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J Bacteriol. 2006 February; 188(3): 834–841.
PMCID: PMC1347362

Regulation and Physiologic Significance of the Agmatine Deiminase System of Streptococcus mutans UA159


We previously demonstrated that Streptococcus mutans expresses a functional agmatine deiminase system (AgDS) encoded by the agmatine-inducible aguBDAC operon (A. R. Griswold, Y. Y. Chen, and R. A. Burne, J. Bacteriol. 186:1902-1904, 2004). The AgDS yields ammonia, CO2, and ATP while converting agmatine to putrescine and is proposed to augment the acid resistance properties and pathogenic potential of S. mutans. To initiate a study of agu gene regulation, the aguB transcription initiation site was identified by primer extension and a putative σ70-like promoter was mapped 5′ to aguB. Analysis of the genome database revealed an open reading frame (SMU.261c) encoding a putative transcriptional regulator located 239 bases upstream of aguB. Inactivation of SMU.261c decreased AgD activity by sevenfold and eliminated agmatine induction. AgD was also found to be induced by certain environmental stresses, including low pH and heat, implying that the AgDS may also be a part of a general stress response pathway of this organism. Interestingly, an AgDS-deficient strain was unable to grow in the presence of 20 mM agmatine, suggesting that the AgDS converts a growth-inhibitory substance into products that can enhance acid tolerance and contribute to the competitive fitness of the organism at low pH. The capacity to detoxify and catabolize agmatine is likely to have major ramifications on oral biofilm ecology.

The virulence of Streptococcus mutans is attributable to constitutive and inducible acid tolerance properties that allow continued growth and glycolysis at low pH values in oral biofilms. Repeated acidification of oral biofilms is not conducive to persistence of a flora associated with dental health and can result in significant tooth demineralization. A primary determinant of acid tolerance in S. mutans is the membrane-bound F1F0-ATPase, but reduction in proton permeability of the cell membrane and induction of DNA repair pathways and stress proteins also contribute in major ways to aciduricity (31). A common theme for acid adaptation in bacteria is the production of basic compounds to neutralize the cytoplasm and surroundings of the organisms (11). While S. mutans has multiple strategies to cope with low pH and gains a competitive advantage over acid-sensitive bacteria at low pH, it has not been considered capable of generating significant quantities of alkali (28). Recently, we described a pathway for ammonia production in S. mutans that had not yet been identified in oral bacteria (14). Alkali production by the agmatine deiminase system (AgDS) may increase the competitive fitness of S. mutans, contributing in major ways to the persistence and pathogenesis of this organism.

Agmatine is a decarboxylated derivative of arginine that can be catabolized by the AgDS (37). The S. mutans AgDS is encoded by the aguBDAC operon (Fig. (Fig.1)1) (14). Agmatine can enter the cell via an agmatine-putrescine antiporter, encoded by aguD, where it is hydrolyzed to N-carbamoylputrescine and ammonia by agmatine deiminase (AgD; EC, encoded by aguA. Putrescine carbamoyltransferase (EC, 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, encoded by aguC, generating ATP, CO2, and NH3. Putrescine is then exchanged for agmatine via the antiporter.

FIG. 1.
Genetic organization of the agu operon in S. mutans UA159. SMU.261c, annotated as a putative transcriptional regulator, has been designated aguR in this study to reflect its role as an activator of the agu genes. PTC, putrescine carbamoyltransferase; ...

Agmatine can be produced by a broad range of organisms, including maize, rice, soybeans, cucumbers, Enterococcus faecalis, Pseudomonas aeruginosa, and Lactobacillus hilgardii (2, 26, 36, 37, 42). NCBI BLAST searches have revealed potential AgD enzymes in Lactobacillus sakei, Lactococcus lactis, Listeria monocytogenes, Yersinia pestis, Streptococcus pneumoniae, Streptomyces spp., Bacillus cereus, and Helicobacter pylori, although functionality of these apparent agu homologues has not been established (14). The role of agmatine catabolism in bacteria appears to vary depending upon the source of the agmatine. In organisms that possess an intracellular arginine decarboxylase (ADC), such as P. aeruginosa, agmatine is produced directly from arginine and is converted to N-carbamoylputrescine by AgD (26). N-Carbamoylputrescine is then hydrolyzed to putrescine, CO2, and NH3 by N-carbamoylputrescine amidohydrolase (26). Putrescine is converted to spermidine by spermidine synthase or broken down to succinate, which enters the tricarboxylic acid cycle. The AgDS in these bacteria may be an important source of polyamines, in addition to providing carbon and nitrogen.

