Stress-responsive proteins are upregulated in Streptococcus mutans during acid tolerance

Alice C. L. Len, Derek W. S. Harty and Nicholas A. Jacques

Institute of Dental Research, Westmead Millennium Institute and Westmead Centre for Oral Health, PO BOX 533, Wentworthville, NSW 2145, Australia

Correspondence
Nicholas A. Jacques
njacques{at}dental.wsahs.nsw.gov.au


   ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Streptococcus mutans is an important pathogen in the initiation of dental caries as the bacterium remains metabolically active when the environment becomes acidic. The mechanisms underlying this ability to survive and proliferate at low pH remain an area of intense investigation. Differential two-dimensional electrophoretic proteome analysis of S. mutans grown at steady state in continuous culture at pH 7·0 or pH 5·0 enabled the resolution of 199 cellular and extracellular protein spots with altered levels of expression. Matrix-assisted laser desorption ionization time-of-flight mass spectrometry identified 167 of these protein spots. Sixty-one were associated with stress-responsive pathways involved in DNA replication, transcription, translation, protein folding and proteolysis. The 61 protein spots represented isoforms or cleavage products of 30 different proteins, of which 25 were either upregulated or uniquely expressed during acid-tolerant growth at pH 5·0. Among the unique and upregulated proteins were five that have not been previously identified as being associated with acid tolerance in S. mutans and/or which have not been studied in any detail in oral streptococci. These were the single-stranded DNA-binding protein, Ssb, the transcription elongation factor, GreA, the RNA exonuclease, polyribonucleotide nucleotidyltransferase (PnpA), and two proteinases, the ATP-binding subunit, ClpL, of the Clp family of proteinases and a proteinase encoded by the pep gene family with properties similar to the dipeptidase, PepD, of Lactobacillus helveticus. The identification of these and other differentially expressed proteins associated with an acid-tolerant-growth phenotype provides new information on targets for mutagenic studies that will allow the future assessment of their physiological significance in the survival and proliferation of S. mutans in low pH environments.


Abbreviations: 2-DGE, two-dimensional gel electrophoresis; ASB-14, amidosulfobetaine-14; D, dilution rate; DE, differential expression (values); IPG, immobilized pH gradient; MALDI-TOF, matrix-assisted laser desorption ionization time-of-flight; PMM, peptide mass mapping


   INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Streptococcus mutans is now well recognized as being associated with the initiation of dental caries, since its acid fermentation by-products can result in the demineralization of tooth enamel (Hamada & Slade, 1980; Harper & Loesche, 1984; Loesche, 1986; van Houte, 1994; van Ruyven et al., 2000). A key to the survival of S. mutans at low pH is its ability to maintain a transmembrane pH gradient ({Delta}pH), with the interior of the cell more alkaline. This is achieved by upregulation of a proton-translocating F1F0-ATPase that extrudes H+ as the external environment becomes more acidic. This results in an increased use of ATP for H+extrusion and a consequent reduction in cell yield (Belli & Marquis, 1991; Hamilton & Buckley, 1991; Dashper & Reynolds, 1992; Quivey et al., 2001). A series of recent physiological, mutagenic and proteome studies (Quivey et al., 1995; Gutierrez et al., 1996, 1999; Jayaraman et al., 1997; Hamilton & Svensäter, 1998; Hahn et al., 1999; Hanna et al., 2001; Kremer et al., 2001; Lemos et al., 2001; Li et al., 2002; Wilkins et al., 2002; Len et al., 2004), however, indicates that S. mutans regulates its phenotype in a far more complex fashion than simply increasing its ability to extrude H+ in response to acid stress. For instance, our recent proteome analysis detected changes in metabolic pathways following acid-tolerant growth. Analysis of the data gave rise to the hypothesis that S. mutans redirects carbon from acidic fermentation by-products to more alkaline catabolites. These changes appear to occur in order to minimize the detrimental effects that result from the uncoupling of carbon flux from catabolism, as a consequence of the use of ATP for H+extrusion (Len et al., 2004). What remains to be elucidated is the breadth of the stress response in S. mutans that allows it to survive and proliferate at low pH.

