### Amino Acid Analysis by Gas Chromatography Mass Spectrometry (GC-MS)

Cells were cultivated in modified BG-11 medium for 1 day (starting from OD730 = 0.2), and equal amounts of cells (50 mL cell culture with OD730 = 1.0) were harvested by rapid filtration. Nitrogen-starved cells were similarly collected by filtration after 4 h of cultivation in BG-110 medium. Amino acids were quantified by GC-MS as previously described (Osanai et al., 2014). All the results are listed in Table S2.

### RESULTS

### Alteration of Oxygen Evolution and Consumption by Hik8 Overexpression

To study the role of Hik8 in photosynthetic electron transport, respiratory and photosynthetic activities were measured. Previous studies showed that Hik8 regulates the expression of genes related to sugar catabolism and the nitrogen-induced sigma factor sigE (Singh and Sherman, 2005; Osanai et al., 2015), and sugar catabolism in Synechocystis 6803 is particularly altered by nitrogen status (Osanai et al., 2006). Thus, we chose both nitrogen-replete and nitrogen-depleted experimental conditions. Respiratory activities of GT and HOX80 under nitrogen-replete conditions were 18.6 and 12.5µmol O2 mg chla −1 h −1 , respectively, and thus, the respiratory activity of HOX80 was two-thirds of that in the GT strain (**Figure 1A**). The respiratory activity in GT increased by 1.3 times after 1 day of nitrogen depletion, whereas that in HOX80 increased by 2.1 times after 1 day of nitrogen depletion (**Figures 1A,B**). Photosynthetic activity was almost the same between GT and HOX80 under nitrogen-replete and nitrogen-depleted conditions for 1 day (**Figure 1B**). After 3 days of nitrogen depletion, respiratory, and photosynthetic activities in HOX80 were slightly higher and lower, respectively, than those in GT (**Figures 1A,B**). Changes in the color of the cultures during nitrogen starvation were similar between GT and HOX80. To confirm this, the absorption spectra were measured using an end-on type spectrophotometer. The transient increase and gradual decrease after prolonged nitrogen starvation in OD623 (which are the peaks representing phycobilisomes) were similar between GT and HOX80 (**Figure 1C**).

Chlorophyll fluorescence was subsequently measured to determine the values of the photosynthetic parameters. The values of Fv/Fm (the maximal photochemical efficiency of PSII), Fv ′ /Fm′ (the photochemical efficiency of open PSII centers), qP (photochemical quenching), and 8II (the effective quantum yield of electron transport through PSII) were decreased by hik8 overexpression under nitrogen-replete conditions (**Figure 2**). The values of Fv ′ /Fm′ and 8II in HOX80 were also lower than in GT under nitrogen-depleted conditions for 1 day (**Figure 2**).

### Alteration of the Expression of Genes Related to Electron Transport

Subsequently, the transcript levels of the genes related to electron transport were measured. The transcript levels of 15 out of 22 PSII genes increased as a result of hik8 overexpression under nitrogen-replete conditions; this increase was statistically significant (**Figures 3**, **4**). The transcript levels of genes encoding the reactive center of PSII (psbAII, psbB, psbC, psbD, and psbD2) increased more than 1.6 times by hik8 overexpression under nitrogen-replete conditions (**Figure 3**). After nitrogen depletion for 1 day, the expression of all the genes except psbAII and psbD2 was repressed in both GT and HOX80, and their levels were similar between the two strains (**Figures 3**, **4**).

The transcript levels of 7 out of 12 PSI genes increased as a result of hik8 overexpression under nitrogen-replete conditions (**Figure 5**). The expression of all PSI genes was down-regulated during nitrogen starvation, and their transcript levels were similar between GT and HOX80 (**Figure 5**).

The transcript analysis of five genes encoding the terminal cytochrome c oxidase showed that the transcript levels of ctaCI, ctaDII, and ctaEII increased by hik8 overexpression under nitrogen-replete conditions, whereas the levels ofctaEI decreased (**Figure 6**). The expression of five cytochrome c oxidase genes was induced by nitrogen depletion, and the levels were similar between GT and HOX80 (**Figure 6**).

nitrogen-depleted conditions. Data are the mean ± SD from seven independent experiments. Differences between GT and hik8-overexpressing cells were analyzed with Student's t-test. Asterisks denote statistical significance at \*P < 0.05. (C) The levels of OD623, representing absorption by phycobilisomes during nitrogen starvation. Data are the mean ± SD from eight independent experiments.

### Increase in Amino Acid Levels after Nitrogen Depletion was Abolished by Hik8 Overexpression

Phycobilisome degradation during nitrogen-starved conditions is thought to provide amino acids as nitrogen sources (Richaud et al., 2001). The 18 amino acids, ornithine, and glutathione were quantified under nitrogen-replete and nitrogen-depleted conditions (**Figure 7**). The increases in valine, leucine, isoleucine, threonine, serine, phenylalanine, glutamine, and tyrosine by nitrogen depletion were abolished by hik8 overexpression (**Figure 7**).

### DISCUSSION

In this study, we examined the involvement of a histidine kinase Hik8 in photosynthetic electron transport, and our genetic analysis revealed that hik8 overexpression modified the expression of PSII and PSI genes (**Figures 3**–**5**). Our analysis demonstrated that photosynthetic electron transport is regulated by a circadian-related protein in this cyanobacterium. Photosynthetic activity was partially decreased under nitrogendepletion by hik8 overexpression (**Figure 1B**), which is empirically consistent with a decrease in 8II and Fv ′ /Fm′

in HOX80 (**Figure 2**). Previous study has showed that overexpression of sigE accelerates sugar catabolism and decreases the values of 8II and Fv ′ /Fm′ (Osanai et al., 2013) and hik8 overexpression activates sugar catabolism (Osanai et al., 2015). Thus, the degree of sugar catabolism and the values of 8II and Fv ′ /Fm′ may be negatively correlated in this cyanobacterium. The value of Fv/Fm also decreased by hik8 overexpression (**Figure 2**), indicating HOX80 strain contains aberrant PSII complexes. The expression of PSII and PSI genes was upregulated as a consequence of hik8 overexpression (**Figures 3**–**5**), and thus, the proper amount of photosynthetic transcripts and/or proteins was important for the activity of oxygen evolution. RpaA is a probable cognate response regulator of Hik8 and inactivation of rpaA resulted in a decrease in the monomeric PSI and D1 protein levels (Majeed et al., 2012). Although the direct involvement of Hik8 in photosynthetic electron transport is unclear, RNA-seq analysis has demonstrated that RpaA does not bind with the promoters of photosynthetic genes (Markson et al., 2013), nevertheless RpaA is important in adaptation to changes in light conditions (Iijima et al., 2015b). We here genetically showed that Hik8 manipulates the expression of genes related to PSI, PSII, and cytochrome c oxidases.

Several groups have shown a transient increase in amino acid levels after nitrogen depletion in Synechocystis 6803

(Hauf et al., 2013; Kiyota et al., 2014; Osanai et al., 2014). Amino acids containing additional nitrogen molecules (glutamine, glutamate, aspartate, and asparagine) decreased after 4 h of nitrogen depletion, whereas other amino acids increased (Osanai et al., 2014). Kiyota et al. (2014) divided amino acids into two groups: NblA-dependent and NblAindependent amino acids. NblA is a protein essential for the degradation of phycobilisome in cyanobacteria (Collier and Grossman, 1994). The NblA-dependent amino acids are glutamine, glutamate, glutathione, glycine, isoleucine, leucine, methionine, phenylalanine, proline, serine, threonine, tyrosine, and valine, and the NblA-independent amino acids are alanine, asparagine, lysine, and tryptophan (Kiyota et al., 2014). All eight amino acids whose induction was modestly decreased during nitrogen starvation by hik8 overexpression (valine, leucine, isoleucine, threonine, serine, phenylalanine, glutamine, and tyrosine) are included in the NblA-dependent group (**Figure 7**). Since the phycobilisome degradation was similar between GT and HOX80 (**Figure 1C**), the reason for the difference of the amino acid levels between GT and HOX80 after nitrogen depletion was unclear. The interaction of photosynthetic electron transport and amino acid metabolism offers intriguing insights into the mechanisms of cell maintenance in cyanobacteria, and we suggest, from this study, that a circadian-related protein is important for this integrity. In this study, we found the metabolite levels and photosynthetic electron transport concomitantly altered in HOX80.

The regulation of photosynthetic genes by a histidine kinase named CSK has been demonstrated in Arabidopsis thaliana (Puthiyaveetil et al., 2010). CSK regulates the activity of bacteriatype RNA polymerase through control of the phosphorylation of sigma factor Sig1 (Puthiyaveetil et al., 2010). Synechocystis 6803 possesses multiple sigma factors, SigA–SigI (Osanai et al., 2008), and light-induced psbAII/AIII gene expression is reduced by sigD knockout (Imamura et al., 2003, 2004). Microarray experiments also indicate the involvement of SigD in the expression of photosynthetic genes (Summerfield and Sherman, 2007). Photosynthetic oxygen evolution is not affected by single or double knockout of group-2 sigma factors, but the double knockout of sigB/sigD leads to sensitivity to photoinhibition because of abolished up-regulation of psbA expression (Pollari et al., 2008, 2009, 2011). We previously showed that Hik8 positively regulates sigma factor SigE, and genetic modification of sigE alters photosynthetic electron transport (Osanai et al., 2013, 2015). In this way, regulation of photosynthesis by combinations of histidine kinases and sigma factors is conserved in both

FIGURE 5 | Transcript levels of 12 genes encoding Photosystem I proteins in GT and hik8-overexpressing (HOX80) cells. Data are the mean ± SD from independent experiments (n = 3∼4). The levels were calibrated relative to the value obtained in the GT strain under nitrogen-replete conditions, which was set at 100%. Differences between GT and hik8-overexpressing cells were analyzed with Student's t-test. Asterisks denote statistical significance at \*P < 0.05.

prokaryotic and eukaryotic photosynthetic organisms. Detailed analysis of the mechanisms controlling photosynthetic electron transport through transcriptional cascades is important, and it may also lead to an understanding of the regulatory mechanisms of primary metabolism in cyanobacteria.

### ACKNOWLEDGMENTS

This work was supported by the Ministry of Education, Culture, Sports, Science, and Technology, Japan, through a grant to TO from CREST from the Japan Science and Technology Agency and

### REFERENCES

ALCA from the Japan Science and Technology Agency (project name "Production of cyanobacterial succinate by the genetic engineering of transcriptional regulators and circadian clocks"). All authors contributed to the work and approved the submission of the manuscript.

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2015.01150

alters amino acid composition and glycogen metabolism. Front. Microbiol. 6:326. doi: 10.3389/fmicb.2015.00326

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Kuwahara, Arisaka, Takeya, Iijima, Hirai and Osanai. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Genetic manipulation of a metabolic enzyme and a transcriptional regulator increasing succinate excretion from unicellular cyanobacterium

#### Edited by:

*Weiwen Zhang, Tianjin University, China*

#### Reviewed by:

*Jiangxin Wang, Shenzhen University, China Min Chen, University of Sydney, Australia*

#### \*Correspondence:

*Takashi Osanai, Department of Agricultural Chemistry, School of Agriculture, Meiji University, 1-1-1 Higashimita, Tama-ku, Kawasaki, Kanagawa 214-8571, Japan<EMAIL_ADDRESS>Masami Y. Hirai, RIKEN Center for Sustainable Resource Science, 1-7-22 Suehiro-cho, Tsurumi-ku, Yokohama, Kanagawa 230-0045, Japan<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Microbial Physiology and Metabolism, a section of the journal Frontiers in Microbiology*

> Received: *21 August 2015* Accepted: *15 September 2015* Published: *06 October 2015*

#### Citation:

*Osanai T, Shirai T, Iijima H, Nakaya Y, Okamoto M, Kondo A and Hirai MY (2015) Genetic manipulation of a metabolic enzyme and a transcriptional regulator increasing succinate excretion from unicellular cyanobacterium. Front. Microbiol. 6:1064. doi: 10.3389/fmicb.2015.01064*

Takashi Osanai 1, 2 \*, Tomokazu Shirai 3 , Hiroko Iijima1, 2, Yuka Nakaya1, 3 , Mami Okamoto3 , Akihiko Kondo3, 4 and Masami Y. Hirai 1 \*

*1 RIKEN Center for Sustainable Resource Science, Yokohama, Japan, 2 Department of Agricultural Chemistry, School of Agriculture, Meiji University, Kawasaki, Japan, 3 Biomass Engineering Program, RIKEN, Yokohama, Japan, 4 Department of Chemical Science and Engineering, Graduate School of Engineering, Kobe University, Kobe, Japan*

Succinate is a building block compound that the U.S. Department of Energy (DOE) has declared as important in biorefineries, and it is widely used as a commodity chemical. Here, we identified the two genes increasing succinate production of the unicellular cyanobacterium *Synechocystis* sp. PCC 6803. Succinate was excreted under dark, anaerobic conditions, and its production level increased by knocking out *ackA*, which encodes an acetate kinase, and by overexpressing *sigE*, which encodes an RNA polymerase sigma factor. Glycogen catabolism and organic acid biosynthesis were enhanced in the mutant lacking *ackA* and overexpressing *sigE*, leading to an increase in succinate production reaching five times of the wild-type levels. Our genetic and metabolomic analyses thus demonstrated the effect of genetic manipulation of a metabolic enzyme and a transcriptional regulator on succinate excretion from this cyanobacterium with the data based on metabolomic technique.

#### Keywords: cyanobacteria, metabolism, metabolomics, sigma factor, succinate

## Introduction

In 2004, the U.S. Department of Energy (DOE) selected the top 12 building block chemicals from a list of more than 300 candidates that were produced from biomass (Werpy and Petersen, 2004). Among these, four-carbon dicarboxylic acids, including succinate, were included. Succinate can be used as a precursor to numerous chemicals such as a biodegradable plastic like polybutylene succinate, fibers, and pigments (Zeikus et al., 1999; Hong and Lee, 2002; Werpy and Petersen, 2004). Succinate is currently derived from petroleum, but it could also be produced using bacteria (McKinlay et al., 2007).

Production of succinate by recombinant heterotrophic bacteria such as Escherichia coli, Corynebacterium glutamicum, Anaerobiospirillum succiniciproducens, Actinobacillus succinogenes, and Mannheimia succiniciproducens has been intensively studied (Samuelov et al., 1991; Guettler et al., 1999; Chatterjee et al., 2001; Hong and Lee, 2001; Hong et al., 2004; Lee et al., 2006). Succinate is an intermediate in the tricarboxylic acid (TCA) cycle and is excreted by succinate-producing cells during anaerobic fermentation (McKinlay et al., 2007). Succinate is produced from phosphoenolpyruvate via the reductive branch of the TCA cycle, in which phosphoenolpyruvate is converted to oxaloacetate by phosphoenolpyruvate carboxylase (PEPC) or phosphoenolpyruvate carboxykinase (PEPCK) under anaerobic conditions (McKinlay et al., 2007). Malate is produced from oxaloacetate when catalyzed by malate dehydrogenase, and fumarate is produced from malate when catalyzed by fumarase. This is followed by the production of succinate when catalyzed by succinate dehydrogenase (SDH; McKinlay et al., 2007). The overexpression of a gene encoding PEPC in E. coli increases succinate production 3.8-fold (Millard et al., 1996). The introduction of PEPC, PEPCK, or malic enzyme (catalyzing a reaction from pyruvate to malate), also enhances succinate production in E. coli (Hong and Lee, 2001; Kim et al., 2004; Lin et al., 2005; Zhang et al., 2009). The deletion of ldhA (encoding Llactate dehydrogenase), adhE (encoding alcohol dehydrogenase), and ack-pta (encoding acetate kinase and phosphotransacetylase, respectively) prevented the production of L-lactate, ethanol, and acetate, that are by-products during anaerobic fermentation, also increases the production of succinate in E. coli (Sánchez et al., 2005a,b; Jantama et al., 2007). In addition, the activation of the glyoxylate pathway by the deletion of iclR, which encodes the transcriptional repressor of the genes related to glyoxylate pathway, increased the succinate productivity in E. coli (Sánchez et al., 2005a,b). Thus, inhibition of by-product formation

combined with additional genetic engineering can up-regulate succinate productivity.

