ABSTRACT
The Sin quorum sensing (QS) system of S. meliloti activates exopolysaccharide and represses flagellum production. The system consists of an N-acyl-homoserine lactone (AHL) synthase, SinI, and at least two LuxR-type regulators, SinR and ExpR. SinR appears to be independent of AHLs for its control of sinI expression, while ExpR is almost completely dependent upon AHLs. In this study, we confirmed 7 previously detected ExpR-DNA binding sites and used the consensus sequence to identify another 26 sites, some of which regulate genes previously not known to be members of the ExpR/AHL regulon. The activities of promoters dependent upon ExpR/AHL were titrated against AHL levels, with varied outcomes in AHL sensitivity. The data suggest a type of temporal expression program whereby the activity of each promoter is subject to a specific range of AHL concentrations. For example, genes responsible for exopolysaccharide production are activated at lower concentrations of AHLs than those required for the repression of genes controlling flagellum production. Several features of ExpR-regulated promoters appear to determine their response to AHLs. The location of the ExpR-binding site with respect to the relevant transcription start within each promoter region determines whether ExpR/AHL activates or represses promoter activity. Furthermore, the strength of the response is dependent upon the concentration of AHLs. We propose that this differential sensitivity to AHLs provides a bacterial colony with a transcription control program that is dynamic and precise.
INTRODUCTION
The ability of bacteria to perceive population density has become known in the world of microbiology as quorum sensing (QS). This widespread bacterial mechanism facilitates the recognition of population density and an appropriate response (1). The best-studied QS signaling systems are based upon the employment of N-acyl-homoserine lactones (AHLs) as the signaling molecule. As might be expected, the genetic determinants of AHL production and perception are frequently integrated in complex regulatory networks and affect numerous aspects of the bacterial lifestyle (2). Typically, quorum-sensing systems operate as transcription networks regulated by a LuxR-type transcription regulator and its cognate AHL (reviewed in reference 3). While many studies on quorum sensing have focused on extending the known regulon (4–13), few have considered the temporary aspects of transcription regulation (12). Interestingly, studies on regulatory networks are revealing the dynamic behavior and precision timing of transcription control (14, 15). This raises the question as to whether quorum sensing-regulatory controlled networks exhibit similar characteristics in response to accumulating AHL levels.
In a previous study on a model bacterium for legume-rhizobium symbiosis, Sinorhizobium meliloti, we found a subset of genes whose expression was dependent not only on the presence of AHLs but also on the level of AHLs supplied to the growth culture of a strain incapable of producing AHLs (16). These genes, sinI, sinR, and expR, are all essential for quorum sensing regulation (see Fig. 1A for the regulatory scheme). sinI encodes an AHL synthase which catalyzes the synthesis of several long-chain AHLs, including oxo-C14-HL, oxo-C16:1-HL, and C16:1-HL (17–19). Upstream of the sinI gene and separated from it by 156 nucleotides is sinR, which encodes a transcription regulator controlling the activity of the promoter of sinI (16–18, 20). The promoter of sinR responds to environmental cues, such as nutrient limitation, by increasing the transcription of sinR (16). SinR protein is necessary for activation of the promoter of sinI, most likely through a SinR binding site immediately preceding the −35 position (21), and an increase in the production of SinR results in an increase in sinI expression (16).
The Sin/ExpR quorum-sensing system regulates its own genes (sinR, sinI, and expR) and a multitude of other genes. (A) Regulatory diagram (based upon reference 16) showing the transcriptional control of sinR and expR by ExpR and AHLs and of sinI transcription by SinR and ExpR and AHLs. ExpR and AHLs also control the expression of genes important for motility, exopolysaccharide production, and many other processes that are less well known. Arrows indicate activation; flat-ended lines indicate repression. OM, outer membrane; IM, inner membrane. (B) A total of 570 genes have been identified as being regulated by ExpR and AHLs based on all three global approaches. The Venn diagram shows the overlap in genes determined to be regulated by both ExpR and AHLs in various studies (references 11 to 13, as indicated in parentheses). Bold type indicates the number of genes in each study identified as regulated. Ratios in parentheses are the number of genes immediately preceded by an ExpR site/number of genes either immediately preceded by an ExpR site or located within an operon-like structure downstream of an ExpR binding site. The numbers in the overlap are the numbers of genes identified in more than one study. Genes in the overlaps are listed; the underlined genes are situated immediately downstream of an ExpR binding site identified either in a previous study or in this study.
The third gene of the Sin system, expR, is disrupted by an insertion element in the S. meliloti strains Rm1021 and Rm2011 (22), which are most intensively studied. A restoration of the gene confers QS capacity upon the bacterium and a strong increase in production of the symbiotically important exopolysaccharides galactoglucan and succinoglycan (11, 13, 20, 22–24). The DNA-binding activity of ExpR depends upon the presence of AHLs (21), and the same is true for its regulatory activity (16). The ExpR-AHL complex regulates a number of promoters throughout the genome. Exactly how many is unknown, but binding has been demonstrated for the promoters of genes controlling galactoglucan production (wgeA and wgaA), genes related to succinoglycan production (exoI and exsH), genes controlling flagellum production (visNR), and the Sin system genes sinR and sinI (16, 20, 25).
From the outset of this study, we suspected that the ExpR-AHL complex binds many more promoters in addition to the seven listed above. This is because evidence from both mRNA (12, 13) and protein (11) accumulation analyses suggests that the Sin/ExpR system regulates a multitude of genes (Fig. 1B). However, for the majority of these genes, the mechanisms underlying the regulation are unknown.
The purposes of this study were to extend the known Sin/ExpR regulon and to understand its mechanisms of regulation. Over 100 promoter regions were tested for binding of ExpR, and we analyzed the regions that were able to bind to ExpR for ExpR/AHL dependent expression activity. We show that the expression of each gene in the ExpR regulon is controlled by a specific AHL concentration range, resulting in a differential expression program which is sensitive to increasing levels of AHLs. Based on this information, we speculate on general regulatory mechanisms involved in the Sin/ExpR QS process and how these might be related to survival strategies.
