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Metabolic Engineering 38 (2016) 217–227 Contents lists available at ScienceDirect Metabolic Engineering journal homepage: www.elsevier.com/locate/ymben Original Research Article Introducing extra NADPH consumption ability significantly increases the photosynthetic efficiency and biomass production of cyanobacteria Jie Zhou a,1, Fuliang Zhang a,b,1, Hengkai Meng a,c, Yanping Zhang a, Yin Li a,n a CAS Key Laboratory of Microbial Physiological and Metabolic Engineering, Institute of Microbiology, Chinese Academy of Sciences, Beijing, China University of Chinese Academy of Sciences, Beijing, China c School of Life Sciences, University of Science and Technology of China, Hefei, China b art ic l e i nf o a b s t r a c t Article history: Received 5 April 2016 Received in revised form 28 June 2016 Accepted 4 August 2016 Available online 4 August 2016 Increasing photosynthetic efficiency is crucial to increasing biomass production to meet the growing demands for food and energy. Previous theoretical arithmetic analysis suggests that the light reactions and dark reactions are imperfectly coupled due to shortage of ATP supply, or accumulation of NADPH. Here we hypothesized that solely increasing NADPH consumption might improve the coupling of light reactions and dark reactions, thereby increasing the photosynthetic efficiency and biomass production. To test this hypothesis, an NADPH consumption pathway was constructed in cyanobacterium Synechocystis sp. PCC 6803. The resulting extra NADPH-consuming mutant grew much faster and achieved a higher biomass concentration. Analyses of photosynthesis characteristics showed the activities of photosystem II and photosystem I and the light saturation point of the NADPH-consuming mutant all significantly increased. Thus, we demonstrated that introducing extra NADPH consumption ability is a promising strategy to increase photosynthetic efficiency and to enable utilization of high-intensity lights. & 2016 International Metabolic Engineering Society. Published by Elsevier Inc. All rights reserved. Keywords: Photosynthetic efficiency Light/dark reactions cycle Energy balance Extra NADPH consumption Metabolic engineering 1. Introduction The continuing decline of arable land and the constantly growing population pose significant challenges to the food supply. It is estimated that global food production must achieve a 50% increase by 2030 to meet the global food demand (Covshoff and Hibberd, 2012; Singh et al., 2014). Photosynthesis is one of the fundamental components determining crop productivity and is perhaps the most important biochemical process on Earth. It converts solar energy into biochemical energy, while converting CO2 into organic compounds. Improving photosynthesis efficiency would significantly contribute to increasing the productivity of crops and other photosynthetic organisms, i.e. microalgae and cyanobacteria, which have shown great potential in production of fuels and chemicals from CO2 (Evans, 2013; Oliver and Atsumi, 2014; Stephenson et al., 2011; Xiong et al., 2015a). Improving photosynthetic efficiency is always of great interest in spite of its challenging nature (Evans, 2013; von Caemmerer and Evans, 2010). On the energy conversion side, researchers have been working on the optimization of light reactions by expanding the photosynthetically active radiation spectrum (Blankenship and n Correspondence to: Institute of Microbiology, Chinese Academy of Sciences, No. 1 Beichen West Road, Chaoyang District, Beijing 100101, China. E-mail address: [email protected] (Y. Li). 1 Contributed equally. Chen, 2013; Gan et al., 2014) and optimizing light-harvesting antenna complexes (LHC) (Blankenship and Chen, 2013; Melis, 2009; Ort et al., 2011; Work et al., 2012). On the carbon fixation side, increasing the catalytic efficiency of ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) has been the most challenging task (Khan, 2007). Increasing the catalytic efficiency of Rubisco through molecular engineering (Cai et al., 2014; Liu et al., 2010) and replacing the native Rubisco in tobacco with a Rubisco from cyanobacteria (Lin et al., 2014) are recent exciting examples. Alternative strategies to increase carbon fixation include introducing CO2 concentrating mechanisms, increasing the regeneration of ribulose-1, 5-bisphosphate to accelerate the Calvin cycle and reducing or engineering photorespiration (Covshoff and Hibberd, 2012; Kebeish et al., 2007; Shih et al., 2014). Photosynthetic reactions are usually divided into light reactions and dark reactions; however, light reactions and dark reactions are not absolutely separated. Light energy is converted into chemical energy, in the form of ATP and reducing equivalent NADPH, via a series of photochemical reactions during light reactions. During dark reactions, CO2 is reduced into organic compounds via the Calvin cycle at the expense of the ATP and NADPH that are generated from the light reactions. The generation and consumption of ATP and NADPH are therefore the key for coupling the light and dark reactions. Theoretical analysis showed that 2.57 ATP/2 NADPH are generated in the light reactions via linear electron transport (Fig. 1), whereas 3 ATP/2 NADPH are required for CO2 http://dx.doi.org/10.1016/j.ymben.2016.08.002 1096-7176/& 2016 International Metabolic Engineering Society. Published by Elsevier Inc. All rights reserved. 218 J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 Fig. 1. Diagram of ATP and NADPH-coupled production in the light reaction and the NADPH consumption pathway-isopropanol synthetic pathway constructed in S. 6803. The isopropanol synthetic pathway is an acetyl-CoA-dependent pathway. From acetyl-CoA to isopropanol, three exogenous enzymes were required: coenzyme A transferase (CTFAB, UniProt accession number P33752 and P23673) and acetoacetate decarboxylase (ADC, UniProt accession number P23670), which were obtained from Clostridium acetobutylicum DSM 1731 to convert acetoacetyl-CoA to acetone; and secondary alcohol dehydrogenase (sADH, UniProt accession number P25984) from Clostridium beijerinckii to convert acetone to isopropanol. fixation in the dark reactions (Kramer and Evans, 2011; Marcus et al., 2011). This indicates that the ATP generated in light reactions (2.57 ATP) is insufficient to meet the energy requirement (3 ATP) for CO2 fixation in dark reactions. Such an imbalanced energy supply/requirement between the light and dark reactions might be one of the rate-limiting factors affecting photosynthetic efficiency (Kramer and Evans, 2011; Marcus et al., 2011). Considering from another perspective, if the ATP generated from the light reactions met precisely the demand of ATP in the dark reactions, the expected ATP/NADPH ratio in light reactions would be 3 ATP/2.33 NADPH, as ATP generation is tightly coupled with NADPH production via linear electron transport in the light reactions (Kramer and Evans, 2011). This would result in an NADPH imbalance, as only 2 NADPH will be consumed in dark reactions. We therefore hypothesized that increasing consumption of NADPH generated from the light reactions might improve the coupling between the light and dark reactions, thus increasing the photosynthetic efficiency. This hypothesis was tested by introducing an NADPH-consuming pathway into an oxygenic, photosynthetic, prokaryote cyanobacterium, Synechocystis sp. PCC 6803 (hereafter termed as S. 6803), with the aim to better balance the ATP/NADPH between light and dark reactions. A series of biochemical analyses demonstrated that this approach significantly increased the photosynthetic efficiency and biomass production of S. 6803. 