Control of proline utilization by the Lrp-like regulator PutR in Caulobacter crescentus.

Cellular metabolism recently emerged as a central player modulating the bacterial cell cycle. The Alphaproteobacterium Caulobacter crescentus appears as one of the best models to study these connections, but its metabolism is still poorly characterized. Considering that it lives in oligotrophic environments, its capacity to use amino-acids is often critical for its growth. Here, we characterized the C. crescentus PutA bi-functional enzyme and showed that it is required for the utilization of proline as a carbon source. We also found that putA transcription and proline utilization by PutA are strictly dependent on the Lrp-like PutR activator. The activation of putA by PutR needs proline, which most likely acts as an effector molecule for PutR. Surprisingly, we also observed that an over-production of PutR leads to cell elongation in liquid medium containing proline, while it inhibits colony formation even in the absence of proline on solid medium. These cell division and growth defects were equally pronounced in a ΔputA mutant background, indicating that PutR can play other roles beyond the control of proline catabolism. Altogether, these findings suggest that PutR might connect central metabolism with cell cycle processes.

Introduction

The proline amino-acid is not only indispensable for protein synthesis, but also often for osmoprotection or heavy metal tolerance[1,2]. In addition, proline can be used as a source of nutrients by bacteria leaving in poor nutrient conditions or in proline-rich host microenvironments[3]. The first step of the proline utilization pathway is the conversion of proline into glutamate, to feed the tricarboxylic acid (TCA) or the nitrogen cycles. This conversion requires the sequential action of a proline dehydrogenase (PRODH) and of a Δ[1]-pyrroline-5-carboxylate dehydrogenase (P5CDH). The first of these two oxidation reactions produces reactive oxygen species as frequent by-products[4-7]. In most but not all bacteria, both activities are carried by a single bi-functional enzyme called PutA[3]. This reaction is not only necessary for proline utilization, but also for a diversity of biological processes by bacteria such as infection, symbiosis and redox homeostasis[3,7-9].

Considering the multiple roles of PutA inside cells, its activity must be tightly controlled. While the enzymes carrying PRODH and P5CDH domains are highly conserved, the control of their levels or activities is much more divergent. It is especially obvious in bacteria, where putA transcription is highly controlled by one or more transcription factors in response to nutrient availability[10,11]. In some bacteria, such as Escherichia coli or Salmonella thyphimurium, PutA has a DNA binding domain and can repress putA transcription[12,13], while other bacteria, such as Agrobacterium tumefaciens or Brucella abortus encode an independent regulator named PutR to control putA transcription[9,14]. putA is sometimes regulated by PutA and PutR such as in Rhodobacter capsulatus[15], illustrating the diversity of pathways regulating proline utilization in bacteria. The PutR regulator belongs to the Lrp/AsnC family of transcriptional regulators that are the most often responding to effector molecules[16,17]. This family includes not only specific regulators, but also global regulators controlling large regulons. Several members of this family respond to amino-acids, such as leucine for the best characterized leucine-responsive protein (Lrp)[16]. As for PutR, it has been shown to respond to proline in bacteria where it has been investigated[11,14,15]. Proline binding to PutR most likely induces a conformational change necessary for its binding to the putA promoter. It nevertheless remains unclear whether PutR proteins are always specific regulators of putA. Interestingly, one study indicated the PutR protein of Agrobacterium tumefaciens has the capacity to condense DNA into larger nucleoprotein complexes[18], suggesting that its mechanism of action is original in Alphaproteobacteria. It may control other genes or affect chromosome conformation as seen for some other Lrp-like factors[16].

Caulobacter crescentus is a well-characterized Alphaproteobacterium, as it serves as an excellent model system to study the bacterial cell cycle and the biology of the bacterial chromosome[19,20]. Interestingly, several studies recently uncovered interesting connections between metabolic pathways and cell cycle progression in this bacterium[21-26]. This might be particularly important for oligotrophic bacteria like C. crescentus, which thrives in aquatic environments such as lakes, rivers or oceans. In such environments, nutrients are rare and very diverse. Then, C. crescentus must be able to use a variety of different carbon sources and to modulate its cell cycle in response to nutrient availability as a strategy to survive in its natural environment. The central metabolism of C. crescentus and its control is however underexplored[27]. Still, it is known that this bacterium can use many sugars and amino-acids, including proline, as carbon sources[28-31].