Here we report the phenotypic changes, previously associated with the maintenance of bacterial viability under a variety of imposed environmental stresses, that were observed when S. mutans was grown at steady state in continuous culture at low pH. Proteins required for the maintenance of DNA integrity, transcriptional fidelity, translational efficiency, and protein folding were uniquely identified during acid-tolerant growth at pH 5·0, or were present at higher levels than those in S. mutans grown at pH 7·0. The mode of action of these proteins is discussed in relation to current knowledge of their roles in responding to stress, particularly in Gram-positive bacteria associated with acidic environments.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
Growth conditions.
Triplicate continuous cultures of Streptococcus mutans LT11 (Tao et al., 1993) were grown under anaerobic conditions at a dilution rate (D) of 0·100±0·001 h–1, at either pH 7·0±0·1 or pH 5·0±0·1, with glucose limitation as previously described (Jacques et al., 1979). DMM medium, devoid of mucin, was used, but modified to include adenine, guanine and uracil at 20 µg ml–1, and both KH2PO4 and K2HPO4 at 15 mM (Sissons et al., 1991).

Preparation of cellular and extracellular proteins.
When steady state had been achieved, the bacterial contents of the culture vessel were harvested, washed and lyophilized before aliquots of 10 mg dry wt of cells were treated with mutanolysin (Len et al., 2003). Proteins that were to be separated on acidic immobilized pH gradient (IPG) strips (pH 4·0–6·7) were extracted as previously described (Len et al., 2003), except that 1 % (w/v) amidosulfobetaine-14 (ASB-14) and 65 mM DTT were added to produce a modified solubilizing solution for two-dimensional electrophoresis (2-DGE). While the addition of these reagents increased the total number of protein spots that could be readily discerned on 2-DGE gels, their inclusion selectively inhibited the extraction or subsequent separation of a small number of weakly expressed proteins that had been previously visualized and/or identified on 2-DGE gels (Len et al., 2003).

Proteins that were to be separated on basic IPG strips (pH 6–11) were obtained from the mutanolysin-treated cells by a two-fraction solubilization procedure. Following centrifugation of the cell lysate (12 000 g, 4 °C, 10 min), the cell pellet was stored at –20 °C, and the proteins in the supernatant precipitated overnight at –20 °C with 15 % (w/v) trichloroacetic acid. After centrifugation and two washes in methanol (12 000 g, 4 °C, 10 min), the precipitated proteins were solubilized in 300 µl of a 1 : 1 mixture of modified solubilization solution (without ASB-14) and Cellular and Organelle Membrane Solubilizing Reagent (Sigma-Aldrich) containing 1 % (v/v) Triton X-100 and 2 mM tributylphosphine. The frozen cell pellet was then thawed and resuspended by sonication (Branson Ultrasonics; 50 W, 10x10 s, 20–22 °C, with cooling on ice between each burst) in 700 µl of the same solubilization solution, before 150 U of exonuclease III was added and the suspension incubated at room temperature (20–22 °C) for 15 min to degrade any DNA. The two cellular fractions were then combined and centrifuged at room temperature (12 000 g, 20–22 °C, 10 min). Prior to IEF, 100 µl 500 mM iodoacetamide was added, and the mixture incubated at room temperature (20–22 °C) for 2 h.

2-DGE and mass spectroscopic analyses of proteins.
Both 2-DGE and mass spectroscopic analyses, using matrix-assisted laser desorption ionization time-of-flight (MALDI-TOF) mass spectrometry, were performed using a PerSeptive Biosystems Voyager DE-STR mass spectrometer, with trypsin autolysis peptide masses of 842·5 and 2211·1 Da as internal standards, as previously described (Len et al., 2003, 2004). All mass spectra were obtained in reflectron-delayed extraction mode. The density/volume, or differential expression (DE) value (arbitrary units), of SyproRuby-stained protein spots of 2-DGE gels was determined using the software package z3 (Compugen). The main source of error associated with this form of quantification is the reproducibility of the 2-DGE displays themselves, as biological variation is minimized when a chemostat is used to culture bacteria. That the 2-DGE displays were the main source of error was confirmed by comparing the DE values of 25 randomly chosen protein spots selected from 2-DGE displays separated on broad-range IPG strips over the pI range 4·0–7·0. Equivalent protein spots from triplicate samples from each of three repeat continuous cultures were analysed. The data confirmed that the main source of error in determining DE values was 2-DGE. As a consequence, triplicate experimental samples from cells grown at each pH were used for all 2-DGE analyses, and the increase in the level of expression of a protein spot was based on the difference in the mean DE values.

