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Previously, it has been demonstrated that the membrane fatty acid composition of Streptococcus mutans is affected by growth pH (E. M. Fozo and R. G. Quivey, Jr., Appl. Environ. Microbiol. 70:929-936, 2004; R. G. Quivey, Jr., R. Faustoferri, K. Monahan, and R. Marquis, FEMS Microbiol. Lett. 189:89-92, 2000). Specifically, the proportion of monounsaturated fatty acids increases when the organism is grown in acidic environments; if the shift to increased monounsaturated fatty acids is blocked by the addition of a fatty acid biosynthesis inhibitor, the organism is rendered more acid sensitive (E. M. Fozo and R. G. Quivey, Jr., Appl. Environ. Microbiol. 70:929-936, 2004). Recently, work with Streptococcus pneumoniae has identified a novel enzyme, FabM, responsible for the production of monounsaturated fatty acids (H. Marrakchi, K. H. Choi, and C. O. Rock, J. Biol. Chem. 277:44809-44816, 2002). Using the published S. pneumoniae sequence, a putative FabM was identified in the S. mutans strain UA159. We generated a fabM strain that does not produce unsaturated fatty acids as determined by gas chromatography of fatty acid methyl esters. The mutant strain was extremely sensitive to low pH in comparison to the wild type; however, the acid-sensitive phenotype was relieved by growth in the presence of long-chain monounsaturated fatty acids or through genetic complementation. The strain exhibited reduced glycolytic capability and altered glucose-PTS activity. In addition, the altered membrane composition was more impermeable to protons and did not maintain a normal ΔpH. The results suggest that altered membrane composition can significantly affect the acid survival capabilities, as well as several enzymatic activities, of S. mutans.
In response to environmental stress, bacteria invoke numerous protective measures to survive. One main mechanism for survival involves alteration of either the fatty acid or phospholipid composition of the organism's membrane (1, 10, 14, 27, 28). For example, increases in the proportion of monounsaturated membrane fatty acids have been linked to cold shock survival by Bacillus subtilis and Escherichia coli (1, 17). In addition, increased proportions of cyclopropane fatty acids have been shown to be crucial for acid shock survival of E. coli (14).
In response to acidification of its environment, the cariogenic organism Streptococcus mutans increases the proportion of monounsaturated membrane fatty acids (19, 33). Using a glucose-shock chemostat model system, we observed that this increase occurs when the organism is allowed to self-acidify through the metabolism of glucose and occurs rapidly over time (19). This shift of membrane composition could be prevented by the addition of the fatty acid biosynthesis inhibitor cerulenin, which in turn, rendered the organism more acid sensitive (19). The significance of monounsaturated membrane fatty acids in low-pH survival of S. mutans UA159 was further explored here.
The enzyme responsible for the generation of monounsaturated membrane fatty acids in S. mutans was unknown. In E. coli, the bacterial model for fatty acid biosynthesis, unsaturated fatty acids are generated through the activity of FabA (13, 36). However, based upon sequence homology searches, FabA was not identifiable in the S. mutans genome (2). Recently, a novel protein was characterized in S. pneumoniae that is capable of isomerizing trans unsaturated bonds to their cis isomers (24). In an extensive biochemical and genetic analysis, the novel enzyme, termed FabM, was shown to be a trans-2, cis-3-decenoyl-ACP isomerase. Using this information, a putative homologue was identified in the S. mutans strain UA159. A mutant with an isogenic mutation in fabM was generated, and its ability to survive acidic pH environments was measured. The inability to produce monounsaturated membrane fatty acids rendered the organism acid sensitive, reiterating the necessity of monounsaturated fatty acids for low-pH survival of S. mutans. Further physiological characterization of the fabM mutant strain demonstrated that altered membrane composition had significant effects on glycolysis, phosphotransferase (PTS) activity, membrane permeability, and F1F0 ATPase activity. Thus, altering membrane fatty acid composition appears to have a significant impact on multiple physiological characteristics and enzymatic activities.
