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Environmental Microbiology (2014)

doi:10.1111/1462-2920.12638

Does 2-phosphoglycolate serve as an internal signal molecule of inorganic carbon deprivation in the cyanobacterium Synechocystis sp. PCC 6803?

Maya Haimovich-Dayan,1 Judy Lieman-Hurwitz,1 Isabel Orf,2 Martin Hagemann3 and Aaron Kaplan1* 1 Department of Plant and Environmental Sciences, Edmond J. Safra Campus, The Hebrew University of Jerusalem, Jerusalem 91904, Israel. 2 Applied Metabolome Analysis, Max Planck Institute of Molecular Plant Physiology, Am Mühlenberg, Potsdam-Golm 114476, Germany. 3 Institut für Biowissenschaften, Abt. Pflanzenphysiologie, Universität Rostock, Einsteinstraße 3, Rostock D-18059, Germany. Summary Cyanobacteria possess CO2-concentrating mechanisms (CCM) that functionally compensate for the poor affinity of their ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) to CO2. It was proposed that 2-phosphoglycolate (2PG), produced by the oxygenase activity of Rubisco and metabolized via photorespiratory routes, serves as a signal molecule for the induction of CCM-related genes under limiting CO2 level (LC) conditions. However, in vivo evidence is still missing. Since 2PG does not permeate the cells, we manipulated its internal concentration. Four putative phosphoglycolate phosphatases (PGPases) encoding genes (slr0458, sll1349, slr0586 and slr1762) were identified in the cyanobacterium Synechocystis PCC 6803. Expression of slr0458 in Escherichia coli led to a significant rise in PGPase activity. A Synechocystis mutant overexpressing (OE) slr0458 was constructed. Compared with the wild type (WT), the mutant grew slower under limiting CO2 concentration and the intracellular 2PG level was considerably smaller than in the wild type, the transcript abundance of LC-induced genes including cmpA, sbtA and ndhF3 was reduced, and the OE cells acclimated slower to LC – indicated by the delayed rise in the apparent photosynthetic affinity to inorganic carbon. Data obtained here implicated 2PG in Received 10 July, 2014; revised 19 September, 2014; accepted 22 September, 2014. *For correspondence. E-mail aaron.kaplan@ mail.huji.ac.il; Tel. 972 2 6585234; Fax 972 2 6584463.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd

the acclimation of this cyanobacterium to LC but also indicated that other, yet to be identified components, are involved. Introduction The affinity of the cyanobacterial CO2-fixing enzyme ribulose-1,5-bisphosphate (RuBP) carboxylase/ oxygenase (Rubisco) to CO2, K1/2(CO2), approximately 200–300 μM, is about 20 to 30-fold lower than the CO2 concentration at equilibrium with water (Badger, 1980; Kaplan et al., 1988). To compensate for this difference, cyanobacteria possess a CO2-concentrating mechanism (CCM) which raises the concentration of CO2 in close proximity to Rubisco, mostly located within the carboxysomes together with carbonic anhydrase (Reinhold et al., 1989; Reinhold et al., 1991; Raven, 1991; Schwarz et al., 1995; Kaplan and Reinhold, 1999; Giordano et al., 2005; Badger et al., 2006; Fukuzawa et al., 2011; Cameron et al., 2013; Mangan and Brenner, 2014). Low activity of the CCM is observed in cyanobacterial cells grown under high levels of CO2 (HC, 1–8% CO2 in air). The CCM activity is strongly upregulated in cells exposed to a limiting CO2 level (LC) such as in equilibrium with CO2 in air, under high illumination, or photomixotrophic growth conditions (Hihara et al., 2001; Haimovich-Dayan et al., 2011). In Synechocystis sp. strain PCC 6803 (hereafter Synechocystis), often used as a model system to study cyanobacterial biology, two systems that convert CO2 to HCO3− are located on the thylakoid membrane, one constitutive and the other induced under LC (Shibata et al., 2001; Ogawa and Kaplan, 2003). In addition, three HCO3− transporters located in the cytoplasmic membrane, two of them induced under LC, were recognized (Shibata et al., 2001; Price et al., 2004). The activities of these systems lead to a large rise in the cytoplasmic inorganic carbon (Ci) level well above that expected at chemical equilibrium. Earlier studies on the acclimation of Synechocystis from HC to LC examined the transcript abundances after shorter (McGinn et al., 2003; Wang et al., 2004; Burnap et al., 2013) or similar (Eisenhut et al., 2007) duration under LC to those used here (Fig. 7, below). The data revealed large variability in the conditions applied and