Cyanobacteria are a group of bacteria that fix carbon dioxide via oxygenic photosynthesis. The potential applications of cyanobacteria in providing renewable energy and resources may reduce the environmental burden. Genome information for cyanobacteria is available (Kanesaki et al., 2012), and genetic engineering is easily performed by homologous recombination in several cyanobacterial strains, including the non-nitrogen fixing cyanobacterium Synechocystis sp. PCC 6803 (hereafter Synechocystis 6803; Ikeuchi and Tabata, 2001). The genome of Synechocystis 6803 was the first sequenced among the cyanobacteria (Kaneko et al., 1996), and it has been used extensively in basic and applied sciences.

There are few reports of succinate production using cyanobacteria. McNeely et al. revealed that five fermentation products, lactate, acetate, succinate, alanine, and hydrogen, were produced under dark, anaerobic conditions by the marine cyanobacterium Synechococcus sp. PCC 7002 (hereafter Synechococcus 7002; McNeely et al., 2010). A knockout of ldhA increased acetate and hydrogen levels, and diminished lactate production (McNeely et al., 2010). Succinate was excreted from the ldhA knockout cells, but it was not detected from the wild-type cells of Synechococcus 7002 (McNeely et al., 2010). The filamentous, non-diazotrophic cyanobacteria Arthrospira

phosphoenolpyruvate carboxylase; PPS, phosphoenolpyruvate synthase; PK, pyruvate kinase; MDH, malate dehydrogenase; ME, malic enzyme; FH, fumarate hydratase; SDH, succinate dehydrogenase; PDH, pyruvate dehydrogenase; DDH, D-lactate dehydrogenase, CS, citrate synthase; PTA, phosphotransacetylase; ACK, acetate kinase; and ACS, acetyl-CoA synthetase.

maxima CS-328 cells produced lactate, acetate, ethanol, formate, and hydrogen under dark, anaerobic conditions, but succinate excretion was not detected (Carrieri et al., 2010, 2011). For Synechocystis 6803, hydrogen is generated under both light and dark, anaerobic conditions (Osanai et al., 2013). Organic acids, including D-lactates, were highly produced by genetically engineered Synechocystis 6803 cells, and succinate was also generated but its levels were only 3% of total carbon excreted, suggesting that genetic and metabolic engineering are necessary to increase succinate production (Angermayr et al., 2014; Hollinshead et al., 2014; McNeely et al., 2014). The genes encoding enzymes for organic acid production exist in the Synechocystis 6803 genome (**Figure 1**).

We reveal here that succinate was excreted from Synechocystis 6803 cells under dark, anaerobic conditions and the succinate levels were enhanced by reducing acetate biosynthesis and overexpressing sigE encoding a sigma factor. These results demonstrated the genetic manipulation of two types of genes increasing the succinate excretion from this cyanobacterium.

### Materials and Methods

#### Bacterial Strains and Culture Conditions

The glucose-tolerant strain of Synechocystis sp. PCC 6803, isolated by Williams (Williams, 1988), was grown in modified BG-11 medium, consisting of BG-110 liquid medium (Rippka, 1988) supplemented with 5 mM NH4Cl (buffered with 20 mM HEPES–KOH, pH 7.8). The GT-I strain, among GT substrains, was used in the current study (Kanesaki et al., 2012). Liquid cultures were bubbled with 1% (v/v) CO2 in air and incubated at 30◦C under continuous white light (∼50–70µmol photons m−2 s −1 ). For the mutant strains, 10, 0.3, and 10µg/mL of kanamycin, gentamycin and chloramphenicol, respectively, were added for preculturing. Modified BG-11 medium (containing 10 mM NH4Cl in liquid medium) was solidified with agar (1.5% w/v) for plate cultures, and similarly incubated in air at 30◦C under continuous white light (∼50–70µmol photons m−2 s −1 ). Cell densities were measured at A730 using a Hitachi U-3310 spectrophotometer (Hitachi High-Tech., Tokyo, Japan).

For succinate production, cells grown in 70 mL modified BG-11 medium (started from A730 = 0.4) for 3 days were concentrated into 10 mL HEPES buffer (20 mM HEPES–KOH, pH 7.8) or modified BG-11 medium to A730 = 20 in a GC vial. The vial was sealed using butyl rubber, and N2 gas was introduced using syringes for 1 h to produce anaerobic conditions. After removing the syringes, the vial was wrapped with aluminum foil and shaken at 30◦C. Cell cultures were then centrifuged at 5800 × g for 2 min, the supernatant was filtrated, and 1 mL supernatant was freeze-dried for 1 day. The dried sample was used for high-performance liquid chromatography analysis.

### Plasmid Construction of Knock-in Vectors pTCP1556, pTCP0542, and pTCP1299

The kanamycin resistance cassette of the pTKP2031V vector (Osanai et al., 2011) was removed by digestion with XhoI and AatII (Takara Bio, Shiga, Japan). The chloramphenicol resistance cassette from pKRP10 (Reece and Phillips, 1995) was amplified by PCR with KOD polymerase (Toyobo, Osaka, Japan) and the specific primers in Table S2, digested with XhoI and AatII, and inserted into the XhoI-AatII sites of pTKP2031V. The resultant plasmid was named pTCP2031. Regions of ddh (slr1556), from −297 to +800 bp, acs (sll0542), from +921

to +1962 bp, and ackA (sll1299), from +270 to +1238 bp, based on the translation initiation codons, were amplified by PCR with KOD plus neo polymerase (Toyobo) and the specific primers in Table S2. The fragments amplified by PCR were digested with SphI and EcoRV (Takara Bio) and inserted into the SphI-SmaI sites of the pUC119 vector (Takara Bio). The resultant plasmid was digested with HincII (for ddh and ackA) or ApaI (for acs), and the region including the chloramphenicol resistance cassette, psbAII promoter, and NdeI-HpaI cloning sites of pTCP2031 was amplified with KOD plus neo polymerase and the specific primers 5′ -TTTGCTTCATCGCTCGAG-3′ and 5′ - ATCCAATGTGAGGTTAAC-3′ , and integrated into the HincII or ApaI site of the plasmid. The resultant plasmids were named pTCP1556, pTCP0542, and pTCP1299 for knockouts of ddh, acs, and ackA, respectively. The sigE ORF was obtained by digestion with NdeI and HpaI from pTGP0945-sigE plasmid (Osanai et al., 2014a) and cloned into the NdeI-HpaI sites of pTCP1556, pTCP0542, and pTCP1299. The plasmids were integrated into the GT-I strain by natural transformation as described previously (Osanai et al., 2011). Knockouts and the insertion of the sigE ORF were confirmed by PCR using GoTaq (Promega, Fitchburg, WI, USA) with the primers in Table S2.

### Immunoblotting

Cells were collected by centrifugation (5800 × g for 2 min), and the supernatant was removed and cells were frozen by liquid nitrogen. Then, cells were dissolved in PBS-T and disrupted by sonication as described previously (Osanai et al., 2014a). Immunoblotting was performed as described previously (Osanai et al., 2014a). Antisera against SigE were generated previously (Osanai et al., 2009).

### Glycogen Measurement

Glycogen levels were measured at the Biotechnology Center of Akita Prefectural University (Akita, Japan), as described in Osanai et al. (2014a).

### LC-MS/MS Analysis

Equal amounts of cells (10 mL cell culture with A730 = 1.0) were harvested by rapid filtration, and metabolites were extracted using a previously described method (Osanai et al., 2014b). Briefly, the cells were filtrated, and then the intermediate metabolites were quenched and extracted in 1.2 mL of solvent mixture (CHCl3:CH3OH:H2O, 2.5:2.5:1, v/v/v) containing 10µg/L D-(+)-camphor-10-sulfonic acid as an

internal standard. After centrifugation at 15,000 × g at 4◦C for 5 min, 400µL of the upper phase was transferred to a new tube and vacuum-dried.

### GC-MS Analysis

Equal amounts of cells (10 mL cell culture with A730 = 1.0) were harvested by rapid filtration as mentioned above. GC-MS was carried out using a GCMS-QP2010 Ultra, and the detailed protocol is described in Osanai et al. (2015).

### Measurement of Organic Acids by High-Performance Liquid Chromatography (HPLC)

Freeze-dried supernatants were resolved in 100µL of filtered 3 mM perchloric acid. The resolved samples were analyzed by HPLC using a LC-2000Plus Systems (JASCO, Tokyo, Japan) with a photodiode array detector and two RSpak KC-811 columns (Showa Denko, Tokyo, Japan). Organic acids were quantified with 0.2 mM bromothymol blue in 15 mM sodium phosphate buffer; peaks were detected at 445 nm. The column temperature was 60◦C, and the flow rates of 3 mM perchloric acid and 0.2 mM bromothymol blue solutions were 1.0 and 1.5 mL/min, respectively.

### Results

### ackA Knockout and sigE Overexpression Enhanced Succinate Production

The identities of the excreted organic acids from the wildtype Synechocystis 6803 (GT) during anaerobic conditions were determined first. After cultivation for 3 days under light, aerobic conditions (1% CO2 in the air), cells were concentrated into 10 mL BG-110 medium or HEPES buffer with or without nitrogen sources (5 mM NH4Cl) in a GC-vial, subjected to anaerobic conditions by introducing N2 gas, and incubated for 3 days under dark conditions with shaking at 30◦C. Organic acids excreted into the medium or buffer were analyzed by HPLC. Succinate, lactate and acetate were detected, and the succinate levels were highest in HEPES buffer with nitrogen

source among the four conditions tested (**Figure 2**). Lactates were not detected in HEPES buffer (**Figure 2**). Acetate levels were higher in BG-11 or HEPES buffer without nitrogen sources than those in BG-11 or HEPES buffer with nitrogen sources (**Figure 2**). To reduce the cost of succinate production, subsequent experiments were performed using HEPES buffer without nitrogen sources.

To increase succinate production, we applied two strategies, decreasing lactate and acetate by knocking out each of the three genes (ddh, acs, or ackA; **Figure 1**) and promoting the sugar catabolic pathway by overexpressing sigE, encoding an RNA polymerase sigma factor, which activates the expression of sugar catabolic enzymes (Osanai et al., 2011). Knock-in vectors, which integrate the region containing the chloramphenicol resistance cassette, the psbAII promoter from the D1 protein of Photosystem II, and NdeI-HpaI cloning sites, were constructed to generate the knockout mutants of ddh, acs, and ackA (**Figure 3A**). The sigE open reading frame (ORF) was cloned into the NdeI-HpaI sites to generate the sigE overexpression strain combined with the ddh, acs, or ackA knockout, and the resultant strains were designated as 1556E, 0542E, and 1299E, respectively (**Figure 3A**). The insertion of these DNA fragments was confirmed by PCR (**Figure 3B**). Immunoblotting confirmed that SigE proteins in the three sigE-overexpressing strains were higher than in GT after 3 days of cultivation under dark, anaerobic conditions (**Figure 3C**).

Although the knockouts of ddh and acs did not increase the succinate levels, the ackA knockout increased the succinate level to 34.8 mg/L compared with the 13.9 mg/L produced by the parental wild-type strain under the same conditions. sigE overexpression (GOX50) alone increased the succinate level to 20.3 mg/L, and an additional knockout of ddh, acs, or ackA enhanced the levels to ∼35.6, 29.4, or 71.5 mg/L, respectively (**Figure 4A**). The wild-type cells produced less than 10 mg/L lactate, while the lactate levels increased in the ackA knockout to 51.7 mg/L, and sigE overexpression with an acs or ackA knockout enhanced the levels to 30.0 or 93.5 mg/L, respectively (**Figure 4A**). Acetate levels were decreased to 62.0 mg/L by the ackA knockout, compared with 294.3 mg/L acetate produced by wild-type cells, and the ratio of succinate and lactate to acetate increased in the ackA knockout mutant (**Figures 4A,B**). The strain lacking ackA and overexpressing sigE (1299E) had the highest succinate levels and ratios among the eight strains (**Figures 4A,B**). A time-course experiment analyzed different lengths of dark, anaerobic incubations and showed that a 3- or 4-day incubation period was long enough to produce sufficient quantities and ratios of succinate in 1299E (**Figure 5**). The succinate production rates from 1299E were 1.38 and 1.18 mg/L/h for 3- and 4-day incubation, respectively (**Figure 5**). Therefore, subsequent experiments were performed using a 3-day incubation period under dark, anaerobic conditions.

### A Metabolome Analysis

A metabolome analysis using the GT and 1299E strains grown under aerobic and anaerobic conditions was then performed to clarify the metabolic profiles. After dark, aerobic cultivation, ADP-glucoses disappeared and the levels of several sugar

*Data represent means* ± *SD results from four independent experiments. Glycogen levels were calibrated relative to that in GT under light conditions (set at 100%). ND, glycogen under detectable levels.*

phosphates increased (**Figure 6** and Table S1). The fructose-1, 6-bisphoshate and dihydroxyacetone phosphate levels increased more than 10 times under anaerobic conditions in the wildtype strain (**Figure 6**). Phosphoenolpyruvate, pyruvate and acetyl-CoA decreased greatly under anaerobic conditions (**Figure 6**). The levels of sugar phosphates, such as glucose-1-phosphate, glucose-6-phosphate, ribulose-5-phosphate, 6-phosphogluconate, fructose-6-phosphate, and fructose-1, 6-bisphosphate, in the 1299E strain under anaerobic conditions, were higher than those in the wild-type strain (**Figure 6**). Phosphoenolpyruvate and acetyl-CoA were lower in the 1299E strain than in the wild-type strain, and organic acids, such as succinate, lactate, malate, and fumarate, were higher in the 1299E strain than in the wild-type strain under anaerobic conditions (**Figure 6**).

The quantification of the glycogen levels before and after anaerobic cultivation for 3 days revealed that 33% of glycogen was consumed in the wild-type strain; however, 70% of glycogen were consumed in the 1299E strains (**Table 1**).

### Discussion

Fermentation is closely related to sugar metabolism. In Synechococcus 7002, the levels of excreted fermentation products were altered by the disruption of glgC, which encodes ADP-glucose pyrophosphorylase (Guerra et al., 2013). Lactate

production in the glgC knockout strain was approximately half of that in the wild-type, while acetate and alanine production were not significantly affected (Guerra et al., 2013). The reason for the decreased lactate excretion may be the slower catabolic rate of reduced sugars in the glgC knockout mutant (Guerra et al., 2013). The rate of sugar catabolism, not the amount of total reduced sugars, was important for the increased lactate production (Guerra et al., 2013). This finding was consistent with our results that sigE overexpression, which accelerated glycogen degradation and glucose catabolism (Osanai et al., 2011), increased succinate production in Synechocystis 6803 (**Figures 4A**, **7**). Glycogen degradation was enhanced by sigE overexpression during dark, anaerobic conditions (**Table 1**), possibly leading to increases in the biosynthesis of intracellular organic acids (**Figure 6** and Table S1), which in turn led to the production of extracellular succinate at a higher level (**Figures 4A**, **7**).

