Glycogen synthesis prevents metabolic imbalance and disruption of photosynthetic electron transport from photosystem II during transition to photomixotrophy in Synechocystis sp. PCC 6803

Glycogen synthesis prevents metabolic imbalance and disruption of photosynthetic electron transport from photosystem II during transition to photomixotrophy in Synechocystis sp. PCC 6803 Pablo Ortega-Martınez 1,2 *, Lauri Nikkanen 3 *, Laura T. Wey 3 , Francisco J. Florencio 1,2 , Yagut Allahverdiyeva 3 and Sandra Dıaz-Troya 1,2 1 Instituto de Bioquımica Vegetal y Fotosıntesis, Universidad de Sevilla, Consejo Superior de Investigaciones Cientıficas, Americo Vespucio 49, Sevilla, 41092, Spain; 2 Departamento de Bioquımica Vegetal y Biologıa Molecular, Facultad de Biologıa, Universidad de Sevilla, Profesor Garcıa Gonzalez s/n, Sevilla, 41012, Spain; 3 Molecular Plant Biology, Department of Life Technologies, University of Turku, Turku, FI-20014, Finland Authors for correspondence: Sandra D ıaz-Troya Email: [email protected] Yagut Allahverdiyeva Email: [email protected] Received: 13 February 2024 Accepted: 17 April 2024 New Phytologist (2024) 243: 162–179 doi: 10.1111/nph.19793 Key words: cyanobacteria, glucose, glycogen, metabolism, mixotrophy, photomixotrophy, photosynthesis. Summary Some cyanobacteria can grow photoautotrophically or photomixotrophically by using simultaneously CO 2 and glucose. The switch between these trophic modes and the role of glycogen, their main carbon storage macromolecule, was investigated. We analysed the effect of glucose addition on the physiology, metabolic and photosynthetic state of Synechocystis sp. PCC 6803 and mutants lacking phosphoglucomutase and ADP-glucose pyrophosphorylase, with limitations in glycogen synthesis. Glycogen acted as a metabolic buffer: glucose addition increased growth and glycogen reserves in the wild-type (WT), but arrested growth in the glycogen synthesis mutants. Already 30 min after glucose addition, metabolites from the Calvin–Benson–Bassham cycle and the oxidative pentose phosphate shunt increased threefold more in the glycogen synthesis mutants than the WT. These alterations substantially affected the photosynthetic performance of the glycogen synthesis mutants, as O 2 evolution and CO 2 uptake were both impaired. We conclude that glycogen synthesis is essential during transitions to photomixotrophy to avoid metabolic imbalance that induces inhibition of electron transfer from PSII and subsequently accumulation of reactive oxygen species, loss of PSII core proteins, and cell death. Our study lays foundations for optimising photomixotrophy-based biotechnologies through understanding the coordination of the crosstalk between photosynthetic electron transport and metabolism. Introduction The biosphere and trophic chains are sustained by photosynthetic organisms, including cyanobacteria, algae and plants. Photoautotrophic growth involves the fixation of CO 2 into metabolism through the Calvin–Benson–Bassham (CBB) cycle, fuelled by photosynthetic processes. Cyanobacteria are unique as prokaryotic oxygenic photosynthetic microorganisms and play a crucial role in the global carbon cycle, contributing c. 25% of annual global productivity (Field et al., 1998). Additionally, cyanobacteria are increasingly used as green cell factories by redirecting their metabolism towards the biosynthesis of desired compounds. Besides photoautotrophic growth, some cyanobacteria such as Synechocystis sp. PCC 6803 (hereafter Synechocystis) have a plastic metabolic network that enables growth under a variety of trophic conditions, including photomixotrophy (Luan et al., 2019; Muth-Pawlak et al., 2022). Under photomixotrophy, cells can supplement photosynthesis by using organic carbon compounds from the environment, such as glucose, sucrose, glycerol or acetate (Matson & Atsumi, 2018). The dual carbon source enhances growth, offering attractive possibilities for biotechnology (Wan et al., 2015; Kanno et al., 2017; Luan et al., 2019). Despite their ecological and biotechnological significance (Mu~noz-Marın et al., 2024), the physiological mechanisms underlying the transition from photoautotrophy to photomixotrophy in cyanobacteria and the role of the main carbon storage macromolecule glycogen in that process remain poorly understood. The CBB cycle is an energy-demanding process that requires both ATP and NADPH produced in the photosynthetic electron transport chain (PETC). Briefly, the photolysis of water by photosystem II (PSII) releases electrons as well as molecular oxygen (O 2 ) as a by-product (Fig. 1). In linear electron transfer (LET) these electrons ultimately reduce ferredoxin (Fd), which, in turn, is used to generate NADPH. The PETC generates a *These authors contributed equally to this work. 162 New Phytologist (2024) 243: 162–179 Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation www.newphytologist.com This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited. Research proton motive force (pmf ) over the thylakoid membrane through different mechanisms, such as water oxidation and proton translocation associated with the plastoquinone (PQ) reductive cycle. Other complexes, such as NAD(P)H dehydrogenase-like (NDH- 1) complex and the respiratory terminal oxidase COX, also contribute to proton gradient formation. The pmf is harnessed by ATP synthase to produce ATP. The electrons from Fd can also be diverted back to the PETC in cyclic electron transfer (CET), facilitating pmf generation. Cells can adjust the rates of LET and CET based on their energetic requirements for the ATP/ NADPH ratio under variable environmental conditions (Nikkanen et al., 2021; Theune et al., 2021; Miller et al., 2022). In contrast to photosynthetic eukaryotes, which perform catabolic and anabolic reactions in separate organelles, cyanobacteria partly Fig. 1 Diagram of the main metabolic pathways and their interconnections to photosynthesis in Synechocystis sp. PCC 6803. Each route is colour-coded for clarity: the glycogen metabolic pathway (grey), the oxidative pentose phosphate (OPP) shunt (blue), the Calvin–Benson–Bassham (CBB) cycle (green), the photosynthetic electron transport chain (PETC, yellow), and the downstream metabolism of 3-phosphoglycerate (3PG, pink). The Entner-Doudoroff pathway, tricarboxylic acid cycle and flavodiiron proteins are not depicted. Glucose assimilation involves its phosphorylation to glucose-6P by hexokinase (HK), followed by conversion to either 6P-gluconate (6PG) through the action of glucose-6P dehydrogenase in the OPP shunt, or fructose-6P (F6P) via phosphoglucose isomerase (PGI) through the PGI