In this study, we characterized the proline utilization pathway of C. crescentus and its control in response to proline availability. In addition, we uncovered a potential connection between proline catabolism and cell division through the Lrp-like PutR regulator.

Results

Caulobacter crescentus needs PutA for proline utilization

Most bacteria use a bi-functional PutA enzyme to catabolize the conversion of proline into glutamate, the first step in proline utilization[3]. In order to study how C. crescentus uses proline as a source of nutrients, we searched for a putA homolog in its genome and found the CCNA_00846 protein that shares ~47% of identity with bi-functional PutA proteins from other Alphaproteobacteria[9,14]. It encodes a protein carrying two domains: one with similarities with the PRODH domain and the other one with similarities with the P5CDH domain, as expected for a bi-functional enzyme[3]. We constructed a complete ΔputA deletion strain and found that the growth rate (Supplementary Fig. S1) and the morphology (Supplementary Fig. S2) of mutant cells cultivated in complex peptone-yeast extract (PYE) medium (in which amino-acids serve as the primary carbon source) were unaffected, compared to isogenic wild-type (WT) cells. We then tested its capacity to grow in minimal media with proline (M2P) or glucose (M2G) as the sole carbon source. As expected, the ΔputA and its isogenic wild-type (WT) strain could both grow at a similar rate in liquid (Fig. 1A) or solid (Fig. 1B) M2G. We also observed that the WT strain could grow on M2P, but at a lower rate than on M2G, showing that glucose is a better carbon source than proline for C. crescentus. As for the ΔputA strain, we could not detect any growth in liquid (Fig. 1A) or on solid (Fig. 1B) M2P media, showing that PutA is required for proline utilization by C. crescentus. This result strongly suggests that C. crescentus PutA can convert proline into glutamate as previously shown for PutA homologs from other Alphaproteobacteria[9,11,14,15].

Figure 1

PutA and PutR are required for proline utilization as a carbon source. (A) Growth of WT (NA1000), ΔputA (JC1695) and ΔputR (JC1040) strains in liquid M2 minimal media with 0.2% glucose (M2G, dashed lines) or 10 mM proline (M2P, solids lines) as carbon sources. Each culture was cultivated over-night in M2G medium and re-diluted into M2G or M2P media at time 0. Growth was tracked by measuring the OD660nm for ~70 hours. Each curve represents the mean of six independent experiments. Error bars correspond to standard deviations. (B) Growth of WT, ΔputA and ΔputR colonies on solid M2 minimal medium with 0.2% glucose (M2G) or 10 mM proline (M2P) as carbon sources. Representative plates (from three independent experiments) incubated for 6 days are shown here.

PutR is critical for putA transcription and proline utilization

C. crescentus must control PutA levels or activity to make sure that it does not catabolize proline when it grows in a medium that does not contain proline, in order to maintain intracellular levels of proline that are sufficient for efficient protein synthesis. Interestingly, a gene (CCNA_00847) encoding a protein sharing ~40% of identity with PutR from other Alphaproteobacteria[9,14] lies next to the putA gene but in an opposite orientation, suggesting that it may encode a regulator of putA transcription. To test if PutR regulates putA transcription, we cloned the putA promoter region upstream of the lacZ gene and introduced this PputA-lacZ reporter into a WT, a ΔputR deletion strain and a strain over-expressing putR. We then used these strains to perform β-galactosidase assays to evaluate the impact of PutR on putA transcription in complex PYE medium. We observed that the putA promoter was ~7-fold less active in ΔputR cells than in WT cells (Fig. 2A), reaching very basal levels that are most likely not sufficient for proline utilization. Indeed, we also observed that ΔputR cells could not grow in M2P medium containing proline as the only carbon source, while they could grow as WT cells in M2G (Fig. 1) or PYE media (Supplementary Fig. S1). In addition, the activity of the putA promoter was induced ~2-fold upon putR-overexpression when cells were cultivated in rich medium for 90 minutes (Fig. 2B). All together, these results show that PutR is a strong activator of putA transcription and that PutR is thus required for proline utilization by C. crescentus.

