A transcriptional regulator linking quorum sensing and chitin induction to render Vibrio cholerae naturally transformable

Media and growth conditions

Vibrio cholerae and Escherichia coli strains were grown in Luria both (LB) medium at 30°C with shaking, unless otherwise stated. Antibiotics were added for plasmid maintenance or transformants/transconjugants selection at concentrations of 50 or 100 μg/ml for ampicillin, 75 μg/ml for kanamycin and at 50 μg/ml for gentamicin. Thiosulfate-citrate-bile salts-sucrose agar plates were used to counter-select E. coli after bi-/triparental mating with V. cholerae; the thiosulfate-citrate-bile salts-sucrose agar plates were prepared according to the manufacturer’s instructions (Sigma-Aldrich/Fluka, Buchs, Switzerland). NaCl-free LB medium containing 6% sucrose was prepared for sucrose-based sacB-counter-selection.

Bacterial strains and plasmids

The V. cholerae strains and plasmids used in this study are indicated in Table 1. The E. coli strains DH5α (43) and S17-1λpir (44) were used for cloning and as a donor strain in bacterial mating experiments, respectively. E. coli Origami 2^TM (DE3) pLysS (Novagen) was used as a host for HapR protein expression.

Construction of V. cholerae mutant strains

The gene qstR (VC0396) was deleted from the parental strain A1552 using the gene disruption method based on the counter-selectable plasmid pGP704-Sac28, as previously described (14). The oligonucleotides used to construct the respective plasmid are indicated in Supplementary Table S1.

Construction of plasmids used to investigate the putative HapR binding site upstream comEA

Most of the plasmids used in this study were derived from the plasmid pBR322 (39) (Table 1). Initially, pBR322 was modified through the partial deletion of the tetracycline resistance cassette and the constitutive promoter P_Tet, resulting in the plasmid pBR-Tet_MCSII (Table 1). The primers P[VC1917]-GFP#1-NotI and VC1917-down-NotI (Supplementary Table S1) were used in polymerase chain reaction (PCR) to amplify comEA preceded by 900 bp of its upstream region. The genomic DNA (gDNA) from the V. cholerae strain A1552 was used as a template. The PCR fragment was digested with NotI and cloned into the equally digested vector pBR-Tet_MCSII, resulting in the plasmid pBR-[own]comEA.

The plasmid pBR-[own]comEA was digested with AatII and BglII to shorten the upstream region of comEA. Self-ligation resulted in the plasmid pBR-[P_wt]comEA, carrying comEA and 200 bp of its upstream region (Table 1). Site-directed mutagenesis using inverse PCR on plasmid pBR-[P_wt]comEA generated the plasmids pBR-[P_{mut L}]comEA, pBR-[P_{mut R}]comEA and pBR-[P_{mut L/R}]comEA, carrying mutations in the putative HapR binding site upstream comEA (see oligonucleotides in Supplementary Table S1).

Construction of complementing plasmids and plasmid pBAD-hapR-N-Strep

The genes hapR, comEA and qstR were amplified with the respective primers indicated in Supplementary Table S1 using gDNA from V. cholerae strain A1552 as a template. The NcoI and EcoRI-digested PCR products were cloned into the equally digested plasmid pBAD/Myc-HisA to generate plasmids pBAD-hapR, pBAD-comEA and pBAD-qstR (Table 1). The Strep-tagII® encoding sequence was added to hapR through inverse PCR using the oligonucleotides hapR-N-Strep-fw and hapR-N-Strep-bw (Supplementary Table S1) and pBAD-hapR as a template, yielding plasmid pBAD-hapR-N-Strep.

Construction of plasmids containing the putative HapR binding sites upstream of dns and qstR

The short DNA segments upstream of dns and qstR, respectively (30–50 bp), were introduced as overhangs in the oligonucleotides indicated in Supplementary Table S1. The primers were used to amplify pBR-Tet_MCSI through inverse PCR to generate the plasmids pBR-[P_dns-100-50], pBR-[P_dns-50-1], pBR-[P_qstR-150-120] and pBR-[P_qstR-150-102] (Table 1). For the electrophoretic mobility shift assay (EMSA) experiments, the respective inserted DNA regions, flanked by DNA sequences derived from the plasmid (∼200-bp fragments), were PCR amplified using the primers pBR-TET_MCS-before and pBR-TET_MCS-after (Supplementary Table S1). The plasmid pBR-[P_qstR-150-102]_mut_AgeI, carrying mutations in the promoter region −150 to −102 upstream of the start codon of qstR (P_qstR; see Figure 6D), was constructed through inverse PCR using the oligonucleotides PVC0396_AgeI_fw and PVC0396_AgeI_bw and the plasmid pBR-[P_qstR-150-102] as a template.