2. Materials and methods 2.1. Strains and growth conditions The strains used are listed in Table S1. Escherichia coli strain DH5a was used as the host for vector construction. Wild-type S. 6803 and its mutants were grown in BG11 medium at 30 °C at an illumination intensity of approximately 100 μmol photons/m2/s, as described previously (Nielsen et al., 2013; Zhou et al., 2012). Chloromycetin (10 μg/ml) and/or kanamycin (10 μg/ml) were added to the medium when necessary. 2.2. Construction of vectors The plasmids used and constructed in this work are listed in Table S2. Vectors were constructed to delete the phaCE and pta genes or to express the ctfAB, adc and sadh genes in S. 6803. All primers used are listed in Table S3. Plasmid pSM5 was constructed by inserting the Pcpc560-sadh expression cassette, which comprised the promoter Pcpc560 and the structural gene sadh, into the Bam HI site of plasmid pSM1 (Zhou et al., 2012). The Pcpc560-sadh expression cassette was synthesized by GENEWIZ. Inc. (China) and the nucleotide sequence of sadh was optimized for the preferred codon usage of S. 6803. Plasmid pSM6 was constructed by inserting the Pcpc560-ctfAB and PrbcL-adc expression cassettes into the Xho I site of pSM2 (Zhou et al., 2012). The Pcpc560-ctfAB and PrbcL-adc expression cassette were synthesized by GENEWIZ. Inc. (China) and the nucleotide sequences of the Pcpc560-ctfAB and PrbcL-adc were optimized for the preferred codon usage of S. 6803. 2.3. Construction of mutant strains Mutants of S. 6803 were constructed by transforming S. 6803 with plasmids listed in Table S2. Transformations were performed as previously described (Lindberg et al., 2010). All constructed strains are listed in Table S1. Briefly, strain SM6 was constructed by integration of plasmid pSM6 at the pta locus of SM2 (Zhou et al., 2012) via double crossover homologous recombination. Strain SM7 was constructed by integration of plasmids pSM5 and pSM6 at the phaCE and pta loci of S. 6803, respectively. Strain SM7* was constructed by integration of plasmids pSM5 at the phaCE loci of S. 6803. 2.4. Enzyme assays Crude cell extracts were prepared as previously described (Zhou et al., 2014c), with some modifications. S. 6803 wild-type and mutant cells grown to mid-exponential growth phase were collected by centrifugation (14,000g, 4 °C, 2 min) and washed with prechilled buffer. The cells were then disrupted using the silicon carbide disruption method (Zhou et al., 2014a) in prechilled buffer J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 for the enzyme assay described below. The Bradford method (Zhou et al., 2014c) was used to determine the total protein in the crude cell extracts. The specific activity of CoA transferase (CTFAB) was determined by monitoring the decrease in absorbance at 310 nm, which corresponded to the disappearance of acetoacetyl-CoA, as previously described (Clark et al., 1989), using a spectrophotometer (Specter MAX 190, Molecular Devices, CA, USA). The reaction mixtures contained 110 mM Tris chloride (pH 7.5), 5.5% (vol/vol) glycerol, 20 mMMgCl2, 0.1 mM acetoacetyl-CoA, 0.32 M potassium acetate, and 1-5 μg of protein in the crude cell extracts. The specific activity assay for acetoacetate decarboxylase (ADC) was performed as described previously (Kusakabe et al., 2013). The reaction mixtures contained 30 mM sodium acetate buffer (pH 5.8), 30 mM lithium acetoacetate and 1-5 μg protein in the crude cell extracts. The consumption of lithium acetoacetate was monitored at 270 nm using a spectrophotometer. The specific activity assay for secondary alcohol dehydrogenase (SADH) was carried out as previously described (Hanai et al., 2007). The total reaction mixture contained 50 mM Tris-HCI (pH 7.5), 1 mM dithiothreitol, 0.2 mM NADPH, crude cell extracts (1– 5 μg protein) and 6.7 mM acetone. The consumption of NADPH was determined by measuring the decrease in absorbance at 340 nm. 219 molecule, propidium iodide, which can permeate the cellular membranes of dead cells and bind to nucleic acids, as previously described (Riccardi and Nicoletti, 2006). Samples were analyzed using a flow cytometer (FACSCalibur, Becton Dickinson, San Jose, CA, USA) equipped with a laser emitting at 488 nm and an optical filter FL1 (530/30 nm). The collected data were analyzed using FlowJo software (Tree Star, San Carlos, CA, USA). 2.8. Carbon fixation rate determination To calculate the net CO2 uptake rate by each strain, the initial and remnant CO2 concentrations after cultivation were measured as previously described (Xiong et al., 2015b) with modification. Briefly, cells in the exponential growth phase were harvested, washed once with BG11, resuspended in 130 ml BG11 at OD730 of 0.2 (approximately 3 mg dry cell weight), and cultivated for an additional 2 h. Then 130 ml cell suspension was transferred to a sealed tube with a volume of 140 ml. The total inorganic carbon of BG11 medium and the culture after two hours cultivation was released by addition of concentrated HCl (Xiong et al., 2015b). The CO2 in 1 ml headspace gas was quantified through the RTX-QBOND capillary column (30 m; 0.32 mm inside diameter; 10 mm film thickness; RESTEK Corporation, Bellefonte, USA) using the GC2014 gas chromatograph (Shimadzu, Kyoto, Japan) equipped with a TCD detector (Gong et al., 2015). 2.5. Isopropanol production assay 2.9. Chlorophyll fluorescence of PS II analysis For isopropanol production assay, the wild-type S. 6803 and its mutants were grown in 250 ml flasks containing 50 ml BG11 medium with an initial OD730 of 0.4. The incubation was carried out in a shaking incubator (30 °C, 200 rpm) under constant illumination intensity of 100 μmol photons/m2/s for about 3 weeks. Growth was monitored by measuring the OD at 730 nm every 2 days. To quantify the isopropanol produced, 0.5 μL of culture supernatant was analyzed using a GC-2014 gas chromatograph (Shimadzu, Kyoto, Japan) equipped with a flame ionization detector and an Rtx-WAX capillary column (30 m, 0.25 mm inside diameter, 0.25 mm film thickness; Agilent Technologies, Santa Clara, CA, USA) as previously described (Kusakabe et al., 2013). Isopropanol was used as the internal standard. To determine the quantum efficiency of PS II, chlorophyll fluorescence analysis of S. 6803 and its mutants grown to exponential growth phase was performed on a Dual-PAM-100 instrument (Walz, Germany), according to the manufacturer's instructions. To exclude the effect of phycobiliprotein, a red light detector (RD) was used. After 4 min of dark adaptation, slow induced curve and light response curve of cell suspension at an OD730 of 3.5 were measured to get parameters of chlorophyll fluorescence kinetics, including the maximum quantum yield (Fv/ Fm), the effective quantum yield (Y(II) ¼Fv’/Fm’) and the relative electron transport rate (rETR(II)) of PS II. The intensity of measure light and saturation pulse were 0.1 μmol/m2/s and 5000 μmol/m2/ s, respectively. 