A metabolome analysis revealed that the levels of fructose-1,6-bisphosphate and dihydroxyacetone phosphate increased more than 10 times after dark, anaerobic cultivation in the wildtype strain (**Figure 6**). Glyceraldehyde-3-phosphate production also increased, but 3-phosphoglycerate/2-phosphoglycerate, phosphoenolpyruvate, pyruvate, and acetyl-CoA decreased after dark, anaerobic cultivation in the wild-type strain (**Figure 6**). These results suggest that important enzymatic reactions exist downstream of glyceraldehyde-3-phosphate under dark, anaerobic conditions. Glyceraldehyde-3-phosphate dehydrogenase, which produces glycerate-1,3-bisphosphate from glyceraldehyde-3-phosphate, is encoded by gap1 and gap2, and the reactions were uniquely catalyzed by the two enzymes in Synechocystis 6803. Gap1 catalyzes the catabolic reactions and Gap2 catalyzes the anabolic reactions (Koksharova et al., 1998). In addition, the gap1 transcript levels are regulated by at least two transcriptional regulators, SigE and a response regulator Rre37 (Osanai et al., 2005; Azuma et al., 2011), indicating the importance of gap1 in the sugar catabolism of Synechocystis 6803. The flux into the TCA cycle in Synechocystis is relatively low compared with other heterotrophic bacteria (You et al., 2015). Current results demonstrate that the flux into succinate can be up-regulated by our genetic modification under dark, anaerobic conditions.

A knockout of ackA reduced the acetate level but a knockout of acs did not affect the acetate level (**Figure 4A**), suggesting the major route of acetate biosynthesis under dark, anaerobic conditions is through an AckA-dependent pathway in this cyanobacterium (**Figure 1**). Prochlorococcus species lack ackA in their genomes (KEGG database URL: http://www.genome.jp/kegg-bin/show\_pathway?syn00620), and thus, acetate biosynthesis pathway may be diverse among cyanobacteria. The mutant lacking ddh showed diminished lactate production, but lactate was produced by sigE overexpression even in the ddh knockout (**Figure 4A**). These results indicate another pathway for lactate biosynthesis exists in this cyanobacterium. Lactate can be synthesized from lactoylglutathione, which is derived from dihydroxyacetone phosphate (KEGG database URL: http://www.genome.jp/kegg-bin/show\_pathway?syn00620), and the pathway may be activated by sigE overexpression. Our metabolome analysis revealed that NADPH disappeared, and that malate, fumarate, and succinate generally increased in the strains producing more succinate (**Figure 6** and Table S1). These

data suggest that succinate is produced through the reverse TCA cycle, as shown in **Figure 1**, as in other heterotrophic bacteria (Lee et al., 2006). Thus, the metabolic flux toward succinate production has been clarified in this cyanobacterium by the metabolome analysis. Pyruvate and phosphoenolpyruvate were severely reduced by light-to-dark transition (Iijima et al., 2015), and thus, the provision of these metabolites may be important to increase organic acid production under dark, anaerobic conditions. The current results demonstrated that a combination of the genetic manipulation of genes encoding a metabolic enzyme and a sigma factor succeeded in upregulating the succinate levels. Future study about the detailed metabolic regulation will contribute to further understanding of the mechanistic implication of succinate excretion from this cyanobacterium.

### References

### Acknowledgments

This work was supported by the Ministry of Education, Culture, Sports, Science, and Technology, Japan, by a grant to TO from ALCA (Project name "Production of cyanobacterial succinate by the genetic engineering of transcriptional regulators and circadian clocks") from the Japan Science and Technology Agency. All authors contributed to the work and approved the submission of the manuscript.

### Supplementary Material

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2015.01064

cyanobacterium Synechocystis sp. PCC 6803. Front. Microbiol. 6:888. doi: 10.3389/fmicb.2015.00888

activities of Anaerobiospirillum succiniciproducens. Appl. Environ. Microbiol. 57, 3013–3019.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Osanai, Shirai, Iijima, Nakaya, Okamoto, Kondo and Hirai. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Manipulation of oil synthesis in Nannochloropsis strain NIES-2145 with a phosphorus starvation–inducible promoter from Chlamydomonas reinhardtii

Masako Iwai 1, 2, Koichi Hori 1, 2, Yuko Sasaki-Sekimoto3 , Mie Shimojima1 and Hiroyuki Ohta1, 2, 3 \*

#### Edited by:

*Yuki Nakamura, Academia Sinica, Taiwan*

#### Reviewed by:

*Mee-Len Chye, The University of Hong Kong, Hong Kong Norihiro Sato, Tokyo University of Pharmacy and Life Sciences, Japan*

#### \*Correspondence:

*Hiroyuki Ohta, Graduate School of Bioscience and Biotechnology, Tokyo Institute of Technology, 4259-B-65 Nagatsuta-cho, Midori-ku, Yokohama, Kanagawa 226-8501, Japan<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Microbiotechnology, Ecotoxicology and Bioremediation, a section of the journal Frontiers in Microbiology*

Received: *30 June 2015* Accepted: *19 August 2015* Published: *07 September 2015*

#### Citation:

*Iwai M, Hori K, Sasaki-Sekimoto Y, Shimojima M and Ohta H (2015) Manipulation of oil synthesis in Nannochloropsis strain NIES-2145 with a phosphorus starvation–inducible promoter from Chlamydomonas reinhardtii. Front. Microbiol. 6:912. doi: 10.3389/fmicb.2015.00912* *1 Graduate School of Bioscience and Biotechnology, Tokyo Institute of Technology, Yokohama, Japan, 2 JST CREST, Tokyo, Japan, 3 Earth-Life Science Institute, Tokyo Institute of Technology, Tokyo, Japan*

Microalgae accumulate triacylglycerols (TAGs) under conditions of nutrient stress. Phosphorus (P) starvation induces the accumulation of TAGs, and the cells under P starvation maintain growth through photosynthesis. We recently reported that P starvation–dependent overexpression of type-2 diacylglycerol acyl-CoA acyltransferase (CrDGTT4) from *Chlamydomonas reinhardtii* using a sulfoquinovosyldiacylglycerol synthase 2 (SQD2) promoter, which has increased activity during P starvation, enhances TAG accumulation in *C. reinhardtii* cells. As a result, the content of C18:1 fatty acid, a preferred substrate of CrDGTT4, is increased in TAGs. Here we isolated genes encoding SQD2 from strain NIES-2145 of the eustigmatophyte *Nannochloropsis* and showed that their expression, like that in *C. reinhardtii*, was up-regulated during P starvation. To enhance oil accumulation under P starvation, we transformed pCrSQD2-CrDGTT4 into *Nannochloropsis* strain NIES-2145. The transformants had a fatty acid composition that was more similar to that of *C. reinhardtii*, which resulted in enhanced TAG accumulation and higher 18:1(9) content. The results indicated that the P starvation–inducible promoter of *C. reinhardtii* was able to drive expression of the *CrDGTT4* gene in *Nannochloropsis* strain NIES-2145 under P starvation. We conclude that the heterologous *CrSQD2* promoter is effective in manipulating TAG synthesis in *Nannochloropsis* during P starvation.

Keywords: algae, Nannochloropsis, phosphorus starvation, inducible promoter, triacylglycerol

## Introduction

Algal biofuel technology exploits algal photosynthesis and biosynthesis processes to produce oils using only sunlight, CO2, water and limited nutrients. Many microalgae accumulate triacylglycerols (TAGs) during nutrient stress (Giroud et al., 1988; Guschina and Harwood, 2006; Hu et al., 2008). It is estimated that the annual oil production from algae is in the range of 40,700–53,200 L ha−1 year−1 (Weyer et al., 2010). Therefore, the potential use of microalgae to provide biofuel feedstock is receiving significant attention.

A eukaryotic microalga, Chlamydomonas reinhardtii, is a model organism for studying algal biodiesel production because of its available whole-genome sequence and the ability to manipulate gene expression within this organism (Harris, 2009). Under stress conditions, such as nitrogen (N) starvation, Chlamydomonas nearly stops its growth and accumulates large amounts of TAGs (Grossman, 2000; Zhang et al., 2004). We reported that C. reinhardtii cells in logarithmic growth phase that were diluted into fresh medium showed substantial TAG accumulation under N or P deprivation (Iwai et al., 2014). P deprivation substantially induced the accumulation of oil droplets in the cytosol but allowed the maintenance of thylakoid membranes in C. reinhardtii (Iwai et al., 2014).

Nannochloropsis species are unicellular photosynthetic microalgae in the class Eustigmatophyceae. Previously known as "marine Chlorella," this class was identified on the basis of its ultrastructure and named Nannochloropsis by Maruyama et al. (1986). Its cells are spherical to slightly ovoid, are 2–4 µm in diameter and contain ovoid or cup-shaped chloroplasts (Maruyama et al., 1986). Nannochloropsis are of interest because of their rapid growth and their ability to produce large quantities of TAGs and be cultured on an industrial scale (Hodgson et al., 1991; Rodolfi et al., 2009; Huerlimann et al., 2010) In recent years, Nannochloropsis strains have been studied for their biomass production and their lipid composition and content under different growth conditions (Hu and Gao, 2006; Converti et al., 2009; Rodolfi et al., 2009; Simionato et al., 2011; Arudchelvam and Nirmalakhandan, 2012; Vieler et al., 2012b; Wang et al., 2014). In Nannochloropsis strains, N deprivation induces lipid accumulation and lipid droplet formation (Rodolfi et al., 2009; Vieler et al., 2012a,b; Martin et al., 2014), and their lipid content also increases with decreasing P concentrations (Hu and Gao, 2006; Rodolfi et al., 2009; Bondioli et al., 2012).

Diacylglycerol acyltransferase (DGAT) catalyzes the last step of TAG synthesis involving sn-1,2 diacylglycerol and acyl-CoA (Lung and Weselake, 2006; Rajakumari et al., 2008) and includes two major types, type 1 DGAT (DGAT1) and type 2 DGAT (DGAT2) (Cases et al., 1998; Lardizabal et al., 2001; Shockey et al., 2006). There are six DGAT genes in C. reinhardtii (encoding one DGAT1 and five DGAT2s) (Miller et al., 2010; Boyle et al., 2012). In contrast, 12 or 13 DGAT genes (encoding one or two DGAT1s and 11 DGAT2s) are present in Nannochloropsis strains (Radakovits et al., 2012; Vieler et al., 2012b; Wang et al., 2014). During N starvation in Nannochloropsis oceanica IMET1, seven DGAT transcripts are up-regulated and six other DGAT transcripts are down-regulated (Li et al., 2014).

Endogenous promoters are mainly used for the nuclear transformation of algae such as C. reinhardtii (Eichler-Stahlberg et al., 2009; Harris, 2009; Brueggeman et al., 2014), although stable nuclear transformation of algae is feasible with promoters from heterologous sources that are categorized in the same class (Hirata et al., 2011; Lerche and Hallmann, 2013, 2014). Transformation experiments using Nannochloropsis with endogenous promoters have recently been reported (Kilian et al., 2011; Vieler et al., 2012b). Additionally, a construct with the C. reinhardtii α-tubulin promoter was used for transforming Nannochloropsis (Vieler et al., 2012b). However, the stable nuclear transformation of Nannochloropsis using an inducible promoter from a heterologous source under specific conditions, in particular an oil-accumulating condition such as nutrient starvation, has not been reported.

In this study, we found that Nannochloropsis strain NIES-2145 has a homolog of the Arabidopsis SQD2 gene. Our results suggest that there is a common expression control system in a wide range of algal species, including primary and secondary microalgae, for adaptation to low P. We used the promoter of sulfoquinovosyldiacylglycerol (SQDG) synthase 2 (SQD2), which encodes the sulfoquinovosyl transferase that catalyzes the second step of sulfolipid biosynthesis (Yu et al., 2002). The transcript levels of the SQD2 gene homologs are increased in C. reinhardtii and Arabidopsis by P starvation concomitant with increases in the SQDG content (Yu et al., 2002; Chang et al., 2005; Okazaki et al., 2009; Iwai et al., 2014). The promoter of SQD2 induced expression of the down stream gene during P starvation, to successfully overexpress C. reinhardtii type-2 diacylglycerol acyl-CoA acyltransferase (CrDGTT4) in Nannochloropsis strain NIES-2145 under P starvation. The total lipid content, neutral lipid content and fatty acid profiles were determined. CrDGTT4 enhanced TAG accumulation under P starvation by changing the fatty acid composition to be more similar to that of C. reinhardtii. Therefore, the heterologous CrSQD2 promoter is effective in manipulating oil synthesis in Nannochloropsis during P starvation.

### Materials and Methods

### Materials and Culture Conditions

Nannochloropsis strain NIES-2145 was obtained from the Microbial Culture Collection of the National Institute for Environmental Studies, Japan. Nannochloropsis strain NIES-2145 was grown photoautotrophically in f/2 medium (Guillard and Ryther, 1962) or F2N50%SW medium (standard medium), which is F2N medium (Kilian et al., 2011) made with 50% artificial seawater (Wako Pure Chemical Industries, Ltd., Japan). Liquid cultures were grown in continuous white light (20– 40µmol photons m−2 s −1 ) at room temperature. Nutrient deficiency was induced by centrifuging the cells for 10 min at 2000 × g, washing them twice with the respective medium and subsequently resuspending them in the standard medium without NaNO3 and NH4Cl (–N) or phosphate (–P) solutions. Agar plates were prepared using 0.8% Bacto agar (Difco, USA) in standard medium. The cells were maintained on these plates at the same light intensity at room temperature.

### Analysis of Differential Gene Expression Levels

The differential expression levels of the CrDGAT gene and SQD2 genes in Nannochloropsis strain NIES-2145 cultured in standard, -P and -N media were determined using quantitative real-time PCR (qPCR). RNA was extracted from standard, -P and -N cultures using the phenol/chloroform method. Total RNA (500 ng) was used for the synthesis of cDNA with an oligo(dT)18 primer, random hexamers and Superscript II reverse transcriptase (Invitrogen, Carlsbad, CA). cDNA was amplified using SYBR Premix Ex Taq II (Takara, Japan). The Thermal Cycler Dice Real Time System and Multiplate RQ software (Takara) were used for the analysis. Expression levels of the CrDGAT gene and SQD2 genes were normalized to TUB mRNA expression. The CrDGTT4 expression levels in Nannochloropsis cultured in -P medium were compared with those of Nannochloropsis cultured in the standard medium (the values of which were set to 1). Primers are listed in **Supplemental Table 1**.

### Construction and Identification of CrDGTT4-overexpressing NIES-2145 Cell Lines

Transformation experiments were performed using a construct developed for the nuclear transformation of Nannochloropsis, pMD20 with a NT7 cassette (Kilian et al., 2011), except that a truncated bidirectional violaxanthin–chlorophyll a binding protein 2 promoter and the violaxanthin–chlorophyll a binding protein 1 3′ UTR from Nannochloropsis strain NIES-2145 were used and that the EcoRV restriction site was inserted between the ble gene and the violaxanthin–chlorophyll a binding protein 1 3′ UTR. To construct the expression plasmid carrying the CrDGTT4 (XM\_001693137.1) gene, pCrSQD2aDGTT4, which contains the C. reinhardtii SQD2 promoter region and the coding sequence of the DGTT4 gene (Iwai et al., 2014), was digested with restriction endonucleases EcoRV and SpeI and was treated using a DNA Blunting Kit (Takara). The resulting blunt-ended DNA was cloned into pMD-NT7, which was digested with restriction endonuclease EcoRV. Electroporation with a supplier was then used to transform the nuclear genome of Nannochloropsis strain NIES-2145 cells with 2–10µg vector construct (Vieler et al., 2012b). The selection of transformants was performed on standard medium supplemented with 2µg mL−1 Zeocin. Approximately 100 positive transformants were identified by growth on the selective medium, and overexpression levels in each resulting line were determined using qPCR.

