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. 2019 Oct 24;182(1):507–517. doi: 10.1104/pp.19.01184

Glycogen Metabolism Supports Photosynthesis Start through the Oxidative Pentose Phosphate Pathway in Cyanobacteria1,[OPEN]

Shrameeta Shinde a, Xiaohui Zhang a,b, Sonali P Singapuri a, Isha Kalra a, Xianhua Liu b, Rachael M Morgan-Kiss a, Xin Wang a,2,3
PMCID: PMC6945877  PMID: 31649110

Glycogen metabolism kick-starts photosynthesis during cyanobacterial diurnal growth.

Abstract

Cyanobacteria experience drastic changes in their carbon metabolism under daily light/dark cycles. During the day, the Calvin-Benson cycle fixes CO2 and diverts excess carbon into glycogen storage. At night, glycogen is degraded to support cellular respiration. The dark/light transition represents a universal environmental stress for cyanobacteria and other photosynthetic lifeforms. Recent studies revealed the essential genetic background necessary for the fitness of cyanobacteria during diurnal growth. However, the metabolic processes underlying the dark/light transition are not well understood. In this study, we observed that glycogen metabolism supports photosynthesis in the cyanobacterium Synechococcus elongatus PCC 7942 when photosynthesis reactions start upon light exposure. Compared with the wild type, the glycogen mutant ∆glgC showed a reduced photosynthetic efficiency and a slower P700+ rereduction rate when photosynthesis starts. Proteomic analyses indicated that glycogen is degraded through the oxidative pentose phosphate (OPP) pathway during the dark/light transition. We confirmed that the OPP pathway is essential for the initiation of photosynthesis and further showed that glycogen degradation through the OPP pathway contributes to the activation of key Calvin-Benson cycle enzymes by modulating NADPH levels. This strategy stimulates photosynthesis in cyanobacteria following dark respiration and stabilizes the Calvin-Benson cycle under fluctuating environmental conditions, thereby offering evolutionary advantages for photosynthetic organisms using the Calvin-Benson cycle for carbon fixation.

Photosynthesis supports life on Earth by converting solar energy into chemical energy stored as organic carbon. The Calvin-Benson cycle, the major carbon fixation pathway in cyanobacteria, algae, and plants, is employed to reduce CO2 into organic carbon molecules. Since its discovery in the 1950s, the enzymes of the Calvin-Benson cycle have been elucidated and the consensus pathways established (Bassham et al., 1954). CO2 is reduced into the photosynthesis output glyceraldehyde-3-phosphate (G3P) in three stages (i.e. carbon fixation, carbon reduction, and carbon regeneration). The Calvin-Benson cycle is an elegant pathway in which the starting substrate ribulose-1,5-bisphosphate (RuBP) is continuously supplied by partially recycling G3P back to RuBP through carbon rearrangement reactions. This carbon regeneration stage resembles the nonoxidative portion of the pentose phosphate pathway. However, the Calvin-Benson cycle uses a key enzyme, sedoheptulose-1,7-bisphosphatase (SBPase), to drive carbon regeneration rather than using transaldolase found in the pentose phosphate pathway (Sharkey and Weise, 2016). The activity of SBPase often determines the photosynthetic capacity and carbon accumulation in downstream metabolic processes (Harrison et al., 1997, 2001; Raines et al., 1999).

In cyanobacteria, a major portion of the photosynthetically fixed carbon is used to synthesize the storage polymer glycogen. Two enzymes, ADP-Glc pyrophosphorylase (AGPase) and glycogen synthase, catalyze the sequential conversion of Glc-1-P (G1P) to 1,4-α-glucan (Preiss, 1984). Another enzyme encoded by a 1,4-α-glucan-branching enzyme gene, glgB, can catalyze the formation of the 1,6-α branches of glycogen (Preiss, 1984). Glycogen synthesis is closely linked to photosynthesis carbon output. The photosynthate 3-phosphoglycerate (3PG) is an allosteric activator of the AGPase (Preiss, 1984; Gómez-Casati et al., 2003). When photosynthetically fixed carbon is in excess, a major carbon flux will be diverted to glycogen storage, serving as the carbon and electron sources for cellular respiration in the dark (Preiss, 1984; Suzuki et al., 2010). More recently, glycogen metabolism has also been recognized for additional roles played in cyanobacterial carbon metabolism. These studies revealed the involvement of glycogen metabolism as an energy-buffering system to maintain homeostasis (Cano et al., 2018) and as the carbon source for rapid resuscitation from nitrogen chlorosis (Doello et al., 2018).

Glycogen metabolism and cellular respiration are critical for cell viability in the dark until light resumes (Lehmann and Wöber, 1976). Research in the past has identified the oxidative pentose phosphate (OPP) pathway (also known as the Glc-6-P [G6P] shunt) as the major route for glycogen degradation in the dark (Smith, 1983; Broedel and Wolf, 1990). On the other hand, the OPP pathway seems to be indispensable only after long periods of darkness (over 24 h), while glycolysis might have been able to compensate for the need of a reducing equivalent during short incubation in the dark (Scanlan et al., 1995). However, it is also important to know that the OPP pathway is indispensable for cyanobacterial dark survival because many reactive oxygen species-detoxifying enzymes require NADPH as cofactors (Welkie et al., 2019). In cyanobacteria, cellular respiration is closely associated with photosynthesis. Many intermediates are shared between the carbon fixation and respiration processes. When photosynthesis reactions start upon light, intermediates in the Calvin-Benson cycle might be limited after dark respiration, leading to stalled carbon fixation reactions. Dynamic metabolic regulation to ensure a smooth transition from dark respiration to photosynthesis reactions is thus essential for the fitness of cyanobacteria during diurnal growth (Puszynska and O’Shea, 2017; Welkie et al., 2018). In this study, we discovered the active participation of glycogen metabolism during the dark/light transition in cyanobacteria. We found that glycogen degradation through the OPP pathway helps activate and stabilize the Calvin-Benson cycle reactions when photosynthesis reactions start upon light. We propose that this strategy helps support photosynthesis during the dark/light transition and ensures the growth advantage of photosynthetic organisms in their natural environment.

