Diatoms are important players in the global carbon cycle. Their apparent photosynthetic affinity for ambient CO2 is much higher than that of ribulose 1,5-bisphosphate carboxylase/oxygenase (Rubisco), indicating that a CO2-concentrating mechanism (CCM) is functioning. However, the nature of the CCM, a biophysical or a biochemical C4, remains elusive. Although 14C labeling experiments and presence of complete sets of genes for C4 metabolism in two diatoms supported the presence of C4, other data and predicted localization of the decarboxylating enzymes, away from Rubisco, makes this unlikely.
We used RNA-interference to silence the single gene encoding pyruvate-orthophosphate dikinase (PPDK) in Phaeodactylum tricornutum, essential for C4 metabolism, and examined the photosynthetic characteristics.
The mutants possess much lower ppdk transcript and PPDK activity but the photosynthetic K1/2 (CO2) was hardly affected, thus clearly indicating that the C4 route does not serve the purpose of raising the CO2 concentration in close proximity of Rubisco in P. tricornutum. The photosynthetic Vmax was slightly reduced in the mutant, possibly reflecting a metabolic constraint that also resulted in a larger lipid accumulation.
We propose that the C4 metabolism does not function in net CO2 fixation but helps the cells to dissipate excess light energy and in pH homeostasis.
Diatoms play an important role in the global carbon cycle and it is estimated that they perform c. 20% of global CO2 fixation (Falkowski & Raven, 2007). Information on the uptake of inorganic carbon (Ci) and its fixation by diatoms is rather limited, as only a few model organisms such as Thalassiosira weissflogii, Thalassiosira pseudonana, Thalassiosira oceanica, and Phaeodactylum tricornutum have been examined. Nevertheless, the K1/2(CO2) of their ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco), c. 28–40 μM (Badger et al., 1998; Whitney et al., 2001), is c. 20- to 40-fold higher than the apparent photosynthetic K1/2(CO2) for ambient CO2 (Burkhardt et al., 2001) and c. three- to fourfold higher than the CO2 concentration in the present-day marine environment (Riebesell et al., 1993). Measurements indicated that in most cases Ci uptake and carbonic anhydrase (CA) activity are strongly affected by the CO2 concentrations experienced by the cells during growth. They both increase significantly with a declining ambient CO2 from a high concentration (1–5% CO2 in air) to the concentration of CO2 in air or lower, and that some of the CA encoding genes are up-regulated when the cells are exposed to low concentrations of CO2 (Tortell et al., 1997, 2008; Burkhardt et al., 2001; Morel et al., 2002; Cassar et al., 2004; Harada et al., 2005; Tanaka et al., 2005; Trimborn et al., 2008; Maberly et al., 2009; Matsuda et al., 2011; Tachibana et al., 2011). These findings have led to the recognition that, like many other phytoplankton species, the model diatoms must rely on CO2 concentrating mechanisms (CCMs) to perform significant rates of CO2 fixation under contemporary CO2 concentrations. These data also supported the notion that a CCM is activated in cells exposed to declining CO2 concentrations (see Kaplan et al., 1991, 1994; Kaplan & Reinhold, 1999; Badger et al., 2002; Giordano et al., 2005; Price et al., 2008; Fukuzawa et al., 2011 for reviews).
The nature of the CCM operating in diatoms is not clear and the extent of its expression may be species-specific and strongly affected by growth conditions (Roberts et al., 2007a,b; Reinfelder, 2011). Although suggestive, induction of genes and activities involved in Ci uptake such as CA cannot and must not be regarded as conclusive evidence for the operation of a biophysical CCM. Here, the light energy is used to actively accumulate Ci within the cells and thereby raise the CO2 concentrations in close proximity to Rubisco, the carboxylating enzyme that is mostly confined within the carboxysomes in cyanobacteria or pyrenoids in eukaryotes (Kaplan & Reinhold, 1999; Giordano et al., 2005; Espie & Kimber, 2011). The presence of pyrenoids and the localization of most of the plastidic Rubisco, together with a beta-CA (PtCA1) within these bodies (Jenks & Gibbs, 2000; Roberts et al., 2007b; Hopkinson et al., 2011; Ma et al., 2011; Matsuda et al., 2011; Reinfelder, 2011; Tachibana et al., 2011), provide additional support to the notion that a biophysical CCM is functioning in at least some of the diatoms.
