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Applied and Environmental Microbiology, May 2009, p. 2629-2637, Vol. 75, No. 9
0099-2240/09/$08.00+0     doi:10.1128/AEM.02145-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

AguR Is Required for Induction of the Streptococcus mutans Agmatine Deiminase System by Low pH and Agmatine

Yaling Liu, Lin Zeng, and Robert A. Burne^*

Department of Oral Biology, University of Florida, Gainesville, Florida

Received 16 September 2008/ Accepted 22 February 2009

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Acidic conditions and the presence of exogenous agmatine are required to achieve maximal expression of the agmatine deiminase system (AgDS) of Streptococcus mutans. Here we demonstrate that the transcriptional activator of the AgDS, AguR, is required for the responses to agmatine and to low pH. Linker scanning mutagenesis was used to create a panel of mutated aguR genes that were utilized to complement an aguR deletion mutant of S. mutans. The level of production of the mutant proteins was shown to be comparable to that of the wild-type AguR protein. Mutations in the predicted DNA binding domain of AguR eliminated activation of the agu operon. Insertions into the region connecting the DNA binding domain to the predicted extracellular and transmembrane domains were well tolerated. In contrast, a variety of mutants were isolated that had a diminished capacity to respond to low pH but retained the ability to activate AgDS gene expression in response to agmatine, and vice versa. Also, a number of mutants were unable to respond to either agmatine or low pH. AguD, which is a predicted agmatine-putrescine antiporter, was found to be a negative regulator of AgDS gene expression in the absence of exogenous agmatine but was not required for low-pH induction of the AgDS genes. This study reveals that the control of AgDS gene expression by both agmatine and low pH is coordinated through the AguR protein and begins to identify domains of the protein involved in sensing and signaling.

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The agmatine deiminase system (AgDS) of Streptococcus mutans is encoded by the aguBDAC operon (11). Agmatine enters the cell via an agmatine-putrescine antiporter (AguD), where it is hydrolyzed to N-carbamoylputrescine and ammonia by agmatine deiminase (AgD; EC 3.5.3.12), encoded by aguA. Putrescine carbamoyltransferase (EC 2.1.3.6), encoded by aguB, mediates the phosphorolysis of N-carbamoylputrescine, yielding putrescine and carbamoylphosphate. Finally, a phosphate group is transferred from carbamoylphosphate to ADP by carbamate kinase (EC 2.7.2.2), the product of the aguC gene, to generate ATP, CO₂, and NH₃. Putrescine is then exchanged for agmatine via the antiporter.

Streptococcus mutans is the etiological agent of dental caries (17). A key virulence attribute of this organism is acid tolerance, since catabolism of dietary carbohydrates lowers the pH of dental plaque to values below 4.0 (3). S. mutans has multiple strategies to cope with low pH and gains a competitive advantage over acid-sensitive bacteria under acidic conditions (19, 21). Although a variety of oral streptococci utilize ammonia generation from urea or from amino acids as a fundamental acid tolerance mechanism (19), alkali production was not considered a mechanism employed by S. mutans to cope with acidification until the AgDS was characterized for S. mutans UA159 (11) and, subsequently, for other mutans streptococci (13). The primary roles of the AgDS in S. mutans and other mutans streptococci appear to be to catabolize agmatine, to produce ATP, and to augment acid tolerance (11, 12).

The aguR gene, which encodes a trans-acting factor required for efficient AgDS expression, is located 239 bp upstream of aguB and is transcribed in the opposite direction (12). AguR is a putative LuxR-like transcriptional regulator belonging to the FixJ-NarL superfamily, which includes two-component response regulators involved in quorum sensing (10). AguR is 318 amino acids long, with a predicted molecular mass of 37,611 Da, and has several noteworthy structural features (Fig. 1). Computer algorithms at http://smart.embl-heidelberg.de/and http://bp.nuap.nagoya-u.ac.jp/sosui/ predict an N-terminal signal sequence and as many as four membrane-spanning domains that may allow for a substantial portion of the protein to be exposed to the environment. A predicted helix-turn-helix (HTH) motif of the LuxR family is located in the C terminus of the protein and likely mediates DNA binding. It was reported previously that induction of the AgDS of S. mutans is optimal at low pH when agmatine is present (12). In this study, we tested the hypothesis that agmatine and low-pH induction of the AgDS of S. mutans is mediated through the AguR transcriptional regulator.

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FIG. 1. Predicted topology and linker insertions for AguR. The four-transmembrane-domain model is based on computer algorithms from http://smart.embl-heidelberg.de/ and http://bp.nuap.nagoya-u.ac.jp/sosui/. Transmembrane domains are indicated by boxes and numbered in the top left corners. The circles indicate that insertions adjacent to these residues resulted in altered responses to pH and agmatine. The square indicates that insertions at this residue resulted only in altered responses to agmatine. The diamond indicates that insertions at this point in AguR altered only pH responsiveness. The hexagons indicate that mutagenesis at these residues resulted in no significant changes in the response to stimuli.

