red and black arrows indicate down-regulated and equally represented proteins at low CO2 relative to high CO2, whereas gray arrow indicates protein was not quantified in both biological replicates. The upper and lower portions represent the oxidative and nonoxidative phases of the pentose phosphate pathway, respectively. Enzymes highlighted here – and their corresponding Uniprot IDs – include NAD-dependent epimerase (EP; B8LBQ0), fructose bisphosphatase (FBP; B8BXS2), fructose bisphosphate aldolase (ALDO; B8CFH1), glucose-6phosphate dehydrogenase (G6PD; B8C3E7), glucose-6-phosphate isomerase (GPI; B8BYC8 or B8LDA5), 6-phosphogluconate dehydrogenase (6PGD; B8BZH6), transketolase (TK; B8BTR4), transaldolase (TA; B8BVA0), triose phosphate isomerase (TPI; B5YLS7, B5YNQ0 or B8C5E1) and ribose-5-phosphate isomerase (RPI; B8BUF5). Abbreviations for substrates: Ery4P, erythrose-4-phosphate; F6P, fructose6-phosphate; GADP, glyceraldehyde phosphate; G6P, glucose-6phosphate; PEP, phosphoenolpyruvate; 6PG, 6-phosphogluconate; R5P, ribose-5-phosphate, Ru5P, ribulose-5-phosphate; S7P, sedoheptulose-7phosphate; Xu5P, xylulose-5-phosphate. The down-regulation of cytoplasmic PPP enzymes may serve to diminish the drawdown of GADP in the cytoplasm and retain more GADP in the chloroplast to ensure sufficient regeneration of Ru5P lost to photorespiration. Plastid PPP enzyme abundances did not change between treatments (Supporting Information Table S4).

significant changes in transcript abundances for either PEPC. Two factors may explain these disparate results. Steady-state differences in transcript abundances are expected to be minimal, whereas, during the rapid shifts from high to low CO2, we expected (and observed) a transient state in which photosynthesis was limited by CO2 availability, followed by recovery after CO2responsive genes were transcribed. Therefore, the results of Roberts et al. (2007) may reflect the conditions of the cultures, rather than the absence of C4-assisted photosynthesis. In addition, their CO2 concentrations (380 and 100 ll l1 CO2) vastly undersaturate RuBisCO carboxylase activity (Badger et al., 1998), raising the possibility that the CCM would be operating at similar rates under both conditions. Short-term changes in PEPC transcription observed in our CO2 shift experiments are corroborated by steady-state proteomic results (Table 2), where abundances of CER- or PPS-targeted PEPC2 increased c. 1.5-fold under low CO2. Increased PEPC2 Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

Research 9

protein levels may elevate OAA production rates in the CER (or PPS). Given the filamentous distribution of the ER, this carboxylation is functionally analogous to the cytoplasmic OAA production in other single-cell C4 systems (Voznesenskaya et al., 2002; Edwards et al., 2004). These increases in PEPC abundance may also be accompanied by other adaptations to low CO2 that serve to increase PEPC activity. For example, the down-regulation of an ER membrane-anchored PGA/glyceraldehyde phosphate (GADP) transporter (B8BSB3) under low CO2 would help to decrease 3-PGA export to the cytosol and promote the production of PEP in the CER (described in detail below). Our results do not support either ME- or PEPCK-mediated decarboxylation for C4 photosynthesis in T. pseudonana. Intracellular targeting analysis suggests that neither protein is localized in the chloroplast, inconsistent with the most parsimonious model of where C4 decarboxylation should occur. Moreover, various transcripts for proteins involved in the putative ME-mediated C4 pathway (Fig. 1) did not increase after the shift to low CO2 (Fig. 4). This suggests that, if ME is responsible for decarboxylation here, it is not regulated at the transcriptional level. The indistinguishable protein abundances for ME (Table S5) further suggest that ME may not be involved in C4-assisted photosynthesis in this species. The absence of transcript abundance differences (Fig. 4) and the five-fold or greater decrease in PEPCK protein abundances (Table 3) at low CO2 are consistent with the decrease in P. tricornutum PEPCK activities at low CO2 (Cassar & Laws, 2007), but not with a role for PEPCK-mediated decarboxylation in C4-assisted photosynthesis here. The identity of the proteins responsible for the decarboxylation step of C4 photosynthesis in these diatoms is unresolved. The independent origins for C4 photosynthesis (Sage, 2004) leave open the possibility that some other enzyme altogether may have been co-opted for the decarboxylation step in diatoms. The chloroplast-localized and reversible PYC (B8CE42) was c. 1.8-fold more abundant under low CO2. The decarboxylation of OAA by PYC, coupled to the synthesis of ATP from ADP, is exothermic at neutral pH, an ATP/ADP ratio of 2.5 and an OAA concentration > 1 mM (calculations not shown), which raises the possibility that this protein carries out the decarboxylation reaction (as has been observed in model systems; Attwood & Cleland, 1986). However, a forward-acting PYC in T. pseudonana would seem to compete with RuBisCO for HCO3/CO2 under the very conditions in which plastid CO2 must be elevated to support rapid growth. Four conditions should be satisfied to support the hypothesized role for PYC. First, OAA formed in the CER or PPS needs to be transported to the chloroplast. Two chloroplast-targeted bestrophin family anion channels were among the most up-regulated proteins under low CO2. One plausible role for these bestrophins is to facilitate diffusion of CER/PPS-localized OAA to the chloroplast, particularly given the recent discovery that some bestrophins are permeable to carboxylated organic anions (Roberts et al., 2011). Our current ability to characterize the substrate specificity of bestrophins in silico is admittedly very limited (Hagen et al., 2005), leaving open the possibility of an altogether different function for these proteins. Second, this mechanism of PYC-mediated decarboxylation also requires the participation of New Phytologist (2014) www.newphytologist.com

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a plastid-localized CA. Tachibana et al. (2011) showed clear experimental evidence of plastid targeting of CA1-green fluorescent protein (CA1-GFP) fusion proteins in T. pseudonana. Yet, bioinformatics analysis suggests that this may be targeted to the CER/PPS (Table S3), as it lacks the canonical signal-transit peptide boundary motif (Kilian & Kroth, 2005; Gruber et al., 2007). This example illustrates a shortcoming of our in silico knowledge of diatom physiology brought about by uncertainties in protein targeting. Third, the regeneration of PEP from PYR is required to sustain the C4 pump. This is most often thought to be achieved by PPDK (Kroth et al., 2008), but transcription of PPDK did not respond to CO2, and PPDK was not detected in our proteome or by probing with antibodies raised towards Zea mays (courtesy of Chris Chastain, Minnesota State University, Moorhead, MN, USA; data not shown). In addition, PPDK has been shown recently to have little effect on growth or photosynthesis in P. tricornutum (Haimovich-Dayan et al., 2013). Fourth, plastid PYR concentrations should be drawn down relative to those of OAA to promote the back-reaction. The proteomic response of T. pseudonana suggests that a PPDK-independent mechanism of PEP regeneration may be at play (Fig. 6), which also satisfies the third and fourth criteria. Greater abundances of cytoplasmic alanine aminotransferase (ALAT), the bifunctional mitochondrial AGAT/serine pyruvate transaminase and mitochondrial GDC at low CO2 suggest an alternative pathway to recover PYR through its transfer to the mitochondria as alanine and its sequential conversion to 3-hydroxypyruvate, glycerate and 3-PGA (Kroth et al., 2008). A potential complete pathway for the regeneration of PEP from PYR is outlined in Fig. 6(b). The unchanged and lower abundances of chloroplast membrane- and CER-localized triose

New Phytologist phosphate translocators (TPT3; B8BSB3 and TPT4; B5YLS2), respectively, at low CO2 would support the retention of 3-PGA in the CER for subsequent conversion to 2-PGA by phosphoglycerate mutase (PGM) (Fig. 6), although the topologies of these translocators are unknown. PEP formed by cytosolic enolase may be imported to the CER by a PEP/inorganic phosphate (PEP/Pi) antiporter (PPT), which can accept either Pi or 2-PGA as a counter-exchange substrate (Fischer et al., 1997). These proteins have been functionally verified and best described in vascular plants (Fischer et al., 1994, 1997; Fl€ugge et al., 2011), but there has been no heterologous expression work in any photosynthetic chromalveolates, and there are currently no clear candidates in the T. pseudonana genome. Three proteins from our limited list of those up-regulated at low CO2 would serve to draw down PYR concentrations and facilitate the net export of PYR from the plastid. These up-regulated proteins are also integral to the regeneration of PEP from PYR. Malate is the principal C4 acid isolated from low-CO2-acclimated T. weissflogii (Roberts et al., 2007), which, at first blush, might suggest that malate is the decarboxylation substrate. However, both malate and OAA equally compete for 14CO2 fixation (Reinfelder et al., 2004), suggesting that diatoms rapidly interconvert these substrates. This proposed model of OAA decarboxylation in T. pseudonana is consistent with these observations, considering that the CER/PPS-localized MDH may serve to buffer the CER/PPS pool of OAA. Without some mechanism to buffer the OAA supply, the PEPC2-mediated production and flux of OAA into the plastid would result in futile CO2 production during, for example, periods of transient low light. The production of malate by MDH when CER/PPS OAA concentrations become transiently elevated, as OAA production exceeds demand,

Fig. 6 Model of C4 photosynthetic carbon assimilation and associated rearrangement of carbon metabolism to support growth at low CO2 in Thalassiosira pseudonana. (a) Core components of the model. Red, green and black arrows indicate down-regulated, up-regulated and equally represented proteins at low CO2 relative to high CO2, respectively, whereas gray arrows indicate that protein was not quantified in both biological replicates. In cases in which more than one candidate protein may be responsible, multiple arrows indicating CO2-responsive status are shown. In the chloroplastic endoplasmic reticulum CER or periplastidic space PPS, PEPC2 converts bicarbonate and PEP to OAA. PEP concentrations are enhanced by the down-regulation of a PEP transporter (TPT3) anchored to the ER membrane. Elevated mitochondrial production of CO2 by photorespiration (only GDC is shown here for simplicity) may be scavenged by mitochondrial CA and PEPC1 (forming OAA), whereas diminution of mitochondrial PEPCK-catalyzed C4 decarboxylation would prevent a futile CO2–C4 cycle in the mitochondrion. The down-regulation of PEPCK should favor a greater flux of OAA decarboxylation through pyruvate carboxylase. Greater abundances of one or both anion channel bestrophin proteins (BEST1/2) may be responsible for facilitating OAA diffusion into the chloroplast at low CO2. Malate dehydrogenase may serve to minimize the futile decarboxylation of OAA under transient conditions of low light by minimizing any accumulation of OAA from PEPC2 under these conditions. OAA decarboxylation by PYC produces bicarbonate, which is dehydrated to CO2 in the proximity of RuBisCO by a chloroplast-localized CA, such as CA1 (reported by Tachibana et al., 2011). Elevated mitochondrial production of CO2 by photorespiration (only GDC is shown here for simplicity) may be scavenged by mitochondrial CA and PEPC1 (forming OAA), whereas PEPCK is down-regulated to prevent a futile cycle and promote the flux of OAA to the chloroplast. NH4+ derived from photorespiration may be sequestered as cytoplasmic carbamoyl phosphate through GS and glutamine-dependent CPScyto, presumably for pyrimidine synthesis. (b) Proposed PPDK-independent pathway to regenerate phosphoenolpyruvate from pyruvate. Plastidic pyruvate may be used to regenerate PEP through the coordinated reaction of ALAT, AGAT/SPT, HPR, GK, PGM and ENO and others (as shown in simplified form in a, where proposed reactions that regenerate glyoxylate and serine are omitted for clarity). Currently, no known homologs for GK and PPT have been identified in the T. pseudonana genome. Enzymes highlighted in (a) – and their corresponding Uniprot IDs – include alanine aminotransferase (ALAT; B8C017), alanine glyoxylate aminotransferase (AGAT; B8BZ35), bestrophins 1 and 2 (BEST1, BEST2; B8C0D8, B8C0D9), carbamoyl phosphate synthetase (CPScyto; B8C8T5), carbonic anhydrase (CA; possibly B8C025), cytosolic CA (CAcyto, CA3 or CA4; B8C250 or B8BTB4), CA6 (B8C250), mitochondrial CA (CAmito, CA8, CA9 or CA13; B8C2H3, B8BRC3 or B8C215), enolase (ENO, B8C355), glutamine synthetase (GS; B8BVI3), glycine decarboxylase (GDC P-protein; B8BX31), hydroxypyruvate reductase (HPR; B8BVI2), malate dehydrogenase (MDH; B8BQC2), PEP carboxykinase (PEPCK; B8C274), PEP carboxylase (PEPC1; B8BYW8 and PEPC2; B8C1R7), phosphoglycerate mutase (PGM; B8C354), pyruvate carboxylase (PYC; B8CE42), RuBisCO (rbcL; A8DP73 and rbcS; A0T0N5), serine pyruvate transaminase (SPT; B8BZ35) and triose phosphate transporters (TPT3; B8BSB3 and TPT4; B5YLS2). Abbreviations for substrates: CP, carbamoyl phosphate; GADP, glyceraldehyde phosphate; GLC, glycerate; GLX, glyoxylate; MAL, malate; OAA, oxaloacetate; OXO, 2-oxoglutarate; 2PGA, 2-phosphoglycerate; PEP, phosphoenolpyruvate; PGA, phosphoglycerate; PYR, pyruvate; RuBP, ribulose bisphosphate. Adenosine phosphates and reductants not shown for simplicity. New Phytologist (2014) www.newphytologist.com

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would prevent this futile CO2 production, potentially by both buffering the OAA pool and by inhibiting PEPC2 activity (Budde & Chollet, 1986). As bestrophins only facilitate diffusion, unutilized chloroplast OAA levels would lead to the net conversion of OAA in the CER/PPS to malate. On return to high light, (a)

ALAT

Ala

OXO Ala

PYR

OAA drawdown in the plastid will lead to the net conversion of malate back to OAA in the CER/PPS (and possibly the de-repression of PEPC2). This model would be entirely consistent with the ecology of marine diatoms, which are particularly adapted to turbulent conditions in which their incident light

cytoplasm

Glu RuBisCO PGA CO2 + RuBP

AGAT

PYR Ser S Gly P GLX T Ala OH-PYR HPR GLC CO2+ OAA PEP PEPCK

GLC PGA GK

PEP + HCO3– OAA PEPC1 CAmito

2PGA

Gly

GDC

PYR + HCO3–

mitochondrion NH4+ GSI Glu

PYC

OAA

BEST1/2

MAL TPT4 PEPC2 PGA MDH – PGM PEP + HCO3 OAA CO2

PEP

PPT CO2 + NH4+

plastid

CA

2PGA PEP ENO CPScyto Gln HCO3–

CA6

CAcyto CO2 HCO3– CP

pyrimidine synthesis

(b)

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12 Research

regime rapidly changes as cells transit throughout the mixed layer (Cerme~ no et al., 2008). If in vivo RuBisCO oxygenase activities represent a substantial sink for fixed carbon in marine diatoms at low CO2, a highly efficient compensatory recycling mechanism must be at play, as growth rates are independent of CO2. Others have suggested that diffusive losses of NH4+ and CO2 may be avoided by scavenging and recycling through the OUC (Parker et al., 2008; Allen et al., 2011). However, of the five OUC proteins quantified, none were up-regulated. Higher levels of mitochondrial C4 carboxylation catalyzed by PEPC1 and various CAs may serve to recover CO2 released from photorespiration, whereas the diminution of PEPCK-catalyzed C4 decarboxylation would prevent a futile CO2–C4 cycle in the mitochondria. Cytoplasmic carbamoyl phosphate synthetase (CPS) was up-regulated at low CO2, suggesting a possible recuperation of NH4+ through glutamine synthetase and CPS activity (Fig. 6). The present results support a model of diatom metabolism in which high carbon fixation and growth rates are maintained at low CO2 by elevated OAA synthesis in the CER, brought about by elevated PEPC2 abundance and elevated PEP concentrations derived from the antiport of PGA out of and PEP into the CER. Our results further indicate that the decarboxylation of OAA may be catalyzed by plastidic PYC (Fig. 6). It should be noted that this model would require and is entirely consistent with the presence of bicarbonate transporters in the plasmalemma (Nakajima et al., 2013) and ER membranes. The up-regulation of ALAT, AGAT and GDC at low CO2 presents a mechanism to recycle pyruvate and complete the proposed C4 pathway. Validation of this model will require further verification of the intracellular localizations of the various enzymes involved, as well as the quantification of their kinetic and regulatory control. These proteomic data provide some insight into both the mechanisms of carbon concentration and compensation for photorespiration in diatoms, which may contribute to their success during blooms when aqueous CO2 is drawn down to levels far below atmospheric equilibrium and well below the levels that saturate the carboxylase activity of RuBisCO.

Acknowledgements This work was supported by National Science Foundation (NSF) grants to J.R.R. and K.D.B. (OCE 0526365), A.J.M. (OCE 0526188) and A.B.K. (OCE 0927733).

References Allen AE, Dupont CL, Obornik M, Horak A, Nunes-Nesi A, McCrow JP, Zheng H, Johnson DA, Hu HH, Fernie AR et al. 2011. Evolution and metabolic significance of the urea cycle in photosynthetic diatoms. Nature 473: 203–210. Amoroso G, Sultemeyer D, Thyssen C, Fock HP. 1998. Uptake of HCO3 and CO2 in cells and chloroplasts from the microalgae Chlamydomonas reinhardtii and Dunaliella tertiolecta. Plant Physiology 116: 193–201. Attwood PV, Cleland WW. 1986. Decarboxylation of oxaloacetate by pyruvate carboxylase. Biochemistry 25: 8191–8196. New Phytologist (2014) www.newphytologist.com

New Phytologist Badger MR, Andrews TJ, Whitney SM, Ludwig M, Yellowlees DC, Leggat W, Price GD. 1998. The diversity and coevolution of Rubisco, plastids, pyrenoids, and chloroplast-based CO2-concentrating mechanisms in algae. Canadian Journal of Botany-Revue Canadienne de Botanique 76: 1052–1071. Badger MR, Price GD, Long BM, Woodger FJ. 2006. The environmental plasticity and ecological genomics of the cyanobacterial CO2 concentrating mechanism. Journal of Experimental Botany 57: 249–265. Beardall J. 1989. Photosynthesis and photorespiration in marine phytoplankton. Aquat. Bot. 34: 105–130. Bender SJ, Parker MS, Armbrust EV. 2012. Coupled effects of light and nitrogen source on the urea cycle and nitrogen metabolism over a diel cycle in the marine diatom Thalassiosira pseudonana. Protist 163: 232–251. Bendtsen JD, Nielsen H, Von Heijne G, Brunak S. 2004. Improved prediction of signal peptides: SignalP 3.0. Journal of Molecular Biology 340: 783–795. Bowes G. 2011. Single-cell C4 photosynthesis in aquatic plants. In: Raghavendra AS, Sage RF, eds. Photosynthesis and related CO2 concentrating mechanisms, advances in photosynthesis and respiration. Dordrecht, the Netherlands: Springer, 32: 63–80. Budde RJ, Chollet R. 1986. In vitro phosphorylation of maize leaf phosphoenolpyruvate carboxylase. Plant Physiology 82: 1107–1114. Burkhardt S, Amoroso G, Riebesell U, Sultemeyer D. 2001. CO2 and HCO3 uptake in marine diatoms acclimated to different CO2 concentrations. Limnology and Oceanography 46: 1378–1391. Cassar N, Laws EA. 2007. Potential contribution of beta-carboxylases to photosynthetic carbon isotope fractionation in a marine diatom. Phycologia 46: 307–314. Cerme~ no P, Dutkiewicz S, Harris RP, Follows M, Schofield O, Falkowski PG. 2008. The role of nutricline depth in regulating the ocean. Proceedings of the National Academy of Sciences, USA 105: 20344–20349. Christin P-A, Besnard G, Samaritani E, Duvall MR, Hodkinson TR, Savolainen V, Salamin N. 2008. Oligocene CO2 decline promoted C4 photosynthesis in grasses. Current Biology 18: 37–43. Colman B, Huertas IE, Bhatti S, Dason JS. 2002. The diversity of inorganic carbon acquisition mechanisms in eukaryotic microalgae. Functional Plant Biology 29: 261–270. Cooper TG, Filmer D, Wishnick M, Lane MD. 1969. The active species of “CO2” utilized by ribulose diphosphate carboxylase. Journal of Biological Chemistry 244: 1081–1083. Crawfurd K, Raven JA, Wheeler GL, Baxter EJ, Joint I. 2011. The response of Thalassiosira pseudonana to long-term exposure to increased CO2 and decreased pH. PLoS ONE 6: e26695. Edwards GE, Franceschi VR, Voznesenskaya EV. 2004. Single-cell C4 photosynthesis versus the dual-cell (Kranz) paradigm. Annual Review of Plant Biology 55: 173–196. Eisenstadt D, Barkan E, Luz B, Kaplan A. 2010. Enrichment of oxygen heavy isotopes during photosynthesis in phytoplankton. Photosynthesis Research 103: 97–103. Einsenhut M, Ruth W, Haimovich M, Bauwe H, Kaplan A, Hagemann M. 2008. The photorespiratory glycolate metabolism is essential for cyanobacteria and might have been conveyed endosymbiontically to plants. Proceedings of the National Academy of Sciences, USA 105: 17199–17204. Fielding AS, Turpin DH, Guy RD, Calvert SE, Crawford DW, Harrison PJ. 1998. Influence of the carbon concentrating mechanism on carbon stable isotope discrimination by the marine diatom Thalassiosira pseudonana. Canadian Journal of Botany-Revue Canadienne de Botanique 76: 1098– 1103. Fischer K, Arbinger B, Kammerer B, Kammerer S, Busch C, Wallmeier H, Sauer N, Eckersorn C, Ulf-lngo F. 1994. Cloning and in vivo expression of functional triose phosphate/phosphate translocators from C3- and C4-plants: evidence for the putative participation of specific amino acid residues in the recognition of phosphoenolpyruvate. Plant Journal 5: 215–226. Fischer K, Kammerer B, Gutensohn M, Arbinger B, Weber A, Hausler RE. 1997. A new class of plastidic phosphate translocators: a putative link between primary and secondary metabolism by the phosphoenolpyruvate/phosphate antiporter. Plant Cell 9: 453–462. Fl€ ugge U. 1999. Phosphate translocators in plastids. Annual Review of Plant Physiology and Plant Molecular Biology 50: 27–45. Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

New Phytologist Fl€ ugge U-I, H€a usler RE, Ludewig F, Gierth M. 2011. The role of transporters in supplying energy to plant plastids. Journal of Experimental Botany 62: 2381–2392. Govindjee 2004. Chlorophyll a fluorescence: a bit of basics and history. In: Papageorgiou GC, Govindjee , eds. Chlorophyll a fluorescence: a signature of photosynthesis. Dordrecht, the Netherlands: Springer, 1–42. Granum E, Roberts K, Raven JA, Leegood RC. 2009. Primary carbon and nitrogen metabolic gene expression in the diatom Thalassiosira pseudonana (Bacillariophyceae): diel periodicity and effects of inorganic carbon and nitrogen. Journal of Phycology 45: 1083–1092. Grouneva I, Jakob T, Wilhelm C, Goss R. 2008. A new multicomponent NPQ mechanism in the diatom Cyclotella meneghiniana. Plant and Cell Physiology 49: 1217–1225. Gruber A, Vugrinec S, Hempel F, Gould SB, Maier UG, Kroth PG. 2007. Protein targeting into complex diatom plastids: functional characterisation of a specific targeting motif. Plant Molecular Biology 64: 519–530. Hagen AR, Barabote RD, Saier MH. 2005. The bestrophin family of anion channels: identification of prokaryotic homologues. Molecular Membrane Biology 22: 291–302. Haimovich-Dayan M, Garfinkel N, Ewe D, Marcus Y, Gruber A, Wagner H, Kroth PG, Kaplan A. 2013. The role of C4 metabolism in the marine diatom Phaeodactylum tricornutum. New Phytologist 197: 177–185. Hatch MD, Slack CR. 1966. Photosynthesis by sugar-cane leaves: A new carboxylation reaction and the pathway of sugar formation. Biochemistry Journal 101: 103–111. Huttlin EL, Hegeman AD, Harms AC, Sussman MR, Subtle W. 2007. Comparison of full versus partial metabolic labeling for quantitative proteomics analysis in Arabidopsis thaliana. Molecular and Cellular Proteomics 6: 860–881. Jeffrey SW, Humphrey GF. 1975. New spectrophotometric equations for determining chlorophylls a, b, c1 and c2 in higher plants, algae, and natural phytoplankton. Biochemie und Physiologie der Pflanzen 167: 191–194. Keller M. 1988. Microwave treatment for sterilization of phytoplankton culture. Journal of Experimental Marine Biology and Ecology 117: 279–283. Kilian O, Kroth PG. 2005. Identification and characterization of a new conserved motif within the presequence of proteins targeted into complex diatom plastids. Plant Journal 41: 175–183. Kolber Z, Prasil O, Falkowski PG. 1998. Measurements of variable fluorescence using fast repetition rate techniques: defining methodology and experimental protocols. Biochimica et Biophysica Acta 1367: 88–106. Kroth PG, Chiovitti A, Gruber A, Martin-Jezequel V, Mock T, Parker MS, Stanley MS, Kaplan A, Caron L, Weber T et al. 2008. A model for carbohydrate metabolism in the diatom Phaeodactylum tricornutum deduced from comparative whole genome analysis. PLoS ONE 3: e1426. Kustka AB, Allen AE, Morel FMM. 2007. Sequence analysis and transcriptional regulation of iron acquisition genes in two marine diatoms. Journal of Phycology 43: 715–729. Lee K, Tong LT, Millero FJ, Sabine CL, Dickson AG, Goyet C, Park G-H, Wanninkhof R, Feely RA, Key RM. 2006. Global relationships of total alkalinity with salinity and temperature in surface waters of the world’s oceans. Geophysical Research Letters 33: L19605. Lewis MR, Smith JC. 1983. A small volume, short-incubation-time method for measurement of photosynthesis as a function of incident irradiance. Marine Ecology Progress Series 13: 99–102. McGinn PJ, Morel FMM. 2008. Expression and inhibition of the carboxylating and decarboxylating enzymes in the photosynthetic C4 pathway of marine diatoms. Plant Physiology 146: 300–309. Milligan AJ, Aparicio UA, Behrenfeld MJ. 2012. Fluorescence and nonphotochemical quenching responses to simulated vertical mixing in the marine diatom Thalassiosira weissflogii. Marine Ecology Progress Series 448: 67–78. Miloslavina Y, Grouneva I, Lambrev PH, Lepetit B, Goss R, Wilhelm C, Holzwarth AR. 2009. Ultrafast fluorescence study on the location and mechanism of non-photochemical quenching in diatoms. Biochimica et Biophysica Acta-Bioenergetics 1787: 1189–1197. Nakamura Y, Kanakagiri S, Van K, He W, Spalding MH. 2005. Disruption of the glycolate dehydrogenase gene in the high CO2-requiring mutant HCR89 of Chlamydomonas reinhardtii. Canadian Journal of Botany 83: 820–833. Ó 2014 The Authors New Phytologist Ó 2014 New Phytologist Trust