On the other hand, the seminal studies of Reinfelder (Reinfelder, 2011 and references therein) suggested the existence of a plant-type biochemical C4 metabolism in diatoms. This was based on short-term 14CO2 pulse-chase labeling experiments, analyses of phosphoenol pyruvate (PEP) carboxylase (PEPC) and PEP carboxykinase (PEPCK) activities, the effects of inhibitors on the photosynthetic rate, and the effect of CO2 concentrations on gene expression (Reinfelder et al., 2000; Morel et al., 2002). However, there are opposing conclusions based on the slow rate of C4 acid accumulation in some diatoms such as T. pseudonana and constitutive expression of C4-related genes, hardly affected by the CO2 concentrations (Roberts et al., 2007a,b). The evidence for C4 metabolism in P. tricornutum rests mainly on the lower rate of photosynthesis following inhibition of PEPC and the constitutive high transcript abundance from genes encoding enzymes essential for this route (see Reinfelder, 2011 and references therein).
The classical plant C4 metabolism rests on cooperation between two cell types, with the mesophyll cell performing the first carboxylation using PEPC and followed by reduction. The C4 acid obtained is then transferred to the bundle sheath cells where decarboxylation takes place, leading to an elevated CO2 concentration in close proximity to Rubisco confined to these cells (Hatch, 1992). However, a C4 metabolism functioning in a single plant cell was already described in Hydrilla verticillata (Rao et al., 2002), Bienertia cycloptera and Borszczowia aralocaspica (Edwards et al., 2004) and was recently proposed for Ulva prolifera (Xu et al., 2012).
A plant-like C4 activity in diatoms is further supported by the presence of a complete set of genes essential for this metabolic route in T. pseudonana and P. tricornutum, for which the entire genomic sequence information is available (Armbrust et al., 2004; Bowler et al., 2008). However, in silico analyses of the subcellular localization of the relevant enzymes predicted a scrambled C4 metabolism (Kroth et al., 2008), possibly a consequence of the evolutionary history of diatoms via secondary endosymbiosis (Armbrust et al., 2004; Kroth et al., 2008; Vardi et al., 2008). Phosphoenol pyruvate carboxylase 1 (PEPC1) is predicted to be located either in the endoplasmic reticulum or in the periplastidic space between the second and the third membranes of the four-membrane-bound diatom plastid, representing the former cytoplasm of the eukaryotic endosymbiont. PEPC2, one pyruvate kinase (PK) isoform, PEPCK, malate dehydrogenase (MDH), malic enzyme (ME) and pyruvate carboxylase (PYC1) are predicted to be targeted to the mitochondria, whereas pyruvate phosphate dikinase (PPDK), another PK isoform, and PYC2 were predicted to be plastid-targeted. As a result of these subcellular localizations, for a functional plant-like C4 metabolism to occur and in the absence of a recognized plastidic decarboxylase, the CO2 generated via decarboxylation of the C4 acid in the mitochondria must cross six membranes in order to reach Rubisco in the plastid (Kroth et al., 2008). Owing to the presence of multiple forms of CAs (nine and 13 CA sequences were detected in the genomes of P. tricornutum and T. pseudonana, respectively) and their localization (Tachibana et al., 2011), the CO2 produced by decarboxylation outside the plastid is most likely being converted back to . Consequently, a futile Ci cycling would be expected where ATP is consumed during PEP formation from pyruvate.
