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Keywords:

  • Ffh;
  • Acid stress;
  • Continuous culture;
  • Streptococcus mutans

Abstract

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

Previously, we described in Streptococcus mutans strain NG8 a 5-gene operon (sat) that includes ffh, the bacterial homologue of the eukaryotic signal recognition particle (SRP) protein, SR54. A mutation in ffh resulted in acid sensitivity but not loss of viability. In the present study, chemostat-grown cells of the ffh mutant were shown to possess only 26% and 39% of the parental membrane F-ATPase activity and 55% and 75% of parental glucose–phosphotransferase (PTS) activity when pH-7 and pH-5-grown cells, respectively, were assayed. Two-dimensional-gel electrophoretic analyses revealed significant differences in protein profiles between parent and ffh-mutant strains at both pH 5 and pH 7. It appears that the loss of active SRP (Ffh) function, while not lethal, results in substantial alterations in cellular physiology that includes acid tolerance.


1Introduction

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

Prokaryotes employ two main strategies for translocating newly synthesized proteins from ribosomes to membranes and extracellular spaces. The first, post-translational translocation, is exemplified in the general secretory pathway (GSP) that is dependent on the chaperone SecB and membrane-associated proteins, SecA/Y/E/D/G (reviewed in [1,2]). The second, co-translational translocation involving the signal recognition particle (SRP) [3–6]. The SRPs described to date in prokaryotes are far simpler than their analogues in eucaryotes and consist of a minimal functional unit of a chaperone (Ffh), and a small cytoplasmic(sc) RNA (reviewed in [6]). It has been proposed that in E. coli the SRP functions in targeting integral membrane proteins [7,8], whereas in Bacillus subtilis the SRP functions as a general targeting factor for most secreted proteins. In each of these species a mutation in ffh and/or fts Y, which encodes the SRP membrane receptor FtsY, as well as depletion of the SRP-associated small cytoplasmic (sc) RNA [9,10] is lethal.

Previously, we [11] described a 5-gene operon in the dental pathogen Streptococcus mutans that we named sat (s ecretion and a cid t olerance; [12,13]) that appears to be regulated by environmental pH levels. Included in this operon is an ffh homologue plus four other genes of unknown functions. Non-polar mutations created individually in each of the five genes yielded viable mutants [11]. The ffh mutant displayed an acid-sensitive phenotype and failed to grow in batch culture upon passage of stationary-phase cultures to fresh medium at pH 5.0. Exponential-phase cultures of the ffh mutant grew, albeit poorly compared to the parent, when acid-shocked to pH 5.0. Viability under non-acid stress conditions was surprising and defied the conventional wisdom that a functioning Ffh-containing SRP was necessary for microbial survival and growth [6]. This suggests that other mechanisms are present in S. mutans responsible for translocating proteins to the membrane while the SRP (Ffh) may be more important during stress response.

The present study describes an assessment of physiologic adjustments made by S. mutans MK4, an ffh mutant, during acid stress. Indeed, data presented below show that this mutant not only survives but also grows well in continuous culture under anaerobic conditions at acidic pH and appears to make significant physiologic changes likely associated with its survival.

2Materials and methods

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

2.1Bacterial strains and continuous culture conditions

Streptococcus mutans parent strain NG8 [14] and ffh-mutant MK4 [11], a non-polar mutant created by insertional inactivation with a promoterless aph3A (kanamycin-resistance marker), were cultured in 1% tryptone/0.5% yeast extract supplemented with 16.6 mM glucose and 11.5 mM K2HPO4 at 37 °C in a New Brunswick Scientific BioFlo model chemostat (New Brunswick Scientific, Edison, NJ) with a 1.2 l culture vessel volume. The dilution rate (D) was 0.1 h−1 and the pH was maintained with 2.0 N KOH using a pH controller until steady-state growth was achieved. The culture vessel was stirred at 200 rpm and sparged continuously (0.8 l m−1) with an anaerobic gas mixture of 85%N2, 10%H, 5%CO2. One-liter samples were collected rapidly through the sampling port, by slightly pressurizing the culture vessel with the gas mixture, and then placed on ice. The sample was centrifuged, washed twice in ice-cold dH2O (or buffers as indicated below), aliquoted and stored at −80 °C. The culture supernatants were filter-sterilized and stored at −20 °C prior to end-product analysis.

Batch culture of both acid- and non-acid-adapted ffh-mutant MK4, as well as parent strain NG8, was performed by inoculation of each strain into THYE broth with subsequent incubation for 16 h at 37 °C. The cultures then were passaged 1:10 (v/v) into warm THYE and incubated at 37 °C until early exponential phase was reached. The cultures were then passaged again, 1:10 (v:v), into warm THYE at pH 7.0 and 5.0 and growth of all cultures was monitored using a Klett–Summerson colorimeter (Klett Manufacturing Co., Inc., NY) until stationary phase was reached. All growth curves were performed in duplicate.

2.2Fermentation end-product analysis

Protein from each culture supernatant was precipitated with 35% perchloric acid for 10 min on ice followed by neutralization with 2.5 N KOH. The resulting precipitate was removed by centrifugation, and fermentation end-product concentrations in the supernatants were determined using an Aminex HPX87H organic acid analysis HPLC column as described by Snoep et al. [15].

2.3Membrane preparations

An aliquot of cells that had been stored as a concentrated slurry in TM buffer (50 mM Tris–maleate, pH 6.0, 20 mM MgCl2, 0.1 mM phenylmethylsulfonylflouride, PMSF) at −80 °C was transferred to a frozen mortar with alumina powder (A-5, Sigma; 3 g powder g wet wt−1 of cells) and ground for 20 min according to the method of Vadeboncoeur et al. [16]. The sample was centrifuged for 5 min at 3000g to remove the alumina and then again at 16,000g for 1 h to remove intact cells and cell debris. The supernatant was centrifuged at 100,000g for 18 h to pellet the membranes. The supernatant (cytoplasm) was placed on ice and used immediately for lactate dehydrogenase (LDH) assays. The membrane pellets were rinsed three times with TM buffer and suspended in TM buffer for F-ATPase assays.

2.4Enzyme assays

2.4.1Lactate dehydrogenase

Cytoplasm samples derived from membrane preparations (above) were assayed for LDH spectrophotometrically according to the method of Hillman et al. [17]. Samples were assayed for protein using the bicinchoninic acid (BCA) protocol (Pierce Chemical Co., Rockford, IL). Specific activity was calculated as nmoles NADH oxidized min−1 mg−1 protein.

2.4.2Membrane F-ATPase

Membrane F-ATPase activity was assayed according to the method of Bender et al. [18]. Briefly, membranes (∼50 μg) in TM buffer (no PMSF added) were incubated for 5–10 min at 37 °C after addition of 5 mM ATP. The reaction was stopped by addition of 0.01 N HCl and the tubes were placed on ice for 10 min. Samples containing buffer plus ATP and membranes minus ATP were included as controls. Membranes were removed by centrifugation at 14,000g for 15 min and supernatants were assayed for inorganic phosphorus by a modification of Weisman and Pillegi [19] that uses the phosphorous determination kit from Sigma Chemical Company (St. Louis, MO). Membranes were assayed for protein using the BCA protocol. Specific activities were calculated as nmoles Pi released min−1 mg−1 total membrane protein.

2.4.3Total cellular-F-ATPase

Cells were thawed and washed once in TM buffer then suspended in 1 ml of TM buffer per 25 ml cell culture. Toluene was added (100 μl ml−1 of cell suspension) and the cells were vortexed for 30 s. After vortexing the cells were placed on ice for 5 min, snap-frozen in dry ice–ethanol and thawed in a 37 °C water bath. Cells underwent a total of three freeze–thaw cycles and then were pelleted. The supernatant was decanted and the cells were resuspended in TM buffer and placed on ice. The assay was performed by adding 100 μl of decryptified cells to 3 ml of TM buffer without PMSF added. ATP (5 mM) was added to the suspension and a 0.5 ml sample was immediately removed and added to 2 ml of 20% trichloracetic acid for the zero time point. Samples were similarly collected at 5 and 10 min and allowed to sit for 10 min at room temperature. The samples were centrifuged and assayed for inorganic phosphorus as above. Samples of the decryptified cell suspensions were assayed for protein as above. Specific activities were calculated as nmoles Pi released min−1 mg−1 total cellular protein.

2.4.4PEP-dependent phosphotransferase (PTS)

Cells were thawed, washed once with ice-cold 100 mM sodium phosphate buffer, pH 7.2, with 5 mM MgCl2 and suspended in 3 ml of ice-cold buffer. About 150 μl of a 1:9 toluene:acetone mix was added and the suspensions were vortexed for 1 min followed by cooling on ice for 30 s. This was repeated three more times. The decryptified cells were assayed for sugar-specific PTS activity essentially according to the method of LeBlanc et al. [20] modified as follows: in a 1 ml reaction volume was added 1 mM NaF, 0.1 mM β-NADH, 0.5 mM PEP, 3 U LDH and 25–100 μl of decryptified cells. Specific sugars (1 mM) were added and the change in O.D. was monitored for 5 min at 340 nm. Decryptified cell suspensions were assayed for protein. Specific activities were expressed as nmoles of PEP-dependent NADH oxidized min−1 mg−1 protein.

2.5Cellular protein extraction for two-dimensional gel electrophoresis

Cell suspensions were frozen and thawed twice and centrifuged at 2500g for 5 min. Pellets were suspended in 0.35 ml of lysis buffer (8 M urea, 2% CHAPS, 62 mM DTT and 2% (v/v) pharmalyte (Amersham Pharmacia Biotech, Piscataway, NJ) and sonicated four times for 5 min each in the presence of 0.2 mm glass beads. Between bursts, cells were placed on ice for 10 min. Sonicated cells were centrifuged for 5 min at 14,000g and the supernatants stored at −20 °C. Protein was measured using the method of Fey et al. [21] with lysis buffer as the diluent for samples and standards.

2.6Two-dimensional gel electrophoresis and Western immunobloting

Proteins in 250 μg of cell extracts were separated by 2D gel electrophoresis essentially as described previously [22]. Gels were stained with Coomassie brilliant blue R-250 or silver and vacuum dried. Previously [23] 2D gels of cellular protein extracts using the same IEF and SDS–PAGE conditions were run and individual protein spots were identified by in-gel proteolytic digestion of resolved proteins and MALDI-TOF MS analysis. Their gel locations were mapped and each protein spot was assigned a number. In this study gels were run in triplicate and visual comparisons were made between protein spots from MK4 and NG8 at each pH. An increase or decrease in the relative amount of a particular protein was concluded if the result was seen in at least two out of three gels.

For Western immunoblotting, 60–120 μg of cellular extracts were subject to 2D-gel electrophoresis as described above. Following electrophoresis proteins were electrophoretically transferred to Immobilon membrane (hydrophobic polyvinylidene difluoride membrane, Millipore Intertech, Bedford, MA). The blots were rinsed three times with Tris-buffered saline, pH 7.4 (TBS) and non-specific binding sites were blocked with 5% non-fat milk in TBS at 37 °C for 1 h. After a wash with 0.1% non-fat milk in TBS, the blots were incubated with each rabbit serum (dilution 1:400) on a shaker at room temperature. A new wash followed and the blots were incubated for 2 h with horse radish peroxidase-conjugated goat–anti-rabbit-IgG (DAKO, Denmark) in 1% non-fat milk and 0.05% Tween 20 in TBS. After being washed in TBS, the blots were developed in the dark for 15 min using 1% peroxidase substrate (3-amino-9-ethylcarbazole in acetone; Sigma Chemical Co.) in 50 mM sodium acetate buffer, pH 5.0, and 0.015% H2O2. Development was stopped by rinsing the blots in water.

3Results and discussion

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

3.1An Ffh-negative mutant can grow under acid conditions in continuous culture

Growth of the ffh-mutant, MK4, in continuous culture, yielded surprising results. We showed previously [11] that MK4 grew slowly in batch culture compared to parent strain NG8 at pH 7 and not at all following passage of a 16 h culture to pH 5.0-adjusted broth. In the present study when MK4 was grown in continuous culture under glucose limitation and the culture pH was allowed to drop gradually from 7.0 to 5.0 as a result of the cell's normal metabolism, not only did the cells survive following continuous culture at pH 5.0 but the steady-state culture density of 159 ± 7.0 measured using the Klett colorimeter was only slightly lower than that of the parent strain measured at 198 ± 11. In pH 7.0-maintained cultures, cell densities reached 230 ± 3.4 and 180 ± 21 in cultures of NG8 and MK4, respectively. Thus, while the mutant did not grow to wild-type levels, it was clearly able to overcome the acid stress and grow at pH 5.0 under conditions of anaerobic continuous culture.

To examine whether the acid-tolerant phenotype displayed by MK4 following continuous culture carried over to growth in batch culture, steady-state cells harvested from continuous culture at pH 5.0 were placed in batch culture at both pH 5.0 and 7.0 alongside non-acid-adapted cells. Both acid-adapted and non-adapted MK4 cells grew identically at both pHs with similar final culture densities (data not shown). The ability to revert to its acid-sensitive phenotype in batch culture at pH 5.0 demonstrates that acid-tolerance during continuous culture does not involve a genetic alteration, such as a spontaneous mutation, but rather a reversible physiologic adaptation.

To determine if depriving S. mutans of Ffh altered its ability to ferment glucose we analyzed supernatants from steady-state cultures of MK4 and NG8 by HPLC. Analysis showed that the mutant and parent strain produced organic acids typical for S. mutans. At pH 7.0, strain NG8 produced an average of 29.8 mM formate, 15.8 mM acetate, 10.1 mM ethanol and 0.15 mM lactate, while MK4 produced an average of 29.6 mM formate, 16.0 mM acetate, 9.15 mM ethanol with no detectable lactate. At pH 5.0 both strains produced lactate as the most abundant organic acid, 19.4 mM for NG8 and 24.4 mM for MK4 as average values. In addition formate was measured at averages of 16.8 mM in NG8 and 10.2 mM in MK4, actetate at 10.7 mM in NG8 and 6.35 mM in MK4 and ethanol at 6.50 mM in NG8 and 3.00 mM in MK4. Carbon dioxide production was not measured. Ethanol is likely to be slightly under represented due to evaporation during cell culture in the chemostat in which anaerobic gases are continuously sparging the culture medium. These results are typical of S. mutans grown anaerobically under glucose limitation at these pHs [24,25] and indicate that a mutation in ffh did not impair the organism's ability to ferment glucose under either neutral or acid conditions although a slightly higher percentage of lactate versus the other organic acids was produced in the mutant at pH 5.0 as compared with the parent strain.

3.2Proton ATPase, glucose–PTS and lactate dehydrogenase activities are altered in an Ffh-negative mutant

F-ATPase activity was measured in both decryptified whole cells and purified cellular membrane preparations from steady-state cultures of NG8 and MK4 maintained at pH 7.0 and pH 5.0. Nearly identical activities were measured in decryptified cells of both the parent and mutant strains at each pH with about 65% increased activity at pH 5.0 in each strain (Table 1). This is typical for an acid tolerance response since F-ATPase is necessary for increased proton extrusion at lower pH values [18,26]. In purified membrane preparations significant decreases in specific activity of F-ATPase of 74% at pH 7.0 and 61% at pH 5.0 were observed for the mutant MK4 when compared with NG8 (Table 1). F-ATPase activities from whole, decryptified cell assays result from the F1 portion of the enzyme (reviewed in [27]), regardless of whether or not it is complexed with the F0 component. The F0 component anchors the F1 component at the membrane and serves as the membrane embedded proton extrusion channel. The decreased enzymatic activity associated with mutant membranes may reflect less efficient assembly of membrane embedded F0 component subunits that may require the SRP system for translocation and insertion. Altered insertion of F-ATPase into the membrane may be a major part of the acid tolerance response and these data may explain the acid-sensitive phenotype of the ffh mutant MK4. Our data therefore suggest that Ffh may play an active role in the assembly of a functional F-ATPase at the membrane. Generation of an antibody to the F0 component to probe Western blots of membrane extracts will help confirm this.

Table 1.  Enzyme activitiesa
Strain/pHLDHbF-ATPasecPTSd
  MembraneWhole cellGlucoseFructose
  1. aAll enzyme assays were performed on the same chemostat cell harvests. Mean values±SE were calculated for each.

  2. bDuplicate assays were performed for each sample and specific activity expressed as μmol NADH oxidized min−1mg−1 whole-cell protein.

  3. cDuplicate decryptified cell aliquots were assayed in duplicate for whole-cell activity. Membrane preparations were assayed in triplicate. Specific activities are expressed as nmoles Pi released min−1mg−1 whole-cell protein.

  4. dAssays were performed in triplicate and specific activity expressed as nmoles NADH oxidized min−1mg−1 whole-cell protein. Reactions were initiated with glucose or fructose as indicated.

NG8–7.01.28 ± 0.04123 ± 4164 ± 9262 ± 8109 ± 1
MK4–7.00.09 ± 0.0133 ± 3165 ± 8144 ± 11117 ± 3
NG8–5.04.68 ± 0.05193 ± 15285 ± 16110 ± 988 ± 9
MK4–5.00.35 ± 0.0475 ± 2268 ± 5182 ± 1272 ± 3

Glucose and fructose–PEP-dependent PTS-specific activities of whole decryptified cells were measured to test whether a lack of Ffh affected this membrane-associated sugar transport system. Reductions in the glucose–PTS activity in the mutant strain as compared to NG8 were measured to be 45% in pH 7.0-maintained cells and 25% in pH 5.0-maintained cells (Table 1). In contrast, there was little if any difference in fructose–PTS activity between parent and mutant. Wen et al. [28] demonstrated that mutations in one or both fructose-dependent PTS loci, fruI and fruCD, encoding the inducible and constitutive EII permeases, respectively, resulted in wild-type levels of growth in 0.5% fructose supplemented media. This suggests that S. mutans has compensatory mechanisms for fructose uptake in the absence of the fruI and fruCD gene products and these alternative mechanisms may not depend on a functional SRP. The significant reductions in glucose–PTS activity in the mutant indicate a clear role for the SRP in translocation of the EIIgluc permease to the membrane.

Interestingly, while fermentation end-product analysis demonstrated the presence of predominantly lactate in both the parent and ffh mutant strains at pH 5.0, LDH activity values from cellular extracts of the mutant and parent were markedly different. Both the parent and mutant strains displayed a 4-fold increase in LDH specific activity following a shift in culture pH from 7.0 to 5.0 in anaerobically grown chemostat cultures of S. mutans, a result also seen by Iwami et al. [25]. However, there was a substantial 15-fold decrease in specific activity in the mutant as compared to the parent strain at both pHs (Table 1). The marked reduction in measurable, soluble LDH specific activity in MK4 cytoplasmic extracts was reproducibly obtained using cells from two different chemostat cultures. This enzymatic assay is driven by fructose 1–6-diphosphate (FDP) and measures oxidation of NADH making it specific for LDH [29]. Lactate is produced only via this pathway in S. mutans and was detected at nearly the same levels by HPLC analysis in both strains grown at pH 5.0. In contrast to the enzymatic data, reproducible two-dimensional gel electrophoresis (Table 2A) did not demonstrate any difference in LDH protein spot intensity between the parent and mutant strains. The reason for this disparity is unclear. Yamada and Carlsson [24] reported inhibition of LDH activity by high intracellular levels of ADP and inorganic phosphorus, even in the presence of optimal levels of FDP in glucose-limited S. mutans cultures. Indeed, the increased cytoplasmic ATPase activity in MK4 could yield such high levels of putative inhibitors. Other physiologic co-factors (e.g. NADH) in unusual abundance in the mutant cytoplasm also could be considered as LDH inhibitors. Yet, the data presented above show that lactate is the predominant organic acid produced in MK4 and NG8 during continuous culture at pH 5.0. LDH, while not assayable in MK4 extracts, clearly is active in vivo. A “compartmentalization” of the enzyme, due to a tight association with mutant, but not wild-type membranes, thereby removing LDH from the cytoplasm (and from derived cellular extracts), could explain our data. Our attempt to measure LDH activity from purified membrane samples however was unsuccessful since there were NAD-reducing reactions present that yielded high “background” levels of NADH. Studies of the protein compositions of MK4 and NG8 membranes are in progress.