### Extraction and Separation of Lipids

Nannochloropsis cells were harvested by centrifugation (10 min at 3000 × g), and total lipids were extracted as described (Bligh and Dyer, 1959). The lipids were dissolved in chloroform/methanol (2:1, v/v) and stored at −20◦C. Lipid classes were separated using two-dimensional thin-layer chromatography (TLC). The first dimension was developed using chloroform/methanol/7 N ammonia water (115:80:8, v/v/v). After the plates were dried for 1 h, the second dimension was developed using chloroform/methanol/acetic acid/water (170:25:15:3, v/v/v/v). TAGs were separated by TLC using the solvent system hexane/diethyl ether/acetic acid (160:40:4, v/v/v). Lipids were visualized under UV light after spraying the TLC plates with 0.001% (w/v) primuline in 80% (v/v) acetone.

### Preparation of Fatty Acid Methyl Esters (FAMEs)

Each lipid was scraped off of the TLC plates. FAMEs were obtained by incubating lipids for 1 h at 85◦C in the presence of 5% (v/v) hydrogen chloride–methanol solution (Wako Pure Chemical Industries) (Iwai et al., 2014). FAMEs were extracted using hexane and were determined by gas chromatography (GC). Fatty acid quantification was performed by comparing samples with the FAMEs derived from the internal standard, pentadecanoic acid (15:0).

### GC Analysis

FAMEs were analyzed using a GC-2014 (Shimadzu Corporation, Kyoto, Japan) with a HR-SS-10 capillary column (length, 25 m; internal diameter, 0.25 mm; Shinwa Chemical Industries, Ltd., Japan). The fatty acid profiles were determined by comparison with a standard reference mix composed of FAMEs (Supelco 18917-1AMP, 18913-1AMP, CRM47885; Sigma-Aldrich, GLC411, GLC462; Funakoshi, Japan).

### GC-mass Spectrometry (MS)

The qualitative composition of FAMEs was studied using a GS-MS (model GCMS-TQ8030; Shimadzu Corporation). Highgrade pure helium was used as the carrier gas. The ionization voltage was 70 eV, and the ionization temperature was 200◦C. Mass spectra were scanned every 0.2 s. For the analysis of the FAMEs, a DB-5 ms column (length, 30 m; internal diameter, 0.25 mm; Agilent Technologies, Inc., CA, USA) was used. For each sample, 1µL was injected on the column into a helium gas flow held constant at 1.4 mL min−1 . The column temperature was elevated from 40◦C to 320◦C at a rate of 6◦C min−1 and then kept at 320◦C for 1.15 min.

### SQD2 cDNA

RNA was extracted from standard and -P cultures using the phenol/chloroform method. Total RNA (500 ng) was used for the synthesis of cDNA with an oligo(dT)18 primer, random hexamers and Superscript II reverse transcriptase (Invitrogen, Carlsbad, CA). The primers used are listed in **Supplemental Table 1**. The gene sequences were then cloned into the pMD20 vector (Takara) for DNA sequence.

### Phylogenetic Analyses

Amino acid sequences were aligned using MAFFT v7.220 (Katoh and Standley, 2013). Gblocks 0.91b (Talavera and Castresana, 2007) was used to remove any poorly conserved regions. The phylogenetic analyses were performed using the maximum likelihood and the neighbor-joining methods in MEGA6 (Tamura et al., 2013), and a Bayesian analysis in MrBayes 3.2.3 (Ronquist et al., 2012). The amino acid substitution model for the maximum likelihood and Bayesian inference methods was selected using Aminosan (Tanabe, 2011). The maximum likelihood and neighbor-joining methods were performed based on the LG model + Gamma (eight categories) with 1000 bootstraps and the JTT model with 1000 bootstraps, respectively. The Bayesian analysis was performed based on the LG model + Gamma (eight categories) for 1,000,000 generations. Every 500th generation was sampled, and the first 200 trees were discarded as burn-in.

### Motifs Analysis

The analysis of the motifs was performed in silico using New PLACE (https://sogo.dna.affrc.go.jp/cgi-bin/sogo.cgi?sid=& lang=en&pj=640&action=page&page=newplace), a database of cis-acting regulatory DNA elements from plants, to find the binding motifs of transcription factors.

### Results

### Cell Growth and Tag Accumulation in Cells under P Deprivation

Nannochloropsis strain NIES-2145 was cultured to the logarithmic phase in f/2 medium. Cultures were then centrifuged and resuspended in fresh f/2 or in N-depleted (-N) or P-depleted (-P) f/2 medium. The cultures derived from logarithmic-phase cultures were inoculated at low cell densities (1 × 106 cells mL−1 ). Under N-deprived conditions, the Nannochloropsis cells were nearly white, but the cells exposed to P starvation were still green (**Figure 1**). Because C. reinhardtii cells diluted into P-depleted medium show substantial TAG accumulation (Iwai et al., 2014), we investigated whether TAG was accumulated in these nutrient-starved cells by quantifying lipid-derived FAMEs using GC. Cells had substantially increased TAG levels under both N and P deprivation (**Supplemental Figure 1**). Compared with cells grown in complete medium, the N-starved cells contained ∼11-fold and the P-starved cells contained ∼6-fold more TAG per cell after 13 days. The TAG accumulation in the N-starved cells is consistent with that in recent reports (Bondioli et al., 2012; Vieler et al., 2012b; Simionato et al., 2013). When the cultures were inoculated at 5 × 106 cells mL−1 in standard medium or –P medium, P-starved cells contained more TAG per cell than the corresponding cells maintained in the standard medium (∼7-fold more at 4 days and ∼47-fold more at 6 days; **Figure 2A**). P-starved cultures contained ∼4.6-fold more TAG per liter at 4 days and ∼4-fold more TAG per liter at 6 days than the corresponding culture maintained in the standard medium (**Figure 2B**).

### Increase in SQDG and SQD2 Homolog Promoter Activity during P Starvation

Lipids extracted from Nannochloropsis strain NIES-2145 were separated by two-dimensional TLC, followed by GCflame ionization detection and GC-MS. P starvation caused

FIGURE 1 | Growth of Nannochloropsis NIES-2145 under standard growth conditions, N starvation and P starvation. *Nannochloropsis* cells cultured to the logarithmic phase were inoculated into standard (control), -N and -P media. Cells were cultured for the indicated times after transfer before being photographed.

significant changes in lipid composition. The glycerolipid composition of Nannochloropsis strain NIES-2145 resembles that of the photosynthetic organism Arabidopsis thaliana (Li-Beisson et al., 2010), being comprised mostly of the prevalent glycoglycerolipids monogalactosyldiacylglycerol (MGDG), DGDG and SQDG, as well as the common phospholipids phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS), phosphatidylinositol (PI), and phosphatidylglycerol (PG). In addition, a betaine lipid diacylglycerol-N,N,N-trimethylhomoserine (DGTS) is present, as in C. reinhardtii (Sato and Furuya, 1985; Giroud et al., 1988). P deprivation resulted in elevated levels of SQDG and DGTS. PG, PC, and PS which contain phosphates in their head groups, decreased after P deprivation (**Figure 3**). This result corroborates previous reports on P deprivation in several species (Benning et al., 1993; Essigmann et al., 1998; Sato et al., 2000; Khozin-Goldberg and Cohen, 2006).

Orthologs of the SQD2 encoding the sulfoquinovosyl transferase (Yu et al., 2002) and a cyanobacterial counterpart, the sqdX, are highly conserved across different groups of algae and plants (**Figure 4**). We expected that the expression of homologs of the SQD2 gene would be induced during P starvation. To determine the expression of SQD2 homologs in Nannochloropsis strain NIES-2145, we searched the N. oceanica genome (Vieler et al., 2012b) using a BLAST algorithm to identify amino acid sequences with similarity to C. reinhardtii and Arabidopsis SQD2 genes. Using consensus sequences of the SQD2 genes, we amplified the corresponding cDNA from Nannochloropsis strain NIES-2145. We sequenced the cDNA and found that Nannochloropsis strain NIES-2145 has a homolog of the Arabidopsis SQD2 gene (**Figure 4**). **Figure 4** is based on protein comparison. Expression of the SQD2 gene homolog, as measured by qPCR, was induced after P deprivation in Nannochloropsis strain NIES-2145 (**Figure 5**). The Chlamydomonas SQD2 promoter is useful for improving transgene expression in C. reinhardtii when P is depleted (Iwai et al., 2014). Therefore, we expected that the Chlamydomonas SQD2 promoter would be available for inducing transgene expression to increase TAG production in Nannochloropsis under P-depleted conditions.

DGAT enzymes are important for TAG accumulation. During N starvation in N. oceanica IMET1, seven DGAT transcripts are up-regulated and six other DGAT transcripts are down-regulated (Li et al., 2014). However, it is unclear how DGAT transcripts are regulated during P starvation in Nannochloropsis strain NIES-2145. We therefore hypothesized that the overexpression of C. reinhardtii DGTT4 in Nannochloropsis would be a convenient way to increase the lipid content or alter the lipid composition of these cells.

### Enhanced Tag Accumulation and Changes in Fatty Acid Profiles in CrDGTT4-overexpressing Lines under P Deprivation

To explore CrDGTT4 as a tool to produce TAGs in Nannochloropsis cells, we expressed the CrDGTT4 coding sequence in Nannochloropsis under the control of the CrSQD2 promoter, which increases transcription levels during P

starvation in C. reinhardtii (Iwai et al., 2014). The relative level of overexpression of CrDGTT4 was measured using qPCR. CrDGTT4 mRNA levels in the overexpressing lines under P starvation were determined to be between 7- and 112-fold higher than in those under standard conditions (**Figure 6**), whereas no transcripts were detected in the wild-type and the empty vector controls. Thus, the SQD2 promoter from C. reinhardtii is quite useful as a heterologous promoter in Nannochloropsis strain NIES-2145. Three overexpressing lines, #8, 9, and 21, were then selected for the subsequent experiments. To confirm the effect of CrDGAT gene overexpression, we compared the lipid content and fatty acid composition of the wild-type and overexpressing lines. All lines were cultured to the logarithmic phase prior to P depletion, as was done in the previous experiments.

TAG accumulation increased in CrDGTT4-overexpressing lines compared with the control and wild-type lines in P-depleted medium. CrDGTT4-overexpressing lines contained ∼1.7 fold and ∼1.3-fold more TAG per cell than the empty vector control cells at 4 and 6 days, respectively (**Figure 7**, **Supplemental Figure 2**). We next investigated if the fatty acid composition of total lipids and TAGs was altered in the

FIGURE 4 | Phylogenetic tree analysis of SQD2. A phylogeny incorporating eukaryotic SQD2, cyanobacterial sqdX and the glycosyl transferase proteins of other organisms was constructed using Bayesian inference, maximum likelihood and neighbor-joining algorithms. It is based on a protein comparison. The topologies and branch lengths were calculated using maximum likelihood. Bootstrap values (maximum likelihood and neighbor-joining) higher than 70 and Bayesian posterior probabilities (Bayesian inference) higher than 0.9 are indicated under each branch (maximum likelihood/neighborjoining/Bayesian inference). The scale bar represents 0.2 amino acid substitutions per site. Sequence accession numbers or sequence resources for the phylogenetic tree are as follows: Archaea glycosyl transferase: *Methanocella paludicola* (YP\_003355097), *Methanocella* sp. (YP\_685710) and *Haloferax volcanii* (YP\_003535220); Alpha-proteobacteria glycosyl transferase: *Caulobacter crescentus* (YP\_002516166), *Novosphingobium aromaticivorans* (WP\_011446186), and *Maricaulis maris* (WP\_011642337); Cyanobacteria sqdX: *Cyanothece* sp. 1 (WP\_015784192), *Cyanothece* sp. 2 (WP\_012629991), *Synechococcus elongatus* (WP\_011243256), *Synechococcus* sp. (WP\_011429929), and *Cyanothece* sp. 3 (WP\_015956855); Other bacterial glycosyl transferases: *Truepera radiovictrix* (WP\_013178429), *Rubrobacter xylanophilus* (WP\_011564279), and *Alicyclobacillus acidocaldarius* (WP\_012810695); Rhodophyta SQD2: *Chondrus crispus* (XP\_005715110), and *Cyanidioschyzon merolae* (XP\_005538341); Stramenopile SQD2: *Nannochloropsis* sp. NIES-2145 (DDBJ Accession LC061442), *Nannochloropsis oceanica* IMET1 (BioProject: PRJNA202418, scaffold00247.g6721), *Nannochloropsis granulata* CCMP529 (BioProject: PRJNA65111, evm.model.NODE\_3998\_length\_11613\_cov\_18.861534.4), *Nannochloropsis salina* CCMP537 (BioProject: PRJNA62503, evm.model.NODE\_9973\_length\_179428\_cov\_24.782236.3), *Nannochloropsis gaditana* (EWM27152), *Phaeodactylum tricornutum* (JGI Protein ID: 50356), *Fragilariopsis cylindrus* 1 (JGI Protein ID: 207999), *Fragilariopsis cylindrus* 2 (JGI Protein ID: 158007), *Pseudo-nitzschia multiseries* 1 (JGI Protein ID: 252457), *Pseudo-nitzschia multiseries* 2 (JGI Protein ID: 145536), *Thalassiosira pseudonana* (JGI Protein ID: 38775), [Chlorophyta] *Ostreococcus tauri* (JGI Protein ID: 3203), *Micromonas* sp. (JGI Protein ID: 58169), *Chlorella* sp. (JGI Protein ID: 33086), *Volvox carteri* (Phytozome Transcript Name: Vocar20009459m), *Chlamydomonas reinhardtii* 1 (Phytozome Transcript Name: Cre01.g038550.t1.3), and *Chlamydomonas reinhardtii* 2 (Phytozome Transcript Name: Cre16.g689150.t1.2); Charophyta: *Klebsormidium flaccidum* 1 (BioProject: PRJDB718, kfl00392\_0070) and *Klebsormidium flaccidum* 2 (BioProject: PRJDB718, kfl00041\_0140), and Embryophyta (Phytozome Transcript Name): *Arabidopsis thaliana* (AT5G01220.1), *Populus trichocarpa* 1 (Potri.006G097600.1), *Populus trichocarpa* 2 (Potri.016G112600.1), *Oryza sativa* 1 (Os07g01030.1), *Oryza sativa* 2 (Os01g04920.1), *Oryza sativa* 3 (Os03g15840.1), *Selaginella moellendorffii* (170091), and *Physcomitrella patens* (Pp1s24\_194V6.1).