Figure 2

PutR is a proline-dependent activator of the putA but not of the relB1 promoters. Promoter activities in Miller Units (MU) were measured by β-galactosidase assays from cells cultivated in exponential phase and carrying the placZ290-putAP (PputA-lacZ) or placZ290-relB1P (PrelB1-lacZ) plasmids. Error bars correspond to standard deviations from minimum six independent experiments. Statistical analysis were performed using student t-tests. Stars (*) highlight significant (p < 0.0005) but also relevant differences between two strains. (A) Activity of the putA promoter in WT (NA1000) or ΔputR (JC1040) strains cultivated in PYE rich medium. (B) Activity of the putA promoter in a WT strain carrying pBXMCS6 (empty vector) or pX-putR. Cells were cultivated over-night and re-diluted into PYE + 0.2% glucose (PYEG). Once cells reached exponential phase again, 0.3% xylose was added into half of the culture (PYEGX) to induce the expression of putR from pX-putR. β-galactosidase assays were done 90 minutes after. (C) Activity of the putA promoter in WT or ΔputR strains cultivated in M2 minimal medium containing 0.2% glucose (M2G) supplemented or not with 10 mM proline. (D) Activity of the putA promoter in a WT strain carrying pBXMCS6 (empty vector) or pX-putR. Cells were cultivated over-night in M2G and re-diluted into M2G supplemented or not with proline. Once cells reached exponential phase again, 0.3% xylose was added into half of the culture to induce the expression of putR from pX-putR. β-glactosidase assays were done 180 minutes after. (E) Activity of the relB1 promoter in WT or ΔputR strains cultivated in M2 minimal medium containing 0.2% glucose (M2G) supplemented or not with 10 mM proline.

PutR is a proline-responsive activator of putA

PutR belongs to the Lrp/AsnC family of transcription factors that often use effector molecules, such as amino-acids, to modulate their activity[16]. Considering that PutR regulates the proline utilization enzyme PutA, we tested if proline may affect the capacity of PutR to activate putA transcription. We grew the WT, ΔputR and putR-over-expression strains carrying the PputA-lacZ reporter in M2G media supplemented or not with proline and performed β-galactosidase assays. In the WT strain, the activity of the putA promoter was ~8-fold higher when proline was added into the medium, showing that proline induces putA transcription (Fig. 2C). Consistent with the proposal that PutR senses proline levels, we observed that proline could not induce the putA promoter in the absence of PutR. Interestingly, proline activated the putA promoter even more efficiently (~11-fold) when putR was over-expressed (Fig. 2D). Overall, these observations show that proline is required for the activation of putA transcription by PutR.

The overproduction of PutR in proline-containing medium leads to cell elongation in a PutA-independent manner

Surprisingly, we observed that the overexpression of putR under the control of the xylX promoter on the medium-copy number vector pBXMCS6 leads to cell morphology defects. Wild-type cells over-expressing putR appeared significantly elongated compared to control cells carrying the empty vector, but only when cultivated in the presence of proline in minimal medium (Fig. 3) or in complex medium (Supplementary Fig. S3). This observation suggests that PutR inhibits cell division in liquid cultures. Since this phenotype is dependent on the presence of proline, we hypothesized that it might be connected with proline catabolism due to excessive transcription of putA when PutR is over-produced. Indeed, previous studies indicated that proline catabolism produces hydrogen peroxides, which can influence redox homeostasis in certain bacterial species[5,6]. Considering that oxidative stress may lead to DNA or protein damage and that such damage can lead to cell cycle arrests[32,33], it could have been an explanation for the phenotype of cells over-expressing putR. To test this hypothesis, we looked at the effect of PutR overproduction in ΔputA cells that cannot catabolize proline. We observed that the cell division defect was similar in wild-type and ΔputA cells (Fig. 3), indicating that PutR can inhibit cell division independently of its function as a transcriptional activator of putA.

Figure 3

Excess of PutR leads to cell division defects in a proline-dependent manner. NA1000 (WT) and isogenic ΔputA cells, or CB15 (WT’) and isogenic ΔrelBE1 cells, carrying the pBXMCS6 (pX; empty vector) or the pX-putR plasmid were cultivated in M2G and diluted the next morning into M2G supplemented or not with proline. Once cultures reached an OD660nm of ~0.25, 0.3% xylose was added to induce the expression of putR from pX-putR over-night. (A) Cells were then imaged by phase-contrast microscopy and representative images from three independent experiments are shown in this figure. (B) Cell size distributions of WT and ΔputA cells carrying pX or pX-putR were compared. The bold grey line indicates the median medial axis length of cells. The limits of the box are the first and third quartiles. The whiskers are the 5% and 95% quantiles. The values were normalized so that the median length of the WT cells carrying pX equals 1 arbitrary unit (a.u.) to facilitate comparisons.