2 M. Haimovich-Dayan et al. consequently the results obtained in the various studies. One example: when the cells were transferred from 3% CO2 in air to CO2-free air, the rise in the transcript abundances of LC-induced genes was completed in less than 2 h (McGinn et al., 2003) but took over 3 h in cells aerated with air (Wang et al., 2004). Although the components involved in the CCM operation have been largely recognized, the nature of the signal sensed by the cells when exposed to a limiting CO2 level is largely unknown and this subject has attracted considerable attention. It was shown over 30 years ago (Marcus et al., 1983) that the acclimation of Anabaena variabilis to LC is strongly affected by the expected rate of the oxygenation reaction of Rubisco under various CO2 and O2 levels. These data supported the possibility that the level of a product of the oxygenation reaction, 2PG, or a metabolite downstream is involved in the acclimation to LC. Two LysR type transcription factors, CcmR (NdhR) and CmpR, and CyAbrB (Sll0822) were implicated in the acclimation of cyanobacteria to low CO2 conditions and the induction of the CCM (Figge et al., 2001; Nishimura et al., 2008; Lieman-Hurwitz et al., 2009; Burnap et al., 2013). Studies by Omata and colleagues (Takahashi et al., 2004; Nishimura et al., 2008) revealed that the binding of the activator CmpR to the promoter of cmp operon (which encodes for ABC-type HCO3− transporter) in Synechococcus sp. PCC 7942 was strengthened significantly by the presence of RuBP or 2-phosphoglycolate (2PG). The levels of these metabolites, expected to rise after transfer of HC-grown cells to LC conditions, may thus act as signal molecules regulating LC induction. Studies by Burnap and colleagues (Wang et al., 2004; Daley et al., 2012; Burnap et al., 2013) on the role of another LysR regulator, CcmR acting as repressor, showed that NADP+ and α-ketoglutarate enhance the binding of CcmR to its own promoter as well as to the DNA region upstream of the operon encoding ndhF3/ndhD3/cupA/sll1735. They also confirmed, in Synechocystis, the impact of 2PG or RuBP additions on the CmpR binding to the cmp operon. Inactivation of ccmR alters the expression of about 20 different genes. Under HC and low illumination conditions, CcmR represses their expression, particularly of those genes that use redox energy to raise the internal Ci concentration such as ndhF3 and sbtA. In cyanobacteria, 2PG is metabolized via three routes initiated by the activity of phosphoglycolate phosphatase (PGPase, hydrolysing 2PG to produce glycolate and Pi). These routes are a plant-like photorespiratory C2 cycle, a bacterial glycerate pathway and a complete decarboxylation via oxalate (Eisenhut et al., 2008; Hagemann et al., 2013). Otherwise 2PG inhibits triose phosphate isomerase in the Calvin-Benson cycle (Norman and Colman, 1991) and phosphofructokinase

(Kelly and Latzko, 1976). Inactivation of these routes led to HC-requiring phenotypes, indicating the importance of 2PG metabolism for the well-being of these organisms (Eisenhut et al., 2008; Hagemann et al., 2013). It was assumed that the activation of the CCM largely eliminates cyanobacterial photorespiration due to the large rise in the intracellular Ci level (Price et al., 1998; Kaplan and Reinhold, 1999; Giordano et al., 2005). However, in an earlier study, we showed the accumulation of glycolate in Synechocystis even when exposed to 3% CO2 in air (Eisenhut et al., 2008). A slower acclimation from HC to LC was observed in a mutant of Synechococcus PCC 7942 overexpressing a PGPase from Ralstonia eutropha (CbbZ) (Kaplan et al., 1998), possibly due to a lower 2PG level, but this was not examined. Thus clear physiological/ biochemical evidence for the role of 2PG in the acclimation of cyanobacteria to LC is missing. Previous studies on mutants impaired in a single gene encoding PGPases showed a HC-requiring phenotype in the green alga Chlamydomonas reinhardtii and two higher plants, Arabidopsis thaliana and Hordeum vulgare (Somerville and Ogren, 1979; Hall et al., 1987; Suzuki et al., 1990; Schwarte and Bauwe, 2007). In sum, the levels of four metabolites, NADP+, α-ketoglutarate, RuBP and 2PG, were implicated in the regulation of two LysR transcription factors and thereby the CCM induction. Of these metabolites, it is likely that only the level of 2PG can be modified without severe consequences to basal cell metabolism. To examine the role of 2PG, we chose to modify its intracellular level by altering the activity of PGPase and examine the consequences on the transcript abundance of LC-induced genes, growth and photosynthetic parameters. Results and discussion Identification of potential phosphoglycolate phosphatase (PGPase) encoding genes in Synechocystis To investigate the role of 2PG in the acclimation of Synechocystis to LC, we aimed at interfering with its internal concentrations. Since 2PG is impermeable to cyanobacterial cells (Marcus et al., 1983), we could not alter its internal level by extracellular supplementation and thus focused on its metabolism in Synechocystis (Eisenhut et al., 2008; Hagemann et al., 2013). The first step in the 2PG degradation is dephosphorylation by PGPase to form glycolate. Earlier studies partly characterized the activity and properties of PGPases from various microorganisms, including cyanobacteria and green algae (Husic and Tolber, 1985; Norman and Colman, 1991), but to the best of our knowledge, the first sequenced and cloned microbial PGPase encoding gene, cbbZ, originated from R. eutropha (formerly Alcaligenes eutropha) (Schäferjohann et al., 1993). Biochemically

