Introduction

Cyanobacteria perform fermentation, using glycolytic products as electron acceptors. (1). Cyanobacteria have multiple fermentation pathways according to the environment. For example, the freshwater living cyanobacterium Synechocystis sp. PCC 6803 (hereafter referred to as Synechocystis) produces acetate, lactate, dicarboxylic acids, and hydrogen (1, 2).

Hydrogen is generated in quantities comparable to lactate and dicarboxylic acids as the result of electron acceptance in the dark microoxic condition (3, 4). Bidirectional hydrogenase is a key enzyme for H₂ production from protons (5) and is commonly found in cyanobacteria (6). Cyanobacterial hydrogenase comprises five subunits (HoxEFUHY) containing nickel and Fe-S clusters (7). This enzyme can utilize NADH, reduced ferredoxin, and flavodoxin as substrates (8). Hydrogenase mainly receives reduced ferredoxin from pyruvate-ferredoxin oxidoreductase (PFOR) in the microoxic condition (8, 9). Although hydrogenase and PFOR are O₂ sensitive, they can work under aerobic conditions (10–12). Therefore, uncontrolled expression of hox operon and nifJ (coding gene of PFOR) may hamper metabolism under photosynthetic conditions. Furthermore, genetic manipulations on Synechocystis have demonstrated that modulating the expression of certain enzymes including hydrogenase can alter fermentative metabolic flow (3, 4, 13). This provides evidence that transcription regulates the fermentative pathway. Thus, transcriptional regulation in response to the environment is essential for optimal energy cost performance.

Promoter recognition by RNA polymerases is an essential step in transcriptional regulation. Sigma factors, subunits of RNA polymerase, recognize core promoter sequences. Transcription factors can also bind to promoter regions to suppress or promote RNA polymerase transcription. As well as recruitment or blocking of RNA polymerase, some transcriptional factors, known as Nucleoid-associated proteins (NAPs), modulate chromosomal conformation to regulate transcription (14). NAPs are common in bacteria, but cyanobacterial NAPs remain unidentified, and higher-order DNA structure in cyanobacteria is yet to be shown. A recent study suggested that the manipulation of chromosomal supercoiling impacts transcriptional properties in cyanobacteria (15). There is room for consideration of NAPs modulating chromosomal conformation and regulating expression in cyanobacteria.

In Synechosysits, the coding genes of HoxEFUHY form a single operon (sll1220–1226), while PFOR is encoded in the nifJ (sll0741) gene. Both hox and nifJ operons are highly expressed under microoxic conditions (16). Genetic analysis has revealed that multiple global transcriptional regulators control hox and nifJ expression. Sigma factor SigE (Sll1689) promotes the expression of hox and nifJ operons (17, 18), while transcription factor cyAbrB2 (Sll0822) represses them (19, 20). Positive regulators for the hox operon include LexA (Sll1626) and cyAbrB1 (Sll0359) (21–23).

SigE, an alternative sigma factor, controls the expression of genes involved in glycogen catabolism and glycolysis, as well as PFOR/nifJ and hydrogenase (17). SigE shows a high amino acid sequence similarity with the primary sigma factor SigA, which is responsible for transcribing housekeeping and photosynthetic genes (24). A ChIP-seq study revealed that, while most SigE-binding sites are the same as SigA, SigE exclusively occupies the promoters of glycogen catabolism and glycolysis (25).

CyAbrB2 and its homolog cyAbrB1 are transcription factors highly conserved in cyanobacteria. For example, cyAbrB homologs in Anabaena sp. PCC7120 is involved in heterocyst formation (26). CyAbrB2 in Synechocystis regulates the expression of several genes involved in carbon metabolism, nitrogen metabolism, and cell division (20, 27, 28). CyAbrB2 binds to the hox promoter in vitro and represses its expression in vivo (19). CyAbrB1, an essential gene, physically interacts with the cyAbrB2 protein (29) and binds the hox promotor region in vitro to promote its expression (21).

To explore the dynamics of those transcription factors governing the expression of hox operon, we conducted a time-course analysis of the transcriptome and ChIP-seq of SigE and cyAbrB2. Our ChIP-seq and transcriptome analysis showed the NAPs-like nature of cyAbrB2, which prompted us to conduct a chromosomal conformation capture assay. 3C analysis explored the physical interaction between the hox promoter region and its downstream and upstream genomic region in the aerobic condition, and some loci changed interaction frequency upon entry to the microoxic condition. Furthermore, some interactions in the ΔcyabrB2 mutant were different from those of the wildtype. From those experiments, we propose that cyAbrB2 modulates chromosomal conformation like NAPs, allowing access to the SigE-containing RNA polymerase complex on the hox promoter, by which the hox operon achieves distinct expression dynamics. Chromosomal conformation of bacteria is a growing area of interest, and the findings of this study have brought insight into the transcriptional regulation of cyanobacteria.