Protein identification.
Peptide mass mapping (PMM) analyses of proteins were undertaken as previously described, making use of the six contigs of the S. mutans UA159 genome downloaded on October 6, 2001 that were translated in all six reading frames (Len et al., 2003). All translated ORFs that matched PMM data were then used to query the annotated S. mutans genome at the Oral Pathogen Sequence Databases (http://www.stdgen.lanl.gov/oragen; Ajdic et al., 2002), using the local BLAST search facility to determine the gene identification number. The original six contigs were used in this manner as some genes identified in these contigs were not present in the final annotated version.

Mass spectroscopic parameters for protein identification included a mass tolerance of 150 p.p.m. and a maximum of one missed cleavage per peptide while taking into consideration methionine sulfoxide and cysteine acrylamide modifications. Matches were defined on the basis of the number of matching peptide masses and the total percentage sequence covered by the peptides. As a general rule, a minimum total sequence coverage of 25 % was taken to match a given translated ORF of a high-Mr protein with confidence, though coverage as high as 80 % was observed with many low-Mr proteins. All translated ORFs that matched PMM data were then used to query the annotated S. mutans genome at the Oral Pathogen Sequence Databases (http://www.stdgen.lanl.gov/oragen), using the local BLAST search facility to determine the gene identification number. All gene names used are those associated with the S. mutans genome at the Oral Pathogen Sequence Database site. Theoretical Mr and pI were determined using MassLynx software version 3.4 (Micromass).

Protein isoforms.
The term ‘isoform’ is used in the text to describe a protein that exists in multiple charged forms on 2-DGE gels, where the mean observed Mr for each form calculated from the second (SDS-PAGE) dimension deviates by up to 5 % and where there is no evidence from peptide mass mapping of any form of truncation or degradation (Len et al., 2004).


   RESULTS AND DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
REFERENCES
 
The increased expression of the proton-translocating F1F0-ATPase that extrudes H+ from Streptococcus mutans (Belli & Marquis, 1991; Hamilton & Buckley, 1991; Dashper & Reynolds, 1992; Quivey et al., 2001) is a key to the survival of the bacterium in an acidic environment. This was reflected in the previously reported increase in the {alpha}- (AtpA) and {gamma}- (AtpC) subunits of the F1 component of the ATPase when the bacterium was grown at pH 5·0 (Fig. 1, Table 1; Quivey et al., 2001; Len et al., 2004). A combination of steady-state continuous-culture technology and narrow-range IPG strips, however, resulted in the resolution of an additional 197 differentially expressed protein spots on SyproRuby-stained 2-DGE gels, following acid-tolerant growth of S. mutans at pH 5·0. Of these, 167 (including all 44 extracellular protein spots) were identified by MALDI-TOF analysis, and 106 found to be associated with metabolism: glycolysis, alternative acid production and branched-chain amino acid synthesis, in particular (Len et al., 2004). The remaining 61 protein spots were associated with regulatory and/or stress-responsive pathways. These proteins included those involved in DNA replication, transcription, translation, protein folding and proteolysis and are discussed below in light of current knowledge of the possible roles they play in Gram-positive bacteria, particularly oral streptococci.



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Fig. 1. Differentially expressed S. mutans proteins involved in replication, transcription and translation from cells grown at pH 7·0 or at pH 5·0. The columns represent the percentage mean DE values for each charged isoform identified on 2-DGE gels, relative to the most highly expressed isoform. Protein spots were either upregulated (cross-hatching), down-regulated (black), or uniquely expressed (dots), relative to the alternative pH. Truncated forms of proteins are not shown, except for those observed in the extracellular milieu.

 

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Table 1. Differentially expressed cellular stress-related proteins from S. mutans grown at pH 7·0 or pH 5·0

Numbers in the Gene ID column are associated with the S. mutans genome at the Oral Pathogen Sequence Database site (http://www.stdgen.lanl.gov/oragen).