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

Cyanobacterial acclimation to changing CO2 level

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Fig. 1. Alignment of putative PGPases: Multiple sequence alignment of the hypothetical PGPases encoding genes in Synechocystis PCC 6803 was produced using the Clustal Omega program (http://www.ebi.ac.uk/Tools/msa/clustalo/). CbbZ from Ralstonia eutropha was used as a reference of a known PGPase; the three conserved regions are highlighted.

purified cyanobacterial PGPase was found to be fairly specific towards dephosphorylation of 2PG (Norman and Colman, 1991; 1992) but, as mentioned below, other phosphatases may be able to degrade 2PG as well. To modify the internal 2PG level in Synechocystis by cleavage, we first identified the relevant PGPase encoding genes; our search using CbbZ identified four candidates in Synechocystis – slr0458, sll1349, slr0586 and slr1762. Noticeably they show very low similarities to CbbZ and to one another, less than 30% in amino acid sequence (Fig. 1 and Fig. S1), but all contain conserved regions of the haloacid dehalogenase (HAD) – like superfamily that comprise L-2-HAD, epoxide hydrolase, phosphoserine phosphatase, phosphomannomutase, PGPases, P-type ATPase and many others. Three noticeable motifs are observed in the HAD-like family. Motif I: DX(D/T/Y)X(T/ V)(L/V); motif II: (S/T) and motif III: K-(G/S)(D/S)XXX(D/N) (Aravind et al., 1998; Collet et al., 1998). In motif I, the first Asp is the most conserved one in the family; the second Asp exists in PGPases and varies among other members of the family. In all the PGPases that we have identified, the second Asp residue exists in the first motif, whereas in motif II, they possess Thr residue with the exception of slr1762 bearing a Ser residue.

Expression and PGPase activity in Escherichia coli We did not aim to study the entire PGPase gene family but focused on Slr0458 since it shows the highest homology to CbbZ from R. eutropha. Furthermore, for some reason,

sll1349, which is almost as homologous to cbbZ, did not express well in E. coli. To examine whether slr0458 encode PGPases, we expressed it in E. coli. Total protein extracts from recombinant cells showed a significant rise in PGPase activity using its specific substrate 2PG (Fig. 2). However, the activity declined completely within 10 h even under 4°C. It was necessary to perform the entire procedure from cell harvesting to activity assays within a few hours and, thus, we do not know how high PGPase activity in vivo was. Compared with control, the extract from E. coli cells expressing the cyanobacterial

Fig. 2. PGPase activity of slr0458: The hypothetical PGPase slr0458 from Synechocystis PCC 6803 was overexpressed in E. coli strain BL-21 RIL using the His-tag parallel 3 plasmid. A crude extract of the E. coli cells was analysed for PGPase activity with His-tag slr0458 insert or without slr0458 insert, bearing only his-tag parallel 3 plasmid (NIC). Pi concentrations were determined using the Ames reagent.

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Fig. 3. The transcript abundance of slr0458 in the WT and the OE mutant. The abundances were normalized to the level of the reference gene rnpB in each sample and to the level of the transcript in HC-grown WT.