CrDGTT4-overexpressing cell line. One of the overexpressing lines, line #9, was analyzed because of TAG accumulation most increased in this line. The major total fatty acids in either control or P-depleted medium were eicosapentaenoic (20:5 ω-3), palmitoleic (16:1 ω-7), and palmitic (16:0) acids (**Figure 8**). This agreed with previous analyses (Maruyama et al., 1986; Patil et al., 2007). The levels of 16:0, 16:1, oleic (18:1 ω-9), and dihomo-gamma-linolenic (20:3 ω-6) fatty acids increased, whereas 16:3, linoleic (18:2 ω-6), arachidonic (20:4 ω-6) and 20:5 fatty acids decreased under P starvation conditions (**Figure 8**). The degree of unsaturation for each fatty acid decreased as the unsaturated fatty acids were replaced with saturated fatty acids. As for the total fatty acid composition, no statistically significant differences could be observed among the wild-type,

control and overexpressing lines (**Figure 8**). In the case of TAG fatty acids in the overexpressing line, the levels of 18:1 and 20:3 fatty acids were higher, whereas 16:0, 16:1, 16:2, and 16:3 fatty acids were lower than those in the wild-type or control lines during P starvation (**Figure 9**). The levels of 20:4 and 20:5 bound to TAGs (∼0.8% and ∼2.3%, respectively) were remarkably reduced under P starvation in comparison with the levels of these fatty acids in total fatty acids (∼1–3% and ∼15%, respectively; **Figure 9**). Under P starvation, the increase in the 20:3 fatty acid in the total lipids was a reflection of its increase in TAG. It is interesting to note that the level of 18:1 fatty acids was higher in the CrDGTT4-overexpressing cell line than in the control line under P starvation conditions (**Figure 9**). This is consistent with our previous report on CrDGTT4-overexpressing C. reinhardtii cell lines during P starvation (Iwai et al., 2014). These results showed that the C. reinhardtii SQD2 promoter is useful as a heterologous promoter in the secondary endosymbiotic alga Nannochloropsis strain NIES-2145.

### P Starvation did not Cause Significant Changes in Membrane Lipid Composition between the Control Line and Overexpressing Line

The overexpressing line showed alteration in TAG, whereas no serious alterations in the major membrane lipids compared with the control line under standard growth conditions or conditions of P stress, except for PS level (**Figure 10**, **Supplemental Figure 3**). PS decreased more in the overexpressing line than in the control line under P starvation (**Supplemental Figure 3**). We investigated whether the fatty

FIGURE 7 | Changes in the TAG content of CrDGTT4-overexpressing cells in response to P starvation. *Nannochloropsis* cells were transformed with pCrSQD2-CrDGTT4 to overexpress *CrDGTT4*. Cells cultured to a logarithmic phase under standard conditions were then inoculated into P-starved medium and cultured for 4 days. Three transformants, the empty vector control (control) and the wild type (WT) are shown. Values represent the mean ± SD from four independent replicates. Asterisks indicate a statistically significant difference compared with the empty vector control based on a two-tailed Student's *t*-test (\**P* < 0.05 and \*\**P* < 0.01).

acid composition of the major membrane lipids was altered in the CrDGTT4-overexpressing cell line (**Figures 11**–**13**, **Supplemental Figures 4**–**7**). During P-depleted conditions, the enhanced incorporation of 18:1(9) fatty acid was observed in the major plastidic membrane lipids MGDG, DGDG and PG and the extra-plastidic membrane lipids PC, PS, and DGTS (**Figures 11**, **12**), whereas the enhanced incorporation of 20:3 fatty acid was observed in MGDG, DGDG, PC, and DGTS (**Figures 11**, **13**). In the contrast, there were decreases of 18:1(9) and 20:3 fatty acids in PE. There was a global decrease in the long polyunsaturated fatty acid, 20:5 (**Figure 11** and **Supplemental Figure 4**), and an increase in saturated fatty acids, for example 16:0 (**Figure 11** and **Supplemental Figure 5**), during P-depleted conditions. Although large changes occur during P depravation, no significant differences could be observed in the major membrane lipids between the control line and overexpressing line, except for the decrease of 18:1 (9) fatty acid in the major plastidic membrane lipid PG.

### Discussion

Whereas TAG amounts increase under N starvation, MGDG, DGDG, SQDG, PG, and PI decrease in C. reinhardtii cells (Siaut et al., 2011; Sakurai et al., 2014) and the polar glycerolipid contents, represented mainly by MGDG, DGDG, and SQDG and phospholipids PE and PC, are reduced in Nannochloropsis strains (Simionato et al., 2013; Li et al., 2014; Martin et al., 2014). However, we found that SQDG and DGTS accumulated, replacing PG and PC, respectively, in Nannochloropsis cells under P starvation conditions (**Figures 3**, **10**, **Supplemental Figure 3**). This result corroborates previous reports on P deprivation in several species (Benning et al., 1993, 1995; Essigmann et al., 1998; Sato et al., 2000; Khozin-Goldberg and Cohen, 2006). In Monodus subterraneus, which is a freshwater microalga in the class Eustigmatophyceae, DGTS and SQDG increased under P starvation conditions (Khozin-Goldberg and Cohen, 2006). In A. thaliana, SQDG is synthesized to replace PG in chloroplasts during P starvation so that the amount of anionic thylakoid lipid is maintained (Essigmann et al., 1998; Yu and Benning, 2003).

and \*\**P* < 0.01).

In C. reinhardtii, mechanisms exist that maintain a constant total amount of SQDG and PG under sulfur- or P-limiting conditions (Sato et al., 2000). Therefore, our results suggest that SQDG substitutes for PG, to some extent, to sustain the functional activity of the decreased chloroplast membranes in Nannochloropsis, as in C. reinhardtii.

When the SQD2 promoter is applied, total TAG levels per cell after 4 days of P starvation were ∼1.7-fold greater in CrDGTT4-overexpressing lines compared with the control line (**Figure 7**). However, the difference between CrDGTT4 overexpressing lines and the control line after 6 days of P starvation was less than after 4 days (**Supplemental Figure 2**). During N starvation in N. oceanica IMET1, seven DGAT transcripts are up-regulated and six other DGAT transcripts are down-regulated (Li et al., 2014). Our results in Nannochloropsis might be explained by the up-regulation of Nannochloropsis's own DGATs, particularly in later stages of P depletion. The levels of 18:1(9) and 20:3 fatty acids bound to TAGs increased during P starvation in comparison with the levels of these fatty acids under standard conditions (**Figure 9**). The levels of these two fatty acids in TAG molecular species were higher in the CrDGTT4-overexpressing cell line than in the control line during P starvation. This may reflect the specificity of CrDGTT4. In contrast to the TAG fraction, the 18:1(9) molecular ratio of PG decreased in the CrDGTT4-overexpressing cell line than in the control line during P starvation (**Figures 11**, **12**). This fatty acid change may be associated with the preferential incorporation of the 18:1(9) molecular species into the TAG fraction.

It was unexpected that 20:3 fatty acid also increased in the CrDGTT4-overexpressing line (**Figures 11**, **13**). This may have been due to the suppression of desaturation in very long chain fatty acids, like 20:4 and 20:5 (**Figure 11** and **Supplemental Figures 6**, **7**), because of accelerated TAG accumulation. Although the exact reason is unclear, 20:3 fatty acid tends to accumulate under P starvation conditions, but not under N starvation conditions (Simionato et al., 2013; Martin et al., 2014). This fact suggests that a specific pathway for fatty acid incorporation is activated during P starvation.

The SQD2 is highly conserved from red algae to plants, as are its primary structures (**Figure 4**), and the algal SQD2 is also conserved in secondary algae, including Nannochloropsis. This may be due to the importance of the acidic membrane lipid SQDG, which is common to various algal species. The importance of SQDG for photosystem II was demonstrated clearly for C. reinhardtii and the cyanobacterium Synechocystis sp. PCC6803 through the characterization of mutants from each species that are deficient in the ability to synthesize SQDG (Minoda et al., 2002; Sato et al., 2003; Aoki et al., 2004; Sato, 2004). Recent work on higher plant SQD2 genes indicated that SQD2 is also involved with another acidic lipid, glucuronosyldiacylglycerol (Okazaki et al., 2013). It is possible that algal SQD2 also produces glucuronosyldiacylglycerol, which would thus be more important under P starvation conditions.

Our results showed that the SQD2 promoter from the green alga C. reinhardtii is useful as heterologous promoter in Nannochloropsis strain NIES-2145 cells. The results of this study suggest that there is a common expression control system in a

wide range of algal species, including primary and secondary microalgae, for adaptation to low P. This result, together with the occurrence of SQDG synthesis genes throughout algal species, suggests that a membrane remodeling system in chloroplasts under low-P stress is itself common among various microalgae and, thus, related genes and cis- and transacting elements are highly conserved. This also suggests that the SQD2 promoter from C. reinhardtii could be applied in various algae.

Arabidopsis SQD2 contains three PHR1-binding sequence (P1BS) motifs in its promoter. The PHR1 transcription factor binds to P1BS elements containing the consensus sequence "GNATATNC" in the promoters of a large number of P starvation–responsive genes (Franco-Zorrilla et al., 2004; Bustos et al., 2010; Pant et al., 2015). PHR1 is a well-described transcriptional regulator of P starvation in the MYB family. Most of the genes and their promoters that are involved in phospholipid degradation and glycolipid biosynthesis contain P1BS motifs in Arabidopsis (Pant et al., 2015). As there are no significant effects of a PHR1 null allele on DGDG synthesis during P limitation in Arabidopsis, a role for PHR1 in lipid remodeling was thus excluded (Gaude et al., 2008). However, glycerolipids, including MGDG, DGDG and SQDG, their composition and the expression of most lipid-remodeling gene transcripts analyzed were all altered in the phr1 mutant under P starvation when compared with the wild type (Pant et al., 2015). We used the C. reinhardtii SQD2 gene (Cre01.g038550) promoter for CrDGTT4 overexpression. Another SQD2 gene homolog (Cre16.g689150) exists in C. reinhardtii. Both C. reinhardtii SQD2 genes were up-regulated under P starvation (data not shown). We searched the promoter regions of these genes, and they do not contain P1BS motifs in their promoters but have binding motifs for many other transcription factors. C. reinhardtii SQD2 has 14 MYB or MYC binding domains in its promoter region, whereas Nannochloropsis oceanica SQD2 has 12 MYB or MYC binding domains in its promoter region (**Supplemental Figure 8**). There is no reliable evidence revealing which motif offers the most effective transcriptional regulation under P starvation. Further work is necessary to understand the regulation of lipid remodeling during P stress in algae.

## Author Contributions

MI prepared samples of Nannochloropsis strain NIES-2145 for each experiment. KH performed in silico analysis. YS performed GC-MS analysis. MI, KH, YS and HO wrote the manuscript. MS and HO planned the project.

## Acknowledgments

This work was partially supported by the KAO Corporation and JST CREST from the Ministry of Education, Sports, Science and Culture in Japan.

## Supplementary Material

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fmicb. 2015.00912

Supplemental Figure 1 | Changes in the TAG content of Nannochloropsis in response to N or P starvation. Cells cultured to the logarithmic phase under f/2 medium were then inoculated into f/2, -N or -P medium and cultured for 13 d.

#### Supplemental Figure 2 | Changes in the TAG content of

CrDGTT4-overexpressing cells in response to P starvation. *Nannochloropsis* cells were transformed with pCrSQD2-CrDGTT4 to overexpress *CrDGTT4*. Cells cultured to the logarithmic phase under standard conditions were then inoculated into control (+P) or -P medium and cultured for 6 days (6 d). The transformants (#8, #9, and #21), the empty vector control (VC) and the wild type (WT) are shown. Values represent the mean ± SD from three independent replicates.

Supplemental Figure 3 | Changes in major lipid classes in P-starved cells.

Black bars, the vector control (VC) 4 days in control medium (+P); white bars, VC 4 days in -P; dark gray bars, pCrSQD2-CrDGTT4 (#9) 4 days in control medium (+P); and light gray bars, pCrSQD2-CrDGTT4 (#9) 4 days in -P. Values are the mean ± SD from three independent experiments. Asterisks indicate a statistically significant difference compared with VC based on a two-tailed Student's *t*-test ( ∗*P* < 0.05). Small letters indicate statistically significant difference compared with the control medium based on a two-tailed Student's *t*-test (a*P* < 0.05 and b*P* < 0.01).

Supplemental Figure 4 | Quantitative analysis and ratio of 20:5 in the various lipids. Cells were cultured in control (+P) or -P medium for 4 days. (A) Each 20:5 in the various lipids is expressed in nmol per 106 cells. (B) The y-axis is ratio of 20:5 in each lipid per 106 cells. Values are the mean ± SD from three independent experiments. Asterisks indicate a statistically significant difference compared with VC based on a two-tailed Student's *t*-test (∗*P* < 0.05).

#### Supplemental Figure 5 | Quantitative analysis and ratio of 16:0 in the various lipids. Cells were cultured in control (+P) or -P medium for 4 days. (A) Each 16:0 in the various lipids is expressed in nmol per 106 cells. (B) The y-axis is ratio of 16:0 in each lipid per 106 cells. Values are the mean ± SD from three independent experiments. Asterisks indicate a statistically significant difference compared with VC based on a two-tailed Student's *t*-test (∗*P* < 0.05).

Supplemental Figure 6 | Quantitative analysis and ratio of 16:1 in the

various lipids. Cells were cultured in control (+P) or –P medium for 4 days. (A) Each 16:1 in the various lipids is expressed in nmol per 106 cells. (B) The y-axis is ratio of 16:1 in each lipid per 106 cells. Values are the mean ± SD from three independent experiments. Asterisks indicate a statistically significant difference compared with VC based on a two-tailed Student's *t*-test (∗*P* < 0.05).

Supplemental Figure 7 | Quantitative analysis and ratio of 20:4 in the various lipids. Cells were cultured in control (+P) or -P medium for 4 days. (A) Each 20:4 in the various lipids is expressed in nmol per 106 cells. (B) The y-axis is ratio of 20:4 in each lipid per 106 cells. Values are the mean ± SD from three independent experiments. Asterisks indicate a statistically significant difference compared with VC based on a two-tailed Student's *t*-test (∗*P* < 0.05).

Supplemental Figure 8 | MYB/MYC binding motif in the SQD2 promoter region of Nannochloropsis oceanica and Chlamydomonas reinhardtii. Yellow character, CTGTTA (+) strand MYB/MYC binding motif; red character, (+) and (−) strand MYB/MYC binding motif; blue character, (−) strand MYB/MYC

binding motif; green character, CDS; pale green character, UTR region. Supplemental Table 1 | Primer sequences.