The overproduction of PutR inhibits colony formation in a PutA-independent manner

In addition to the observed cell division defect, we noticed that wild-type cells over-expressing putR while cultivated on plates could rarely form colonies (Fig. 4). When growth could nevertheless be detected, colonies appeared as much smaller than colonies of cells carrying the empty vector. Interestingly, this additional phenotype associated with the over-production of PutR was neither dependent on the presence of proline in the medium, nor on the capacity of cells to catabolize proline using PutA. Thus, it appeared as independent of the effect of PutR on putA regulation or on cell division.

Figure 4

Excess of PutR inhibits colony formation and growth in a proline-independent manner. NA1000 (WT) and isogenic ΔputA cells, or CB15 (WT’) and isogenic ΔrelBE1 cells, carrying the pBXMCS6 (empty vector) or the pX-putR plasmid were cultivated in M2G and diluted the next morning into M2G supplemented or not with 10 mM proline. Once cultures reached stationary phase, 10-fold serial dilutions in M2G medium were prepared and 5 μL of each dilution was spotted onto M2GA plates containing 0.3% xylose (M2GAX) to induce the expression of putR from pX-putR over-night. Proline was also added into plates when indicated. Representative images from three independent experiments are shown in this figure.

PutR and proline levels do not influence relBE1 expression

Interestingly, we noticed that the relBE1 genes, encoding a functional toxin-antitoxin (TA) module[34], are located right after the putA gene on the C. crescentus genome (Fig. 5A). In addition, an RNA-sequencing experiment suggested that putA and relBE1 may belong to the same polycistronic transcript[35,36]. Then, an excess of PutR might result in an excessive transcription of relBE1, potentially leading to RelE1-mediated toxicity on plates or to cell division defects. It is however important to mention that the relBE1 genes are also transcribed from their own promoter located just upstream of the relB1 gene (Fig. 5A)[34]. To test if this second promoter is dependent on PutR and/or proline like the putA promoter, we constructed a transcriptional fusion between the relB1 promoter and the lacZ gene, and introduced this construct into wild-type and ΔputR strains. These strains were then cultivated in M2G medium supplemented or not with proline and promoter activity was evaluated by β-galactosidase assays. We found that the relB1 promoter was neither dependent on PutR, nor on the presence of proline in the culture medium (Fig. 2E). If relBE1 and putA do belong to the same operon[35], it was however still possible that relBE1 transcription might be affected by PutR and proline through their regulation of the putA promoter (Fig. 2C,D). To test this possibility, we performed quantitative reverse transcription PCR (qRT-PCR) with four different probes allowing us to distinguish transcripts carrying putA, relB1, relE1 or the intergenic region between putA and relB1 (IG) independently (Fig. 5A). First, these experiments confirmed that putA transcription is strongly dependent on PutR and proline (Fig. 5B), as previously shown using transcriptional reporters (Fig. 2C,D). Second, it showed that the IG region is much less transcribed than the putA gene in all tested conditions (Fig. 5B), indicating that transcription starting at the putA promoter mostly ends before the relB1 gene (Fig. 5A). All together, these qRT-PCR results demonstrate that the relBE1 TA module is only transcribed from the PutR-independent relB1 promoter under standard growth conditions. Then, it is unlikely that the overproduction of PutR will affect relBE1 expression, leading to defects in colony formation (Fig. 4) or in cell division (Fig. 3). Supporting this proposal, we observed that the over-production of PutR in a relBE1 deletion strain still inhibits colony formation on solid media (Fig. 4) and cell division in liquid media (Fig. 3). All together, these results suggest that PutR has other targets than the putA promoter and that PutR may not always need proline as an effector.

Figure 5

putA and relBE1 are mostly transcribed from independent and differentially regulated promoters. (A) Schematic showing the organization of the putA chromosomal region including the relBE1 genes. Grey arrows represent open reading frames. The blue and pink arrows represent the putA and the relB1 promoters, respectively. Green lines show the position of each region (160–200 base pairs) amplified during quantitative RT-PCR (qRT-PCR) assays. The grey shade highlights the IG region amplified by qRT-PCR. Blue and pink lines are a representation of transcripts (mRNA) detected by qRT-PCR and controlled by the putA or relB1 promoters, respectively, when WT cells were cultivated in minimal medium containing 0.2% glucose (M2G) supplemented or not with proline. The weight of blue and pink lines is proportional to the levels of detected transcripts. (B) Transcripts from the putA-relBE1 region detected by qRT-PCR from WT (NA1000) or ΔputR (JC1040) cells cultivated in M2G supplemented or not with 10 mM proline. On each graph, the relative abundance of the relB1 mRNA was arbitrarily set to 1 (A.U.). The other values represent the abundance of specific transcripts relative to the level of relB1 mRNA in the same strain and growth condition. Error bars represent standard deviations from minimum three independent experiments performed in triplicates.