2.6. NADPH and ATP Assays 2.10. Measurement of P700 signal Wild-type Synechocystis and mutants cells grown to exponential growth phase were collected by centrifugation (14,000g, 4 °C, 2 min) and washed with prechilled PBS buffer. The cells were then lyzed using the lysis buffer associated with the Enhanced ATP Assay Kit, S0027 (Beyotime Biotechnology, Shanghai, China). The NADPH content was determined by using an NADP/NADPH Quantification Colorimetric Kit (Biovision, Catalog-K347-100), according to the manufacturer's instructions. The concentration of NADPH was calculated according to an NADPH standard curve and expressed as pmol/OD730. The ATP content was determined by using an Enhanced ATP Assay Kit, S0027 (Beyotime Biotechnology, Shanghai, China) according to the manufacturer's instructions. The concentration of ATP was calculated according to an ATP standard curve and expressed as nmol/OD730. To determine the quantum efficiency of PS I, the redox state of the reaction center chlorophyll of PS I (P700) was determined from its absorbance change in the near-infrared (wavelength 870– 830 nm) simultaneously with the measurement of chlorophyll fluorescence of PS II, using a Dual-PAM-100 instrument (Walz, Germany), as previously described (Klughammer and Schreiber, 2008; Wientjes and Croce, 2012). After 4 min of dark adaptation, slow induced curve and light response curve of cell suspension at an OD730 of 3.5 were measured to obtain PS I maximum quantum yield (Pv/Pm), PS I effective quantum yield (Y(I) ¼ Pv’/Fm’), the relative electron transport rate (rETR(I)) via PS I, and PS I complementary quantum yields of non-photochemical energy dissipation, Y(ND) and Y(NA). The intensity of measure light and saturation pulse were 0.1 μmol/m2/s and 5000 μmol/m2/s, respectively. Far-red light of 820 nm was used to measure the absorbance and far-red light of 875 nm was used as control as previously described (Klughammer and Schreiber, 2008; Pfündel et al., 2008). 2.7. Flow cytometry Flow cytometry was used to analyze the ratio of dead cells in the cultures of S. 6803 and its mutants. Cells grown to stationary phase were collected, washed once and resuspended in 1 ml of PBS at 1–2 106 cells/ml. Cells were stained with a fluorescent 2.11. Oxygen evolution The oxygen evolution rate of all strains was measured using a LED1/W-1503illuminator (Hansatech) and an Oxygraph Plus Clark- 220 J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 type electrode (Hansatech) as previously described (Gonzalez-Esquer et al., 2015) with modification. To determine the light response curves of oxygen evolution rate, cultures grown to exponential growth phase were harvested and resuspended in BG11 at an OD730 of 6.0. Two microliter cell suspension was supplemented with 100 mM bicarbonate and the steady state rate of oxygen evolution was determined at different light intensity at 30 °C for 2 min 3. Results 3.1. Design and introduction of extra NADPH consumption ability into S. 6803 Two criteria need to be considered when designing the experiments. One is that such perturbation should directly target the consumption of NADPH to drive the coupling of the light and dark reactions, while minimally affecting the cellular physiology. The other consideration is that such perturbation should be sufficiently strong to disturb the existing ATP/NADPH ratio in S. 6803, which has been established through long-term evolution and supposes to be quite stable. To meet the first criterion, we proposed to introduce a synthetic pathway that only consumes NADPH but does not affect energy metabolism. The pathway selected was the isopropanol biosynthetic pathway, which does not exist in S. 6803, but is found in some heterotrophic microbes, such as Clostridium beijerinckii (Dai et al., 2012; Ismaiel et al., 1993). To connect the heterogeneous isopropanol biosynthetic pathway with the central metabolism of S. 6803, three exogenous enzymes, namely ctfAB, encoding coenzyme A transferase and adc, encoding acetoacetate decarboxylase from Clostridium acetobutylicum DSM 1731 (Bao et al., 2011), and sadh, encoding secondary alcohol dehydrogenase from C. beijerinckii NRRLB593 (Dai et al., 2012), need to be introduced (Fig. 1). The assembled isopropanol pathway would connect with central metabolism at the acetoacetyl-CoA node, which is already present in S. 6803. This pathway only consumes NADPH and does not consume ATP as the secondary alcohol dehydrogenase is solely dependent on NADPH (Dai et al., 2012). We calculated the ΔG of the proposed isopropanol biosynthetic pathway (from acetyl-CoA to isopropanol). The ΔG is 67.6 KJ/ mol, suggesting this pathway is thermodynamically highly favorable as a decarboxylation reaction is included in the pathway. In addition, we planned to use a strong promoter that was developed in a previous study (Zhou et al., 2014c) to express the genes involved in this pathway, so as the second criterion can be met. To introduce a functional isopropanol biosynthetic pathway, we constructed two mutants, SM6 with two genes (ctfAB þadc), which was expected to produce acetone only and SM7 with three genes (ctfABþ adc þsadh), which was expected to produce isopropanol at the expense of NADPH. As indicated in Fig. 2A, mutant SM6 was constructed by placing the codon-optimized adc and ctfAB genes under a promoter PrbcL (Takeshima et al., 1994) and a strong promoter Pcpc560 (Zhou et al., 2014c), respectively. Because knockout pta gene would not completely block acetate production, this DNA fragment was inserted into the pta site of the mutant SM2 (Zhou et al., 2012). Mutant SM7 was constructed by placing the codon- Fig. 2. The genetic modifications in SM strains for extra NADPH consumption. (A) Genetic modifications. (B) Whole cell PCR using primers for the recombinant cassette at the phaCE and pta sites; one primer which is 100 bp upstream the recombinant cassette and the other primer which is located in the recombinant cassette were used to confirm the complete segregation of all mutants. (C) Whole cell PCR with specific primers demonstrating the integration of each gene into the chromosome of individual mutants of S. 6803. J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 221 Fig. 3. Growth profile analysis of all strains and the time course of isopropanol production by NADPH consumption strain SM7. (A) Growth profile. (B) Cell death rate analysis by flow cytometry. (C) CO2 fixation rate. (D) Time course of isopropanol production by strain SM7. Error bars indicate standard deviation (SD) of the data from three independent experiments. For each experiment, three technical replicates were performed. optimized sadh under Pcpc560, and the Pcpc560-sadh was inserted into the phaCE site of mutant SM6. Therefore, the sole genomic difference between mutant SM7 and its direct control mutant SM6 was that the phaCE gene (Zhou et al., 2012) in mutant SM6 was replaced with the Cmr gene, whereas in mutant SM7, the phaCE gene was replaced with the Cmr gene and sadh. Complete segregation and gene insertions were verified by PCR and sequencing (Fig. 2B and C). Metabolite spectrum analysis indicated that mutant SM6 produced 108.5 mg/L acetone, while mutant SM7 produced 226.9 7 5.4 mg/L isopropanol, when incubated in BG11 medium at 30 °C under constant white light for 20 days (Figs. 2A and 3D). Enzyme assays showed that the specific activities of CTFAB, ADC and SADH in mutant SM7 were 8.6 72.4, 140727 and 528 779 μmol/min/mg crude extract, respectively. These results demonstrated that the introduced isopropanol pathway was functional. Subsequently, a series of physiological and biochemical analyses were performed to investigate whether the introduced isopropanol biosynthetic pathway increased the photosynthetic efficiency. 