MATERIALS AND METHODS
Bacterial strains and growth conditions.Strains used in this work are shown in Table S1 in the supplemental material. Unless otherwise specified, S. meliloti strains were cultivated at 30°C on solid or liquid TY medium (26) or a modified MOPS (morpholinepropanesulfonic acid)-buffered minimal medium containing 48 mM MOPS (adjusted to pH 7.2 with KOH), 55 mM mannitol, 21 mM sodium glutamate, 1 mM MgSO4, 250 mM CaCl2, 37 mM FeCl3, 48 mM H3BO3, 10 mM MnSO4, 1.0 mM ZnSO4, 0.6 mM NaMoO4, 0.3 mM CoCl2, 4.1 mM biotin and 0.1 mM K2HPO4 (27). E. coli strains were cultivated at 37°C in Luria-Bertani (LB) medium (28). When required, antibiotics were added at the following concentrations: 10 μg ml−1 nalidixic acid, 10 μg ml−1 tetracycline, and 200 μg ml−1 kanamycin for S. meliloti and 100 μg ml−1 ampicillin, 10 μg ml−1 tetracycline, and 50 μg ml−1 kanamycin for E. coli.
Plasmid construction.The construction of plasmids pLK64, pLK65, and pLK66, in which the promoters regions of sinI, sinR, and expR plus the translation start were fused to the gene encoding enhanced green fluorescent protein (egfp) using pPHU231, was described previously (16, 20). In similar manner, the other plasmids in the pLK series were generated using the primers listed in Table S2 in the supplemental material. Typically, the promoter regions included in the plasmid constructs contained the first 300 bp upstream of the target gene, the ATG of the target gene and ensuing ≤15 bp, and these regions were fused to the ATG of egfp.
Binding site cloning and DNA labeling.The promoter regions used in the electrophoretic mobility shift assays were prepared via PCR in two steps. The DNA fragments were first amplified from genomic DNA using a promoter-specific forward primer which contains a linker, GTGAGCGGATAACAATTTCACACAGGA, together with a promoter-specific reverse primer. (These primers were also used for the construction of pLK plasmids.) The resulting PCR product was then used as a template for a second PCR using the Cy3-labeled primer GTGAGCGGATAACAATTTCACACAGGA (pUC18-unifwd-Cy3) together with an unlabeled promoter-specific reverse primer.
PCR-based mutation of the ExpR binding site upstream of expR.For the replacement of specific nucleotides within the ExpR binding site upstream of expR, internal complementary primers were designed based around the region of interest. In the first round of PCRs, each of the complementary primers was included with the matching peripheral primer carrying a specific DNA restriction site for cloning. In the second round of PCRs, both PCR products from the primary PCRs were combined and used as a template with the peripheral primers. The resulting product depended upon the annealing of the PCR products from the primary PCR round and resulted in a modified promoter region upstream of expR. The wggR promoter region was modified in a similar manner.
Expression and purification of His6-ExpR.The expression and purification of recombinant His6-ExpR were performed as described previously (21) with modifications. LB (250 ml) was inoculated with an overnight culture (1:100 dilution) and grown at 37°C until an optical density at 600 nm (OD600) of 0.6 was reached. The Escherichia coli M15 culture was then induced with 0.5 mM IPTG (isopropyl-β-d-thiogalactopyranoside) and grown at 21°C overnight. The cell pellet was resuspended in 5 ml of lysis buffer containing 50 mM MOPS, 0.5 M NaCl, and 20 mM imidazole (pH 7.5). Cell breakage was performed by passing the cell suspension three times through a French pressure cell at 1,000 lb/in2. Cell debris was removed by centrifugation at 10,000 × g for 10 min at 4°C. The supernatant was filtered with a 0.5-μm filter and then loaded onto a 1-ml nickel-nitrilotriacetic acid (Ni-NTA) affinity column. The column was washed with lysis buffer to remove nonspecifically bound proteins. Elution of the protein His6-ExpR was achieved with an imidazole gradient (0.02 to 1.0 M). Fractions (0.5 ml) were collected and analyzed with sodium dodecyl sulfate-polyacrylamide gel electrophoresis, and protein concentration was measured using a NanoDrop1000 (PEQLAB). Purified protein was mixed with glycerol (1:1, vol/vol) and stored at −20°C for up to 2 weeks.
Electrophoretic mobility shift assay (EMSA).The EMSA protocol was as described previously (16, 21) with slight modifications. The Cy3-labeled DNA fragments were mixed with purified His6-ExpR in a reaction buffer containing approximately 2.5 A260 units ml−1 of sonicated salmon sperm DNA (GE Healthcare) and 1.0 mg ml−1 of bovine serum albumin (Sigma) in a final volume of 10 μl of DNA binding buffer (10 mM Tris-HCl, pH 8.5, and 50 mM KCl). In the 10-μl reaction mixture, the Cy3-labeled DNA was included at 0.05 pmol (1 ng μl−1); His6-ExpR was included at 85 pmol (0.25 μg μl−1); AHL (C16:1-HL) was included at 100 pmol (10 μM). The reaction mixture was incubated at room temperature for 15 min. Loading buffer (2.5 μl; 20% TAE buffer, 80% glycerol) was added, and the reaction mixtures were loaded onto a 2% agarose gel. Following electrophoresis at 5 V cm−1 and room temperature for 1.5 h, gel images were scanned using a Typhoon 8600 variable-mode imager (Amersham Bioscience).
