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• W2801974322 abstract "•ATP levels in a cyanobacterium vary according to growth phase and environment•A glycogen-deficient strain shows difference in ATP levels and photosynthesis•Glycogen synthesis/degradation is an important cellular energy buffer•Metabolite overflow is an alternative energy dissipation mechanism Understanding how living cells manage high-energy metabolites such as ATP and NADPH is essential for understanding energy transformations in the biosphere. Using light as the energy input, we find that energy charge (ratio of ATP over ADP+ATP) in the cyanobacterium Synechocystis sp. PCC 6803 varies in different growth stages, with a peak upon entry into the rapid growth phase, as well as a positive correlation with light intensity. In contrast, a mutant that can no longer synthesize the main carbon storage compound glycogen showed higher energy charge. The overflow of organic acids in this mutant under nitrogen depletion could also be triggered under high light in nitrogen-replete conditions, with an energy input level dependency. These findings suggest that energy charge in cyanobacteria is tightly linked to growth and carbon partition and that energy management is of key significance for their application as photosynthetic carbon dioxide-assimilating cell factories. Understanding how living cells manage high-energy metabolites such as ATP and NADPH is essential for understanding energy transformations in the biosphere. Using light as the energy input, we find that energy charge (ratio of ATP over ADP+ATP) in the cyanobacterium Synechocystis sp. PCC 6803 varies in different growth stages, with a peak upon entry into the rapid growth phase, as well as a positive correlation with light intensity. In contrast, a mutant that can no longer synthesize the main carbon storage compound glycogen showed higher energy charge. The overflow of organic acids in this mutant under nitrogen depletion could also be triggered under high light in nitrogen-replete conditions, with an energy input level dependency. These findings suggest that energy charge in cyanobacteria is tightly linked to growth and carbon partition and that energy management is of key significance for their application as photosynthetic carbon dioxide-assimilating cell factories. Understanding how living cells manage high-energy metabolites such as ATP and NADPH is essential for understanding energy transformations in the biosphere, including the primary energy-converting process, photosynthesis, and for developing carbon dioxide-assimilating cell factories. ATP and NADPH are generated in photosynthetic organisms by converting solar energy or releasing energy stored in chemical bonds and used to drive biosynthetic reactions and other life processes. Their generation and subsequent use need to be regulated under ever changing environmental conditions, such as variation of light and nutrient availability. Photosynthetic organisms have evolved myriad energy-balancing mechanisms at the level of the photosynthetic electron transport chain in order to regulate light sensing and harvesting through photosynthetic rearrangements and photoprotection capabilities (Eberhard et al., 2008Eberhard S. Finazzi G. Wollman F.-A. The dynamics of photosynthesis.Annu. Rev. Genet. 2008; 42: 463-515Crossref PubMed Scopus (475) Google Scholar, Kirilovsky and Kerfeld, 2013Kirilovsky D. Kerfeld C.A. The orange carotenoid protein: a blue-green light photoactive protein.Photochem. Photobiol. Sci. 2013; 12: 1135-1143Crossref PubMed Scopus (130) Google Scholar, Montgomery, 2014Montgomery B.L. The regulation of light sensing and light harvesting impacts the use of cyanobacteria as biotechnology platforms.Front. Bioeng. Biotechnol. 2014; 2: 22Crossref PubMed Scopus (17) Google Scholar) and to modulate energy production through cyclic electron flow (CEF) and alternative electron flow (AEF; also called pseudo-cyclic electron transfer) pathways (Allahverdiyeva et al., 2013Allahverdiyeva Y. Mustila H. Ermakova M. Bersanini L. Richaud P. Ajlani G. Battchikova N. Cournac L. Aro E.M. Flavodiiron proteins Flv1 and Flv3 enable cyanobacterial growth and photosynthesis under fluctuating light.Proc. Natl. Acad. Sci. U S A. 2013; 110: 4111-4116Crossref PubMed Scopus (218) Google Scholar) that complement the linear electron flow. Synthesis and use of carbon reserves, such as glycogen in cyanobacteria and starch and lipids in plants and eukaryotic algae, is also a common mechanism of adaptation to variations in light and nutrient availability (Zilliges, 2014Zilliges