Natural transformation assay (chitin-dependent and chitin-independent)

Chitin-dependent natural transformation assays were done on chitin flakes as previously described by Marvig and Blokesch (37,45). Chitin-independent induction of natural competence and scoring of transformants was performed using V. cholerae strains carrying an inducible chromosomal copy of TfoX as described by Lo Scrudato and Blokesch (8). Statistical analyses of transformation data were done on log-transformed data (46) using a two-tailed Student’s t-test.

Sodium dodecyl sulfate–polyacrylamide gel electrophoresis and western blotting

The proteins were separated through sodium dodecyl sulfate–polyacrylamide gel electrophoresis and then either stained for protein using Coomassie blue or subjected to western blotting as previously described (8). Primary antibodies against HapR (8) and against the Strep-tagII sequence (α-Strep-MAB classic, IBA GmbH, Göttingen, Germany) were diluted at 1:10 000 and 1:1000, respectively. Goat anti-rabbit horseradish peroxidase (HRP) (diluted 1:20 000; Sigma-Aldrich, Switzerland) and goat anti-mouse HRP (diluted 1:5000; Sigma-Aldrich, Switzerland) were used as secondary antibodies. Lumi-Light^PLUS western blotting substrate (Roche, Rotkreuz, Switzerland) or Western Lightning®-ECL (PerkinElmer, Schwerzenbach, Switzerland) were used as HRP substrates, and the luminescence signals were detected using chemiluminescence-detecting films (Amersham Hyperfilm ECL, GE Healthcare via VWR, Dietikon, Switzerland).

Purification of HapR-N-Strep

HapR-N-Strep protein was expressed in the E. coli strain Origami^TM 2 (DE3) pLysS (Novagen) carrying the plasmid pBAD-hapR-N-Strep. The cells were grown in LB medium at 30°C until the culture reached an optical density of ∼0.8 at 600 nm. At that time, the expression was induced on the addition of 0.2% arabinose to the culture medium. After induction, the bacteria were further cultivated at 16°C overnight. The cells were collected through centrifugation and resuspended in lysis buffer (50 mM sodium phosphate pH 8, 125 mM NaCl and 1% Triton) containing a protease inhibitor cocktail (Roche, Rotkreuz, Switzerland). The cells were disrupted using several passages through a French press cell. The crude extract was clarified through centrifugation (17 000 rpm for 30 min at 4°C), and the supernatant was loaded onto two individual Strep-Tactin® Sepharose columns (1 ml of column volume; IBA GmbH, Göttingen, Germany). Each column was washed with 5 volumes of washing buffer (100 mM Tris–HCl, pH 8.0, 150 mM NaCl and 1 mM ethylenediaminetetraacetic acid) to remove the unbound proteins. The protein was eluted in six fractions (each at 0.5× column volume) of the same buffer containing 2.5 mM D-desthiobiotin. The respective fractions obtained from both columns were combined and loaded onto a 15% sodium dodecyl sulfate-polyacrylamide gel for electrophoretic separation. The proteins within the gel were stained with Coomassie blue (BIO-RAD). The fraction with the highest amount of protein was selected and concentrated using centrifugal filter columns (Roti®-Spin MINI-10, Carl Roth, Germany). The protein concentration was determined using the Pierce BCA Protein Assay kit (Thermo Scientific). The HapR-N-Strep protein was stored at −20°C in elution buffer containing 10% glycerol and 0.3 mM Dithiothreitol (DTT).