3.2. Introduction of extra NADPH consumption ability increased the ATP/NADPH ratio First, the intracellular concentrations of the two photochemical products of the light reactions, ATP and NADPH, were determined in the wild-type (WT), SM6, and SM7, respectively (Table 1). Concentration of NADPH in strain SM7 was the lowest among all the strains tested, which was approximately 40% lower than that in control strain SM6 and WT. Moreover, the concentration of ATP in strain SM7 was 77% and 59% higher than that in strain SM6 and WT, respectively. Taken together, the ATP/NADPH ratio of strain SM7 (0.55) was significant higher than that of SM6 (0.18) and WT Table 1 Intracellular concentration of NADPH and ATP in WT and mutant cells of S. 6803. Strain NADPH (pmol/OD730) ATP (nmol/OD730) WT SM6 SM7 193.5 7 4.27 190.83 7 7.06 112.83 7 3.94 39.28 7 1.39 35.23 7 1.63 62.53 7 0.89 WT, wild-type; Standard deviation (7 SD) of the data from three independent experiments. For each experiment, three technical replicates were performed. (0.20). These data suggested that the introduced synthetic isopropanol pathway not only decreased the intracellular NADPH level, but also increased intracellular ATP level, therefore resulted in a significantly increased ATP/NADPH ratio in strain SM7. 3.3. Introduction of extra NADPH consumption ability increased the biomass production of S. 6803 Growth curves of different mutants were assessed under the same growth conditions as those described previously (Fig. 3A). Strain SM7 grew significantly faster than the other strains. At the first day of cultivation, the growth rate of mutant SM7 was 81%, 42%, and 33% higher than that of the WT strain, mutant SM2 (in which the pta gene was replaced with the Kmr gene and phaCE gene was replaced with the Cmr gene) (Zhou et al., 2012), and mutant SM6, respectively (Fig. S1). At 20 days of cultivation, the dry cell concentration of strain SM7 was 84%, 62%, and 48% higher than that of strains WT, SM2, and SM6, respectively (Table S4). To exclude the possibility that the high OD730 and dry cell weight of mutant SM7 was resulted from an increased amount of dead cells, the ratio of dead cells over total cells in each culture was determined by flow cytometry. The results showed that strain 222 J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 SM7 had the least dead cells among the four strains tested during the whole culture process (Fig. 3B). Notably, the ratio of dead cells in the cultures of the acetone-producing mutant SM6, the control strains SM2, and the WT, all increased sharply after 20 days of cultivation. Rather, the ratio of dead cells in the culture of the NADPH-consuming strain SM7 only increased slightly after 20 days of cultivation, while the isopropanol concentration also ceased to increase (Fig. 3D). These suggest that increasing NADPH consumption can increase biomass production and cellular activity. Furthermore, the carbon-fixation rates of all strains were determined according to the amount of net CO2 uptake during a period of cultivation (Xiong et al., 2015b). Fig. 3C showed that the carbon-fixation rate of the NADPH-consuming strain SM7 was 38%, 40%, and 36% higher than that of the control strains WT, SM2 and SM6, respectively. 3.4. Introduction of extra NADPH consumption ability increased photochemical efficiency of PS II in S. 6803 Light reactions take place in photosystem II (PS II, P680 as the reaction-center) and photosystem I (PS I, P700 as the reactioncenter). The photochemical efficiency is dependent on the activity of both PS II and PS I. Chlorophyll fluorescence kinetics indicates the activity of PS II (Shen and Luo, 2011; Singh et al., 2005; Thamatrakoln et al., 2013). To investigate the effect of introducing extra NADPH consumption ability on the activity of PS II, parameters of chlorophyll fluorescence kinetics, including the maximum quantum yield (Fv/Fm), the effective quantum yield (Y(II) ¼ Fv’/Fm’), and the relative electron transport rate (rETR(II)) of PS II, were determined (Table 2). Fv/Fm of all strains were approximately 0.42, showing there was no significant difference among the maximum quantum yield of PS II of strain SM7 and its control strains WT, SM2 and SM6. Due to state transitions, the cyanobacterial Fv/Fm is not as accurate as the Fv/Fm of higher plants to reflect the maximum quantum yield of PS II (Campbell et al., 1998). Therefore, the Y(II) and rETR(II) of each strain were measured under normal light condition of 100 μmol/m2/s (Table 2). Y(II) and rETR(II) of strain SM7 were 16.2% and 15.8% higher than that of control strains WT, SM2 and SM6, respectively, indicating that extra NADPH consumption increased the activity of PS II. To further analyze the effect of higher light intensity on NADPH consumption strain SM7 and its control strains WT, SM2 and SM6, the light response curve of rETR(II) was investigated (Fig. 4B). The rETR(II) of strain SM7 was significant higher than that of its control strains WT, SM2 and SM6 under the tested light intensity (ranging from 142 to 2292 μmol photons/m2/s). Interestingly, the rETR(II) of all control strains reached the highest value (approximately 21) under 588 μmol photons/m2/s; whereas the rETR(II) of SM7 kept increasing and reached the highest value (38.68 71.45) under 1182 μmol photons/m2/s. This means that the saturation light points of all control strains were around 600 μmol photons/m2/s, whereas the saturation light point of SM7 increased to 1200 μmol photons/m2/s. Introducing extra NADPH consumption ability thus resulted in a significant increase of the saturation light point and a 1.84 fold increase of the highest rETR(II) value (rETR(II) of strain SM7 under 1182 μmol photons/m2/s versus rETR(II) of control strains under 588 μmol photons/m2/s). Notably, the rETR(II) of all control strains decreased to approximately 10 under 2292 μmol photons/m2/s, whereas the rETR(II) of strain SM7 was still significantly higher than the highest rETR(II) of all control strains. Subsequently, the light response curves of Y(II) of all strains were plotted under different light intensity (Fig. S2). The Y(II) of SM7 was 1.8-fold and 2.4-fold that of the control strains WT, SM2 and SM6 under 588 μmol photons/m2/s and 1182 μmol photons/ m2/s, respectively (Fig. S2). Moreover, the Y(II) of all control strains decreased to nearly 0 under 2292 μmol photons/m2/s, whereas strain SM7 still exhibited an effective quantum yield, which was higher than that of all control strains under 1182 μmol photons/ m2/s. These suggested that introducing extra NADPH consumption ability not only increased the photosynthetic efficiency of PS II, but also enabled strain SM7 to utilize light with higher intensity. Photosynthetic oxygen is another sensitive indicator of photosynthetic activity of PS II, and an increased oxygen evolution may indicate the increased activity of the light reactions (Xiong et al., 2015b). Light response curve of rETR(II) showed that the saturation light point of SM7 was approximately 1200 μmol photons/m2/s. We therefore investigated the effect of introducing extra NADPH consumption ability on photosynthetic oxygen evolution under 600 μmol photons/m2/s, the semi-light saturation point of SM7. Fig. 4A shows that there are no significant difference among the oxygen evolution rate of strain WT, SM2, and SM6, under 600 μmol photons/m2/s. Interestingly, the oxygen evolution rates of strain SM7 were approximately 86% higher than the rest of the strains. This shows that strain SM7 exhibits a significantly higher photosynthetic oxygen evolution activity, indicating the light reactions of strain SM7 was improved. 