EGFP fluorescence assay.The EGFP fluorescence assay was modified from a previous study (16). S. meliloti strains were grown in MOPS minimal medium containing 0.1 mM phosphate at 30°C. Starter cultures were grown for 20 h in MOPS minimal medium and then diluted to an OD600 of 0.002 in fresh MOPS minimal medium. For measurement of promoter activity, bacteria were grown in 0.1-ml volumes (4 biological replicates) in a 96-well microtiter plate (Greiner) with shaking (800 rpm) at 30°C. Measurements were made at 8 h, 16 h, 24 h, and 40 h. Background fluorescence (≈600 to 700 fluorescence units/OD unit) was determined from strains carrying the pLK vector with a promoterless egfp. This background has not been removed from any of the promoter activity profiles (Fig. 5A). For all fluorescence assays, optical density (OD600) and fluorescence (excitation, 485 nm; emission, 538 nm) were measured using the Tecan Infinite M200 reader (Tecan Trading AG, Switzerland). For measurement of promoter sensitivity to C16:1-HL, the sinI mutant strain carrying one of the pLK series vectors was grown in MOPS minimal medium supplemented with various concentrations of C16:1-HL (Cayman Chemical Company). Expression of expR is ≈1.5- to 2-fold higher in the wild-type strain than in the expR and sinI mutant strains (16). In order to compensate for this lower level of expR expression in the sinI mutant, a low level of ectopically expressed expR from the vector pBSexpR (16) was induced via the addition of 0.1 mM IPTG. We have tested the level of IPTG needed to induce ectopic expression of expR from pBSexpR (in which expR expression is controlled by IPTG-inducible lacp). This estimation was based upon the concentration of IPTG which restored sinIp activity in a sinI mutant strain supplemented with 1 μM AHL to those observed in the wild type (see Fig. S2 in the supplemental material). We selected this concentration of C16:1-HL because AHL concentrations in cultures of the wild-type strain were previously estimated at ≈1.3 μM (16). C16:1-HL and IPTG were added at the point of culture inoculation. Measurements were made at 24 h and 40 h.
Determination of expR transcription start site.Rapid amplification of cDNA 5′ ends (5′-RACE) PCR experiments were performed using tobacco acid pyrophosphatase (Epicentre Biotech) treatment of purified mRNA followed by ligation to the 3′end of the RNA primer GUAUGCGCGAAUUCCUGUAGAACGAACACUAGAAGAAA using T4-RNA ligase (Fermentas). After reverse transcription, the forward primer GCGCGAATTCCTGTAGAACG (based on the RNA primer above) and the expR specific reverse primer GTCCGGCCAGAAGAAGTCTC were used to amplify cDNA fragments in a PCR. These fragments were then cloned into the TOPO-TA vector (Invitrogen) and sequenced.
RESULTS
SinR activation of sinIp is independent of AHLs.LuxR-type proteins, such as SinR and ExpR, typically contain a C-terminal domain which binds to DNA and affects transcription activity, plus an N-terminal domain which binds to a ligand (3). Previous studies have shown that the loss of sinR results in a loss of detectable promoter activity of sinI, both in the presence and absence of expR or AHLs. Thus, SinR is necessary for a basal activity of sinIp, and this activity is modulated in the presence of both ExpR and AHLs (16, 20, 22, 24). Furthermore, the expR mutant produces significant levels of AHLs (17–20). However, the question of whether SinR requires AHLs for its activation has not been rigorously addressed. Since SinR is encoded immediately upstream of sinI, it has been typically assumed to be dependent upon AHLs, but this has not been demonstrated. The results in this study show that in an expR mutant, sinIp remained active in the absence of sinI (Fig. 2). Thus, neither expR nor sinI is necessary for the basal activity of sinIp. Furthermore, addition of AHLs to the double mutant (expR sinI) did not alter sinIp activity (Fig. 2). This indicates that while ExpR is dependent upon AHLs for its activation of the sinIp, SinR-dependent activation is unaffected by the presence of AHLs. Also, ExpR/AHLs cannot activate sinIp in a sinR mutant, consistent with previous observations (16, 20), even with ectopic expression of expR from pBSexpR (data not shown). However, ectopic expression of sinR strongly activates sinIp activity, even in the absence of AHLs or ExpR (16). Therefore, SinR is necessary for the AHL-independent basal activity of sinIp, and ExpR enhances this activity in an AHL-dependent manner (see Fig. 1A for the regulatory scheme). Since ExpR, but not SinR, is dependent upon AHLs for its regulatory activity, we focused upon the regulatory targets of ExpR.
SinR activates promoter activity of sinI independently of AHLs. Strains carrying the vector pLK64, in which the promoter of sinI is fused to egfp, were measured after 24 h growth in phosphate-limiting MOPS minimal medium. The promoter activity of sinI is dependent upon the presence of SinR and ExpR/AHLs but in the absence of ExpR is unaffected by the presence of SinI or AHLs. The measurements were made at least three times. Error bars show errors from 4 biological replicates.
ExpR binds to multiple binding sites.ExpR binds to a DNA sequence of imperfect or degenerate dyad symmetry, as noted previously (16). From this alignment of seven previously detected ExpR DNA-binding regions delimited to 20 nucleotides (nt), we deduced a consensus sequence which was then used with PatScan (29) at http://iant.toulouse.inra.fr/bacteria/annotation/cgi/rhime.cgi to identify additional sites in the genome of the S. meliloti strain 1021. A list of hits was compiled using the following criteria: a hit must consist of 18 to 20 nucleotides with some similarity to the ExpR consensus sequence (CCCANNATTNTATTGGGG) and be located within a promoter region (≈250 bp upstream of the translation start) of a gene. Particular attention was paid to genes that were previously identified as being regulated by ExpR/AHLs, by either protein or mRNA accumulation (11–13). To test for ExpR-DNA binding, the DNA surrounding the hit was included in a gel shift binding assay together with purified His6-ExpR and AHLs (Fig. 3). Table 1 lists the binding site sequences and the genes located downstream of each ExpR binding site.
Electrophoretic mobility shift assays (EMSA) using purified His6-ExpR and Cy3-labeled DNA fragments derived from the promoter regions, except for SMb20391 (asterisk), which contains an ExpR site inside the coding region. For each set of three lanes, the first lane is the negative control, which includes only the Cy3-labeled DNA from the promoter region. The second contains both the Cy3-DNA and His6-ExpR. The third contains Cy3-DNA, His6-ExpR, and AHLs (10 μM, C16:1-HL). (A) EMSAs of promoter regions known to bind to ExpR from previous studies. (B) Promoter regions identified as binding to ExpR in this study. (C) Promoter regions that do not bind to ExpR, included here as negative controls. (D) Promoter regions with modifications depicted in panels E (expR upstream region) and F (wggR upstream region). The gel shift assays were carried out at least three times.