RESULTS

Proteomic Analysis Suggests a Supportive Role of Glycogen Metabolism for Photosynthesis

Glycogen biosynthesis is a major carbon assimilatory pathway in the cyanobacterium Synechococcus elongatus PCC 7942. In this study, we generated a glycogen synthesis mutant (∆glgC) by deleting the AGPase gene (glgC) in the S. elongatus genome (Fig. 1A). When wild-type and ∆glgC mutant cells were grown under continuous light (50 µmol photons m−2 s−1), the ∆glgC mutant exhibited a longer lag phase relative to wild-type cells (Fig. 1B). This observation suggests that glycogen metabolism might play a supportive role for the rapid start of photosynthesis (i.e. a short lag phase to quickly start the carbon fixation process).

Figure 1.

Figure 1.

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Impact of glycogen metabolism on S. elongatus growth. A, Glycogen synthesis pathway in S. elongatus. The gene glgC was deleted from the S. elongatus genome to create the glycogen mutant (glgC::kan). B, Growth curves of the S. elongatus wild type (WT) and ΔglgC mutant under continuous light. C, Comparative proteomic analysis of the S. elongatus wild type and ΔglgC mutant. Enzymes for both glycogen biosynthesis (AGPase [glgC], glycogen synthase [glgA], and 1,4-α-glucan-branching enzyme [glgB]) and degradation (α-1,4-glucan phosphorylase [glgP]) were in higher abundance in wild-type compared with ΔglgC mutant cells. Bar graphs indicate enzyme expression levels based on the normalized spectral abundance factor. Error bars indicate the sd; two-tailed Student’s t test was used to compare protein abundances: **, P < 0.01 and ***, P < 0.001.

To understand how glycogen biosynthesis supports photosynthesis, we conducted a comparative proteomic analysis on the wild-type and ∆glgC mutant cells. The AGPase was not detected in the ∆glgC mutant, showing the success of constructing a clean mutant. All three enzymes related to glycogen synthesis (i.e. AGPase [glgC], glycogen synthase [glgA], and 1,4-α-glucan-branching enzyme [glgB]), were found to be in significantly higher abundance in the wild type compared with the ∆glgC mutant (Fig. 1C). Interestingly, enzymes related to glycogen degradation were also found to be more abundant in wild-type cells. The α-1,4-glucan phosphorylase (glgP) that removes Glc from the glycogen chain was expressed over 4-fold higher in wild-type cells (Fig. 1C; Supplemental Table S1). However, if glycogen degradation was to continue through the glycolytic Embden-Meyerhof-Parnas pathway, it would be a futile cycle to carbon fixation reactions. We hypothesize that the glycogen hydrolysis might proceed through other glycolytic pathways to support the rapid photosynthesis start in wild-type cells. We began to test our hypothesis by first validating the impact of abolished glycogen synthesis on photosynthetic functions.

The Glycogen Mutant Has Lower Photosynthetic Efficiency When Photosynthesis First Starts

To test our hypothesis that glycogen metabolism can support a higher photosynthetic efficiency when photosynthesis first starts, we measured the PSII efficiency of wild-type and ∆glgC mutant cells using the Walz Phyto PAM II fluorometer. When cyanobacterial cells were grown under continuous light, the maximum PSII efficiency of both the wild type and the ∆glgC mutant were relatively low in the first few hours of growth but gradually reached above 0.4 (Supplemental Fig. S1), consistent with values reported for S. elongatus (Campbell et al., 1998). The effective PSII efficiency (ΦII) exhibited a similar trend, with values ranging from 0.25 to 0.35 in the first 20 h growing under continuous light conditions (Fig. 2A). On the other hand, the ΦII of the ∆glgC mutant was significantly lower than that of wild-type cells in the beginning of the growth phase and only rose to a comparable level at later stages of growth (Fig. 2A). Thus, the differential levels of ΦII between the wild type and the ∆glgC mutant during the lag phase is consistent with our hypothesis.

Figure 2.

Figure 2.

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PSII efficiency of S. elongatus wild-type (WT) and glycogen mutant cells. A, ΦII measured under continuous light. Paired Student’s t test showed a P value of 0.0193 between the wild type and the ΔglgC mutant. B, ΦII in dark-incubated cells. Both ΦII following the dark incubation (0 h) and after light was turned on for 4 h were measured to monitor the ΦII during photosynthesis restart. C, Oxygen evolution rates of cells grown in light/dark (8 h/16 h) cycles. After a few cycles of light/dark growth, the oxygen evolution rates were measured at the end of the dark period (0 h) and after the light was turned on for 2 h. Error bars indicate the sd; two-tailed Student’s t tests were used to compare PSII efficiencies and oxygen evolution rates: *, P < 0.05 and ***, P < 0.001.

To further validate the supportive role of glycogen metabolism in starting photosynthesis, we monitored ΦII following prolonged dark incubation. Cellular respiration in the dark consumes the Calvin-Benson cycle intermediates, gradually lowering or depleting the C6, C5, and C3 sugar intermediates pool (Iijima et al., 2015; Diamond et al., 2017). Without the support from glycogen metabolism, we thus should observe a lower photosynthetic efficiency in ∆glgC cells when photosynthesis restarts after the light cycle resumes. To ensure the depletion of phosphate sugars, we incubated cells in the dark for 30 h. Following dark incubation, the ΦII was significantly lower in the ∆glgC mutant compared with the wild type during the first few hours of light (Fig. 2B).