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Bacterial strains, growth conditions, and reagents.
S. mutans UA159 and its derivatives were maintained in brain heart infusion broth (BHI; Difco Laboratories, Detroit, MI) at 37°C in 5% CO₂ and 95% air, with antibiotics added at the following concentrations when necessary: kanamycin (Km), 1 mg ml^–1; erythromycin (Em), 5 µg ml^–1; and spectinomycin (Sp), 1 mg ml^–1. To monitor AgDS expression, batch cultures of S. mutans were grown in a tryptone-vitamin (TV) base medium (5) supplemented with 25 mM galactose, with or without 10 mM agmatine, to an optical density at 600 nm (OD₆₀₀) of 0.5. Escherichia coli strains were grown in Luria-Bertani (LB) medium supplemented with antibiotics at the following concentrations: ampicillin (Ap), 100 µg ml^–1; Km, 50 µg ml^–1; Em, 500 µg ml^–1; and Sp, 50 µg ml^–1. Chemical reagents and antibiotics were obtained from Sigma (St. Louis, MO).

For studies on pH- and agmatine-dependent regulation of AgDS expression, S. mutans strains were grown in continuous culture in a Biostat i Twin Controller chemostat (Braun Biotech, Inc., Allentown, PA) in a tryptone-yeast extract (TY) medium (29) supplemented with 25 mM galactose at a dilution rate (D) of 0.3 h^–1. Cultures were maintained at pH 5.5 or pH 7.0 by the addition of 2 M KOH. Samples were harvested at steady state, which was defined as cultivation under given conditions for at least 10 generations. Where indicated, cultures were pulsed with 10 mM agmatine for 1 h prior to sampling.

DNA manipulation.
Genomic DNA was isolated from S. mutans UA159 as previously described (7). Plasmid DNA used in sequencing reactions was prepared from E. coli by use of a miniprep kit (Qiagen, Valencia, CA). Restriction and DNA-modifying enzymes were purchased from Life Technologies Inc. (Rockville, MD) or New England Biolabs (Beverly, MA).

Construction of reporter gene fusions.
A 157-bp fragment upstream of the aguB start codon (P_aguB) was amplified using primers PaguB-S and PaguB-AS (Table 1), with inserted SacI and BamHI restriction sites to facilitate cloning. The promoter region was fused to a promoterless lacZ gene derived from Streptococcus salivarius on pMC195 (6). After confirming the correct sequence of the promoter fusion by DNA sequencing, the P_aguB-lacZ construct was cloned into plasmid pMC340B (30), which allows for integration of the gene fusion in S. mutans as a single copy at the mtlA-phnA locus, and the construct was transformed into S. mutans UA159 to create strain WTZ. Transformants were selected on BHI agar with Km and screened by PCR to confirm that the desired integration had occurred. Strain WTZ was transformed with plasmid pMSP3535 (4), which allows for nisin-inducible expression from the nisA promoter, to generate WTZ/V, or with a nisR/nisK-deficient derivative of pMSP3535 (pMSP-nisRK) to generate WTZ/V-RK. pMSP-nisRK was constructed by removal of an EcoRV fragment from pMSP3535 to delete both the nisR and nisK genes. Plasmid transformants were selected on BHI agar containing Em.

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TABLE 1. Primers used in this study

Deletion of aguR.
Primers aguR1-S and aguR1-AS (Table 1) were used to amplify a 2.2-kbp aguR-containing DNA fragment. The PCR product was then digested with BsrGI and HindIII and ligated with an Sp resistance cassette. The ligation mixture was introduced into strains WTZ, WTZ/V, and WTZ/V-RK (Table 2) by natural transformation to generate R^–, R^–/V, and R^–/V-RK, respectively, and transformants were selected on BHI agar with Sp. The correct mutations were confirmed by DNA sequencing.

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TABLE 2. S. mutans strains used in this study

Construction of aguR-complemented strains.
The expression vector pMSP3535, which allows for controlled expression of inserted genes by addition of subinhibitory concentrations of nisin to the growth medium (4, 20), was used for complementation studies with the aguR gene of S. mutans. To construct an aguR-complemented strain, a 1.1-kbp fragment of aguR, containing the aguR coding sequence and the native ribosome binding site, was amplified with primers aguR2-S and aguR2-AS to introduce BamHI and XhoI restriction sites for subsequent cloning (Table 1). The PCR product was then digested with BamHI and XhoI and cloned into pMSP3535, ensuring that transcription of aguR was in the same orientation as that of the nisin promoter, resulting in pMSP-aguR. Faithful amplification of the gene was confirmed by DNA sequencing with primers pMSP3535F and pMSP3535R (Table 1). The complemented strain (R^–/R) was constructed by introducing pMSP-aguR into the AguR-deficient mutant of S. mutans (R^–) by natural transformation, and transformants were selected on BHI agar with Em.

Deletion of aguD.
Primers aguD-S and aguD-SmaI-AS (Table 1) were used to amplify a 0.6-kbp region upstream of aguD. Primers aguD-SmaI-S and aguD-AS (Table 1) were used to amplify a 0.6-kbp region downstream of aguD. The PCR products were digested with SmaI and ligated to a nonpolar Km resistance cassette from pALH124 (2). The ligation mixture was introduced into S. mutans UA159 by natural transformation to generate an aguD mutant, and bacteria were plated on BHI agar with Km. Double-crossover mutants were confirmed by PCR.