Y. Glycogen, a dynamic cellular sink and reservoir for carbon.in: Herrero A. The Cell Biology of Cyanobacteria. Caister Academic Press, 2014: 189-210Google Scholar). Yet a systematic understanding of the mechanisms of photosynthetic and metabolic adaptations to variations of energy supply, or how photosynthetic cells manage their energy, is still missing. In the model cyanobacterium Synechocystis sp. PCC 6803 (hereafter Synechocystis), glycogen synthesis occurs in the daytime according to light availability (Zilliges, 2014Zilliges Y. Glycogen, a dynamic cellular sink and reservoir for carbon.in: Herrero A. The Cell Biology of Cyanobacteria. Caister Academic Press, 2014: 189-210Google Scholar) and involves the key enzyme ADP-glucose pyrophosphorylase (AGPase) encoded by the glgC gene (slr1176). A strain devoid of this gene, ΔglgC, is unable to produce glycogen and exhibits striking phenotypes under nitrogen starvation, where biomass growth is halted and photosynthetically fixed carbon is directed toward the synthesis and excretion of organic acids, mainly pyruvate and 2-oxoglutarate (Carrieri et al., 2012Carrieri D. Paddock T. Maness P.-C. Seibert M. Yu J. Photo-catalytic conversion of carbon dioxide to organic acids by a recombinant cyanobacterium incapable of glycogen storage.Energy Environ. Sci. 2012; 5: 9457-9461Crossref Scopus (63) Google Scholar). This “overflow metabolism” has also been observed under mixotrophic growth conditions (Gründel et al., 2012Gründel M. Scheunemann R. Lockau W. Zilliges Y. Impaired glycogen synthesis causes metabolic overflow reactions and affects stress responses in the cyanobacterium Synechocystis sp. PCC 6803.Microbiology. 2012; 158: 3032-3043Crossref PubMed Scopus (182) Google Scholar) and in similar mutants in other cyanobacteria (Benson et al., 2016Benson P.J. Purcell-Meyerink D. Hocart C.H. Truong T.T. James G.O. Rourke L. Djordjevic M.A. Blackburn S.I. Price G.D. Factors altering pyruvate excretion in a glycogen storage mutant of the cyanobacterium, Synechococcus PCC7942.Front. Microbiol. 2016; 7: 475Crossref PubMed Scopus (14) Google Scholar, Davies et al., 2014Davies F.K. Work V.H. Beliaev A.S. Posewitz M.C. Engineering limonene and bisabolene production in wild type and a glycogen-deficient mutant of Synechococcus sp. PCC 7002.Front. Bioeng. Biotechnol. 2014; 2: 21Crossref PubMed Scopus (175) Google Scholar, Hickman et al., 2013Hickman J.W. Kotovic K.M. Miller C. Warrener P. Kaiser B. Jurista T. Budde M. Cross F. Roberts J.M. Carleton M. Glycogen synthesis is a required component of the nitrogen stress response in Synechococcus elongatus PCC 7942.Algal Res. 2013; 2: 98-106Crossref Scopus (51) Google Scholar), and it has been exploited to produce valuable chemicals from CO2 by enhancing flux into native (Qi et al., 2013Qi F. Yao L. Tan X. Lu X. Construction, characterization and application of molecular tools for metabolic engineering of Synechocystis sp.Biotechnol. Lett. 2013; 35: 1655-1661Crossref PubMed Scopus (21) Google Scholar) or heterologous pathways (Davies et al., 2014Davies F.K. Work V.H. Beliaev A.S. Posewitz M.C. Engineering limonene and bisabolene production in wild type and a glycogen-deficient mutant of Synechococcus sp. PCC 7002.Front. Bioeng. Biotechnol. 2014; 2: 21Crossref PubMed Scopus (175) Google Scholar, Li et al., 2014Li X. Shen C.R. Liao J.C. Isobutanol production as an alternative metabolic sink to rescue the growth deficiency of the glycogen mutant of Synechococcus elongatus PCC 7942.Photosynth. Res. 2014; 120: 301-310Crossref PubMed Scopus (84) Google Scholar, Work et al., 2015Work V.H. Melnicki M.R. Hill E. Davies F.K. Kucek L. Beliaev A. Posewitz M.C. Lauric acid production in a glycogen-less Synechococcus sp. PCC 7002 mutant.Front. Bioeng. Biotechnol. 2015; 3: 48Crossref PubMed Scopus (22) Google Scholar). The molecular mechanisms that direct the cells into this mode of metabolism were investigated, and they were hypothesized to occur at the metabolic level instead of transcriptional or translational level (Carrieri et al., 2017Carrieri D. Lombardi T. Paddock T. Cano M. Goodney G.A. Nag A. Old W. Maness P.-C. Seibert M. Ghirardi M. et al.Transcriptome and proteome analysis of nitrogen starvation responses in Synechocystis 6803 ΔglgC, a mutant incapable of glycogen storage.Algal Res. 2017; 21: 64-75Crossref Scopus (18) Google Scholar). The observation that the ratio of pyruvate and 2-oxoglutarate excreted during nitrogen starvation can be altered with changes in light intensity (Carrieri et al., 2015Carrieri D. Broadbent C. Carruth D. Paddock T. Ungerer J. Maness P.C. Ghirardi M. Yu J. Enhancing photo-catalytic production of organic acids in the cyanobacterium Synechocystis sp. PCC 6803 ΔglgC, a strain incapable of glycogen storage.Microb. Biotechnol. 