Discussion

This study aimed at characterizing proline catabolism and its control in C. crescentus. Such a control is critical to ensure that cells can use proline as carbon or nitrogen sources, but that they do not degrade proline when endogenous levels are too limiting. This is all the more important for environmental bacteria like C. crescentus that thrive in oligotrophic environments where amino-acids often serve as major carbon sources. Also, the catabolism of proline by PutA enzymes can generate hydrogen peroxide, which can be deleterious to cells[4-7]. We showed that the PutR regulator plays a key role in this process and unexpectedly also discovered that it may affect other PutA-independent cellular functions.

The putA gene of C. crescentus encodes a bi-functional proline dehydrogenase (Figs 1 and 6). Although indispensable for growth on minimal media containing proline as the only carbon source (Fig. 1), we observed that the absence of PutA does not affect growth or cell morphology in complex PYE media where amino-acids serve as major carbon sources (Supplementary Figs S1 and S2). However, C. crescentus may frequently find conditions where proline becomes an important source of nutrients in its natural oligotrophic environment. Indeed, Alphaproteobacteria have been shown to be often exposed to environments where proline catabolism by PutA becomes indispensable, such as during nodulation or infection by Sinorhizobium meliloti and B. abortus, respectively[8,9].

Figure 6

Model for the various roles of PutR in C. crescentus. The activation of putA transcription by PutR is necessary for the catabolism of proline into glutamate, corresponding to the first step of the proline utilization pathway. In addition, excess of PutR can inhibit cell division and colony formation through still unknown pathways (dashed arrows). PutR requires proline to activate putA transcription and inhibit cell division, while proline is dispensable for its effect on colony formation.

As in many Alphaproteobacteria, the putA gene of C. crescentus is located immediately next to a gene that is divergently transcribed and which encodes a putative DNA binding protein of the Lrp/AsnC family. We found that this gene encodes a strong activator of putA transcription in the presence of proline (Figs 2 and 5) and thus named it PutR for proline utilization regulator. Without PutR, the levels of PutA are so low that C. crescentus can no more use proline as a carbon source (Fig. 1). The fact that PutR requires proline to activate putA transcription (Figs 2 and 5) suggests that proline acts as an effector molecule to stimulate PutR activity. Transcription factors from the AsnC/Lrp family often require such effector molecules to bind to some of their promoter targets, most likely through conformational changes[16] that may also take place with C. crescentus PutR.

Our observation that the putA promoter is efficiently activated by proline but still efficiently repressed without proline (Fig. 2C), suggests that this promoter could be used as a useful genetic tool for inducible protein expression in C. crescentus cells cultivated in minimal media. Compared to the two other promoters currently available for such applications (from the xylX and vanA genes)[37], the putA promoter appears to have properties similar to the xylX promoter[38] in terms of efficiency of induction and repression. Moreover, the putA promoter does not respond to xylose levels (Fig. 2B), making the combined use of xylX and putA promoters, to induce the expression of two different proteins, potentially feasible.

Interestingly, several members of the Lrp/AsnC family of transcription factors are so-called global regulators controlling large regulons. The best characterized example is the leucine-responsive protein Lrp of E. coli that directly regulates ~130 genes encoding proteins involved in various processes such as metabolism, pili synthesis or adhesion to host cells[39-41]. Other members of this family are more specific regulators, like PutR homologs from a few other Alphaproteobacteria such as A. tumefaciens, S. meliloti and R. capsulatus[9,11,14,15]. In C. crescentus, we observed that an overproduction of PutR leads to cell division and colony formation defects, which are independent of the activation of putA by PutR (Figs 3 and 4). This observation indicates that PutR may have other activities in C. crescentus, despite no apparent defects in growth or cell morphology for ΔputR cells cultivated with or without proline (Fig. 1 and Supplementary Figs S1 and S2). Future experiments should notably aim at determining under which conditions C. crescentus may use this other activity to block or slow down cell division in response to proline levels. Interestingly, recent observations made comparing the phenotypes of ΔputA and ΔputR strains of B. abortus also lead to conclude that PutR may have additional functions beyond simply regulating putA transcription, such as adaptation to oxidative stress[9].