From measurements of Ci fluxes and comparisons of the apparent photosynthetic affinity to CO2 with that of Rubisco, we conclude that a CCM is functioning in diatoms. However, we are unable to determine whether a biophysical or a biochemical CCM route is taking place. Therefore, we adopted a genetic approach using RNAi technology to down-regulate the single copy gene encoding PPDK in P. tricornutum. Regardless of whether the predicted localizations of the various enzymes potentially involved in the C4 route (Kroth et al., 2008) are correct, PPDK that catalyzes the formation of PEP from pyruvate using ATP performs an essential step for net CO2 fixation via a biochemical CCM (Hatch & Kagawa, 1973); its down-regulation thus enables us to assess the role of C4 in P. tricornutum.
Materials and Methods
Strains and media
Phaeodactylum tricornutum Bohlin (Bacillariophyceae) strain UTEX646 (available at UTEX Culture Collection of Algae, University of Texas, Austin, TX; http://www.bio.utexas.edu/research/utex/) was grown in 22°C with continuous illumination at 75 μmol photons m−2 s−1 in seawater-enriched f/2 media (Guillard & Ryther, 1962) and 2 mM Tris buffer, pH = 8.0. Solid media contained 1.2% Bacto Agar (Difco, Becton Dickinson, France). Cell cultures were maintained in Erlenmeyer flasks without any forced aeration or bubbling in order to impose extreme CO2 limitations, conditions under which various genes essential for both types of CCM are likely up-regulated. All the experiments were performed in triplicate, using at least three independent cultures of the wildtype (WT) and transformed cell lines.
Construction of plasmids and PCR
Standard cloning procedures were used for plasmid constructions (Sambrook et al., 1989). PCR was performed with a conventional thermocycler (GenePro, Bioer, Arlington, MA, USA) using a PrimeSTAR™ HS DNA polymerase (TaKaRa, Tokyo, Japan) according to the manufacturer's instructions. The transformation vector pPha-T1 (GenBank accession AF219942.1) was used (Zaslavskaia et al. 2000). It contains a she ble gene for Zeocin resistance, thus allowing screening for positive colonies of P. tricornutum and an amp gene encoding ampicillin resistance for bacterial selection and amplification. A sequence of 469 bp, sense PPDK and a sequence of 189 bp, antisense PPDK were amplified from P. tricornutum cDNA (Supporting Information, Fig. S1a) and cloned on each side of an 874-bp loop containing the eGFP gene into the pPha-T1 vector, giving rise to pPha-T1-PPDK-RNAi (Fig. S1b).
Cells were bombarded using the Bio-Rad Biolistic PDS-1000/He Particle Delivery System (Bio-Rad) fitted with 1350 psi rupture discs as described in Kroth (2007). After transformation, cells were allowed to recover for 24 h before being plated onto an f/2 medium containing 75 μg ml−1 Zeocin (Invitrogen). The plates were incubated at 22°C under constant illumination (75 μmol photons m−2 s−1).
Genomic DNA isolation
Cells were harvested by centrifugation (3000 g, 10 min) and resuspended in a lysis buffer containing 0.2 M Tris-HCl, pH = 9, 0.4 M LiCl and 25 mM EDTA. Glass beads (diameter = 212–300 μm; Sigma) were added to the cells before they were mechanically broken for 1 min in a bead beater. The cell lysate was centrifuged for 5 min at maximum speed and the supernatant was transferred to a clean Eppendorf tube containing isopropanol at equal volume, followed by a second centrifugation for 10 min at maximal speed. The DNA pellet was air-dried after an ethanol wash and resuspended in 30 μl double-distilled water (DDW). Primers, binding to the transformation vector, were used to screen for genomic integration of the RNAi construct in Zeocin-resistant colonies.