In ADC-deficient bacteria, such as S. mutans and E. faecalis, agmatine is derived from exogenous sources via an agmatine-putrescine antiporter (10, 14). In S. mutans and E. faecalis, the AgDS closely resembles the arginine deiminase system (ADS), a pathway that generates ammonia, CO2, and ATP from arginine (4, 6, 13, 23, 33, 37, 39). The ADS is considered a primary mechanism of acid tolerance used by some oral bacteria to survive the frequent cycles of acidification encountered in dental plaque, but it is not present in the genome of S. mutans (9). Biochemical analyses in E. faecalis (33) showed that the AgDS is highly analogous to the ADS, suggesting a role in acid tolerance, but the genes encoding this pathway or their mode of regulation have not been characterized.

Previously, we demonstrated that the agu operon in S. mutans is induced in the presence of agmatine and is regulated by carbon catabolite repression (CCR) (14). Overall, the AgDS is expressed at a relatively low level compared to other ammonia-generating pathways of oral streptococci, and it is unlikely that agmatine catabolism results in significant environmental alkalinization. However, ammonia production by the AgDS under acidic conditions would increase ΔpH and provide ATP, thereby contributing to acid tolerance and growth at low pH, which would substantively augment the virulence of the organism. In order to better understand the role of the AgDS in the physiology and virulence of S. mutans, we have initiated a study of the factors regulating the AgDS in this organism. We have also described a novel mechanism by which S. mutans copes with the production of an antagonistic compound generated by competing organisms in response to environmental acidification of oral biofilms.


Bacterial strains, growth conditions, and reagents.

S. mutans UA159 was grown in brain heart infusion broth (BHI; Difco Laboratories, Detroit, MI) at 37°C in 5% CO2 and 95% air. To monitor AgDS expression, batch cultures of S. mutans were grown in a low-carbohydrate tryptone-vitamin (TV) medium (7) containing 25 mM glucose or galactose, with or without 10 mM agmatine, to an optical density at 600 nm (OD600) of 0.5. Recombinant Escherichia coli DH10B strains were maintained on L agar supplemented, where indicated, with 100 μg ml−1 kanamycin (Km) or 300 μg ml−1 erythromycin (Erm). Chemical reagents were obtained from Sigma (St. Louis, MO). Growth curves of S. mutans UA159 and mutant strains were generated using a Bioscreen C (Oy Growth Curves AB Ltd., Helsinki, Finland). Optical density at 600 nm was recorded every 30 min, with shaking for 10 s before each reading.

For studies on pH-dependent regulation of AgDS expression, steady-state continuous cultures of S. mutans were grown in a Biostat i Twin Controller chemostat (B. Braun Biotech, Inc., Allentown, PA) in a tryptone-yeast extract (TY) medium (41) supplemented with 25 mM glucose at a dilution rate (D) of 0.3 h−1. Where indicated, cultures were pulsed with 3 mM agmatine for 1 h prior to sampling. Cultures were maintained at pH 5 or pH 7 by the addition of 2 M KOH.

DNA manipulation and construction of mutant strains.

Genomic DNA was isolated from S. mutans UA159 as previously described (8). Plasmid DNA used in sequencing reactions was prepared from Escherichia coli DH10B by the method of Birnboim and Doly (5). Cloning and electrophoretic analysis of DNA fragments were carried out according to established protocols (3). Southern hybridization and high-stringency washes were performed as previously described (19). Restriction and DNA-modifying enzymes were purchased from Life Technologies Inc. (Rockville, MD) or New England Biolabs (Beverly, MA).