shunt. Another route to G6P is its interconversion to glucose-1P by the phosphoglucomutases (PGM and PMM/PGM) and its incorporation to glycogen by the action of ADP-glucose pyrophosphorylase (AGP) and glycogen synthases (GlgAs) with ATP consumption. In the CBB cycle, CO 2 is incorporated into the intermediate metabolite ribulose 1,5-bisphosphate (RuBP) by RuBisCO releasing two molecules of 3-phosphoglycerate (3PG). Some 3PG is used for biosynthetic processes, while the majority is redirected to the CBB cycle to replenish the RuBP pools and ensure efficient CO 2 fixation. In this cycle, 3PG undergoes phosphorylation by 3-phosphoglycerate kinase and subsequent reduction by glyceraldehyde-3-phosphate dehydrogenase generating glyceraldehyde-3P (GAP), with the consumption of ATP and NADPH, respectively. The PETC consists of protein complexes that transport electrons through linear (LET, black arrows) or cyclic (CET, blue arrows) pathways: Within photosystem II (PSII) and photosystem I (PSI) the electrons are excited by photons harnessed by phycobilisome (PBS) antennae and Chls. Respiration (RETC, blue arrows) also takes place in thylakoids. The green and orange crosses depicted on upper panel of the PGM and AGP reactions arrows represent the knock-out mutations used to study glycogen metabolism. c 6 , cytochrome c 6 ; COX, cytochrome coxidase; cyd, cytochrome bd-quinol oxidase; Cyt b 6 f, cytochrome b 6 f; DCBQ, 2,6-Dichloro-1,4-benzoquinone; DCMU, 3-(3,4-dichlorophenyl)-1,1-dimethylurea; DHAP, Dihydroxyacetone-P; Fd, Ferredoxin; FNR, Ferredoxin-NADPH reductase; Fum, Fumarate; NDH-1, NADPH dehydrogenase complexes; OEC, oxygen-evolving complex; PC, plastocyanin; PFOR, Pyruvate-Ferredoxin oxidoreductase; PQ/PQH2, plastoquinone pool; PSI, photosystem I; PSII, photosystem II; Ru5P, Ribulose-5P; RuBisCO, Ribulose-1,5-P Carboxylase Oxidase; SDH, succinate dehydrogenase; Suc, Succinate. Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation New Phytologist (2024) 243: 162–179 www.newphytologist.com New Phytologist Research 163 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License share the PETC for respiration (Fig. 1, Mullineaux, 2014; Lea-Smith et al., 2016). Exogenous organic compounds such as glucose are incorporated via membrane transporters, imposing reorganisations of several metabolic processes, the details of which remain unclear (Kahlon et al., 2006; Trautmann et al., 2012; Burnap et al., 2015;Zavrel et al., 2017). First, sharing PETC components, such as the PQ pool, cytochrome (Cyt) b 6 fcomplex, plastocyanin (PC) or Cyt c 6 between photosynthesis and respiration requires adjustments to fulfil cellular demands while preventing overreduction (Mullineaux, 2014). Second, glucose supplementation rearranges metabolism to optimise resource utilisation. For example, fluxomic studies have revealed that carbon metabolic flux is largely redirected towards increasing ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO) activity, thus boosting CO 2 fixation in the CBB cycle (Schulze et al., 2022). The remaining carbon is predominantly channelled to glycogen, which serves as a carbon and energy buffer and plays a crucial role in the plasticity of cell metabolism (Cano et al., 2018;Luanet al., 2019;Makowkaet al., 2020; Ortega-Martınez et al., 2023). Glycogen is synthesised from ADP-glucose in a pathway widely conserved in bacteria (Fig. 1). ADP-glucose synthesis requires G1P, which is produced in a bidirectional reaction using glucose-6P (G6P) by the phosphohexomutase enzymes, such as phosphoglucomutase (PGM), and to a lesser extent by phosphomannomutase/phosphoglucomutase (PMM/PGM). G1P is then coupled with ATP to produce ADP-glucose by ADP-glucose pyrophosphorylase (AGP), which is used by glycogen synthase(s) (GlgA) to elongate the glycogen chains (Ball &Morell,2003;Cifuenteet al., 2019). Strains deficient in either AGP or PGM, which contain null or reduced glycogen levels, respectively, are viable under permissive conditions (Gr€undel et al., 2012;Doelloet al., 2022; Ortega-Martınez et al., 2023). However, under stress conditions, such as high light intensity, nitrogen deprivation or extended darkness, they exhibit impaired growth, metabolic overflow, and cell death (Carrieri et al., 2012; Cano et al., 2018; Ortega-Martınez et al., 2023). Furthermore, the sensitivity of glycogen-less Synechocystis mutants to photomixotrophic conditions (Gr€undel et al., 2012) suggests that glycogen is crucial in the transition between trophic modes. In this study we investigated the photosynthetic and metabolic response of Synechocystis during transitions from photoautotrophy to photomixotrophy. We aimed to understand how the ability to store excess carbon in glycogen influences these responses. Towards these aims, we analysed glycogen synthesis mutant strains ΔPGM and ΔAGP using metabolomics and a suite of biophysical and biochemical techniques to study photosynthesis. We found that in glycogen synthesis mutants, the presence of an organic carbon source caused accumulation of most metabolites in the intertwined pathways of central carbon metabolism, as well as a swift decline in photosynthetic electron transport due to restriction of electron transfer from PSII. Our study provides valuable physiological insights into the metabolic transition from photoautotrophic to photomixotrophic conditions, shedding light on the mechanisms of coordinating the intertwined bioenergetics of photosynthesis and metabolism. Materials and Methods Strains and culture conditions Glucose-tolerant WT strain of Synechocystis sp. PCC 6803 and mutants in phosphoglucomutase (PGM) (Δpgm::CK1, Ortega-Martınez et al., 2023) and ADP-glucose pyrophosphorylase (AGP) (Δagp::C.C1, Dıaz-Troya et al., 2014) were cultivated in either BG11 buffered to pH 7.5 with 20 mM HEPES-KOH (BG11) or BG11 with 12 mM bicarbonate (BG11C) (Rippka et al., 1979). No differences were observed in growth and oxygen evolution rates between media. Cell growth was monitored by OD 750 nm . Precultures in mid-logarithmic phase grown at 30°C under constant light at 40 5lmol photons m 2 s 1 , and with continuous shaking at 100 rpm under atmospheric CO 2 were adjusted to 0.8 OD 750 nm 24 h before the experiments. On the day of the experiment, cultures were centrifuged, resuspended in fresh medium at 1 OD 750 nm or at the Chl concentration described for each experiment. WT, ΔPGM and ΔAGP cultures adjusted to 1 OD 750 nm had Chl contents of 4.68 0.46, 4.95 0.38 and 4.79 0.48 (mean SD) lg Chl ml 1 , respectively. Cultures were then acclimated to the same growth conditions described above for 2 h before adding 2 mM glucose (Acros Organics, Geel, Belgium). Glycogen content and glucose consumption determination Glycogen was purified and digested to glucose as described in Ortega-Martınez et al.