 
DNA replication and chromosome integrity
One of the consequences of intracellular acidification is the loss of purines and pyrimidines from DNA, since deoxyribonucleotides are acid labile (Lindahl & Nyberg, 1972). Unrepaired non-instructive DNA damage blocks DNA replication and can be lethal. Aborted replication, however, exposes single-stranded DNA at replication forks and results in the binding of the recombinase RecA. The DNA-bound RecA protein undergoes a conformational transition to its active form, RecA*. In both Escherichia coli and Streptococcus pneumoniae, RecA* induces an SOS response that can lead to mutation of the DNA or acquisition of pre-evolved functions by horizontal gene transfer (Taddei et al., 1997; Horst et al., 1999; Steffen & Bryant, 2000; Volkert & Landini, 2001; Bjedov et al., 2003; Katz & Bryant, 2003).

In S. mutans, RecA was found to be upregulated 6·8-fold when grown at pH 5·0 (Fig. 1, Table 1). The physiological effects of low pH on a RecA-deficient strain of S. mutans have previously indicated that RecA is required for survival in cells grown at neutral pH when subjected to a rapid drop in pH, but that cells grown at pH 5·0 can diminish the sensitizing effects of RecA deficiency (Quivey et al., 1995). This physiological observation is a consequence of the induction, in S. mutans during growth at pH 5·0, of an abasic site-specific endonuclease activity which apparently acts independently of the RecA protein (Hahn et al., 1999; Quivey et al., 2001) in a similar manner to an error-prone form of DNA polymerase I (PolA) during the SOS response in E. coli (Lackey et al., 1985; Wandt et al., 1997). Even though a DNA polymerase I protein spot was identified in the 2-DGE proteome of S. mutans, it represented only the C-terminus of the protein, and was absent at pH 5·0 (Table 1).

Along with the enhanced expression of RecA, one of the two single-stranded DNA-binding proteins coded in the S. mutans genome, Ssb (Ajdic et al., 2002), was uniquely expressed by growth of S. mutans at pH 5·0 (Fig. 1, Table 1). Both RecA and Ssb are essential for homologous genetic recombination as well as recombinational rescue and DNA repair of chromosomal replication. To the best of our knowledge, the only studies of this protein in streptococci relate to S. pneumoniae (Steffen et al., 2002; Katz & Bryant, 2003), making this the first report of a role for Ssb in the acid tolerance of S. mutans. From the different roles for recombinational repair and the SOS response currently recognized in Gram-negative and Gram-positive bacteria, and the lack of information available on streptococci, it is clear that further research is needed to understand these processes in the acid-tolerant growth of S. mutans.

RNA synthesis and degradation
Under imposed stress conditions, it is self-evident that the DNA repair and protection responses needed for the survival of the cell require the transcription of appropriate genes. While the nature of these events has been well documented in E. coli, little is known of the mechanism(s) coordinating these events in streptococci, particularly oral streptococci (Volkert & Landini, 2001). Acid tolerance in S. mutans, however, led to upregulation of the transcription proteins, DNA-directed RNA polymerase {alpha} subunit, RpoA, and two isoforms of the transcription elongation factor (cleavage stimulatory factor) GreA, by 5·4-, 7·5- and 5·1-fold, respectively (Fig. 1, Table 1). In addition, one isoform of GreA was found to be uniquely expressed in the extracellular milieu at pH 5·0 (Fig. 1, Table 2). RNA polymerase forms an elongation complex with its template DNA and the nascent RNA product. While this complex is completely processive, it is responsive both to extrinsic regulatory factors and to intrinsic signals associated with the DNA and RNA that can alter the rate of elongation and lead to a transient pause or arrest of the complex (Nakasone et al., 1998; Erie, 2002). This may be as simple as a lack of nucleotide substrates: a situation that may readily occur in S. mutans under acidic conditions, due to the disturbance of anabolic functions (Len et al., 2004). The main function of GreA is to reactivate RNA polymerase once such a halt has occurred. This is achieved by enhancing the intrinsic cleavage activity of RNA polymerase, thus releasing RNA from the elongation complex, and preventing backward translocation and hydrolysis of the RNA (Fish & Kane, 2002; Opalka et al., 2003). Other than in the current study, elevated levels of GreA have been noted in the Gram-positive bacterium Staphylococcus aureus, when challenged with oxacillin (Singh et al., 2001).