3.5. Introduction of extra NADPH consumption ability contributed to energy conversion of PS I in S. 6803 To further investigate the effects of introducing extra NADPH consumption ability on energy conversion efficiency of PS I, the maximum quantum yield (Pv/Pm), the effective quantum yield Y (I), and relative electron transport rate rETR(I) were analyzed (Klughammer and Schreiber, 2008). Pv/Pm of all strains were approximately 0.94, showing there were no significant difference among the quantum yield of strain SM7 and control strains WT, SM2 and SM6. However, Y(I) and rETR(I) of strain SM7 were approximately 14.9% and 15.6% higher than the Y(I) and rETR(I) of control strains WT, SM2 and SM6, respectively, under normal light condition of 100 μmol/m2/s (Table S5). Similarly, we investigated the light intensity response curve of Table 2 Measurement of chlorophyll fluorescence kinetics of PS II, Fv/Fm and Y(II), rETR(II) of NADPH consumption strain (SM7) and control strains (SM6, SM2 and WT) under different light intensity. Light intensity (μmol/m2/s) Parameter WT 0 100 100 588 588 Fv/Fm Y(II) rETR(II) Y(II) rETR(II) 0.420 0.264 10.97 0.097 24.04 7 7 7 7 7 0.014 0.014 0.261 0.003 1.615 SM2 SM6 SM7 0.435 7 0.021 0.263 7 0.025 10.86 7 1.237 0.094 7 0.002 23.18 7 1.386 0.430 7 0.000 0.281 7 0.007 11.77 7 0.049 0.090 7 0.003 22.17 7 0.484 0.440 7 0.028 0.326 7 0.020 13.63 7 0.778 0.153 7 0.003 37.82 7 0.572 Fv/Fm, maximum quantum efficiency of PS II. Y(II), effective quantum efficiency of PS II. rETR(II), relative electron transport rate of PS II. Standard deviation ( 7 SD) of the data from three independent experiments. For each experiment, three technical replicates were performed. J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 223 Fig. 4. Analyses the activity of PS II and PS I of NADPH consumption strain (SM7) and control strains (SM6, SM2 and WT). (A) O2 evolution rate under 600 μmol/m2/s. (B) Light response curve of rETR(II), relative electron transport rate of PS II. (C) Light response curve of rETR(I), relative electron transport rate of PS I. Error bars indicate standard deviation (SD) of the data from three independent experiments. For each experiment, three technical replicates were performed. rETR(I) of all strains (Fig. 4C). The rETR(I) of strain SM7 was also higher than that of its control strains WT, SM2 and SM6 under the tested light intensity. The rETR(I) of all control strains reached the highest value (approximately 38) under 588 μmol photons/m2/s, whereas the rETR(I) strain SM7 reached the highest value (86.08 73.05) under 1182 μmol photons/m2/s, which was 2.3 fold that of all control strains. The light intensity response curve of Y(I) (Fig. S3) showed that the Y(I) of strain SM7 was approximately 1.3fold and 2.8-fold that of control strains WT, SM2 and SM6 under 588 μmol photons/m2/s and 1182 μmol photons/m2/s, respectively (Fig. S3). These suggested that introducing extra NADPH consumption ability increased the energy conversion efficiency of PS I. The quantum yield of PS I can be divided into Y(I), the effective quantum yield and non photochemical dissipation which includes Y(ND), the quantum yield of non-photochemical energy dissipation due to donor side limitation; and Y(AD), the quantum yield of non-photochemical energy dissipation due to acceptor side limitation. Thus, Y(I) þ Y(NA) þ Y(ND) ¼1. To understand the fate of quantum yield of PS I, the non photochemical dissipation of PS I quantum yield was analyzed (Table 3). Under normal light intensity of 100 μmol photons/m2/s, both quantum yields of nonphotochemical energy dissipation due to donor side limitation Y (ND) and acceptor side limitation Y(NA) of strain SM7 were much lower than that of the control strains, indicating that the introducing extra NADPH consumption ability decreased the quantum yields of non-photochemical energy dissipation of PS I. As a result, the photochemical energy was significantly increased. Under higher light intensity of 588 μmol photons/m2/s, although the total quantum yields of non-photochemical energy dissipation of PS I (Y(ND) þY(NA)) of strain SM7 was much lower than that of the control strains, Y(ND) of strain SM7 decreased 34% while Y(NA) of SM7 increased 82%, compared to Y(ND) and Y(NA) of all control strains, respectively (Table 3). 4. Discussion In this study, we hypothesized and tested a new strategy for Table 3 Measurement of redox state of P700 to detect Y(I), rETR(I), Y(ND) and Y(NA) under light intensity 588 μmol/m2/s of NADPH consumption strain (SM7) and control strains (SM6, SM2 and WT). Light intensity (μmol/m2/s) Parameter WT 588 588 588 588 Y(I) rETR(I) Y(ND) Y(NA) 0.153 37.70 0.853 0.013 7 7 7 7 0.005 1.516 0.043 0.023 SM2 SM6 SM7 0.150 7 0.005 37.16 7 5.666 0.845 7 0.024 0.013 7 0.017 0.148 7 0.010 36.52 7 2.485 0.854 7 0.025 0.005 7 0.012 0.341 84.16 0.637 0.024 Y(I), effective quantum efficiency of PS I. rETR(I), relative electron transport rate of PS I. Y(ND), quantum yield of non-photochemical energy dissipation due to donor side limitation. Y(NA), quantum yield of non-photochemical energy dissipation due to acceptor side limitation. Standard deviation ( 7 SD) of the data from three independent measurements. For each experiment, three technical replicates were performed. 7 7 7 7 0.006 1.516 0.016 0.015 224 J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 improving the photosynthetic efficiency of a photoautotrophic organism, cyanobacterium S. 6803. The core idea of this strategy was to better couple the light reactions and dark reactions by introducing extra NADPH consumption capability. Using this strategy, the activity of the light reactions was remarkably increased, resulting in a significantly increased photosynthetic growth rate. Strikingly, the photosynthetic oxygen evolution and photochemical efficiency all significantly increased, at a much higher light intensity, indicating strain SM7 has not only increased its photosynthesis efficiency, but also acquired ability for utilizing stronger light. To address the insufficient supply of ATP in dark reactions, the first attempt was to increase the absorption of light energy by increasing light intensity or optimizing the light capture system to produce more ATP (Blankenship and Chen, 2013; Melis, 2009). As ATP synthesis is tightly coupled with NADPH production via linear electron transport chain in a fixed ratio (Fig. 1) (Kramer and Evans, 2011), an increase in ATP synthesis is accompanied by an increase in NADPH production. However, the demand for NADPH is much less than that for ATP in the dark reactions. Increasing ATP production using the above approach would result in the accumulation of NADPH, which is toxic to cells. Therefore, directly increasing ATP synthesis via increasing absorption of light energy would have a limited positive impact on increasing the photosynthetic efficiency (Kozaki and Takeba, 1996; Nogales et al., 2012). Alternatively, some photosynthetic organisms contain an accessorial cyclic electron transport chain, from which ATP can be generated without forming NADPH (Allen, 2003). In such a case, three ATP and two NADPH could be generated in the light reactions via both linear and cyclic electron transport chains (Allen, 2003; Kramer and Evans, 2011), which would possibly satisfy the energy needs in the dark reactions. However, considering that photorespiration, nitrogen assimilation and other anabolic processes all consume ATP, the extra ATP generated via the cyclic electron transport chain might not meet the ATP requirement for cell growth (Allen, 2003; Kramer and Evans, 2011; Noctor and Foyer, 1998). In this work, extra ATP generation in light reactions was achieved by introducing the capability to consume extra NADPH. The introduced extra NADPH consumption capability not only resulted in a significant increase of oxygen revolution, relative electron transfer rate and effective quantum yield of both PS II and PS I, but also resulted in a significant increase of cell growth rate, CO2 fixation rate and dry cell weight. Compared to the directed control strain SM6, the introduced extra NADPH consumption capability resulted in an approximately 0.6 g/L extra biomass formation (Table S4). Generally, for photoautotrophic microorganisms, formation of 1 g biomass requires approximately 100 mmol ATP (Kliphuis et al., 2012). In