We further grew cyanobacterial cells under light/dark cycles and measured the whole-cell oxygen evolution rate to further validate the photophysiology results. Both wild-type and ∆glgC mutant cells were grown under a light/dark (8 h/16 h) regime for a few cycles, then the oxygen evolution rates were measured at the end of the dark period and after light was turned on for 2 h. When photosynthesis restarted following dark incubation, the oxygen evolution rate was significantly lower in the ∆glgC mutant compared with the wild type (Fig. 2C), supporting our hypothesis on the role of glycogen metabolism in supporting a rapid start of photosynthesis.

PSI-Mediated Cyclic Electron Flow Is Lower in the Glycogen Mutant

Cyclic electron flow (CEF) around PSI contributes largely to additional ATP requirements in cyanobacterial carbon metabolism (Mullineaux, 2014). We measured the CEF rate through P700 photooxidation under far-red (FR) light (Klughammer and Schreiber, 1994; Cook et al., 2019). Oxidized PSI (P700+) absorbs strongly at wavelengths between 810 and 820 nm (A820), while reduced P700 has minimal absorbance at this range. Both wild-type and ∆glgC mutant cells exhibited a rapid increase in A820 after the FR light was turned on, indicating the oxidation of PSI (P700 to P700+; Supplemental Fig. S2). Once a stable A820 was attained, the FR light was turned off and the rate of rereduction of P700+ to P700 (t1/2red) was calculated as an estimate of P700+ rereduction from alternative electron donors, of which CEF is the major pathway (Yu et al., 1993). We monitored CEF rates in the wild type and the ∆glgC mutant under both dark and light conditions. P700 kinetics were monitored in the log-phase S. elongatus wild type and ∆glgC mutant both at the end of dark incubation and after light was turned on for 2 and 24 h. The ratio ΔA820/A820 was comparable for both strains under all conditions (Fig. 3A), indicating that, unlike PSII, the amount of photooxidizable P700 was not affected by the mutation in the glycogen synthesis pathway. After dark incubation for 16 h, both wild-type and ∆glgC mutant cells exhibited relatively slow t1/2red, suggesting that CEF is low in the cyanobacterial cultures following dark incubation (Fig. 3B; Supplemental Fig. S2). However, the wild-type cells exhibited more than a twofold faster t1/2red after 2 h of incubation in the light. After 24 h in the light, the t1/2red of both the wild type and the ∆glgC mutant recovered to comparable levels (Fig. 3B; Supplemental Fig. S2).

Figure 3.

Figure 3.

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P700+ reduction in S. elongatus wild-type and glycogen mutant cells following dark incubation. A, The oxidizable P700 pool (∆A820/A820). B, t1/2red. Cultures were incubated in the dark for 16 h. The P700+ reduction was measured right after dark adaption (0 h), 2 h into the light (2 h), and 24 h into the light (24 h). The data were plotted from the means of nine replicates (three biological × three technical replicates). Error bars indicate the se; two-tailed Student’s t tests were used to compare the oxidizable P700 pool and P700+ rereduction rates: **, P < 0.01.

Glycogen Metabolism Stimulates Photosynthesis through the OPP Pathway

To determine how glycogen metabolism can initiate photosynthesis more rapidly, we conducted another comparative proteomics study on dark-incubated cyanobacterial cells. Log-phase wild-type and ∆glgC mutant cells were dark incubated for 16 h before the light was turned on. After 2 h in the light, cells were collected for proteomic analysis. Similar to log-phase cells collected under continuous light (Fig. 1), the enzymes for glycogen metabolism (i.e. AGPase [glgC], glycogen synthase [glgA], 1,4-α-glucan-branching enzyme [glgB], and α-1,4-glucan phosphorylase [glgP]) were found in significantly lower abundances in the ∆glgC mutant compared with wild-type cells (Fig. 4A). Interestingly, the enzyme 6-phosphogluconate dehydrogenase (gnd), a key enzyme of the OPP pathway, was also in significantly lower abundance in ∆glgC cells (Fig. 4A). The proteomic analysis indicates that the glycogen degradation could proceed through the OPP pathway.

Figure 4.

Figure 4.

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Glycogen metabolism stimulates photosynthesis through the OPP pathway. A, Brown arrows indicate the proposed mechanism in which glycogen degrades through the OPP pathway to recharge photosynthesis during the start of photosynthesis. Bar graphs indicate the protein levels from proteomic analysis. Error bars indicate the sd. B, Growth profile and P700+ rereduction analysis of the wild type (WT) and the OPP pathway mutant ∆gnd. The bar graph indicates the calculated t1/2red. Error bars indicate the se. C, Light/dark cycle growth profile of the wild type, the ∆glgC mutant, and the ∆gnd mutant. Cells were grown in quadruplet at 30°C with 50 μmol photons m−2 s−1 illumination in the Multi-Cultivator MC-1000-OD-MULTI. Two-tailed Student’s t tests were used to compare protein abundances and P700+ rereduction rates: *, P < 0.05; **, P < 0.01; ***, P < 0.001; and ****, P < 0.0001. 6PGL, 6-Phosphogluconolactone; Ru5P, ribulose-5-phosphate.

To validate the involvement of the OPP pathway in stimulating a rapid photosynthesis start, we generated an OPP pathway mutant strain (Δgnd) by deleting the gnd gene in the S. elongatus genome. The selection of gnd over the G6P dehydrogenase gene (zwf) for the deletion was to minimize the potential impact on other metabolic processes such as the Entner-Doudoroff pathway, which depends on Zwf (Chen et al., 2016). When photosynthesis starts after dark incubation, the Δgnd strain showed impaired growth compared with the wild type (Fig. 4B), indicating a critical role of the OPP pathway in supporting photosynthesis. If the glycogen degradation proceeds through the OPP pathway, a higher ATP/NADPH ratio would be required during carbon fixation (Sharkey and Weise, 2016). We thus measured the P700+ rereduction rate as a proxy for the CEF. The wild-type and Δgnd cells were grown to log phase under moderate light levels (50 µmol photons m−2 s−1) before the light was turned off for 16 h. Following the transition to light, the P700+ rereduction rates of both wild-type and Δgnd cells were slow after dark incubation (Fig. 4B). However, the t1/2red of wild-type cells quickly recovered after 2 h of incubation in the light, whereas the t1/2red of the Δgnd cells was significantly slower (Fig. 4B). Even after 24 h in light, the t1/2red of the Δgnd cells was still relatively slow compared with wild-type cells (Fig. 4B). This observation further supports our hypothesis that the OPP pathway is crucial for the rapid start of photosynthesis after dark incubation.

We further conducted light/dark cycle growth experiments to evaluate the role of glycogen metabolism and the OPP pathway in supporting the rapid start of photosynthesis. The S. elongatus wild type and two mutants (∆glgC and ∆gnd) were grown under two light/dark cycle (12 h/12 h and 8 h/16 h) conditions. Compared with the wild type, ∆glgC and ∆gnd mutants had much slower photosynthesis start after the dark period under both conditions (Fig. 4C). Under the 12-h/12-h light/dark condition, the photosynthesis restart rate of the ∆glgC mutant was lower in the first few light/dark cycles but gradually reached similar levels to the wild type at later cycles of growth. The ∆gnd mutant showed similar growth to the ∆glgC mutant for the first few light/dark cycles but completely abolished its growth afterward, indicating the crucial role of the OPP pathway in the photosynthesis-starting process during diurnal growth. The phenotype was even more obvious in the 8-h/16-h light/dark cycle experiment. Throughout the growth cycle, the ∆glgC mutant had much slower photosynthesis start after each dark period compared with the wild type, suggesting limited availability of intermediates needed for photosynthesis restart after a 16-h dark incubation. On the other hand, the growth of the ∆gnd mutant was completely abolished when growing under the 8-h/16-h light/dark cycles (Fig. 4C). The growth experiment strongly supports our hypothesis that glycogen metabolism stimulates photosynthesis through the OPP pathway during cyanobacterial diurnal growth.

The OPP Pathway Contributes to the Activation of the Calvin-Benson Cycle

Upon restarting of photosynthesis reactions following dark incubation, the intermediate pool needed to regenerate RuBP in the Calvin-Benson cycle may be limited or depleted (Iijima et al., 2015). Continuous supply of RuBP during the dark/light transition is likely critical for maintaining homeostatic balance between energy production by the light reactions and energy consumption by the Calvin-Benson cycle. To evaluate intermediate availability during cyanobacterial diurnal growth, we analyzed the intermediates of central metabolism in the wild type and the ∆glgC mutant grown under light/dark (8 h/16 h) cycles. After 16 h of dark incubation, several intermediates in the Calvin-Benson cycle, including Fru-6-P (F6P), ribose-5-phosphate (R5P), and Ru5P, were found in low abundances in both the wild type and the ∆glgC mutant; however, after short exposure in the light, these intermediates were quickly replenished in the wild type but not in the ∆glgC mutant (Fig. 5A). In the mean time, the glycogen degradation products, G1P and G6P, increased to much higher levels in the wild type compared with the ∆glgC mutant after short exposure in the light (Fig. 5A), suggesting robust glycogen degradation in the wild type to replenish the Calvin-Benson cycle intermediates. Moreover, the level of the carbon fixation product 3PG stayed at similar levels during the dark/light transition period in the wild type, whereas the ∆glgC mutant had slightly decreased levels (Fig. 5A). It is also worth mentioning that 3PG was severely depleted in the ∆glgC mutant compared with the wild type, which is in line with the lower photosynthetic efficiency in the ∆glgC mutant (Fig. 2).

Figure 5.

Figure 5.

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Glycogen metabolism contributes to activation of the Calvin-Benson cycle during the dark/light transition. After a few light/dark (8 h/16 h) cycles, wild-type (WT) and ∆glgC mutant cells were collected right after the dark period (0 h), 30 min into the light (0.5 h), and 2 h into the light (2 h) for the metabolomics analysis (A), PRK activities at 0 and 2 h (B), and NAD(P)H measurements at 0 and 2 h (C). Error bars indicate the sd; *, P < 0.05; **, P < 0.01; ***, P < 0.001; and ****, P < 0.0001.

During the dark/light transition, glycogen degradation through the Embden-Meyerhof-Parnas pathway would form a futile cycle to carbon fixation reactions. The obvious route for glycogen degradation is thus through the OPP pathway to replenish C5 carbon pools. This hypothesis is supported by the observation of increased expression of 6-phosphogluconate dehydrogenase in the wild type relative to the ∆glgC mutant during the start of photosynthesis (Fig. 4A). Ru5P derived from the OPP pathway can then be phosphorylated to RuBP by phosphoribulokinase (PRK), replenishing the starting substrate for Rubisco to continuously fix CO2 in the Calvin-Benson cycle. However, PRK is inactive in the dark and requires redox regulation to become active in the light (Tamoi et al., 2005). To validate whether glycogen degradation through the OPP pathway contributes to the activation of the Calvin-Benson cycle during the dark/light transition, we measured PRK activities in cells grown under light/dark (8 h/16 h) cycles. After a few light/dark cycles, both wild-type and ∆glgC cells were collected at the end of the dark period and when light was turned on for 2 h. After light exposure, PRK activity in the wild type increased significantly whereas the increase was limited in the ∆glgC mutant (Fig. 5B). We further measured the NAD(P)H levels in both the wild type and the ∆glgC mutant. When photosynthesis reactions restart after the dark period, the NADPH level was significantly lower in the ∆glgC mutant compared with the wild type (Fig. 5C). Collectively, these results support our hypothesis that glycogen metabolism helps modulate NADPH levels to activate the Calvin-Benson cycle for a rapid start of photosynthesis.