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Table 2. Differentially expressed extracellular stress-related proteins from S. mutans grown at pH 7·0 or pH 5·0

 
Two isoforms of the RNA-degrading enzyme polynucleotide phosphorylase [polyribonucleotide nucleotidyltransferase (PNPase; PnpA)] were also upregulated by at least 3·1-fold in S. mutans in acid-tolerant growth. (Fig. 1, Table 1). In E. coli, two 3'– 5' exonucleases, RNase II and PNPase, are involved in mRNA decay, since the loss of both activities results in the cessation of growth (Donovan & Kushner, 1986). Recent analysis of RNA decay in E. coli (Mohanty & Kushner, 2003) has shown that even though RNase II constitutes about 90 % of the exonucleolytic activity in the cell (Deutscher & Reuven, 1991), PNPase plays a greater role in the degradation of mRNA, since it forms part of a multi-protein complex, the ‘degradosome’, that contains an RhlB RNA helicase capable of removing secondary structure that would impede PNPase activity (Mohanty & Kushner, 2003).

While the level of understanding of RNA decay in Gram-positive bacteria is not as advanced as that of Gram-negative bacteria, in vitro RNA decay does not appear to be as severely compromised by the absence of a PNPase activity (Wang & Bechhofer, 1996). In Bacillus subtilis, however, a pnpA deletion mutant shows pleiotrophic effects, including a cold-sensitive-growth phenotype, sensitivity to growth in the presence of tetracycline and multiseptate, filamentous growth (Wang & Bechhofer, 1996). It has been hypothesized that defective processing of specific RNAs in the pnpA mutant of B. subtilis results in these phenotypes, though there is no direct evidence in support of this contention (Wang & Bechhofer, 1996). To date, no studies of the role of PNPase in oral streptococci appear to have been undertaken, though a 1·9-fold down-regulation of the enzyme has been observed in Streptococcus oralis when inoculated and grown in batch culture under aerobic conditions at low pH (Wilkins et al., 2001). Our observation of an increase in PNPase is clearly at odds with this observation. Whether the aerobic conditions used to cultivate S. oralis engender an additional oxidative stress that further influences PNPase expression requires investigation.

Translation
The incorporation of correctly encoded amino acids into proteins depends on the attachment of each amino acid to an appropriate tRNA by aminoacyl tRNA synthases (Cooper, 2000). Acid tolerance in S. mutans resulted in the upregulation of phenylalanyl- (PheS), alanyl- (AlaS) and two isoforms of threonyl- (ThsS) tRNA synthases, as well as subunits A (GatA) and B (GatB) of glutamyl-tRNA amidotransferase, which is required for the transamidation of misacylated Glu–tRNAGln to form Gln–tRNAGln in all Gram-positive bacteria (Curnow et al., 1997; Harpel et al., 2002; Table 1). In contrast, arginyl-tRNA synthase (ArgS) was down-regulated 20-fold by growth at pH 5·0 (Table 1). The other four aminoacyl tRNA synthases previously identified (Len et al., 2003) did not show any differential expression at pH 5·0 (data not shown).

In S. mutans, the 50S and 30S ribosomal subunits are composed of 51 proteins (Ajdic et al., 2002). Of these, five were identified as being upregulated by growth at pH 5·0, four of which existed in more than one charged isogenic form and possessed an observed Mr higher than that predicted from their gene sequence (Table 1). While 16 ribosomal proteins have previously been mapped on 2-DGE gels (Len et al., 2003), the inability to detect all 51 proteins is most likely due to their low Mr and/or very basic pI (>10·5), which place them at the limit of the resolving power of current 2-DGE technology. Our data imply that the number of ribosomes may increase by a factor of four in a low-pH environment – a suggestion that will require independent confirmation. Protein S1 and the ribosomal protein L10 were also found in the culture medium, but at significant levels at pH 7·0 only (Table 2).