### References

a key enzyme in triacylglycerol synthesis. Proc. Natl. Acad. Sci. U.S.A. 95, 13018–13023. doi: 10.1073/pnas.95.22.13018

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Iwai, Hori, Sasaki-Sekimoto, Shimojima and Ohta. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Changes in primary metabolism under light and dark conditions in response to overproduction of a response regulator RpaA in the unicellular cyanobacterium *Synechocystis* sp. PCC 6803

*Hiroko Iijima1†, Tomokazu Shirai2†, Mami Okamoto2, Akihiko Kondo2,3, Masami Yokota Hirai2 and Takashi Osanai1,2\**

*Edited by: Weiwen Zhang, Tianjin University, China*

#### *Reviewed by:*

*Lei Chen, Tianjin University, China Gopal K. Pattanayak, The University of Chicago, USA*

#### *\*Correspondence:*

*Takashi Osanai, School of Agriculture, Meiji University, 1-1-1, Higashimita, Tama-Ku, Kawasaki, Kanagawa 214-8571, Japan<EMAIL_ADDRESS>†These authors have contributed equally to this work.*

#### *Specialty section:*

*This article was submitted to Microbiotechnology, Ecotoxicology and Bioremediation, a section of the journal Frontiers in Microbiology*

> *Received: 10 July 2015 Accepted: 14 August 2015 Published: 26 August 2015*

#### *Citation:*

*Iijima H, Shirai T, Okamoto M, Kondo A, Hirai MY and Osanai T (2015) Changes in primary metabolism under light and dark conditions in response to overproduction of a response regulator RpaA in the unicellular cyanobacterium Synechocystis sp. PCC 6803. Front. Microbiol. 6:888. doi: 10.3389/fmicb.2015.00888* *1 School of Agriculture, Meiji University, Kawasaki, Japan, 2 RIKEN, Center for Sustainable Resource Science, Yokohama, Japan, 3 Department of Chemical Science and Engineering, Graduate School of Engineering, Kobe University, Kobe, Japan*

The study of the primary metabolism of cyanobacteria in response to light conditions is important for environmental biology because cyanobacteria are widely distributed among various ecological niches. Cyanobacteria uniquely possess circadian rhythms, with central oscillators consisting from three proteins, KaiA, KaiB, and KaiC. The two-component histidine kinase SasA/Hik8 and response regulator RpaA transduce the circadian signal from KaiABC to control gene expression. Here, we generated a strain overexpressing *rpaA* in a unicellular cyanobacterium *Synechocystis* sp. PCC 6803. The *rpaA*-overexpressing strain showed pleiotropic phenotypes, including slower growth, aberrant degradation of an RNA polymerase sigma factor SigE after the lightto-dark transition, and higher accumulation of sugar catabolic enzyme transcripts under dark conditions. Metabolome analysis revealed delayed glycogen degradation, decreased sugar phosphates and organic acids in the tricarboxylic acid cycle, and increased amino acids under dark conditions. The current results demonstrate that in this cyanobacterium, RpaA is a regulator of primary metabolism and involved in adaptation to changes in light conditions.

Keywords: amino acids, cyanobacteria, *Synechocystis*, response regulator, sugar metabolism

### Introduction

Cyanobacteria are organisms performing oxygenic photosynthesis that exist in various environmental niches such as fresh water, seawater, soil, and hot springs. Studying the regulatory mechanism of cyanobacterial metabolism is important in environmental biology and biotechnology. One of the most widely studied cyanobacteria is the non-nitrogen fixing species *Synechocystis* sp. PCC 6803 (hereafter *Synechocystis* 6803). *Synechocystis* 6803 cells grow fast and are naturally transformable with homologous recombination (Berla et al., 2013).

Cyanobacteria have a circadian rhythm and their central oscillator consists of three proteins KaiABC, first found in the unicellular cyanobacterium *Synechococcus* sp. PCC 7942 (hereafter *Synechococcus* 7942; Ishiura et al., 1998). An important issue for study in the circadian system is metabolic compensation, which is the persistence and entrainment of circadian rhythms in response to various nutrient conditions (Johnson and Egli, 2014). KaiC is an enzyme that phosphorylates and dephosphorylates its own residues in a strict order (Nishiwaki et al., 2007; Rust et al., 2007). ATP is a substrate of KaiC phosphorylation, and the ATP/ADP ratio is important for entrainment of the circadian clocks (Rust et al., 2011). KaiC phosphorylation is inhibited by ADP and integrates metabolic information into KaiC through the kinase-stimulation activity of KaiA (Rust et al., 2011). ATP is decreased under dark conditions and treatment with a dark pulse leads to a phase shift in the circadian clock (Rust et al., 2011). Oxidized quinones also input signals into the circadian clock in *Synechococcus* 7942 via the inhibition of KaiC phosphorylation (Kim et al., 2012). Thus, information on the availability of biochemical energy and light/dark conditions is transduced into circadian clocks. Glycogen metabolism entrains the circadian oscillator by providing ATP, and the mutants deficient in glycogen synthesis show the phenotype of a circadian clock that is hypersensitive to dark pulses (Pattanayak et al., 2014).

SasA is a histidine kinase interacting with KaiC in *Synechococcus* 7942 (Iwasaki et al., 2000). SasA is autophosphorylated by its histidine residue and the phosphate moiety is transferred to the cognate response regulator RpaA (Takai et al., 2006). The gene *rpaA* is the "regulator of phycobilisome association," whose protein is involved in energy transfer from phycobilisome to Photosystem I (Ashby and Mullineaux, 1999). The phosphorelay from SasA to RpaA is enhanced in the presence of KaiC (Takai et al., 2006). The knockout mutants of *sasA* or *rpaA* in *Synechococcus* 7942 grow normally under continuous light conditions, but the growth is severely retarded under light/dark cycle conditions (Takai et al., 2006). The knockout of *rpaA* alters the gene expression in *Synechococcus* 7942 widely; the genes, whose peaks of the expression are subjective dusk or dawn, are down-regulated or up-regulated, respectively, in the *rpaA* mutant (Markson et al., 2013). Chromatin-immunoprecipitation with high throughput sequencing (ChIP-Seq) analysis reveals 110 binding sites in the *Synechococcus* 7942 genome, which has A/T-rich motif included in the promoters of *kaiBC* and *rpoD6* (encoding an RNA polymerase sigma factor). *In vitro* assay demonstrates that phosphorylated RpaA binds with the promoter regions, while non-phosphorylated RpaA does not (Hanaoka et al., 2012; Markson et al., 2013).

The mechanistic implications of a circadian clock in *Synechocystis* 6803 have remained obscure due to the redundancy of *kaiABC* genes, with one *kaiA*, three *kaiB* (*kaiB1*–*B3*), and three *kaiC* (*kaiC1*–*3*; Kanesaki et al., 2012). The *kaiAB1C1* and *kaiC2B2* genes constitute an operon in the *Synechocystis* 6803 genome. KaiC1 phosphorylation is dependent on KaiA and KaiC directly interacting with KaiA (Wiegard et al., 2013). The phosphorylation of KaiC2 and KaiC3 is not dependent on KaiA, and therefore KaiAB1C1 proteins seem to be the central oscillator in *Synechocystis* 6803 (Wiegard et al., 2013). *Synechocystis* 6803 contains SasA (Hik8, sll0750) and RpaA (Rre31, slr0115) orthologs. Hik8 interacts with KaiC1, but not KaiC2 *in vivo* (Osanai et al., 2015). The knockout of *hik8* results in pleiotropic phenotypes, with the gene expression of enzymes in the glycogen catabolism, glycolysis, and the oxidative pentose phosphate (OPP) pathway altered (Singh and Sherman, 2005). The *hik8* overexpression also leads to changes in primary metabolism (Osanai et al., 2015). Glycogen and sugar phosphate levels are decreased under light conditions and amino acid levels such as glycine and lysine are increased by *hik8* overexpression (Osanai et al., 2015). The involvement of RpaA in salt and hyperosmotic stress has been shown by microarray, with the knockout of *rpaA*/*rre31* down-regulating the salt-induced gene expression (Shoumskaya et al., 2005). Nevertheless, in these studies, the involvement of RpaA in the regulation of primary metabolism and the effect of *rpaA* modification on metabolic alteration has remained unclear due to lack of metabolome data.

Here, we generated a *Synechocystis* 6803 strain overexpressing *rpaA*. The *rpaA*-overexpressing strain showed pleiotropic and similar phenotypes with the *hik8*-overexpressing strain. Genetic and metabolomic analyses indicate that RpaA plays pivotal roles in metabolic regulation under both light and dark conditions.

## Materials and Methods

### Bacterial Strains and Culture Conditions

The glucose-tolerant (GT) strain of *Synechocystis* sp. PCC 6803, isolated by Williams (1988), and the *rpaA*-overexpressing strain, designated as ROX310, were grown in modified BG-11 medium, which consisted of BG-110 liquid medium (Rippka, 1988) containing 5 mM NH4Cl (buffered with 20 mM HEPES-KOH, pH 7.8). Among GT substrains, the GT-I strain was used in this study (Kanesaki et al., 2012). Liquid cultures were bubbled with 1% (v/v) CO2 in air and incubated at 30◦C under continuous white light (ca. 50–70 µmol photons m−2 s−1). Growth and cell densities were measured at OD730 with a Hitachi U-3310 spectrophotometer (Hitachi High-Tech., Tokyo, Japan). Kanamycin (10 µg/mL) was added to ROX310 during pre-culture.

### Construction of Plasmids for *rpaA* Overexpression

A region of the *Synechocystis* 6803 genome encoding the *rpaA* (slr0115, *rre31*) ORF was amplified by PCR using KOD Plus Neo polymerase (Toyobo, Osaka, Japan) and the specific primers 5 - GAATTATAACCATATGCCTCGAATACTGATC-3 (forward) and 5 -ATCCAATGTGAGGTTAACCTACGTTGGACTACCG CC-3 (reverse). The amplified PCR fragment was inserted into the *Nde*I-*Hpa*I sites of the pTKP2031V vector, using an In-Fusion HD cloning kit (Takara Bio, Shiga, Japan). The resultant plasmid was confirmed by sequencing and transformed into GT-I as described previously (Osanai et al., 2011).

### Construction of Plasmids for Protease-Knockout Mutants

To construct protease-knockout mutants, the coding regions of proteases were amplified by PCR with KOD Plus Neo and the following primer sets: *clpB1*(slr1641)

5 -GGGAATTCTGCGGGATCGCAAACTA-3 (forward) and 5 - GCGCATGCGAGCGTTGAATGGCTTCG-3 (reverse); *clpB2* (slr0156) 5 -GGGAATTCTCCGCGCGTTTAACCTTG-3 (forward) and 5 -GCGCATGCCCCGCTCAGCTTTTTCT-3 (reverse); *clpC*(sll0020) 5 -GGGAATTCGCTTCCTGCCCGAT AAG-3 (forward) and 5 -GCGCATGCCACGTCTTCCAACAGG C-3 (reverse); *clpX*(sll0535) 5 -GGGAATTCGAAGGAACGG TGGCCAA-3 (forward) and 5 -GCGCATGCCCGTCGTTG TCCAACCA-3 (reverse); *degP*(slr1204) 5 -GGGAATTCGTG CTGGGGGGACATTT-3 (forward) and 5 -GCGCATGCCAGT TTGCCCACTAGGG-3 (reverse); *degQ*(sll1679) 5 -GGGAATT CCTTGGTTACGCCGCATC-3 (forward) and 5 -GCGCATG CACTATGCGCTGTAGGCG-3 (reverse); and *degS*(sll1427) 5 -GGGAATTCGTGGCCGTGCTTTTACT-3 (forward) and 5 -GCGCATGCCTTCCACCCGTTCTTGA-3 (reverse). PCR fragments were isolated with the Wizard SV Gel and PCR Cleanup System (Promega, Madison, WI, USA) and the fragments were digested with *Eco*RI and *Sph*I. Each of the resulting fragments was cloned into pUC119 (Clontech) digested with *Eco*RI and *Sph*I. The chloramphenicol-resistant cassette obtained by digesting pKRP10 (Reece and Phillips, 1995) with *Sma*I or *Pst*I was inserted into the *Hinc*II sites of *clpB1* and *clpC*, the

grown under light and dark conditions. (C) Growth of GT and ROX310

SmaI sites of *clpB2* and *clpX*, and the *Pst*I sites of *degP*, *degQ*, and *degS*. The resultant vectors were transformed into GT. The chloramphenicol-resistant (20 µg/mL) cells were isolated and streaked on plates several times.

### Immunoblotting

except for 15 min light per day).

Cells grown under light or dark conditions were collected by centrifugation (5,800 × *g* for 2 min), and the supernatant was removed by pipetting. The cells were frozen by liquid nitrogen. Cells were dissolved in PBS-T (137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, 1.5 mM KH2PO4, 0.1% Tween-20) and disruption by sonication and immunoblotting was performed as described previously (Osanai et al., 2014a). Antisera against SigE and GlgP(sll1356) were generated previously (Osanai et al., 2009, 2011).

#### RNA Isolation and Quantitative Real-Time PCR

RNA isolation and cDNA synthesis were performed as described previously (Osanai et al., 2014b). The cDNAs were synthesized with the SuperScript III First-Strand Synthesis System (Life Technologies Japan, Tokyo, Japan) with 2 µg of total RNA. Quantitative real-time PCR was performed with the

StepOnePlus Real-Time PCR System (Life Technologies) according to the manufacturer's instructions, using the primers listed in Supplementary Table S2. The transcript level of *rnpB*, which encodes RNaseP subunit B, was used as an internal standard as previously described (Osanai et al., 2015).

#### Glycogen Measurement

Glycogen levels were measured at the Biotechnology Center of Akita Prefectural University (Akita, Japan), as described previously (Osanai et al., 2014b).

#### LC–MS/MS Analysis

Equal amounts of cells (10 mL of cell culture with OD730 = 1.0) were harvested by rapid filtration. LC–MS/MS analysis was performed using a 100-mL aliquot of the upper phase as previously described (Osanai et al., 2014b). All metabolite analyses were performed with the cells grown without external carbon sources except CO2.

#### Amino Acid Analysis by GC–MS

Equal amounts of cells (50 mL of cell culture with OD730 = 1.0) were harvested by rapid filtration. Amino acids were quantified by GC-MS as previously described (Osanai et al., 2014a).

### Organic Acid Analysis by GC–MS

Equal amounts of cells (10 mL of cell culture with OD730 = 1.0) were harvested by rapid filtration using a previously described method (Osanai et al., 2014b). GC-MS was carried out using a GCMS-QP2010 Ultra equipped with a CP-Sil 8 CB-MS capillary column (30 m × 0.25 mm × 0.25 µm; Agilent, Palo Alto, CA, USA) as previously described (Osanai et al., 2014b).

### Results

#### Slower Growth of *rpaA*-Overexpressing Strain

We generated a strain overexpressing *rpaA* by fusing the promoter of *psbAII* (encoding Photosystem II D1 protein; **Figure 1A**) and the strain was named ROX310. Quantitative real-time PCR confirmed that the expression levels of *rpaA* in ROX310 were higher than in the wild-type, glucose-tolerant (GT) strain under both light and dark conditions (**Figure 1B**). Under both photoautotrophic and photomixotrophic conditions, the *rpaA*-overexpressing strain grew more slowly than did GT (**Figure 1C**). ROX310 grew similarly to GT under light-activated heterotrophic growth (LAHG) conditions until 1 day, although it lost viability under prolonged LAHG conditions (**Figure 1C**).

### Altered Protein and Transcript Levels of SigE in ROX310

Previously, the *hik8*-overexpressing strain showed aberrant protein degradation of SigE, RNA polymerase sigma factors activating sugar catabolism that is essential for dark/heterotrophic conditions, after the light-to-dark transition (Osanai et al., 2015). Immunoblotting demonstrated that *rpaA* overexpression reduced the degradation of SigE proteins under dark conditions (**Figure 2A**). To identify the proteases of SigE, we tested seven mutants lacking orthologous proteinases related to dark conditions in *Synechococcus* 7942 (Hosokawa et al., 2011), but these knockouts did not affect SigE protein levels (**Figure 2A**). The level of *sigE* transcripts was higher in ROX310 than in GT under both light and dark conditions, although the levels were similarly decreased in both strains by the lightto-dark transition (**Figure 2B**). We then quantified the levels of sugar catabolic enzymes by immunoblotting. The protein levels of GlgP(sll1356), one of two glycogen phosphorylases

involved in glycogen degradation in *Synechocystis* 6803, were not induced by the *rpaA*-overexpressing strain under dark conditions (**Figure 2C**).