PGPase did not show a rise in phosphatase activity using 4-methylumbelliferyl phosphate (4-MUP, Sigma-Aldrich, MO, USA), commonly applied as a substrate to examine various phosphatase activities (Bar-Yosef et al., 2010). Although we did not test a large array of potential substrates, these data confirmed that slr0458 encodes for PGPase activity and suggested rather high specificity towards 2PG as also described for the purified PGPase from Chlamydomonas reinhardtii (Husic and Tolber, 1985). A mutant overexpressing slr0458 encoding a PGPase: transcript abundance and growth In order to examine a possible role of 2PG in the acclimation of Synechocystis to LC, we raised a mutant where

controlled overexpression (hereafter OE) of slr0458 was obtained by cloning it under the high-light-induced promoter region of psbA2. Quantitative polymerase chain reaction (PCR) was used to assess the transcript abundance of slr0458 (Fig. 3). The transcript levels of the other putative PGPase encoding genes in this strain are shown in Fig. S2. To induce slr0458 transcription, OE cells grown under HC and a light intensity of 35 μmol photons m−2 s−1 were transferred to a higher illumination, 95 μmol photons m−2 s−1 for 2 h, identical to that experienced by the WT during growth. Aliquots were transferred to LC under the same light intensity and temperature conditions, for various durations, prior to RNA extraction. In the WT, the transcript abundance of slr0458 increased twofold to threefold after transfer from HC to LC (Fig. 3). In HC-grown OE mutant, the slr0458 transcript level was 800-fold higher than in HC-grown WT but contrary to the WT, in the OE mutant, the transcript abundance declined significantly by about 10-fold within 6 h of exposure to LC (Fig. 3). These data showed that the OE mutant indeed overexpress the slr0458 but also suggested activation of a mechanism whereby its level is thereafter downregulated. Nevertheless, the slr0458 transcript level remained considerably higher in the OE mutant than in WT cells under all the growth conditions examined here. Similar growth rates were obtained for the WT and the OE mutant under HC but the mutant grew significantly slower when exposed to LC (Fig. 4). If the 2PG level serves as the signal for the induction of the CCM and thus acclimation of HC-grown cells to LC, a slower 2PG accumulation due to faster de-phosphorylation in the OE mutant could have slowed the growth of this mutant under LC. However, the reasons for a prolonged slowing of growth as observed here, despite significant physiological

Fig. 4. Growth curve of Synechocystis WT and the OE mutant at HC (A) and LC (B). Growth experiments were performed in the initial OD730 nm of 0.05. Mean values represent at least three independent experiments.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

Cyanobacterial acclimation to changing CO2 level

Fig. 5. Phosphoglycolate phosphatase activity in the WT and OE mutant. The activity was measured in protein extracts at HC after 1.5 h and 3 h of treatment of LC. Pi concentrations were calculated using a calibration curve of Pi and normalized to protein concentrations. Mean values represent at least three independent experiments.

acclimation to LC (see below), are unknown. Alternative explanations are provided in the Appendix S1. PGPase activity and the levels of 2PG and glycolate Despite many attempts, we were unable to raise a mutant lacking PGPase activity. Apparently, the cells produced 2PG even under very high level of CO2 such as used here. The purpose of slr0458 overexpression in the OE mutant was to raise the PGPase activity and thereby lower the 2PG level and its expected rise after transfer of HC-grown cells to LC conditions. In their pioneering study,

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Norman and Colman (1991) measured the activity of PGPase in Coccochloris peniocystis but a protocol to optimize the assessment of the activity of this essential enzyme in cyanobacteria is missing. As indicated, optimization of the PGPase assay was not the focal point of the present study but, following numerous experiments using various combinations of extraction and activity media, the maximal in vitro measured PGPase activities observed here were about 4 μmol Pi released mg−1 protein h−1 in HC grown WT and it increased slightly in cells transferred to LC. The PGPase activity was higher in HC-grown OE cells, up to about 7.5 μmol Pi released mg−1 protein h−1 after exposure to LC (Fig. 5). Although these activities are about fourfold higher than previously reported (Norman and Colman, 1991), it may not be large enough to dephosphorylate the 2PG formed when Synechocystis cells are transferred from a HC to a LC environment. This is based on the kinetic properties of Rubisco and the expected rate of 2PG formation at the given CO2 and O2 levels, calculated as in the study of Marcus and colleagues (1983), see Appendix S1. Based on the following three observations, we propose that the PGPase activity in vivo is significantly higher than observed in vitro and that it is probably under strict control: (i) In both E. coli Synechocystis, the observed PGPase activity decayed very fast after extraction. (ii) Compared with the large rise in slr0458 transcription in the OE mutant, the change in PGPase activity is negligible. (iii) The fact that 2PG was hardly detected in the OE mutant but was abundant in the WT (Fig. 6, below), whereas there was little difference in the observed PGPase activities between the two, supports the suggestion that the activities measured in vitro do not represent that in vivo. Clearly, additional research

Fig. 6. 2PG (A) and glycolate (B) levels in the WT and OE mutant. Time-scale experiment at LC of the three strains is presented (OE-slr0458 OE). The response was normalized to an internal standard (Sorbitol) and optical density at 730 nm. The Δ represents a significant change in comparison to WT using the Wilcoxon rank-sum test (P < 0.05). Mean values represent at least three independent experiments.