DISCUSSION

Photosynthetic organisms experience drastic changes in their carbon metabolism during diurnal growth. Recent research has highlighted the contribution of glycogen metabolism and the OPP pathway to the fitness of cyanobacteria during diurnal growth (Diamond et al., 2017; Puszynska and O’Shea, 2017; Welkie et al., 2018). In this study, we provided strong evidence of the active participation of glycogen metabolism and the OPP pathway in supporting photosynthesis during the dark/light transition period of diurnal growth. This supportive role is likely essential for the survival of cyanobacteria in their natural environment.

Glycogen Degradation through the OPP Pathway Serves as the Perfect Buffering System for the Stalled Calvin-Benson Cycle

Cellular respiration from glycogen degradation helps maintain cell viability in the dark. When light resumes, many enzymes for cellular respiration are inactivated, shifting cell metabolism to carbon fixation (Udvardy et al., 1984; Gleason, 1996; Sharkey and Weise, 2016). However, several recent studies suggest that glycogen metabolism plays an additional supportive role for the start of photosynthesis. A glycogen phosphorylase (ΔglgP) mutant of Synechocystis sp. PCC 6803 retards dark respiration of glycogen and had a much lower photosynthetic oxygen evolution rate (Shimakawa et al., 2014). Another study conducted in the ΔglgC mutant of Synechocystis sp. PCC 6803 exhibited a delayed activation of the Calvin-Benson cycle and diminished photochemical efficiency when cultured under high-carbon conditions (Holland et al., 2016). These studies indicate a tight link between glycogen metabolism and photosynthesis. In this study, we showed that glycogen metabolism can stabilize photosynthesis through the OPP pathway in the unicellular cyanobacterium S. elongatus when photosynthesis reactions start upon light (Fig. 4). When cells were grown under light/dark cycles, we found that levels of several intermediates (e.g. F6P, R5P, and Ru5P) in the Calvin-Benson cycle were low after dark respiration (Fig. 5A). The replenishment of these intermediates is essential for RuBP regeneration in the Calvin-Benson cycle. However, due to limited RuBP availability upon photosynthesis restart, newly synthesized triose phosphate would be insufficient to replenish the Calvin-Benson cycle intermediates, leading to stalled carbon fixation reactions. The alternative carbon flow from glycogen degradation through the OPP pathway is thus essential for RuBP regeneration to ensure the continuous carbon fixation. The importance of this alternative carbon flow was shown in the ∆glgC mutant in that cells could not replenish these intermediates efficiently without the glycogen synthesis pathway (Fig. 5A). The importance of this pathway is further manifested by the lack of efficiency to synthesize triose phosphates via phosphofructokinase and phosphoglucoisomerase, both of which require high levels of substrate (Knowles and Plaxton, 2003; Preiser et al., 2018). Glycogen metabolism thus serves as a perfect buffering system to supply G6P for the OPP pathway through glycogen degradation (Gómez-Casati et al., 2003). In addition, our proteomics results showed that transaldolase, a nonessential enzyme in the nonoxidative phase of the pentose phosphate pathway, was also in significantly higher abundance in the wild type compared with the ΔglgC mutant when photosynthesis reactions first start (Supplemental Table S1). Interestingly, a recent genome-wide fitness study in S. elongatus showed a similar result in that the transaldolase mutant was sensitive to light/dark cycles (Welkie et al., 2018). These results suggest an underappreciated role for transaldolase during the operation of the OPP pathway.

Our findings echo with the proposed mechanism by Sharkey and Weise (2016) for the involvement of the OPP pathway to stabilize photosynthesis in plants. They suggested that the OPP pathway (the G6P shunt) could account for 10% to 20% of Rubisco activity and that the enzyme SBPase plays an important role in controlling the carbon flow between the Calvin-Benson cycle and the shunt (Sharkey and Weise, 2016). Their recent evidence showed that the G6P shunt was activated in a photorespiration mutant of Arabidopsis (Arabidopsis thaliana) in which the activity of triose phosphate isomerase (TPI) was inhibited (Li et al., 2019). TPI is responsible for interconverting G3P and dihydroxyacetone phosphate, a key reaction for RuBP regeneration in the Calvin-Benson cycle. During photosynthesis, it is critical that RuBP can be continuously regenerated for CO2 fixation, which limits the carbon regeneration reactions to a small margin of error. Perturbation to carbon regeneration reactions (e.g. TPI inhibition) would lead to a temporarily impaired Calvin-Benson cycle. An alternative mechanism such as the G6P shunt is thus essential to stabilize photosynthesis.