ExpR binding sites and their relevant surrounding features
To increase the confidence in the correct identification of each ExpR binding site, we sequentially reduced the size of the original 300-bp fragment from both ends to either include or exclude the hit and tested each derivative fragment in a gel shift assay (data not shown). In this way, the relevance of the hit for the binding between ExpR and the 300-bp fragment was established. Using this method, 26 novel binding sites were confirmed in this study. We also tested some 93 additional promoter regions in gel shift assays, selected on the basis of containing some similarity to the ExpR binding site consensus or having promoter activity which was dependent upon ExpR and AHLs. However, these did not bind to His6-ExpR under our conditions. As negative controls, the promoters of glgP, mucR, and pstS, which do not contain any similarity to the ExpR binding site in their promoter regions, were included. Their lack of binding to His6-ExpR also indicates the specificity of the ExpR-DNA interaction (Fig. 3C).
Previous attempts to identify an ExpR binding site in the promoter region of expR were unsuccessful, despite a weak but clear positive regulatory effect of ExpR and AHLs on this promoter (16). However, several factors prompted us to reexamine this. First, we identified a single transcription start for expR at 88 bp upstream of the ATG (see Fig. S1 in the supplemental material). Assuming a classical promoter structure, this places the −35 region immediately downstream of a DNA sequence that weakly resembles the ExpR binding site consensus. Second, improvements in the protein-purifying procedure provided a more active fraction of His6-ExpR. Thus, when the improved purified His6-ExpR was included with the 300-bp fragment upstream of the expR gene in an EMSA, a clear shift was observed (Fig. 3B). When a single nucleotide was exchanged (T→C) within the binding site sequence to improve resemblance to the ExpR binding site consensus, a stronger binding to ExpR was not apparent (Fig. 3D). However, when another nucleotide within this sequence was modified (C→T) to decrease similarity to the ExpR binding site consensus, the result was an almost complete lack of binding to ExpR (Fig. 3D). Together with corresponding changes in expR promoter activity associated with the nucleotide exchanges (see below), these experiments increase the confidence in the location and function of this ExpR binding site.
In each gel shift assay, the same amount of His6-ExpR and target DNA was used. This allows a comparison of relative binding strengths encoded by the DNA fragments, which is apparent in Fig. 3. Addition of 10 μM C16:1-HL to the assay (which represents a saturation concentration in our assay) did increase shift strength in some cases, particularly for the weaker binding sites. Additions of >10 μM C16:1 did not increase shift strength (data not shown). However, the majority of the promoter regions bound to His6-ExpR even in the absence of AHLs. This is perhaps best explained by the amount of His6-ExpR included in the assay, which was set at a concentration that was optimal for the observance of the weakest shifts but was in excess for the stronger binding sites. Generally, a closer resemblance to the ExpR binding consensus (Table 1) correlates positively with a stronger shift. For example, binding sites located upstream of sinI, exoI, SMb21681, SMc01711, SMc02378, SMc02726, SMc03149, SMc03150, SMc04232, and phrR are among those exhibiting the stronger shifts and are therefore better indicators of the binding consensus. In contrast, binding sites upstream of cspA3, nesR, SMb21135, SMc01524, SMc04246, and SMc04258 are among the weakest. For the SMc04258 and nesR promoters, a weak shift is observable upon addition of His6-ExpR and AHLs (Fig. 3), although we could not detect any expression activity from these promoters. Conversely, such a weak shift was also observed with the promoter region of SMc01524 (Fig. 3), and yet this appears to be adequate for a ≥2-fold reduction in promoter activity (see Fig. S2 in the supplemental material). This was not the case with the control promoters (mucR, pstS, and glgP) and those of wggR and SMb20911, where addition of His6-ExpR and AHLs clearly did not induce a shift (Fig. 3). Typically, sites with 3 or 4 Cs on the gene-distal side and 3 or 4 Gs on the gene-proximal side separated by 10 to 12 nucleotides rich in A and T are better suited for binding (Table 1). The presence of Gs or Cs in the A/T-rich regions appears to weaken the shift, e.g., SMb21543 and SMb21135. Similarly, the presence of As or Ts in the G or C regions may weaken the shift, e.g., wgeA and wgaA. A new 19-nucleotide consensus sequence was derived from all 33 binding sites tested in this study. It can be represented as CCCCAAAAATTTTTTGGGG (Table 1). This is comparable to a recent study on the Pseudomonas aeruginosa LuxR-type regulator, LasR (5), where the consensus sequence is based upon binding sites in 14 promoters.
Promoter-egfp reporter assays reveal effects of ExpR and AHLs on promoter activity.ExpR in the presence of AHLs regulates the promoter activity of sinI, sinR, expR, wgaA, wgeA, wggR (12, 16, 20, 31), exoI, exsH, exoH (23), and visN (12, 25), as determined by real-time PCR and promoter-reporter fusions. Good congruence between promoter activities reported here (Fig. 4 and 5; also, see Fig. S2 in the supplemental material) and those in other studies (12, 13, 16, 20) provides for a high confidence in these results, despite variation in the use of strains and culture conditions. Three strains, SM2B3001 (wild type, with a functional expR), SM2B4001 (sinI) and Rm2011 (expR), were used to establish the role of ExpR and AHLs in the regulation of promoter activity. All promoter regions which bound to His6-ExpR in the gel shift assay were fused to egfp in the plasmid pPHU231. As controls, promoter regions which do not bind to His6-ExpR were also fused to egfp. Two examples of these are those of glgP (glycogen production/breakdown) and pstS (phosphate transport during phosphate limiting growth), selected as examples of active promoters that are not affected by QS.
Selected examples of promoters with various dependence upon the presence of ExpR and AHLs. Promoter regions were fused to egfp in plasmid pPHU231 and grown in three S. meliloti strains: SM2B3001 (wild type), SM2B4001 (sinI), and Rm2011 (expR). Fluorescence units per unit of optical density (F/OD; y axis) were measured for each promoter at the indicated time points (hours; x axis). Category 1, ≥2-fold-upregulated promoters in the presence of ExpR and AHLs which bind to His6-ExpR; category 2, ≥2-fold-downregulated promoters in the presence of ExpR and AHLs which bind to His6-ExpR; category 3, promoter region (nolR) which binds to His6-ExpR but is not regulated by ExpR and AHLs; control, promoter region (pstS) which neither binds to His6-ExpR and is not regulated by ExpR and AHLs. The measurements were made at least three times. Error bars show errors derived from 4 biological replicates.