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M. Haimovich-Dayan et al. Fig. 7. Transcript abundance of genes involved in the CO2-concentrating mechanism (CCM). Quantitative PCR of cmpA (A), sbtA (B) and ndhF3 (C) was analysed at 1.5 h, 6 h and 24 h after transferring the cells from HC to LC. These genes, involved in CO2 and in bicarbonate uptake, were examined as an indication of CCM expression. Expression levels were calculated relative to the reference gene rnpB, and to the HC-grown WT. The levels of these genes were up to twofold higher in high CO2-grown OE mutant cells. Mean values represent at least three independent experiments.

is necessary to clarify the reasons and the mechanisms involved but this was not the purpose of this study aimed at reducing the internal level of 2PG. The metabolite levels were examined using gas chromatography coupled to electron impact ionization/time-offlight mass spectrometry (GC-EI/TOF-MS). It is important to note that the data (Fig. 6) were normalized to the level of each metabolite observed in HC-grown WT cells just before transfer to LC. In the WT, the 2PG level increased by about 50% within 10 min after exposure of the cells to LC and then started to decline, reaching a very low level at steady state LC conditions (1440 min time point). In contrast, we could hardly detect 2PG in the OE mutant even after transfer of HC-grown cells to LC (Fig. 6). Correspondingly, relatively higher amounts of glycolate, the PGPase reaction product, were observed in the OE mutant than in the WT (Fig. 6). These data confirmed that, unlike the WT, the PGPase activity in the OE mutant is sufficient to dephosphorylate the forming 2PG despite the rather similar enzyme activity levels measured in crude extracts (see Fig. 5). Given the relatively high level of CO2 during growth, 8% CO2 in air and the kinetic properties of cyanobacterial Rubisco (Badger, 1980), it is likely that the oxygenase activity of Rubisco was largely depressed under steady state HC conditions and that the 2PG level was low (Fig. 6A). This is consistent with our earlier observation of a low 2PG level in HC Synechocystis cell and the rising level during 3 h exposure to LC (Schwarz et al., 2014). Most likely the relatively low 2PG and glycolate levels observed under steady state LC, compared with that observed shortly after exposure of HC grown cells to

LC, reflect the induction of the CCM leading to an elevated CO2 level in close proximity of Rubisco within the carboxysomes thereby depressing the oxygenase activity. Expression of LC-induced genes Earlier in gel studies showed that 2PG affects the binding of CmpR but not of CcmR to promoters of CCM-related genes (Takahashi et al., 2004; Nishimura et al., 2008; Daley et al., 2012; Burnap et al., 2013). It was therefore expected that the lowered 2PG level in the OE mutant would inhibit the transcription of cmpA (regulated by CmpR) but not of sbtA and ndhF3 controlled by CcmR. However, this was clearly not the case (Fig. 7). In agreement with earlier reports, the transcript abundance of cmpA, ndhF3 and sbtA examined here, representing the high affinity CO2 uptake and HCO3− transport systems, increased significantly in the WT, although with different time courses for the various genes. The sbtA was the first to respond, followed by cmpA and ndhF3 (see also McGinn et al., 2003; Wang et al., 2004; Eisenhut et al., 2007; Burnap et al., 2013). In the OE mutant on the other hand, 1.5 h after the transfer from 8% CO2 to LC the transcript abundance of cmpA was lower than in the WT but thereafter, at the 6 h time point, it was much higher (Fig. 7A). Contrary to our expectation the transcript, abundances of sbtA and ndhF3 (Fig. 7B and C, respectively) were much lower in the OE mutant than in the WT. There are numerous possible explanations of this contrast and, as much as it is tempting to speculate about it, at this time, we do not know the reason. We can only propose that in

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Cyanobacterial acclimation to changing CO2 level Table 1. The photosynthetic Vmax (μmol O2 evolved × mg chlorophyll × h−1) and apparent affinity to extracellular inorganic carbon of WT and OE mutant grown under HC and after exposure to LC for various duration. −1

WT

OE

HC 1.5 h LC 6 h LC 24 h LC HC 1.5 h LC 6 h LC 24 h LC

Vmax

K1/2 (μM Ci)

243 ± 22 390 ± 12 317 ± 5 387 ± 7 337 ± 12 285 ± 54 283 ± 0.17 260 ± 73

219 ± 7 116 ± 11 74 ± 14 26 ± 3 190 ± 10 135 ± 4 188 ± 2 49 ± 10

The cells were placed in the O2 electrode chamber and allowed to reach O2 compensation point, utilizing the Ci in the medium. They were then provided with known concentrations of NaHCO3 as their Ci source, and O2 evolution was measured. The data are from three biologically independent experiments.

reality the regulation of the LysR type transcription factors is far more complex than our initial assumption and that additional component(s) must be involved.