Glycogen Metabolism Is Critical for the Optimal Photosynthesis Performance

Glycogen metabolism and the OPP pathway are critical for optimal photosynthetic performance during the start of photosynthesis. This can be observed through our photochemistry measurements of the wild type and the ∆glgC mutant. Compared with the wild type, the photosynthetic efficiency of the ∆glgC mutant was significantly lower when photosynthesis starts (Fig. 2), suggesting that PSII is likely down-regulated through a feedback mechanism in the ∆glgC mutant. The temporally impaired PSII in the ∆glgC mutant could be partially explained by the proteomics data. Proteomic analysis of the dark-adapted wild type and ∆glgC mutant showed differential expression of several proteins associated with PSII (Supplemental Table S1). Most notably, the abundance of the ATP-dependent zinc metalloprotease FtsH was over 2-fold higher in the ∆glgC mutant in comparison with the wild type. In both plants and cyanobacteria, the FtsH protease plays important roles in repairing damaged PSII by degrading the D1 protein (Lindahl et al., 2000; Silva et al., 2003). Increased FtsH abundance in the ∆glgC mutant suggests an active PSII repair cycle when light was turned on. Without the support from glycogen metabolism, the photosynthetic electron transport chain is likely overreduced due to an imbalance of energy production and consumption, leading to PSII photodamage in the ∆glgC mutant. This hypothesis is supported by the finding of additional key proteins involved in PSII repair. Both carotene isomerase and the protein Psb28 were in higher abundance in the ∆glgC mutant compared with the wild type (Supplemental Table S1). Carotene isomerase is an important enzyme for carotenoid synthesis. Past studies in cyanobacteria have shown the active participation of carotenoids during PSII assembly (Masamoto et al., 2004; Tóth et al., 2015). Moreover, Psb28 was found to be involved in protecting the PSII RC47 assembly intermediate during its conversion to functional PSII in Synechocystis sp. PCC 6803 (Dobáková et al., 2009; Weisz et al., 2017).

For PSI performance, we applied P700+ rereduction rates to evaluate the alternative electron flows returning to PSI as a proxy for CEF. When light was turned on following dark incubation, we observed different P700+ rereduction rates in the wild type compared with both mutants (∆glgC and ∆gnd). Both wild-type and ∆glgC mutant cells had comparably slow P700+ rereduction rates after dark incubation; however, the wild type exhibited rapid recovery within 2 h in the light, while it took 24 h for the ∆glgC mutant to reach a comparable level to the wild-type cells (Fig. 3). When comparing the wild type and the ∆gnd mutant, the P700+ rereduction rate showed a similar trend (i.e. a much faster rereduction rate in the wild type after a short incubation in the light; Fig. 4B). This strongly suggests a higher ATP/NADPH ratio requirement when photosynthesis first starts and is directly linked to glycogen metabolism and the OPP pathway. The action of the OPP pathway alongside the Calvin-Benson cycle exacerbates the deficit of ATP/NADPH ratio needed for carbon fixation (Sharkey and Weise, 2016). It is well accepted that the additional ATP requirement in photosynthetic carbon metabolism can be fulfilled through the CEF (Kramer and Evans, 2011). The higher CEF rate observed in the wild-type cells fits well with the additional ATP requirements for operating the OPP pathway during the start of photosynthesis. Unlike the ΔglgC mutant, the ∆gnd mutant did not recover to a comparable level of P700+ rereduction rate even after 24 h in the light, indicating that the OPP pathway is essential for the initiation of photosynthesis and the fitness of cells (Fig. 4B). The additional light/dark cycle growth experiments further showed the importance of the OPP pathway in supporting the start of photosynthesis. When growing under 12-h/12-h light/dark cycles, the ∆gnd mutant showed similar growth to the ∆glgC mutant during the first few cycles but stopped growing afterward. On the other hand, growth of the ∆gnd mutant was completely abolished under 8-h/16-h light/dark growth cycles (Fig. 4C), suggesting depletion of the Calvin-Benson cycle intermediates during long periods of darkness and the necessity of the OPP pathway to start photosynthesis.

Glycogen Metabolism Participates in Activation of the Calvin-Benson Cycle during Cyanobacterial Diurnal Growth

Glycogen degradation through the OPP pathway could replenish RuBP to ensure the continuous operation of carbon fixation during the start of photosynthesis. However, it is important to know that the OPP pathway does not lead to any net carbon fixation. CO2 fixed in the Calvin-Benson cycle is quickly lost when G6P is oxidized to Ru5P in the shunt. It is thus intriguing to understand the role of the OPP pathway in supporting photosynthesis. We propose that the OPP pathway could generate additional NADPH required for the redox regulation of the Calvin-Benson cycle during the start of photosynthesis. Previous research showed that the activities of two key enzymes in the Calvin-Benson cycle, PRK and G3P dehydrogenase (GADPH), are suppressed in the dark through the formation of a CP12/PRK/GAPDH protein complex (Tamoi et al., 2005). The reversible dissociation of the complex was controlled by the pyrimidine nucleotide levels in S. elongatus (Tamoi et al., 2005). The operation of the OPP pathway during the start of photosynthesis thus could contribute largely to the increase of NADPH levels when light resumes, releasing PRK and GAPDH from the complex to activate the Calvin-Benson cycle. This hypothesis is supported by the measurements of both PRK activities and NADP(H) levels during the start of photosynthesis. Following light/dark (8 h/16 h) cycles, both PRK activities and NADPH levels were significantly higher in the wild type relative to the ∆glgC mutant after the light was turned on (Fig. 5, B and C). However, NAPDH generation from the OPP pathway relative to that from the light reactions is worthy of further investigation.