EMSA

The protocol for EMSA was established based on similar previously described protocols (12,13). Briefly, DNA fragments were obtained through the PCR amplification of gDNA from V. cholerae or plasmid DNA sequences using the primers listed in Supplementary Table S1. The amplicons were visualized on a 1.2% agarose gel. The DNA fragments were (gel-) purified using either GenElute^TM PCR Clean-Up or Gel Extraction kits (Sigma-Aldrich, Switzerland). The in vitro binding of HapR to the DNA probes was facilitated through incubation of the purified protein with the DNA fragments in reaction buffer (10 mM Tris–HCl pH 8.0, 2.5% glycerol, 0.5 mM ethylenediaminetetraacetic acid, 100 mM KCl, 1 mM MgCl₂ and 2 mM DTT) for 20 min at 30°C. After the incubation, bromophenol blue and 2.5% glycerol were added to the reactions. The samples were loaded onto a pre-run (in 0.5× TBE) 8% polyacrylamide gel. The electrophoretic separation occurred for 1 h 40 min at 100 V. The proteins were transferred from the gel to a polyvinylidene difluoride membrane for western blotting where indicated. All other gels were soaked for 30 min in a solution containing 0.2 μg/ml of ethidium bromide to stain DNA. The pictures were captured with an UV transilluminator (G:BOX, Syngene). The images were rotated, cropped and uniformly adjusted for brightness and contrast using Adobe Photoshop.

Quantitative reverse transcription PCR (qRT-PCR)

The V. cholerae strains were grown for several hours in LB medium, as previously described (8). Where indicated, 0.02% arabinose was added to the medium to artificially induce natural competence (as TfoX is driven by the P_BAD promoter in this system) (8). The cell harvest, RNA preparation, and reverse transcription followed by quantitative PCR (qRT-PCR) (LightCycler® Nano, Roche) were performed as described earlier (8), except the cDNA was prepared using the Transcriptor Universal cDNA Master mix (Roche, Rotkreuz, Switzerland). The expression values are given relative to the expression of the reference gene gyrA (8). The gene-specific primers used for qRT-PCR are indicated in reference (8) and in Supplementary Table S1.

The artificial expression of comEA increases natural transformation in a HapR-negative strain

Natural competence and transformation of V. cholerae occurs at a high cell density, as described earlier in the text. We previously showed that the expression of comEA is dependent on HapR, whereas many of the other competence genes, such as pilA, are expressed independently of this regulatory protein (8) (Figure 1). Based on this finding, we wanted to determine whether the transformability of a V. cholerae HapR mutant strain could be rescued solely through mimicking the HapR-mediated activation of comEA. Thus, strain ΔhapR-TntfoX carrying the inducible plasmid pBAD-comEA was subjected to a chitin-independent natural transformation assay, as previously described (8). As shown in Figure 2, the artificial expression of comEA partly rescued the transformation phenotype in the hapR minus strain (lane 4). This result is consistent with that of a previous study showing that the natural transformation in HapR mutant strains could be partially rescued through deletion of dns, the gene repressed by HapR (7). However, it is unknown whether HapR or intermediate regulators directly influence comEA and dns expression.

A putative HapR binding site upstream of comEA is not essential for natural transformation

The direct binding of HapR to the promoter regions of the virulence regulator gene aphA and the gene encoding the hemagglutinin protease (HA protease) hapA has been demonstrated in vitro (12,13) (Figure 1). Furthermore, Tsou et al. (13) identified two distinct HapR-binding motifs, motif 1 and motif 2, using bioinformatics tools and experimental validation. We wanted to determine whether a similar motif would also be present in the upstream region of comEA, indicating the direct regulation of comEA through HapR. Indeed, we identified a DNA region located between 122 and 103 bp upstream of the comEA start codon, which highly resembled motif 1 (13) (Supplementary Figure S1A). To characterize the significance of this motif, we generated the plasmid pBR-[P_WT]comEA, carrying comEA preceded by 200 bp of its indigenous upstream region. This plasmid was used as a template for the site-directed mutagenesis of the left, right or both segments of the putative HapR binding site. The resulting plasmids, pBR-[P_{mut L}]comEA, pBR-[P_{mut R}]comEA and pBR-[P_{mut L/R}]comEA, were assessed for their ability to complement V. cholerae strains lacking the comEA gene on the chromosome in a chitin-independent transformation assay. As shown in Supplementary Figure S1, the transformation frequencies of all strains carrying mutations in the putative HapR binding site (panel B, lanes 4–6) were comparable with the frequency of the strain complemented with comEA preceded by its indigenous upstream region (lane 3). Similar results were obtained when natural competence was induced on a chitin surface using a previously described method (37). Indeed, in this chitin-dependent transformation assay, the complementation worked even better (Supplementary Figure S1C). The most likely explanation for this difference in complementation efficiency is that the plasmids used in this study were derivatives of pBR322, which changes copy number under different growth conditions/growth rates (47). These results demonstrate that the putative HapR binding site is negligible for the regulation of comEA. However, it remains unknown whether HapR binds to a different sequence within the upstream region of comEA or whether HapR indirectly regulates this competence gene.