In translation, three elongation factors, EF-Tu, EF-Ts and EF-G, are responsible for escorting aminoacyl tRNAs to the ribosome and for translocation of the ribosome along the mRNA (Fig. 1; Cooper, 2000). A fourfold increase in the expression of the four charged isogenic forms of EF-Tu was observed by growing S. mutans at pH 5·0. In each case, the observed Mr of the protein was greater than that predicted from the gene sequence (Table 1). Two charged isogenic forms of EF-G were also upregulated at pH 5·0. Most notable, however, was the finding that the reduced amounts of multiple truncated forms of EF-Tu and a C-terminal fragment of EF-G in cells grown at pH 5·0 could be measured at all, as this meant that each protein spot had the same 2-DGE coordinates, irrespective of the growth pH (Table 1). This suggested that specific cleavage events had occurred, rather than random proteolysis. A similar observation has been made with Salmonella enterica serovar Typhimurium, where two specific proteinases are believed to be responsible for forming elongation factor artefacts (Adams et al., 1999). Even if this were the case with S. mutans, the data in Table 1 indicate that the cumulative level of both EF-Tu and EF-G would be at least threefold higher during acid-tolerant growth at pH 5·0. Interestingly, no change in the third elongation factor, EF-Ts, was observed, except in the extracellular milieu, where its level was substantially upregulated at pH 7·0 (Fig. 1, Table 2).

The increase in EF-Tu and EF-G during acid-tolerant growth is also of interest from another standpoint, since it has recently been shown that both proteins behave like chaperones towards unfolded and denatured proteins in E. coli (Kudlicki et al., 1997; Caldas et al., 1998, 2000). EF-Tu, for example, recognizes the same hydrophobic binding motifs in proteins as the chaperone DnaK (see below; Malki et al., 2002). Furthermore, EF-Ts has been shown to act as a folding template in a chaperone-like manner towards its substrate protein, EF-Tu (Krab et al., 2001).

Molecular chaperones and degradation of misfolded proteins
Early pulse–chase experiments demonstrated an increase in DnaK levels during thermal stress in S. mutans, thus confirming the existence of a functional heat-shock response-system in this species (Jayaraman & Burne, 1995). The DnaK chaperone machinery prevents the misfolding and aggregation of ribosome-bound polypeptides (Szabo et al., 1994; Rudiger et al., 1997; Bukau & Horwich, 1998; Agashe & Hartl, 2000). In the current study, proteome analysis identified three isoforms of DnaK in S. mutans, which were upregulated 4·6-, 10·9- and 5·0-fold, respectively, at pH 5·0 (Fig. 2, Table 1). This confirmed a similar observation made by Jayaraman et al. (1997), who used the same values of D and pH to culture S. mutans.



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Fig. 2. Differentially expressed S. mutans molecular chaperones and proteinases from cells grown at pH 7·0 or at pH 5·0. The columns represent the percentage mean DE values for each charged isoform identified on 2-DGE gels, relative to the most highly expressed isoform in a given compartment. Protein spots were either upregulated (cross-hatching), down-regulated (black), or uniquely expressed (dots), relative to the alternative pH. Truncated forms of proteins are not shown, except for those observed in the extracellular milieu.

 
It is now clear that the function of DnaK overlaps with that of another component, trigger factor, RopA (Deuerling et al., 1999; Teter et al., 1999). RopA is a major ATP-independent molecular chaperone with prolyl isomerase activity, which binds to the large ribosomal subunit proteins L23 and L29 near the polypeptide exit site and interacts with nascent polypeptide chains prior to DnaK (Hesterkamp et al., 1996; Kramer et al., 2002). A 2·4-fold increase in RopA during steady-state growth at pH 5·0 was observed in S. mutans (Fig. 2, Table 1). Other molecular chaperones, such as the bacterial group I chaperonin, GroEL, are indispensable for cell viability (Fayet et al., 1989; Kubota et al., 1995). GroEL is able to capture and refold non-native substrate proteins up to 50–60 kDa, while protecting them from aggregation with other non-native proteins (Braig et al., 1994; Mayhew et al., 1996; Weissman et al., 1996; Xu et al., 1997). Proteome analysis showed the existence of three isoforms of GroEL in S. mutans, which were enhanced 20·0-, 6·9- and 7·0-fold, respectively, by growth in an acidic environment (Fig. 2, Table 1). While similar results have been reported for a 60 kDa chaperonin (most probably GroEL) in aerobic batch cultures of S. mutans and S. oralis (Wilkins et al., 2001, 2002), the current results differed from those previously reported in continuous culture (Lemos et al., 2001). In the previous study, S. mutans groEL mRNA was induced 2·5-fold by an acid shock from pH 7·0 to pH 5·0, but no significant differences in the levels of groEL mRNA or GroEL (determined by Western blot analysis) were observed, once steady state was achieved at D=0·1 h–1 (Lemos et al., 2001).