Photosynthetic performance One of the main features of acclimation to LC is the marked rise in the apparent photosynthetic affinity of the cells to extracellular Ci, reflecting the induction of a whole set of genes (Kaplan and Reinhold, 1999; Giordano et al., 2005; Badger et al., 2006; Eisenhut et al., 2007; Fukuzawa et al., 2011). Data presented in Table 1 summarize the photosynthetic parameters during acclimation of the WT and the OE mutant from HC to LC. The initial apparent affinity of the OE mutant to Ci was slightly higher (lower K1/2 Ci), for an unknown reason, but the rate of its acclimation to LC was significantly slower than the WT. This is most pronounced during the first 6 h. Thereafter, with the rising transcript abundances of the relevant genes (Fig. 7) the OE mutant was able to close the gap. The physiological analyses (Table 1) showed that under our growth conditions, 8% CO2 in air, the acclimation of the cells to LC was slower than from 3% CO2 to CO2-free air (McGinn et al., 2003). It suggested that through its effect on the expression of LC-induced genes, the 2PG level affects the acclimation to LC. But, it also showed that when the cells are deprived of CO2 – the primary photosynthetic electron acceptor – they activate other means, the nature of which is unknown to upregulate the performance of the CCM.

Conclusions In conclusion, reducing the level of 2PG in Synechocystis by means of overexpression of a gene encoding PGPase altered the abundance of LC-induced genes and slowed

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the acclimation to LC, thus implicating 2PG in this process. The fast shift from a very high CO2 level to CO2 deprivation is stressful and the cells activate an array of responses. We draw the attention of the reader to the following four observations that support this notion. (i) The apparent affinity of HC-grown OE mutant to CO2 is somewhat higher than observed in the WT (Table 1), suggesting that the OE cells might be at least partly acclimated to LC even under 8% CO2. (ii) Analysis of the ΔccmM mutant that accumulates 2PG and glycolate already at HC does not show the expected transcriptional changes in CmpR (or CcmR) controlled genes (Hackenberg et al., 2012). (iii) Mutant E1 of Anacystis nidulans (also impaired in the structural organization of the carboxysomes) accumulates the 42 kDa polypeptide (thereafter identified and designated CmpA) under 0.3% CO2, a concentration where the WT showed typical HC characteristics (Omata et al., 1987). (iv) Earlier studies revealed the involvement of Sll0822, an AbrB homologue, in the acclimation of Synechocystis to LC and nitrogen status (Ishii and Hihara, 2008) and it remains to be seen whether it is also involved in the response to 2PG level. Finally, an earlier study showed that a mutation in Synechocystis’s glycolate dehydrogenases led to an increased accumulation of glycolate and of the number of carboxysomes when grown in HC, despite marginal changes in the expression of CCM-related genes (Eisenhut et al., 2007). It remains to be seen whether, in addition to 2PG, the elevated glycolate level due to PGPase activity could partly explain the differences in the rate of acclimation to LC between WT and the OE mutant. Experimental procedures Growth conditions and mutant construction The glucose-tolerant strain of Synechocystis was used in this research. The cells were grown at 30°C with continuous illumination of 95 μmol photons m−2 s−1 in BG11 medium (Stanier et al., 1971) supplemented with 20 mM HepesNaOH pH 7.8. Liquid cultures were bubbled with air enriched with 8% CO2 (high CO2, HC) or air (low CO2, LC). Mutant OE that overexpresses slr0458 was constructed by inserting it into pT7Blue plasmid bearing a kanamycin-resistant cassette, 300 bp of the promoter region of psbA2 from Synechocystis and the coding region of slr2031. The psbA2 promoter is fused to slr2031 at the initial codon where NdeI site was introduced and an HpaI site at position 277 bp downstream. Notably, the psbA2 is strongly upregulated under high light and was therefore used to raise the level of Slr0458 in the OE mutant. However, since the promoter also function to some extent under low light, all the data shown here with respect to the OE mutant are after exposure to high light. The double mutant was created by insertion of a kanamycinresistant cassette into slr0458 and a spectinomycin-resistant cassette into sll1349. Complete segregation of the mutant was confirmed by PCR.