Direct and indirect regulation of competence genes by HapR

To determine whether HapR directly regulates certain genes that play a role in competence and transformation, we assessed the binding of HapR to the respective promoter regions of these genes in vitro. First, we purified the HapR protein using a tagged version of HapR through the cloning of plasmid pBAD-hapR-N-Strep (fusion between the sequence encoding for the Strep-tagII peptide and hapR). We then determined whether hapR-N-Strep could complement the transformation-negative phenotype of a V. cholerae hapR minus strain. As depicted in Supplementary Figure S2, the transformation frequencies of the strain ΔhapR-TntfoX carrying hapR (lane 3) or hapR-N-Strep (lane 4) were almost identical, highlighting the in vivo functionality of this translational fusion.

Assured that HapR-N-Strep was fully functional in vivo, we heterologously expressed this gene in E. coli strain Origami 2^TM (DE3) pLysS. The successful but low-level expression of HapR was verified through PAGE and western blot analyses (Supplementary Figure S2B). The N-terminally tagged protein was purified using a Strep-Tactin® Sepharose column (see ‘Material and Methods’ section for details; Supplementary Figure S2C) and stored at −20°C.

Next, we characterized the in vitro binding of the purified HapR protein to specific DNA fragments using an EMSA. We first tested the in vitro binding ability of HapR to the comEA promoter region (the features of this and other tested DNA probes are indicated in Supplementary Tables S2–S4). As indicated in Figure 3A (lanes 5–8), HapR did not shift the comEA promoter (P_comEA)-specific DNA fragment, even at the highest tested protein concentration (lane 8). The same result was obtained for another V. cholerae competence gene, comEC (21), whose expression is also influenced through HapR in vivo (8) (Figure 3A, lanes 9–12). In contrast, the promoter region of aphA, to which HapR binds in vitro (12), was successfully used as positive control in this assay (Figure 3A, lanes 1–4). We also examined the promoter of the competence gene pilA, which is not a component of the QS regulon in V. cholerae (8) (Figure 1). In this experiment, we used a longer DNA fragment of ∼600 bp, as this fragment was previously used to report pilA transcription using transcriptional reporter fusions (8). Consistent with the HapR-independent expression in vivo, HapR did not bind the pilA promoter in vitro (Figure 3B, lanes 5–8). A similar-sized fragment spanning the aphA promoter region (aphA*) served as a positive control (lanes 1–4) to facilitate the proper comparison with the P_pilA probe. From these data, we concluded that HapR neither binds to the comEA and comEC promoter regions nor to the pilA promoter region.

The other QS-regulated gene that plays a major role in natural transformation is dns (7) (Figure 1). To determine whether HapR directly or indirectly mediates the repression of dns, we repeated the mobility shift assays using the dns promoter region as a DNA template (Figure 4). An ∼250 bp fragment covering the region −203 bp to +48 bp from the start codon of dns was first tested (Figure 4A). Interestingly, HapR was able to shift the DNA fragment, although only at higher HapR protein concentrations than those sufficient to shift the positive control fragment (Figure 3A). We also included a non-related negative control in this assay [the recA promoter region; (8)], which did not show any mobility shift (data not shown). A western blot analysis of the dns-specific EMSA gel followed by the detection of HapR with α-HapR-specific antibodies confirmed that HapR protein co-localizes with the shifted DNA fragment (data not shown).