Isolation of RNA and cDNA synthesis
Fifty milliliters of logarithmic phase cells (c. 106 cells ml−1) were harvested by centrifugation at 5500 g for 3 min at room temperature. The pellet was resuspended in 1 ml phosphate-buffered saline (PBS) and centrifuged at 5500 g, 4°C, for 1 min. Cell pellets were resuspended in 1 ml RNAzol (Molecular Research Center, Inc., Cincinnati, OH, USA) and then homogenized by mechanical breakdown using acid-washed glass beads (diameter = 212–300 μm) for 20 s in a bead-beater. Further RNA isolation steps were performed according to the RNAzol protocol. Contaminating DNA was digested with Turbo-DNase (Turbo DNA-free™; Ambion) according to the manufacturer's instructions. The RNA obtained was reverse transcribed using ImProm- II™ Reverse Transcription System (Promega) according to the manufacturer's instructions. Complete removal of genomic DNA from RNA samples was verified after cDNA synthesis by quantitative PCR (qPCR) amplification of histon H4 (h4) gene.
Quantitative PCR assays
These were performed using a Rotor-Gene™ 6000 Thermal Cycler (Corbett Research, Brisbane, Australia). The primers used are shown in Table S1. Five microliters of diluted cDNA, corresponding to 10 ng total RNA, were used in the following program: DNA polymerase activation at 95°C for 15 min, followed by 40 cycles of denaturation at 95°C for 10 s, annealing at 56°C for 15 s, product elongation at 72°C for 20 s, and signal acquiring at 79°C. Amplifications were carried out in a total volume of 15 μl using the Absolute Blue SYBR Green ROX Mix (Thermo Scientific, ABgene, Rockford, IL, USA) according to the manufacturer's instructions. Transcript abundances were examined relative to the level of the gene transcript for histone 4 (h4) (Siaut et al., 2007). All samples were analyzed in three replicates per experiment and each experiment was repeated independently at least three times.
PPDK activity assay
Pyruvate phosphate dikinase activity was measured essentially as described in Ashton et al. (1990) but with several modifications. The procedures developed for the higher plant enzyme were used as a starting point because of the presence of a plant-like regulatory protein of PPDK (see the 'Results and Discussion' section). Cells maintained in growth conditions or exposed to a higher light intensity, 300 μmol photons m−2 s−1, for 1 h were broken as described earlier in extraction medium containing 50 mM HEPES-NaOH, pH 8.0, 10 mM MgCl2, 1 mM dithiothreitol (DTT), 10 mM EDTA, 1% casein, 1% polyvinylpyrrolidone, 0.25 M mannitol, 0.05% Triton X-100 and a mix of protease inhibitors 1:100 (Sigma). The assay buffer contained 100 mM HEPES-NaOH, pH 8.0, 15 mM MgCl2, 0.15 mM EDTA pH 8.0, 5 mM NaHCO3, 5 mM NH4Cl, 2.5 mM K2HPO4, 5 mM DTT and 1 mM glucose 6-phosphate. Freshly prepared 0.3 mM NADH, 1.5 mM ATP, 10.5 U MDH, 1.25 mM pyruvate and 0.5 U of PEPC were added directly to the quartz cuvette and the reaction was initiated by the addition of 20 μl of protein extract; the overall reaction volume was 1 ml. The changing absorption at 340 nm was recorded by the spectrophotometer (Cary 300 bio, Varian) and the results were normalized to the protein content in the reaction. An earlier published alternative protocol developed to assess PPDK activity in certain algae, including P. tricornutum, also used the coupling to PEPC reaction but relied on its intrinsic value (Mukerji, 1980), which might be rate-limiting for the coupled reaction.
CO2- and light intensity-dependent O2 evolution
The rate of CO2-dependent O2 evolution as a function of Ci concentrations was determined using a Clark type O2 electrode (PS2108, PASPORT dissolved O2 sensor; Pasco, Roseville, CA, USA) essentially as described in Kaplan et al. (1988). Approx. 108 cells were harvested by centrifugation for 10 min at 3000 g in a swinging-bucket rotor and resuspended in 0.5–1 ml CO2-free f/2 medium containing 20 mM Hepes. The pH was adjusted to 7.5 with saturated NaOH. Two hundred microliters of CO2-free cells were then diluted in 4 ml of the same media and incubated in the O2 electrode chamber at 22°C and 1000 μmol photons m−2 s−1. Cells were allowed to utilize the Ci in their medium until the CO2 compensation point was reached. Aliquots of NaHCO3 of known concentrations were injected to raise the Ci concentration by known increments while measuring the resulting rise in the rate of O2 concentration in the chamber. For light intensity-dependent O2 evolution measurements, cells were incubated in darkness for at least 5 min followed by exposure to increasing light of known intensities. A saturating NaHCO3 (10 mM) concentration was added to ensure that the cells were not Ci-limited.