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Induction of PGPase in E. coli The slr0458 was amplified from genomic DNA of the WT and cloned into his-tagged plasmid parallel 3 that contain N-terminal His (pET22). Transformations were carried out into E. coli strain BL21 (DE3) using CodonPlus-RIL (Stratagene, USA) in order to provide the synthesis of unique amino acids. Inductions of PGPase was performed in autoinduction medium in 37°C overnight, and cell free protein extracts were achieved by sonication in an extraction buffer containing 20 mM Tris-HCl pH 7.5, 10 mM MgCl2, 1 mM dithiothreitol, 1:100 protease inhibitor cocktail (Sigma). For the PGPases assay, protein extracts were dialysed against the PGPases activity assay buffer (40 mM MOPS pH 7, 10 mM MgCl2 at 4°C). The protein levels were quantified by the Bradford method.

Sampling procedure and extraction for metabolite profiling A modified version of the protocol described previously (Eisenhut et al., 2008) was used for sampling procedure and extraction. Samples of 10 ml cells corresponding to about OD730nm = 0.5 were withdrawn from the cultivation vessels and the cells were separated from the medium by filtration (0.45 μm nitrocellulose filters, Sigma-Aldrich Chemie GmbH, Munich, Germany) in light without any subsequent washing. The filters bearing the cells were placed into 2 ml Eppendorf tubes, frozen in liquid nitrogen and stored at −80°C. Metabolites were extracted by premixing with 630 μl methanol, 60 μl non-adeconoic acid methylester (2 mg ml−1 chloroform), 30 μl of 13C-sorbitol (0.2 mg ml−1 in methanol) and D4-2,3,3,3Alanine (1 mg ml−1, in water). Each sample was mixed thoroughly at least 1 min and incubated in a ultra-sonification bath for 5 min for complete suspension, agitated 15 min at 70°C, brought to room temperature, mixed with 400 μl chloroform and agitated again 5 min at 37°C. Finally, phase separation was induced by 800 μl distilled water and the upper polar fraction retrieved by 5 min centrifugation at 14 000 rpm using an Eppendorf 5417 microcentrifuge. A total volume of 600 μl was concentrated and dried in 1.5 ml microtubes by 12 to 18 h of vacuum centrifugation.

Metabolite profiling analysis This was performed as detailed previously (Wagner et al., 2003; Erban et al., 2007) by GC-EI/TOF-MS. An Agilent 6890N24 gas chromatograph (Agilent Technologies, Böblingen, Germany) with splitless injection onto a FactorFour VF-5 ms capillary column, 30 m length, 0.25 mm inner diameter, 0.25 μm film thickness (Varian-Agilent Technologies) was connected to a Pegasus III time-of-flight mass spectrometer (LECO Instrumente GmbH, Mönchengladbach, Germany). Metabolites were methoxyaminated and trimethylsilylated manually prior to GC-EI/TOF-MS analysis (Erban et al., 2007). Retention indices were calibrated by addition of a C10, C12, C15, C18, C19, C22, C28, C32 and C36 n-alkane mixture to each sample (Strehmel et al., 2008). GC-EI/TOF-MS chromatograms were acquired, visually controlled, baseline corrected and exported in NetCDF file format using ChromaTOF software (Version 4.22; LECO, St.

Joseph, USA). GC-MS data processing into a standardized numerical data matrix and compound identification were performed using TagFinder software (Luedemann et al., 2008; Allwood et al., 2009). Compounds were identified by mass spectral and retention time index matching to the reference collection of the Golm metabolome database [http://gmd .mpimpgolm.mpg.de/; (Kopka et al., 2005; Schauer et al., 2005; Hummel et al., 2010)]. Guidelines for manually supervised metabolite identification were based on the presence of at least three specific mass fragments per compound and a retention index deviation < 1.0% (Strehmel et al., 2008). All mass features of an experiment were normalized by sample volume, OD750nm, internal standard and maximum scaled. For quantification purposes, all mass features were evaluated for best specific, selective and quantitative representation of observed analytes. Laboratory and reagent contaminations were evaluated by non-sample control experiments. Metabolite levels were routinely assessed by relative changes.

PGPase activity in Synechocystis Synechocystis cultures were concentrated and washed in 5 mM Hepes pH 7.5. Pellets were resuspended in PGPase assay buffer (5 mM MgCl2, 20 mM MES-KOH pH 6.3) and sonicated, centrifuged at 20 000 g, 4°C for 30 min. Total proteins levels were quantified by the Bradford method. Crude protein extracts (20 μg) were used for PGPase activity assay essentially as described in the study of Suzuki (1995) for Synechocystis and in the study of Schäferjohann and colleagues (1993) for the expressed CbbZ in E. coli, using the Ames reagent (Ames, 1966) for the quantification of the phosphatase depended release of inorganic phosphate (Pi).