We measured the transcript levels of genes related to sugar catabolism in ROX310 (**Figure 3**). The transcript levels of *pfkA*(sll0765) and *fbaII* were enhanced by *rpaA* overexpression under light conditions, while that of *pfkA*(sll1196) was repressed (**Figure 3**). The transcript levels of all 12 genes were decreased at 1 day after the light-to-dark transition in both GT and ROX310; however, there were higher transcript levels of these genes in ROX310 than in GT (**Figure 3**).

### Metabolome Analysis using the Cells Grown under Light and Dark Conditions

The glycogen levels were quantified under light and dark conditions. Glycogen rapidly disappeared in GT after the lightto-dark transition (**Table 1**). In ROX310, glycogen decreased after

TABLE 1 | Relative values of glycogen in GT and ROX310.

*Data represent mean* ± *SD from four independent experiments. Glycogen levels were calibrated relative to that in the GT strain under light conditions (set at 100%). ND indicates glycogen under detectable levels.*

the light-to-dark transition, but the glycogen degradation was slower than GT (**Table 1**).

LC–MS/MS analysis revealed that the levels of sugar phosphates (glucose-6-phosphate, ribose-5-phosphate, sedoheptulose-7-phosphate, fructose-6-phosphate, ribulose-5-phosphate, fructose-1,6-bisphosphate) and dihydroxyacetone phosphate were lower in ROX310 than in GT under both light and dark conditions (**Table 2**). Phosphoenolpyruvate levels were higher in ROX310 than in GT under light and dark conditions (**Table 2**). Fumarate and isocitrate levels were lower in ROX310 than in GT under light conditions (**Table 2**). Malate could not be detected in ROX310 under light conditions (**Table 2**). Organic acids in the TCA cycle were lesser in ROX310 than in GT under dark conditions (**Table 2**). NADP levels increased by *rpaA* overexpression, but other nucleotides did not (**Table 2**).

Amino acid analyses showed that the levels of glycine and proline were higher in ROX310 than in GT under light conditions (**Figure 4**). The levels of alanine, glycine, threonine, and lysine were higher in ROX310 than in GT under dark conditions (**Figure 4**). The level of ornithine was lower in ROX310 than in GT under dark conditions (**Figure 4**).

#### TABLE 2 | Relative levels of metabolites in primary metabolism in GT and ROX310.

*Data represent mean* ± *SD from four independent experiments. Metabolite levels were calibrated relative to that of the corresponding metabolite in GT under light conditions (set at 100%). Asterisks indicate statistically significant differences between GT and ROX310 (Student's t-test;* ∗*P < 0.05,* ∗∗*P < 0.005). The data from the GT strain under light and dark conditions were cited from our previous study (Osanai et al., 2015). P, phosphate; DHAP, dihydroxyacetone phosphate. Metabolites were quantified by LC–MS/MS, except for those marked by #, which were quantified by GC–MS.*

### Alteration in Transcript Levels of Genes Encoding Circadian Clocks

Finally, expression of *kaiABC* genes was quantified. The transcript levels of *kaiA, B1*, *C1*, *B2,* and *C2* were doubled by *rpaA* overexpression under light conditions (**Figure 5**). The transcript levels of *kaiB3* and *kaiC3* were marginally increased in ROX310 under light conditions (**Figure 5**). All the transcripts of *kaiABC* were decreased after 1 day of dark cultivation, and the levels remained higher in ROX310 than those in GT (**Figure 5**).

## Discussion

Here, we have performed genetic and metabolomic analyses using an *rpaA*-overexpressing strain and revealed that RpaA is involved in the regulation of primary metabolism in this unicellular cyanobacterium. The mechanism of circadian clocks in *Synechocystis* 6803, which could be different from *Synechococcus* 7942, has been less studied except Drs. Axmann's and Hellingwerf's groups (Wiegard et al., 2013; Beck et al., 2014; van Alphen and Hellingwerf, 2015), and we proceeded metabolome analyses with the circadian-related mutants of *Synechocystis* 6803. ChIP-Seq analysis in *Synechococcus* 7942 shows that RpaA binds the promoters of *kaiBC*, *sasA*, *rpaA*, and genes encoding sigma factors (*rpoD2, D5, D6*) and sugar catabolic enzymes (*glgP*, *malQ*, *zwf*, *opcA*, *gap1*, *fbaII*), activating their gene expression at night (Markson et al., 2013). These results are consistent with our analysis: the *rpaA* overexpression altered the gene expression of *sigE*, a sugar catabolic enzyme, and *kaiABC* (**Figures 2, 3,** and **5**). SigE is a sigma factor activating glycogen catabolism, glycolysis, and the OPP pathway (Osanai et al., 2005, 2011), and its expression peaks before night (Kucho et al., 2005). Combining the results of previous and current genetic analyses, the signal transduction from the circadian clock to sugar metabolism consists of the cascade of the proteins KaiABC-Hik8-RpaA-SigE in *Synechocystis* sp. PCC 6803, although further promoter analysis is required. Since RpaA altered the expression of *kaiABC* genes (**Figure 5**), feedback or feedforward regulation from RpaA to the central circadian oscillator may exist to entrain the clock by metabolic information.

The *rpaA*-overexpressing strain of *Synechocystis* 6803 exhibited several phenotypes. Results from the *rpaA*overexpressing strain of *Synechocystis* 6803 were similar to the *rpaA-*null mutant of *Synechococcus* 7942, which showed decreased glycogen catabolism (Diamond et al., 2015). The *rpaA* knockout mutant in *Synechocystis* 6803 exhibited high light sensitive phenotype (Majeed et al., 2012), which is consistent with our results that RpaA is important in light acclimation. A previous study demonstrated that introduction of RpaA(D53E; which mimics phosphorylated RpaA) restored the RpaA function, but that the introduction of RpaA(D53A; which mimics non-phosphorylated RpaA) could not (Markson et al., 2013). Therefore, the *rpaA* overexpression in our study may have increased non-phosphorylated RpaA in the *Synechocystis* 6803 cells, leading to phenotypes that were the mixture of gain-of-function and loss-of-function of RpaA. Introduction of a phospho-mimic RpaA into *Synechocystis* 6803 may be intriguing to distinguish these phenotypes.

Overexpression of *rpaA* led to phenotypes similar to those of the *hik8* overexpressor (Osanai et al., 2015); that is, growth defects under light-activated heterotrophic conditions (**Figure 1**), aberrant degradation of SigE after the light-todark transition (**Figure 2A**), accumulated transcripts during darkness (**Figure 3**), decreased levels of sugar phosphates (**Table 2**), and increased levels of several amino acids under dark conditions (**Figure 4**). These similarities may be due to the overexpression of *hik8* accelerated the phosphorylation of RpaA proteins. The glycogen metabolism mutant could not grow under dark/heterotrophic conditions (Osanai et al., 2005; Singh and Sherman, 2005; Tabei et al., 2007), or treatment with high salt and oxidative stress (Suzuki et al., 2010). Thus, the growth phenotypes of ROX310 under LAHG conditions may be caused by the changes in primary metabolism (**Figure 1C**). Genetic analyses have suggested that the mutants of Clp proteases alter circadian oscillation in *Synechococcus* 7942 (Holtman et al., 2005; Imai et al., 2013). Nevertheless the aberrant degradation of SigE proteins under dark conditions in the *rpaA*-overexpressing strain (**Figure 2A**), glycogen catabolism was slowed under dark conditions (**Table 1**). These results suggest that proteins other than SigE concertedly determine the degree of glycogen degradation in *Synechocystis* 6803. Several sugar catabolic regulators, including Hik31, Rre37, and AbrB are known in *Synechocystis* 6803 (Kahlon et al., 2006; Tabei et al., 2007; Yamauchi et al., 2011). For example, Rre37 preferentially activates the gene expression of glycogen catabolic and glycolytic enzymes such as *pfkA*(sll1196; Azuma et al., 2009), and thus, further study of the relationships among several sugar catabolic regulators is necessary to elucidate the regulatory mechanism of sugar catabolism in *Synechocystis* 6803.

Control of primary carbon metabolism by a circadian clock is an important theme in cyanobacteria (Diamond et al., 2015). Decrease in glycogen and sugar phosphates was observed during dark conditions (**Tables 1** and **2**), which indicates glycogen and sugar phosphates are positively correlated in this condition. On the other hand, organic acids in the TCA cycle kept higher levels under dark conditions (**Table 2**). Thus, organic acids in the TCA cycle were not correlated with the metabolites in glycolysis and the OPP pathway. The *rpaA*-overexpressing strain showed decreased levels of sugar phosphates under both light and dark conditions (**Table 2**), which is consistent with the fact that RpaA is important for the expression of genes related to glycogen catabolism in unicellular cyanobacteria (Diamond et al., 2015). Our immunoblotting showed that GlgP(sll1356) proteins were decreased with *rpaA* overexpression (**Figure 2C**), which may be one reason for the down-regulation of glycogen catabolism in this mutant. The metabolomic analysis also revealed that organic acids in the TCA cycle (fumarate, malate, and oxaloacetate) were lowered by *rpaA* overexpression (**Table 2**). The organic acids in the TCA cycle are an important pool of carbon sources in this cyanobacterium (Osanai et al., 2014a). Thus, the data also indicates that RpaA widely regulates primary metabolism related to the carbon sinks in this cyanobacterium. KaiC regulates the production of lysine, which has been shown to be lowered during light/dark cycles in *Synechococcus* 7942 (Diamond et al., 2015). The analysis showed lysine and glycine levels were up-regulated by *rpaA*-overexpression during the light-to-dark transition (**Figure 4**), demonstrating the involvement of RpaA in amino acid metabolism in response to light conditions. In summary,

### References

our metabolome analyses have revealed the RpaA-regulation in primary sugar and amino acid metabolism of *Synechocystis* 6803.

### Acknowledgments

This work was supported by the Ministry of Education, Culture, Sports, Science, and Technology, Japan, by a grant to TO from ALCA (Project name "Production of cyanobacterial succinate by the genetic engineering of transcriptional regulators and circadian clocks") from the Japan Science and Technology Agency.

### Supplementary Material

The Supplementary Material for this article can be found online at: http://journal*.*frontiersin*.*org/article/10*.*3389/fmicb*.* 2015*.*00888

*Synechococcus elongatus* PCC 7942 defective in glycogen synthesis. *Appl. Environ. Microbiol.* 76, 3153–3159. doi: 10.1128/AEM.00397-08

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2015 Iijima, Shirai, Okamoto, Kondo, Hirai and Osanai. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

## Characterization of *Chlamydomonas reinhardtii* phosphatidylglycerophosphate synthase in *Synechocystis* sp. PCC 6803

*Chun-Hsien Hung1, Kaichiro Endo2, Koichi Kobayashi2, Yuki Nakamura1,3\* and Hajime Wada2,4*

*1 Institute of Plant and Microbial Biology, Academia Sinica, Taipei, Taiwan, 2 Department of Life Sciences, Graduate School of Arts and Sciences, The University of Tokyo, Tokyo, Japan, 3 PRESTO, Japan Science and Technology Agency, Saitama, Japan, 4 CREST, Japan Science and Technology Agency, Saitama, Japan*

#### *Edited by:*

*Weiwen Zhang, Tianjin University, China*

#### *Reviewed by:*

*Jiangxin Wang, Arizona State University, USA Junbiao Dai, Tsinghua University, China Norihiro Sato, Tokyo University of Pharmacy and Life Sciences, Japan*

#### *\*Correspondence:*

*Yuki Nakamura, Institute of Plant and Microbial Biology, Academia Sinica, 128 Section 2, Academia Road, Nankang, Taipei 11529, Taiwan<EMAIL_ADDRESS>
#### *Specialty section:*

*This article was submitted to Microbiotechnology, Ecotoxicology and Bioremediation, a section of the journal Frontiers in Microbiology*

> *Received: 02 June 2015 Accepted: 31 July 2015 Published: 24 August 2015*

#### *Citation:*

*Hung C-H, Endo K, Kobayashi K, Nakamura Y and Wada H (2015) Characterization of Chlamydomonas reinhardtii phosphatidylglycerophosphate synthase in Synechocystis sp. PCC 6803. Front. Microbiol. 6:842. doi: 10.3389/fmicb.2015.00842* Phosphatidylglycerol (PG) is an indispensable phospholipid class with photosynthetic function in plants and cyanobacteria. However, its biosynthesis in eukaryotic green microalgae is poorly studied. Here, we report the isolation and characterization of two homologs (CrPGP1 and CrPGP2) of phosphatidylglycerophosphate synthase (PGPS), the rate-limiting enzyme in PG biosynthesis, in *Chlamydomonas reinhardtii*. Heterologous complementation of *Synechocystis* sp. PCC 6803 *pgsA* mutant by *CrPGP1* and *CrPGP2* rescued the PG-dependent growth phenotype, but the PG level and its fatty acid composition were not fully rescued in the complemented strains. As well, oxygen evolution activity was not fully recovered, although electron transport activity of photosystem II was restored to the wild-type level. Gene expression study of *CrPGP1* and *CrPGP2* in nutrient-starved *C. reinhardtii* showed differential response to phosphorus and nitrogen deficiency. Taken together, these results highlight the distinct and overlapping function of PGPS in cyanobacteria and eukaryotic algae.

Keywords: *Chlamydomonas reinhardtii*, chloroplast, glycerolipid, phosphatidylglycerol, photosynthesis

### Introduction

Photosynthetic membranes are highly specialized biological membranes that contain distinct yet conserved classes of polar glycerolipids MGDG, DGDG, SQDG, and PG among cyanobacteria, eukaryotic microalgae, and land plants (Omata and Murata, 1983; Dorne et al., 1990; Sakurai et al., 2006).  Biosynthesis of these lipids may be critical for photosynthesis.

**Abbreviations:** Chl, chlorophyll; DAG, *sn*-1,2-diacylglycerol; DCMU, 3-(3,4-dichlorophenyl)-1,1-dimethylurea; DGDG, digalactosyldiacylglycerol; MGDG, monogalactosyldiacylglycerol; MGlcDG, monoglucosyldiacylglycerol; PG, phosphatidylglycerol; PGPS, phosphatidylglycerophosphate synthase; PS, photosystem; SQDG, sulfoquinovosyldiacylglycerol.

Except for variation in fatty acid composition, the structural features of these four lipid classes are highly similar; however, the biosynthetic pathways may be diverse among cyanobacteria, eukaryotic algae and higher plants. For example, the most abundant lipid class, MGDG, is synthesized in *Synechocystis* sp. PCC 6803 by two steps (Sato and Murata, 1982): first DAG is glucosylated with UDP-glucose to form MGlcDG by MGlcDG synthase (Awai et al., 2006), which is then isomerized to MGDG by an epimerase (Awai et al., 2014). However, in *A. thaliana* and other seed plants, MGDG is synthesized by onestep galactosylation with UDP-galactose by MGDG synthases (Shimojima et al., 1997). In contrast, DGDG is produced by the further galactosylation of MGDG by DGDG synthases in both *Synechocystis* sp. PCC 6803 (Awai et al., 2007) and *A. thaliana* (Kelly and Dormann, 2002; Kelly et al., 2003).