Response of expRp to ExpR and AHLs. The expR promoter region was fused to egfp in plasmid pPHU231. Growth conditions were comparable to those used in the experiment whose results are shown in Fig. 4. The gray line indicates background fluorescence (see Materials and Methods). The effects of the CCC and TTT alterations (see Fig. 3E for details) are shown (A); also, the effects of various concentrations of supplemented AHL are shown (B). The measurements were made at least three times. Error bars show errors derived from 4 biological replicates.
The expR promoter (Fig. 5) responds to ExpR and AHLs, as was previously reported (16). Nucleotide replacements in the binding sequence correlated well with changes in promoter activity. The effect of the T→C change was not obvious in a gel shift assay (Fig. 3B and D), but it did result in a significant promoter activity increase in response to ExpR and AHLs (Fig. 5). Furthermore, the C→T change almost completely removed not only the binding to ExpR (Fig. 3D) but also the activating effect from ExpR and AHLs (Fig. 5A).
Other promoter regions containing ExpR binding sites showed a variety of responses to the presence of ExpR and AHLs (see Fig. S2 in the supplemental material). These fall into several categories. In the first are those promoters which are upregulated in the presence of ExpR and AHLs ≥2-fold. In this category are the promoters of sinI, wgeA, wgaA, exoI, and exsH, to which we add those of expR, exoH, SMb21543, SMc04237, SMc03150, and SMa2111. For these promoters, maximal activation under such growth conditions requires the presence of both ExpR and AHLs (Fig. 4 and 5; also, see Fig. S2). For example, the exoH promoter responds to a combination of ExpR and AHLs by a 3- to 4-fold increase in activity, but not if either ExpR or AHL is lacking. In the case of SMc03150, promoter activity (see Fig. S2 in the supplemental material) was observable only when a ribosome binding site from another gene (sinI) was inserted between the promoter of SMc03150 and egfp. This may be because our methods of detection are not sensitive enough, or because the native ribosome binding site and translation start were misidentified.
In the second category are those promoters where ExpR and AHLs downregulate activity ≥2-fold. In this category are the promoters of sinR and visNR plus the promoters identified in this study which include those of SMc01110 (phrR), SMc01524, SMc02378, SMc03864, and SMc04059 (see Fig. S2 in the supplemental material). As was the case of the ExpR/AHL activated promoters, repression of these promoters requires the presence of both ExpR and AHLs.
In the third category are promoters which contain a binding site but whose activity is affected by ExpR and AHLs, either negatively or positively, <2-fold. In this category are the promoters of SMc05009 (nolR), SMb21135, SMc04246, SMc01585 (cspA3), and SMb21245 (exoF3) (see Fig. S2 in the supplemental material). In the case of the promoter region of nolR, expression activity was observed only when a downstream (+24 bp) sequence (which included a likely ribosome binding site and alternative ATG) was included in the promoter-egfp fusion. In the S. meliloti strains Rm1021 and Rm2011 (used in this study), nolR is inactivated by a frameshift mutation (32). In the presence of a functional copy of nolR, activity from this promoter region was reduced (data not shown), consistent with the negative autoregulation properties of NolR as reported previously (33). Despite a previous study reporting variation in nolR expression with increasing population density in S. meliloti AK6321 (34) and the presence of an ExpR binding site upstream of nolR in S. meliloti 2011, ExpR did not greatly affect the activity detected from the nolR promoter region in our study. One possibility is that this region of DNA carries multiple promoters, which may depend upon some interaction between ExpR and AHLs and signals not present in our growth conditions. In S. meliloti AK631, nolR expression was affected by a number of environmental stimuli, such as nutrients, pH, and oxygen (34).
In a fourth category are DNA regions located upstream of an annotated gene which contain a binding site but do not contain detectable promoter activity under our conditions. These binding sites are located upstream of the genes SMb21025, SMb21071 (exoP2), SMc01711, SMc03149, SMc04032 (nesR), SMc04232, and SMc04258. For regions containing an ExpR binding site but lacking detectable promoter activity, it is possible that their downstream genes have falsely annotated translation starts (as was the case for nolR) or that mutations accumulating in our lab strain Rm2011 have rendered their promoters inactive. Another possibility is that these promoters are dependent upon external signals not present in these growth conditions. Yet another possibility is that these binding sites are components of ExpR/AHL-regulated promoters but that their activity is too weak for our methods of detection. For example, Patankar and González (35) reported a phenotype associated with nesR (SMc04032) disruption. However, we were unable to detect any activity from the nesR promoter, in either the presence or absence of ExpR or AHLs.
In a fifth category is the occurrence of an ExpR binding site located inside a coding region. The gene SMb20391 is annotated as a cellulose synthase. Using PatScan, we located an ExpR binding site downstream of the annotated ATG translation start for this gene. This site binds to His6-ExpR in a gel shift assay, but we have not tested for promoter activity surrounding the binding site. In one other case, the ExpR binding site controlling activity of the promoter of exoH is located inside the coding region of the upstream gene SMb20953.
Several promoter regions do not appear to bind to His6-ExpR in our gel shift assays despite a significant dependence upon ExpR/AHL for expression activity. These include the promoters of wggR, SMc04171, and SMb20911 (see Fig. S2 in the supplemental material). Expression of these genes was previously reported to be controlled by ExpR and AHLs (12), and we have reproduced those results. (See also a recent study by Gao et al. [36] for the dependence of wggRp on various concentrations of AHLs.) We do not know why these promoters do not bind to His6-ExpR in our gel shift assays (see Fig. 3C for wggR and SMb20911) despite strong promoter activation (>10-fold for wggR and >3-fold for SMc04171) and repression (>30-fold for SMb20911) dependent upon ExpR and AHLs. Nonetheless, we selected the wggR promoter region as a case study and looked for any possible location in the sequence that might contain a weak ExpR binding site. Similarly to the approach taken for expRp, we replaced selected nucleotides at two of the best-fitting locations to increase similarity to the ExpR binding site consensus. While these alterations did allow binding between His6-ExpR and the altered promoter region of wggR (Fig. 3D), they did not affect the activity of wggRp (data not shown). Therefore, for the promoters of wggR, SMc04171, and SMb20911, it is possible that the control exerted by ExpR/AHLs is through an intermediate(s) or that their binding to ExpR is too weak to be detected in our assay or dependent on factors absent from the assays.