Research 13 Nakajima K, Tanaka A, Matsuda Y. 2013. SLC4 family transporters in a marine diatom directly pump bicarbonate from seawater. Proceedings of the National Academy of Sciences, USA 110: 1767–1772. Ogren WL. 1994. Energy utilization by photorespiration. In: Tolbert NE, Press J, eds. Regulation of atmospheric CO2 and O2 by photosynthetic carbon metabolism. Oxford, UK: Oxford University Press, 115–125. Parker MS, Armbrust EV, Piovia-Scott J, Keil RG. 2004. Induction of photorespiration by light in the centric diatom Thalassiosira weissflogii (Bacillariophyceae): molecular characterization and physiological consequences. Journal of Phycology 40: 557–567. Parker MS, Armbrust EV. 2005. Synergistic effects of light, temperature, and nitrogen source on transcription of genes for carbon and nitrogen metabolism in the centric diatom Thalassiosira pseudonana (Bacillariophyceae). Journal of Phycology 41: 1142–1153. Parker MS, Mock T, Armbrust EV. 2008. Genomic insights into marine microalgae. Annual Review of Genetics 42: 619–645. Pfaffl MW. 2001. A new mathematical model for relative quantification in real-time RT–PCR. Nucleic Acids Research 29: e45. Platt T, Gallegos CL. 1980. Modeling primary production. In: Falkowski PG, ed. Primary production in the sea. New York, NY, USA: Plenum Press, 339–362. Reinfelder JR. 2011. Carbon concentrating mechanisms in eukaryotic marine phytoplankton. Annual Review of Marine Science 3: 291–315. Reinfelder JR, Kraepiel AML, Morel FMM. 2000. Unicellular C4 photosynthesis in a marine diatom. Nature 407: 996–999. Reinfelder JR, Milligan AJ, Morel FMM. 2004. The role of the C4 pathway in carbon accumulation and fixation in a marine diatom. Plant Physiology 135: 2106–2111. Reiskind JB, Bowes G. 1991. The role of phosphoenolpyruvate carboxykinase in a marine macroalga with C4-like photosynthetic characteristics. Proceedings of the National Academy of Sciences, USA 88: 2883–2887. Riebesell U, Wolf-Gladrow DA, Smetacek VS. 1993. Carbon dioxide limitation of marine phytoplankton growth rates. Nature 361: 249–251. Roberts K, Granum E, Leegood RC, Raven JA. 2007. C3 and C4 pathways of photosynthetic carbon assimilation in marine diatoms are under genetic, not environmental, control. Plant Physiology 145: 230–235. Roberts SK, Milnes J, Caddick M. 2011. Characterisation of AnBEST1, a functional anion channel in the plasma membrane of the filamentous fungus, Aspergillus nidulans. Fungal Genetics and Biology 48: 928–938. Rotatore C, Colman B, Kuzma M. 1995. The active uptake of carbon dioxide by the marine diatoms Phaeodactylum ticornutum and Cyclotella sp. Plant, Cell & Environment 18: 913–918. Sage RF. 2002. C4 photosynthesis in terrestrial plants does not require Kranz anatomy. Trends in Plant Science 7: 283–285. Sage RF. 2004. The evolution of C4 photosynthesis. New Phytologist 161: 341–370. Sunda W, Price N, Morel FMM. 2005. Trace metal ion buffers and their use in culture studies. In: Andersen RA, ed. Algal culturing techniques. Amsterdam, the Netherlands: Academy Press/Elsevier, 35–63. Svensson P, Blasing OE, Westhoff P. 2003. Evolution of C4 phosphoenolpyruvate carboxylase. Archives of Biochemistry and Biophysics 414: 180–188. Tachibana M, Allen AE, Kikutani S, Endo Y, Bowler C, Matsuda Y. 2011. Localization of putative carbonic anhydrases in two marine diatoms, Phaeodactylum tricornutum and Thalassiosira pseudonana. Photosynthesis Research 109: 205–221. Tortell PD, DiTullio GR, Sigman DM, Morel FMM. 2002. CO2 effects on taxonomic composition and nutrient utilization in an Equatorial Pacific phytoplankton assemblage. Marine Ecology-Progress Series 236: 37–43. Voznesenskaya EV, Franceschi VR, Kiirats O, Artyusheva EG, Freitag H, Edwards GE. 2002. Proof of C4 photosynthesis without Kranz anatomy in Bienertia cycloptera (Chenopodiaceae). Plant Journal 31: 649– 662. Zelitch I, Schultes NP, Peterson RB, Brown P, Brutnell TP. 2009. High glycolate oxidase activity is required for survival of maize in normal air. Plant Physiology 149: 195–204. New Phytologist (2014) www.newphytologist.com

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14 Research

Supporting Information Additional supporting information may be found in the online version of this article. Table S1 Analysis of actin transcript abundance and intra-treatment cycle threshold variability Table S2 Thalassiosira pseudonana proteome 15N : 14N ratios grown under high CO2 with either 14N or 15N nitrate

Table S4 Thalassiosira pseudonana proteome grown under low and high CO2

15

N : 14N ratios

Table S5 Relative quantification of proteins from pathways of interest Please note: Wiley Blackwell are not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office.

Table S3 Targeting predictions and assigned cellular localization for proteins of interest

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