The photosynthetic rate of cells exposed to 10 mM Ci and 500 μmol photons m−2 s−1 was measured as described earlier followed by exposure to excess light of 2000 μmol photons m−2 s−1 for 1 h. The photosynthetic O2 evolution was then measured after lowering the light intensity back to 500 μmol photons m−2 s−1. To inhibit recovery of photosystem II during photoinhibition, the cells were exposed to 100 μg ml−1 lincomycin for 15 min before exposure to excess light.
Fluorescence emitted by photosystem II was measured by pulse amplitude-modulated (PAM) kinetics using a PAM-101 (Walz, Effertlich, Germany). The light intensity (measured at the surface of the chamber) of the modulated measuring beam (at 1.6 kHz frequency) was 0.1 μmol photons m−2 s−1. White actinic light was delivered by a projector lamp at 1000 μmol photons m−2 s−1. Maximal fluorescence (Fm) was measured with saturating white light pulses of 4000 μmol photons m−2 s−1 for 1 s, with 1 min intervals.
Cells were centrifuged for 5 min at 12 000 g. The pellet was resuspended in 100 μl methanol followed by 900 μl acetone and a short vortex for a final 90% acetone extraction. The extract was centrifuged again for 10 min at 12 000 g, and the supernatant was measured in a spectrophotometer at 664, 639 and 750 nm. Chla concentration was calculated according to Jeffrey & Humphrey (1975).
Measurement of lipids content using Nile red
To estimate lipid content, 200 μl of logarithmic-phase cells were loaded in triplicate on a 96-well plate and the O.D. 750 nm was measured before staining. Ten microliters of Nile red (500 μg of 9-diethylamino-5Hbenzo[α]phenoxazine-5-one per 1 ml acetone (Sigma)), a fluorescent probe of intracellular lipids and hydrophobic domains of proteins were added to the cells. Fluorometric analysis occurred 10 min after staining using a Sequoia-Turner Model 450 Digital Fluorometer with a 485 nm narrow-band excitation filter and a 590 nm narrow-band emission filter. With this technique, cellular storage or neutral lipids display yellow-golden fluorescence (Greenspan et al., 1985; McGinnis et al., 1997).
Fourier transform infrared (FTIR) spectroscopy and macromolecular composition
For FTIR spectroscopy analysis, cells were harvested by centrifugation at 4000 g for 1 min (Centrifuge 5415D; Eppendorf, Hamburg, Germany) and washed with distilled water. After centrifugation, the pellet was freeze-dried (Christ Alpha 1-4, B. Braun Biotech International, Allentown, PA, USA) and stored at −20°C before measurement. Cells were resuspended in c. 10 μl distilled water to obtain a final cell density of 1.8 × 106 cells μl−1. A volume of 2 μl of this cell suspension was placed on a 384-well silicon microplate (with n > 5) and dried in a cabinet dryer at 40°C for at least 10 min. FTIR spectra were measured using a HTS-XT microtiterplate module (Bruker, Berlin, Germany) with a DTGS detector as described in Wagner et al. (2010). Transmission spectra were recorded in the range between 4000 and 700 cm−1 with 32 scans per sample and at a resolution of 4 cm−1. The spectra were analyzed using OPUS Lab Software (version 5.0, Bruker), corrected to the background spectra and baseline-corrected by the rubber band method. The carbohydrate, lipid and protein contents were calculated from the spectra according to Wagner et al. (2010).