Correlating promoter response to AHL concentrations in growth medium.In a previous study, it was shown that the ExpR-dependent promoter activities of sinR, sinI, and expR are not only dependent upon the presence of AHLs but also sensitive to the level of AHLs (16). In that study, it was demonstrated that the promoter of sinI responds to very low concentrations (<5 nM) of supplemented AHLs in a culture of a mutant incapable of producing its own AHLs. In contrast, the promoters of expR and sinR required substantially higher concentrations (50 nM and 200 nM, respectively) of supplemented AHLs before a response was observable. In this study, we applied the same approach to all of the promoters that respond to the presence of ExpR and AHLs. A strain incapable of producing AHLs (SM2B4001, sinI mutant) was grown in the presence of various concentrations of supplemented C16:1-HL. The resulting change in promoter activity relative to the concentration of C16:1-HL, as determined by promoter-egfp fusions, is presented in Fig. S3 in the supplemental material and summarized in Fig. 6, showing the minimal and maximal AHL concentrations that induced a change in promoter activity after 40 h. Interestingly, each promoter responded to a specific range of AHL concentrations. For example, similar to sinIp, the promoters of phrR and SMb20911 responded to very low levels of AHLs (5 to 10 nM). However, unlike sinIp, their response to the presence of AHLs was negative. All other promoters required higher levels of AHLs for a response. Addition of 50 to 100 nM was sufficient to induce regulation of the promoters of expR, SMc04237, SMb21543, SMa2111, and genes controlling exopolysaccharide production (exoH, exsH, exoI, wgeA, and wgaA) (see Fig. S3 in the supplemental material). Interestingly, all of these promoters respond positively to the addition of AHLs. In contrast, most of the promoters that are repressed by AHLs required a higher level (100 to 1,000 nM) before a response was observable (see Fig. S3). These include promoters of SMc04059, sinR, SMc01524, SMc02378, and SMc03864, which responded at 100 to 200 nM, and SMc04246, visN, SMb21135, and exoF3, which responded at 500 nM. Thus, from these data, a fascinating pattern emerges: promoters repressed by AHLs tend to require higher levels of AHLs for this response, while promoters activated by AHLs tend to begin responding at lower levels. Three exceptions to this pattern are the promoters of phrR, SMc04059, and SMb20911, which respond negatively to AHLs.
Overview of the sensitivity of ExpR-regulated promoters in the sinI mutant supplemented with different concentrations of AHL. The source data from which this overview is derived are shown in Fig. S3 in the supplemental material. Solid lines indicate a positive response of a promoter to AHL. Broken lines indicate a negative response. Line beginnings and endings indicate the range of AHL concentrations to which the promoter responded, indicated by asterisks in Fig. S3. The following concentrations (nM) were used: 0, 5, 10, 50, 100, 200, 500, 1,000, 2,000, 5,000, and 10,000.
Interestingly, the majority of promoters did not respond to changes in AHL levels above 2,000 nM. This suggests that most promoters have a specific lower and a common upper limit in their sensitivity to AHLs. Between these limits, promoter activity varies with AHL concentrations.
Another interesting observation regards the opposing effects on promoter activity from lower versus higher levels of AHL. For example, the promoter of sinI is activated by low levels of AHL while higher levels of AHL reduced activity (16). In a similar fashion, the activity profiles of the promoters of wgaA and wgeA were almost inactive in the absence of AHLs (Fig. 4; also, see Fig. S2 in the supplemental material) and increasingly active with increasing levels of AHL to a maximal activity at 500 nM (see Fig. S3 in the supplemental material). Intriguingly, as the levels of AHL were further increased to 2,000 nM, these promoters responded by decreasing in activity, so that both exhibited a lower activity (wgaAp, 5-fold; wgeAp, 3.5-fold) at 2,000 nM compared to that at 500 nM. Of all the promoters measured in this study, only those controlling the wga and wge operons and sinI showed such clear double-response effects that depend on AHL levels. In the case of sinIp, the second (negative) response is mediated not by an ExpR binding site located upstream of sinI itself but by another site upstream of sinR. Binding of ExpR to the site upstream of sinR results in a decrease of sinR expression and thus a decrease in SinR-dependent sinI expression (16). Likewise, in the case of wga and wge promoters, the second response may be due to the ExpR/AHL-dependent regulation of other genes related to the activity of these promoters.
DISCUSSION
QS is a social strategy that facilitates the detection of population density and the appropriate regulation of the production of common goods and other survival-related strategies. The interest in defining AHL-based QS regulons is evident from the number of studies that have focused on regulon determination and include organisms such as Burkholderia (4), Pseudomonas (5), Vibrio (6–9), Agrobacterium (10), and Sinorhizobium (11–13). S. meliloti contains genes which code for at least 8 LuxR-type proteins but only one AHL synthase, SinI. ExpR, SinI, and SinR, together with the long-chain AHLs produced by SinI, are essential for the Sin QS system. To date, only one of the LuxR-type proteins, ExpR, has been shown to be dependent upon the long-chain AHLs produced by SinI. The results in this study strongly indicate that SinR is independent of the long-chain AHLs for its activation of the promoter of sinI (Fig. 2). Likewise, VisN and VisR are also LuxR-type regulators whose activities are independent of AHLs (25). Interestingly, the N-terminal receiver domains of 11 other LuxR-type proteins, including TraR of A. tumefaciens, LasR of P. aeruginosa, and LuxR of V. fischeri, have been aligned and the conserved AHL-interacting residues highlighted (3). Compared to that analysis, ExpR contains most of the conserved AHL-interacting residues, while SinR, VisN, and VisR appear to differ from the consensus at the most conserved region (residues 58 to 77) upon alignment using CLUSTALW (data not shown). This fits with our observations that ExpR binds to an AHL and controls expression of the genes coding for VisN, VisR, and SinR, placing them under QS regulation, while the SinR, VisN, and VisR proteins have activities that are independent of AHLs. VisN, VisR, and SinR have not been characterized with respect to the presence or nature of an activating ligand. This is partly due to the highly insoluble nature of these proteins upon overexpression (our unpublished data). ExpR, on the other hand, is soluble upon overexpression, even in the absence of AHLs, making it suitable for this study.