Table 2A.  Differential expression of carbohydrate metabolism enzyme
inline image

The alteration in soluble LDH activity, coupled with measurable decreases in specific activities of both membrane F-ATPase and cellular glucose–PTS points to broad changes in the cellular physiology of mutant MK4 following a loss of SRP function. Presumably such changes could allow for growth of the mutant strain under both acid and non-stress conditions.

3.3Two-dimensional electrophoretic and Western immunoblot analyses of whole-cell lysates from an ffh mutant further demonstrate changes in cellular protein composition

Whole cell lysates prepared from aliquots of the same steady-state cultures used for the enzymatic analyses described above were visually evaluated by 2D gel electrohoresis and silver-staining. Cellular samples from at least two individual chemostat runs were extracted and run for analysis on gels in triplicate. Fig. 1 shows a sample comparison of 2D gels of NG8 (panels A and C) and MK4 (panels B and D) extracts of steady-state cells cultured at pH 7.0 (panels A and B) and pH 5.0 (panels C and D). Comparisons were also made between gels of NG8 extracts of steady-state cells cultured at pH 7.0 and 5.0 and between MK4 extracts of steady-state cells cultured at pH 7.0 and 5.0 (not shown). Tables 2A and 2B are the collated results of the comparative analyses. There was a decrease in the amount of several enzymes involved in carbohydrate metabolism (Table 2A), including glyceraldehyde-3-phosphate dehydrogenase, pyruvate kinase and acetate kinase, in strain MK4 as compared with NG8 grown at pH 7. A decrease in the PTS component EII(AB) was also observed in MK4 compared to NG8 at both pH 7.0 and 5.0 in agreement with enzymatic assays (Table 1). EII complexes are the sugar specific protein complexes of the PTS system and include a membrane-embedded sugar specific permease involved in transport and ultimate phosphorylation of incoming sugars (reviewed in [30]). Western immunoblot analysis using rabbit-anti-EIIman serum demonstrated a reduction in EII(AB) spot intensity in MK4 as compared with NG8 (Fig. 2B). In addition, when blots of replicate gels were developed with rabbit anti-EI serum (Fig. 2A) a slight decrease in reactivity was seen in the mutant at both pHs compared with the parent.

image

Figure 1. Two-dimensional gel electrophoresis of whole-cell proteins. Cellular lysate proteins from S. mutans parent strain NG8 (A, C) and ffh mutant strain MK4 (B,D) cells grown to steady-state at pH7.0 (A, B) or 5.0 (C, D) were focused over a pH range of 4.0–7.0, electrophoresed through 14% polyacrylamide gels and silver-stained. Filled arrowheads in panels B and D indicate a relative increase, and unfilled arrowheads a relative decrease, in spot intensity compared to those same spots in panels A and C, respectively. Regular arrows point to HPr isoforms. The spots circled in panel B represent a cluster of unidentified acidic proteins present only in cellular extracts of MK4.

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Table 2B.  Additional protein differences between NG8 and MK4
inline image
image

Figure 2. Western immunoblot comparisons of Enzyme I and Enzyme II (A, B) from whole-cell lysates of NG8 and MK4 (ffh). Cells were grown to steady state at pH 7.0 or 5.0 and developed with antibodies to Enzyme I (panel A) and Enzyme IImann (panel B) from Streptococcus salivarius. Proteins were focused over a pH range of 4.0–7.0, electrophoresed through 14% polyacrylamide gels and blotted onto PVDF membranes. Blots were developed with antiserum at a 1:400 dilution.