(This work was conducted in partial fulfillment of the Ph.D. requirement of the University of Rochester for E.M.F.)
S. mutans UA159 and its derivatives were maintained on brain heart infusion (BHI; Difco) agar plates. Organisms were grown on solid medium at 37°C in atmosphere containing 5% (vol/vol) CO2-95% air. Cultures were grown in TY medium (3% tryptone, 0.1% yeast extract, 0.5% KOH, 1 mM H3PO4). Antibiotics were added to the following final concentrations for S. mutans: erythromycin, 5 μg ml−1; and kanamycin, 1 mg ml−1. cis-Vaccenic acid (C18:1) or cis-eicosenoic acid (C20:1) was added as indicated to a final concentration of 10 μg ml−1 (Sigma).
Escherichia coli was maintained on Luria-Bertani agar and supplemented when needed with the following antibiotics: erythromycin, 500 μg ml−1; kanamycin, 50 μg ml−1; tetracycline, 10 μg ml−1; and ampicillin, 100 μg ml−1 (Sigma).
Chromosomal DNA was isolated from S. mutans as previously described (34). Small-scale DNA preparations were also utilized (15). Plasmid DNA from E. coli was isolated by rapid plasmid preps (11) or with the QIAprep Spin Miniprep kit (QIAGEN, Chatsworth, Calif.). PCR was carried out with Vent DNA polymerase (New England Biolabs). PCR products were isolated and purified via gel electrophoresis as previously described (22). Restriction and DNA-modifiying enzymes were purchased from Invitrogen (Gaithersburg, Md.). E. coli DH10B was transformed by electroporation (22); S. mutans UA159 was transformed by the method of Murchison (29). Colony hybridization of E. coli was performed by standard techniques (34). Southern analysis was performed as previously described (34).
Primers were designed to amplify the predicted fabM coding sequence (792 bases) plus additional flanking regions from the UA159 genome (2). Splice-overlap extension PCR (20) was performed to insert a unique BglII site at position +396 of the predicted coding region to allow for insertion of an antibiotic marker. PCR primers (Invitrogen) were designed as follows to amplify the 5′ 530-bp portion of the coding region and contained restriction sites as underlined to aid in cloning: NsiFwd (5′ ATAAAATGTATGCATTGGAACTTTGATTTTCAAAC 3′) and BglRev (5′ CATTCACAAAGGCTTGAATAAGATCTGTTTTGTCAC 3′). A 605-bp PCR product containing the carboxyl terminal of the coding sequence and flanking region was generated with the following primers: BglFwd (5′ GTGACAAAACAGATCTTATTCAAGCCTTTGTGAATG 3′) and EcoRev (5′ CACCCACAAAATAGAATTCAATTTATAAGATTACGTA 3′). The two PCR products from the above reaction mixtures were combined (10 ng each), and using the primers NsiFwd and EcoRev, a 1,135-bp amplicon was generated. This resulting product contained a unique BglII site at position +591 (position +396 in the predicted fabM coding sequence). The fabM PCR product was digested with EcoRI and NsiI and cloned into the compatible EcoRI and PstI sites of pBR322, replacing part of the ampicillin cassette (37). E. coli DH10B was transformed with the ligation mix, and transformants were selected by their ability to grow on tetracycline-containing medium. Positive clones were confirmed by sequencing at the Functional Genomics Center at the University of Rochester Medical Center. The resulting plasmid, pBRB3, was linearized with BglII and ligated to an erythromycin cassette from pTS19E (4) containing compatible BamHI ends. This new construct, pBRBErm 4, was used to transform E. coli, and transformants were selected by their ability to grow on erythromycin-containing medium and confirmed by restriction digestion.
UA159 was transformed with pBRBErm 4, and transformants were selected on BHI medium containing erythromycin. Putative mutant strains were confirmed by Southern hybridization. One such strain was designated as S. mutans UR117.
To confirm that the observed phenotype was due to disruption of fabM and not due to downstream polar effects, a fabM complement strain was generated. A 1,290-bp PCR product was amplified from wild-type UA159 genomic DNA containing 430 bases upstream of the predicted start codon and an additional 68 bases downstream of the predicted stop codon for fabM. Primers were designed as follows and contained SmaI sites as underlined: FabMSmaFwd (5′ GATTATTTTGACCCGGGTTTATCGGGAG 3′) and FabMSmaRev (5′ GCACAATAAAAACCCGGGACATTTTTGTCCC 3′). The resulting PCR product was cloned into the unique SmaI site of pBGK, an S. mutans vector that integrates into the gtfA locus (39). E. coli DH10B was transformed with this ligation mix, and transformants were selected on Luria-Bertani medium contain kanamycin. Positive clones were identified by colony hybridization (34) and subsequent SmaI digestions. Nucleotide sequencing was performed to verify the clone, and the resulting plasmid was designated pMC4-10. S. mutans UR117 was transformed with pMC4-10, and transformants were selected on kanamycin-containing medium. Southern hybridization was performed to confirm integration of pMC4-10 (39), and one such strain was designated as UR119.
The membrane fatty acid content of the cultures was determined by Avanti Polar Lipids, Inc. (Alabaster, Ala.) as previously described (19, 33). Briefly, 100 ml of cultures was grown overnight in TY supplemented with 1% glucose, the appropriate antibiotic, and 10 μg of the indicated fatty acid ml−1. Cultures were harvested in the morning by centrifugation. The cells were washed with 25 ml of deionized water and recollected by centrifugation, and pellets were stored at −80°C prior to fatty acid analysis. Two independent cultures per strain or growth condition were assayed.
Total lipids (approximately 5 mg obtained per sample) were extracted by the method of Bligh and Dyer (12). Fatty acid esters were prepared through the addition of 0.2 ml of toluene and 0.4 ml 1% H2SO4 in methanol and heated for 30 min. Fatty acids were extracted via the addition of 1 ml of hexane and 1 ml of H2O. The hexane phase containing the fatty acid esters was evaporated under nitrogen gas. The fatty acid methyl esters were reconstituted in hexane, and gas chromatography was performed, using nitrogen as a carrier gas. Nu-Chek Prep standard 68A was used to determine retention times and then used to determine the identity of fatty acids derived from the strains.
An established acid sensitivity assay (5, 19) was utilized to determine whether the inability to produce monounsaturated membrane fatty acids affected the low-pH survival capabilities of S. mutans. Overnight cultures of S. mutans UA159, UR117, and UR117 supplemented with 10 μg of either cis-vaccenic acid (C18:1) or cis-eicosenoic acid (C20:1) ml−1 and UR119 were harvested by centrifugation at 2,500 × g for 10 min. Cell pellets were resuspended in 3 ml of 0.1 M glycine-HCl (pH 2.5) and were continuously stirred for 1 h at room temperature. Aliquots (0.1 ml) of the cell suspension were removed at 0, 15, 30, and 60 min; serially diluted into BHI medium (Difco); and plated in duplicate on BHI agar. UA159 and UR119 plates were incubated 48 h at 37°C, 5% CO2-95% air. All UR117 plates were incubated for 72 h at 37°C in 5%CO2-95% air. Viable cell counts were enumerated and used to calculate log (N/N0).