Regulatory targets of ExpR.A variety of methods have been used to detect S. meliloti genes that are regulated by QS, including transcriptomics and proteomics (Fig. 1B). Microarray approaches found over 100 genes (13) and almost 500 genes (12) regulated by ExpR in the presence of AHLs. A protein two-dimensional gel separation approach found 38 proteins affected at least 2-fold in their accumulation (11) by ExpR in the presence of AHLs. Curiously, the overlap of genes identified as being regulated by ExpR in different studies is surprisingly low (Fig. 1B), prompting some speculation over differences in bacterial strains and culture conditions (37). Furthermore, each promoter differs in its response to QS, both in degree and in timing (Fig. 4; also, see Fig. S2 in the supplemental material). Some genes respond early, for example, because of a higher sensitivity to AHL accumulation, while others require higher levels of AHLs before a response is observable (Fig. 6) (see also Table S1 in reference 12). Therefore, genes detected as controlled by ExpR in the presence of AHLs will depend upon the point in the growth phase at which the mRNA or proteins are harvested and the method used for the harvest.
The total number of ExpR binding sites discovered thus far in S. meliloti is 33, dispersed among all three replicons. Most of the ExpR binding sites in S. meliloti also appear in S. medicae and S. fredii (see Table 1; also, see Table S3 in the supplemental material), although to our knowledge these have never been tested. In S. meliloti, eight sites are located upstream of operon-like arrangements. When genes within operon-like arrangements are taken into account, these 33 sites appear to control 66 to 71 genes. The study by Gurich and González (12) is the most sensitive study to date, revealing a total of 473 genes whose expression is dependent upon ExpR in the presence of AHLs, slightly less than 8% of the genome (38). Within this group of 473, only 25 genes are located downstream of an identified ExpR binding site, either within an operon-like structure or monocistronic. But perhaps this should be expected given that some genes directly regulated by ExpR are themselves characterized as transcription regulators.
One prominent example is phrR encoding a global transcription regulator. Expression activity of this gene was previously identified as responding to various stress conditions, such as low pH and high concentrations of ethanol, Zn2+, Cu2+, or H2O2 (39). More recently, in Rhizobium leguminosarum, a gene highly similar to phrR, praR, was shown to be integrated in QS regulation (40), including repression of praR expression by the R. leguminosarum ExpR homologue. According to our data, the S. meliloti ExpR also negatively regulates the promoter of phrR in the presence of AHLs.
Perhaps the most important novel ExpR site is located upstream of expR, which presumably confers the property of self-regulation. Together with the other previously reported sites located upstream of sinR (16) and sinI (20, 21), these three sites may explain the positive feedback loop (at low AHL levels) and the negative feedback loop (at high AHL levels) by which the Sin/ExpR system appears to control AHL levels. Most genes preceded by a novel ExpR binding site are not yet characterized for their function, such as SMc04237 (unknown function), SMc03150 (uncharacterized transcription regulator), SMb21543 (putative adenylate cyclase), and SMa2111 (putative hemolysin-type Ca+-binding protein).
Characteristics of promoter regulation by ExpR.Of considerable interest are the molecular mechanisms by which QS regulates its target genes. A theoretical model of bacterial transcription found regulatory logic functions of plausible complexity by varying only two factors: strength of interaction between regulatory proteins and the relative positions of the relevant protein-binding DNA sequences in the cis-regulatory region (41). One example of this is the TyrR protein of E. coli (reviewed in reference 42), which can act as a repressor or activator of transcription for its eight known target promoters. Transcription activation and repression by TyrR are effected by binding to its TyrR box, and the direction of regulation is determined by the location of the TyrR box relative to the promoter. Tyrosine is the most important ligand which controls multimerization states of TyrR and affects binding to the TyrR box. TyrR boxes are present in two basic classes. Strong TyrR boxes can bind to TyrR even in the absence of tyrosine, but weaker-affinity boxes require the presence of tyrosine. The mechanism for repression can involve the exclusion of RNA polymerase from the promoter or interference with the ability of bound RNA polymerase to form open complexes or to exit the promoter. For transcription activation, TyrR can bind upstream of a promoter and interact with the α-subunit of the RNA polymerase. Finally, intracellular levels of TyrR protein are thought to be critical for determining regulatory outcomes.
Somewhat analogous to the E. coli TyrR paradigm, there are at least three factors that determine the strength of the regulatory effect of the S. meliloti ExpR/AHL combination on its regulon: (i) the abundance of ExpR, (ii) the abundance of AHLs, and (iii) the DNA sequence in and around each ExpR binding site. Evidence for factor 1 was reported in a previous study (16), where levels of ExpR were controlled via expression from an IPTG promoter. In that study, various levels of ExpR intensified or weakened the promoter responses correspondingly.
Evidence for the abundance of AHLs as a determinant of gene expression was revealed in this study when AHL levels were varied in cultures carrying a promoter-egfp fusion (see Fig. S3 in the supplemental material). Promoter activity clearly depends upon the concentration of AHLs. However, in many cases, the effect of AHL addition on the ExpR-induced shift was only weakly apparent, if at all (Fig. 3). Furthermore, a previous study using atomic force spectroscopy found that the strength of interaction between ExpR and its DNA binding site upstream of sinI was significantly increased upon the addition of AHLs (21). Evidence for the DNA sequence within and surrounding the ExpR binding site as one determinant of gene expression is suggested by the banding patterns in Fig. 3. We have not presented data in this study showing the titration of His6-ExpR against target DNA in the gel shift assays or any other measurement of ExpR-DNA binding strength. However, we believe that such data could provide important and interesting verifications of the conclusions drawn from this study. ExpR-DNA binding is only one step in a multistep process of transcription activation and is therefore not necessarily a good indication of the strength of transcription activation. Examples of this are the ExpR binding sites upstream of the genes wgeA and wgaA. These promoters are strongly activated by ExpR in their transcription activity but do not appear to produce strong shifts in the gel shift assay. However, in both transcription activation and repression, the strength of the ExpR-DNA interaction is arguably one of the most critical steps in the regulation. This is supported by our study of expRp, in which alterations in the ExpR binding site in expRp affected not only the strength of the shift in a gel assay (Fig. 3D) but also promoter activity in the presence of ExpR (Fig. 5). Based on these data, we propose a testable hypothesis: at least one determinant of varying promoter sensitivity to AHLs is the DNA sequence to which ExpR binds, in which binding strength is stronger for sites that are more similar to the ExpR consensus.