Linker scanning mutagenesis.
Mutagenesis was performed with the GPS-LS linker scanning system (New England Biolabs) according to the supplier's instructions. The GPS-LS system allows for random insertion of 15 nucleotides into the gene of interest, with only two of the six possible reading frames creating stop codons. In the other four reading frames, the insertion introduces five amino acids into the protein of interest. A transposon (Tp) donor plasmid (pGPS5; Km resistant) and the TnsABC transposase were used to mutagenize plasmid pMSP-aguR in vitro. The mutagenized plasmids were electroporated into E. coli by selection on LB agar containing Km, and the colonies were pooled for isolation of plasmids. The Tp-containing XbaI-SphI fragments harboring the mutated aguR genes from pools of targeted plasmids were cloned back into pMSP3535 to construct pMSP-aguR::Tp plasmids. One hundred individual pMSP-aguR::Tp plasmids were each digested with PmeI to release the Tp and then were religated. The resulting clones were selected on BHI agar containing Em and screened for Km sensitivity. Insertions were located by DNA sequencing with the primers pMSP3535F and pMSP3535R (Table 1). Selected pMSP-aguR::Tp plasmids were used to transform the R^– strain, and transformants were selected on BHI agar containing Em.

β-Galactosidase assay.
Measurements of β-galactosidase activity were obtained using a modification of the Miller protocol (31). Briefly, cells were harvested by centrifugation, washed once with Z buffer (60 mM Na₂HPO₄, 40 mM NaH₂PO₄, 10 mM KCl, 1 mM MgSO₄, and 5 mM β-mercaptoethanol, pH 7.0), and resuspended in 1/10 of the original culture volume in the same buffer. One milliliter of the suspension was used to determine the OD₆₀₀. Another 500 µl of the sample was vortexed with 25 µl of a toluene-acetone mix (1:9) for 2 min and then kept at 37°C. A reaction was initiated by adding 100 µl of ONPG (o-nitrophenyl-β-D-galactopyranoside) solution (4 mg/ml) and terminated by the addition of 500 µl of 1 M Na₂CO₃. Samples were centrifuged at maximum speed for 2 min using a tabletop centrifuge, and the OD of the supernatant fluid was measured at 420 and 550 nm. Activity was expressed in Miller units (31).

Expression and purification of recombinant AguR in E. coli.
A 324-bp fragment encoding the C-terminal one-third (amino acids 210 to 318) of AguR was amplified by PCR using primers aguR3-S and aguR3-AS, with BamHI and PstI sites added (Table 1). The PCR product was then digested with BamHI and PstI and inserted in frame behind the malE gene on pMAL-p2X (New England Biolabs) to create a maltose binding protein fusion. Expression of the recombinant AguR protein was induced with 1 mM IPTG (isopropyl-β-D-thiogalactopyranoside) for 3 h, and protein lysates were prepared by homogenization with a BeadBeater (Biospec, Bartlesville, OK). The recombinant protein was purified using an amylose column as recommended by the supplier. Polyclonal AguR antibody was elicited in rabbits with the purified recombinant AguR protein at Lampire Biological Laboratories (Pipersville, PA).

Western blot assays of AguR protein.
Mid-exponential-phase (OD₆₀₀ 0.5 to 0.6) cells were centrifuged and washed twice with Tris-buffered saline (10 mM Tris, 0.9% NaCl, pH 7.4). Whole-cell lysates were obtained by homogenization in sodium dodecyl sulfate (SDS) boiling buffer (60 mM Tris, pH 6.8, 10% glycerol, and 5% SDS) in the presence of glass beads, followed by centrifugation at 2,000 x g for 10 min (8). Protein samples were separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE), blotted onto polyvinylidene difluoride membranes, and incubated with affinity-purified (23) AguR antibody, followed by peroxidase-conjugated goat anti-rabbit immunoglobulin G (KPL, Gaithersburg, MD). Signals were developed using a SuperSignal West Femto chemiluminescence kit (Thermo, Waltham, MA).

Real-time PCR.
Extraction of RNA, reverse transcription-PCR, real-time PCR, and copy number normalizations were performed as previously described (1). The primers used for reverse transcription-PCR and real-time PCR for the aguA gene are described elsewhere (12). The aguR-specific primers (aguR-RT-S and aguR-RT-AS) were designed using Beacon Designer 4.0 software (Premier Biosoft International, Palo Alto, CA) and are detailed in Table 1.

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Involvement of AguR in induction by pH and agmatine.
In a previous study using batch cultivation of cells, agmatine deiminase activity was optimal in cells growing in acidified medium in the presence of agmatine (12). To assess whether AguR exerted transcriptional control over the agu operon in response to pH and agmatine, the wild-type strain and strain R^– carrying the aguB promoter (P_aguB) fused to a Streptococcus salivarius lacZ gene were examined. The strains were cultured batchwise in TV medium containing 25 mM galactose, with or without 10 mM agmatine. Cells were grown to mid-exponential phase, and LacZ activity was measured. The expression of P_aguB was induced 2.5-fold by growth in the presence of 10 mM agmatine in the wild-type background, whereas the R^– strain displayed a basal level of P_aguB expression and no agmatine induction was evident (Fig. 2). WTZ and R^– mutants were also grown to mid-exponential phase in TY medium that was acidified using HCl to pH 5.5 or buffered at pH 7.0 with 50 mM potassium phosphate buffer, and LacZ activity was measured. In the wild-type background, cells expressed twofold higher LacZ activity from P_aguB at pH 5.5 than at pH 7.0, whereas no significant difference in P_aguB expression was detected in the R^– strain at pH 5.5 and pH 7.0 (Fig. 3).