2015; 8: 275-280Crossref PubMed Scopus (15) Google Scholar) prompted us to investigate high-energy metabolites through changing environmental conditions in the wild-type (WT) and ΔglgC Synechocystis strains. Intracellular ATP and ADP levels were estimated using a fast-performing luciferase-based assay. We observed a correlation between the energy charge (EC) and the growth phase in WT cultures: under non-saturating light at 50 μE·m−2·s−1, the EC increased gradually while cells adapted to new growth conditions (lag phase), peaked transiently, and steadily decreased as cells went through the early exponential phase (Figure 1A; see also Figure S1). Both volumetric ATP and ADP levels increased during the lag phase and subsequently decreased during the exponential phase (Figure 1B), with ATP levels decreasing faster than ADP levels, resulting in an overall decline in the EC. This rise and fall trend in the EC measured was consistently observed in numerous independent experiments, albeit with variations in absolute values and timing depending on the age of the inoculum (Figure S1). The increase in the magnitude and length of the initial EC buildup appeared to be sensitive to the age of the inoculum (Figures S1B and S1C). Duration of the initial EC buildup in WT cells was seemingly related to the length of lag phase (Figure S1), when cells prepare for exponential growth (Rolfe et al., 2012Rolfe M.D. Rice C.J. Lucchini S. Pin C. Thompson A. Cameron A.D.S. Alston M. Stringer M.F. Betts R.P. Baranyi J. et al.Lag phase is a distinct growth phase that prepares bacteria for exponential growth and involves transient metal accumulation.J. Bacteriol. 2012; 194: 686-701Crossref PubMed Scopus (364) Google Scholar), while entry into the exponential phase coincided with the peak of the EC level measured. Given the significant influence of the age of the inoculum on the EC values measured and the lag phase, we used similar standardized inoculum (3-day-old inoculum with similar recent inoculation history) to grow the cultures used in the remainder of this study. When exposed to extended periods of darkness, WT cultures’ turbidity as measured by A730nm reached a plateau (Figure 2A), and the EC increased in the first 6 hr of dark exposure up to a steady level. This rise in the EC upon shifting from light to dark is directly related to higher levels of ATP and to a lesser extent to changes in ADP levels (Figure 2B). When shifted back to light after 12 hr of dark exposure, the EC resumes its decrease, with ATP and ADP levels continuing to decline as before the transition to dark (Figure S2). In addition, when WT Synechocystis cells in mid-exponential phase (corresponding to A730nm ≈ 0.5–0.7 in a typical growth curve as presented in Figure 1A) were moved from moderate light intensity (50 μE·m−2·s−1) to either higher light intensity (HL; 250 μE·m−2·s−1) or lower light intensity (LL; 15 μE·m−2·s−1), the EC increased up to 50% and decreased by 10%, respectively (Figure 3A), indicating a dependency to the energy input in WT Synechocystis cells.Figure 2EC in the Dark in WT and ΔglgCShow full caption(A) Evolution of the EC (solid lines) through 24 hr of darkness after transitioning from light cultures of WT Synechocystis sp. PCC 6803 (blue diamond) and ΔglgC mutant (red circles) strains in exponential phase at 0 hr. Values are expressed as percentage of the EC in respective strains at time 0 hr. Growth was monitored by A730nm (dotted lines) under atmospheric air conditions supplemented with 5% CO2.(B) Relative amounts of ATP (WT, blue; ΔglgC, red) and ADP (WT, light blue; ΔglgC, light red) expressed as a percentage of the total adenylate pool levels ([ATP]+[ADP]) measured in WT (asterisk) and ΔglgC (double asterisk) respectively at time 0 hr.Vertical error bars represent SD of biological replicates (n = 3). See also Figure S2.View Large Image Figure ViewerDownload Hi-res image Download (PPT)Figure 3Overflow Metabolism Is an Energy-Balancing MechanismShow full caption(A) EC values measured 24 hr after switching cultures from 50 μE·m−2·s−1 illumination to either higher light (HL; 250 μE·m−2·s−1) or lower light (LL; 15 μE·m−2·s−1) intensities. EC is expressed as a percentage of the value measured in either strain under 50 μE·m−2·s−1 illumination at the same time point in cell growth (Figure S1). Average values are labeled on bars.