Level of Rubisco active sites
The amount of Rubisco was assessed both by western analyses and by the amount of RuBP binding sites using 14C-carboxypentitolbisphosphate (CPBP) binding as described in Marcus et al. (2005). CPBP is an analog of the intermediate Rubisco's substrate ribulose 1,5-bisphosphate.
Results and Discussion
Silencing of PPDK in P. tricornutum
As indicated, we took a genetic approach to assess whether C4 metabolism plays an important role in the growth and photosynthetic performance of low-CO2-grown P. tricornutum. Recent developments of gene silencing in P. tricornutum using constructs that express either antisense or inverted repeat RNAs (De Riso et al., 2009) enable the application of this approach for down-regulation of genes in this organism. We applied RNAi technology to silence the single ppdk gene copy (Phatr v2.0 Protein ID: 21988) that encodes PPDK (Fig. S1). This enzyme is predicted to be located within the plastid (Kroth et al., 2008) where it converts pyruvate to PEP at the expense of ATP, an essential step in net CO2 fixation via the C4 route. To silence the ppdk gene in P. tricornutum, we constructed a transformation vector containing sense and antisense fragments of this gene (see the 'Materials and Methods' section for a detailed description of the plasmid construction) and biolistically transformed P. tricornutum with this construct. Colonies growing on selective plates containing Zeocin were collected for further analysis and screened for ppdk down-regulation.
Transcript abundance of ppdk and PPDK activity
Two of the Zeocin-resistant cell lines were selected for further analyses and had a significantly (70–80%) lower abundance of the ppdk transcript, as determined by qPCR. Data from one sPPDK cell line are presented as an example (Fig. 1a). In all the aspects examined here, the two mutants we investigated in parallel showed similar responses. Although various approaches were applied, we could not raise useful antisera against the P. tricornutum PPDK and thus were unable to examine whether the PPDK level is lower in the silenced transformant strains. However, as the purpose of the gene silencing was to reduce PPDK activity, we focused on this parameter, because it was more important for the sake of this study than the enzyme level. Our analysis clearly shows that the PPDK activity was much lower in the sPPDK mutants (Fig. 1b), which was the goal of our genetic transformation efforts.
In C4 plants, the activity of PPDK is strongly affected by reversible phosphorylation of a threonine residue in the active site by the bifunctional protein kinase/phosphatase PPDK regulatory (PR) protein (see Chastain et al., 2011 for a recent review and references therein). The phosphorylation of this threonine blocks PPDK activity in the dark, thereby avoiding futile ATP consumption in darkness. To the best of our knowledge, analyses of PPDK activation and/or involvement of a regulatory protein were not examined in diatoms. Analysis of the P. tricornutum genome database identified a single gene (JGI Prot-ID 49027), bearing a plastid targeting sequence, that shows significant amino acid homology (38% identity) to the plant type enzyme and contains the typical DUF299 domain of the bacterial PPDK-PR protein (Chastain et al., 2011). On the other hand, we could not detect a PPDK-PR encoding gene in the T. pseudonana genome, although other genes essential for the C4 metabolism were found. This is consistent with the suggestion that T. pseudonana does not perform a biochemical CCM (Roberts et al., 2007b). Clearly, if an active PPDK is present in T. pseudonana, it would be important to down-regulate its activity in the dark to avoid futile ATP consumption. We did not perform a detailed study on the activation of PPDK in P. tricornutum, but the presence of the plant type PPDK regulatory protein may suggest that PPDK activation in P. tricornutum is similar to that observed in the plant type protein.
