JOURNAL OF BACTERIOLOGY, Feb. 1990, p. 556-563
Vol. 172, No. 2
0021-9193/90/020556-08$02.00/0
Characterization of Transmembrane Movement of Glucose and Glucose Analogs in Streptococcus mutans Ingbritt STUART G. DASHPER AND ERIC C. REYNOLDS* Biochemistry and Molecular Biology Unit, Faculty of Medicine and Dentistry, University of Melbourne, 711 Elizabeth Street, Melbourne, 3000, Australia Received 1 August 1989/Accepted 23 October 1989
The transmembrane movement of radiolabeled, nonmetabolizable glucose analogs in Streptococcus mutans Ingbritt was studied under conditions of differing transmembrane electrochemical potentials (A*) and pH gradients (ApH). The ApH and A* were determined from the transmembrane equilibration of radiolabeled benzoate and tetraphenylphosphonium ions, respectively. Growth conditions of S. mutans Ingbritt were chosen so that the cells had a low apparent phosphoenolpyruvate (PEP)-dependent glucose:phosphotransferase activity. Cells energized under different conditions produced transmembrane proton potentials ranging from -49 to -103 mV but did not accumulate 6-deoxyglucose intracellularly. An artificial transmembrane proton potential was generated in deenergized cells by creating a A* with a valinomycin-induced K+ diffusion potential and a ApH by rapid acidification of the medium. Artificial transmembrane proton potentials up to -83 mV, although producing proton influx, could not accumulate 6-deoxyglucose in deenergized cells or 2-deoxyglucose or thiomethylgalactoside in deenergized, PEP-depleted cells. The transmembrane diffusion of glucose in PEP-depleted, KF-treated cells did not exhibit saturation kinetics or competitive inhibition by 6-deoxyglucose or 2-deoxyglucose, indicating that diffusion was not facilitated by a membrane carrier. As proton-linked membrane carriers have been shown to facilitate diffusion in the absence of a transmembrane proton potential, the results therefore are not consistent with a proton-linked glucose carrier in S. mutans Ingbritt. This together with the lack of proton-linked transport of the glucose analogs suggests that glucose transmembrane movement in S. mutans Ingbritt is not linked to the transmembrane proton potential.
The oral bacterium Streptococcus mutans and dietary sugar have been implicated in the development of dental caries (7, 26). Fundamental to the existence of S. mutans in dental plaque is its ability to compete effectively for sugar substrates under differing environmental conditions. Competition is at the level of cellular transport. It is well established that S. mutans transports glucose, 2-deoxyglucose, sucrose, lactose, mannose, and fructose by a phosphoenolpyruvate (PEP)-dependent sugar:phosphotransferase system (PTS) (4, 8, 12, 25, 42). Substrate specificity is conferred by the membrane-bound enzymes II, with each protein recognizing a series of structurally related sugars (36, 41). A second glucose transport system in S. mutans has been proposed by Hamilton and St. Martin (9) and Keevil et al. (22) to involve a carrier-mediated proton-glucose symport driven by the transmembrane electrochemical proton potential (Ap) or proton motive force (PMF) (33). Keevil et al. (22) have suggested that sugar transport in S. mutans at the low pH of cariogenic plaque would occur predominantly by this PMF-driven transporter, and therefore, this system could be an important virulence determinant. However, the evidence to support the existence of a PMF-driven glucose transporter is equivocal and is based largely on the findings that acid production from glucose by S. mutans is inhibited by certain PMF-dissipating ionophores and that the glucose-PTS activity of toluenized cells is not commensurate with the rate of acid production by whole cells with glucose as the substrate. To interpret these results as evidence for a PMF-driven glucose transporter, it must be assumed that glucose uptake is rate limiting for glycolysis, that the PTS activity of toluenized cells is a quantitative measure of whole-cell *
activity, and that ionophores do not affect the PTS or glycolysis through perturbation of intracellular H+ and K+ concentrations. Transport of a nonmetabolizable glucose analog in S. mutans Ingbritt has been investigated by Keevil et al. (21). They used radiolabeled 6-deoxyglucose, which is an ideal substrate as it cannot be actively translocated by the glucose-PTS. However, the results of this study were again equivocal owing to the low level of accumulation and the high level of cellular adsorption of the radiolabel. Further, the accumulation of 6-deoxyglucose was not correlated with Ap nor was a link between proton and 6-deoxyglucose transmembrane movement demonstrated. In an approach to characterize glucose transport in S. mutans Ingbritt, we attempted to demonstrate a PMF-driven glucose transport system by correlating intracellular accumulation of radiolabeled, nonmetabolizable glucose analogs with Ap. The relative contributions of the PMF-driven system and the PTS were then to be determined under different environmental conditions. However, proton movement across the cell membrane did not result in an intracellular accumulation of 6-deoxyglucose or 2-deoxyglucose. Further, the diffusion of ['4C]glucose in PEP-depleted, metabolically poisoned cells did not exhibit saturation kinetics, indicating the absence of a membrane carrier capable of facilitating diffusion. The results therefore are not consistent with a PMF-linked glucose carrier in S. mutans Ingbritt. MATERIALS AND METHODS Bacterial strains. S. mutans Ingbritt (24) and variant 162 (3) were kindly supplied by B. Krasse (Karolinska Institutets, Stockholm, Sweden) and K. Knox (Institute of Dental Research, Sydney, Australia), respectively. The bacteria were stored as lyophilized cultures in sealed ampoules or in