DnaK and RopA were also identified in the S. mutans culture fluid. Both proteins were down-regulated, by factors of 34 and 313, respectively, at pH 5·0 (Fig. 2, Table 2). These levels of DnaK and RopA were equivalent to 39 % and 36 %, respectively, of the steady-state cellular levels at pH 7·0, but were equivalent to less than 0·2 % of the steady-state cellular levels at pH 5·0, implying a loss (or secretion) rate of approximately 4·0 % h–1 of the steady-state cellular levels at pH 7·0, but a negligible rate of loss at pH 5·0 (Jacques et al., 1985). Although previous studies have not considered the extracellular milieu as a source of these proteins (Lemos et al., 2001; Wilkins et al., 2001, 2002), DnaK has recently been identified on the surface of both Streptococcus agalactiae (Hughes et al., 2002) and Haemophilus influenzae (Hartmann et al., 2001), and is known to be highly immunogenic in S. pneumoniae (Hamel et al., 1997). Whether S. mutans has surface-bound molecular chaperones is a matter of conjecture, as our technique of protein extraction did not discriminate between the various cellular compartments. Irrespective of this, the increase in the steady-state cellular levels of DnaK, RopA and GroEL in S. mutans at pH 5·0 is consistent with a need for an enhanced complement of molecular chaperones to counteract the denaturing properties of an acidic cytosol when cytosolic pH falls below 6·5 (Dashper & Reynolds, 1992).

Proteins which cannot be folded by molecular chaperones may be targeted for degradation, in order to recycle amino acids for de novo protein synthesis (Jenal & Hengge-Aronis, 2003). Among the proteins that can carry out such functions are the ATP-dependent proteases of the Clp family, which possess a dual chaperone/proteinase role. Proteolysis by Clp requires a serine type peptidase, the ClpP subunit, and a regulatory ATPase subunit consisting of several orthologues (Gottesman et al., 1997; Porankiewicz et al., 1999; Lemos & Burne, 2002). In S. mutans, only one ATP-binding subunit, ClpL, was expressed, and solely in growth at pH 5·0 (Fig. 2, Table 1). ClpL homologues appear to be specific to Gram-positive bacteria, as they have not been found in Gram-negative bacteria (Lemos & Burne, 2002; Kwon et al., 2003). To date, information on the role of ClpL is limited to a single study of the heat-shock response in S. pneumoniae, in which mutations in the clpL and clpP genes were found to modulate virulence-gene expression, and purified recombinant ClpL was shown to possess molecular chaperone properties (Kwon et al., 2003). Our data suggest that ClpL is upregulated and maintained under low pH conditions, with the implication that it plays a vital role in pH tolerance. It remains to be seen whether ClpL operates solely as a molecular chaperone, or whether it interacts with ClpP to initiate proteolysis.

Two other differentially expressed proteinases were identified by MALDI-TOF, both belonging to the pep gene family of proteinases, for which 13 different genes exist in the S. mutans genome (Ajdic et al., 2002). One of these proteinases, a truncated version of a putative PepB, was down-regulated by a factor of 4·2 in growth at pH 5·0 (Fig. 2, Table 1). In Group B streptococci, PepB has oligopeptidase activity and has been shown to degrade a variety of small bioactive peptides, as well as the synthetic collagen-like substrate N-(3-[2-furyl]acryloyl)-Leu-Gly-Pro-Ala in vitro (Lin et al., 1996). The second proteinase was homologous to the dipeptidase PepD of Lactobacillus helveticus (Ajdic et al., 2002). This enzyme does not appear to be similar to the cytoplasmic PepD isolated from S. mutans and S. sanguis which catalyses the hydrolysis of X-Pro dipeptides (Cowman & Baron, 1997), since the Lactobacillus enzyme is not able to hydrolyse di- and tripeptides containing proline (Vesanto et al., 1996). PepD was upregulated 12·3-fold at pH 5·0 (Fig. 2, Table 1).