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FIG. 2. LacZ specific activities in AguR variants of S. mutans carrying PaguB-lacZ gene fusions. Cells were grown in TV medium containing 25 mM galactose, with or without 10 mM agmatine, to mid-exponential phase, and LacZ specific activities were measured. The values shown represent the means and standard deviations for three independent experiments.

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FIG. 3. LacZ specific activities of AguR variants of S. mutans carrying PaguB-lacZ. Cells were grown to mid-exponential phase in TY broth containing 25 mM galactose with 10 mM agmatine that had been acidified to pH 5.5 with HCl (TY-HCl) or in TY broth buffered at pH 7.0 (TY-KPB), and the LacZ specific activities were measured. The values shown represent the means and standard deviations for three independent experiments.

There are a number of limitations to the use of batch cultures for studies of the effect of pH, including the fact that growth rates can be substantially different in cells growing at pH 5.5 and at pH 7.0, as can the amount of available carbohydrate, which may affect agu gene expression through catabolite repression (12). In contrast, continuous chemostat culture allows for tight control over growth rates and the availability of limiting nutrients. To test if there was indeed a specific effect of pH that was independent of carbon catabolite repression or growth rate, WTZ and R^– mutants were cultured in a chemostat at pH 7.0 and pH 5.5 to steady state, followed by a 10 mM agmatine pulse for 60 min. The results, shown in Table 3, revealed that the effects of agmatine and low pH on P_aguB expression were similar to those observed during batch cultivation (Fig. 2 and 3), reinforcing the observation that AguR is required for activation of the agu operon by low pH and agmatine.

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TABLE 3. LacZ activity expressed from PaguB in S. mutans strains growing in continuous chemostat culturec

Generation of linker scanning mutants of AguR.
To begin to define structure-function relationships in AguR, linker scanning mutagenesis was used to introduce insertions of five amino acids randomly throughout the protein (Table 4). A total of 34 unique mutants were characterized, with 12 insertions located in the predicted transmembrane domains, 15 in predicted extracellular domains, and 7 in predicted intracellular domains, including 2 within the HTH motif (Fig. 1). The mutants (RLS) were named according to the amino acid number preceding the site of insertion (Table 4). In addition, nine clones were characterized that contained nonsense codons that were predicted to lead to production of a truncated AguR protein (Table 4). All of the mutant proteins were expressed in a host with a complete deletion of the aguR gene, as detailed in Materials and Methods.

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TABLE 4. Linker mutation locations, inserted residues, and responsiveness of mutants to pH and agmatine in agu operon induction

To test whether the linker scanner insertions caused decreases in the levels of the AguR derivatives expressed in the R^– strain, possibly through decreased translation or stability of the altered proteins, clarified cell lysates from the wild-type and mutant strains were subjected to SDS-PAGE and Western blot analysis with an affinity-purified polyclonal antibody raised against the C-terminal third of AguR (Fig. 4). A 37-kDa band, which matches the predicted molecular mass of AguR, was detected in the WTZ strain. A lower-molecular-mass species, which we believe may be a cleavage product of AguR, was consistently observed in the Western blots. As expected, higher levels of AguR protein were observed in the R^–/R strain (Fig. 4), in which aguR was carried on pMSMP3535, a multicopy plasmid that can drive gene expression in gram-positive bacteria (4, 9, 14). The AguR protein could be detected in all RLS mutants at levels similar to that in the R^–/R strain (Fig. 4), which indicated that the linker scanning insertions did not have any major effects on AguR biogenesis or stability. It was also noteworthy that when protein samples from the WTZ strain were loaded in the same amount as that from the R^–/R strain and RLS mutants, only a very faint band could be detected, which is consistent with our observations that AguR is expressed at extremely low levels in the wild-type strain. By adding fivefold more protein from the WTZ strain, AguR protein and the lower-molecular-weight species could be detected (Fig. 4C).

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FIG. 4. Western blot analysis of wild-type, WTZ, R–/V, R–/R, and RLS mutant strains of S. mutans. Following SDS-PAGE, proteins were transferred to a nitrocellulose membrane and subjected to Western blotting using an affinity-purified AguR antibody at a dilution of 1:100. Lanes are marked with the strains examined. (A) Western analysis of various RLS insertion mutants and control strains (see Table 2). (B) The truncated derivative RLS 286 (lane 3) expresses a protein of the correct predicted molecular mass at a level comparable to that of the intact AguR protein (lane 1). Lane 2 contains an equivalent amount of protein from an aguR deletion mutant. (C) Western blot showing the presence of full-length AguR (37 kDa) and the potential processed form of AguR expressed as a single copy in the wild-type background, except that fivefold more protein was loaded in that lane (5xWTZ) (lane 1). Lane 2 harbors a control showing no reactive bands for an AguR-deficient mutant. Lanes 3 and 4 harbor one-fifth the amount of protein in lanes 1 and 2, but extracts were prepared from strains expressing plasmid-borne derivatives of mutant aguR genes (see Table 4).