(2023). Glucose derived from purified glycogen and glucose content in the media were measured using a colorimetric assay based on the oxidation of 200 mg ml 1 of O-Dianisidine (Sigma) by 5 U ml 1 of peroxidase (from horseradish, Sigma) using the H 2 O 2 released from glucose by 25 U ml 1 glucose oxidase (from Aspergillus niger, Sigma) for 30 min at 37°C. The reaction was quenched by adding H 2 SO 4 to a final concentration of 4.8 N, and absorbance was measured at 540 nm using a Varioskan multiplate reader (Thermo Fisher Scientific, Erlangen, Germany). Standard curves of known amounts of amyloglucosidase-digested glycogen or glucose were employed for quantification. Protein extraction and immunoblotting Ten milliliter cultures were harvested and frozen at 20°C. Pellets were resuspended in 50 mM Tris–HCl (pH 8), 25 mM NaCl, and 1 mM phenylmethylsulphonyl fluoride (PMFS) with glass beads. Cells were lysed by mechanical disruption at 6 m s 1 for 30 s in a FastPrep-24 5G (MP Biomedicals, Santa Ana, CA, USA). Tubes were centrifuged at 4°C for 5 min at 1500 gand supernatants collected. Total protein content was determined a Lowry assay (Markwell et al., 1978). Fifteen micrograms of protein was resolved by SDS–PAGE (10% acrylamide/bis-acrylamide) containing 6 M urea and New Phytologist (2024) 243: 162–179 www.newphytologist.com Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation Research New Phytologist 164 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License transferred to a polyvinylidene difluoride membrane (Immobilon-P; Millipore). Blocking, probing and detection was performed as in Ortega-Martınez et al.(2023) using primary antibodies against D1 (1 : 10 000, AS111786; Agrisera, V€ann€as, Sweden) CP-47 (1 : 10 000, AS04038; Agrisera), PsaB (1 : 10 000, AS10695; Agrisera), ATPb(1 : 10 000, AS05085; Agrisera) or GroEL (1 : 75 000, G6532; Sigma-Aldrich) diluted in blocking solution. Reactive oxygen species measurement Reactive oxygen species levels were measured as outlined in Lee et al.(2018) with the following modifications. Samples were centrifuged at 10 000 gfor 5 min. Pellets were resuspended in 0.8 ml of 19phosphate-buffered saline (PBS) containing 50 lM of the fluorescent probe 20,70-dichlorofluorescein diacetate (DCFH-DA, D6883 Sigma-Aldrich). Samples were incubated for 1 h in darkness at 30°C with agitation. To eliminate free DCFH-DA, the samples were centrifuged at 10 000 gfor 5 min and washed once with PBS. Cells were resuspended in 200 ll PBS and transferred to a fluorescent 96-well microplate. Fluorescence of dichlorofluorescein (DCF), the oxidised form of DCFH, was measured at 525 nm after excitation at 488 nm in a Varioskan multiplate reader. Glutathione pools quantification Glutathione extraction and quantification was performed as outlined in Mallen-Ponce et al.(2022) using 10 ml culture harvested (5000 g, 5 min, 4°C) at the indicated times. Standards with known glutathione concentrations were used for quantification. Targeted metabolomics (LC/MS) Extraction and analysis of intracellular metabolites was performed using 2 OD 750 nm culture as described in Ortega-Martınez et al.(2023). For quantification of the total amount of metabolites, known concentrations of standards were used. Metabolites with concentration values outside the standard curve or not available for quantification are reported as fold change in relation to the WT value at the initial time point. Gas exchange measurements Real-time gas fluxes were measured from intact cells at a concentration of 10 lg Chl ml 1 under 50 lmol photons m 2 s 1 of white actinic light using membrane inlet mass spectrometry (MIMS) as described in Solymosi et al.(2020). Cultures were first dark-adapted for 20 min, purged with N 2 , and supplemented with 1.5 mM NaHCO 3 and 18 O 2 at an equivalent concentration to 16 O 2 , enabling the differentiation of O 2 uptake from O 2 evolution. Gas exchange rates were calculated according to (Beckmann et al., 2009). After 15 min of recording 0.5 mM 2,6-Dichloro-1,4-benzoquinone (DCBQ) in dimethyl sulfoxide (DMSO) was introduced into the sample chamber and measurements continued for 10 min. Spectrometry Chlorophyll fluorometry Induction curves of Chl fluorescence from intact cells were measured using a DUAL-PAM-100 fluorometer as outlined in Ortega-Martınez et al.(2023). Cells at 5lg Chl ml 1 and dark-adapted for 10 min were exposed to red actinic light at 40 lmol photons m 2 s 1 and multiple saturating pulses, followed by a postillumination recovery. Reoxidation kinetics of Q A  The relaxation kinetics of single-turnover flash-induced Chl fluorescence were monitored using an FL 3500 fluorometer (PSI Instruments, Drasov, Czech Republic). Cells were adjusted to 7.5 lgChlml 1 and dark-adapted for 10 min before measurements. Measurements were performedwithorwithout10lM 3-(3,4-dichlorophenyl)-1, 1-dimethylurea (DCMU). Curves were baselined to the fluorescencebeforetheflashandnormalised to the maximum fluorescence after the flash. The fluorescence decay traces were fitted to a three-component exponential decay function (y=A1 (x/t1) + A2 (x/t2) +A3 (x/t3) +y 0 )usingORIGINPRO 2023b. PC, P700 and Fd redox changes Redox changes of PC, P700 and Fd in cells adjusted to 20 lgChlml 1 were obtained by NIR absorption difference spectrometry using a DUAL-KLAS-NIR spectrophotometer (Walz) and a NIRMAX script modified for cyanobacteria (3 s AL with a MT at 200 ms to obtain maximal values of Fd reduction and PC and P700 oxidation) (Schreiber & Klughammer, 2016). The model spectra for deconvolution of Synechocystis PC, P700, and Fd signals were measured as in Nikkanen et al.(2020). LET and CET quantification The rates of LET and CET through PSI were quantified using analysis of dark interval relaxation kinetics (DIRK) in the absence or presence of 10 lMDCMU as described in Theune et al.(2021). Cells were preilluminated for 2 min under 50 lmol photons m 2 s 1 and subjected to 100 dark intervals of 20 ms with 1 s illumination between each repetition. Traces from all 100 dark intervals were then averaged. Cytochrome b 6 fredox kinetics Cytochrome fand bheme redox kinetics were measured as described in Solymosi et al.(2020) using a JTS-10 spectrophotometer (BioLogic). NAD(P)H fluorescence Light-induced NAD(P)H redox kinetics were measured from 7.5 lg Chl ml 1 samples at 30°C, monitoring fluorescence changes between 420 and 580 nm after excitation at 365 nm using the NADPH/9-AA module of a DUAL-PAM (Walz, Germany) (Nikkanen et al., 2020). Results Glucose causes growth arrest, oxidative stress, and decrease in PSII levels in strains with impaired glycogen metabolism To understand the role of glycogen metabolism during the transition to photomixotrophic conditions, we analysed the effect of Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation New Phytologist (2024) 243: 162–179 www.newphytologist.com New Phytologist Research 165 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License 2 mM glucose addition to photoautotrophically grown cultures of a WT Synechocystis strain and mutants lacking either PGM or AGP, which exhibit reduced or no glycogen synthesis, respectively (Ortega-Martınez et al., 2023, Fig. 2c). WT cells transitioned to photomixotrophic conditions displayed sixfold higher increase in OD 750 nm than cells kept under photoautotrophic conditions within the first 4 h (Fig. 2a,b). Concomitantly, glycogen rapidly accumulated, reaching levels in 4 h that exceeded those in photoautotrophic cultures after 24 h by >50% (Fig. 2c). Enhanced growth and glycogen synthesis were accompanied by complete consumption of the provided glucose within 24 h (Fig. 2d). By contrast, in ΔPGM and ΔAGP cultures, the addition of glucose induced an immediate growth arrest (Fig. 2b) and pigmentation bleaching within 48 h (Fig. 2a). Minimal glucose consumption was observed, with over 75% and 90% of the initial amounts remaining in the media after 24 h for ΔPGM and ΔAGP, respectively (Fig. 2d). The ΔPGM mutant showed reduced amounts of glycogen, reaching <20% of WT levels after 24 h, in both photomixotrophy and photoautotrophy (Fig. 2c). Glycogen was undetectable in the ΔAGP mutant, regardless of the availability of glucose in the medium (Fig. 2c). We measured the activities of hexokinase (HK) and other enzymes involved in the glucose-6P intersection, such as phosphoglucomutase (PGM), phosphoglucose isomerase (PGI) and glucose-6P dehydrogenase (G6PDH), to test if differences in glucose consumption rates between strains were caused by inactivity (Supporting Information Methods S1). The activity of these enzymes in cell extracts was similar between all strains (except for the absence of PGM activity in DPGM) and remained unchanged 4 h after glucose addition in all strains (Fig. S1). The progressive bleaching observed in both mutant cultures (Fig. 2a) prompted us to assess the relative levels of PSII, photosystem I (PSI), and ATP synthase by immunoblotting. Initially, there were no significant changes in the levels of the proteins examined during the first 4 h. However, while WT maintained similar protein levels, the abundance of D1 and CP47 PSII core proteins decreased 75–80% in the glycogen synthesis mutant strains between four and 24 h after glucose addition (Figs 2e, S2). The level of the PSI core protein PsaB decreased by 20% in ΔPGM and by 50% in ΔAGP between four and 24 h (Fig. 2e). ATPb(or GroEL, probed as a housekeeping protein) levels did not change in the mutants after glucose addition (Fig. 2e). A lower accumulation of photosystem proteins in the mutants upon a shift to photomixotrophy could possibly be linked to the generation of reactive oxygen species (ROS) (Johnson & Pakrasi, 2022). In vivo monitoring of ROS levels, using the probe 20,70- dichlorofluorescin diacetate (DCFH-DA) demonstrated elevated ROS by a four- and sixfold in ΔPGM and ΔAGP mutants, respectively, after 24 h under photomixotrophy (Fig. 2f). To investigate the role of oxidative stress protection upon a shift to photomixotrophy, we measured the in vivo levels of the key ROS scavenger glutathione using LC-MS. We detected similar glutathione pool sizes for all the strains in the absence of glucose (Fig. 2g). Upon the addition of glucose, WT cells gradually increased their glutathione pool size, indicating an oxidative protective response. Contrarily, the glycogen synthesis mutants emptied their glutathione pools by 24 h (Fig. 2g). Removing glucose from the photomixotrophic mutant cultures 4 h postglucose addition prevented the adverse effects of glucose on the fitness of the cultures. However, after 24 h of glucose exposure, the mutant strains could not be recovered (Fig. S3). Management of glucose-induced immediate metabolic alterations requires glycogen metabolism To dissect the metabolic responses of WT and glycogen synthesis mutant cells during transitions to photomixotrophy, we employed targeted metabolomics to monitor the key carbon pathways, including the OPP shunt, the CBB cycle, the Entner– Meyerhof–Parnas (EMP) pathway that shares some intermediates with the CBB cycle, and the tricarboxylic acid (TCA) cycle, as well as the pool sizes of detectable amino acids (Figs 3,4,S4; Dataset S1). Before glucose addition (0 min), all strains presented similar levels of most examined metabolites, indicating that glycogen deficiency did not lead to major metabolic alterations under photoautotrophic conditions (Figs 3, 4,S4). Glucose addition induced rapid alteration in the levels of nearly all metabolites in all strains, with detectable changes already 5 min after glucose addition. In WT, two distinct patterns of metabolic changes were observed (Figs 3, 4,S4). First, levels of the RuBisCO substrate ribulose-1,5-BP (RuBP), some of the metabolites shared by the CBB cycle and the EMP pathway (e.g. fructose-1,6-BP (F1,6BP), dihydroxyacetone-P (DHAP), 3PG), the last steps of the EMP pathway (e.g. phosphoenolpyruvate (PEP), pyruvate), and most TCA cycle intermediates underwent transient fluctuations, peaking 5–15 min after glucose addition (Figs 3, 4,S4a). These metabolites typically more than doubled their initial amounts before returning to photoautotrophic levels. Amino acids alanine and serine, derived from pyruvate and 3PG, respectively, increased more than fivefold, similarly to threonine and valine. Second, levels of metabolites from the OPP shunt (e.g. G6P, 6P-gluconate (6PG)), those from the CBB cycle (e.g. erythrose-4P (E4P) and sedoheptulose-7P (S7P)) and others in the interconnection of this cycle and the gluconeogenic EMP pathway (e.g. glyceraldehyde-3P (GAP), fructose-6P (F6P)) increased linearly three- to fivefold until 120 min (Figs 3,S4a). In the glycogen synthesis mutants, glucose addition induced faster and more pronounced intermediate accumulation than in WT (Figs 3, 4,S4b). The first metabolites involved in glucose assimilation (G6P, 6PG, F6P) through the OPP shunt and PGI shunt increased rapidly and dramatically (by 5 min more than 40-, 10-, and 25-fold, respectively). This increase is key to enhance CO 2 fixation by providing CBB cycle intermediates under photomixotrophy (Schulze et al., 