Results

Transcriptomes on entry to dark microoxic conditions

To understand transcriptional regulation under microoxic conditions, we conducted a time-course transcriptome capturing light aerobic and dark microoxic conditions at 1-, 2-, and 4-hours time points (Figure 1A). Gene set enrichment analysis based on KEGG pathway revealed that many biological pathways, including photosynthesis and respiration (oxidative phosphorylation), were downregulated by the transition to dark microoxic conditions from light aerobic conditions (Figure 1B). Upregulated pathways included butanoate metabolism and two-component systems. The enrichment in the butanoate metabolism pathway indicates the upregulation of genes involved in carbohydrate metabolism. We further classified genes according to their expression dynamics. Within 1 hour of switching from aerobic to microoxic conditions, the expression levels of 508 genes increased more than 2-fold. Furthermore, genes with increased expression levels were classified into four groups based on the time course (Figures 1C and S1). Of the 508 genes, 28 were termed “transiently upregulated genes” due to their decreased expression upon the comparison of 1- and 4-hours incubation under microoxic conditions (log2 fold change < −0.5), and 119 were termed “continuously upregulated genes.”, which continuously increased between 1- and 4-hours incubation under microoxic conditions (log2-fold change > 0.5). Other than 508 genes 2-fold upregulated within 1 hour, 28 genes showed more than 4-fold increment within 4 hours but not upregulated within 1 hour. We combined those “Late upregulated genes” with 508 genes and referred to as “All upregulated genes” in the subsequent analysis (Figure S1). Mapping the classified genes to central carbon metabolism revealed that nifJ encoding PFOR and hox operon encoding a bidirectional hydrogenase complex were transiently upregulated (Figure 1D and Table 1). RTqPCR verified the transient expression of hoxF, hoxH, and nifJ (Figure S2).

Figure 1

Table 1

SigE and cyAbrB2 control the expression of transiently upregulated genes

The functional correlation between hydrogenase and PFOR, encoded by the hox operon and nifJ, suggests that transient upregulation has physiological significance. We focused on transiently upregulated genes and attempted to reveal the regulatory mechanism underlying transient upregulation. While SigE promotes the expression of hox and nifJ, cyAbrB2 represses them (17, 19, 20). We confirmed that the deletion of sigE and cyabrb2 (ΔsigE and Δcyabrb2, respectively) affected the expression of hoxF, hoxH, and nifJ by RT-qPCR (Figure S1). Thus, we conducted a time-course transcriptome analysis of ΔsigE and Δcyabrb2 under aerobic conditions and after 1- and 2-hours cultivation in dark microoxic conditions (Figures 2A and S3). The transcriptome data confirmed that SigE and cyAbrB2 regulated hox operon expression (Figure 2B). At each time point, we searched for differentially expressed genes (DEGs) between mutants and wildtype with a more than 2-fold expression change and a false discovery rate (FDR) less than 0.05. We found that deleting sigE or cyabrb2 preferentially affected the expression of transiently upregulated genes, not limited to hox and nifJ operons (Figures 2C and 2D). Interestingly, cyabrb2 deletion resulted in the upregulated expression of transient genes under aerobic conditions, in contrast to 1-hour cultivation under microoxic conditions (Figure 2C).

Figure 2

Genome-wide analysis of cyAbrB2, cyAbrB1, and SigE via ChIP-seq

To decipher the regulatory mechanism of transiently upregulated genes, we must first comprehend the fundamental features and functions of these transcriptional regulators. Therefore, a genome-wide survey of cyAbrB2 and SigE occupation (Figure S4) combined with transcriptome data was done. Specifically, we generated a Synechocystis strain in which cyAbrB2 was epitope-tagged and performed a ChIP-seq assay under aerobic and microoxic conditions (Figure S4A). SigE-tagged strains previously constructed and analyzed elsewhere were also employed (25). The primary sigma factor SigA was also analyzed to determine SigE-specific binding. In addition to cyAbrB2, we tagged and analyzed cyAbrB1, which is the interactor of cyAbrB2 and positively regulates the hox operon.

CyAbrB2 binds to long tracts of the genomic region and suppresses genes in the binding region

The ChIP-seq data showed that cyAbrB2 bound to long tracts of the genomic region with lower GC content than the whole genome Synechocystis (Figures 3A and 3B). Vice versa, regions exhibiting lower GC contents showed a greater binding signal of cyAbrB2 (Figure 3C). This correlation was not a systematic bias of next-generation sequencing because the binding signals of SigE, SigA, and control showed no negative correlation to GC contents (Figures S5A and S5B). The binding regions of cyAbrB2 called by peak-caller covered 15.7% of the entire genome length. 805 of 3614 genes overlapped with cyAbrB2 binding regions, and almost half (399 of 805 genes) were entirely covered by cyAbrB2 binding regions. The cyAbrB2 binding regions included 80 of 125 insertion sequence elements (Figure 3D). Comparison with the transcriptome of ΔcyabrB2 revealed that cyAbrB2 tended to suppress the genes overlapping with its binding regions under aerobic conditions (Figures 3A and 3E). A survey of the genomic localization of cyAbrB1 under aerobic conditions revealed that cyAbrB1 and cyAbrB2 shared similar binding patterns (Figures 3A and S5C). Due to the essentiality of cyAbrB1, we did not perform transcriptome analysis on a cyAbrB1-disrupted strain. Instead, we referred to the recent study performing transcriptome of cyAbrB1 knockdown strain, whose cyAbrB1 protein amount drops by half (30). Among 24 genes induced by cyAbrB1 knockdown, 12 genes are differentially downregulated genes in cyabrb2Δ in our study (Figure S5D).

Figure 3

CyAbrB2 binds to transiently upregulated genes

The binding regions of cyAbrB2 overlapped 17 of 28 transiently upregulated genes, showing significant enrichment from all upregulated genes (Figure 4A). The transiently upregulated genes belong to 17 transcriptional units (TUs), according to the previous study(31), and cyAbrB2 tends to bind TUs with transiently upregulated genes (Figure 4B). While cyAbrB2 covered the entire length of insertion sequences and unknown proteins, its binding positions on other transient genes were diverse (Figure 4C). Specifically, the hox and nifJ operons had two distinct binding regions located at the transcription start sites (TSSs) and middle of operons, the pntAB operon had two binding regions in the middle and downstream of the operon, and the nrtABCD operon had one binding region downstream of the operon (Figure 4C).

Figure 4

Localization of cyAbrB2 became blurry under the microoxic condition

When cells entered microoxic conditions, the relative ChIP-seq signals in the cyAbrB2 binding regions slightly declined (Figures 5A and 5B). Notably, the total quantities of precipitated DNA by tagged cyAbrB2 did not decrease (Figure S6A), and qPCR confirmed that the cyAbrB2 binding signal increased in all positions tested (Figure 5C). ChIP-seq data and ChIP-qPCR data indicate that the boundary between cyAbrB2 binding region and cyAbrB2-free region became obscured when the cells entered microoxic conditions due to increased binding of cyAbrB2 to both cyAbrB2 binding and cyAbrB2-free region. The protein amount of cyAbrB2 was not altered on entry to the microoxic condition (Figure S6B). The cyAbrB2 binding signal around the transiently upregulated genes became less specific upon entry into microoxic conditions, consistent with the general tendency (Figure 5B). The amount of DNA immunoprecipitated by cyAbrB1 was also increased in the microoxic condition, and the protein amount was not increased (Figures S6C and S6D).

Figure 5

Sigma factors SigE and SigA are excluded from cyAbrB2 binding regions regardless of GC contents

We searched for SigE and SigA binding sites under aerobic and microoxic conditions (Figures S7A and S7B, left and right, respectively). The SigE and SigA peaks identified in this study predominantly covered the previously identified peaks (Figure S7C), reproducing the previous study’s conclusion (25); i.e., SigE and the primary sigma factor SigA share localization on the promoters of housekeeping genes, but SigE exclusively binds to the promoters of its dependent genes. SigE and SigA binding peaks were significantly excluded from the cyAbrB2 binding regions (Figures 6A and 6B). SigE preferred the cyAbrB2-free region in the aerobic condition more than SigA did (Odds ratios of SigE and SigA being in the cyAbrB2-free region were 4.88 and 2.74, respectively). CyAbrB2 prefers AT-rich regions, but no correlation was found between the GC content and binding signals of SigE and SigA (Figures S5A and S5B). Thus, SigA and SigE are excluded from cyAbrB2 binding regions regardless of GC contents.

Figure 6

Dynamics of sigma factors upon exposure to the microoxic condition

When cells entered microoxic conditions, the binding signals of SigA and SigE were changed, although most of their peaks observed under aerobic conditions were present under microoxic conditions (Figure S7A). The preference of SigE for the cyAbrB2-free region was alleviated in the microoxic condition (Figure 6A). Next, we focused on sigma factor dynamics in transiently upregulated genes. SigE, but not SigA, binds at the TSS of pntAB under aerobic and microoxic conditions (Figure 6C top). SigE binding summits were not identified at the TSSs of the hox and nifJ operons under aerobic conditions. However, the SigE-specific binding summit appeared at the TSS of nifJ when cells entered microoxic conditions (Figure 6C middle). A bimodal peak of SigE was observed at the TSS of the hox operon in a microoxic-specific manner (Figure 6C bottom panel). The downstream side of the bimodal SigE peak coincides with the SigA peak and the TSS of TU1715. Another side of the bimodal peak lacked SigA binding and was located at the TSS of the hox operon (marked with an arrow in Figure 6C), although the peak caller failed to recognize it as a peak. SigE binding without SigA on the promoters of hox, nifj, and pntAB is consistent with their SigE-dependent expression (Figure 2B).

Chromatin conformation around hox operon and nifJ operon

We have shown that cyAbrB2 broadly binds to AT-rich genomic regions, including insertion element sequences, and represses expression (Figure 3). This is functionally similar to the NAPs (14), which makes us hypothesize that cyAbrB2 modulates chromosomal conformation. Therefore, we conducted the chromatin conformation capture (3C) assay against wildtype and cyabrb2Δ strains at aerobic and microoxic conditions. qPCR was performed with unidirectional primer sets, where the genomic fragment containing hox operon and nif operon (hereinafter hox fragment and nifJ fragment, respectively) were used as bait (Figure 7).

Figure 7

First, focusing on the aerobic condition of wildtype (Figure 7B, solid line), the hox fragment interacted with its proximal downstream loci (loci (f) to (g)) and proximal upstream locus (locus (j)). The hox fragment also interacts with the distal downstream locus (locus (c)). Meanwhile, the nifJ fragment shows high interaction frequency with proximal upstream and downstream loci (Figure 7G, loci (i’) and (j’)), and a distal downstream locus (locus (g’)) showed higher interaction frequency with nifJ fragment than proximal locus (h’) did. The upstream regions of nifJ (loci (l’) to (n’) and (p’)) showed comparable frequency with locus (g’).

The chromatin conformation is changed in cyabrb2Δ in some loci

Then we compared the chromatin conformation of wildtype and cyabrb2Δ. Although overall shapes of graphs did not differ, some differences were observed in wildtype and cyabrb2Δ (Figures 7B and 7G); interaction of locus (c) with hox region were slightly lower in cyabrb2Δ and interaction of loci (f’) and (g’) with nifJ region were different in wildtype and cyabrb2Δ. Note that the interaction scores exhibit considerable variability and we could not detect statistical significance at those loci.

Changes of chromatin conformation upon microoxic condition

When the cells entered the microoxic condition, proximal loci interacted more frequently (Figure 7D; loci (f)-(h) and Figure 7I loci (j’) and (k’)). This tendency was more apparent in cyabrb2Δ (Figures 7E and 7J). Furthermore, the interaction of nifJ upstream loci (l’) – (n’) increased in the microoxic condition in cyabrb2Δ but not wildtype (Figures 7I and 7J). The locus (c) and locus (j) interacted less frequently with hox fragment upon entry to the microoxic condition in the wildtype. While the interaction scores exhibit considerable variability, the individual data over time demonstrate declining trends of the wildtype at locus (c) and (j) (Figure S8). In Δcyabrb2, by contrast, the interaction frequency of loci (c) and (j) was unchanged in the aerobic and microoxic conditions (Figure 7E). The interaction frequency of locus (c) in Δcyabrb2 was as low as that in the microoxic condition of wildtype, while that of locus (j) in Δcyabrb2 was as high as that in the aerobic condition of wildtype (Figures 7B and 7C). In summary, 3C analysis demonstrated cyAbrB2-dependent and independent dynamics of chromosomal conformation around the hox and nifJ operon in response to the microoxic condition (Figure 8).

Figure 8

Discussion

Physiological significance of transient upregulation of hox and nifJ operons

As the transcriptional change can alter the metabolic flow, the transcriptional upregulation of fermentative genes in response to the microoxic condition is expected to be adaptive for energy acquisition and the maintenance of redox balance. Our time-course transcriptome showed upregulation of several genes involved in catabolism upon exposure to the microoxic condition. The transient upregulation of hox and nifJ operons is distinctive among them (Figure 1D).

One reason for transient upregulation is probably the resource constraints of inorganic cofactors. Hydrogenase and PFOR (the product of nifJ gene) have iron-sulfur clusters, and hydrogenase requires nickel for its activity (32, 33). Overexpression of the hox operon should be futile under physiological conditions without an adequate nickel supply (34).

Another significance for transient upregulation may be the reusability of fermentative products. Hydrogen, lactate, and dicarboxylic acids can be reused as the source of reducing power when cells return to aerobic conditions (12, 35, 36). The substrate proton is abundant, but hydrogen is diffusive and difficult to store. Therefore, hydrogenase may favor fermentation initiation, and the reductive branch of TCA-producing dicarboxylic acids may become active subsequently. In fact, citH/mdh (sll0891) encoding a key enzyme of the reductive branch of TCA was classified as continuously upregulated genes in this study (Figures 1C and 1D).

Mechanisms for transient expression mediated by SigE and cyAbrB2

SigE and cyAbrB2 can independently contribute to the transient transcriptional upregulation. This is evident as the single mutants, ΔsigE or Δcyabrb2, maintained transient expression of hoxF and nifJ (Figure S2). We first discuss cyAbrB2 as the potential NAPs, and then the mechanism of transient upregulation mediated by cyAbrB2 and SigE will be discussed.

cyAbrB2 is a novel nucleoid-associated protein of cyanobacteria

We have shown that cyAbrB2 broadly binds to AT-rich genomic regions, including IS elements (Figure 3). This is functionally similar to the histone-like nucleoid protein H-NS family; including H-NS in Enterobacteriaceae (37, 38), and Lsr2 in Mycobacteria (39). Like H-NS and Lsr2, cyAbrB2 may defend against exogenous DNA elements, which often have different GC content. Interestingly, Lsr2 controls genes responding to hypoxia, showing a functional analogy with cyAbrB2 (40).

The biochemistry of cyAbrB2 will shed light on the regulation of chromatin conformation in the future

H-NS proteins often cause bound DNA to bend, stiffen, and/or bridge (14). DNA-bound cyAbrB2 is expected to oligomerize, based on the long tract of cyAbrB2 binding region in our ChIP-seq data. However, no biochemical data mentioned the DNA deforming function or oligomerization of cyAbrB2 in the previous studies, and preference for AT-rich DNA is not fully demonstrated in vitro (19, 27, 41). Moreover, our 3C data did not support bridging at least in hox region and nifJ region, as the high interaction locus and cyAbrB2 binding region did not seem to correlate (Figure 7). Therefore, direct observation of the DNA-cyAbrB2 complex by atomic force microscopy is the solution in the future.

Not only DNA structural change but also the effect of the post-translational modification can be investigated by biochemistry. The previous studies report that cyAbrB2 is subject to phosphorylation and glutathionylation (42, 43), and pH and redox state alters cyAbrB1’s affinity to DNA (41). Those modifications might respond to environmental changes and be involved in transient expression. Overall, the biochemistry integrating assay conditions (PTM, buffer condition, and cooperation with cyAbrB1) and output (DNA binding, oligomerization, and DNA structure) will deepen the understanding of cyAbrB2 as cyanobacterial NAPs.

Cooperative and antagonistic function of cyAbrB1 and cyAbrB2

CyAbrB1, the homolog of cyAbrB2, may cooperatively work, as cyAbrB1 directly interacts with cyAbrB2 (29), their distribution is similar, and they partially share their target genes for suppression (Figures 3A S5C and S5D). The possibility of cooperation would be examined by the electrophoretic mobility shift assay of cyAbrB1 and cyAbrB2 as a complex. Despite their similar repressive function, cyAbrB1 and cyAbrB2 regulate hox expression in opposite directions, and their mechanism remains elusive. The stoichiometry of cyAbrB1 and cyAbrB2 bound to DNA fluctuates in response to the environmental changes (28), but there was no difference in the behavior of cyAbrB1 and cyAbrB2 around the hox region on entry to the microoxic condition.

Localization pattern and function of cyAbrB2

Herein, we classified three types of binding patterns for cyAbrB2. The first is that cyAbrB2 binds a long DNA tract covering the entire gene or operon, represented by the insertion sequence elements. CyAbrB2 suppresses expression in this pattern (Figure 3E). In the second pattern, cyAbrB2 binds on promoter regions, such as hox operon and nifJ. The binding on those promoters fluctuates in response to environmental changes, thus regulating expression. This pattern also applies to the promoter of sbtA (Na⁺/HCO₃⁻ symporter), where cyAbrB2 is bound in a CO₂ concentration-dependent manner (28). The last one is cyAbrB2 binding in the middle or downstream of operons. The middle of hox, pntAB, and nifJ operons and the downstream of nrt operon are the cases (Figure 4C). Our data show that genes in the same operon separated by the cyAbrB2 binding region behave differently. In particular, pntB is classified as the transiently upregulated gene, while pntA is not, despite being in the same operon. This might be explained by the recent study which reported that cyAbrB2 affects the stability of mRNA transcribed from its binding gene (41). The cyAbrB2-mediated stability of mRNA may also account for the decrease in transcript from transient upregulated genes at 4 hours of cultivation. Hereafter, we will focus on the mechanism of the second pattern, regulation by cyAbrB2 on the promoter.