Phosphatidylglycerol is the only major phospholipid class present in the photosynthetic membrane. It has a distinct contribution to photosynthesis. Gene knockout study revealed the crucial role of PG biosynthesis: disruption of *pgsA*, encoding PGPS in *Synechocystis* sp. PCC 6803, affects cell growth and photosynthetic activity unless PG is supplemented exogenously (Hagio et al., 2000). Arabidopsis possesses two PGPS, PGP1 and PGP2; knocking out *PGP1* severely impairs chloroplast biogenesis but not mitochondrial function (Hagio et al., 2002; Babiychuk et al., 2003), and double knockout of *PGP1* and *PGP2* further reduces PG levels to a trace amount, and causing an embryonic-lethal phenotype (Tanoue et al., 2014).

Much less is known about the biosynthesis of photosynthetic membrane lipids in *C. reinhardtii*. However, a distinct feature of chloroplast lipid metabolism has been shown: an involvement of chloroplastic galactoglycerolipid lipase in triacylglycerol production in nitrogen-starved *C. reinhardtii* (Li et al., 2012). Because the possible contribution of chloroplastic glycerolipids in triacylglycerol production is uniquely observed in *C. reinhardtii*, dissecting the similarity and distinctiveness of chloroplastic lipid biosynthesis in *C. reinhardtii* with reference to *Synechocystis* sp. PCC 6803 and *A. thaliana* is important.

In this study, we isolated two *PGPS* genes in *C. reinhardtii*, *CrPGP1* and *CrPGP2*, and assessed the molecular function by transforming them into a *pgsA* mutant of *Synechocystis* sp. PCC 6803. Moreover, gene expression profiles were examined in *C. reinhardtii* under phosphorus or nitrogen-starved conditions. The result showed distinct and overlapping functions of PGPS between cyanobacteria and eukaryotic algae.

## Materials and Methods

### Strains and Growth Conditions

The wild-type and *pgsA* cells of *Synechocystis* sp. PCC 6803 were grown photoautotrophically at 30◦C in BG-11 medium supplemented with 20 µM PG as described previously (Sakurai et al., 2003). Growth of cultures was monitored by determining optical density at 730 nm (OD730). Light was provided by fluorescent lamps with approximately 50–60 µmol photons m−2 s −1. *C. reinhardtii* strain CC-4351 was obtained from the Chlamydomonas Resource Center and transformed with empty pChlamiRNA2 plasmid (Molnar et al., 2009). Cells were photoheterotrophically grown in TAP medium (Gorman and Levine, 1965) at 22◦C. Nutrient deficiency was induced by collecting the cells by centrifugation (5 min at 3000 × *g*), washed twice with the respective media and subsequently resuspended in TAP medium, TAP medium without nitrogen (TAP-N), or phosphorus (TAP-P) by omitting NH4Cl, or replacing potassium phosphate with 1.5 mM KCl, respectively (Quisel et al., 1996).

## Molecular Cloning *CrPGP1* (Cre03.g162601)

A 937-bp fragment was amplified using cDNA synthesized from the total RNA of *C. reinhardtii* strain CC-503 (cw92 mt+) as the template with the primers CH223 and CH224 and cloned into pENTR/D\_TOPO to construct pCH067. Then, the cloned fragment was amplified with the primers CH772 and CH773 and inserted into *Nde*I and *Hpa*I sites of pTCP2031V to construct pCH167.

## *CrPGP2* (Cre02.g095106)

A 790-bp fragment was amplified using cDNA synthesized from the total RNA of *C. reinhardtii* strain CC-503 (cw92 mt+) as the template with the primers CH225 and CH226 and cloned into pENTR/D\_TOPO to construct pCH068. Then, the cloned fragment was amplified with the primers CH774 and CH775 and inserted into *Nde*I and *Hpa*I sites of the pTCP2031V vector which was designed to incorporate a gene of interest at a neutral site (*slr2031*) with the *psbA2* (*slr1311*) promoter and a chloramphenicol-resistance cassette (Satoh et al., 2001). The resulting plasmid pCH160 was used to transform the *pgsA* mutant of *Synechocystis* sp. PCC 6803 by homologous recombination. The strains, plasmids, and oligonucleotide primers used in this study are described in Supplementary Tables 1–3, respectively.

### Complementation Assay of the *Synechocystis* sp. PCC 6803 *pgsA* Mutant by *CrPGP1* and *CrPGP2*

For culture on BG-11 agar plates, 5 µl of liquid culture was used with serial 10-fold dilution for spotting from left to right starting at OD730 0.05 onto the plate with or without 20 µM PG. Plates were incubated photoautotrophically under 50∼60 µmol photons m−2 s−1 for 5 days at 30◦C to observe the growth phenotype. For liquid culture, cells were photoautotrophically grown in BG-11 media. Initial growth was started at OD730 0.1 by the stirring culture at 180 rpm, 50∼60 µmol photons m−2 s−1 at 30◦C.

### Genotype Analysis

Genomic DNA was isolated from cells of the wild type, *pgsA CrPGP1* and *pgsA CrPGP2* of *Synechocystis* sp. PCC 6803. The primers CH784 and CH785 were used for PCR analysis of genetic background; a 1,141-bp fragment could be amplified from wild-type genomic DNA, whereas a 2,341-bp fragment could be amplified from genomic DNA of *pgsA CrPGP1* or *pgsA CrPGP2*. PCR analysis of the insertion at *slr2031* involved the primers for CH982 and CH1000, for an expected 575-bp fragment from the wild type, 2,938-bp fragment from *pgsA CrPGP1* and 2,791-bp fragment from *pgsA CrPGP2*.

### Lipid Analysis

Lipids were extracted from intact cells as described in (Bligh and Dyer, 1959) and analyzed previously (Nakamura et al., 2003).

### RNA Extraction and qRT-PCR Analysis

Total RNA from *Synechocystis* sp. PCC 6803 cells grown in BG-11 media by stirring culture at 180 rpm, 50∼60 µmol photons m−2 s −1 at 30◦C was extracted using TRI reagent (Ambion) including DNase treatment and reverse-transcribed with SuperScript III (Invitrogen, Carlsbad, CA, USA) for cDNA synthesis. Quantitative RT-PCR involved the ABI 7500 Real Time PCR System (Applied Biosystems) with the oligonucleotide primers for *pgsA* (*slr 1522*; CH1028 and CH1029), *PGP1* (Cre03.g162601; CH919 and CH920), *PGP2* (Cre02.g095106; CH953 and CH954) and *RNase P subunit B* (*rnpB*; CH947 and CH948). Gene expression was normalized to that of *rnpB* (Yuzawa et al., 2014). Data were averaged by three technical replicates in the same run and three biological replicates in separate runs.

Total RNA extraction and cDNA synthesis from *C. reinhardtii* cells grown by stirring culture at 200 rpm, 50∼60 µmol photons m−2 s−1 at 22◦C, follow the method as described above. Oligonucleotide primers used are: *Chlamydomonas G-protein beta subunit-like polypeptide* (*CBLP*; Cre06.g278222; CH1076 and CH1077), *PGP1* (Cre03.g162601; CH919 and CH920), *PGP2* (Cre02.g095106; CH953 and CH954), *monogalactosyldiacylglycerol synthase1* (*MGD1*; Cre13.g585301; CH890 and CH891), *digalactosyldiacylglycerol synthase1* (*DGD1*; Cre13.g583600; CH1060 and CH1061), *UDP-sulfoquinovose synthase* (*SQD1*; Cre16.g656400; CH1062 and CH1063), *sulfoquinovosyldiacylglycerol synthase* (*SQD2*; Cre01.g038550; CH1064 and CH1065), *nitrate reductase1* (*NIT1*; Cre09.g410950; CH1072 and CH1073) and *phosphorus starvation response protein* 1 (*PSR1*; Cre12.g495100; CH1066 and CH1067). Gene expression was normalized to that of *CBLP* (Schloss, 1990; Chang et al., 2005). Data were averaged by three technical replicates in the same run and two biological replicates in separate runs. Annotation of genes for the lipid metabolism is according to (Li-Beisson et al., 2015). The primers used are listed in Supplementary Table 3.

### Assay of Photosynthetic Parameters

Photosynthetic oxygen-evolving activity from H2O to CO2 of intact cells was measured with a Clark-type oxygen electrode (Hansatech Instruments, Kings Lynn, UK) as described in (Gombos et al., 1991). Light from an incandescent lamp through a red optical filter was used for all oxygen evolution measurements at the light intensity 900 µmol photons m−2 s−1. The Chl concentration of cells, determined by the method of Porra et al. (1989) with 100% methanol extraction, was adjusted to 5 µg mL−1.

For measurements of 77 K fluorescence emission spectra, intact cells were suspended in BG-11 at 10 µg Chl mL−1. Fluorescence emission spectra of intact cells under 435 or 600 nm excitation were recorded at 77 K with a spectrofluorometer (RF-5300PC; Shimadzu) as described (Sakurai et al., 2007a).

Measurements of relaxation of flash-induced Chl fluorescence yield were performed in intact cells (5 µg Chl mL−1) by profiling the spectra following single flash excitation with or without 10 µM DCMU (Vass et al., 1999). Samples were incubated in darkness for 10 min before DCMU was added at a final concentration of 10 µM. The levels of *F*o and *F*m were normalized.

For measurements of absorption spectra, cells were grown by stirring culture at 180 rpm, 50∼60 µmol photons m−2 s−1 at 30◦C. Absorption spectra of pigments were determined directly in intact cell suspension (OD730≈0.1) by spectrophotometry (Beckman Coulter DU 800 Spectrophotometer). Spectra were normalized at 625 nm, the maximum absorption of phycobiliproteins, as described (Gombos et al., 2002).

### Results

### Isolation of Genes Encoding Putative PGPS from *C. reinhardtii*

To isolate genes encoding functional PGPS for the biosynthesis of PG in *C. reinhardtii*, we performed a homology search with the amino acid sequence of AtPGP1 (At2g39290), which plays a major role in biosynthesis of PG and is essential for thylakoid membrane development in *A. thaliana* (Hagio et al., 2002; Xu et al., 2002; Babiychuk et al., 2003). Two genes were found as putative genes of PGPS in *C. reinhardtii* and named *CrPGP1* (Cre03.g162601) and *CrPGP2* (Cre02.g095106), encoding 32.1 and 28.7 kDa protein, respectively. We compared the amino acid sequence of CrPGP1 and CrPGP2 with characterized PGPs in *A. thaliana* (AtPGP1 and AtPGP2) (Hagio et al., 2002; Xu et al., 2002; Babiychuk et al., 2003) and *Synechosystis* sp. PCC 6803 (PgsA) (Hagio et al., 2000) by creating multiple amino acid sequence alignment (**Figure 1A**). As can be seen, the amino acid residues conserved in all sequences of proteins with the CDP-OH-P motif (PF01066.9) indicated by asterisks were conserved in CrPGP1 and CrPGP2, suggesting that these proteins are functional PGPs. Interestingly, CrPGP1 has longer N-terminal sequence than CrPGP2, as is found between AtPGP1 and AtPGP2. Next, we compared the identity and similarity of these PGPs (**Figure 1B**). The value ranges from 31.8 to 81.1% in identity and from 62.6 to 91.0% in similarity. The identity and similarity of PGPs were higher between *C. reinhardtii* and *A. thaliana* than CrPGP1 and CrPGP2, and those with *Synechocystis* PgsA were similar between *C. reinhardtii* and *A. thaliana*.

To examine whether these two genes encode functional PGPS, we cloned *CrPGP1* and *CrPGP2* under the control of the *psbA2* promoter and stably transformed them into a neutral genomic site (*slr2031*) (Satoh et al., 2001) of the *Synechocystis* sp. PCC 6803 *pgsA* mutant, which is defective in PGPS activity (Hagio et al., 2000) (**Figure 2A**). To confirm the successful homologous recombination of *CrPGP1* and *CrPGP2* into the neutral genomic site in the *pgsA* mutant, we performed PCR-based genotyping for both *pgs*A (*slr1522*) and *slr2031* (**Figure 2B**). Compared to

among PGPs aligned in (A).

the wild type, both *pgsA CrPGP1* and *pgsA CrPGP2* mutants maintained the *pgsA* background and the introduced genes were fully segregated at the *slr2031* locus by the homologous recombination. We further investigated whether *CrPGP1* and *CrPGP2* were expressed in *pgsA* by comparing mRNA levels of *CrPGP1* in *pgsA CrPGP1, CrPGP2* in *pgsA CrPGP2*, and *pgsA* in wild type. As shown in **Figure 2C**, both *CrPGP1* and *CrPGP2* were expressed, whose levels were fivefold and eightfold higher than that of *pgsA*. Thus, under the *psbA2* promoter control, both *CrPGP1* and *CrPGP2* were expressed in *pgsA CrPGP1* and *pgsA CrPGP2*.

PGPs of *Arabidopsis thaliana* (AtPGP1 and AtPGP2) and PgsA of

### Complementation of *Synechocystis* sp. PCC 6803 *pgsA* Mutant with *CrPGPs*

Previous study showed that the *pgsA* mutant of *Synechocystis* sp. PCC 6803 requires exogenous supplementation of PG for growth (Hagio et al., 2000), so we compared the growth of the wild type, *pgsA*, *pgsA CrPGP1*, and *pgsA CrPGP2* on solid BG-11 media with or without 20 µM PG. The wild-type cells grew similarly in the presence or absence of PG, whereas the *pgsA* mutant showed rescued growth only with PG (**Figure 3A**). The *pgsA CrPGP1* and *pgsA CrPGP2* showed rescued growth even in the absence of PG, which suggests that *CrPGP1* and *CrPGP2* are functional PGPS to rescue the growth defect of *pgsA*. To further investigate the growth profiles of *pgsA CrPGP1* or *pgsA CrPGP2*, we monitored the growth in liquid BG-11 media without PG. Although the growth rate was significantly restored in the transgenic strains, it was inferior to that of the wild type, with the *pgsA CrPGP1* showed slightly better growth than *pgsA CrPGP2* (**Figure 3B**). Therefore, *CrPGP1* and *CrPGP2* could functionally complement the lethal phenotype of the *pgsA* mutant, although their growth remain slightly retarded as compared with the wild type.

### Lipid Composition of *pgsA CrPGP1* and *pgsA CrPGP2*

To examine whether the levels of PG and other polar glycerolipids are restored in *pgsA CrPGP1* or *pgsA CrPGP2*, we analyzed the composition of membrane lipid classes and their fatty acid composition. PG composition in the wild type was 11 mol% but was 8 and 5 mol% in *pgsA CrPGP1* and *pgsA CrPGP2*, respectively (**Figure 4A**). Moreover, composition of DGDG was

slightly higher in the transgenic strains than the wild type. Next, we analyzed the fatty acid composition of these lipid classes (**Figures 4B–E**). The fatty acid composition of MGDG, DGDG, and SQDG was fairly similar among the three strains, but that of PG was markedly decreased in 18:3 and 18:2 composition and increased in 16:0 in *pgsA CrPGP1* and *pgsA CrPGP2* (**Figure 4E**). Hence, *CrPGP1* and *CrPGP2* produced a significant level of PG in *pgsA,* but the level was slightly lower and fatty acid composition differed from that of the wild type.

### Pigment Content and Oxygen Evolution Activity of *pgsA CrPGP1* and *pgsA CrPGP2*

Previous studies have shown that PG deficiency reduces Chl content in *Synechocystis* sp. PCC 6803 and other cyanobacteria (Gombos et al., 2002; Wu et al., 2006; Bogos et al., 2010). To assess whether the expression of *CrPGPs* can restore the Chl content in *pgsA*, we examined absorption spectra of pigments in the whole cells of the wild type, *pgsA CrPGP1* and *pgsA CrPGP2* (**Figure 5**). The spectra was normalized at 625 nm, the maximum absorption