this work, formation of 0.6 g/L extra biomass would require 60 mmol/L ATP, indicating introducing extra NADPH consumption capability stimulated ATP production. Notably, although the specific growth rate at early stage of cultivation and the biomass concentration after 20 days cultivation of strains SM2 and SM6 were significantly higher than that of WT (Fig. S1; Table S4), the CO2 fixation rates of strains WT, SM2 and SM6 were the same (Fig. 3C). This indicates that there was no difference among the photosynthetic efficiency of WT, SM2 and SM6. The better growth of strain SM2 and SM6 might be attributed to the disruption of acetate and PHB synthetic pathways in these mutants, which may contribute to cell growth due to redistribution of carbon flux (Zhou et al., 2014b). Moreover, through analysis of quantum yields of non-photochemical energy dissipation of PS I, we learned that the introduced extra NADPH consumption ability mainly decreased the donor side non-photochemical energy dissipation under high-intensity light conditions. The increase of acceptor side non-photochemical energy dissipation in strain SM7 under high-intensity light conditions indicates the energy generated from PS I cannot be completely utilized by carbon fixation, thus extra energy was dissipated as heat in PS I. Namely, it was the acceptor side (carbon fixation) that limited the energy conversion of PS I. Thus, if carbon fixation rate of strain SM7 can be improved, the efficiency of light reactions of strain SM7 under high-intensity light conditions can be further increased. Photosynthetic microorganisms usually utilize light with low or medium intensity (below 600 μmol photons/m2/s). Light intensity higher than 600 μmol photons/m2/s usually leads to photoinhibition or photodamage (Tikkanen et al., 2014; Vinyard et al., 2014). However, the light intensity in nature often fluctuates. For instance, the maximum light intensity in daytime often fluctuates from 990 μmol photons/m2/s to 1486 μmol photons/m2/s (He et al., 2015). Therefore, capabilities for utilizing fluctuating and high intensity lights are desired features for photosynthetic organisms. To utilize high-intensity light, the D1 protein of PS II of Synechococcus elongatus PCC 7942 was engineered by point mutagenesis (Vinyard et al., 2014). Although the oxygen evolution rate of each mutant strain were all lower than that of wild type, the authors provided fundamental design principles for engineering photosynthesis with optimal photochemical efficiencies for growth under low versus high light intensities (Vinyard et al., 2014). In this work, the mutant strain SM7 functions well in a broad range of light intensity, and its saturation light point, 1200 μmol photons/m2/s, falls perfectly in the range of the maximum light intensity in nature. This indicates that introducing extra NADPH consumption ability into photosynthetic organisms could be a useful strategy for utilizing fluctuating and high-intensity lights. The isopropanol synthetic pathway was constructed in Synechococcus sp. PCC 7942; however, the effects of NADPH consumption on growth and photosynthesis was not investigated (Hirokawa et al., 2015; Kusakabe et al., 2013). Because the production of isopropanol under light and aerobic growth conditions was too low to be detected, isopropanol was produced under dark and anaerobic conditions and 26.5 mg/L isopropanol was produced (Kusakabe et al., 2013). Subsequently, the isopropanol production was increased to 146 mg/L by optimizing isopropanol production conditions, shifting cells from dark and anaerobic conditions to light and aerobic conditions (Hirokawa et al., 2015). In this study, we optimized the expression of the introduced three enzymes in S. 6803 and the production of 226.9 mg/L isopropanol was solely achieved under light and aerobic conditions, while dark and anaerobic conditions were required for isopropanol production in previous isopropanol works (Hirokawa et al., 2015; Kusakabe et al., 2013). In terms of isopropanol production, this study is an extension of our previous acetone work (Zhou et al., 2012). In our previous acetone work, the production of acetone was very low and only 36 mg/L acetone was detected under dark and anaerobic conditions. It is known that codon optimization and application of strong promoters for expression of key enzymes can significantly increase the production of target chemicals (Angermayr and Hellingwerf, 2013; Guerrero et al., 2012; Lan and Liao, 2012; Zhou et al., 2014b). Therefore, in this work, the three genes required for construction of the isopropanol pathway were codon-optimized and placed under a strong promoter Pcpc560 before introducing into in S. 6803 (Zhou et al., 2014c). Consequently, acetone production was increased to 108.5 mg/L and 226.9 mg/L isopropanol production was achieved, under light and aerobic growth conditions. The actual titer of isopropanol could be higher, as the evaporated isopropanol during 20 days aerobic cultivation was not taken into account. To further investigate whether the increased biomass J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 225 Fig. 5. The time course of isopropanol conversion from acetone by NADPH consumption strain SM7* (A) and growth profile analysis of all strains (B). All strains were supplied with 2 g/L acetone. In A, square represented acetone assay; circle represented isopropanol assay. Error bars indicate standard deviation (SD) of the data from three independent experiments. For each experiment, three technical replicates were performed production was due to the introduced extra NADPH consumption ability, we constructed a mutant strain SM7*, in which only the Pcpc560 controlled codon-optimized sadh was inserted into the phaCE site of S. 6803. Acetone, the direct substrate of the sadh encoded NADPH dependent secondary alcohol dehydrogenase (SADH), was added to the medium of mutant strain SM7*, WT and its direct control strain SM1, in which the phaCE gene was replaced with the Cmr gene (Zhou et al., 2012) to a final concentration of 1800 mg/L. Isopropanol was detected in the medium of strain SM7* but not in that of WT and SM1 (Fig. 5A), suggesting acetone can be converted to isopropanol by strain SM7*. The gradually decreased concentration of acetone in the medium of strain WT and SM1 (Fig. 5A) was simply due to evaporation, as the blank medium containing 1800 mg/L acetone but free of S. 6803 cells also showed similar acetone profile (data not shown). This also suggested the depletion of acetone in the culture of strain SM7* includes evaporated acetone. Growth profile under light and aerobic conditions showed strain SM7* grew significantly faster than WT and its direct control strain SM1 (Fig. 5B). At 14 days of cultivation, the dry cell concentration of strain SM7* was 39% and 49% higher than that of strains WT and SM1, respectively (Table S6). These data further confirmed that consumption of extra NADPH stimulated the growth of S. 6803 cells. As NADPH is the major form of reducing equivalent in cyanobacteria (Takahashi et al., 2008), NADPH was used as a cofactor for chemicals production in engineered cyanobacteria by introducing NADPH-dependent enzymes or transhydrogenases to convert NADPH into NADH to aid the NADH-dependent enzymes (Angermayr et al., 2014; Atsumi et al., 2009; Lan and Liao, 2012; Liu et al., 2011; Luan et al., 2015; Oliver et al., 2013; Savakis et al., 2013). In previous efforts to produce chemicals at the expense of NADPH, for instance, isobutyraldehyde (Atsumi et al., 2009), butanol (Lan and Liao, 2012), fatty acids (Liu et al., 2011), lactate (Angermayr et al., 2014), 2,3-butanediol (Oliver et al., 2013; Savakis