2022). G1P, the first metabolite in the glycogen synthesis pathway, steadily increased by up to 10-fold in ΔAGP mutant but remained at low levels in ΔPGM, due to limited G6P/G1P interconversion in that mutant (Fig. S1, Ortega-Martınez et al., 2023). New Phytologist (2024) 243: 162–179 www.newphytologist.com Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation Research New Phytologist 166 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License Fig. 2 Phenotypic characterisation of WT Synechocystis and strains with deficient glycogen metabolism after 2 mM glucose supplementation. Cultures grown under photoautotrophic conditions in BG11C were adjusted to a final optical density of 1 at 750 nm (OD 750 nm ), divided into two flasks, and supplemented (photomixotrophy) or not (photoautotrophy) with 2 mM glucose. (a) Photographs of WT, ΔPGM and ΔAGP strains cultivated in parallel under either photoautotrophic or photomixotrophic conditions for 48 h. (b) Growth curves of WT, ΔPGM and ΔAGP strains measured as OD 750 nm in both photoautotrophy (control, dash lines) and photomixotrophy (solid lines). (c) Time course glycogen content per optical density (OD 750 nm )ofWT,ΔPGM and ΔAGP strains in the presence (solid lines) or absence (dashed lines) of glucose. (d) Glucose concentration in the media for 24 h after the initial addition of 2 mM of glucose. (e). Western blot analysis of proteins from the PETC (D1 and CP47 proteins of PSII and PsaB of PSI) and ATP synthase (ATP b ,bsubunit of ATP synthase) in the WT, ΔPGM and ΔAGP strains at 0, 4 and 24 h after glucose addition. A representative western blot of six biological replicates is shown Quantifications of band intensity of all blots are shown in Supporting Information Fig. S2. (f) Quantification of cellular reactive oxygen species (ROS) accumulation measured by DFC fluorescence in the WT, ΔPGM and ΔAGP strains at 0, 4 and 24 h after glucose addition. (g) Quantification of glutathione pools in the WT, ΔPGM and ΔAGP strains at 0, 4 and 24 h after glucose addition. The data represent the mean SE from six (c–e), five (f) and three (g) biological replicates, respectively. Statistical significance levels are denoted by *,P<0.05; ****,P<0.0001; otherwise, not significant (two-way ANOVA). Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation New Phytologist (2024) 243: 162–179 www.newphytologist.com New Phytologist Research 167 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License Fig. 3 Metabolic changes in upper metabolism of WT, DPGM and DAGP Synechocystis strains after glucose addition. Cultures of WT, ΔPGM and ΔAGP strains grown in BG11C were adjusted to a final optical density (OD 750 nm ) of 1 and harvested before (0) and 5, 15, 30 and 120 min postglucose supplementation (2 mM). The time course displays 14 metabolites involved in glucose assimilation, the shared Oxidative Pentose Phosphate shunt (blue arrows), the Calvin–Benson–Bassham cycle (green arrows), glycogen metabolism (grey arrows) and the Entner–Meyerhof–Parnas pathway. Metabolite names highlighted with coloured rectangles refer to molecules shared with different pathways (indicated by dashed arrows). 3PG, 3P-glycerate; DHAP, dihydroxyacetone-P; E4P, erythrose-4P; GAP, glyceraldehyde-3P; R5P, ribose-5P; Ru5P/Xu5P, ribulose-5P/xylulose-5P pool; RuBP, ribulose-1,5P; S1,7BP, sedoheptulose-1,7P; S7P, sedoheptulose-7P. Values represent the mean SE or fold change SE relative to WT value before glucose addition of 6 independent biological replicates. New Phytologist (2024) 243: 162–179 www.newphytologist.com Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation Research New Phytologist 168 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License Fig. 4 Metabolic changes in 3PG downstream metabolism of WT, DPGM and DAGP Synechocystis strains after glucose addition. Cultures of WT, ΔPGM and ΔAGP strains grown in BG11C were adjusted to a final optical density (OD 750 nm ) of 1 and harvested before (0) and 5, 15, 30 and 120 min postglucose supplementation (2 mM). The time course display 17 metabolites involved in lower glycolysis, tricarboxylic acid cycle and ammonium assimilation. 2-OG, 2-oxoglutarate; AcCoA, acetyl-CoA; OAA, oxalacetate; PEP, phosphoenolpyruvate; Pyr, pyruvate; SSAL, succinic semialdehyde. Dashed arrows indicate steps involving several reactions. Values represent the mean SE or fold change SE relative to WT value before glucose addition of 6 independent biological replicates. Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation New Phytologist (2024) 243: 162–179 www.newphytologist.com New Phytologist Research 169 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License We also detected photomixotrophy-induced changes in CBB cycle metabolites in the glycogen synthesis mutants. The concentrations of the ribulose-5P (Ru5P)/xylulose-5P (Xu5P) pool and ribose-5P (R5P) both doubled within 5 min of glucose addition and remained stable throughout the experiment. RuBP, however, increased >10-fold. The metabolites S7P, E4P, and GAP exhibited similar glucose-induced patterns to G6P or F6P in the mutants, accumulating 50-, 30-, and 10-fold, respectively. By contrast, DHAP, an isomer of GAP, showed a similar trend to F1,6BP and correlated with WT values. Finally, 3PG, the product of RuBisCO, increased only transiently in WT and accumulated steadily in glycogen synthesis mutants up to twofold, which constitutes a substantial increase due to a high initial concentration. Downstream from 3PG, the first metabolite measured was PEP (Fig. 4). PEP was reduced fivefold in ΔPGM and ΔAGP 5 min after glucose addition and remained at very low levels throughout the experiment. The decrease in PEP was coupled to an increase in pyruvate, reaching >0.25 nmol OD 750 1 120 min after glucose addition. Levels of acetyl coenzyme A (AcCoA) remained steady in the mutants upon transition to photomixotrophy (Fig. 4). AcCoA plays a central role in cellular metabolism, serving as a key intermediate compound into the TCA cycle. Levels of most intermediates in the TCA cycle doubled, with citrate accounting for most of the carbon allocation. Finally, amino acid pools underwent several photomixotrophy-induced changes in the glycogen synthesis mutants (Figs 3, 4,S4). Alanine and serine increased c. sixfold, and levels remained higher than 5 nmol OD 750 1 in contrast to the transient accumulation observed in WT. Aspartate, threonine, valine and tryptophan showed transient accumulation peaking between two- and fourfold photoautotrophic levels at 30 min after glucose addition in both glycogen synthesis mutants, compared to unchanged levels in WT. Conversely, glutamate, an amino acid essential to nitrogen assimilation in the GS-GOGAT cycle, decreased to <2 nmol OD 750 1 despite being one of the most abundant metabolites measured (>10 nmol OD 750 1 under photoautotrophic conditions). Other amino acids, such as tyrosine, phenylalanine, methionine and leucine-isoleucine were considerably depleted in ΔPGM and ΔAGP mutants, while WT showed similar levels or slight increases compared to photoautotrophy (Fig. S4b). In summary, Synechocystis exhibits metabolic flexibility upon glucose addition, channelling carbon through the PGI and OPP shunts to fuel the CBB cycle, coupled with transient fluctuations in some glycolytic and TCA cycle intermediates. The impact was more pronounced in ΔPGM and ΔAGP mutants, showing rapid and sustained increases in glucose assimilation metabolites and the CBB cycle within 5 min, highlighting the crucial role of glycogen synthesis in photomixotrophic acclimation. Photosynthesis is impaired in the glycogen synthesis mutant strains cultured with glucose Considering the impaired growth (Fig. 2b) and decreased levels of photosynthetic proteins of the glycogen-deficient mutants under photomixotrophy (Fig. 2e), we next monitored photosynthetic performance in both photoautotrophy and 30 min postglucose addition, capturing the metabolic overflow in both glycogen-deficient mutants (Fig. S5), but before PSII levels decreased (Fig. 2e) and before long-term acclimation to photomixotrophy. Real-time measurement of photosynthetic CO 2 gas flux (Fig. 5) by MIMS revealed similar rates and kinetics of CO 2 Fig. 5 Effect of glucose supplementation on the rate of CO 2 fluxes in WT, ΔPGM and ΔAGP Synechocystis strains determined by MIMS. (a) CO 2 exchange rates in photoautotrophy and 30 min after 2 mM glucose addition to 10 lg Chl ml 1 WT, ΔPGM and ΔAGP cultures grown in BG11 measured in a Membrane Inlet Mass Spectrometer (MIMS). The yellow rectangles represent an exposure to a light intensity of 50 lmol photons m 2 s 1 while the white rectangles represent dark periods. In photoautotrophic conditions, upon the onset of light, the carbon concentration mechanism (CCM) triggered an initial concentration step absorbing high amounts of CO 2 (phase I). Subsequently, during the rest of the light period, consumption reaches a steady state (phase II). Conversely, when the lights are switched off, CCM activity ceases, leading to the release and efflux of the intracellular inorganic carbon pool back into the media (phase III). (b) Quantification of CO 2 fluxes during the steady-state in the light (phase II) in photoautotrophy or 30 min after 2 mM glucose addition. The data represent the mean SE from four biological replicates. Statistical significance levels are denoted by *,P<0.05; **,P<0.01; ns, not significant (two-way ANOVA). New Phytologist (2024) 243: 162–179 www.newphytologist.com Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation Research New Phytologist 170 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License Foundation project ‘PhotoCat’ project (no. NNF20OC0064371 to YA). PO-M is a recipient of grant FPU18/06580 from the Ministerio de Universidades and an EMBO Scientific Exchange Grant. We thank Dr Tuomas Huokko and Michal Hubacek for assistance with PO-M’s research visit to University of Turku, and Dr Luis Lopez-Maury for critical reading of the manuscript. The photosynthetic electron transport studies (except Chl fluorescence analysis) were conducted in the PhotoSYN Finnish Infrastructure for Photosynthesis Research. Competing interests None declared. Author contributions SD-T and FJF conceived the research, all the authors designed the research, PO-M: performed all the experiments, and wrote the manuscript draft; LN performed MIMS measurements and drafted the photosynthesis section; SD-T supervised physiological and metabolomics experiments; LTW developed photosynthesis experiment analysis scripts; FJF and YA, funding acquisition; all the authors revised and approved the manuscript. PO-M and LN contributed equally to this work. ORCID Yagut Allahverdiyeva https://orcid.org/0000-0002-9262- 1757 Sandra Dıaz-Troya https://orcid.org/0000-0002-5878-9062 Francisco J. Florencio https://orcid.org/0000-0002-2068- 7861 Lauri Nikkanen https://orcid.org/0000-0002-7192-9322 Pablo Ortega-Martınez https://orcid.org/0000-0001-5508- 5636 Laura T. Wey https://orcid.org/0000-0003-2345-0699 Data availability The data that support the findings of this study are available in the article and its Supporting Information. References Baikie TK, Wey LT, Lawrence JM, Medipally H, Reisner E, Nowaczyk MM, Friend RH, Howe CJ, Schnedermann C, Rao A et al. 2023. Photosynthesis rewired on the pico-second timescale. Nature 615: 836–840. Ball SG, Morell MK. 2003. From bacterial glycogen to starch: understanding the biogenesis of the plant starch granule. Annual Review of Plant Biology 54: 207– 233. Beckmann K, Messinger J, Badger MR, Wydrzynski T, Hillier W. 2009. Online mass spectrometry: membrane inlet sampling. Photosynthesis Research 102: 511–522. Berthomieu C, Hienerwadel R. 2001. Iron coordination in photosystem II: interaction between bicarbonate and the Q B pocket studied by Fourier transform infrared spectroscopy. Biochemistry 40: 4044–4052. Brinkert K, De Causmaecker S, Krieger-Liszkay A, Fantuzzi A, Rutherford AW. 2016. Bicarbonate-induced redox tuning in Photosystem II for regulation and protection. Proceedings of the National Academy of Sciences, USA 113: 12144– 12149. Burnap RL, Hagemann M, Kaplan A. 2015. Regulation of CO 2 concentrating mechanism in cyanobacteria. Life 5: 348–371. Cano M, Holland SC, Artier J, Burnap RL, Ghirardi M, Morgan JA, Yu J. 2018. Glycogen synthesis and metabolite overflow contribute to energy balancing in cyanobacteria. Cell Reports 23: 667–672. Carrieri D, Paddock T, Maness PC, Seibert M, Yu J. 2012. Photo-catalytic conversion of carbon dioxide to organic acids by a recombinant cyanobacterium incapable of glycogen storage. Energy and Environmental Science 5: 9457–9461. Cifuente JO, Comino N, Trastoy B, D’Angelo C, Guerin ME. 2019. Structural basis of glycogen metabolism in bacteria. Biochemical Journal 476: 2059–2092. Dıaz-Troya S, Lopez-Maury L, Sanchez-Riego AM, Roldan M, Florencio FJ. 2014. Redox regulation of glycogen biosynthesis in the cyanobacterium Synechocystis sp. PCC 6803: analysis of the AGP and glycogen synthases. Molecular Plant 7:87–100. Doello S, Neumann N, Forchhammer K. 2022. Regulatory phosphorylation event of phosphoglucomutase 1 tunes its activity to regulate glycogen metabolism. FEBS Journal 289: 6005–6020. Fantuzzi A, Haniewicz P, Farci D, Loi MC, Park K, B€uchel C, Bochtler M, Rutherford AW, Piano D. 2023. Bicarbonate activation of the monomeric photosystem II-PsbS/Psb27 complex. Plant Physiology 192: 2656–2671. Field CB, Behrenfeld MJ, Randerson JT, Falkowski P. 1998. Primary production of the biosphere: integrating terrestrial and oceanic components. Science 281: 237–240. Fufezan C, Gross CM, Sj€odin M, Rutherford AW, Krieger-Liszkay A, Kirilovsky D. 2007. Influence of the redox potential of the primary quinone electron acceptor on photoinhibition in photosystem II. Journal of Biological Chemistry 282: 12492–12502. Gr€undel M, Scheunemann R, Lockau W, Zilliges Y. 2012. Impaired glycogen synthesis causes metabolic overflow reactions and affects stress responses in the cyanobacterium Synechocystis sp. PCC 6803. Microbiology 158: 3032–3043. Haimovich-Dayan M, Kahlon S, Hihara Y, Hagemann M, Ogawa T, Ohad I, Lieman-Hurwitz J, Kaplan A. 2011. Cross-talk between photomixotrophic growth and CO2-concentrating mechanism in Synechocystis sp. strain PCC 6803. Environmental Microbiology 13: 1767–1777. Heifetz PB, F€o B, Osmond CB, Giles LJ, Boynton JE. 2000. Effects of acetate on facultative autotrophy in Chlamydomonas reinhardtii assessed by photosynthetic measurements and stable isotope analyses 1. Plant Physiology 122:1439–1445. Ishikita H, Galstyan A, Knapp EW. 2007. Redox potential of the non-heme iron complex in bacterial photosynthetic reaction center. Biochimica et Biophysica Acta –Bioenergetics 1767: 1300–1309. Johnson VM, Pakrasi HB. 2022. Advances in the understanding of the lifecycle of photosystem II. Microorganisms 10:1–19. Kahlon S, Beeri K, Ohkawa H, Hihara Y, Murik O, Suzuki I, Ogawa T, Kaplan A. 2006. A putative sensor kinase, Hik31, is involved in the response of Synechocystis sp. strain PCC 6803 to the presence of glucose. Microbiology 152: 647–655. Kanno M, Carroll AL, Atsumi S. 2017. Global metabolic rewiring for improved CO 2 fixation and chemical production in cyanobacteria. Nature Communications 8: 14724. Latifi A, Ruiz M, Zhang CC. 2009. Oxidative stress in cyanobacteria. FEMS Microbiology Reviews 33: 258–278. Lea-Smith DJ, Bombelli P, Vasudevan R, Howe CJ. 2016. Photosynthetic, respiratory and extracellular electron transport pathways in cyanobacteria. Biochimica et Biophysica Acta (BBA) –Bioenergetics 1857: 247–255. Lee HW, Park BS, Joo JH, Patidar SK, Choi HJ, Jin ES, Han MS. 2018. Cyanobacteria-specific algicidal mechanism of bioinspired naphthoquinone derivative, NQ 2-0. Scientific Reports 8: 11595. Lee S, Ryu JY, Soo YK, Jeon JH, Ji YS, Cho HT, Choi SB, Choi D, De Marsac NT, Il PY. 2007. Transcriptional regulation of the respiratory genes in the cyanobacterium Synechocystis sp. PCC 6803 during the early response to glucose feeding. Plant Physiology 145: 1018–1030. Luan G, Zhang S, Wang M, Lu X. 2019. Progress and perspective on cyanobacterial glycogen metabolism engineering. Biotechnology Advances 37: 771–786. Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation New Phytologist (2024) 243: 162–179 www.newphytologist.com New Phytologist Research 177 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License Makowka A, Nichelmann L, Schulze D, Spengler K, Wittmann C, Forchhammer K, Gutekunst K. 2020. Glycolytic shunts replenish the Calvin– Benson–Bassham cycle as anaplerotic reactions in cyanobacteria. Molecular Plant 13: 471–482. Mallen-Ponce MJ, Marıa Esther Perez-Perez MP, Crespo JL. 2022. Photosynthetic assimilation of CO 2 regulates TOR activity. Proceedings of the National Academy of Sciences, USA 119: e2115261119. Markwell MAK, Haas SM, Bieber LL, Tolbert NE. 1978. A modification of the lowry procedure to simplify protein determination in membrane and lipoprotein sample. Analytical Biochemistry 87: 206–210. Matson MM, Atsumi S. 2018. Photomixotrophic chemical production in cyanobacteria. Current Opinion in Biotechnology 50:65–71. Miller NT, Ajlani G, Burnap RL. 2022. Cyclic electron flow-coupled proton pumping in Synechocystis sp. PCC6803 is dependent upon NADPH oxidation by the soluble isoform of Ferredoxin:NADP-Oxidoreductase. Microorganisms 10: 855. Mullineaux CW. 2014. Co-existence of photosynthetic and respiratory activities in cyanobacterial thylakoid membranes. Biochimica et Biophysica Acta (BBA) – Bioenergetics 1837: 503–511. Mu~noz-Marın MDC, Lopez-Lozano A, Moreno-Cabezuelo J  A, Dıez J, Garcıa- Fernandez JM. 2024. Mixotrophy in cyanobacteria. Current Opinion in Microbiology 78: 102432. Murata N, Takahashi S, Nishiyama Y, Allakhverdiev SI. 2007. Photoinhibition of photosystem II under environmental stress. Biochimica et Biophysica Acta (BBA) –Bioenergetics 1767: 414–421. Muth-Pawlak D, Kreula S, Gollan PJ, Huokko T, Allahverdiyeva Y, Aro EM. 2022. Patterning of the autotrophic, mixotrophic, and heterotrophic proteomes of oxygen-evolving cyanobacterium Synechocystis sp. PCC 6803. Frontiers in Microbiology 13: 891895. Nakajima T, Kajihata S, Yoshikawa K, Matsuda F, Furusawa C, Hirasawa T, Shimizu H. 2014. Integrated metabolic flux and omics analysis of Synechocystis sp. PCC 6803 under mixotrophic and photoheterotrophic conditions. Plant and Cell Physiology 55: 1606–1612. Nikkanen L, Santana Sanchez A, Ermakova M, R€ogner M, Cournac L, Allahverdiyeva Y. 2020. Functional redundancy between flavodiiron proteins and NDH-1 in Synechocystis sp. PCC 6803. The Plant Journal 103: 1460– 1476. Nikkanen L, Solymosi D, Jokel M, Allahverdiyeva Y. 2021. Regulatory electron transport pathways of photosynthesis in cyanobacteria and microalgae: recent advances and biotechnological prospects. Physiologia Plantarum 173: 514–525. Ogawa T, Sonoike K. 2016. Effects of bleaching by nitrogen deficiency on the quantum yield of Photosystem II in Synechocystis sp. PCC 6803 revealed by Chl fluorescence measurements. Plant and Cell Physiology 57: 558–567. Ogawa T, Suzuki K, Sonoike K. 2021. Respiration interacts with photosynthesis through the acceptor side of photosystem I, reflected in the dark-to-light induction kinetics of chlorophyll fluorescence in the cyanobacterium Synechocystis sp. PCC 6803. Frontiers in Plant Science 12: 717968. Ortega-Martınez P, Roldan M, Dıaz-Troya S, Florencio FJ. 2023. Stress response requires an efficient connection between glycogen and central carbon metabolism by phosphoglucomutases in cyanobacteria. Journal of Experimental Botany 74: 1532–1550. Rippka R, Deruelles J, Waterbury JB, Herdman M, Stanier RY. 1979. Generic assignments, strain histories and properties of pure cultures of cyanobacteria. Microbiology 111:1–61. Roach T, Sedoud A, Krieger-Liszkay A. 2013. Acetate in mixotrophic growth medium affects photosystem II in Chlamydomonas reinhardtii