RNA extraction Cells were harvested from 20 ml cultures by centrifugation at 4°C and treated with Tri-Reagent (Molecular Research Center, USA) according to the provided protocol. The RNA pellets were dried at 55°C, dissolved in 20–40 μl DNase/ RNase/Protease free water (Sigma) and then stored at –80°C. DNase (Turbo DNA-free, Ambion, Austin, TX, USA) reaction was carried out on the RNA samples according to the manufacturer’s instructions.

cDNA synthesis and real-time PCR (qPCR) For qPCR analysis, cDNA syntheses were carried out using the ImProm-II Reverse Transcription System (Promega, Madison, WI, USA) according to manufacturer’s instructions. Five microlitres of diluted cDNA, corresponding to 10 ng total RNA, was used for qPCR carried out on a Rotor-Gene 6000 Thermal Cycler (Corbett Research, Australia). Amplifications were performed in a total volume of 15 μl using the KAPA SYBR® FAST qPCR Kit (Kapa Biosystems) according to the manufacturer’s instructions. Forty-five cycles of amplification were performed under the following conditions: DNA polymerase activation at 95°C for 3 min, denaturation at 95°C for 3 s, annealing at 52–60°C (according to the primers used) for 30 s, product extension at 72°C for 1 s and signal acquiring at

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology

Cyanobacterial acclimation to changing CO2 level

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Table 2. Primers used for qPCR. Primer

Forward (5′ to 3′)

Reverse (5′ to 3′)

rnpB sbtA ndhF3 cmpA slr0458 sll1349 slr0586 slr1762

GAGTTGCGGATTCCTGTCAC TGGTGCTCTGTATCCCTTTATG AAACCTTTGTGGACGGAGTG CGAGCGGATTGGGTAGATAA CGCTCCCTACGATTACCAAA CTGCTCAACCAAAAGGGCTA CGGATAGTGTGTGGATGGTG GGAGTGTGGAACGGTTCATT

TGCACCCTTACCCTTTTCAG TGCTGAACGATTCCTCAATACT AAAGGCGCCAATTAAAGTCA CCGGACACAATCTGTACAAC GATGATCCACTGGCAAATCC CCCGATTGATAATGCGGT ACCCCATGGTGTTGGTAAAA CAATCATCAAGGTGTGCTC

72°C. The reactions were carried out in triplicates. The primers used in this study are shown in Table 2.

CO2 depended O2 evolution The rates of CO2-dependent O2 evolution as a function of Ci concentration were determined using a Clark type O2 electrode (PS2108, Passport dissolved O2 sensor Roseville, CA, USA), essentially as described in the study of Kaplan and colleagues (1988). The cells were harvested by centrifugation and resuspended in a CO2-free medium containing 10 mM NaCl and 20 mM Hepes-NaOH, pH 7.8 and then placed in the O2 electrode chamber at 30°C. The cells were exposed to 450 μmol photons m−2 s−1 and allowed to utilize the Ci in their medium until they reached the CO2 compensation point. 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.

Acknowledgements We thank the German Research Foundation (Deutsche Forschungsgemeinschaft) (Project FE 218/14-2, HA2002/ 12), The Israel Science Foundation and the Israeli Ministry of Science and Technology for funding this research.

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Supporting information Additional Supporting Information may be found in the online version of this article at the publisher’s web-site: Fig. S1. Phylogenetic tree of the hypothetical PGPases from Synechocystis PCC 6803 compared with known PGPases or hypothetical PGPases or members of the HAD-superfamily from different organisms was produced using MEGA software (http://www.megasoftware.net/). The PGPases from Synechocystis and Ralstonia eutropha (CbbZ) are highlighted. Fig. S2. Fold change in transcript abundance of putative PGPases encoding genes as affected by time under LC. Quantitative PCR of slr0458 (A), sll1349 (B), slr0586 (C) and slr1762 (D) was analysed 1.5 h, 6 h and 24 h after transferring the cells from HC to LC. Expression levels were calculated relative to the reference gene rnpB, and to the level of the transcript under HC conditions. Mean values represent at least three biologically independent repeats. Fig. S3. Glycogen content in the WT, OE and DM. Cells from HC were transferred to LC for 3 h, then harvested and glycogen content measured. Mean values represent three independent experiments. Glycogen levels in Synechocystis were carried out as described in the study of Singh and Sherman (2005) using the anthrone reagent and a glucose concentration standard curve. Appendix S1. Transcript abundance of other putative PGPase encoding genes.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology