As the mild cell cycle defect of cells lacking the TipN polarity factor (∆tipN) is not well understood, we sought Tn mutations that enhance the defect. To this end, we compared the Tn insertion sequencing (Tn-Seq) profiles of wild-type and ∆tipN cells, seeking Tn insertions that specifically undermine the viability or fitness of cells lacking TipN. This analysis revealed Tn insertions in cpdR, rcdA or popA genes that are underrepresented in ∆tipN cells when compared with WT cells (Figure 1B and Figure 1—figure supplement 1A, Figure 1—source data 1). These three genes encode a hierarchical proteolytic adaptor cascade that coordinates the delivery of a range of substrates, including CtrA, to the ClpXP protease for proteolytic removal during the G₁→S transition (Duerig et al., 2009; Iniesta et al., 2006; Joshi et al., 2015; Joshi and Chien, 2016; McGrath et al., 2006). A converse Tn-Seq comparison between WT and ∆cpdR cells also revealed an underrepresentation of Tn insertions in the tipN gene (Figure 1—figure supplement 1B, Figure 1—source data 1). To confirm the genetic relationship between tipN and cpdR, rcdA or popA, we created double mutants by introducing the ∆cpdR, ∆rcdA or ∆popA mutations into ∆tipN cells and found that all of the resulting double mutants exhibit a reduction in viability by three orders of magnitude on a logarithmic scale, as determined by efficiency of plating (EOP) assays (Figure 1C; Figure 1—figure supplement 1C and D).

Examination of ∆tipN; ∆cpdR double mutant cells by phase-contrast microscopy showed that they are 70% more elongated on average than WT and ∆cpdR, ∆rcdA or ∆popA single mutant cells (Figure 1D and Figure 1—figure supplement 1E and F). Flow cytometry analysis of exponentially growing ∆tipN; ∆cpdR double mutant cells using a fluorescence activated cell sorter (FACS) revealed a massive reduction in the number of G₁-phase cells and an increase in the frequency of cells with multiple (>2) chromosomes compared to WT, whereas ∆cpdR and ∆tipN single mutants only showed a slight decrease in the G₁ population (Figure 1D). Importantly, the ∆tipN; ∆rcdA and ∆tipN; ∆popA double mutants show a similar accumulation of elongated cells and reduction in the number of G₁-phase cells (Figure 1—figure supplement 1E and F). Thus, the proteolytic adaptors promote efficient cell cycle progression in cells that lack TipN.

A reduction in the proportion of G₁ cells is often correlated with reduced activity or abundance of CtrA, the principal G₁-phase transcriptional regulator. To assess whether this is also the case in cells that lack both TipN and CpdR, we introduced a translational pilA::P_pilA-GFP promoter probe reporter into the pilA locus of WT cells, ∆tipN and ∆cpdR single mutant cells, and ∆tipN; ∆cpdR double mutant cells. In this reporter, the CtrA-dependent pilA promoter (P_pilA) that fires in G₁-phase along with the PilA start codon is translationally fused to a start codon-less variant encoding the green fluorescent protein (GFP). GFP expression from this reporter can be conveniently observed and quantified by live-cell fluorescence microscopy (Figure 2A). In agreement with the FACS profiles shown in Figure 1E, GFP fluorescence intensity is only slightly lower in ∆cpdR cells than in WT cells, but clearly reduced in ∆tipN cells. Importantly, a further strong decrease in GFP fluorescence is observed in ∆tipN; ∆cpdR double mutant cells, indicating a strong downregulation in CtrA-dependent reporter activity. Likewise, transcription from a reporter in which P_pilA is fused to the promoter-less nptII gene (conferring resistance to kanamycin) at the pilA locus (pilA::P_pilA-nptII) is strongly reduced in ∆tipN; ∆cpdR double mutant cells when compared to WT cells, precluding growth on plates containing 20 µg/mL kanamycin (Figure 2B). We conclude that cells that lack both TipN and CpdR suffer from an insufficiency of CtrA.

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Paradoxically, inactivation of CpdR, RcdA or PopA should enhance CtrA abundance in ∆tipN cells, since the proteolytic removal of CtrA at the G₁→S transition should be blocked in the absence of each of these adaptors. We therefore reasoned that another, indirect, effect underlies the crippled CtrA activity in ∆tipN ; ∆cpdR cells. It could be that this effect is mediated by an inhibitor of the CtrA pathway that is also degraded by the CpdR-RcdA-PopA pathway, which would accumulate in the absence of CpdR. We sought to uncover this gene by comparative Tn-Seq analyses in ∆tipN; ∆cpdR double mutant cells versus WT or ∆tipN and ∆cpdR single mutant cells, and we found a 19-fold increase in Tn insertions in the kidO gene (Figure 1—figure supplement 2A, Figure 1—source data 1). KidO is a bifunctional oxidoreductase-like negative regulator of cell division and the CtrA pathway (Radhakrishnan et al., 2010). Akin to CtrA, KidO is degraded during the G₁→S transition by (CpdR/RcdA/PopA) adaptor-primed ClpXP. As KidO is stabilized in cells lacking CpdR, RcdA or PopA (Radhakrishnan et al., 2010), we asked whether the stabilization of KidO confers the cell defect of ∆tipN; ∆cpdR cells. To test this idea, we expressed the kidO^AA::DD allele from the xylX locus in ∆tipN cells. This allele encodes a mutant form of KidO in which the two penultimate alanine residues are both substituted by aspartic acid residues, a double substitution that prevents degradation of KidO by the ClpXP protease at the G₁→S transition, akin to the ∆cpdR mutation (Radhakrishnan et al., 2010). The resulting ∆tipN xylX::kidO^AA::DD cells are highly filamentous, even without induction of the xylX promoter by xylose: with more than two chromosomes recapitulating the phenotype of the ∆tipN; ∆cpdR double mutant cells (Figure 1—figure supplement 2B). Conversely, an in-frame deletion in kidO (∆kidO) restores a near WT cell division phenotype to ∆tipN; ∆cpdR cells (Figure 1—figure supplement 2B).

Taken together, these experiments support the conclusion that stabilization of KidO strongly impairs progression of the cell division cycle in cells lacking TipN.

The impaired activity of the pilA::P_pilA-nptII reporter in ∆tipN; ∆cpdR cells (Figure 2A) offered a convenient opportunity to isolate Tn insertions that restore or elevate P_pilA activity. Towards this goal, we mutagenized ∆tipN; ∆cpdR; pilA::P_pilA-nptII reporter cells with a mini-himar1 Tn (Mar2xT7) encoding gentamycin resistance, and selected for growth on plates containing kanamycin and gentamycin. Among several isolated mutants, we found one mutant harboring a Tn insertion in the middle of the CCNA_01983 (henceforth citA) gene, whose gene product is annotated as a type II citrate synthase (PRK05614). After confirming by backcrossing that the citA::Tn mutation did indeed confer kanamycin resistance to ∆tipN; ∆cpdR; pilA::P_pilA-nptII reporter cells, we engineered an in-frame deletion of citA (∆citA) and found that this mutation also supports growth of ∆tipN; ∆cpdR; pilA::P_pilA-nptII reporter cells on kanamycin plates, indicating that inactivation of citA augments P_pilA activity (Figure 2B). Moreover, the citA::Tn or the ∆citA mutations both correct the abnormal cell size distribution (cell filamentation) and augment the G₁ population of ∆tipN; ∆cpdR double mutant cells (Figure 2C and Figure 2—figure supplement 1A).

In sum, inactivation of citA gene causes a strong increase of P_pilA activity and promotes the accumulation of G₁ cells in the joint absence of TipN and CpdR.

The primary structure of CitA resembles that of citrate synthases, which execute the first enzymatic reaction in the Krebs (tricarboxylic, TCA) cycle in which the acetyl group from acetyl-CoA is condensed onto oxaloacetate to form citrate (Figure 3—figure supplement 1A; Figure 3A). C. crescentus CitA has 65% amino acid identity to the GltA citrate synthase from Escherichia coli K12 (strain MG1655) and 32% identity to CitA from Bacillus subtilis (strain 168). To confirm that C. crescentus CitA does indeed have citrate synthase activity, we probed for heterologous complementation of glutamate auxotrophy in E. coli ∆gltA cells that lack citrate synthase activity (Lakshmi and Helling, 1976). To this end, we engineered E. coli ∆gltA cells expressing either C. crescentus CitA or E. coli GltA from a multicopy plasmid. As expected, E. coli ∆gltA cells harboring the empty vector were unable to grow in (M9) minimal medium without glutamate, but ∆gltA cells grew well in the presence of either the gltA- or the citA-expression plasmid (Figure 3B). Thus, C. crescentus citA encodes a functional citrate synthase.

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Metabolomic data sets showing the metabolites detected (sheet 1) and statistically significant changes in relative metabolite abundance between WT and citA::Tn mutant cells (sheet 2) or between WT and ∆citA mutant cells (sheet 3), presented as volcano plots.

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Next, we conducted metabolic profiling experiments using liquid chromatography coupled to high-resolution mass spectrometry (LC-HRMS) to quantify the abundance of intracellular metabolites in C. crescentus WT and citA::Tn or ∆citA cells grown in PYE (Pezzatti et al., 2019a). Robust quantitation of 103 metabolites (Figure 3—source data 1) revealed that the metabolomic profile of citA::Tn resembles that of ∆citA cells. Surprisingly, these metabolomic analyses did not show any significant difference in many TCAs such as citrate and isocitrate when comparing WT and citA mutant cells (Figure 3—figure supplement 1B). An indication that TCA cycle flux is nevertheless affected in the absence of CitA comes from the observation that there is a small increase in the levels of acetyl-CoA, as would be expected for citrate synthase mutant cells (Figure 3C).

The relatively modest effect of the ∆citA mutation on the TCA cycle activity might result from the presence of a protein(s) other than CitA that has citrate synthase activity. Unlike other TCA cycle enzymes, CitA is not essential for the viability of C. crescentus cells on PYE (Christen et al., 2011). Therefore, we reasoned that CitA is not the only citrate-synthase-like protein encoded in the C. crescentus genome. Indeed, BLAST searches revealed the presence of two other putative citrate synthase genes: CCNA_03757 and CCNA_03758 (Figure 3—figure supplement 1A) (henceforth citB and citC, respectively), which were also annotated as non-essential for viability on PYE (Christen et al., 2011). The citB and citC genes encode proteins with 30% and 32% identity to CitA from C. crescentus, 30% and 33% identity to GltA from E. coli K12 (MG1655), and 37% and 32% identity to CitA from Bacillus subtilis strain 168. We therefore tested the ability of citB and citC to support citrate synthase function by heterologous complementation of the glutamate auxotrophy of E. coli ∆gltA cells on minimal medium lacking glutamate, and found that expression of CitB, but not CitC, supported growth (Figure 3B). Thus, C. crescentus citB also encodes a functional citrate synthase and citA mutants are probably able to grow on PYE because of residual citrate synthase activity conferred by CitB. To test whether CitA is essential in cells lacking both citB and citC, we first created a strain with in-frame deletions in citB and citC (∆citBC) and then attempted to introduce citA::Tn (which encodes gentamycin resistance) or ∆citA (tagged with a kanamycin resistance marker, ∆citA::pNPTS138) by ϕCr30-mediated generalized transduction. Unlike WT cells, ∆citBC cells do not accept citA::Tn or ∆citA::pNPTS138 generalized transducing particles (Figure 3D), but accept generalized transducing particles harboring another genomic locus marked with either the gentamycin or the kanamycin resistance gene with efficiency similar to that of WT cells. We conclude that C. crescentus encodes at least two functional citrate synthases, one of which is absolutely required for growth on PYE.

To determine how loss of CitA signals G₁ cell accumulation, we combined population-based and single -cell approaches. First, EOP assays and growth curve measurements indicate that the absence of CitA leads to a slow growth phenotype in PYE and that CitA is required for growth on minimal M2G medium (Figure 4A). Phase contrast microscopy of citA::Tn or ∆citA mutant cells revealed that ∆citA cells are shorter and narrower than WT cells (areas of 0.42 ± 0.009 µm and 0.43 ± 0.007 µm, respectively, for the citA::Tn and ∆citA compared to 0.69 ± 0.01 µm for WT cells; Figure 4B), perhaps because they spend more time in the non-growing G₁ phase. Indeed, FACS profiles revealed a strong increase in the G₁-phase population in the absence of CitA: 68.3 ± 1.25% and 69.3 ± 1.22 of citA::Tn and ∆citA cells, respectively, reside in G₁ phase compared to 36.1 ± 0.6% of WT cells (Figure 4B). Importantly, these phenotypes of citA mutant cells cannot be corrected by the addition of exogenous glutamine and, therefore, are not related to glutamine auxotrophy. Indeed, the addition of glutamine to PYE or to M2G (minimal medium) does not ameliorate growth or division, as determined by EOP assays (Figure 4—figure supplement 1A). Moreover, the addition of glutamine does not restore a normal FACS profile to citA mutant cells (Figure 4—figure supplement 1B). The citA mutant phenotypes are not corrected by complementation of citA mutant cells with a multi-copy plasmid harboring C. crescentus citB (pMT335-citB) or E. coli gltA (pMT335-gltA), arguing that these functions probably depend on the presence of the CitA protein rather than on citrate synthase enzymatic activity (Figure 4C). However, these deficiencies are corrected when a WT copy of citA is expressed in trans on a multi-copy plasmid (pMT335-citA) (Figure 4C). Thus, CitA promotes the G₁→S transition, a function that other citrate synthases such as CitB and GltA cannot provide.

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Further support for the conclusion that CitA fulfills a regulatory role that is independent of its catalytic activity came from the discovery that catalytically inactive CitA can still control the cell cycle. Residue H306 of E. coli GltA is critical to bind oxaloacetate, and its substitution impairs the catalytic activity of GltA (Handford et al., 1988; Pereira et al., 1994). We thus engineered variants in which the corresponding residue (H303) in C. crescentus CitA is substituted either by a tryptophan or by an alanine, giving rise to the H303W and H303A CitA variants. As expected, expression of the CitA^H303W or CitA^H303A variant in E. coli ∆gltA cells no longer correct the glutamate auxotrophy on minimal medium, as determined by EOP assays (Figure 4—figure supplement 1C). Immunoblotting using polyclonal antibodies to CitA revealed that these variants are produced to the same levels as WT CitA (Figure 4—figure supplement 1D). We therefore conclude that CitA^H303W and CitA^H303A have lost enzymatic activity. When these variants are expressed in C. crescentus ∆citA mutant cells to similar levels as WT CitA (Figure 4—figure supplement 1E), a normal FACS profile and cell size distribution is observed by phase-contrast microscopy (Figure 4D). As these results show that the catalytic activity of CitA is dispensable for its developmental function, CitA must fulfill a specific regulatory role in promoting the G₁→S transition.

To establish that CitA is required for the G₁→S transition, we performed cell cycle studies using synchronized WT and citA mutant cells. FACS profiles revealed that WT G₁ cells initiate DNA replication 30 minutes after their release into PYE, whereas citA::Tn or ∆citA G₁ cells do not enter S-phase before 90 minutes after their release into PYE (Figure 4—figure supplement 1G). We also discovered that a fraction of citA::Tn or ∆citA cells remain in G₁ phase, with only approximately half entering S-phase. To confirm this observation at the single-cell level, we conducted time-lapse microscopy experiments with synchronized WT and citA::Tn or ∆citA G1 cells expressing GFP-ParB as a marker for DNA replication (Figure 4E). ParB is a chromosome partitioning protein that specifically binds near the origin of replication (C_ori) and is translocated with a duplicated copy of C_ori to the daughter cell pole once DNA replication commences (Mohl and Gober, 1997; Thanbichler and Shapiro, 2008). In synchronized WT G1 cells expressing ParB-GFP, we observed a single, polarly localized C_ori, represented by a single GFP-ParB focus. After 40 minutes, ~80% (n = 39) of the cells have a duplicated GFP-ParB focus, one of which is segregated to the opposite pole. Finally, cell division is completed by 120 minutes. By contrast, in citA::Tn (n = 35) or ∆citA (n = 29) G₁ cells, a duplicated GFP-ParB focus only appeared in some cells after 100 minutes. Importantly, we noticed that even after 260 minutes, ~60% of the population still exhibit only one GFP-ParB focus. Thus, a large fraction of the population remains in G₁-phase and only part of the citA mutant population enters S-phase.

While chromosome duplication is delayed, citA G₁ cells harbor a long stalk, as indicated by live-cell fluorescence imaging with the stalk marker StpX-GFP (Hughes et al., 2010), which coincides with the presence of a SpmX-mCherry focus, a marker of the stalked pole (Figure 4F). Knowing that SpmX is normally absent from the G₁ cells, we hypothesized that polar remodeling and chromosome replication might be uncoupled in the absence of CitA (Figure 5—figure supplement 1C). Snapshot analyses of a population of synchronized WT cells expressing MipZ-YFP (a marker of chromosome origin) and SpmX-mCherry (a marker of polar remodeling) revealed that chromosome duplication occurs before the appearance of SpmX-mCherry. Contrary to that, 60 minutes after synchronization, citA::Tn cells have a MipZ-YFP and SpmX-mCherry focus at the same pole, suggesting that chromosome replication and polar remodeling is uncoupled in the absence of CitA. This is typically a phenotype observed in cells that have hyperactivation of CtrA (Hung and Shapiro, 2002).

As inactivation of citA augments P_pilA activity, we hypothesized that the citA mutation elevates CtrA activity and/or abundance. To test this hypothesis, we asked whether the activity of other CtrA-activated promoters is also elevated in ∆citA cells compared to WT cells. LacZ-based promoter probe assays indeed revealed elevated activity of CtrA-dependent promoters (Figure 5A). To explore whether loss of CitA alters the levels of active CtrA, we used immunoblotting of Phos-tag PAGE to confirm that the levels of phosphorylated CtrA (CtrA ~P) in extracts of ∆citA cells are elevated relative to those in WT cell extracts (Figure 5B and C). This result prompted us to investigate whether this increase of CtrA steady-state levels was caused by increased stability of CtrA. Chloramphenicol chase experiments (Figure 5D and E) revealed that CtrA is indeed more stable in ∆citA cells than in WT cells, with the stability levels being similar to that of a non-degradable version of CtrA (ctrA::ctrA-M2) (Domian et al., 1997).

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ChIP-Seq data set showing RNAP peak abundance measured as sequencing reads of a 20-bp window across the genome of WT, ∆citA and xylX::P_xyl-relA’-FLAG cells (in sheet 1). Sheet two shows the peaks sorted for CtrA-activated promoters that fire in G₁-phase, and sheet three shows the peaks for CtrA-activated promoters that fire in late S-phase.

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To correlate these indirect reporter assays directly and specifically with increased transcription at CtrA-dependent promoters on a genome-wide scale, we quantified the occupancy of RNA polymerase (RNAP) on the genome using chromatin-immunoprecipitation followed by deep-sequencing (ChIP-Seq) experiments (Figure 5F and G, Figure 5—source data 1). This quantification revealed an increase of RNAP occupancy at many, but not all, CtrA-dependent promoters in cells lacking CitA when compared with WT cells. Interestingly, the promoters with the highest change in abundance of RNAP in citA cells when compared with WT cells are those that are activated by CtrA in G₁-phase (Figure 5F, Figure 5—figure supplement 1D; Delaby et al., 2019; Fumeaux et al., 2014; Schrader et al., 2016). This is exemplified by traces of RNAP occupancy in WT and citA mutant cells on well characterized G₁-promoters of sciP, pilA, and hfsJ (Figure 5G).

To see whether this effect is specific to the citA mutant phenotype or a generalized response of a cellular G₁ block, we used a control strain harboring a relA’-FLAG under the control of a promoter that is inducible by xylose. When induced, the resulting cells exhibit a G₁ arrest similar the ∆citA strain . Tracking RNAP occupancy by ChIP-Seq revealed an increase of binding on CtrA-regulated promoters, but without preference for the G₁-phase and S-phase promoter classes (Figure 5F). It is also important to note that global analysis of RNAP binding on all promoters in pairwise comparison between relA’-FLAG cells and WT cells or ∆citA cells (with a R² of 0.83 and 0.81, respectively) showed poor correlation (Figure 5F; Figure 5—figure supplement 1D). Thus, although RNAP occupancy under ectopic (p)ppGpp production leads to an enrichment on both classes of CtrA-dependent promoters, the changes induced by the citA mutation, at least on the level of RNAP occupancy, are restricted to G₁-phase promoters. Interestingly, the LacZ promoter probe assays suggest that the citA mutation might also promote a transcriptional event after RNAP promoter recruitment.

Immunoblotting experiments, using polyclonal antibodies to CitA, revealed that CitA is present at a constant level throughout the cell cycle (Figure 4—figure supplement 1F). This is consistent with RNA-Seq and ribosome profiling data showing that the levels of the citrate synthase (citA, citB and citC) transcripts and their association with ribosomes does not vary considerably during the cell cycle and that the citA transcript is more abundant than those of citB and citC (Schrader et al., 2016). As the cell cycle control function of CitA is not explained by changes in CitA abundance, other cell cycle signals or events probably affect CitA action. As the delay in the G₁→S transition of citA mutant cells probably confers a reduced growth rate of the population (Figure 6—figure supplement 1A), we anticipated that the isolation of fast-growing suppressor mutants would reveal how the cell cycle defect of citA cells can be overcome. We therefore isolated spontaneous suppressor mutants by serially diluting ∆citA or citA::Tn cultures. After three days of serial dilution, we plated cells on PYE and isolated large colonies from the background of slower-growing citA::Tn or ∆citA colonies. The growth and morphology of these mutant cells is like that of WT cells (Figure 6—figure supplement 1A). Whole-genome sequencing of two citA::Tn and one ∆citA suppressor mutant revealed a different frameshift mutation in the same domain of the PEP-phosphotransferase protein encoded by the ptsP gene (CCNA_00892) (Ronneau et al., 2016; Sanselicio et al., 2015). PtsP resembles the first enzyme of a nitrogen-related PEP-phosphotransferase (PTS) protein homolog (EI^Ntr in Enterobacteria) and was shown to inhibit the hydrolase activity of SpoT, the bifunctional synthase/hydrolase of the (p)ppGpp alarmone (Ronneau et al., 2016).

We hypothesized that the PtsP frameshift mutation in the citA suppressor mutants eliminates or decreases PtsP function by affecting (p)ppGpp levels. Indeed, when the citA::Tn mutation was introduced into ∆ptsP or ∆spoT cells, the resulting double mutants grow faster in PYE broth than the ∆citA single mutant and have a higher EOP (Figure 6—figure supplement 1B). Importantly, the FACS profile of ∆ptsP citA::Tn or ∆spoT citA::Tn double mutant cells mirrors that of WT cells, indicating that loss of (p)ppGpp production indeed mitigates the effects caused by loss of CitA (Figure 6A), including the enhanced levels of CtrA ~P that are restored to WT levels in citA; ptsP double-mutant cells (Figure 5B, C, D and E). Quantification of radiolabeled (p)ppGpp extracted from WT and citA mutant cells grown in PYE did not reveal an increase in (p)ppGpp levels (Figure 6B), suggesting that inactivation of citA and the (p)ppGpp pathways converge on the same target. In addition, artificial (p)ppGpp production by expressing RelA’-FLAG enhances swarming motility in soft (0.3%) agar (Figure 5—figure supplement 1B), but the citA mutant does not show a commensurate increase, further reinforcing the notion that (p)ppGpp levels are not elevated in citA mutant cells when compared to WT cells.

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We conclude that CitA acts negatively on CtrA ~P and that this action depends on the presence of (p)ppGpp.

Ectopic induction of (p)ppGpp in WT cells delays the G₁→S transition and prevents the concomitant turnover of CtrA. The augmentation of the G₁ population when citA is inactivated and the stabilization of CtrA turnover perfectly mirror these effects. However, (p)ppGpp levels are not altered in the citA mutant, yet (p)ppGpp is absolutely required for the manifestation of the citA phenotype. On the basis of these results, we propose that CitA and the (p)ppGpp pathways converge to cause an increase in active CtrA (Figure 6C). Although we cannot exclude the hypothesis that CitA acts downstream of (p)ppGpp, a well-known transcriptional target of (p)ppGpp is RNAP (Hauryliuk et al., 2015). RNAP also seems to be a target of (p)ppGpp in alpha-proteobacteria (Delaby et al., 2019; Wells and Long, 2003), suggesting that the effect of (p)ppGpp on CtrA is mediated by a change in transcription. An active site in the cavity created by the alpha, beta' and omega subunits of RNAP binds (p)ppGpp (Hauryliuk et al., 2015). The observed accumulation of CtrA ~P in citA mutant cells and the increase in occupancy of RNAP at CtrA target promoters could result in effects that are comparable to the ectopic induction of (p)ppGpp.

As control of CtrA stability is mediated by the proteolytic adaptors CpdR, RcdA and PopA (Joshi and Chien, 2016), it is tempting to speculate that the citA mutation influences this pathway. However, the fact that the citA::Tn mutation was isolated as a suppressor from a ∆tipN ∆cpdR background already argues against this possibility. Moreover, we observed that the activities of the CtrA-dependent transcriptional reporters P_pilA-lacZ and P_hfsJ-lacZ are diminished in ∆popA and ∆cpdR mutant cells relative to those in WT cells (Figure 5—figure supplement 1E), whereas reporter activities in the citA mutant exceed WT levels. Thus, if the effects on CtrA in citA cells are mediated via CpdR/RcdA/PopA, then another pathway must also be affected to explain the observed effects on CtrA ~P.

By affecting CtrA, arguably the master regulator of the C. crescentus cell cycle, CitA is perfectly positioned to integrate central energy metabolism with cell cycle transcriptional regulation. This function is unique to CitA, as expression of the paralog CitB from C. crescentus or the GltA ortholog from E. coli in ∆citA cells does not restore cell cycle control, even though both enzymes exhibit efficient citrate synthase activity in an E. coli reporter system. Our finding that addition of glutamine does not rescue the developmental problem of a ∆citA strain and that metabolite extractions from citA mutant cells grown on PYE do not reveal a major perturbance in the levels of tricarboxylic acids, provide further support for the conclusion that the citA mutant phenotype is not simply caused by a metabolic deficiency of blocked citrate production. Complementation analysis with catalytically inactive CitA variants revealed that they still confer cell cycle control functions. Moreover, other TCA cycle enzymes are essential for viability in C. crescentus (Christen et al., 2011) Figure 3, presumably because TCA products are essential during growth on PYE. Thus, the apparent redundancy in citrate synthase activities may have allowed the appropriation of CitA to control the cell cycle, as a checkpoint regulator and ‘moonlighting’ or ‘trigger’ enzyme.

Other bacterial lineages also encode multiple citrate synthases, and even within the Rhizobia, symbiotic relatives of C. crescentus, genera with three citA paralogs are often found (Figure 3—figure supplement 1C). The requirement of citrate synthase in virulence and development has been noted in other bacteria. Recently, the GltA citrate synthase from Klebsiella pneumoniae was identified as a virulence determinant (Vornhagen et al., 2019) that is required for replication in different organs, despite the presence of a GltA paralog in the genome. In addition, TCA cycle proteins have been implicated in conferring persister (non-growing) traits in Staphylococcus aureus cells that protect them against bacteriocidal antibiotics (Zalis et al., 2019). This finding aligns well with our observation that the non-replicative G₁-phase population is increased in C. crescentus citA mutants.

Developmental roles have also been described for citrate synthase in other bacteria. B. subtilis cells that lack citrate synthase sporulate poorly (Ireton et al., 1995) and a citrate synthase mutant of Streptomyces coelicolor is impaired in aerial mycelium formation (Viollier et al., 2001). Importantly, while the growth defect of the citrate synthase mutant in S. coelicolor on minimal medium was suppressed by the addition of glutamate, development remains perturbed. Thus, developmental events in bacteria may be controlled by switches, and central metabolic enzymes serve as ideal checkpoint mechanisms that couple developmental gene expression to central energy metabolism.

Eukaryotic cells, such as those of Saccharomyces cerevisiae, restrict citrate synthase paralogs to different compartments of the cell. CIT1 is located in the mitochondria where it participates in the TCA cycle, while CIT2 is located in the peroxisome, where it acts in the glyoxylate cycle (Kim et al., 1986; Rosenkrantz et al., 1986). In Podospora anserina, a citrate synthase mutant strain exhibits a developmental phenotype that impairs meiosis independently of its catalytic citrate synthase activity (Ruprich-Robert et al., 2002), reminiscent of our finding highlighting alternate functions of citrate synthases in cell cycle control of another domain of life.

The regulatory role of moonlighting enzymes raises the question of whether metabolic changes occur during the bacterial cell cycle to direct developmental changes, perhaps using moonlighting enzymes as sensors. Evidence has been provided that the cellular redox potential changes as a function of the C. crescentus cell cycle (Narayanan et al., 2015), and a recent study revealed that glutathione and many unknown metabolites fluctuate in accordance with the cell cycle (Hartl et al., 2020). The oxidoreductase homolog KidO is an NADH-binding protein that is present in the G₁-phase and during cell constriction. KidO is bifunctional, acting as cell division inhibitor that binds FtsZ and as a negative regulator of the CtrA activation pathway (Radhakrishnan et al., 2010). Interestingly, another division regulator that functions as a moonlighting enzyme and that is degraded in a ClpXP and CpdR-dependent manner has been identified: the glutamate dehydrogenase GdhZ whose activity is modulated by glutamate and NADH (Beaufay et al., 2015). The fact that KidO and GdhZ, two NAD(H) binding proteins, and CitA, an active citrate synthase, execute cell cycle control functions, indicates that C. crescentus, and probably other bacteria, integrate cell cycle control with central cellular metabolism (at multiple levels) using different checkpoint regulators derived from metabolic enzymes.

The moonlighting functions of KidO, GdhZ and CitA are not apparent by analysis of their primary structure. Although GdhZ was discovered as an interaction partner of FtsZ in a yeast two-hybrid screen (Beaufay et al., 2015), the genes encoding KidO and CitA both emerged from a forward genetic selection for cell cycle mutants that restore activity of the CtrA-regulated promoter P_pilA that fires in G₁-phase (Radhakrishnan et al., 2010). As CtrA targeted promoters include not only those controlling expression of envelope and developmental functions, but also those controlling metabolic functions (Fiebig et al., 2014; Fumeaux et al., 2014; Laub et al., 2000), it is appealing to speculate that the molecular and genetic interplay between metabolism and cell cycle control is much more profound than anticipated, at least in C. crescentus and related bacteria.

Strains, plasmids and oligos are listed in Supplementary files 1, 2 and 3. C. crescentus NA1000 (Marks et al., 2010) and derivatives were cultivated at 30°C in peptone yeast extract (PYE)-rich medium (2 g/L bactopeptone, 1 g/L yeast extract, 1 mM MgSO₄, and 0.5 mM CaCl₂) or in M2 minimal salts supplemented with 0.2% glucose (M2G, 0.87 g/L Na₂HPO₄, 0.54 g/L KH₂PO₄, 0.50 g/L NH₄Cl, 0.2% [wt/vol] glucose, 0.5 mM MgSO₄, 0.5 mM CaCl₂, and 0.01 mM FeSO₄) (Ely, 1991). E. coli S17-1 λpir (Simon et al., 1983) and EC100D (Epicentre Technologies, Madison, WI) cells were grown at 37°C in Lysogeny Broth (LB)–rich medium (10 g/L NaCl, 5 g/L yeast extract, and 10 g/L tryptone). When appropriate, media were supplemented with antibiotics at the following concentrations (µg/mL in liquid/solid medium for C. crescentus strains; μg.mL⁻¹ in liquid/solid medium for E. coli strains): kanamycin (5/20 μg.mL⁻¹; 20/20 μg.mL⁻¹), tetracycline (1/1 μg.mL⁻¹; not appropriate), spectinomycin and streptomycin 5 μg.mL⁻¹ (in solid medium for C. crescentus only) (25/25 μg.mL⁻¹; 30/90 μg.mL⁻¹), gentamycin (1/1; 10/25 μg.mL⁻¹), aztreonam (in solid medium only) (2.5 μg.mL⁻¹) and colistin (4 μg.mL⁻¹). PYE plates containing 3% sucrose were used to select for loss of pNTPS138-derived plasmids by recombination when constructing mutants by double recombination. When needed, for C. crescentus, D-xylose was added at 0.3% final concentration, glucose at 0.2% final concentration. Glutamine was used at 9.3 mM final in liquid and solid medium.

Swarmer cell isolation, electroporations, biparental matings (intergeneric conjugations) and bacteriophage ϕCr30-mediated generalized transductions were performed as described previously (Ely and Johnson, 1977) with slight modifications. Briefly, swarmer cells were isolated by Percoll density-gradient centrifugation at 4°C, followed by three washes and final re-suspension in pre-warmed (30°C) PYE. Electroporation was done from 1 mL overnight culture that had been washed three times in sterile water. Biparental mattings were done using exponential phase E. coli S17-1 donor cells and C. crescentus recipient cells washed in PYE and mixed at 1:3 ratio on a PYE plate. After 4–5 hr of incubation at 30°C, the mixture of cells was plated on PYE harboring aztreonam (to counter select E. coli) and the antibiotic that the conjugated plasmid confers resistance to. Generalized transductions using ϕCr30 were carried out by mixing 50 μL ultraviolet-inactivated ϕCr30 lysate with 500 μL stationary phase recipient cells, incubating for 2 hours, followed by plating on PYE-containing antibiotic to select for the transduced DNA.

For metabolite extraction, C. crescentus cells were grown overnight at 30°C in PYE medium and diluted to reach an OD_600nm ~ 0.4. Ten mL of cell culture were centrifuged at 2000 g for 5 minutes at 4°C. Metabolism was then quenched by resuspending the pellet in 1 mL of precooled methanol/H₂O (80:20 [vol/vol], kept at ~ −20°C). Cells were subjected to lysis by five thaw/freeze (40°C /–80°C) cycles. Cellular debris was removed by centrifugation at 17,000 g for 20 minutes at 4°C. Metabolite extracts were kept at −80°C prior to analysis on LC-MS. Bacterial biomass of individual samples was determined for normalization. The supernatants were completely evaporated using a SpeedVac (ThermoFisher, Langenselbold, Germany) and metabolite extracts were reconstituted in 100 µL acetonitrile:H₂O 50:50. Quality control (QC) and diluted QC (dQC, diluted by 50%) samples were prepared by pooling equivalent volumes of all reconstituted samples and injected at a regular interval of five samples to assess analytical variability.

LC experiments were performed on a Waters H-Class Acquity UPLC system composed of a quaternary pump, an auto-sampler including a 15 μL flow-through-needle injector and a two-way column manager (Waters, Milford, USA) for which temperatures were set at 7°C and 40°C, respectively. The injected volume was 10 μL. Samples were analyzed with a hydrophilic liquid interaction chromatography (HILIC) SeQuant Zic-pHILIC column (150 × 2.1 mm, 5 μm) and the appropriate guard kit. For mobile phases, solvent A was acetonitrile and solvent B was H₂O containing 2.8 mM ammonium formate adjusted to pH 9. Column flow rate was set at 300 µL.min⁻¹. The following gradient was applied: 5% B for one minute, increased to 51% B over 9 minutes, holding for 3 minutes at 51% B and then returning back to 5% B in 0.1 min and re-equilibrating the column for 6.9 min. The UPLC system was coupled to a TWIMS-QTOF high resolution HRMS (Vion, Waters, Manchester, UK) through an electrospray ionization (ESI) interface. Analyses were performed in negative ESI mode, and continuum data in the range of 50–1000 m/z were acquired with a scan time of 0.2 s. The ESI parameters were set as follows: capillary voltage was −2.0 kV, source and desolvation temperatures were set at 120°C and 500°C, respectively, cone and desolvation gas flow were 50 and 800 L/h, respectively. Velocity and height of StepWave1 and StepWave2 were set to 300 m/s and 5 V and to 200 m/s and 30 V, respectively. The high definition MS^E (HDMS^E, using ion mobility) settings consisted of trap wave velocity at 100 m/s; trap pulse height A at 10 V; trap pulse height B at 5 V; IMS wave velocity at 250 m/s; IMS pulse height at 45 V; wave delay set at 20 pushes; and gate delay at 0 m/s. Gas flows of ion mobility instrument were set to 1.60 L/minute for trap gas, and 25 mL/min for IMS gas. Buffer gas was nitrogen. Fragmentation was performed in HDMS^E mode. For the collision energy, 6.0 eV was used for low energy and high energy was a ramp from 10 to 60 eV. Nitrogen was used as collision gas. Leucine-encephalin served as a lock-mass (554.2615 m/z for ESI-) infused at 5 minute intervals. The CCS and mass calibration of the instrument were done with the calibration mix ‘Major mix IMS-TOF calibration’ (Waters, Manchester, UK). UNIFI v1.9.3 was used for data acquisition and data treatment.

Run alignment, peak picking, adduct deconvolution and feature annotation were sequentially performed on Progenesis QI v2.3 (Nonlinear Dynamics, Waters, Newcastle upon Tyne, UK). Detected peaks were annotated with regard to a set of pure reference standards (MSMLS Library of Standards, Sigma-Aldrich) measured under the experimental conditions described previously (Pezzatti et al., 2019b). The following tolerances were used: 2.5 ppm for precursor and fragment mass, 10% for retention time (Rt), and 5% in the case of collisional cross section (CCS). Data processing was achieved by SUPreMe, which is in-house software with capabilities for drift correction, noise filtering and sample normalization. Finally, data were transferred to SIMCA-P 15.0 software (Umetrics, Umea, Sweden) for multi-variate analysis (MVA).

Exponential phase C. crescentus cells cultivated in PYE were immobilized on a thin layer of 1.2% agarose. For C. crescentus time-lapse imaging, cells were first synchronized by Percoll density-gradient centrifugation and then immobilized on a thin layer of 1.2% agarose in PYE. Fluorescence and contrast microscopy images were taken with a phase contrast objective (Zeiss, alpha plan achromatic 100X/1.46 oil phase 3) on an Axio Imager M2 microscope (Zeiss), with appropriate filter (Visitron Systems GmbH) and a cooled CCD camera (Photometrics, CoolSNAP HQ2) controlled through Metamorph (Molecular Devices). Images were acquired and processed with ImageJ via Fiji software (Schindelin et al., 2012; Schneider et al., 2012). To perform cell segmentation and tracking, images were processed using MicrobeJ (Ducret et al., 2016). Statistics were performed on experiments performed in triplicate representing more than 300 cells.

Mid-log phase cells were cross-linked in 10 mM sodium phosphate (pH 7.6) and 1% formaldehyde at room temperature (RT) for 10 minutes and on ice for 30 minutes thereafter, washed three times in phosphate-buffered saline (PBS) and lysed in a Ready-Lyse lysozyme solution (Epicentre Technologies) according to the manufacturer’s instructions. Lysates were sonicated in an ice-water bath (15 cycles of 30 seconds ON, 30 seconds OFF) to shear DNA fragments to an average length of 0.3–0.5 kbp and cleared by centrifugation at 14,000 rpm for 2 minutes at 4°C. Lysates were normalized by protein content, diluted to 1 mL using ChIP buffer (0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, 16.7 mM Tris-HCl [pH 8.1]), 167 mM NaCl plus protease inhibitors (Roche, Switzerland) and pre-cleared with 80 μl of protein-A agarose (Roche) and 100 μg BSA. To immunoprecipitate the chromatin, 2 µL of a mixed of RNA polymerase antibody sampler kit (ratio 1:1:1:1, Biolegend) were added to the supernatant, incubated overnight at 4°C with 80 μL of protein-A agarose beads pre-saturated with BSA. The immunoprecipitate was washed once with low salt buffer (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris-HCl [pH 8.1] and 150 mM NaCl), high salt buffer (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris-HCl [pH 8.1] and 500 mM NaCl) and LiCl buffer (0.25 M LiCl, 1% NP-40, 1% sodium deoxycholate, 1 mM EDTA and 10 mM Tris-HCl [pH 8.1]), and twice with TE buffer (10 mM Tris-HCl [pH 8.1] and 1 mM EDTA). The protein–DNA complexes were eluted in 500 μL freshly prepared elution buffer (1% SDS and 0.1 M NaHCO₃), supplemented with NaCl to a final concentration of 300 mM and incubated overnight at 65°C to reverse the crosslinks. The samples were treated with 2 μg of Proteinase K for 2 hours at 45°C in 40 mM EDTA and 40 mM Tris-HCl (pH 6.5). DNA was extracted using phenol:chloroform:isoamyl alcohol (25:24:1), ethanol precipitated using 20 μg of glycogen as carrier and resuspended in 100 μL of water.

Immunoprecipitated chromatin was used to prepare barcoded libraries for deep-sequencing at Fasteris SA (Geneva, Switzerland). ChIP-Seq libraries were prepared using the DNA Sample Prep Kit (Illumina) following the manufacturer’s instructions. Single-end runs were performed on an Illumina Genome Analyzer IIx or HiSeq2000, 50 cycles were read and yielded several million reads. The single-end sequence reads stored in FastQ files were mapped against the genome of Caulobacter crescentusNA1000 (NC_011916) and converted to SAM using BWA and SAM tools from the galaxy server (https://usegalaxy.org/). The resulting SAM was imported into SeqMonk (http://www.bioinformatics.babraham.ac.uk/projects/seqmonk/, version 0.21.0) to build sequence read profiles. The initial quantification of the sequencing data was done in SeqMonk: the genome was subdivided into 20-bp probes, and for every probe, we calculated a value that represents the number of reads that occur within the probe (using the Read Count Quantitation option). The heatmaps represent the abundance of RNA polymerase 200 bp upstream and 200 bp after the beginning of the CDS of the gene that belongs to the CtrA regulon, as determined in previous studies (Fumeaux et al., 2014; Schrader et al., 2016). Sequence data have been deposited in the Gene Expression Omnibus (GEO) database (GSE144533).

(p)ppGpp levels were determined using a protocol adapted from a previous study by Lesley and Shapiro (2008). Briefly, strains were grown in PYE medium, and 1 mL normalized at OD₆₀₀ of 0.4 was centrifuged for 3 minutes at 10,000 RPM and resuspended in 250 μl of PYE. A final concentration of 20 μCi of H₃³²PO₄ (Hartmann Analytic) was added to the cultures, and the cells were incubated for 120 minutes at 30°C with agitation, before being fixed by the addition of 2M formic acid. As a positive control, a culture of NA1000 expressing relA’-FLAG under the control of the xylX promoter was grown in PYE for 3 hours, then xylose or glucose was added to induce or repress, respectively, the expression of relA’-FLAG for 120 minutes. All cell extracts were kept on ice for 30 min and then centrifuged for 5 minutes, and 8 × 2 μl of the supernatant (16 μl total) was spotted onto a polyethyleneimine (PEI) plate (Sigma-Aldrich). The PEI plate had been soaked in sterile distilled water overnight and dried at room temperature before spotting. The plate was developed in 1.5 M KH₂PO₄ (pH 3.4) in a saturated thin-layer chromatography (TLC) chamber for approximately 180 min and dried at RT. Nucleotides were detected via a phosphorimaging system (Tritium screen). ppGpp, and GTP were identified on the basis of their retardation factor (R_f) and by comparison with a standard for GTP. Spots were quantified using ImageJ software.

To determine the in vivo phosphorylation of CtrA, strains were grown to mid‐log phase (OD_600nm around 0.4), and 1 mL of cells were pelleted at 20,000 g at 4°C for 5 minutes. Pellets were resuspended in 75 µL TE buffer (10 mM Tris-HCl [pH 8.0] and 1 mM EDTA) followed by the addition of 75 µL loading buffer 2X (0.25 M Tris [pH 6.8], 6% [wt/vol] SDS, 10 mM EDTA, 20% [vol/vol] glycerol) containing 10% (vol/vol) β-mercaptoethanol. Samples were normalized for equivalent loading using OD_600nm. Some samples were boiled by heating at 90°C for 10 minutes. Samples were stored on ice for a short time (<10 minutes) prior to loading onto Phos‐tag acrylamide gels.

Phos‐tag SDS‐PAGE gels were prepared with 25 μM Phos‐tag acrylamide and 50 μM MnCl₂. All gels were run at 4°C under constant voltage (80 V). Before transfer by blotting, gels were washed three times for 10 minutes in transfer buffer containing 10 mM EDTA at 4°C to remove Mn²⁺ from the gel and once with transfer buffer without EDTA at 4°C. Blots were revealed by immunodetection using Western Blot Signal Enhancer (Thermo Pierce) after incubation with rabbit anti‐CtrA (1:5000) primary antibodies and a polyclonal donkey anti-rabbit HRP conjugated secondary antibody (Jackson ImmunoResearch). Band intensities were analyzed using ImageJ. The total CtrA content was determined using a control gel that did not contain Phos‐tag, whereas phosphorylated and non-phosphorylated forms of CtrA were estimated from the gel containing Phos‐tag using two independent biological replicates.

To measure protein stability in vivo, cells were grown to mid‐log phase (OD_600nm of ca. 0.4). Protein synthesis was blocked by the addition of 50 μg/mL chloramphenicol. Samples were taken every 15 min and frozen immediately at −80°C before being analyzed by immunoblotting.

100 μL of cells at OD_600nm = 0.1–0.4 were lysed with chloroform and mixed with 700 μl of Z buffer (60 mM Na₂HPO₄, 40 mM NaH₂PO₄, 10 mM KCl and 1 mM MgSO₄ heptahydrate). 200 μL of ONPG (4 mg ml⁻¹ o-nitrophenyl-β-D-galactopyranoside in 0.1 M KPO₄[pH 7.0]) were added and the was reaction timed. When a medium-yellow color developed, the reaction was stopped with 500 μL of 1 M Na₂CO₃. The OD_420nm of the supernatant was determined and the units were calculated with the equation: U = (OD_420nm × 1000)/(OD_660nm × time [in min]×volume of culture [in mL]). The assays were done in triplicate and normalization was performed by conversion of the Miller Units (absolute values) of one arbitrarily chosen WT construct or WT background as reference, set to 100%. All absolute values were then converted to relative values, averaged and the error was determined by calculation of the standard deviation (s.d.). Data are from three biological replicates.

Pools of >100,000 Tn mutants were isolated as kanamycin-aztreonam or kanamycin-colistin resistant clones in the NA1000 (WT), ΔtipN, ∆cpdR::Ω backgrounds, using the previously described protocol involving a mini-himar1 Tn encoding kanamycin resistance (Viollier et al., 2004). For each Tn pool, chromosomal DNA was extracted and used to generate a Tn-Seq library sequenced on an Illumina HiSeq 2500 sequencer (Fasteris, Geneva, Switzerland). The single-end sequence reads (50 bp) stored in FastQ files were mapped against the genome of the Caulobacter crescentus NA1000 (NC_011916) (Marks et al., 2010) genome and converted to BED files using BWA-MEM and bedtools BAM to BED tools, respectively, from the Galaxy server (https://usegalaxy.org/). The resulting BED file was imported into SeqMonk (http://www.bioinformatics.babraham.ac.uk/projects/seqmonk/) to build sequence read profiles. The initial quantification of the sequencing data was done in SeqMonk: the genome was subdivided into 50-bp probes, and for every probe, we calculated a value that represents a normalized read number per million. A ratio of the reads obtained in the ∆tipN or ∆cpdR strains to the WT reads was calculated for each 50-bp position. This file was used to generate the zoomed panels of the popA, rcdA and cpdR loci (Figure 1B) or the tipN locus (Figure 1—figure supplement 1A and B).