(B) Extracellular pyruvate and 2-oxoglutarate in ΔglgC cultures after changing the light intensity from 50 μE·m−2·s−1 to higher light intensities of 150 μE·m−2·s−1 (left) or 250 μE·m−2·s−1 (right). Time 0 corresponds to 1-day-old cultures. An average growth curve from biological triplicates is represented by a solid red line.(C) Molar ratio of 2-oxoglutarate/pyruvate deduced from (B).Vertical error bars represent SDs of biological replicates (n = 3). See also Figures S3–S5.View Large Image Figure ViewerDownload Hi-res image Download (PPT) (A) Evolution of the EC (solid lines) through 24 hr of darkness after transitioning from light cultures of WT Synechocystis sp. PCC 6803 (blue diamond) and ΔglgC mutant (red circles) strains in exponential phase at 0 hr. Values are expressed as percentage of the EC in respective strains at time 0 hr. Growth was monitored by A730nm (dotted lines) under atmospheric air conditions supplemented with 5% CO2. (B) Relative amounts of ATP (WT, blue; ΔglgC, red) and ADP (WT, light blue; ΔglgC, light red) expressed as a percentage of the total adenylate pool levels ([ATP]+[ADP]) measured in WT (asterisk) and ΔglgC (double asterisk) respectively at time 0 hr. Vertical error bars represent SD of biological replicates (n = 3). See also Figure S2. (A) EC values measured 24 hr after switching cultures from 50 μE·m−2·s−1 illumination to either higher light (HL; 250 μE·m−2·s−1) or lower light (LL; 15 μE·m−2·s−1) intensities. EC is expressed as a percentage of the value measured in either strain under 50 μE·m−2·s−1 illumination at the same time point in cell growth (Figure S1). Average values are labeled on bars. (B) Extracellular pyruvate and 2-oxoglutarate in ΔglgC cultures after changing the light intensity from 50 μE·m−2·s−1 to higher light intensities of 150 μE·m−2·s−1 (left) or 250 μE·m−2·s−1 (right). Time 0 corresponds to 1-day-old cultures. An average growth curve from biological triplicates is represented by a solid red line. (C) Molar ratio of 2-oxoglutarate/pyruvate deduced from (B). Vertical error bars represent SDs of biological replicates (n = 3). See also Figures S3–S5. Contrary to the WT strain, the EC was found to be high and relatively steady in cultures growing under light in the glycogen-deficient strain ΔglgC. Under moderate light (50 μE·m−2·s−1), the EC values oscillated between 0.4 and 0.7 and stayed relatively stable during early exponential phase (Figures 1A and S1). Similar EC values were obtained for mid-exponential WT and ΔglgC cultures following liquid chromatography-mass spectrometry (LC-MS) detection on extracts obtained from cold methanol quenched cells (0.45 ± 0.05 for WT, 0.75 ± 0.05 for ΔglgC). This high and steady EC measured in cells impaired in glycogen synthesis is due mainly to higher levels of ATP than those in WT cells and, to a lesser extent, to a slower increase in ADP levels (Figure 1B) as the total adenylate pool measured is higher in the ΔglgC strain at similar cell density (A730nm). Interestingly, total adenylate pool measured kept increasing with cell density in ΔglgC, whereas it started decreasing in WT later in growth (Figure 1B), possibly indicating a decrease of the pool size per cell related to an increased storage of glycogen. The EC in the mutant decreased continuously in the dark over a period of 12 hr (Figure 2A; see also Figure S2). This energy depletion is connected to a decrease in cell viability as ΔglgC cultures left in darkness over extended periods of time fail to recover after shifting back to light (data not shown). Following a period of 12 hr in the dark, ATP levels could be rapidly restored when the cultures were exposed to light again (Figure S2). Under higher (250 μE·m−2·s−1) or lower (15 μE·m−2·s−1) light intensities, the EC remains rather similar in ΔglgC compared with normal conditions (50 μE·m−2·s−1), in contrast with the WT strain (Figure 3A). Management of adenylate levels is thus greatly affected in cells incapable of storing glycogen; ΔglgC strain seem to live at a high EC in the light, with less modulations of the EC compared with WT cells, and a fast depletion in the dark, when ATP generation by the photosynthetic chain stops and glycogen is absent for energy remobilization. Given that ΔglgC cells are incapable of storing excess carbon and energy in the form of glycogen, we then addressed the question of how cells manage an excess of energy input. ΔglgC cells grew slightly faster when light intensity was increased from 50 to 200 μE·m−2·s−1 (Figures S3 and S4) but bleached after 6 days under high light conditions, consistent with previous observations (Gründel et al., 2012Gründel M. Scheunemann R. Lockau W. Zilliges Y. Impaired glycogen synthesis causes metabolic overflow reactions and affects stress responses in the cyanobacterium Synechocystis sp. PCC 6803.Microbiology. 