Regions required for agmatine induction.
To begin to identify domains in AguR that may be required for agmatine induction, strains were cultured to mid-exponential phase in TV medium containing 25 mM galactose, with or without 10 mM agmatine, and LacZ activity was measured. As shown in Fig. 1 and Table 4, the RLS 46, 52, 53, 60, 65, 66, 67, 81, 88, 104, 105, 110, 168, 175, 209, 287, and 297 mutants did not respond to the presence of agmatine with changes in expression from the aguB promoter, similar to the strain lacking AguR (R^–/V). The RLS 41, 121, 158, 161, and 201 mutants showed around a 2.5-fold increase in P_aguB expression when they were grown with agmatine, which was similar to the level of induction seen in the complemented (R^–/R) strain (Fig. 1 and 2). Also, an increase in P_aguB expression of about fivefold in response to growth in agmatine was observed for the RLS 73, 103, and 106 mutants (Fig. 1 and 2).

Regions of AguR required for low-pH induction.
To determine whether mutagenesis of different domains of AguR would affect pH-dependent expression from the aguB promoter, the various strains were grown in TY broth containing 10 mM agmatine that was acidified to pH 5.5 or buffered at pH 7.0 (Fig. 1 and 3). Cells were harvested in mid-exponential phase, and LacZ activity was measured. Unlike the strain carrying a wild-type copy of the AguR protein, the RLS 41, 46, 52, 53, 73, 81, 88, 168, and 175 mutants showed no difference in P_aguB transcription at pH 5.5 or pH 7.0. An increase of about 1.3-fold in P_aguB transcription at low pH was detected in the RLS 60, 65, 66, and 67 mutants, compared with the 2-fold induction seen at pH 5.5 in the R^–/R strain. A twofold increase in P_aguB expression at low pH was observed for the RLS 103, 104, 105, 106, 110, 121, 158, 161, 201, and 209 mutants (Fig. 1 and 3).

Chemostat studies with AguR variants.
Since the effects of the mutated versions of aguR on aguB promoter expression were studied in batch culture, we examined whether a subset of the mutants did in fact have an aberrant response to low pH and agmatine in continuous culture. LacZ activity was measured in cells that were cultivated to steady state at pH 7.0 or pH 5.5 and then pulsed with 10 mM agmatine for 60 min. The RLS 110 strain expressed approximately threefold higher LacZ activity in response to agmatine, but no induction at pH 5.5 was noted (Table 3). Conversely, a twofold increase in P_aguB expression of RLS 41 was stimulated by the addition of 10 mM agmatine, but no difference in P_aguB expression in this strain was seen at pH 7.0 and pH 5.5 (Table 3). The RLS 46 and 175 mutants, in which responses to low pH and agmatine were eliminated, showed a basal level of P_aguB expression, and no induction by agmatine or low pH was detected (Table 3). Therefore, the results using chemostat-grown cells confirmed that AguR is required for the response to low pH and agmatine and that mutations in different regions of the predicted membrane-spanning or extracellular domains can affect pH and agmatine induction independently or can eliminate both responses.

Mutations in the HTH motif.
AguR is a LuxR-like transcriptional regulator in which the C-terminal HTH domain is implicated in DNA binding (28). However, the conserved acylated homoserine lactone binding region, typical of LuxR, could not be found in AguR. To confirm that the C-terminal HTH motif of AguR was involved in AguR-DNA binding, the RLS 102, 162, 174, 205, and 286 mutants, which contain truncated AguR proteins with termination codons before the HTH motif (Table 2; Fig. 1), were analyzed. First, real-time PCR was used to confirm that the truncated aguR genes could be expressed at a level comparable to that of the parental copy, and the data showed that the truncated aguR derivatives RLS 102, 162, 174, 205, and 286 were transcribed at a level similar to that of wild-type aguR in the R^–/R strain (data not shown). The RLS 286 mutant was the only mutant in which the truncated AguR protein had an overlapping region to the C-terminal one-third of AguR that was used to elicit the AguR antibody. By use of Western blot analysis, the truncated AguR protein in RLS 286 could be detected at levels comparable to that of wild-type AguR in the R^–/R strain (Fig. 4B). All of these mutants showed a basal level of P_aguB transcription, and no induction by agmatine or low pH could be observed (data not shown), suggesting that the agu operon could not be activated by inducing signals through AguR derivatives lacking the HTH motif.

Involvement of AguD in regulation of the AgDS.
AguD is a predicted agmatine-putrescine antiporter. Recently, CadC, which activates the transcription of the lysine decarboxylase genes of E. coli in the presence of low pH and lysine, was shown to sense the presence of extracellular lysine indirectly through the LysP permease. S. mutans UA159 and an AguD-deficient derivative were cultured in TV medium containing 25 mM galactose, with or without 10 mM agmatine. Expression of the AgDS was monitored by real-time PCR with the aguA (AgD) gene as a target. The data (Fig. 5) showed that in the absence of exogenous agmatine, aguA mRNA was about 100-fold more abundant in the aguD mutant than in the wild-type strain. When the strains were grown in the presence of 10 mM agmatine, a slight but statistically significant increase (1.5-fold) in aguA expression was seen in the AguD-deficient strain (Fig. 5A). Thus, in the absence of exogenous agmatine, AguD acts as a negative regulator of agu gene expression.

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FIG. 5. Real-time PCR monitoring of aguA mRNA. Reverse transcription was initiated from 1 µg of total RNA from the wild-type (WT) and AguD-deficient strains of S. mutans cultured in TV medium containing 25 mM galactose, with or without 10 mM agmatine (A), or in TY-KPB or TY-HCl containing 25 mM galactose and 10 mM agmatine (B). The amount of aguA cDNA was determined by real-time PCR using SYBR green. The data represent means ± standard deviations obtained from three different RNA preparations and reverse transcription reactions. Asterisks indicate that aguA gene expression in the mutant differed significantly from that in the wild-type strain under the same culture conditions (P < 0.05; Student's t test).