Recombinant PCR was used to construct a nonpolar aguC mutant (aguCΔKan). The 5′ half of aguC was amplified using primer pairs aguCSPstI (5′-GTCTAGAGACTGCAGTGCCAAAGCACA-3′) and aguCASSmaI (5′-TTTTCGCCAACCTCTCCCGGGATCTACTTT-3′), which inserted PstI and SmaI restriction sites (boldface) to facilitate cloning. The remaining portion of aguC was amplified using primer pairs aguCSSmaI (5′-AAAGTAGATCCCGGGAGAGGTTGGCGAAAA-3′) and aguCASSstI (5′-GGCTTTTCCACTGAGCTCTGCTTCAAC-3′), which inserted SmaI and SstI restriction sites. The two primary PCR products were then used in a reaction with primers aguCSPstI and aguCASSstI to generate a secondary PCR product corresponding to the entire length of aguC. The PCR fragment was then cloned onto pGEM5zf(+) and electroporated into E. coli DH10B. A promoterless kanamycin (Km) resistance cassette, from Tn1545 and lacking a terminator (29), was inserted at the SmaI restriction site to disrupt aguC. The construct was integrated into the S. mutans chromosome via natural transformation (30) selecting for growth on BHI agar containing 1.0 mg ml−1 kanamycin, and correct integration was confirmed by Southern blotting. The polar aguB mutant (aguBΔΩKan) was constructed in a previous study via insertion of ΩKm (14).

The aguR deletion mutant (aguRΔErm) was constructed by PCR ligation mutagenesis (18). Primers aguRS (5′-CGTTCTTTTCCTGCAGGACTCTCAAG-3′) and aguRASHindIII (5′-CGTAAATTGAAGCTTTTCCTAAACTGAC-3′) were used to amplify a 600-bp region upstream of aguR. Primers aguRSSstI (5′-CTCCTTTAATTTGAGCTCAATATCTATAGT-3′) and aguRAS (5′-GATATCATCCAATCTAGAAAGAACAGTTG-3′) were used to amplify a 600-bp region downstream of aguR. The PCR products were digested with HindIII and SstI, respectively, and ligated to the erythromycin (Em) resistance cassette derived from Tn916 delta E (34). The ligation mixture was introduced into S. mutans UA159 by natural transformation, and bacteria were plated on BHI agar containing 8 μg ml−1 erythromycin. Double-crossover mutants were confirmed by Southern blotting and PCR.

RNA extraction and analyses.

RNA was prepared from batch or chemostat cultures using methods described elsewhere (8) and immediately treated with the RNAprotect reagent from QIAGEN (QIAGEN Inc., Valencia, CA). The RNA was further purified and treated with DNase I, using the RNeasy RNA Clean Up mini kit from QIAGEN, and stored at −80°C.

Primer extension analysis was used to map the aguB and aguR transcription initiation sites. Primer AguBAS (5′-TCCTCTGTCGTAATATAATCTGT-3′) encoded the antisense sequence of aguB located 30 bases downstream from the translational start site. Primer AguRAS (5′-ATAGATTATAGATATAGATGAGTTC-3′) encoded the antisense sequence of aguR located 25 bases downstream from the translational start site. Incubation of radiolabeled primers with 50 μg of total RNA at 42°C for 90 min was followed by reverse transcription, and the products were separated by electrophoresis and disclosed by autoradiography. DNA sequencing reactions using the same primers were included on the gel to allow identification of the start sites.

Real-time reverse transcriptase PCR (RT-PCR) was used to monitor expression of aguA in response to growth at pH 5 versus pH 7. cDNA was generated from 1 μg of total RNA using an aguA-specific primer as recommended by the supplier (SuperScript First-Strand Synthesis System for RT-PCR; Invitrogen, Carlsbad, CA). The aguA-specific primers (forward, 5′-ATGCTTGGATTCGTGACTGTGG-3′; reverse, 5′-AAGACCATCGACTAAGCCTCCC-3′) were designed using Beacon Designer 2.0 software (Premier Biosoft International, Palo Alto, CA). Standard curves for each gene, prepared as described by Yin et al. (43), were used in every run. A range of 101 to 108 copies was found to be adequate for all genes examined.

Agmatine deiminase assays.