et al., 2013) and ethanol (Luan et al., 2015), the growth of mutant cells all decreased or did not change as compared with the wild type cells. Among these efforts, Oliver and Machado reported an increased oxygen evolution of an engineered cyanobacterium, associated with production of 2.38 g/L 2,3-butanediol (Oliver et al., 2013). The author simply ascribed the increased oxygen evolution to the overproduction of the target chemical (Oliver et al., 2013). However, increased cell growth was not observed in the 2,3-butanediol production work (Oliver et al., 2013). In another work of sucrose production in cyanobacteria, enhanced biomass accumulation and photosynthetic activity was observed in sucrose-exporting cells under high osmolality conditions (Ducat et al., 2012). Although the author speculated that consumption of excess reducing equivalents during sucrose production might be one of factors (Ducat et al., 2012), the growth of sucrose-exporting cells was still much lower than the growth of wild type cells under light and aerobic growth conditions. Our work represents the first report of having a significant increase in biomass accumulation upon expressing a pathway that utilizes NADPH. Improving photosynthetic efficiency would have a great impact on agriculture. Our study demonstrated that improving energy imbalance between light reactions and dark reactions is a simple but useful strategy to improve the photosynthetic efficiency and to enable utilization of high-intensity light for cyanobacteria. Compared with other strategies, such as optimization of Rubisco or introducing carbon concentrating mechanisms into C3 plants, we propose that up-regulating the activity NADPH-dependent enzymes in C3 plants might contribute to establishing a new ATP/ NADPH balance that would improve its photosynthetic efficiency, biomass production and utilization of high-intensity light. The idea of establishing a new ATP balance could be further tested in other oxygenic, photoautotrophic microorganisms and higher plants to check whether similar improvements in photosynthesis could be achieved. Authors’ contributions Y.L. and J.Z. designed the research. F.Z., J.Z., H.M. and Y.P.Z. performed the research. Y.L. and J.Z. wrote the manuscript. All authors analyzed the data and discussed the results. Acknowledgments This work was supported by the Key Research Program of the Chinese Academy of Sciences (ZDRW-ZS-2016-3), the Strategic Priority Research Program of the Chinese Academy of Sciences (XDA07040405), and the National Natural Science Foundation of China (31470231). We thank Feng Guo, the technical expert of Dual-PAM 100, for assistance with the chlorophyll fluorescence and P700 signal analyses. 226 J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 Appendix A. Supplementary material Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.ymben.2016.08. 002. References Allen, J.F., 2003. Cyclic, pseudocyclic and noncyclic photophosphorylation: new links in the chain. Trends Plant Sci. 8, 15–19. Angermayr, S., van der Woude, A., Correddu, D., Vreugdenhil, A., Verrone, V., Hellingwerf, K., 2014. Exploring metabolic engineering design principles for the photosynthetic production of lactic acid by Synechocystis sp. PCC 6803. Biotechnol. Biofuels 7, 99. Angermayr, S.A., Hellingwerf, K.J., 2013. On the use of metabolic control analysis in the optimization of cyanobacterial biosolar cell factories. J. Phys. Chem. B 117, 11169–11175. Atsumi, S., Higashide, W., Liao, J.C., 2009. Direct photosynthetic recycling of carbon dioxide to isobutyraldehyde. Nat. Biotech. 27, 1177–1180. Bao, G., Wang, R., Zhu, Y., Dong, H., Mao, S., Zhang, Y., Chen, Z., Li, Y., Ma, Y., 2011. Complete genome sequence of Clostridium acetobutylicum DSM 1731, a solvent producing strain with multi-replicon genome architecture. J. Bacteriol. 193, 5007–5008. Blankenship, R.E., Chen, M., 2013. Spectral expansion and antenna reduction can enhance photosynthesis for energy production. Curr. Opin. Chem. Biol. 17, 457–461. Cai, Z., Liu, G., Zhang, J., Li, Y., 2014. Development of an activity-directed selection system enabled significant improvement of the carboxylation efficiency of Rubisco. Protein Cell 5, 552–562. Campbell, D., Hurry, V., Clarke, A.K., Gustafsson, P., Öquist, G., 1998. Chlorophyll fluorescence analysis of cyanobacterial photosynthesis and acclimation. Microbiol. Mol. Biol. Rev. 62, 667–683. Clark, S.W., Bennett, G.N., Rudolph, F.B., 1989. Isolation and characterization of mutants of Clostridium acetobutylicum ATCC 824 deficient in acetoacetylCoenzyme A: acetate/butyrate: coenzyme A-transferase (EC 2.8.3.9) and in other solvent pathway enzymes. Appl. Environ. Microbiol. 55, 970–976. Covshoff, S., Hibberd, J.M., 2012. Integrating C4 photosynthesis into C3 crops to increase yield potential. Curr. Opin. Biotechnol. 23, 209–214. Dai, Z., Dong, H., Zhu, Y., Zhang, Y., Li, Y., Ma, Y., 2012. Introducing a single secondary alcohol dehydrogenase into butanol-tolerant Clostridium acetobutylicum Rh8 switches ABE fermentation to high level IBE fermentation. Biotechnol. Biofuels 5, 44. Ducat, D.C., Avelar-Rivas, J.A., Way, J.C., Silver, P.A., 2012. Rerouting carbon flux to enhance photosynthetic productivity. Appl. Environ. Microbiol. 78, 2660–2668. Evans, J.R., 2013. Improving photosynthesis. Plant Physiol. 162, 1780–1793. Gan, F., Zhang, S., Rockwell, N.C., Martin, S.S., Lagarias, J.C., Bryant, D.A., 2014. Extensive remodeling of a cyanobacterial photosynthetic apparatus in far-red light. Science 345, 1312–1317. Gong, F., Liu, G., Zhai, X., Zhou, J., Cai, Z., Li, Y., 2015. Quantitative analysis of an engineered CO2-fixing Escherichia coli reveals great potential of heterotrophic CO2 fixation. Biotechnol. Biofuels 8, 86. Gonzalez-Esquer, C.R., Shubitowski, T.B., Kerfeld, C.A., 2015. Streamlined construction of the cyanobacterial CO2-fixing organelle via protein domain fusions for use in plant synthetic biology. Plant Cell 27, 2637–2644. Guerrero, F., Carbonell, V., Cossu, M., Correddu, D., Jones, P.R., 2012. Ethylene synthesis and regulated expression of recombinant protein in Synechocystis sp. PCC 6803. PLoS One 7, e50470. Hanai, T., Atsumi, S., Liao, J.C., 2007. Engineered synthetic pathway for isopropanol production in Escherichia coli. Appl. Environ. Microbiol. 73, 7814–7818. He, Q., Yang, H., Xu, L., Xia, L., Hu, C., 2015. Sufficient utilization of natural fluctuating light intensity is an effective approach of promoting lipid productivity in oleaginous microalgal cultivation outdoors. Bioresour. Technol. 180, 79–87. Hirokawa, Y., Suzuki, I., Hanai, T., 2015. Optimization of isopropanol production by engineered cyanobacteria with a synthetic metabolic pathway. J. Biosci. Bioeng. 119, 585–590. Ismaiel, A.A., Zhu, C.X., Colby, G.D., Chen, J.S., 1993. Purification and characterization of a primary-secondary alcohol dehydrogenase from two strains of Clostridium beijerinckii. J. Bacteriol. 175, 5097–5105. Kebeish, R., Niessen, M., Thiruveedhi, K., Bari, R., Hirsch, H.-J., Rosenkranz, R., Stabler, N., Schonfeld, B., Kreuzaler, F., Peterhansel, C., 2007. Chloroplastic photorespiratory bypass increases photosynthesis and biomass production in Arabidopsis thaliana. Nat. Biotech. 25, 593–599. Khan, M.S., 2007. Engineering photorespiration in chloroplasts: a novel strategy for increasing biomass production. Trends Biotechnol. 25, 437–440. Kliphuis, A.M.J., Klok, A.J., Martens, D.E., Lamers, P.P., Janssen, M., Wijffels, R.H., 2012. Metabolic modeling of Chlamydomonas reinhardtii: energy requirements for photoautotrophic growth and maintenance. J. Appl. Phycol. 