Interestingly, we observed that the inhibition of cell division by PutR was dependent on proline (Fig. 3), while the inhibition of colony formation on solid media was not (Fig. 4). Also, the proline and PutR-dependent inhibition of cell division (Fig. 3) in liquid media is not associated with a drop of cell viability as shown by live/dead staining assays and microscopy (Supplementary Fig. S4). Then, PutR may carry minimum two other functions unrelated with putA activation, with only one of these that is dependent on proline (Fig. 6). Another possibility is that the putA-independent target(s) of PutR have more impact when C. crescentus is cultivated on solid rather than in liquid media. Importantly, we ruled out the possibility that PutR inhibits cell division through a polar effect on the transcription of the downstream relBE1 genes (Figs 3 and 5) encoding a functional toxin-antitoxin system in C. crescentus[34]. Then, how PutR can inhibit cell division in liquid medium or colony formation on plates through direct or indirect effects remains to be clarified (Fig. 6). As a DNA binding protein, it may bind to other promoter regions to regulate the transcription of other genes or, alternatively, have more general effects on chromosome condensation as other members of the Lrp/AsnC family of transcription factors[16,18]. Consistent with this last possibility, results from flow cytometry experiments looking at the DNA content of elongated C. crescentus cells over-expressing PutR, suggest that the elongation of DNA replication may be slowed down or frequently arrested in these cells (Supplementary Fig. S5). This may then interfere with cell division through SOS-dependent or SOS-independent mechanisms that tend to block late steps of the cell division process in C. crescentus[32,33]. In agreement with this proposal, we found that the FtsK core divisome protein could still localize at mid-cell in elongated cells over-expressing PutR (Supplementary Fig. S3), suggesting that cell constriction, rather than core divisome assembly is inhibited when PutR is over-expressed. Considering that the proteolysis of the CtrA response regulator, which regulates the initiation of DNA replication and the transcription of several cell division proteins in C. crescentus[42], is induced by proline uptake in the intracellular Alphaproteobacterium Ehrlichia chaffeensis[43], we also checked the intracellular levels of CtrA in elongated C. crescentus cells over-expressing PutR, but observed no significant difference compared to control cells (Supplementary Fig. S6). Then, although a common point between C. crescentus and E. chaffeensis is that proline can affect their cell cycles, it appears to do so using different mechanisms in different Alphaproteobacteria.

Our findings on PutR are reminiscent of so-called “moonlighting” proteins that are multifunctional proteins that perform multiple autonomous functions without partitioning these functions into different protein domains[44]. Interestingly, moonlighting proteins are the most often involved in the glycolytic pathway or in the TCA cycle. An example found in Alphaproteobacteria is the GdhZ moonlighting enzyme that not only converts glutamate into α-ketoglutarate, but also inhibits FtsZ polymerization to modulate cell division in response to glutamate levels[21,45]. Although an excess of PutR in cells exposed to high proline levels may lead to an imbalance in glutamate intracellular levels through PutA, we ruled out the possibility that this is responsible for the observed cell division defect since PutR-overexpressing cells lacking putA were still elongated (Fig. 3).

Altogether, one of our surprising finding was that C. crescentus PutR may connect the catabolism of proline with other cellular functions such as cell division, but in a PutA-independent manner (Fig. 6). Recent discoveries also highlighted other unexpected connections between cell division and the central metabolism in diverse bacteria[25]. Beyond the example of the GdhZ enzyme[21], it has, for example, been shown that intracellular α-ketoglutarate levels can influence peptidoglycan synthesis and cell division[22] or that the transcription of the ftsZ cell division gene is influenced by the glutamine-sensitive phosphotransfer system (PTSNtr)[23,24] in C. crescentus. Also, the pyruvate metabolite has been shown to modulate Z-ring assembly for the division of the Gram-positive Bacillus subtilis bacterium[46]. Such connections may play an important role in coordinating bacterial growth with cell division, potentially participating to cell size homeostasis[26].