To identify the putative HapR binding site within the dns promoter region, we PCR amplified several fragments around the dns start codon as indicated in Figure 4B (fragments A–H; details about the fragments can be found in Supplementary Table S3). All fragments were tested for HapR-dependent mobility shifts, and the results are indicated in the table on the left of Figure 4B. Apart from the outermost fragments (fragments A, G and H), all other probes remained bound to the HapR protein (Figure 4B). These fragments contained either the DNA sequence from −100 to −50 bp and/or the region from −50 to −1 bp, with respect to the start codon. Thus, we cloned these two sub-regions into vector pBR-Tet_MCSI (Table 1). The resulting plasmids were then used as PCR templates to obtain DNA probes of ∼200 bp in length (Figure 4B, fragments I and J). To ensure that HapR did not bind to the vector sequence, we also used a DNA fragment solely derived from vector pBR-Tet_MCSI as a negative control (Figure 4B, fragment K). The resulting EMSA patterns, after in vitro incubation with HapR, are depicted in Figure 4C. These data indicated that HapR binds to at least two separate HapR binding sites within the dns promoter in vitro. Although we did not find any obvious HapR binding site for the dns sub-region of DNA fragment J, we identified a putative HapR binding site resembling motif 2 (13) within DNA fragment I (−71 bp to −56 bp from the dns start codon but on the complementary strand; Supplementary Figure S3).

Despite both comEA and dns being HapR-dependent in vivo (8), the in vitro binding of HapR to the respective promoter region could only be demonstrated for dns. Thus, we concluded that HapR regulates comEA (and most likely also comEC) in an indirect manner.

A QS- and tfoX-dependent regulatory protein plays a major role in the natural competence and transformation of V. cholerae

Based on the results described earlier in the text, we hypothesized that an intermediary regulatory protein might exist between HapR and the QS-dependent expression of comEA. Based on previous microarray expression-profiling data (2), we identified VC0396, which potentially encodes an intermediate protein in the competence regulatory network. VC0396 was initially annotated as a ‘transcriptional regulator of the LuxR family’ (42). Hereafter, we refer to this gene as QS and TfoX-dependent regulator (qstR). We validated the expression of qstR in our chitin-independent experimental model using qRT-PCR. Comparable with other competence genes, qstR was induced upon artificial low-level tfoX expression (Figure 5, highlighted results). This TfoX-dependent induction was not observed in a hapR-deficient background strain (Figure 5), which is consistent with the HapR-mediated regulation of qstR. Furthermore, a comparison of the expression pattern of qstR revealed high similarities with the expression pattern observed for comEA (Figure 5). Thus, these results confirmed our hypothesis that QstR could be an intermediary regulatory protein that signals between HapR and comEA.

The qRT-PCR data suggested the involvement of QstR in natural competence and transformation; therefore, we deleted the qstR gene in the V. cholerae wild-type strain A1552 and in the strain A1552-TntfoX. The deletion of qstR did not result in any obvious competence-independent phenotype [e.g. growth in rich medium, colony morphology, chitin colonization (9), etc.]. We therefore examined whether the lack of qstR influences chitin-induced or chitin-independent natural transformation (Table 2). Indeed, for the qstR minus strain, transformation was below (chitin-dependent) or at the limit of detection (chitin-independent assay) and therefore reduced by at least four orders of magnitude compared with the respective parental strain (Table 2), highlighting the importance of QstR for natural competence and transformation. The extremely rare transformants observed under chitin-independent competence-inducing conditions (Table 2) were most likely a reflection of low levels of comEA transcription even in the absence of QstR, whereas a comEA knockout strain was never transformable in our assay.

Next, we complemented the deletion strain by providing the qstR gene in trans, which restored natural transformation (Table 2). Consistent with our hypothesis that QstR acts as an upstream regulator of comEA (Figure 1), the exogenous expression of comEA also significantly increased natural transformability in the ΔqstR genetic background (Table 2). The transformation frequency in this setting (e.g. ΔqstR/pBAD-comEA) was ∼10-fold higher than in strain ΔhapR/pBAD-comEA (Figure 2), which is consistent with HapR-mediated repression of dns in the qstR mutant. To demonstrate that QstR acts downstream of HapR, we also artificially expressed qstR from a plasmid in the hapR minus strain. Indeed, the exogenous expression of qstR in trans in the ΔhapR-TntfoX strain rescued natural transformation to the same extent as the hapR gene itself (Table 2). We therefore propose that HapR directly or indirectly influences qstR expression and that QstR regulates QS-dependent competence and transformation genes (Figure 1). Furthermore, as the overexpression of qstR in a hapR background apparently also alleviated the transformation barrier exerted by the Dns nuclease, we suggest that QstR might also be involved in the repression of dns (Figure 1).