Proton permeabilities were determined by a previously established protocol (9, 23). Briefly, 200-ml cultures of S. mutans UA159, UR119, UR117, and UR117 supplemented with either 10 μg of cis-vaccenic acid (C18:1) or cis-eicosenoic acid (C20:1) ml−1 were grown overnight in TY broth with 1% glucose and were harvested by centrifugation for 10 min at 9,000 × g at 4°C. The cells were washed once with 50 ml of 5 mM MgCl2. Cells were suspended at 5 mg ml−1 in 20 mM potassium phosphate buffer (pH 7.2)-50 mM KCl-1 mM MgCl2 and incubated for 2 h at 37°C in 5%CO2-95% air. Cells were harvested by centrifugation following this starvation period and resuspended to 20 mg ml−1. A 1-ml aliquot of each suspension was titrated to pH 4.7 with either 100 mM HCl-50 mM KCl or 10 mM HCl-50 mM KCl, depending on the pH of the cell suspension. The pH of the cells was then quickly lowered by approximately 0.2 pH units by addition of 10 mM HCl-50 mM KCl. pH values were recorded over time. After 50 min, butanol was added to 10% (vol/vol) to disrupt the cell membranes and allow for equilibrium between the external and cytoplasmic pHs. pH equilibrium was reached 80 min after the start of the experiment. Using the recorded pH values, an estimate of ΔpH over time could be determined by calculating the pH difference between the time of interest and the 80-min end point. The ΔpH at 50 min (immediately before butanol addition) is reported in the text. The experiment was done in duplicate with two independent cultures from each strain or growth condition.
Two-hundred-milliliter cultures were grown overnight in TY with 1% glucose plus the appropriate antibiotic and 10 μg of the indicated fatty acid ml−1. The strains were permeabilized as described previously (22) with the following modifications. Briefly, the strains were divided into two 100-ml aliquots and harvested by centrifugation. One aliquot was used to determine dry weight; the other was washed once in membrane buffer (75 mM Tris, pH 7, 10 mM MgSO4). These cells were resuspended in 4 ml of membrane buffer and treated with a 1% final volume of toluene. The suspensions were vortexed for 30 s, placed on ice for 5 min, and subjected to 2 rounds of freeze-thaw. The permeabilized cells were collected by centrifugation, resuspended in 4 ml of membrane buffer, and stored as 100-μl aliquots at −80°C for up to 1 month.
Glycolytic minima were determined as described previously (6, 16) with the following modifications. Briefly, overnight cultures were harvested by centrifugation and washed twice with a 50 mM KCl-1 mM MgCl2 salt solution. The cells were resuspended in the salt solution to a final concentration of 10 mg ml−1. Aliquots (5 ml) were titrated to a pH of 7.2 by addition of 0.5 M KOH. Glucose was added to a final concentration of 1% (wt/vol), and the pH was monitored continuously for 2 h. After the pH stabilized, approximately 2 h after the addition of glucose, the minimal glycolytic pH was recorded. The results of these experiments represent three independent overnight cultures, each assayed in triplicate.
Glucose-specific PTS assay conditions were as stated previously (6, 23). Fifty microliters of permeabilized cells was incubated at 37°C for 10 min in 100 mM Tris-maleate (pH 7), 20 mM MgCl2, 40 mM glucose, and 1 mM NaF. Reactions were initiated by the addition of 100 μl of 50 mM phosphoenolpyruvate (PEP). At 0 and 30 min postaddition of PEP, 500-μl aliquots were chilled on ice to stop the reaction. The suspensions were cleared, and the supernatants were assayed for pyruvate content. Two hundred microliters of cleared supernatant was added to 300 μl of double-distilled H2O-500 μl of NADH solution (0.21-mg/ml NADH, 1.5 M Tris, pH 7, 0.021% NaN3). The A350 was recorded. Lactate dehydrogenase (LDH) at 3 μl (3 U/μl) was added, and the mixture was incubated for 5 min. A340 decreases due to the utilization of NADH by LDH. The change in A340 of the 30-min sample versus the zero time point reflects the amount of pyruvate produced and thus the amount of glucose phosphorylated. The data are presented as millimoles of pyruvate produced per minute per milligram of cell dry weight. Results of these experiments reflect data from three independent overnight cultures assayed three times each in triplicate.
A previously described ATPase assay (31) was used to measure the release of inorganic phosphate from ATP in permeabilized cells. Thirty microliters of permeabilized cells was incubated with 50 mM Tris-maleate (pH 6)-10 mM MgSO4 for 20 min. ATP was added to final concentration of 5 μM. Released phosphate was measured by the method of Bencini (7, 8) and is expressed as millimoles of Pi per minute per milligram of cell dry weight.
Student's t test was used to evaluate data in the experiments involving proton-permeability, PTS assays, and ATPase assays.
Nucleotide sequence accession numbers. Nucleotide sequence data reported are available in the Third Party Annotation Section of the DDBJ/EMBL/GenBank databases under the accession number TPA: BK005411.
Previously, we had demonstrated that the membrane fatty acid profile of S. mutans shifts to include higher percentages of monounsaturated fatty acids when cultures are grown under acidic conditions (19, 33). If the shift to monounsaturated membrane fatty acids was blocked, the organism was rendered more acid sensitive (19). However, the mechanism responsible for the synthesis of the monounsaturated fatty acids was unclear. Recent work with Streptococcus pneumoniae has led to the identification and characterization of a novel trans-2, cis-3-decenoyl-ACP isomerase designated as FabM (24). This enzyme is capable of generating unsaturated fatty acids (24). Using the sequence of the S. pneumoniae FabM, we performed a BLAST search (3) against the completed S. mutans UA159 genome (2) from the Advanced Center for Genome Technology at the University of Oklahoma. The search identified an open reading frame, phaB, which was predicted to encode an enoyl-coenzyme A (CoA) hydratase. A ClustalW alignment of PhaB to the S. pneumoniae FabM is shown in Fig. Fig.1A.1A. It was interesting to note that the genetic organization surrounding S. mutans phaB (which was renamed fabM to follow convention) is remarkably similar to that of the gene organization surrounding S. pneumoniae fabM (24) (Fig. (Fig.1B).1B). This observation provided the impetus for our attempt to insertionally inactivate S. mutans fabM, which we hypothesized to be involved in the pH-dependent shift to a monounsaturated fatty acid membrane profile.
An isogenic mutant strain was generated by the insertion of an erythromycin resistance cassette approximately 400 bp downstream of the predicted start codon of S. mutans UA159 fabM. Standard techniques were utilized to identify a mutant strain, designated UR117 (Materials and Methods). The strain, UR117, exhibited a small-colony phenotype, unlike the wild-type strain, UA159 (data not shown). In addition, UR117 exhibited an increased doubling time compared to the parental strain (Table (Table1).1). However, the growth defect could be overcome by supplementing the media with either cis-vaccenic acid (C18:1) or cis-eicosenoic acid (C20:1), the principle monounsaturated fatty acids found in the membrane of UA159 (19, 33). In addition, a complemented UR117 strain, designated UR119, exhibited a similar doubling time to the wild-type strain (Table (Table11).
To confirm that fabM was essential for the production of monounsaturated membrane fatty acids, the total membrane fatty acid content of S. mutans UR117 was determined (Materials and Methods). As shown in Table Table2,2, the mutant strain, UR117, did not have detectable levels of monounsaturated fatty acids, unlike the parental strain UA159 or UR119, the fabM complement. If cultures of the mutant strain were grown in the presence of either cis-vaccenic or cis-eicosenoic acid overnight, the exogenous fatty acids were incorporated into the membrane (Table (Table2).2). Each exogenously supplied fatty acid was incorporated equally well into the membrane of UR117, with each comprising approximately 20% of the total membrane composition. Although UR117 integrated the exogenously supplied monounsaturated fatty acids, it did not elongate either C18:1 or C20:1, as evident by the membrane fatty acid analysis. Thus, by insertionally inactivating fabM, the organism was unable to generate monounsaturated fatty acids.
Previously, when the pH-dependent membrane acid shift was blocked by cerulenin, the organism was rendered more sensitive to extreme acidification (19). Since this suggested that monounsaturated membrane fatty acids are important for acid survival, UR117 was predicted to be more sensitive to acidification. Also, because the mutant strain was capable of incorporating exogenous fatty acids into its membrane (38) (Table (Table2),2), we hypothesized that supplementing UR117 with monounsaturated fatty acids would lead to improved acid resistance.
In response to acid challenge, wild-type cells were still viable 60 min after treatment with 0.1 M glycine-HCl, pH 2.5 (Fig. (Fig.2).2). However, UR117, which did not produce detectable levels of monounsaturated membrane fatty acids, yielded no colonies beyond 15 min of treatment. When UR117 was supplemented with either cis-vaccenic acid (C18:1) or cis-eicosenoic acid (C20:1), survivors were observed after 60 min of low-pH treatment. UR119, which had a membrane composition similar to UA159 (Table (Table2),2), was also viable after acid challenge. Thus, UR117, which did not produce monounsaturated membrane fatty acids, was more susceptible to extreme acidification than the wild-type organism, and this acid-sensitive phenotype could be relieved through growth of the strain in the presence of exogenous monounsaturated fatty acids or by complementation.
UA159 produces organic acid metabolically and is capable of surviving low-pH environments (35). Interestingly, we observed that UR117 cultures did not lower the final pH of their growth media to the same extent as wild-type cultures (data not shown). We hypothesized that if the mutant strain was deficient in acid production, this would account for the higher final pH value of UR117 cultures in comparison to UA159. Therefore, the glycolytic capability of UR117 was examined (Materials and Methods).
The wild-type strain (UA159) and UR119, the complementation strain, lowered the pH of the medium to an approximate value of 3.7 30 min after the addition of glucose (Fig. (Fig.3).3). UR117, at the same time point, had only lowered the pH of the medium to a value of 4.8. However, if UR117 cultures were grown in the presence of exogenous monounsaturated fatty acids, the organism reached a pH value of 4.1 after 30 min, lower than that of the UR117 cultures grown without the addition of monounsaturated fatty acids.
Glycolytic minima were recorded 2 h following the addition of glucose, after the pH of the suspensions had stabilized. The average minimum recorded for S. mutans UR117 was a pH of 4.2 ± 0.1, which was statistically different from the value observed for UA159, a pH value of 3.3 ± 0.1. The complement of UR117, UR119, was capable of glycolysis at a much lower pH than the mutant strain, 3.2 ± 0.2. When supplemented with 10 μg of cis-vaccenic acid (C18:1) ml−1 in its growth medium, UR117 was capable of glycolysis to a pH of 3.5 ± 0.2. A similar minimal pH of 3.6 ± 0.1 was obtained when UR117 was grown in the presence of 10 μg of cis-eicosenoic acid (C20:1) ml−1. Thus, the altered membrane profile did have effects on the glycolytic capability of UR117.
Growth of S. mutans in the presence of exogenously supplied fatty acids has been shown to impact glucose-specific PTS activity (23). This observation was likely due to the incorporation of the exogenous fatty acids into the membrane leading to altered membrane compositions (38), which have, consequently, affected the activities of membrane-bound proteins. The altered membrane composition of UR117 may also have led to changes in glucose-specific PTS activity, which could have impacted the ability of the organism to perform glycolysis under low-pH conditions. In order to address this possibility, we examined the effect of the fabM mutation on glucose-specific PTS activity by using overnight cultures grown in the presence of glucose and 10 μg of either cis-vaccenic acid or cis-eicosenoic acid ml−1 (Materials and Methods).
Surprisingly, despite the absence of monounsaturated membrane fatty acids in S. mutans UR117, glucose-PTS activity in this strain was higher than that observed in UA159 or UR119 (Table (Table3).3). The activity observed in UR117 was 2.4-fold-higher than that seen in UA159. Supplementing cultures of UR117 with exogenously-supplied fatty acids led to various degrees of activity; UR117 grown in the presence of cis-vaccenic acid (C18:1) had roughly 1.4-fold-higher activity than UA159. When grown in cis-eicosenoic acid (C20:1), UR117 had glucose-specific PTS activity approximately twofold higher than that observed in the wild type. These results suggested that the altered membrane fatty acid composition did not decrease the ability of UR117 to take up glucose and that glucose uptake was not likely responsible for the inability of UR117 to perform glycolysis at pH values as low as those observed for the wild type.