Insight into the regulation of hox and nifJ operon by cyAbrB2

Genome-wide analysis indicates that the cyAbrB2-bound region blocks SigE and SigA (Figures 6A and 6B). This is presumably because sigma factors recognize the promoter as a large complex of RNA polymerase. CyAbrB2 binds to the hox and nifJ promoter region and may inhibit access to RNA polymerase complex under aerobic conditions. When cells entered microoxic conditions, the boundaries of the cyAbrB2 binding region and cyAbrB2-free region became obscure (Figure 5), and SigE binding peaks on those promoters became prominent (Figure 6C). Notably, cyAbrB2 ChIP efficiency at the hox promoter is higher in the microoxic condition than in the aerobic condition (Figure 5). Hence, while the exclusion by cyAbrB2 occupancy on promoter inhibits containing RNA polymerase in the aerobic condition, it is also plausible that chromosomal conformation change governed by cyAbrB2 provides SigE-containing RNA polymerase with access to the promoter region (Figure 8). Our 3C result demonstrated that cyAbrB2 influences the chromosomal conformation of hox and nifJ region to some extent (Figure 7).

A recent study demonstrated that manipulating the expression of topoisomerase, which influences chromosomal conformational change through supercoiling, affects transcriptional properties in cyanobacteria (15). Moreover, Song et al. pointed out that DNA looping may inhibit transcription in cyanobacteria (41) because artificial DNA looping by the LacI repressor of E. coli can repress cyanobacterial transcription (44). Thus, we infer conformation change detected by the present 3C experiment regulates expression of hox operon.

Generality for chromosomal conformation in cyanobacteria

Our 3C analysis revealed that local chromosomal conformation changes upon entry to the microoxic conditions (Figure 8). As cyAbrB2 occupies about 15% of the entire genome and globally regulates gene expression, cyAbrB2 likely governs the whole chromosomal conformation. Furthermore, the conformational changes by deletion of cyAbrB2 were limited, suggesting there are potential NAPs in cyanobacteria yet to be characterized. It is speculated that conformational change of the entire chromosome occurs to deal with many environmental stresses.

The sigE-mediated mechanism for the transient expression

One possible SigE-mediated mechanism for transient expression is the post-transcriptional activation and degradation of SigE in the dark; i.e., SigE is sequestered by anti-sigma factor under light conditions and released under dark (45), enabling acute transcription of hox operon and nifJ. Transcripts of sigE were continuously downregulated in our time-course transcriptome, while sigB (sll0306) and sigC (sll0184) were classified as continuous upregulated genes (Supplemental Table S1). It is possible that upregulated SigB and SigC outcompete SigE in prolonged incubation under microoxic conditions. Finally, SigE is degraded under dark within 24 hours (46).

Another reason for the microoxic specific expression may exist in the sequence of the hox promoter. We previously determined the consensus sequence of −10 element for SigE regulon in the aerobic condition as “TANNNT,” where N is rich in cytosine (25). The −10 sequence of the hox promoter “TAACAA” (23) deviates from the consensus, and no hexamer precisely fitting the consensus is found in the nifJ promoter. This deviation can inhibit SigE from binding during aerobic conditions, aside from cyAbrB2-mediated inhibition. Under the microoxic condition, transcription factors LexA (23) and Rre34 (16) may aid SigE binding to the promoter of hox and nifJ, respectively.

Moreover, SigE seems susceptible to the blocking from cyAbrB2 during the aerobic condition compared with SigA. This is supported by the odds ratio of SigE being in the cyAbrB2-free region was higher than that of SigA in the aerobic condition (Figures 6A and 6B). The higher exclusion pressure of cyAbrB2 on SigE may contribute to sharpening the transcriptional response of hox and nifJ on entry to microoxic conditions. Overall, multiple environmental signals are integrated into the hox and nifJ promoter through the cyAbrB2 and SigE dynamics.

Materials and Methods

Bacterial strains and plasmids

The glucose-tolerant strain of Synechocystis sp. PCC 6803 (47) was used as a wildtype strain in this study. The sigE (sll1689)-disrupted strain (G50), SigE FLAG-tagged strain, and SigA FLAG-tagged strain were constructed in a previous study (17, 25). Disruption and epitope-tagging of cyabrb1(sll0359) and cyabrb2(sll0822) were performed by homologous double recombination between the genome and PCR fragment (47). The resulting transformants were selected using three passages on BG-11 plates containing 5 µg/mL kanamycin. Genomic PCR was used to confirm the insertion of epitope tag fragments and gene disruption (Figure S9). Supplemental Tables S2, S3, and S4 contain the cyanobacterial strains, oligonucleotides, and plasmids used in this study.