AgD activity was measured by colorimetric determination of N-carbamoylputrescine production from agmatine (1) as previously described (14). Results were expressed as nmol N-carbamoylputrescine min−1 mg of protein−1.


Role of the putative S. mutans regulatory protein SMU.261c.

Previously, we reported that efficient expression of the S. mutans AgDS requires agmatine (14). To date, the only reported trans-acting factor involved in AgDS regulation is AguR, a transcriptional repressor encoded by an open reading frame (ORF) located immediately upstream of the P. aeruginosa aguBA operon. In P. aeruginosa, the presence of 1 mM agmatine significantly inhibits AguR-DNA interaction, facilitating induction of aguBA in the presence of agmatine (26). Although there were no ORFs that showed substantial similarity to P. aeruginosa AguR in the S. mutans genome, a putative LuxR-like transcriptional regulator, SMU.261c (NCBI database), was identified 239 bases upstream of aguB and transcribed in the opposite direction. LuxR-type proteins belong to the FixJ-NarL superfamily, which is mainly comprised of two-component response regulators involved in quorum sensing (12). Using NCBI Blastp, it was determined that SMU.261c shared the highest levels of similarity with putative LuxR-like regulators that are encoded approximately 399 and 234 bases upstream of the putative agu operons of E. faecalis and L. lactis subsp. lactis (62% and 56% similarity, respectively). A C-terminal helix-turn-helix domain, implicated in DNA binding by members of the LuxR family, was identified in SMU.261c, as well as in the E. faecalis and L. lactis ORFs. However, the conserved acylated homoserine lactone binding region, typical of LuxR, could not be found in any of the three proteins. Interestingly, three transmembrane domains were also predicted to occur in the protein encoded by SMU.261c. Consequently, it is possible to predict a topology for the protein that could result in exposure of a substantial portion of the molecule to the external environment.

The SMU.261c gene was mutated by allelic exchange with the insertion of an Ermr marker (aguRΔErm), using the PCR ligation mutagenesis method described by Lau et al. (18). When increasing concentrations of agmatine were added to the growth medium, AgD activity increased proportionately in UA159, whereas aguRΔErm displayed a low basal level of AgD activity, regardless of agmatine concentration (Fig. (Fig.2).2). Thus, SMU.261c was named aguR to reflect a role in agmatine induction of the AgDS.

FIG. 2.
AgD enzyme activity in S. mutans UA159 and aguRΔErm, in response to increasing concentrations (mM) of agmatine. Results shown are the average and standard deviations (error bars) of a minimum of nine separate cultures for each strain and condition. ...

Localization of paguB and paguR.

Primer extension was used to map the aguB and aguR transcription initiation sites (TIS) (Fig. (Fig.3).3). A single band corresponding to a G residue 22 bases upstream of the aguB start codon was observed. Examination of the upstream sequence revealed a putative σ70-like promoter. The −10 region (TATGAT) shared 5 out of 6 bases with the consensus sequence (bold), whereas the −35 region (TAGTAA) identified 18 bases upstream of the −10 region shared only 3 bases with the consensus (bold). A single band corresponding to an A residue 34 bases upstream of the aguR start codon was observed. Examination of the upstream sequence revealed a putative σ70-type promoter. The −10 region (TATAAT) was identical to the consensus sequence (bold), whereas the −35 region (TTCAAT) identified 18 bases upstream of the −10 region shared only 3 bases with the consensus (bold).

FIG. 3.
The aguRB intergenic sequence and primer extension analysis of S. mutans aguB and aguR. Arrowheads indicate initiation sites of aguB and aguR transcription at G and A residues, respectively. The transcriptional start sites are marked with round arrowheads ...


We have demonstrated that the S. mutans agu genes are transcriptionally repressed in the presence of the preferred carbohydrate source, glucose (14). In AT-rich gram-positive bacteria, CCR is mediated by the trans-acting catabolite control protein A (CcpA), which binds to highly conserved, cis-acting catabolite response elements (cre) during growth on preferred carbohydrate sources to regulate the expression of catabolic genes and operons (35). Two potential CcpA-dependent cre sites were identified at −48 (TGTAATCGTTTACA) and −137 (TGAAAAGGCTTTGT) relative to the aguB transcriptional start site, with bases matching the consensus shown in bold (Fig. (Fig.3)3) (16).