To test if AguD could also play a role in induction of agu gene transcription at low pH, the cells were cultured in TY-KPB or TY-HCl medium (pH 5.5) containing 25 mM galactose and 10 mM agmatine. The aguA gene was expressed at similar levels in the AguD-deficient and wild-type strains growing at either pH 7.0 or pH 5.5 (Fig. 5B), suggesting that AguD is not required for low-pH induction of the AgDS.

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Induction of the AgDS at low pH and in the presence of agmatine is believed to contribute to the competitive fitness of S. mutans in complex oral biofilms (11). Here we have shown that the AguR protein is required for responsiveness of agu gene transcription to low pH and to agmatine. The predicted extracellular domain (Fig. 1) between transmembrane regions 1 and 2 and the transmembrane domains themselves appear to be essential for the response to both pH and agmatine. Not surprisingly, the C-terminal HTH motif of AguR was absolutely required for activation of agu gene transcription. In many ways, the regulation of the AgDS of S. mutans is reminiscent of that of the lysine decarboxylase (cadAB) operon of E. coli (25), whose primary role appears to be to protect the organism from acid damage (26). Induction of cad gene expression occurs only in the presence of the inducer lysine under acidic conditions and requires the transcriptional activator CadC. CadC has a single transmembrane domain, and a substantial portion of the protein resides outside the cytoplasmic membrane. Conformational changes induced by low pH and binding of lysine are transduced to the N-terminal cytoplasmic portion of the protein, which contains the HTH domain (25). Recently, though, it was reported that sensing of lysine requires the LysP permease, which appears to interfere with the ability of CadC to activate transcription when external lysine concentrations are low (16).

The mutational analysis conducted here demonstrates that it is possible to isolate derivatives of AguR that lose the ability to activate transcription in response to pH but not agmatine, and vice versa (Fig. 2 and 3). Likewise, we were able to isolate mutant forms of AguR that lost the ability to respond to either signal. The introduced mutations likely either disrupt a binding site or cause conformational changes that inhibit normal AguR function, since our data show that diminished production or stability of the proteins is not responsible for the observed effects. The simplest interpretation of these findings is that AguR can bind to agmatine, which stimulates AguR binding to its target sequence, and that acidic conditions favor a conformation for AguR that makes signal transduction to the DNA binding domain more efficient. Such a model would be reasonable were it not for the fact that deletion of the aguD gene, which encodes the predicted agmatine-putrescine antiporter, dramatically altered agu gene expression in the absence of agmatine. It seems, therefore, that the mechanism for regulation of agu gene expression by agmatine may be similar to what was recently reported for regulation of cadAB expression by lysine, CadC, and LysP. Specifically, the lack of exogenous agmatine may result in an association between AguD and AguR that interferes with the interaction of AguR with its target. The addition of exogenous agmatine could destabilize this association. An alternative explanation is that internalized agmatine is the actual inducing signal and that the loss of the antiporter inhibits accumulation of agmatine in cells. However, if the predicted topology and orientation of AguR are correct, then induction by intracellular agmatine seems unlikely since the mutations that affect induction by agmatine do not map to intracellular domains. Further analyses of the topology of AguR and its ability to directly interact with agmatine are needed to reach a firm conclusion. Notably, we have attempted to produce a full-length AguR protein to explore the latter possibility but have not been able to express the N-terminal two-thirds of the protein in E. coli. It is also noteworthy that AguR is produced in S. mutans at extremely low levels, so an analysis of the binding characteristics of AguR will require a breakthrough in expression of this protein in sufficient quantities to conduct the necessary biochemical studies.

A recent report suggested that the regulation of cadAB gene expression by CadC may be even more complex than previously appreciated. In particular, site-specific proteolysis of the periplasmic domain of CadC, which is generated in cells exposed to acidic conditions, causes the release of a biologically active cytoplasmic form of the N-terminal DNA binding domain of CadC that can stimulate target gene activation (18). It is clear from our Western blots that a smaller species of AguR is commonly observed in strains expressing full-length wild-type or mutated AguR proteins. The observation that this lower-molecular-weight protein is indeed a derivative of AguR is supported by the fact that the protein is absent in strains lacking AguR and that all Western blotting was done with an antibody that was raised against a highly purified recombinant AguR protein that was expressed in E. coli. Furthermore, the antibodies used in the Western blots were affinity purified using the purified recombinant AguR protein. There is one open reading frame (SMu0441) in S. mutans with limited homology to the C-terminal DNA binding protein of AguR, but inactivation of the gene for that protein did not alter the results of Western blotting (data not shown). We are in the process of determining where the potential cleavage of AguR occurs and whether the processing is stimulated by growth under particular conditions, e.g., low pH.