The citA::Tn insertion was identified using a modification of the kanamycin resistance suppressor screen (Radhakrishnan et al., 2010). Briefly, we screened for mini-himar1 Tn insertions that restore P_pilA firing to ΔtipN; ∆cpdR double mutant cells harboring the P_pilA-nptII transcriptional reporter, which confers kanamycin resistance to 20 μg mL⁻¹ when PpilA is fully active. The Tn encodes gentamycin resistance on plasmid pMar2xT7 delivered from E. coli S17-1 λpir (Liberati et al., 2006) to ΔtipN; ∆cpdR; pilA::P_pilA-nptII C. crescentus cells by selection on plates with gentamycin (1 μg mL⁻¹), kanamycin (20 μg mL⁻¹) and aztreonam (2.5 μg mL⁻¹, to counter-select E. coli). This screen gave rise to one isolate Φ40 with the desired resistance profile. The Tn insertion in Φ40 was mapped to the uncharacterized CCNA_01983 gene at nucleotide (nt) position 1061847 of the C. crescentus NA1000 genome sequence using arbitrarily primed PCR (Liberati et al., 2006).

Two independent clones of C. crescentus NA1000 freshly transduced with ∆citA::kan or citA::Tn were inoculated in 3 mL of PYE. Stationary phase cultures were diluted in 3 mL PYE to OD_600nm ~0.02. After 2 days, the four cultures were re-diluted to OD_600nm ~0.001 in 3 mL PYE. The phenotype of each strain was checked by phase-contrast microscopy and FACS analysis. Each culture was streaked on a PYE plate and one single colony from each culture was grown overnight and chromosomal DNA was extracted. Three suppressors were subjected to whole-genome sequencing. Library preparation and sequencing were performed by the Genomic platform iGE3 at the university of Geneva on a HiSeq 2500 with 50-bp paired-end reads. Data analysis to identify mutations was done using freebayes v1.1.0–3 (Garrison and Marth, 2012) against the C. crescentus NA1000 reference genome (NC_011916.1).

The overnight cultures were started in PYE or in M2G. The cultures were diluted to obtain an OD_600nm of 0.1 in PYE or M2G and were incubated at 30°C with continuous shaking in a microplate reader (Synergy H1, Biotek). The OD_600nm was recorded every 30 min for 30 hours. The graph represents the trend of the growth curve of three independent experiments.

Cells in exponential growth phase (OD₆₀₀0.3 to 0.6) were fixed 1:10 (vol/vol) in ice-cold 70% ethanol solution and stored at −20°C until further use. For rifampicin treatment, the mid-log phase cells were grown in the presence of 20 µg/mL rifampicin at 30°C for 3 hours. Cells were fixed as mentioned above. Fixed cells were centrifuged at 6200 g for 3 minutes at room temperature and washed once in FACS staining buffer (10 mM Tris-HCl, 1 mM EDTA, 50 mM Na-citrate, 0.01% Triton X-100 [pH 7.2]). Then, cells were centrifuged at 6200 g for 3 minutes at RT, and resuspended in FACS staining buffer containing RNase A (Roche) at 0.1 mg.mL⁻¹ for 30 minutes at RT. Cells were stained in FACS staining buffer containing 0.5 µM of SYTOX green nucleic acid stain solution (Invitrogen) and then analyzed using a BD Accuri C6 flow cytometer instrument (BD Biosciences, San Jose, CA, United States). Flow cytometry data were acquired and analyzed using the CFlow Plus v1.0.264.15 software (Accuri Cytometers Inc). A total of 20,000 cells were analyzed from each biological sample, performed in triplicates. The green fluorescence (FL1-A) parameters was used to determine cell chromosome contents. Flow cytometry profiles within one figure were recorded in the same experiment, on the same day with the same settings. The scales of the y- and x-axes of the histograms within one figure panel are identical. Each experiment was repeated independently three times and representative results are shown. The relative chromosome number was directly estimated from the FL1-A value of NA1000 cells treated with 20 µg/mL rifampicin for 3 hours at 30°C. Rifampicin treatment of cells blocks the initiation of chromosomal replication but allows ongoing rounds of replication to finish.

500 μL of an exponential Caulobacter or E. coli cells (OD_600nm = 0.4 and 0.8, respectively) were harvested with 20,000 g at 4°C for 5 minutes. Whole-cell extracts were prepared by resuspension of cell pellets in 75 µL TE buffer (10 mM Tris-HCl [pH 8.0] and 1 mM EDTA) followed by addition of 75 µL loading buffer 2X (0.25 M Tris [pH 6.8], 6% [wt/vol] SDS, 10 mM EDTA, 20% [vol/vol] glycerol) containing 10% (vol/vol) β-mercaptoethanol. Samples were normalized for equivalent loading using OD_600nm and were heated for 10 min at 90°C prior to loading.

Protein samples were separated by SDS–polyacrylamide gel electrophoresis and blotted on polyvinylidenfluoride membranes (Merck Millipore). Membranes were blocked overnight with Tris-buffered saline 1X (TBS) (50 mM Tris-HCl, 150 mM NaCl [pH 8]) containing, 0.1% Tween-20% and 8% dry milk and then incubated for an additional three hours with the primary antibodies diluted in TBS 1X, 0.1% Tween-20, 5% dry milk. The different polyclonal antisera to CitA (1:5000) and to CtrA (1:5000) were used. Primary antibodies were detected using HRP-conjugated donkey anti-rabbit antibody (Jackson ImmunoResearch) with ECL Western Blotting Detection System (GE Healthcare) and a luminescent image analyzer (Chemidoc MP, Biorad).

Recombinant CitA protein was expressed as an N-terminally His₆-tagged variant from pET28a in E. coli BL21(DE3)/pLysS and purified under native conditions using Ni²⁺ chelate chromatography. Cells were grown in LB at 37°C to an OD_600nm of 0.6, induced by the addition of IPTG to 1 mM for 3 hr, and harvested at 5000 RPM at 4°C for 30 minutes. Cells were pelleted and re-suspended in 25 mL of lysis buffer (10 mM Tris HCl [pH 8], 0.1 M NaCl, 1.0 mM β-mercaptoethanol, 5% glycerol, 0.5 mM imidazole Triton X-100 0.02%). Cells were sonicated in a water–ice bath (15 cycles of 30 s ON, 30 s OFF). After centrifugation at 5000 g for 20 minutes at 4°C, the supernatant was loaded onto a column containing 5 mL of Ni-NTA agarose resin (Qiagen, Hilden, Germany) pre-equilibrated with lysis buffer. The column was rinsed with lysis buffer, 400 mM NaCl and 10 mM imidazole, both prepared in lysis buffer. Fractions were collected (in 300 mM imidazole buffer, prepared in lysis buffer) and used to immunize New Zealand white rabbits (Josman LLC).

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Justine Collier, Sean Crosson, Martin Thanbichler, Michael Laub, Urs Jenal and Lucy Shapiro for materials, Julien Prados for help with Tn-Seq, ChIP-Seq and suppressors analyses, and Gaël Panis, Nicolas Kint for critical reading of the manuscript. We thank especially Benjamin Albert and Maksym Shyian from the David Shore lab for critical help with the phos-tag experiment. This work was supported by the Swiss National Science Foundation grant 31003A_182576 to Patrick H Viollier.

1. Received: September 27, 2019
2. Accepted: March 4, 2020
3. Accepted Manuscript published: March 9, 2020 (version 1)
4. Version of Record published: March 20, 2020 (version 2)

© 2020, Bergé et al.

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