Materials and Methods

Bacterial strains and growth conditions

C. crescentus strains used in this study are described in Table 1. The TOP10 E. coli strain (Invitrogen, USA) was used for plasmid constructions. C. crescentus was cultivated at 28 °C (liquid cultures, with constant shaking) or 30 °C (solid media) in peptone yeast extract (PYE) complex medium or in M2 minimal medium containing 0.2% glucose (M2G), 10 mM proline (M2P) or both (M2GP). For solid media, 1.5% agar (A) was added. E. coli was cultivated at 37 °C in LB broth or on LBA. When required, antibiotics where used at the following concentrations (µg.mL−1) in liquid media (LM) or solid media (SM) for C. crescentus: oxytetracycline (OxyTet):1 in LM and 2 on SM; chloramphenicol (Cam): 1 in LM and on SM; kanamycin (Km): 5 in LM and 25 on SM; spectinomycin (Spec): rich medium = 25 in LM and 100 on SM, minimal medium = 400 in LM and 800 on SM; streptomycin (Strep): rich medium = 5 in LM and on SM, minimal medium = 20 in LM and 40 on SM.

Table 1

Strains and plasmids used in this study.

Strain	Genotype	Description	Source or reference
Caulobacter crescentus
CB15	WT’	Wild-type Caulobacter crescentus strain	[51]
NA1000	WT	CB15N wild-type strain derivative of CB15	[28]
JC1040	ΔputR	NA1000 strain with deletion of putR	This study
JC1695	ΔputA	NA1000 strain with in frame deletion of putA	This study
FC917	ΔrelBE1	CB15 strain with in frame deletion of relBE1	[34]
LS4200	xylX::ftsK-gfp	NA1000 strain with ftsK-gfp under the control of the native xylX promoter	[52]
Plasmids
pNPTS138		Suicide vector containing the sacB gene, KmR	D. Alley, unpublished
pNPTS138-putA		Carrying the regions upstream and downstream of putA to create the ΔputA strain	This study
pNPTS138-putR		Carrying the regions upstream and downstream of putR
pBOR		Vector carrying a Spectinomycin/Streptomycin cassette (Ω), derivative of pHP45Ω	[53]
pNPTS138-putR:Ω		Carrying the Ω cassette in between the regions upstream and downstream putR to create the ΔputR strain	This study
pRXMCS6		Low copy number vector with the xylX promoter (Pxyl), CamR	[37]
pRX-putR		putR gene under the control of Pxyl inserted into pRXMCS6 plasmid	This study
pBXMCS6		Medium copy number vector with the xylX promoter (Pxyl), CamR	[31, 37]
pX-putR		putR gene under the control of Pxyl inserted into pBXMCS6 plasmid	This study
placZ290		Low copy number vector carrying the lacZ gene, OxytetR	[54]
placZ290-putAP		lacZ reporter gene controlled by the putA promoter on the placZ290 plasmid	This study
placZ290-relB1P		lacZ reporter gene controlled by the relB1 promoter on the placZ290 plasmid	This study

Plasmid constructions

Plasmids used in this study are described in Table 1 and their construction is described below. All PCR amplifications were done using NA1000 genomic DNA and the KOD hot start DNA polymerase (Merck, Germany).

Construction of placZ290-putAP and placZ290-relB1P

The promoter regions of putA (445 base pairs upstream of the annotated translational start site) or relB1 (568 base pairs upstream of the annotated translational start site) were amplified using primers described in Supplementary Table S1, carrying EcoRI, PstI or XbaI restriction sites. PCR products and the placZ290 vector were digested by EcoRI and XbaI or PstI. Promoters were then ligated into the vector, giving placZ290-putAP and placZ290-relB1P.

Construction of pX-putR and pRX-putR

The putR coding sequence (464 base pairs) was amplified using primers described in Supplementary Table S1, carrying NdeI, EcoRI or NheI restriction sites. The amplified putR sequence and the pBXMCS6 or the pRXMCS6 vectors were digested by NdeI and NheI or EcoRI. The putR sequences and the vectors were then ligated together, giving pX-putR and pRX-putR. With these plasmids, putR expression is controlled by the xylX promoter[31], which is induced in the presence of xylose in the medium and repressed in the presence of glucose in the medium.

Construction of pNPTS138-putA, pNPTS138-putR and pNPTS138-putR:Ω

From 500 to 730 base pairs upstream (UP) and downstream (DOWN) of the putA and putR coding sequences were amplified using primers described in Supplementary Table S1. PCR products were digested by EcoRI, BamHI and NheI (putA) or PstI, BamHI and NheI (putR). Digested UP and DOWN sequences were then both ligated into a pNPTS138 vector digested with the same enzymes, giving pNPTS138-putA and pNPTS138-putR. An Ω cassette extracted from pBOR digested by BamHI and encoding Spec/Strep resistances was then introduced at the BamHI restriction site between the UP and the DOWN sequences of putR, giving pNPTS138-putR:Ω.