An unexpected finding that arose from this study was that aguB promoter activity in the presence of inducing concentrations of agmatine was altered in strains carrying only the vector pMSP3535. Furthermore, deletion of the nisRK two-component system (TCS) from the vector eliminated this effect on agu expression (Fig. 2). NisRK is a TCS that senses environmental nisin and activates nisA promoter expression by using a classical TCS phosphorelay circuit (15). Notably, we have seen an effect of NisRK on one other unrelated gene in S. mutans (L. Zeng and R. Burne, unpublished data). While this observation may serve primarily as a cautionary note on the use of pMSP3535 for certain gene regulation studies, the finding may have some importance in terms of agu gene expression. In particular, the regulation of certain genes by pH, as well as the general property of acid tolerance in S. mutans, has been shown to be influenced in strains lacking a number of different TCS constituents, including CovR, CiaRH, VicRK, and ComDE (2, 22, 24, 27). The influence of heterologous expression of a TCS in S. mutans, in this case NisRK, that may potentially cross talk with endogenous TCSs raises the possibility that pH-dependent expression of the agu operon could be, at least in part, controlled by a TCS. Efforts are ongoing to probe in more detail the basis for low-pH-dependent enhancement of agu gene expression.

Elucidation of the genetics, physiology, and biochemistry of agmatine catabolism is an important first step in understanding the role of the agmatine deiminase in oral biofilm ecology and disease. The AgDS conveys bioenergetic advantages to S. mutans and other oral streptococci through enhancement of the pH and through generation of ATP (12), so it should contribute in major ways to the persistence and virulence of these organisms. Moreover, the complexity of regulation of the AgDS by substrate, catabolite control, and relevant environmental stresses, including pH, along with the relatively broad distribution of the system in pathogens and commensals, makes this an excellent model for the study of gene regulation and physiology in streptococci.

 
We thank Ann R. Griswold for providing an AguD-deficient strain of S. mutans.

This work was supported by Public Health Service grant DE10362 from the National Institute of Dental and Craniofacial Research.

 
* Corresponding author. Mailing address: Department of Oral Biology, University of Florida, P.O. Box 100424, Gainesville, FL 32610-0424. Phone: (352) 392-4370. Fax: (352) 392-7357. E-mail: rburne{at}dental.ufl.edu