Aerobic and microoxic culture conditions

For aerobic conditions, cells were harvested after 24 hours cultivation in HEPES-buffered BG-11₀ medium (48), which was buffered with 20 mM HEPES-KOH (pH 7.8) containing 5 mM NH₄Cl under continuous exposure to white light (40 µmol m⁻²s⁻¹) and bubbled with air containing 1% CO₂ (final OD₇₃₀ = 1.4–1.8). For the dark microoxic culture, the aerobic culture cell was concentrated to an OD₇₃₀ of 20 with the centrifuge and resuspended in the culture medium. The concentrated cultures were poured into vials, bubbled with N₂ gas, and sealed. The sealed vials were shaded and shaken at 30°C for the described times.

Antibodies and immunoblotting

Sample preparation for immunoblotting was performed as previously described (25), and FLAG-tagged proteins were detected by alkaline-phosphatase-conjugated anti-FLAG IgG (A9469, Sigma Aldrich, St. Louis, MO) and 1-Step NBT/BCIP substrate solution (Thermo Fisher Scientific, Waltham, MA).

RNA isolation

Total RNA was isolated with ISOGEN (Nippon gene, Tokyo, Japan) following the manufacturer’s instructions and stored at −80°C until use. The extracted RNA was treated with TURBO DNase (Thermo Fisher Scientific) for 1 hour at 37°C to remove any genomic DNA contamination. We confirmed that the A260/A280 of the extracted RNA was >1.9 by NanoDrop Lite (Thermo Fisher Scientific). We prepared triplicates for each timepoint for the RNA-seq library. RT-qPCR was performed as described elsewhere (46).

ChIP assay

Two biological replicates were used for each ChIP-seq experiment, and one untagged control ChIP was performed. ChIP and qPCR analyses were performed using the modified version of a previous method (25). FLAG-tagged proteins were immunoprecipitated with FLAG-M2 antibody (F1804 Sigma-Aldrich) conjugated to protein G dynabeads (Thermo Fisher Scientific).

Library preparation and next-generation sequencing

For the ChIP-seq library, input and immunoprecipitated DNA were prepared into multiplexed libraries using NEBNext Ultra II DNA Library Prep Kit for Illumina (New England Biolabs, Ipswich, MA). For the RNA-seq library, isolated RNA samples were deprived of ribosomal RNA with Illumina Ribo-Zero Plus rRNA Depletion Kit (Illumina, San Diego, CA) and processed into a cDNA library for Illumina with the NEBNext Ultra II Directional RNA Library Prep Kit for Illumina (New England Biolabs). Dual-index primers were conjugated with NEBNext Multiplex Oligos for Illumina (Set1, New England Biolabs). We pooled all libraries, and the multiplexed libraries were dispatched to Macrogen Japan Inc. and subjected to paired-end sequencing with HiSeqX. Adapter trimming and quality filtering of raw sequence reads were conducted with fastp (ver. 0.21.0) (49) under default conditions. The paired-end sequences were mapped onto the Synechocystis genome (ASM972v1) using Bowtie2 (50) (ver. 2.4.5 paired-end). Supplementary Table S5 contains the read counts that passed via fastp quality control and were mapped by Bowtie2.

RNA-seq analysis

Mapped reads were counted by HT-seq count (ver. 2.0.2) (51) for the GFF file of ASM972v1, with the reverse-strandedness option. EdgeR package (ver. 3.40.1) (52) was used to perform the differential expression analysis. Fold changes in expression and FDR were used for gene classification. Supplemental Table S6-S8 contains fold change in gene expression calculated by edgeR.

Genome-wide analyses

Peaks were called using the MACS3 program (ver. 3.0.0b1) (53). For paired-end reads for SigE, SigA, and untagged control ChIP, narrow peaks were called with <1e−20 of the q-value cut-off and “--call-summits” options. The peak summits from two replicates and the untagged control were merged if summits were located within 40 bp of each other. Peak summits identified in both replicates but not in the control, were considered for further analysis. The midpoint of the peak summits for the two merged replicates was further analyzed.

Broad peak calling methods were applied to paired-end reads for cyAbrB2, cyAbrB1, and untagged control ChIP using the “–broad” option, with a q-value cut-off of < 0.05 and a q-value broad cut-off of < 0.05. The intersection of broad peaks from two replicates, excluding those called by the control, was used in subsequent analyses.

The positions of the TSS, including internal start sites, were obtained as reported by (31). The read count, merging, and intersection of the binding region were calculated using BEDTools (ver. 2.30.0) (54). Supplemental Tables S9–S15 contain SigA and SigE peaks and the broad binding regions of cyAbrB2 and cyAbrB1, respectively.

Binding signals in every 100 bp bin for scatter plots were calculated as (IP read counts within 100 bp window) / (input read counts within 100 bp window) * (total input read counts/total IP read counts). GC contents were calculated within 500 bp in 100 bp sliding windows by seqkit (ver. 2.3.0) (55).

Genome extraction, digestion, and ligation for 3C assay

A 3C assay was conducted based on the previous prokaryotic Hi-C experiment (56, 57), with certain steps modified. To begin, Synechocystis were fixed with 2.5% formaldehyde for 15 minutes at room temperature. Fixation was terminated by adding a final concentration of 0.5M of glycine, and cells were stored at −80°C until use. Fixed cells were disrupted using glass beads and shake master NEO (Bio Medical Science, Tokyo, Japan), following the previous study’s instructions for preparing cell lysate for ChIP. The lysates were incubated with buffer containing 1mM Tris-HCl (pH7.5), 0.1mM EDTA, and 0.5% SDS for 10min at room temperature, and 1% Triton X-100 quenched SDS. Genomes in the cell lysate were digested by 600U/mL of HindIII (Takara Bio, Shiga, Japan) for 4 hours at 37°C, and RNA in the lysate was simultaneously removed by 50 µg/mL of RNaseA (Nippon genetics, Tokyo, Japan). The digestion was terminated by adding 1% SDS and 22 mM EDTA. The fill-in reaction and biotin labeling steps were omitted from the procedure. The digested genomes were diluted by ligation buffer containing 1% triton-X100 to the final concentration of approximately 1µg/mL and incubated for 10 min at room temperature. Ligation was performed with 2 U/mL of T4 DNA ligase (Nippon Gene) overnight at 16°C. Crosslinking was reversed under 65°C for 4 hours in the presence of 2.5 mg/mL of proteinase K (Kanto Chemical, Tokyo, Japan), and DNA was purified with the phenol-chloroform method and ethanol precipitation method.

Preparation of calibration samples for 3C qPCR

Based on a previous study, calibration samples for possible ligated pairs were prepared in parallel with 3C ligation (58). In brief, the purified genome of Synechocystis was digested by HindIII, and DNA was purified with the phenol-chloroform and ethanol precipitation. Purified DNA was dissolved into the ligation buffer at a concentration of about 600ng/µL and ligated with 2 U/mL of T4 DNA ligase at 16°C overnight.

Quantification of crosslinking frequency for 3C assay

Before the real-time PCR assay, we confirmed that each primer set amplified single bands in a ligation-dependent manner by GoTaq Hot Start Green Master Mix (Promega, Madison, WI) (Figure S10). Real-time PCR was performed with StepOnePlus (Applied Biosystems, Foster City, CA) and Fast SYBR Green Master Mix (Thermo Fisher Scientific) according to the manufacturer’s instructions. Interaction frequency was calculated by ΔΔCt method using dilution series of calibration samples described above. We confirmed each primer set amplified DNA fragment with a unique Tm value. The amount of the bait fragment containing hox operon were quantified and used as an internal control. Supplemental Table S4 contains the list of primers used in the 3C quantification. Interaction frequency for each primer position was calculated as the relative abundance of ligated fragments against the calibration samples and normalized among samples by internal control.

Statistical analysis

Statistical analyses were performed with R version 4.2.2 (59). The “fisher.test” function was used for Fisher’s exact test, and p-values < 0.05 were denoted as asterisks in the figure. Multiple comparisons of Fisher’s exact test were conducted using “fisher. Multcomp” function in the RVAideMemoire package (60), where p-values were adjusted by the “fdr” method and FDRs <0.05 are shown in the figures. Multiple comparisons of the Wilcoxon-rank test were conducted by “pairwise.wilcox.test,” and p-values were adjusted by the “fdr” method. Adjusted p-values < 0.05 are shown in the figure. The correlation coefficient was calculated with the “cor” function. Gene set enrichment analysis (GSEA) was performed by culsterPlofiler package (61) in R with p-value cut-off of 0.05. The enriched pathways detected by GSEA are listed in Supplemental Table S16.

Manuscript writing

We employed GPT-3.5 to refine the expression of the manuscripts.

Accession Numbers

Raw ChIP sequencing and RNA sequencing reads were deposited in the Sequence Read Archive (accession ID: PRJNA956842).

Declaration of interest

The authors declare no competing interests.

Acknowledgements

This study was supported by the following grants to T.O.: Grant-in-Aid for Scientific Research (B) (grant no. 20H02905), JST-ALCA of the Japan Science and Technology Agency (grant number JPMJAL1306), the Asahi Glass Foundation, and JST-GteX (grant number JPMJGX23B0). We thank Dr. Kohki Yoshimoto for providing laboratory instruments and Ms. Kaori Iwazumi for the support of bacterial culture and the medium preparation.

Author contributions

R.K. and T.O. designed the study; R.K. conducted the experiment; R.K. analyzed the data; R.K. and T.O. wrote the paper.