2012; 158: 3032-3043Crossref PubMed Scopus (182) Google Scholar). The EC seems to plateau under an illumination as low as 15 μE·m−2·s−1 (Figures 3A and S3). In the absence of glycogen to buffer additional incoming energy, other compensatory mechanisms might be functioning to maintain cellular energy homeostasis. As previously observed in those cells under N-deprivation, we suspected that “overflow metabolism” could be triggered under excess energy input. Indeed, analysis of extracellular medium revealed the excretion during cell growth of organic acids identified as 2-oxoglutarate and pyruvate (Figure S5) in the absence of nitrogen starvation (Figure 3B). The molar ratio of those two compounds was also shown to be affected by light intensity (Figure 3C), as previously reported for nitrogen-depleted conditions (Carrieri et al., 2015Carrieri D. Broadbent C. Carruth D. Paddock T. Ungerer J. Maness P.C. Ghirardi M. Yu J. Enhancing photo-catalytic production of organic acids in the cyanobacterium Synechocystis sp. PCC 6803 ΔglgC, a strain incapable of glycogen storage.Microb. Biotechnol. 2015; 8: 275-280Crossref PubMed Scopus (15) Google Scholar). In this study we explored the intracellular energy management in WT Synechocystis and ΔglgC strains by monitoring the EC through changing environmental conditions and identified energy levels as a possible driving force for growth and carbon partition. The EC was found to be variable in WT correlated to growth phases and peaks around the start of fast growth. This observation has potential in predicting and manipulating growth behavior of algal cultures. The EC also varied in WT under changing environmental conditions such as light intensity. In the dark, WT cells restored a high EC that correlated to increased levels of ATP (Figure 2), largely supported by catabolism of stored glycogen. In the absence of glycogen storage, cells live at a high EC even at low light intensities (Figure 3) and buffer increased energy input with excretion of organic carbon (Figure 3). Furthermore, the mutant displayed no obvious oxidative or high light stress response (Cassier-Chauvat and Chauvat, 2014Cassier-Chauvat C. Chauvat F. Responses to oxidative and heavy metal stresses in cyanobacteria: recent advances.Int. J. Mol. Sci. 2014; 16: 871-886Crossref PubMed Scopus (68) Google Scholar, Huang et al., 2002Huang L. McCluskey M.P. Ni H. LaRossa R.A. Global gene expression profiles of the cyanobacterium Synechocystis sp. strain PCC 6803 in response to irradiation with UV-B and white light.J. Bacteriol. 2002; 184: 6845-6858Crossref PubMed Scopus (164) Google Scholar), suggesting that at least under these conditions, a high EC does not lead to a general stress response. However, ΔglgC cultures do not seem to go through a lag phase, in accordance with high EC (Figure S1A). The relationship between EC and growth phases remains to be further studied. The observation of organic carbon excretion by the ΔglgC mutant under nitrogen-replete conditions is particularly interesting. Under our experimental conditions (moderate light at 50 μE·m−2·s−1), growth, photosynthetic capacity (Figure S6A), and O2 evolution rates (Carrieri et al., 2012Carrieri D. Paddock T. Maness P.-C. Seibert M. Yu J. Photo-catalytic conversion of carbon dioxide to organic acids by a recombinant cyanobacterium incapable of glycogen storage.Energy Environ. Sci. 2012; 5: 9457-9461Crossref Scopus (63) Google Scholar) of WT and ΔglgC are similar, despite major differences in their EC levels. There was no obvious impact on the EC in ΔglgC as the light intensity was increased from 15 to 50 μE·m−2·s−1 and up to 200 μE·m−2·s−1, with only a modest impact on growth (Figure S3). Presence of organic acids in the extracellular medium was seen sometimes in low amounts even under moderate light (50 μE·m−2·s−1); thus metabolite overflow is most likely a continuous process to balance excess energy when the glycogen buffer is not present. Along with metabolic rearrangement, photosynthetic adaptations are taking place in this high energetic context. With a similar photosynthetic capacity (similar quantum yield of PSII in the dark [Fv/Fm] or in the light [Fv′/Fm′] compared with the WT), PSII operating efficiency ([F′m − F]/F′m) is decreased (Figure S6A) under growth illumination (50 μE·m−2·s−1), as indicated by a higher steady-state fluorescence and a more reduced first electron acceptor of PSII QA (Figure S6B). CEF is also decreased, as shown by P700 oxidation-reduction kinetics assays in the presence of DCMU to block PSII-dependent linear electron flow (Figure S6C), and was also observed previously in similar growth conditions (3% CO2 bubbled in cultures, light intensity of 80 μE·m−2·s−1) (Holland et al., 2016Holland S.C. Artier J. Miller N.T. Cano M. Yu J. Ghirardi M.L. Burnap R.L. Impacts of genetically engineered alterations in carbon sink pathways on photosynthetic performance.Algal Res. 2016; 20: 87-99Crossref Scopus (14) Google Scholar). This is in accordance with a high EC observed with the mutant, where further stimulation of ATP synthesis is not needed. The ratio of [ATP]/[NADPH] is assumed to be critical for adjusting cells’ photosynthetic activity to meet metabolic demands (Kramer and Evans, 2011Kramer D.M. Evans J.R. The importance of energy balance in improving photosynthetic productivity.Plant Physiol. 2011; 155: 70-78Crossref PubMed Scopus (318) Google Scholar, Walker et al., 2014Walker B.J. Strand D.D. Kramer D.M. Cousins A.B. The response of cyclic electron flow around photosystem I to changes in photorespiration and nitrate assimilation.Plant Physiol. 2014; 165: 453-462Crossref PubMed Scopus (99) Google Scholar), so the relative production levels of ATP and NADPH need to be tightly regulated to match their use. Our attempts to measure the redox charge [NADPH]/([NADP+]+[NADPH]) using an enzymatic assay (luciferase-based) revealed no changes between WT and ΔglgC across three different light intensities (Figure S3). However, we had previously measured higher levels of NADPH in ΔglgC compared with WT (values were normalized to chlorophyll content) using fluorescence assays (Holland et al., 2016Holland S.C. Artier J. Miller N.T. Cano M. Yu J. Ghirardi M.L. Burnap R.L. Impacts of genetically engineered alterations in carbon sink pathways on photosynthetic performance.Algal Res. 2016; 20: 87-99Crossref Scopus (14) Google Scholar). This is interesting to put in light of a previous study in Saccharomyces cerevisiae that showed that a higher redox potential (primarily determined by the [NADH]/[NAD+] ratio in this organism) was linked to overflow metabolism (Vemuri et al., 2007Vemuri G.N. Eiteman M.A. McEwen J.E. Olsson L. Nielsen J. Increasing NADH oxidation reduces overflow metabolism in Saccharomyces cerevisiae.Proc. Natl. Acad. Sci. U S A. 2007; 104: 2402-2407Crossref PubMed Scopus (267) Google Scholar). Future efforts toward the determination of the redox charge and the [ATP]/[NADPH] ratio, perhaps with better methods, are needed for a more complete understanding of energy balancing in photosynthesis. Glycogen metabolism helps balance energy homeostasis in WT Synechocystis even under conditions in which glycogen does not over-accumulate (i.e., no nitrogen limitation) (Carrieri et al., 2012Carrieri D. Paddock T. Maness P.-C. Seibert M. Yu J. Photo-catalytic conversion of carbon dioxide to organic acids by a recombinant cyanobacterium incapable of glycogen storage.Energy Environ. Sci. 2012; 5: 9457-9461Crossref Scopus (63) Google Scholar), suggesting the possible existence of a dynamic cycle of synthesis and degradation whereby each time a glucose unit goes through the cycle, one ATP is consumed. The operation of such cycles has been shown in rumen bacteria (Hackmann and Firkins, 2015Hackmann T.J. Firkins J.L. Maximizing efficiency of rumen microbial protein production.Front. Microbiol. 2015; 6: 465Crossref PubMed Scopus (84) Google Scholar, Russell, 2007Russell J.B. The energy spilling reactions of bacteria and other organisms.J. Mol. Microbiol. Biotechnol. 2007; 13: 1-11Crossref PubMed Scopus (143) Google Scholar). A previous study in Synechocystis also identified a small but non-zero carbon flux to and from glycogen under autotrophic growth conditions (Young et al., 2011Young J.D. Shastri A.A. Stephanopoulos G. Morgan J.A. Mapping photoautotrophic metabolism with isotopically nonstationary (13)C flux analysis.Metab. Eng. 2011; 13: 656-665Crossref PubMed Scopus (264) Google Scholar). In the absence of glycogen, cells rely on decreased CEF and overflow metabolism to balance excess energy (Figure 4). Slower growth was observed in ΔglgC under higher light intensities. Interestingly, introduction of a heterologous pathway for isobutanol production in a Synechococcus ΔglgC mutant partially rescued the retardation growth phenotype observed (Li et al., 2014Li X. Shen C.R. Liao J.C. Isobutanol production as an alternative metabolic sink to rescue the growth deficiency of the glycogen mutant of Synechococcus elongatus PCC 