It is perhaps pertinent to note that proteolysis has another role additional to the recycling of amino acids for de novo protein synthesis, and that is to regulate cellular events by degrading regulatory proteins, thereby controlling the response of the cell to an imposed stress. A readjustment in the composition of the cellular proteolytic machinery would therefore most likely have pleiotropic consequences for the cell, as any change in the nature or complement of specific proteinases would be expected to affect proteins that are subject to regulation by proteolytic events (Jenal & Hengge-Aronis, 2003).

Concluding remarks
This study has made use of the steady-state conditions enabled by anaerobic continuous culture in a chemostat to study alterations in the stress-response proteome of S. mutans, following adaptation and tolerance to growth at low pH. The proteome literature relating to acid adaptation and tolerance in oral streptococci contains disparate findings. For instance, proteome analysis of S. mutans during acid adaptation, using an aerobic batch-culture model without pH control, identified eight of the 28 differentially expressed proteins found in the current study. The levels of six of these were down-regulated by an average of 2·6-fold, while DNA-directed RNA polymerase was down-regulated 33-fold. Only a 60-kDa chaperonin was upregulated to a similar extent to that observed with GroEL in the current study. It is difficult to assess whether the aerobic conditions, the use of a different culture medium and/or the dissimilar generation times of 1·0 h and 6·6 h, respectively, for the control and the experimental batch cultures influenced the outcome (Wilkins et al., 2002). Since an arbitrary ratio of the levels of expression has been evaluated in both studies, it may be that, in batch culture, the control bacteria are adapting to a fall in the extracellular pH from a starting value of 7·0 to 6·2 at harvest (heading for a final of pH 5·3 at stationary phase) by initially overexpressing stress-related proteins. This would contrast with the steady-state levels measured in a chemostat, once S. mutans had adapted to the prevailing pH conditions. Such an overshoot in enzyme levels as an initial response to change has been noted previously, albeit in continuous culture (Carlsson & Elander, 1973; Koplove & Cooney, 1978), and if occurring in batch culture would explain the apparent reduction in the ratio between control and experimental protein values observed. Whatever the reason for the disparity, one must conclude that the two models are reflecting dissimilar events.

Although a number of stress-related proteins, in both Gram-positive and Gram-negative bacteria, are either well characterized or the subject of concerted ongoing study, others are not. This study has shown the involvement of at least three of these proteins in the acid-tolerant growth of S. mutans: the transcription elongation factor, GreA; the ATPase protease, ClpL; and the single-stranded DNA-binding protein, Ssb. The role of each of these proteins warrants further examination in light of the paucity of information regarding their mode of action. The subtle differences in homologous recombinational repair recently observed between S. pneumoniae and E. coli, along with the apparent differences in the complement of genes associated with the SOS-induced response in Gram-positive and Gram-negative bacteria (Steffen et al., 2002; Katz & Bryant, 2003), emphasize this point, particularly as the regulons involved do not appear to have been studied in relation to acid tolerance or other stress responses in oral streptococci such as S. mutans.


   ACKNOWLEDGEMENTS
 
This research was supported by Grant no. R01 DE 013234 from the Institute of Dental and Craniofacial Research, National Institutes of Health (NIH), USA, and was facilitated by access to the Australian Proteome Analysis Facility (APAF), established under the Australian government Major National Research Facility program. We wish to thank Dr S. J. Cordwell from APAF for his continued advice on all matters relating to 2-DGE proteomics and Dr K Byth Wilson from Westmead Hospital for the statistical analyses. A. C. L. L. was the recipient of an Australian Postgraduate Award.


   REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS AND DISCUSSION
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Received 31 December 2003; revised 14 February 2004; accepted 17 February 2004.