Strain constructions

Plasmids were introduced into NA1000 or CB15 C. crescentus strains as published in[47].

To construct the ΔputA deletion strain, the pNPTS138-putA plasmid was integrated into the C. crescentus chromosome at the putA locus by single homologous recombination, selecting for Km-resistant colonies. The resulting strain was grown to stationary phase in PYE medium lacking Km. Cells were plated on PYEA + sucrose 3% and incubated at 28 °C or 30 °C. Single colonies were picked and transferred in parallel onto PYEA plates containing or not Km. Km-sensitive clones, which had lost the integrated plasmid due to a second recombination event were isolated. Colonies were further tested for their capacity to grow on M2GA medium but not on M2PA medium.The ΔputA deletion was then verified by colony-PCR using specific primers.

To construct the ΔputR deletion strain, the pNPTS138-putR:Ω plasmid was integrated at the putR locus of the chromosome of a C. crescentus NA1000 strain by single homologous recombination, selecting for Km-resistant colonies. The pRX-putR plasmid was then introduced into this strain. The resulting strain was grown to stationary phase in PYE medium supplemented or not with 0.3% xylose and Cam, but lacking Km. Cells were plated on PYEA + sucrose 3% + xylose 0.3% or 0.1% + Cm and incubated at 28 °C. Single colonies were picked and transferred in parallel onto PYEA + xylose 0.3% + Cam + Strep + Spec or onto PYEA + xylose 0.3% + Cam + Km. Km-sensitive and Spec/Strep-resistant clones, which had lost the integrated plasmid due to a second recombination event were isolated. These were supposedly ΔputR::Ω. The ΔputR::Ω deletion was then verified by colony-PCR using specific primers and transduced into a NA1000 WT strain using ΦCR30 phages[48] selecting for Strep/Spec-resistances, giving the so-called ΔputR strain.

β-galactosidase assays

β-galactosidase assays were done using standard procedures as previously described[49].

RNA extraction

Total RNA were extracted from cultures at an OD660 of 0.6, using the RNeasy miniprep kit (Qiagen, GER) including the recommended DNase I treatment. A second DNase I treatment was done to remove the remaining gDNA using the Turbo DNase I (Ambion, USA) at 37 °C for 30 minutes and RNA were then purified using the RNeasy MinElute Kit (Qiagen, GER). RNA samples were tested for purity by PCR. RNA quantity was measured using a Nanodrop fluorospectrometer, while quality was checked by agarose gel electrophoresis.

Quantitative Real-Time PCR (qRT-PCR) analysis

cDNA were synthetized from 5 µg of total RNA using the Superscript III reverse transcriptase and 250 ng of random hexamers according to the manufacturer’s instructions (Invitrogen, USA). The qRT-PCR was performed using the GoTaq qPCR master mix (Promega, USA), 2 μl of cDNA (diluted 1:100) and 0.5 µM of specific primers (Supplementary Table S1) for each gene. The enzyme was activated by heating at 95 °C for 2 minutes. Then, the cDNA was amplified by qPCR (40 cycles) with a denaturation step at 95 °C for 15 seconds, an annealing step at 60 °C for 60 seconds. A final step of dissociation was achieved by increasing the temperature by 0.3 °C every 5 seconds from 60 °C to 95 °C. qPCRs were performed in triplicates using a Rotor-GeneQ instrument (Qiagen, GER). Melting curves were analyzed and the presence of a single peak was checked. For each pair of primers, primer efficiency was evaluated using standard curves from gDNA from 5 ng/µL to 0.001 ng/µL. Absolute quantifications x were determined using the following formula: x = (y − b)/m; y corresponds to the Ct value, while m and b are automatically calculated using standard curves.

Microscopy

Cells were immobilized on slides using a thin layer of PYE + 1.5% agar and imaged immediately. Phase contrast microscopy images were taken with a Plan-Apochromat 100X/1.45 oil Ph3 objective on an AxioImager M1 microscope (Zeiss) with a Prime-95B back-illuminated CMOS camera (Photometrics) controlled by the VisiView software (Visitron Systems, Germany). Images were processed using Adobe Photoshop. The MicrobeJ software[50] was used to measure the length of cells on images using default parameters. Measurements from 100–200 cells were used to evaluate cell size distributions.

Supplementary information