The identification of putative cre sites upstream of paguB, as well as the AgDS regulation patterns observed previously, prompted us to investigate the role of CcpA-like proteins (RegM) (38) and CcpB-like proteins (SMU.105; RegA) (40) of S. mutans in AgDS regulation. AgD activity was measured in S. mutans UA159 and in otherwise isogenic mutants of this strain lacking the ccpA or ccpB genes that were constructed previously in our laboratory (40). The strains were grown in TV broth containing 25 mM glucose or the nonrepressing sugar, galactose, with or without 10 mM agmatine, to mid-exponential phase prior to measurement of enzyme activity. Mutation of ccpA or ccpB did not alleviate catabolite repression of the AgDS (Fig. (Fig.4),4), although AgD activity was slightly higher in the ccpA and ccpB strains following growth in either glucose or galactose, supplemented with agmatine. This observation is consistent with studies of other metabolic pathways in S. mutans that showed that even though cre sequences were tightly linked to the regulatory regions, CcpA (RegM) and CcpB are not primary factors controlling CCR in this organism (40).

FIG. 4.
AgD enzyme activity in S. mutans UA159 and in otherwise isogenic ccpA and ccpB strains in response to growth in the presence of the catabolite-repressing sugar, glucose, or the nonrepressing sugar, galactose. Ag, agmatine. Results shown are the average ...

Regulation of the AgDS by growth phase and environmental stress.

Identification of environmental factors that regulate expression of bacterial genes can provide much insight into the role of the gene products. Thus, we began our investigation of the environmental factors regulating the S. mutans AgDS by examining AgD activity in relation to growth phase. Batch cultures of S. mutans UA159 were grown in TV medium containing 25 mM galactose, with or without 10 mM agmatine, and samples were taken during early exponential phase, mid-exponential phase, and 2 h after the cultures entered stationary phase (OD600 = 0.35, 0.5, and 0.65, respectively). The pH of each sample was measured, and AgD assays were performed. AgD activity was 1.5-fold higher in the stationary phase compared with the mid-exponential phase when agmatine was included in the growth media (Fig. (Fig.5A).5A). Notably, the pH of stationary-phase cultures of S. mutans can be considerably lower than that of cells in the exponential phase, so the post-exponential-phase enhancement of AgD expression could be due to nutrient depletion or to the reduced pH of the growth medium. When batch cultures were grown in TV medium buffered at pH 7, stationary-phase AgD induction of approximately 1.5-fold was still observed (Fig. (Fig.5B),5B), but enzyme activity levels in all cases were lower than those in unbuffered medium. Thus, some induction of the AgDS occurs during stationary phase, probably due to partial alleviation of CCR or another pathway that senses nutrient depletion. However, AgDS induction is clearly enhanced at low pH values (Fig. 5C and D). It is also noteworthy that we previously reported that ammonia production from agmatine by intact cells of S. mutans is optimal at pH 4 (14).

FIG. 5.FIG. 5.FIG. 5.FIG. 5.FIG. 5.
AgD enzyme activity in relation to growth domain (A), in mid-exponential-phase or stationary-phase cultures grown in pH 7-buffered medium (B), in continuous cultures maintained at pH 5 or pH 7 (C), real-time RT-PCR of aguA expression in continuous cultures ...

To examine the effects of low pH on AgD activity under conditions where growth phase, growth rate, and glucose availability could be tightly controlled, steady-state continuous cultures of S. mutans were maintained in a Biostat i chemostat at pH 5 or 7 as detailed in Materials and Methods. AgD activity was up-regulated fourfold at pH 5 compared to pH 7 in agmatine-induced cultures (Fig. (Fig.5C).5C). Of note, AgD activity was significantly lower in chemostat cultures than in batch cultures. This discrepancy was most likely caused by differences in the methods of AgD induction between the two experiments. Specifically, in the chemostat studies, the vessel was pulsed with a final concentration of 3 mM agmatine for 1 h prior to sampling, whereas batch cultures were grown in medium containing 10 mM agmatine. This was done because the presence of an ammonia-yielding compound during the fermentation makes it difficult to maintain acidic pH in the vessel, and it is prohibitively expensive to include agmatine in the continuous cultures. Also of interest is that AgD activity was observed in continuous cultures in the absence of agmatine, whereas the substrate was required for AgD expression in batch cultures. This observation may be attributable to alleviation of residual CCR under glucose-limiting conditions in the chemostat. Also, the slower growth rate in the chemostat (tg = 2.3 h) may also contribute to derepression of the AgDS, which could in part explain the induction of the AgDS during stationary phase.