7942.Photosynth. Res. 2014; 120: 301-310Crossref PubMed Scopus (84) Google Scholar), highlighting the delicate balance between photosynthate production and use, which is precarious in ΔglgC. Glycogen synthesis in cyanobacteria was previously suggested to help prevent reduction of photosynthetic activity from feedback regulation caused by glucose accumulation (Miao et al., 2003Miao X. Wu Q. Wu G. Zhao N. Changes in photosynthesis and pigmentation in an agp deletion mutant of the cyanobacterium Synechocystis sp.Biotechnol. Lett. 2003; 25: 391-396Crossref PubMed Scopus (24) Google Scholar). Here we propose that it specifically enables efficient ATP consumption, thus mitigating excessive reduction of electron carriers and subsequent congestion in the photosynthetic electron transport chain. ATP consumption is an important metabolic driving force: introduction of additional ATP-consuming steps in a non-native pathway in Synechococcus elongatus PCC 7942 allowed direct photosynthetic production of 1-butanol (Lan and Liao, 2012Lan E.I. Liao J.C. ATP drives direct photosynthetic production of 1-butanol in cyanobacteria.Proc. Natl. Acad. Sci. U S A. 2012; 109: 6018-6023Crossref PubMed Scopus (291) Google Scholar). Thus energy management in cyanobacteria is of key significance for their application as photosynthetic carbon dioxide-assimilating cell factories. Overflow metabolism is commonly defined as a metabolic phenomenon induced when the rate of glycolysis exceeds a critical value in aerobic conditions and that generally results in the secretion in the extracellular medium of end products (e.g., ethanol in yeast, acetate in Escherichia coli, and lactate in mammalian cells). From an energy generation point of view, this appears to be a wasteful strategy in which cells use fermentation instead of the more efficient (higher ATP-yielding) oxidative phosphorylation (respiration) to generate energy, despite the availability of O2. Termed the Warburg effect in the context of cancer growth (Vander Heiden et al., 2009Vander Heiden M.G. Cantley L.C. Thompson C.B. Understanding the Warburg effect: the metabolic requirements of cell proliferation.Science. 2009; 324: 1029-1033Crossref PubMed Scopus (10142) Google Scholar) and the Crabtree effect in yeast (De Deken, 1966De Deken R.H. The Crabtree effect: a regulatory system in yeast.J. Gen. Microbiol. 1966; 44: 149-156Crossref PubMed Scopus (566) Google Scholar), this phenomenon occurs ubiquitously for fast-growing cells. In E. coli it has been described as an efficient allocation of energy, as the proteome cost of energy biogenesis by respiration exceeds by 2-fold that by fermentation (Basan et al., 2015Basan M. Hui S. Okano H. Zhang Z. Shen Y. Williamson J.R. Hwa T. Overflow metabolism in Escherichia coli results from efficient proteome allocation.Nature. 2015; 528: 99-104Crossref PubMed Scopus (350) Google Scholar). The molecular mechanism that regulates overflow metabolism in cyanobacteria has been proposed to occur at the metabolic level instead of at the transcriptional or translational levels (Carrieri et al., 2017Carrieri D. Lombardi T. Paddock T. Cano M. Goodney G.A. Nag A. Old W. Maness P.-C. Seibert M. Ghirardi M. et al.Transcriptome and proteome analysis of nitrogen starvation responses in Synechocystis 6803 ΔglgC, a mutant incapable of glycogen storage.Algal Res. 2017; 21: 64-75Crossref Scopus (18) Google Scholar). This work suggests that excess photosynthate is the driving force for metabolite overflow and thus provides more in-depth understandings about the relationship between carbon sink and energy management. More detailed analysis of the relationship between photosynthate supply and carbon partition will be needed to determine the regulatory nodes and mode of action that govern carbon flux in cyanobacterial photosynthesis. Wild-type Synechocystis sp. PCC 6803 and ΔglgC strains (Carrieri et al., 2012Carrieri D. Paddock T. Maness P.