Photosynthetic and growth characteristics
Since PPDK is a key player in net CO2 fixation in organisms that perform a C4 metabolism (Matsuoka et al., 2001; Edwards et al., 2004), decreasing its cellular activity by silencing the gene would be expected to lower the carbon assimilation abilities, particularly under low Ci concentrations. CO2-dependent O2 evolution was measured as an indicator of CO2 assimilation (Fig. 2a) and to assess the apparent photosynthetic affinity to Ci. In both the sPPDK transformants and the WT, the calculated K1/2(CO2) was c. 0.6 μM CO2. This value was calculated from the K1/2(Ci) obtained in Fig. 2, based on the assumption that the Ci species in the medium are at chemical equilibrium according to the experimental pH 7.5. This value is within the range reported previously (Reinfelder, 2011). The rise in photosynthetic performance with increasing light intensity was also similar in the WT and sPPDK transformed cell line (Fig. 2b). The similar photosynthetic performance (Fig. 2) in the mutants, despite the much lower PPDK activity (Fig. 1b), provides strong evidence that photosynthetic net CO2 fixation in P. tricornutum does not take the C4 route. The cultures were grown under limiting Ci level and thus must have exploited a biophysical CCM to perform efficient photosynthesis where the apparent photosynthetic affinity to extracellular CO2 is c. 50- to 70-fold higher than that of Rubisco. These data are consistent with the fact that the growth rates of the WT and sPPDK transformants were identical, even when grown without forced aeration where the sole supply of CO2 is via diffusion from the atmosphere into the flasks (Fig. S2).
The photosynthetic Vmax (at saturating CO2 concentration and light intensity) was somewhat lower in the transformants but the reasons are not clear. We did not detect significant differences in the abundance of rbcl (encoding the large subunit of Rubisco) transcripts between the WT and the sPPDK mutants (not shown) and in the amount of Rubisco. The latter was measured by CPBP binding (Fig. S3). CPBP is an analog of the intermediate Rubisco's substrate, ribulose 1,5-bisphosphate; it binds essentially irreversibly to the reaction center, thereby allowing very accurate (far better than western blots) assessment of Rubisco binding site concentration (Marcus et al., 2005). A metabolic imbalance as a result of reduced consumption of pyruvate in the sPPDK mutant, for instance, could lead to lower photosynthetic Vmax. This may be reflected in the change in cell constituents, a rise in carbohydrates and lipid levels at the expense of proteins in the sPPDK transformed cell line (Figs 3, S4), presumably because of the higher pyruvate availability.
The photosynthetic Vmax in the WT cells is c. 250 μmol O2 evolved mg−1 Chla h−1 (Fig. 2b), whereas that of PPDK activity (Fig. 1b) is significantly (c. 40-fold) lower. Please note that PPDK activity is provided per protein and that photosynthesis activity is per Chla; the ratio of Chla/total soluble protein, measured during the enzyme assays, is close to 15 in P. tricornutum. We do not present all the various modifications examined to get maximal PPDK activity (the focal point of this study was the role of the C4, not optimization of PPDK). One of the modifications applied was exposure to various light intensities before rapidly breaking the cells (see the 'Materials and Methods' section). Of the 25 conditions performed on independent cultures, the PPDK activity increased in some (up to c. 10% of the photosynthetic Vmax) after pretreatments with 300 μmol photons m−2 s−1 for 1 h. However, this activity was only a small fraction of the photosynthetic Vmax and the ratios of PPDK activities in the WT and mutant were not affected by the light treatment. However, we cannot rule out the possibility that the PPDK activity observed here is below the maximal rate in situ. Most importantly, an identical procedure was applied to both the WT and the transformed cell lines, strengthening our conclusion that PPDK activity in the latter is considerably reduced.