Published ahead of print on 6 March 2009.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Ahn, S. J., J. A. Lemos, and R. A. Burne. 2005. Role of HtrA in growth and competence of Streptococcus mutans UA159. J. Bacteriol. 187:3028-3038.[Abstract/Free Full Text]
2
2. Ahn, S. J., Z. T. Wen, and R. A. Burne. 2006. Multilevel control of competence development and stress tolerance in Streptococcus mutans UA159. Infect. Immun. 74:1631-1642.[Abstract/Free Full Text]
3
3. Bender, G. R., S. V. Sutton, and R. E. Marquis. 1986. Acid tolerance, proton permeabilities, and membrane ATPases of oral streptococci. Infect. Immun. 53:331-338.[Abstract/Free Full Text]
4
4. Bryan, E. M., T. Bae, M. Kleerebezem, and G. M. Dunny. 2000. Improved vectors for nisin-controlled expression in gram-positive bacteria. Plasmid 44:183-190.[CrossRef][Medline]
5
5. Burne, R. A., Z. T. Wen, Y. Y. Chen, and J. E. Penders. 1999. Regulation of expression of the fructan hydrolase gene of Streptococcus mutans GS-5 by induction and carbon catabolite repression. J. Bacteriol. 181:2863-2871.[Abstract/Free Full Text]
6
6. Chen, Y. Y., M. J. Betzenhauser, and R. A. Burne. 2002. cis-Acting elements that regulate the low-pH-inducible urease operon of Streptococcus salivarius. Microbiology 148:3599-3608.[Abstract/Free Full Text]
7
7. Chen, Y. Y., and R. A. Burne. 1996. Analysis of Streptococcus salivarius urease expression using continuous chemostat culture. FEMS Microbiol. Lett. 135:223-229.[CrossRef][Medline]
8
8. Chen, Y. Y., C. A. Weaver, D. R. Mendelsohn, and R. A. Burne. 1998. Transcriptional regulation of the Streptococcus salivarius 57.I urease operon. J. Bacteriol. 180:5769-5775.[Abstract/Free Full Text]
9
9. Eichenbaum, Z., M. J. Federle, D. Marra, W. M. de Vos, O. P. Kuipers, M. Kleerebezem, and J. R. Scott. 1998. Use of the lactococcal nisA promoter to regulate gene expression in gram-positive bacteria: comparison of induction level and promoter strength. Appl. Environ. Microbiol. 64:2763-2769.[Abstract/Free Full Text]
10
10. Fuqua, C., M. R. Parsek, and E. P. Greenberg. 2001. Regulation of gene expression by cell-to-cell communication: acyl-homoserine lactone quorum sensing. Annu. Rev. Genet. 35:439-468.[CrossRef][Medline]
11
11. Griswold, A. R., Y. Y. Chen, and R. A. Burne. 2004. Analysis of an agmatine deiminase gene cluster in Streptococcus mutans UA159. J. Bacteriol. 186:1902-1904.[Abstract/Free Full Text]
12
12. Griswold, A. R., M. Jameson-Lee, and R. A. Burne. 2006. Regulation and physiologic significance of the agmatine deiminase system of Streptococcus mutans UA159. J. Bacteriol. 188:834-841.[Abstract/Free Full Text]
13
13. Griswold, A. R., M. N. Nascimento, and R. A. Burne. 2009. Distribution, regulation and role of the agmatine deiminase system in mutans streptococci. Oral Microbiol. Immunol. 24:79-82.[CrossRef][Medline]
14
14. Kleerebezem, M., M. M. Beerthuyzen, E. E. Vaughan, W. M. de Vos, and O. P. Kuipers. 1997. Controlled gene expression systems for lactic acid bacteria: transferable nisin-inducible expression cassettes for Lactococcus, Leuconostoc, and Lactobacillus spp. Appl. Environ. Microbiol. 63:4581-4584.[Abstract]
15
15. Kuipers, O. P., M. M. Beerthuyzen, P. G. de Ruyter, E. J. Luesink, and W. M. de Vos. 1995. Autoregulation of nisin biosynthesis in Lactococcus lactis by signal transduction. J. Biol. Chem. 270:27299-27304.[Abstract/Free Full Text]
16
16. Kuper, C., and K. Jung. 2005. CadC-mediated activation of the cadBA promoter in Escherichia coli. J. Mol. Microbiol. Biotechnol. 10:26-39.[CrossRef][Medline]
17
17. Kuramitsu, H. K. 1993. Virulence factors of mutans streptococci: role of molecular genetics. Crit. Rev. Oral Biol. Med. 4:159-176.[Abstract/Free Full Text]
18
18. Lee, Y. H., J. H. Kim, I. S. Bang, and Y. K. Park. 2008. The membrane-bound transcriptional regulator CadC is activated by proteolytic cleavage in response to acid stress. J. Bacteriol. 190:5120-5126.[Abstract/Free Full Text]
19
19. Lemos, J. A., J. Abranches, and R. A. Burne. 2005. Responses of cariogenic streptococci to environmental stresses. Curr. Issues Mol. Biol. 7:95-107.[Medline]
20
20. Lemos, J. A., T. A. Brown, Jr., and R. A. Burne. 2004. Effects of RelA on key virulence properties of planktonic and biofilm populations of Streptococcus mutans. Infect. Immun. 72:1431-1440.[Abstract/Free Full Text]
21
21. Lemos, J. A., and R. A. Burne. 2002. Regulation and physiological significance of ClpC and ClpP in Streptococcus mutans. J. Bacteriol. 184:6357-6366.[Abstract/Free Full Text]
22
22. Levesque, C. M., R. W. Mair, J. A. Perry, P. C. Lau, Y. H. Li, and D. G. Cvitkovitch. 2007. Systemic inactivation and phenotypic characterization of two-component systems in expression of Streptococcus mutans virulence properties. Lett. Appl. Microbiol. 45:398-404.[CrossRef][Medline]
23
23. Levin, P. 2002. Light microscopy techniques for bacterial cell biology. Methods Microbiol. 31:115-132.[CrossRef]
24
24. Li, Y. H., P. C. Lau, N. Tang, G. Svensater, R. P. Ellen, and D. G. Cvitkovitch. 2002. Novel two-component regulatory system involved in biofilm formation and acid resistance in Streptococcus mutans. J. Bacteriol. 184:6333-6342.[Abstract/Free Full Text]
25
25. Neely, M. N., C. L. Dell, and E. R. Olson. 1994. Roles of LysP and CadC in mediating the lysine requirement for acid induction of the Escherichia coli cad operon. J. Bacteriol. 176:3278-3285.[Abstract/Free Full Text]
26
26. Park, Y. K., B. Bearson, S. H. Bang, I. S. Bang, and J. W. Foster. 1996. Internal pH crisis, lysine decarboxylase and the acid tolerance response of Salmonella typhimurium. Mol. Microbiol. 20:605-611.[CrossRef][Medline]
27
27. Senadheera, M. D., B. Guggenheim, G. A. Spatafora, Y. C. Huang, J. Choi, D. C. Hung, J. S. Treglown, S. D. Goodman, R. P. Ellen, and D. G. Cvitkovitch. 2005. A VicRK signal transduction system in Streptococcus mutans affects gtfBCD, gbpB, and ftf expression, biofilm formation, and genetic competence development. J. Bacteriol. 187:4064-4076.[Abstract/Free Full Text]
28
28. Slock, J., D. VanRiet, D. Kolibachuk, and E. P. Greenberg. 1990. Critical regions of the Vibrio fischeri LuxR protein defined by mutational analysis. J. Bacteriol. 172:3974-3979.[Abstract/Free Full Text]
29
29. Wexler, D. L., M. C. Hudson, and R. A. Burne. 1993. Streptococcus mutans fructosyltransferase (ftf) and glucosyltransferase (gtfBC) operon fusion strains in continuous culture. Infect. Immun. 61:1259-1267.[Abstract/Free Full Text]
30
30. Zeng, L., Z. T. Wen, and R. A. Burne. 2006. A novel signal transduction system and feedback loop regulate fructan hydrolase gene expression in Streptococcus mutans. Mol. Microbiol. 62:187-200.[CrossRef][Medline]
31
31. Zubay, G., D. E. Morse, W. J. Schrenk, and J. H. Miller. 1972. Detection and isolation of the repressor protein for the tryptophan operon of Escherichia coli. Proc. Natl. Acad. Sci. USA 69:1100-1103.[Abstract/Free Full Text]

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Applied and Environmental Microbiology, May 2009, p. 2629-2637, Vol. 75, No. 9
0099-2240/09/$08.00+0     doi:10.1128/AEM.02145-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

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• Liu, Y., Burne, R. A. (2009). Multiple Two-Component Systems Modulate Alkali Generation in Streptococcus gordonii in Response to Environmental Stresses. J. Bacteriol. 191: 7353-7362 [Abstract] [Full Text]  

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