HapR binds to the qstR promoter region

Next, we examined whether HapR regulates qstR in a direct or indirect manner. We repeated the in vitro HapR binding assay as described earlier in the text, using the upstream region of qstR as a probe. As indicated in Figure 6, HapR bound to the DNA fragment causing a shift in mobility.

To identify the location of the putative HapR binding site(s), we followed the strategy described for Figure 4, examining a plethora of DNA fragments derived from −359 bp upstream to +105 bp downstream of the start codon of qstR (Figure 6B). The probes (fragments A–G; details in Supplementary Table S4) were tested for in vitro binding to HapR (EMSA), as indicated in the figure. All shifted fragments shared the sequence spanning the region from −150 to −102 bp upstream qstR (Figure 6B and Supplementary Table S4). Thus, we subcloned two fragments, i.e. the regions spanning either −150 to −120 bp or −150 to −102 bp upstream of the ATG start codon into vector pBR-Tet_MCSI (Table 1). The resulting plasmids were used as PCR templates to obtain linear DNA fragments of ∼200 bp in length (fragments H and I in Figure 6B). Fragment H, containing the shorter stretch of the P_qstR region, did not exhibit in vitro binding to HapR (Figure 6C). In contrast to fragment H, the ∼50 bp of the qstR upstream region within fragment I, were sufficient to facilitate HapR binding. On further examination of this particular DNA region, we identified a sequence resembling in silico predicted motif 2 (13) [Figure 6D; note that the simplified consensus was slightly modified from (13) to facilitate alignment with the putative motif upstream of qstR]. Indeed, after site-directed mutagenesis of two conserved bases within this motif (Figure 6D), the capability of HapR to shift the DNA fragment in the assay was lost (Figure 6C). Taken together, these results demonstrate that HapR directly binds to the qstR upstream region, and that this binding occurs in a sequence-specific manner, despite the fact that the motif does not perfectly match the in silico predicted consensus (13).

QstR is essential for the TfoX-dependent induction of QS-dependent competence genes

The data provided earlier in the text indicated that HapR binds to the qstR promoter region. However, which competence genes does QstR subsequently regulate? To answer this question, we compared the expression levels of a plethora of competence and chitin metabolism genes, both under competence non-inducing and competence-inducing conditions in three different strains. Apart from the tfoX-carrying transposon TntfoX (8) the genetic backgrounds of these strains were either wild-type (A1552-TntfoX), hapR-negative (ΔhapR-TntfoX) or qstR-negative (ΔqstR-TntfoX) (Figure 5). The results showed that the expression of comEA and comEC is significantly lower under competence-inducing conditions in the two knockout strains compared with that observed in the corresponding wild-type strain, although the expression levels were comparable between the two strains. In contrast, the competence-independent background expression of comEA and comEC did not differ among any of the three strains tested (Figure 5). The results were completely different for dns. The expression of dns was significantly reduced in the wild-type background strain upon tfoX induction, although the hapR transcript levels did not change (Figure 5). Furthermore, the expression level of hapA, as a competence-unrelated but HapR-dependent gene and therefore a direct reflection of HapR protein activity (Figure 1), did not change on competence induction. Consistent with HapR acting as repressor of dns, the dns expression levels were significantly increased in the hapR minus background strain, independently of the status of tfoX (induced or not) (Figure 5). For the newly identified regulator QstR, we observed a phenotype with respect to dns expression, which neither reflected the observations of the wild-type strain (e.g. TfoX-dependent repression of dns) nor those of the hapR minus strain (constitutively high expression of dns). In contrast, under competence-non-inducing conditions, dns was repressed through HapR in the qstR negative strain, but further repression on TfoX induction did not occur (Figure 5). We therefore speculate that QstR might exert additional repression on dns (as depicted with a dashed repression arrow in Figure 1). In contrast to the expression signatures of the QS-dependent genes comEA, comEC and dns (2,7,8), none of the other tested genes involved in competence [pilA, pilM, VC0047, dprA; (2,21)] or in the metabolism of chitin in V. cholerae [chiA-1, chiA-2, VCA0700; (9,14)] showed any significant difference between the three tested strains under competence-inducing conditions (Supplementary Figure S4).