Previous work has demonstrated that S. mutans possesses a membrane that is rather impermeable to protons (9) and that growth in exogenously supplied fatty acids can cause alterations in the permeability of S. mutans to protons (23). Because the mutant strain did not survive acidification of its environment well (Fig. (Fig.2),2), it suggested that the altered membrane fatty acid composition may have affected the permeability of the membrane to protons. If the mutant strain did have an altered proton permeability and could not maintain a normal ΔpH across its membrane, it could account for the observed acid sensitivity phenotype.
We measured the proton permeabilities of cultures of S. mutans strains UA159, UR119, UR117, and UR117 grown in the presence of 10 μg of cis-vaccenic acid or cis-eicosenoic acid ml−1 (Materials and Methods). The rise in external pH of UR117 was not as sharp as that seen in UA159 (Fig. (Fig.4);4); thus, the membrane of the mutant was more impermeable to protons. However, when the cells were permeabilized by the addition of 10% butanol at 50 min, UR117 did not reach as high a final pH value as the wild type. UR117 maintained an estimated ΔpH at 50 min of −0.5 ± 0.04, whereas ΔpH of UA159 was −0.8 ± 0.01 (statistically different at P < 0.0001). Thus, it appears that the ΔpH of the mutant strain was not as great as in the wild-type strain.
When UR117 was grown in the presence of exogenous monounsaturated fatty acids, the results obtained were dependent upon the fatty acid supplied. As shown, when grown in the presence of cis-vaccenic acid (C18:1), UR117 gave a permeability profile similar to that seen with the wild type, UA159; the ΔpH at 50 min was −0.7 ± 0.03, statistically different (P < 0.0009) from that observed with UR117 grown without supplementation (Fig. (Fig.4).4). However, when grown in the presence of cis-eicosenoic acid (C20:1), the permeability profile of UR117 was similar to that of unsupplemented cultures of UR117, with a very slow rise in the external pH. Likewise, the ΔpH of UR117 grown in the presence of cis-eicosenoic acid was not as large as seen with UA159; the supplemented strain at 50 min had a ΔpH of −0.6 ± 0.2, smaller than that observed with UA159.
The rise in the pH of the complement, UR119, was similar to that seen in UA159. However, the ΔpH at 50 min was only −0.6 ± 0.02, which was smaller than that observed with the wild type. It appears that despite genetic complementation of the mutant, there do exist some differences between the measured proton permeabilities of UR119 and UA159, which may be due to slight alterations in membrane fatty acid composition.
One complication involved with measuring proton permeabilities is that the assay does not separate the proton-pumping activity of the F1F0 ATPase from the diffusion of protons across the membrane. The ΔpH of S. mutans is maintained through the activity of this enzyme (9, 35). Thus, the observed altered proton permeability and ΔpH of the mutant may be due to differences in the ATPase activity levels.
We found that the ATPase activity levels in UA159 and UR119 were comparable and not statistically different (Table (Table4).4). The mutant strain UR117, however, showed significant differences in ATPase activity as compared to the wild type, with the fabM strain having approximately twofold-higher levels of activity. Supplementing UR117 with exogenous monounsaturated fatty acids led to lower ATPase activity levels in comparison to those in unsupplemented cultures, although they were above the wild-type levels. Thus, the altered membrane fatty composition of UR117 did affect the activity of F1F0 ATPase.
Membrane fatty acid and phospholipid adaptation in bacteria in response to environmental stresses has been explored most extensively in E. coli, B. subtilis, Bacillus stearothermophilus, and Acholeplasma laidlawii (1, 10, 14, 17, 27, 28). Until recently, the phenomena had not been explored as extensively in S. mutans, an organism that inhabits a low-pH environment, dental plaque. Our interests have been focused on elucidating whether changes in membrane fatty acid composition occur (33) and how and when they may occur (19). From these studies, it was observed that the membrane composition of S. mutans is altered in response to acidification; this shift can occur through the self-generation of acid by the organism, and it can be blocked by the fatty acid biosynthesis inhibitor cerulenin (19). However, the mechanism by which the changes occurred was unclear.
Using results from a recent, elegant study in which the biochemical activity of FabM in S. pneumoniae was examined (24), we were able to identify and disrupt the homologue in S. mutans UA159. The resultant mutant strain was extremely acid sensitive, but the mechanism by which monounsaturated fatty acids protect against acid is not yet understood. It may be possible that unsaturated bonds serve as a sink for protons; this, in conjunction with the activity of the F1F0 ATPase, could serve to protect the cytoplasmic contents from damage during growth under low-pH conditions.
S. mutans must survive low-pH environments because it produces acid via its metabolism of carbohydrates. The inability of the UR117 strain to metabolize glucose at pH values as low as those observed for UA159 may prevent the organism from acid adapting as fully as the wild type; thus, this may render the mutant strain more sensitive to extreme acidification. The simple addition of monounsaturated fatty acids to the growth medium of UR117 increased its ability to carry out glycolysis at low pH levels, most likely due to the incorporation of the fatty acids into the membrane.
Differences in glucose-specific PTS activity were expected, as it has been shown that the presence of exogenously supplied fatty acids can alter the PTS activity of S. mutans (23). Enzyme IIC components of PTS are in the membrane; thus, shifting membrane composition could alter protein interactions and, consequently, affect activity. Despite increased glucose-specific PTS activity in UR117, these cultures were unable to perform glycolysis at pH values as low as UA159. The decreased glycolytic ability of UR117 is likely due to other enzymes further downstream of glucose transport.
The enhanced ATPase activity observed in UR117 was unexpected. The activity of F1F0 ATPase was higher in cells grown under low-pH conditions, where the membrane composition would have larger amounts of monounsaturated fatty acids (5, 19, 33). In addition, we and others have shown evidence of transcriptional regulation of F-ATPase in streptococci during growth at low pH (21, 26). It is possible that the effect of a narrower ΔpH in UR117 had led to disregulation of the F-ATPase operon. We are currently investigating this possibility.
Differences in the membrane fatty acid composition may have additional effects on the phospholipid composition, peptidoglycan production, and other forms of membrane modifications. In addition, the effects of incorporation of exogenous fatty acids (in nutritionally supplemented strains) on other membrane components and the impact of this on de novo fatty acid biosynthesis, as well as other metabolic activities, are not completely understood. The presence of a putative transcriptional regulator, Smu1592 (2), immediately upstream of fabH leads to speculation of possible fatty acid biosynthesis regulation, which may occur when the organism is grown in an environment rich in fatty acids. Thus, many questions regarding the regulation of fatty acid biosynthesis and how membrane composition alterations may affect other membrane constituents need to be explored further to completely understand our observations.
Currently, our database searches as well as those done previously (24) have indicated that FabM homologues exist only in streptococcal and staphylococcal species, with the exception of a homologue found in the gram-negative bacterium Fusobacterium nucleatum (data not shown). Several recent articles have suggested the development of antibiotics to target fatty acid biosynthesis (13, 25, 30, 32). Bacterial fatty acid biosynthesis is classified as type II, in that each reaction is carried out by a separate enzyme. The fatty acid biosynthesis of eukaryotes, however, is classified as type I, in that all the reactions are carried out by a large enzyme complex (18, 25). The differences between the two fatty acid biosynthesis systems might be exploited, in that new drugs could be designed to target FabM enzymes (i.e., bacterial) that would probably not affect the enzymes used in eukaryotic cells. Thus, there is potential use in developing therapies that target ACP-isomerases, such as FabM.
We wish to thank Robert Marquis for helpful discussion throughout. We also thank Roberta Faustoferri and Matthew Betzenhauser for technical expertise.
This work was supported by grants from the NIH, National Institute for Dental and Craniofacial Research DE-11549 and DE-01627. E.M.F. was supported by the Rochester Training Program in Oral Infectious Diseases, T32-DE07165.