Lastly, it is also important to understand the activation mechanism of the OPP pathway. The expression of many genes in glycogen metabolism and the OPP pathway are controlled by the circadian clock output protein RpaA (Markson et al., 2013; Welkie et al., 2018). Our proteomic analysis confirmed that many enzymes in these two metabolic processes were in much higher abundance in wild-type cells in the beginning of photosynthesis (Fig. 4A; Supplemental Table S1). Previous research also reported that the first enzyme in the OPP pathway, G6P dehydrogenase, is redox regulated and is inactive in its reduced form (Udvardy et al., 1984; Gleason, 1996). However, G6P could also stabilize and activate G6P dehydrogenase to reverse the reduction inhibition in cyanobacteria (Cossar et al., 1984). It is thus likely that glycogen degradation would lead to increased G6P levels and activate the G6P dehydrogenase. Our metabolite analysis supports this hypothesis in that G6P levels quickly increased during the dark/light transition period in the wild type compared with the ∆glgC mutant (Fig. 5A). Future studies toward understanding these mechanisms could help fully appreciate the roles played by the OPP pathway in supporting photosynthesis.

In conclusion, we found that glycogen metabolism in cyanobacteria helps activate photosynthesis through the OPP pathway during the start of photosynthesis reactions. The OPP pathway is generally considered for its role to generate reducing equivalents for dark respiration. However, evolution empowers cells with the ability to leverage existing metabolic processes for the adaptation of constantly changing environments. We have witnessed an interesting strategy implemented by cyanobacteria to ensure their growth advantage in their natural environment.

MATERIALS AND METHODS

Growth Conditions

Seed cultures of Synechococcus elongatus wild-type and mutant strains were grown in BG11 medium supplemented with 20 mm sodium bicarbonate and 10 mm TES (pH 8.2) at 30°C under 30 μmol photons m−2 s−1 illumination. The growth medium of the mutant strains was also supplemented with 5 mg L−1 kanamycin. Cultures for measurements were grown at 30°C under 50 μmol photons m−2 s−1 illumination in the Multi-Cultivator MC-1000-OD-MULTI (Photon Systems Instruments). Cyanobacterial growth was monitored using the built-in spectrophotometer of Multi-Cultivator by measuring OD720. S. elongatus wild-type and ΔglgC mutant cells used for oxygen evolution measurement, metabolite analysis, PRK enzyme activity, and NADP(H) levels were grown in triplicate under light/dark (8 h/16 h) cycles at 30°C with 50 μmol photons m−2 s−1 illumination in the Multi-Cultivator MC-1000-OD-MULTI.

Plasmid and Strain Construction

The gene fragments for plasmid construction were amplified using primers designed through the NEBuilder Assembly tool (New England Biolabs). The glgC and gnd knockout plasmids were constructed using HiFi DNA Assembly master mix (New England Biolabs). S. elongatus wild-type cells were transformed with knockout plasmids to generate ΔglgC and Δgnd mutant strains. The strains and plasmids along with primers used in this study are listed in Supplemental Tables S2 and S3 and Supplemental Data S1 and S2.

Proteomics

Log-phase S. elongatus wild-type and ΔglgC mutant cells were used for proteomic analysis following our previously described method (Wang et al., 2016). For dark-incubated samples, log-phase cells were taken after light was turned on for 2 h following dark incubation. Briefly, biological triplicates of approximately 10 OD of cells were used for protein extraction. Cell pellets were resuspended in 1 mL of Tris buffer (50 mm Tris-HCl, 10 mm CaCl2, and 0.1% [w/v] n-dodecyl β-d-maltoside, pH 7.6) followed by cell lysis using a homogenizer with bead-beating cycles of 10 s on and 2 min off on ice for a total of six cycles. Protein extract in the supernatant was collected by centrifugation at 13,000g for 30 min at 4°C, followed by protein concentration measurement using the Bradford assay (Thermo Scientific).

For each sample, 100 µg of total protein was digested by Trypsin Gold (Promega) with a 1:100 (w/w) ratio at 37°C for 18 h. The digested peptides were cleaned up using a Sep-Pak C18 column (Waters) followed by peptide fractionation using the Pierce High pH Reverse-Phase Peptide Fractionation Kit (Thermo Scientific). Eight fractions of peptides from each sample were subjected to liquid chromatography-tandem mass spectrometry (MS/MS) analysis in a Thermo LTQ Orbitrap XL mass spectrometer. The mass spectrometer was operated under the data-dependent mode scanning the mass range of 350 to 1,800 mass-to-charge ratio at a resolution of 30,000. The 12 most abundant peaks were subjected to MS/MS analysis by collision-induced dissociation fragmentation. The raw data collected from MS/MS analysis was searched and analyzed using the pipeline programs integrated in the PatternLab for Proteomics (version 4.1.0.17; Carvalho et al., 2016). The normalized spectral abundance factor was used to compare protein abundance in different groups (Zybailov et al., 2006). The mass spectrometry proteomics data have been deposited to both the MassIVE repository with the data set identifier MSV000083575 and the ProteomeXchange Consortium (Vizcaíno et al., 2016) with the data set identifier PXD013099.

PSII Analysis

Cyanobacterial cells grown under continuous light (50 μmol photons m−2 s−1) and dark-incubated cells were subjected to PSII measurement using the Phyto-PAM Phytoplankton Analyzer (Heinz Walz). When measuring PSII efficiency, the dark adaption that normally oxidizes the PSII center in green algae does not work in S. elongatus. Instead, cells would transit to state II after dark adaptation (Campbell et al., 1998). To get a proper measurement of PSII efficiency, cells were illuminated at moderate actinic light (25 µmol m−2 s−1) for 5 min to lock the photosystems in state I and to oxidize PSII reaction centers. The blue light was used as the measuring light to minimize the background fluorescence from phycobilisome light absorption. Standard induction curves by exposing cells to saturation pulses were recorded to determine the maximum PSII efficiency and the ΦII.

Oxygen Evolution

S. elongatus wild-type and ΔglgC mutant cells were collected following a few light/dark cycles. Two milliliters of cells was harvested both at the end of the dark period and after light was turned on for 2 h. Fifty microliters of cells were used for cell counting in an Attune NxT Flow Cytometer (Invitrogen). After a short incubation in the dark, whole-cell oxygen evolution was measured for 2 min at room temperature with saturation light illumination in a Clark-type oxygen electrode (Hansatech Instruments). Since the growth medium was supplemented with 20 mm NaHCO3, no additional bicarbonate was added in the electrode chamber. The final oxygen evolution rate was corrected with the dark respiration rate and normalized to cell numbers.

PSI Analysis

Log-phase S. elongatus wild-type and mutant cells were incubated in the dark for 16 h before the PSI analysis. Five milliliters of the culture was vacuum filtered through Whatman GF/C 25-mm filter discs (catalog no. 1822-025). FR light-induced P700 oxidation-reduction kinetics measurements were performed using a dual-wavelength pulse amplitude-modulated fluorescence monitoring system (Dual-PAM-100; Heinz Walz) with a leaf attachment. The proportion of photooxidizable P700 (ΔA820/A820) was determined as the change in A820 after turning on the FR light (λmax = 715 nm, 10 W m−2; Scott filter RG 715). The t1/2red was calculated after the FR light was turned off and used as an estimate of alternative electron flow around PSI.

Quantification of Central Metabolites

The metabolite analysis was conducted at the West Coast Metabolomics Center at the University of California, Davis. Ten milliliters of S. elongatus wild-type and ΔglgC cells was quenched in 15 mL of cold phosphate-buffered saline (PBS), followed by centrifugation at 7,800g for 15 min at 4°C to collect the cell pellet. Intermediates of the primary metabolism were analyzed by gas chromatography-time of flight-mass spectrometry and identified using the BinBase algorithm (Fiehn et al., 2008). The metabolite levels were normalized to cell numbers and used for comparison.

PRK Enzyme Assay

Following a few light/dark cycles, 10 mL of S. elongatus wild-type and ΔglgC mutant cells was harvested by injecting into 15 mL of cold PBS both at the end of the dark period and after light was on for 2 h. Cells pellets were collected by centrifugation at 7,800g for 15 min at 4°C, followed by resuspension in 500 μL of 50 mm Tris-HCl (pH 8). Cells were then disrupted in a bead beater for a total of 1 min with 10 s on and 2 min off on ice. The supernatant was collected by centrifugation at 15,000 rpm for 20 min at 4°C. An aliquot of the supernatant was used to determine the protein concentration by the Bradford assay (Thermo Scientific). The PRK assay was carried out by monitoring NADH A340 on a 96-well plate by modifying a previously described method (Kanno et al., 2017). Briefly, cell lysate was preincubated at 30°C for 30 min to obtain the baseline absorbance. After the incubation, an enzyme mixture containing 0.5 mm Ru5P, 2 mm ATP, 2.5 mm phosphoenolpyruvate, 10 mm MgCl2, 0.3 mm NADH, 4 units of pyruvate kinase, and 4 units of lactate dehydrogenase was added to the cell lysate to make the final reaction mixture to 100 μL. NADH absorbance was recorded every 1 min for a total of 30 min. The ∆340 nm/min was not corrected to cuvette absorbance for enzyme unit calculation. Specific PRK activity was obtained by normalizing the rate to total protein levels of each sample.

Measurement of NAD(P)H Levels

Following a few light/dark cycles, 10 mL of S. elongatus wild-type and ΔglgC cells was collected both at the end of the dark period and after light was turned on for 2 h. Cells were quickly transferred to 15 mL of cold PBS after collection, followed by centrifugation at 7,800g for 15 min at 4°C. Cell pellets were resuspended in 500 μL of PBS and lysed by the addition of 500 μL of base solution (0.2 n NaOH) with 1% (w/v) dodecyltrimethylammonium bromide. Following centrifugation at 15,000 rpm for 20 min at 4°C, the supernatant was collected and passed through a Pierce Concentrator spin column (catalog no. PI88513; Fisher Scientific) to remove enzymes that could affect the NADP(H) assay. The NADP(H) levels in the flow-throughs were determined by the NADP/NADPH-Glo assay (catalog no. G9071; Promega) following the manufacturer’s instructions. Fifty microliters of cells was also collected for cell counting in an Attune NxT Flow Cytometer (Invitrogen). The amount of NADP(H) in each sample was calculated from a NADPH standard curve and normalized to cell numbers.

Statistical Analyses

The statistical analyses for all experiments were performed using the software Prism. Two-tailed Student’s t tests were used to compare wild-type and mutant cells for differences in protein abundances, PSII and PSI efficiencies, oxygen evolution rates, PRK enzyme activities, and NADP(H) levels. One-way ANOVA was used to analyze the temporal changes of metabolite abundance in the wild type and the ΔglgC mutant during the start of photosynthesis. A value of P < 0.05 between comparing groups was considered statistically different.

Supplemental Data

The following supplemental materials are available.

Acknowledgments

We thank Spencer Diamond (University of California, Berkeley), James Golden, and Susan Golden (University of California, San Diego) for the original plasmid used to generate the glycogen mutant strain in the preliminary study. We thank Theresa Ramelot and Kundi Yang of the Chemistry Department at Miami University for their help in the preliminary metabolite analysis. We thank the Center for Bioinformatics and Functional Genomics at Miami University for providing access to several instruments used in this study. We thank Jianping Yu (U.S. National Renewable Energy Laboratory) and Graham Peers (Colorado State University) for careful reading of the article.

Footnotes

1

This work was supported by Miami University startup funds to X.W. and by Department of Energy-BES grant DE-SC0019138 to X.W. and R.M.M.-K.

[OPEN]

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