-C. Seibert M. Yu J. Photo-catalytic conversion of carbon dioxide to organic acids by a recombinant cyanobacterium incapable of glycogen storage.Energy Environ. Sci. 2012; 5: 9457-9461Crossref Scopus (63) Google Scholar) were grown photoautotrophically in modified BG11 medium supplemented with 100 mM NaHCO3, 20 mM TES (pH 8), in 250 or 50 mL flasks. ΔglgC inoculum was maintained in the presence of 25 μg·mL−1 gentamycin, while the antibiotic was not used in cultures grown to perform physiological assays. Nitrogen starvation was not used in this work. Flasks were maintained on an orbital shaker (120 rpm), placed inside a growth chamber at 30°C with 5% CO2 under continuous moderate light (50 μE·m−2·s−1) provided by cool white fluorescent lamps. Cell growth was monitored by absorbance assays at 730 nm (A730nm) with a Beckman Coulter DU 800 spectrophotometer; cell counting was performed using a Cellometer Auto X4 automated cell counter (Nexcelom Biosciences). Cultures were systematically inoculated at an initial A730nm of 0.09–0.10. Chlorophyll was assayed following extractions with 100% methanol as described previously (Ritchie, 2006Ritchie R.J. Consistent sets of spectrophotometric chlorophyll equations for acetone, methanol and ethanol solvents.Photosynth. Res. 2006; 89: 27-41Crossref PubMed Scopus (737) Google Scholar). Intracellular ATP and ADP concentration were determined using a luciferin/luciferase bioluminescence assay kit (MAK135; Sigma-Aldrich). Assays were performed in 96-well white microplates at room temperature under conditions similar to those used for cell growth (same light intensity or dark throughout the assay) (Figure S7A). Ten microliters of cells were lysed in the buffer containing the luciferase supplied in the kit. Complete permeabilization of cells was achieved after 1 min, as confirmed by SYTOX Green staining assays (Figure S7B), allowing intracellular metabolites to diffuse out of cells. The initial bioluminescence readings, which correspond to ATP levels, were performed immediately after permeabilization with a Tecan Infinite M200 PRO luminometer (Tecan Trading). A mix containing pyruvate kinase was then rapidly added to convert ADP to ATP, and luminescence was once again measured, yielding the total size of the (ATP+ADP) pool. Minimization of time delays along with preservation of the environmental conditions (light intensity/darkness) throughout the experiment was critical to carrying out reliable and consistent EC assays. Intracellular NADP and NADPH levels were determined using a bioluminescence assay kit (G9081; Promega) following the manufacturer’s instructions. Sample removal and metabolite extraction performed for LC-MS/MS analysis were carried out as previously described (Young et al., 2011Young J.D. Shastri A.A. Stephanopoulos G. Morgan J.A. Mapping photoautotrophic metabolism with isotopically nonstationary (13)C flux analysis.Metab. Eng. 2011; 13: 656-665Crossref PubMed Scopus (264) Google Scholar). One milliliter of culture was harvested and centrifuged at 13,000 × g. Supernatants were filtered through 0.20 μm filters and analyzed by high-performance liquid chromatography (HPLC) as previously described (Carrieri et al., 2012Carrieri D. Paddock T. Maness P.-C. Seibert M. Yu J. Photo-catalytic conversion of carbon dioxide to organic acids by a recombinant cyanobacterium incapable of glycogen storage.Energy Environ. Sci. 2012; 5: 9457-9461Crossref Scopus (63) Google Scholar). Chlorophyll fluorescence and P700 oxidation/reduction kinetic assays were performed as previously described (Holland et al., 2016Holland S.C. Artier J. Miller N.T. Cano M. Yu J. Ghirardi M.L. Burnap R.L. Impacts of genetically engineered alterations in carbon sink pathways on photosynthetic performance.Algal Res. 2016; 20: 87-99Crossref Scopus (14) Google Scholar). Project planning, sample preparation, physiological characterization, data analysis, and preparation of the manuscript were supported by the U.S. Department of Energy (DOE), Office of Science Basic Energy Sciences (M.C., M.G., and J.Y. under contract DE-AC36-08-GO28308 and S.C.H., J.A., and R.B under grant DE-FG02-08ER15968). Sample preparation and metabolomics analysis was also supported by DOE BER award DE-SC0008628 (J.A.M.). We wish to thank Peter Ciesielski for assistance with microscopy and Mike Seibert and Longyun Guo for comments on the manuscript. The U.S. government retains and the publisher, by accepting the article for publication, acknowledges that the U.S. government retains a nonexclusive, paid up, irrevocable, worldwide license to publish or reproduce the published form of this work, or allow others to do so, for U.S. government purposes. M.C., M.G., and J.Y. planned the project. M.C., S.C.H., and J.A. performed the experiments. M.C., S.C.H., J.A., R.L.B., J.A.M., M.G., and J.Y. interpreted data and drafted the manuscript. All authors edited and approved the manuscript. The authors declare no competing interests. Download .pdf (.93 MB) Help with pdf files Document S1. Figures S1–S7" @default.
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• W2801974322 title "Glycogen Synthesis and Metabolite Overflow Contribute to Energy Balancing in Cyanobacteria" @default.
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