Consistent with measurements of AgD enzyme activity, the expression of aguA, as measured by real-time RT-PCR, was approximately 3.7-fold higher in cells grown at pH 5 than that in cells grown at pH 7 (Fig. (Fig.5D).5D). Thus, transcription of the aguBDAC operon is activated at acidic pH levels frequently encountered in dental plaque. Induction of AgDS at low pH supports the notion that the system may be a component of the adaptive acid tolerance response by S. mutans. Also of note, many bacteria induce arginine decarboxylase expression when exposed to acid stress (15, 22). Consequently, agmatine would likely be in greater abundance at low pH in vivo, and the combination of low pH and agmatine could result in optimal AgD expression, similar to what has been observed for lysine decarboxylase and arginine decarboxylase in E. coli (24, 32).

To determine if the AgDS could be part of a general stress response pathway in S. mutans, the effects of heat, oxidative, and salt stresses were examined. AgD activity was fourfold higher in S. mutans grown at 42°C than in S. mutans grown at 37°C (Fig. (Fig.5E),5E), implying that this system is responsive to environmental stresses other than low pH. However, oxidative or salt stress had no effect on AgD activity (data not shown). Consensus binding sites for the heat shock regulators HrcA and CtsR (20, 21) were not identified in the aguB promoter region. Whether heat stress acts through aguR-specific regulators or through other pathways remains to be determined.

AgDS and biofilm ecology.

Oral biofilms are complex ecosystems with hundreds of metabolically and physiologically diverse species. The ability of S. mutans to catabolize agmatine at low pH could impart a selective advantage to this organism through generation of ATP for growth and alkalinization of the cytoplasm by ammonia, which would reduce the investment of ATP in proton extrusion. Under such conditions, S. mutans would gain a competitive advantage over less-acid-tolerant species in oral biofilms, particularly since the system is a low-activity system that would not profoundly affect environmental alkalinization. To test this hypothesis, a competition experiment was performed using S. mutans UA159 and its AgDS-deficient derivative, aguBΔΩKan. Individual cultures were grown separately to an OD600 of 0.4 and combined in equal volumes in TV media containing galactose and 0 mM or 20 mM agmatine (data not shown). The strains grew equally well in the absence of agmatine. Surpisingly, in the presence of 20 mM agmatine, the doubling time of the wild-type strain was significantly slower and the aguBΔΩKan strain was unable to grow.

To further investigate the mechanism of growth inhibition by agmatine, the gene for carbamate kinase was mutated (aguCΔKan), which would allow the organisms to degrade agmatine, while they would be unable to produce ATP, carbon dioxide, and the second mole of ammonia from agmatine. The aguCΔKan strain was able to grow in the presence of agmatine, albeit not as rapidly as the wild type, suggesting that inhibition of growth by agmatine is the primary explanation for the phenotype displayed by the aguBΔΩKan strain (Fig. (Fig.6).6). Agmatine inhibition was dose dependent in the wild-type, aguBΔΩKan, and aguCΔKan strains (data not shown).

FIG. 6.
Growth of S. mutans UA159, aguBΔΩKan, aguCΔKan, and strains in TV medium containing 25 mM galactose and either 0 mM or 20 mM agmatine (Ag). WT, wild type. Optical density at 600 nm was determined every 30 min for 50 h using a Bioscreen ...