The role of the C4 route in P. tricornutum
The results presented here clearly indicate that the C4 route does not play a significant part in carbon acquisition in P. tricornutum. The possibility that a biophysical CCM compensates for the reduced ppdk expression in the sPPDK mutant, minimizing the effect of the gene silencing on the CO2 response curves (Fig. 2), should be considered. However, in view of the very low PPDK activity, it is unlikely that a C4 mechanism plays an important part in net CO2 fixation even in the WT. Nevertheless, a significant amount of 14C fixation into C4 acids has been observed in several diatoms, and inhibition of PEPC lowered the rate of photosynthesis in P. tricornutum (see Reinfelder, 2011 and references therein). This raises the question of the biological role of the C4 route in P. tricornutum, if any. In view of the subcellular localization of the proteins involved (Kroth et al., 2008), it is possible that the C4 route is a futile cycle dissipating excess light energy and ATP (see the 'Introduction'). To examine possible involvement in protection against photoinhibition, P. tricornutum WT and sPPDK transformed cells were exposed to 2000 μmol photons m−2 s−1 for 1 h (c. threefold higher than required to saturate the photosynthetic rate). This was followed by measurements of O2 evolution at saturating Ci concentrations. As a result of this treatment, the rate of photosynthesis declined to 85 and 82% (from the initial rate) in the WT and sPPDK mutants, respectively. Treatments with 100 μg ml−1 lincomycin, which inhibits protein synthesis in plastids and thereby replacement of damaged D1 protein in the photosynthetic reaction center II (the main reason for photoinhibition), caused a faster decline of the photosynthetic rate. It reached 40 and 30% in the WT and the sPPDK transformant, respectively, within 1 h. Clearly, the sPPDK mutant is somewhat more sensitive to excess light than the WT, but the differences are very small.
It should be noted that diatoms possess an array of mechanisms that enable them to cope with varying abiotic factors such as excess light or fast-changing light intensities in the water body (Lavaud et al., 2007; Eisenstadt et al., 2008; Wu et al., 2011; Lepetit et al., 2012). Our measurements (Fig. 4) showed that the rate of fluorescence quenching (as indicated by the slope of the fluorescence decline from the maximal obtained after turning on the actinic light) is faster in the sPPDK mutant than in the WT, presumably through nonphotochemical quenching (NPQ). This may indicate that the transformed cell lines are able to compensate for the reduced ability to dissipate excess energy by the PPDK-dependent C4 route. Such mechanisms may involve the activation of the xanthophyll cycle and reaction center events, both leading to the apparent rise in NPQ (Eisenstadt et al., 2008; Lepetit et al., 2012) and possibly minimizing the expected photoinhibitory damage in the transformed cell line. Alternatively, the cells may use the PEPCK route of C4, whereby decarboxylation of oxaloacetate leads to the formation of PEP with little, if any, net CO2 fixation but significant energy dissipation.
The extent of up- or down-regulation of genes presumably involved in one of the CCM modes following a shift in CO2 concentration was used as supporting criteria for the occurrence of C4 in diatoms (see Reinfelder, 2011 and references therein), but this may not be the case. One example is the significant rise in the PEPC transcript abundance shortly after an upshift of the pH in the culture media of T. pseudonana from 7.6 to 8.5, in an attempt to lower the concentration of dissolved CO2. Whether these data demonstrate a rapid up-regulation of PEPC by the lower CO2 concentration, as proposed by Reinfelder (2011), remains to be elucidated. One possibility is that the rise in PEPC transcript and in the activity of the C4 route is the result of the increasing ambient pH. Formation and conversion of dicarboxylic acids could serve as important means of pH homeostasis (Raven et al., 1990). This may be particularly necessary under conditions where the cells are intensively cycling Ci species between the cytoplasm and the surrounding media (Tchernov et al., 1997, 2003). The extent of this cycling is strongly affected by the ambient conditions and by the extent to which they were acclimated to low concentrations of CO2 (and induced the Ci transport capabilities; Fukuzawa et al., 2011). As an example, Ci cycling rises significantly with light intensity and may reach values much higher than the photosynthetic rate and thereby causing a very large load on pH homeostasis (Tchernov et al., 1997, 2003).
This study was supported by the German-Israel Science Foundation (GIF; A.K. and P.G.K.), the Deutsche Forschungsgemeinschaft (German Research Foundation, DFG) trilateral program (A.K.), the Israel Science Foundation (ISF; A.K.), and by DFG grant number INST 268/149-1. The authors further thank Doris Ballert for help in biolistic transformation and the Universität Konstanz for financial support.