24, 253–266. Klughammer, C., Schreiber, U., 2008. Saturation Pulse Method for assessment of Energy Conversion in PS I. PAM Appl. Notes 1, 11–14. Kozaki, A., Takeba, G., 1996. Photorespiration protects C3 plants from photooxidation. Nature 384, 557–560. Kramer, D.M., Evans, J.R., 2011. The importance of energy balance in improving photosynthetic productivity. Plant Physiol. 155, 70–78. Kusakabe, T., Tatsuke, T., Tsuruno, K., Hirokawa, Y., Atsumi, S., Liao, J., Hanai, T., 2013. Engineering a synthetic pathway in cyanobacteria for isopropanol production directly from carbon dioxide and light. Metab. Eng. 20, 101–108. Lan, E.I., Liao, J.C., 2012. ATP drives direct photosynthetic production of 1-butanol in cyanobacteria. Proc. Natl. Acad. Sci. USA 109, 6018–6023. Lin, M.T., Occhialini, A., Andralojc, P.J., Parry, M.A.J., Hanson, M.R., 2014. A faster Rubisco with potential to increase photosynthesis in crops. Nature 513, 547–550. Lindberg, P., Park, S., Melis, A., 2010. Engineering a platform for photosynthetic isoprene production in cyanobacteria, using Synechocystis as the model organism. Metab. Eng. 12, 70–79. Liu, C., Young, A.L., Starling-Windhof, A., Bracher, A., Saschenbrecker, S., Rao, B.V., Rao, K.V., Berninghausen, O., Mielke, T., Hartl, F.U., Beckmann, R., Hayer-Hartl, M., 2010. Coupled chaperone action in folding and assembly of hexadecameric Rubisco. Nature 463, 197–202. Liu, X., Sheng, J., Curtiss, R., 2011. Fatty acid production in genetically modified cyanobacteria. Proc. Natl. Acad. Sci. USA 108, 6899–6904. Luan, G., Qi, Y., Wang, M., Li, Z., Duan, Y., Tan, X., Lu, X., 2015. Combinatory strategy for characterizing and understanding the ethanol synthesis pathway in cyanobacteria cell factories. Biotechnol. Biofuels 8, 1–12. Marcus, Y., Altman-Gueta, H., Wolff, Y., Gurevitz, M., 2011. Rubisco mutagenesis provides new insight into limitations on photosynthesis and growth in Synechocystis PCC 6803. J. Exp. Bot. 62, 4173–4182. Melis, A., 2009. Solar energy conversion efficiencies in photosynthesis: minimizing the chlorophyll antennae to maximize efficiency. Plant Sci. 177, 272–280. Nielsen, A.Z., Ziersen, B., Jensen, K., Lassen, L.M., Olsen, C.E., Møller, B.L., Jensen, P.E., 2013. Redirecting photosynthetic reducing power toward bioactive natural product synthesis. ACS Synth. Biol. 2, 308–315. Noctor, G., Foyer, C.H., 1998. A re-evaluation of the ATP: NADPH budget during C3 photosynthesis: a contribution from nitrate assimilation and its associated respiratory activity? J. Exp. Bot. 49, 1895–1908. Nogales, J., Gudmundsson, S., Knight, E.M., Palsson, B.O., Thiele, I., 2012. Detailing the optimality of photosynthesis in cyanobacteria through systems biology analysis. Proc. Natl. Acad. Sci. USA 109, 2678–2683. Oliver, J.W., Atsumi, S., 2014. Metabolic design for cyanobacterial chemical synthesis. Photosynth. Res. 120, 249–261. Oliver, J.W.K., Machado, I.M.P., Yoneda, H., Atsumi, S., 2013. Cyanobacterial conversion of carbon dioxide to 2,3-butanediol. Proc. Natl. Acad. Sci. USA 110, 1249–1254. Ort, D.R., Zhu, X., Melis, A., 2011. Optimizing antenna size to maximize photosynthetic efficiency. Plant Physiol. 155, 79–85. Pfündel, E., Klughammer, C., Schreiber, U., 2008. Monitoring the effects of reduced PS II antenna size on quantum yields of photosystems I and II using the DualPAM-100 measuring system. PAM Appl. Notes 1, 21–24. Riccardi, C., Nicoletti, I., 2006. Analysis of apoptosis by propidium iodide staining and flow cytometry. Nat. Protoc. 1, 1458–1461. Savakis, P.E., Angermayr, S.A., Hellingwerf, K.J., 2013. Synthesis of 2,3-butanediol by Synechocystis sp. PCC 6803 via heterologous expression of a catabolic pathway from lactic acid and enterobacteria. Metab. Eng. 20, 121–130. Shen, J., Luo, W., 2011. Effects of monosulfuron on growth, photosynthesis, and nitrogenase activity of three nitrogen-fixing cyanobacteria. Arch. Environ. Contam. Toxicol. 60, 34–43. Shih, P.M., Zarzycki, J., Niyogi, K.K., Kerfeld, C.A., 2014. Introduction of a synthetic CO2-fixing photorespiratory bypass into a cyanobacterium. J. Biol. Chem. 289, 9493–9500. Singh, J., Pandey, P., James, D., Chandrasekhar, K., Achary, V.M.M., Kaul, T., Tripathy, B.C., Reddy, M.K., 2014. Enhancing C3 photosynthesis: an outlook on feasible interventions for crop improvement. Plant Biotechnol. J. . http://dx.doi.org/ 10.1111/pbi.12246 Singh, M., Yamamoto, Y., Satoh, K., Aro, E.M., Kanervo, E., 2005. Post-illuminationrelated loss of photochemical efficiency of Photosystem II and degradation of the D1 protein are temperature-dependent. J. Plant Physiol. 162, 1246–1253. Stephenson, P.G., Moore, C.M., Terry, M.J., Zubkov, M.V., Bibby, T.S., 2011. Improving photosynthesis for algal biofuels: toward a green revolution. Trends Biotechnol. 29, 615–623. Takahashi, H., Uchimiya, H., Hihara, Y., 2008. Difference in metabolite levels between photoautotrophic and photomixotrophic cultures of Synechocystis sp. PCC 6803 examined by capillary electrophoresis electrospray ionization mass spectrometry. J. Exp. Bot. 59, 3009–3018. Takeshima, Y., Takatsugu, N., Sugiura, M., Hagiwara, H., 1994. High-level expression of human superoxide dismutase in the cyanobacterium Anacystis nidulans 6301. Proc. Natl. Acad. Sci. USA 91, 9685–9689. Thamatrakoln, K., Bailleul, B., Brown, C.M., Gorbunov, M.Y., Kustka, A.B., Frada, M., Joliot, P.A., Falkowski, P.G., Bidle, K.D., 2013. Death-specific protein in a marine diatom regulates photosynthetic responses to iron and light availability. Proc. Natl. Acad. Sci. USA 110, 20123–20128. Tikkanen, M., Mekala, N.R., Aro, E.M., 2014. Photosystem II photoinhibition-repair cycle protects Photosystem I from irreversible damage. Biochim. Biophys. Acta 1837, 210–215. Vinyard, D.J., Gimpel, J., Ananyev, G.M., Mayfield, S.P., Dismukes, G.C., 2014. Engineered photosystem II reaction centers optimize photochemistry versus photoprotection at different solar intensities. J. Am. Chem. Soc. 136, 4048–4055. von Caemmerer, S., Evans, J.R., 2010. Enhancing C3 photosynthesis. Plant Physiol. 154, 589–592. Wientjes, E., Croce, R., 2012. PMS: Photosystem I electron donor or fluorescence J. Zhou et al. / Metabolic Engineering 38 (2016) 217–227 quencher. Photosynth. Res. 111, 185–191. Work, V.H., D’Adamo, S., Radakovits, R., Jinkerson, R.E., Posewitz, M.C., 2012. Improving photosynthesis and metabolic networks for the competitive production of phototroph-derived biofuels. Curr. Opin. Biotechnol. 23, 290–297. Xiong, W., Lee, T.C., Rommelfanger, S., Gjersing, E., Cano, M., Maness, P.C., Ghirardi, M., Yu, J., 2015a. Phosphoketolase pathway contributes to carbon metabolism in cyanobacteria. Nat. Plants 2, 15187. Xiong, W., Morgan, J.A., Ungerer, J., Wang, B., Maness, P., Yu, J., 2015b. The plasticity of cyanobacterial metabolism supports direct CO2 conversion to ethylene. Nat. Plants 1, 15053. Zhou, J., Zhang, F., Meng, H., Bao, G., Zhang, Y., Li, Y., 2014a. Development of a silicon carbide disruption method enables efficient extraction of proteins from 227 cyanobacterium Synechocystis sp. PCC 6803. Process Biochem. 49, 2199–2202. Zhou, J., Zhang, H., Meng, H., Zhang, Y., Li, Y., 2014b. Production of optically pure Dlactate from CO2 by blocking the PHB and acetate pathways and expressing Dlactate dehydrogenase in cyanobacterium Synechocystis sp. PCC 6803. Process Biochem. 49, 2071–2077. Zhou, J., Zhang, H., Meng, H., Zhu, Y., Bao, G., Zhang, Y., Li, Y., Ma, Y., 2014c. Discovery of a super-strong promoter enables efficient production of heterologous proteins in cyanobacteria. Sci. Rep. 4, 4500. Zhou, J., Zhang, H., Zhang, Y., Li, Y., Ma, Y., 2012. Designing and creating a modularized synthetic pathway in cyanobacterium Synechocystis enables production of acetone from carbon dioxide. Metab. Eng. 14, 394–400.