The basis for inhibition of growth by agmatine has yet to be disclosed, although it may involve competitive inhibition of amino acid transport or perhaps interference with translation. Inhibition by agmatine was less evident when the organisms were grown in peptide- and carbohydrate-rich media that favored overall higher growth rates, lending credence to the idea that agmatine may compete with the structurally related amino acid, arginine, either for uptake or for charging of tRNAs. Specifically, inhibition was alleviated when the wild-type and aguBΔΩKan strains were grown in a tryptone yeast-based medium or BHI, both of which are abundant in peptides and carbohydrates (Fig. 7A to D). We posit that agmatine may compete with amino acid transporters and that provision of an abundant amino acid source in peptide form allows the organisms to bypass competitive inhibition for transport. It is also noteworthy that agmatine inhibition was most severe when the strains were grown in TV medium supplemented with agmatine and the nonrepressing sugar, galactose (Fig. (Fig.6).6). However, the doubling time of the aguBΔΩKan strain was similar to that of UA159 when the cells were grown in TV medium containing agmatine and the repressing sugar, glucose (Fig. (Fig.7A).7A). In B. subtilis and P. aeruginosa, certain amino acid and peptide transporters are negatively regulated by CCR (25, 27). Repression of the transporters during growth in glucose could reduce agmatine uptake and allow growth of the aguBΔΩKan strain. Further experiments are under way to uncover the exact mechanism of agmatine inhibition.

FIG. 7.
Growth of S. mutans UA159 and aguBΔΩKan in the presence and absence of agmatine (Ag). (A) TV medium containing 25 mM glucose. (B) TY medium containing 25 mM galactose. (C) TY medium containing 25 mM glucose. (D) BHI medium. Each point ...

Importantly, the finding that agmatine inhibits the growth of S. mutans reveals a novel type of antagonistic interaction that may be displayed by competing oral biofilm organisms. Specifically, many bacteria respond to acidification by inducing ADC to produce agmatine, which is pumped out of the cell and alkalinizes the environment, enhancing the growth of acid-sensitive organisms (15, 17, 22). In secreting agmatine, the organisms are concomitantly excreting a compound that inhibits the growth of S. mutans, a primary generator of acids in oral biofilms. In response, S. mutans induces the AgDS, deriving ATP and ammonia from an inhibitory compound (Fig. (Fig.88).

FIG. 8.
Proposed role of the AgDS in virulence. As S. mutans lowers the pH of dental plaque, acid-sensitive bacteria in the oral biofilm induce arginine decarboxylase to produce agmatine, which inhibits the growth of S. mutans. At low pH and in the presence of ...

BLAST searches were performed to find oral bacteria that carry ADC genes. The results demonstrated that Wolinella, Prevotella, Bacteroides, Desulfovibrio, and Neisseria species possess the ADC enzyme, while we were unable to generate evidence of the ADC enzyme in Porphyromonas gingivalis, oral streptococci, Actinobacillus actinomycetemcomitans, Actinomyces naeslundii, Fusobacterium nucleatum, Treponema denticola, and Lactobacillus species, albeit not all of these genomes are complete. Preliminary experiments employing high-performance liquid chromatography on derivatized pooled human oral samples obtained from healthy laboratory volunteers revealed approximately 750 nmol of agmatine mg of protein−1 from pooled human plaque and approximately 200 nmol of agmatine mg of protein−1 from pooled human saliva. Collectively, these results demonstrate that agmatine is present in physiologically significant amounts in dental plaque and tentatively identify species that could contribute to the total agmatine pools. These observations support our hypothesis that the AgDS may be involved in agmatine detoxification by S. mutans while concomitantly generating ATP and ammonia. Additional studies to explore agmatine production and utilization by oral microrganisms and biofilm communities are under way.


In summary, the AgDS of S. mutans is subject to complex regulation by substrate, catabolite control, and relevant environmental stresses. AguR is a major regulator of AgDS gene induction in response to agmatine, and perhaps pH, but it is likely that other global regulatory factors orchestrate differential expression of the system in response to CCR and stress. The physiological role of the AgDS in S. mutans is similarly complex, conveying bioenergetic advantages through enhancement of ΔpH and generation of ATP, as well as detoxifying agmatine produced by acid-sensitive organisms in response to acidification by S. mutans. Agmatine catabolism may thereby increase the competitive fitness of S. mutans, contributing in major ways to the persistence and virulence of this organism.


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


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