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Silver-stained 2D gels of parent and mutant cellular extracts included a densely staining protein spot (#615) running at ∼16 kDa (Fig. 1A–D and Fig. 3A), identified by antibody (Fig. 3B) as the PTS-associated phosphocarrier protein, HPr. HPr is differentially phosphorylated and thought to be a key regulatory component of the PTS [30] and has been shown to be cell-surface associated in S. mutans[31]. HPr protein spot #615 was observed to migrate at three different isoelectric points relative to the 10 kDa chaperone (Fig. 1, spot #644). There was no observable variation in spot intensity of the three spot 615 isoforms however. Another less intensely staining isoform of HPr (spot #615a) demonstrated a decrease in intensity in MK4 cells maintained at pH 5.0 when compared to NG8 and MK4 cells maintained at pH 7.0 (Table 2A). The various isoforms of HPr present may be representative of the different states of phosphorylation that HPr exists in depending on the stage of glucose translocation. Considerably more work remains to be done to understand the significance of these results, although they suggest an altered regulatory role for HPr in sugar metabolism in MK4.

image

Figure 3. Western immunoblot identification of HPr. Whole-cell lysates of NG8 were focused over a pH range of 4.0–7.0, electrophoresed through 14% polyacrylamide gels and either silver stained (panel A) or blotted onto PVDF membrane (panel B). The blot was developed with rabbit anti-HPr serum from Streptococcus salivarius at a 1:400 dilution.

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The relative amounts of 14 proteins were increased in mutant MK4 in response to acid stress (Table 2B; MK4, pH 5.0 vs. MK4, pH 7.0) but not in parent NG8 under the same growth conditions. The proteins included two chaperones, DnaK (low MW isoform) and the 10 kDa chaperone as well as alkylhydroperoxidase and SSU ribosomal protein S6P. These proteins were identified previously [32] on similarly run 2D gels using MALDI-TOF mass spectrometry. The identity of the remaining 10 spots is unknown at present. Ribosome recycling factor (RRF), which releases ribosomes from mRNA, was increased in MK4 at pH 7.0 compared to NG8 at pH 7.0 but decreased at pH 5.0 compared to pH 7.0-grown MK4 cells and pH 5.0-grown NG8 cells (Table 2B). Without RRF, protein synthesis would be decreased or inhibited [33]. There were also a number of unidentified acidic proteins that were increased in MK4 at both pH 5.0 and 7.0 compared to NG8 (see circled area of Fig. 1B) and may reflect induction of proteins important or necessary for survival that compensate for the loss of Ffh. Alternatively, some increases in spot intensity may also represent a “back-up” of non-translocated proteins in this mutant. This apparent pattern of adaptation involving increased biosynthesis of chaperones to aid in disposal or translocation of proteins dependent on the SRP and a general decrease in protein synthesis is consistent with changes in patterns of transcription observed for S. cerevisae in response to shut-down of its SRP [34].

Results from experiments performed in our present study provide an overview of the physiological consequences of a mutation in the Ffh gene as they relate to acid tolerance in this organism. Overall, a mutation in ffh resulted in alterations in several important metabolic processes involved with pH homeostasis, pyruvate dissimilation and sugar transport during growth in continuous culture. This mutation also results in salt sensitivity (Crowley and Brady, unpublished data) suggesting that the SRP in S. mutans may play an important role in generalized stress response. Future studies will involve identification of SRP substrates and their roles in membrane biogenesis and stress response in S. mutans.

Acknowledgements

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

We thank Dr. Christian Vadeboncoeur (Laval University, Québec, Canada) for the gift of antibodies to Streptococcus salivarius HPr, EI, and EIImann proteins. Our work was supported by PHS Grant DE08007 from the National Institute of Dental and Craniofacial Research.

References

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References
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