and protects against photoinhibition. Biochimica et Biophysica Acta –Bioenergetics 1827: 1183–1190. Rodríguez-Gil T, Torrado A, Iniesta-PallarésM,Álvarez C, Mariscal V, Molina-Heredia FP. 2021. Cytochrome c M is probably a membrane protein similar to the C subunit of the bacterial nitric oxide reductase. Applied Sciences 11: 9396. Schreiber U, Klughammer C. 2016. Analysis of photosystem I donor and acceptor sides with a new type of online-deconvoluting kinetic LED-array spectrophotometer. Plant and Cell Physiology 57: 1454–1467. Schuller JM, Birrell JA, Tanaka H, Konuma T, Wulfhorst H, Cox N, Schuller SK, Thiemann J, Lubitz W, Setif P et al. 2019. Structural adaptations of photosynthetic complex I enable ferredoxin-dependent electron transfer. Science 363: 257–260. Schulze D, Kohlstedt M, Becker J, Cahoreau E, Peyriga L, Makowka A, Hildebrandt S, Gutekunst K, Portais JC, Wittmann C. 2022. GC/MS-based 13C metabolic flux analysis resolves the parallel and cyclic photomixotrophic metabolism of Synechocystis sp. PCC 6803 and selected deletion mutants including the Entner-Doudoroff and phosphoketolase pathways. Microbial Cell Factories 21: 69. Shevela D, Klimov V, Messinger J. 2007. Interactions of photosystem II with bicarbonate, formate and acetate. Photosynthesis Research 94: 247–264. Shi LX, Hall M, Funk C, Schr€oder WP. 2012. Photosystem II, a growing complex: updates on newly discovered components and low molecular mass proteins. Biochimica et Biophysica Acta –Bioenergetics 1817:13–25. Solymosi D, Nikkanen L, Muth-Pawlak D, Fitzpatrick D, Vasudevan R, Howe CJ, Lea-Smith DJ, Allahverdiyeva Y. 2020. Cytochrome CM decreases photosynthesis under photomixotrophy in Synechocystis sp. PCC 6803. Plant Physiology 183: 700–716. Solymosi D, Shevela D, Allahverdiyeva Y. 2022. Nitric oxide represses photosystem II and NDH-1 in the cyanobacterium Synechocystis sp. PCC 6803. Biochimica et Biophysica Acta –Bioenergetics (BBA) 1863: 148507. Sun Z, Zhao X, Li G, Yang J, Chen Y, Xia M, Hwang I, Hou H. 2023. Metabolic flexibility during a trophic transition reveals the phenotypic plasticity of greater duckweed (Spirodela polyrhiza 7498). New Phytologist 238: 1386–1402. Theune ML, Hildebrandt S, Steffen-Heins A, Bilger W, Gutekunst K, Appel J. 2021. In-vivo quantification of electron flow through photosystem I –cyclic electron transport makes up about 35% in a cyanobacterium. Biochimica et Biophysica Acta –Bioenergetics 1862: 148353. Thiel K, Vuorio E, Aro EM, Kallio PT. 2017. The effect of enhanced acetate influx on Synechocystis sp. PCC 6803 metabolism. Microbial Cell Factories 16: 21. Trautmann D, Voß B, Wilde A, Al-Babili S, Hess WR. 2012. Microevolution in cyanobacteria: re-sequencing a motile substrain of Synechocystis sp. PCC 6803. DNA Research 19: 435–448. Vass I, Kirilovsky D, Etienne AL. 1999. UV-B radiation-induced donor- and acceptor-side modifications of photosystem II in the cyanobacterium Synechocystis sp. PCC 6803. Biochemistry 38: 12786–12794. Wan N, Abernathy M, Tang JKH, Tang YJ, You L. 2015. Cyanobacterial photo-driven mixotrophic metabolism and its advantages for biosynthesis. Frontiers of Chemical Science and Engineering 9: 308–316. Wang Y, Chen X, Spengler K, Terberger K, Boehm M, Appel J, Barske T, Timm S, Battchikova N, Hagemann M et al. 2022. Pyruvate : ferredoxin oxidoreductase and low abundant ferredoxins support aerobic photomixotrophic growth in cyanobacteria. eLife 11: e71339. Wydrzynski T, Govindjee. 1975. A new site of bicarbonate effect in photosystem II of photosynthesis: evidence from chlorophyll fluorescence transients in spinach chloroplasts. Biochimica et Biophysica Acta 387: 403–408. Yamada M, Nagao R, Iwai M, Arai Y, Makita A, Ohta H, Tomo T. 2018. The PsbQ0protein affects the redox potential of the Q A in photosystem II. Photosynthetica 56: 185–191. Yoshikawa K, Hirasawa T, Ogawa K, Hidaka Y, Nakajima T, Furusawa C, Shimizu H. 2013. Integrated transcriptomic and metabolomic analysis of the central metabolism of Synechocystis sp. PCC 6803 under different trophic conditions. Biotechnology Journal 8: 571–580. Zavrel T, OcenasovaP,  Cerveny J. 2017. Phenotypic characterization of Synechocystis sp. PCC 6803 substrains reveals differences in sensitivity to abiotic stress. PLoS ONE 12: e0189130. Supporting Information Additional Supporting Information may be found online in the Supporting Information section at the end of the article. Dataset S1 Quantification of metabolites in WT, DPGM and DAGP Synechocystis strains after glucose addition. New Phytologist (2024) 243: 162–179 www.newphytologist.com Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation Research New Phytologist 178 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License Fig. S1 Measurements of enzymatic activities at the glucose-6P metabolic crossroads in WT, ΔPGM and ΔAGP Synechocystis cell extracts. Fig. S2 Effect of glucose on the photosynthetic machinery integrity in WT, ΔPGM and ΔAGP Synechocystis strains. Fig. S3 Growth recovery of WT, ΔPGM and ΔAGP Synechocystis strains after glucose removal from the media. Fig. S4 Time course metabolic profile extension. Fig. S5 Comparison of metabolite levels under autotrophy and 30 min after glucose addition to WT, ΔPGM and ΔAGP Synechocystis cultures. Fig. S6 Quantification of CO 2 fluxes in WT, ΔPGM and ΔAGP Synechocystis strains before and 30 min after glucose supplementation. Fig. S7 Photosystems abundance ratio in photoautotrophy and 30 min after glucose addition to WT, ΔPGM and ΔAGP Synechocystis strains. Fig. S8 Analysis of the effect of glucose on the PSII effective yield in WT, ΔPGM and ΔAGP Synechocystis strains. Fig. S9 Measurements of O 2 fluxes in MIMS in WT, ΔPGM and ΔAGP Synechocystis strains. Fig. S10 Real-time effects of glucose addition on oxygen evolution and CO 2 uptake in ΔAGP cultures determined by MIMS. Fig. S11 Redox state changes of P700 (PSI), its donor side Plastocyanin and its acceptor side Ferredoxin in photoautotrophy and 30 min after glucose addition to WT, ΔPGM and ΔAGP Synechocystis cultures. Methods S1 Enzyme activity assays. Methods S2 Photosystems abundance ratio. Please note: Wiley is not responsible for the content or functionality of any Supporting Information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office. Ó2024 The Authors New Phytologist Ó2024 New Phytologist Foundation New Phytologist (2024) 243: 162–179 www.newphytologist.com New Phytologist Research 179 14698137, 2024, 1, Downloaded from https://nph.onlinelibrary.wiley.com/doi/10.1111/nph.19793 by Universidad De Sevilla, Wiley Online Library on [13/06/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License