Also relevant is the location of the binding site with respect to the promoter and transcription start, which may determine whether the regulation is positive or negative (Table 1; also, see Fig. S1 in the supplemental material), as is the case for TyrR (42). For our analysis, the experimentally determined transcription starts were reported previously (30) and additionally independently determined in the case of sinR (20), sinI (16), and expR (this study). All of the ExpR-binding promoter regions which were activated by ExpR contained a binding site either covering or upstream of the −35 region (see Fig. S1). An example of this is sinIp, where the ExpR site is not at −35 but at −75 (see Fig. S1). Activation of this promoter via ectopic expression of sinR, although dependent upon an intact −35 region, does not require either ExpR (16) or the ExpR binding site (unpublished data). This fits with the current model, in which SinR binds at or close to the −35 region and is necessary for a basal activity of sinIp, while ExpR binds at −75 and enhances sinIp activity. In contrast to the ExpR-activated promoters, if the ExpR binding site is downstream of the −35 region, ExpR represses promoter activity. It is likely that the mechanism of repression is via ExpR covering the −10 region or the transcription start (+1). Two exceptions to these generalizations are the promoters of SMc04059 and SMc01524. In the case of SMc04059, the ExpR binding site is upstream of the −35 region. We cannot at this stage exclude the possibility of an alternative promoter with a transcription start that is closer to this binding site. In the case of SMc01524, the ExpR binding site covers the −35 region in a manner similar to the positively regulated promoters. We do not know why repression occurs in this case. One possibility is that ExpR represses activity via competition with the RNA polymerase, or that the binding site covers the transcription start of an alternative promoter.
Biological function of a quorum sensing program.Although the long-chain AHLs are suspected not to cross the double membrane barrier of Gram-negative bacteria via diffusion (43–45), the presumptive transport systems and processes involved remains elusive. An individual which relies solely upon imported AHLs might be expected to have a weaker response to QS than an individual that engages in both AHL importation and production. Therefore, mutant cultures incapable of producing their own AHLs cannot be directly compared to wild-type cultures. However, the use of an AHL mutant strain in detecting the sensitivities of selected promoters to supplemented AHLs reveals a variety of AHL sensitivities and suggests that these promoters are organized in a program of QS regulation. Data from this study suggest that as AHLs accumulate in a growing colony, positively regulated promoters are programmed to respond prior to the negatively regulated promoters. The clearest example of this is the activation of the expression of genes controlling exopolysaccharide production and the repression of genes controlling motility. Such inverse regulation appears to be a general feature of many bacteria (46), including Pseudomonas (47), where regulation is achieved via the signal molecule cyclic di-GMP (c-di-GMP). Not only does the quorum sensing program in S. meliloti fit nicely with the pattern of inverse regulation, it also provides a fascinating glimpse of how the QS regulation exhibits dynamic behavior and precision timing. For example, individuals on the periphery of a colony might be expected to encounter lower levels of AHLs than those nearer the center, depending upon parameters such as rates of AHL diffusion and degradation. The lower level of AHLs (e.g., <500 nM) encountered by the peripheral individuals may be insufficient to repress the promoter for the visNR master regulator of the motility regulon (48). Our results indicated that visNR expression is not repressed until AHL concentrations are ≥500 nM (Fig. 6; also, see Fig. S3 in the supplemental material). This would allow these individuals to continue producing more flagella and so enhance mobility. Furthermore, these individuals would encounter sufficient AHLs to activate exopolysaccharide production (50 to 500 nM), which also enhances sliding mobility that is not necessarily dependent upon the presence of flagella (49). In contrast to the peripheral individuals, those near the center of the colony may encounter higher levels (≥500 nM) of AHLs and respond by reducing flagellum production. In a similar way, galactoglucan production appears to be reduced in response to higher levels of AHLs (Fig. 6; also, see Fig. S3 in the supplemental material). Thus, lower levels of AHLs stimulate an increase in exopolysaccharide production but do not affect flagellum production. Conversely, higher AHL levels would result in a simultaneous reduction in the production rates of galactoglucan and flagella. This is presumably a finely tuned solution to avoid unnecessary and costly productions by individuals in the center of a colony with limited access to nutrients. Thus, the rates of galactoglucan and flagellum production may be tightly bound to local AHL concentrations, which are in turn determined by such factors as the shape and size of the colony. This may serve to ensure the production of sufficient levels of galactoglucan and flagella at the appropriate localities within a colony.
In this study, we have explored the mechanisms of the S. meliloti ExpR QS transcriptional regulatory network. The data suggest a type of quorum-sensing program whereby variation in a single input, AHL concentration, is sufficient to generate a tremendous diversity in the sensitivity, direction, and extent of promoter response. Although the details required to account for all the variation in this network appear to be multiple and complex, there are at least two features of the ExpR regulon which could at least partially account for the variation: variations within the DNA sequence of the ExpR binding site, which suggest different binding strengths, and the location of the ExpR binding site with respect to the promoter and transcription start. Both explanations could be tested and provide attractive focal areas for future studies. It would also be interesting to see if QS regulatory networks in other bacteria make use of similar regulatory strategies.
ACKNOWLEDGMENTS
We thank Elizaveta Krol for providing the pLK101 and pLK121 plasmid constructs and, along with other members of the Becker lab, for helpful discussions.
We acknowledge financial support from the LOEWE program of the State of Hesse, Germany (in the framework of the Center for Synthetic Microbiology, SYNMIKRO, Marburg), from the German Research Foundation (SPP 1617), and from the Development and Promotion of Science and Technology talents project (DPST), the Royal Thai Government.
FOOTNOTES
- Received 1 March 2013.
- Accepted 7 May 2013.
- Accepted manuscript posted online 17 May 2013.
Supplemental material for this article may be found at http://dx.doi.org/10.1128/JB.00234-13.
- Copyright © 2013, American Society for Microbiology. All Rights Reserved.