Corresponding author. 556
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30% glycerol broth at -20°C. The strains were checked regularly according to the criteria of Hardie and Bowden (10). Growth conditions. Bacteria were grown as stationary batch cultures or in continuous culture at 37°C. The growth medium for the stationary batch cultures contained 1% (wt/vol) tryptone, 0.5% (wt/vol) yeast extract, 1.62 mM MgSO4, 0.04 mM FeSO4, 0.06 mM MnCl2, 9.70 mM KH2PO4, 15.31 mM K2HPO4, 35 mM KCl, and 55 mM glucose and was adjusted to pH 7.0. The glucose and the phosphate salts were individually filter sterilized and added to the medium before inoculation. Batch-grown cells were harvested in the mid-exponential phase. Continuous cultures were grown in a BioFlo model C-30 chemostat (New Brunswick Scientific Co., Inc., Edison, N.J.) with a working capacity of 365 ml. The temperature was maintained at 37°C, and the pH was controlled at 7.0 or 5.5 as specified by the automatic addition of 5 M KOH. Cultures were gassed with 5% CO2 in nitrogen. The dilution rate was set at D = 0.1 h-1, which is equivalent to a mean generation time of 6.9 h, and cultures were allowed to equilibrate for 10 generations before sampling. The growth medium was the same as that for the batch cultures except that the glucose was 11 mM, which was limiting. Culture purity was checked regularly by microscopic examination and by the criteria of Hardie and Bowden (10). Chemostatgrown cells were collected from the overflow at 4°C for less than 1 h. Chemostat- or batch-grown cells were centrifuged at 1,000 x g for 15 min at 4°C and washed twice with fermentation minimal medium (FMM) containing 50 mM KCl, 5 mM NaCl, 2 mM MgSO4, 2 mM MnCl2, 8 mM (NH4)2SO4, 1.5 ,uM thiamine, and 8 p,M niacin at pH 7.0. The cells were resuspended in FMM to give a cell density of 1.25 mg (dry weight) of cells per ml. Measurement of glycolytic activity. Glycolytic activity was determined in a pH stat (Radiometer Copenhagen, Copenhagen, Denmark) essentially as described by Marsh et al. (32), using a volumetric standard titrant (0.1 M KOH) and a 10 mM final glucose concentration, which was saturating (39). Before the addition of the prewarmed glucose solution, the cells were depleted of endogenous reserves of carbohydrate by preincubation at pH 7.0 in the pH stat for approximately 22 min. Glycolytic activity was measured as nanomoles of H' neutralized per milligram (dry weight) of cells per minute. The dry weight of cells was determined by filtering 1-ml samples of the cell suspension through dry, preweighed, 0.2-,um-pore-size polycarbonate filters (Nuclepore Corp., Pleasanton, Calif.) supported on Whatman GF/ A filters. The ionophore gramicidin D (Sigma Chemical Co., St. Louis, Mo.) was added as a 10 mM solution in 95% ethanol to give a final concentration of 10 p.M. All controls had equivalent amounts of ethanol added, which had no effect on acid production. PTS assay. The PEP-dependent glucose-PTS was assayed by a modification of the method of Kornberg and Reeves (23). Batch- or chemostat-grown cells were centrifuged at 1,000 x g for 10 min at 4°C. The cell pellet was washed twice in 100 mM potassium phosphate buffer (pH 7.2) containing 5 mM dithiothreitol and then resuspended in the same buffer to a cell density of 10 mg (dry weight) of cells per ml. Tolueneacetone (1:9) was used to permeabilize the cells. The PTS assay reaction mixture contained 100 mM potassium phosphate buffer (pH 7.2), 5 mM MgSO4, 5 mM PEP, 5 mM dithiothreitol, 10 mM KF, 0.3 mM NADH (Sigma), 5 U of
557
L-lactic dehydrogenase (EC 1.1.1.27; Sigma) per ml (700 to 1,400 U/mg of protein), and 0.1 mg (dry weight) of cells. Intracellular water volume determination. Intracellular water volume was measured by a modification of the procedure of Maloney et al. (29) with [1,2-3H]taurine as the extracellular marker (15). Batch-grown cells were washed once in 0.1 M Tris hydrochloride (pH 7.2)-100 mM KCl and resuspended in the same buffer to give a cell density of 4.0 mg (dry weight) of cells per ml. [1,2-3H]taurine (50 p.Ci/pmol) was added to the cell suspension to give a final concentration of 100 ,uM. Samples were filtered through dry, preweighed, 0.2-p.m-pore-size polycarbonate filters (Nuclepore) without washing. Total water on the filter was determined gravimetrically, and radioactivity was determined after filter solubilization (Amersham Australia Pty. Ltd.). Filtrates were also collected, and their radioactivity was measured. The polycarbonate filters retained only approximately 30% of the total water as extracellular water, which allowed an accurate determination of total, extracellular, and intracellular water by the [3H]taurine-gravimetric method. In fact, the [3H] taurine-gravimetric method proved to be slightly more reproducible (coefficient of variation, 4.1%) than the ['4C]taurine-3H20 method (coefficient of variation, 8.8%). The intracellular water volume of S. mutans Ingbritt from four separate determinations was 1.69 ± 0.07 ,ul/mg (dry weight) of cells. This compares well with the value obtained by Noji et al. (34) of 1.60 ± 0.14 p.l/mg (dry weight). Uptake of radiolabeled sugar analog in endogenously energized cells. Washed cells were suspended in 4 ml of either 100 mM Tris-citrate buffer (pH 7.0) or FMM containing 5 mM dithiothreitol to give a cell density of 5 mg (dry weight) per ml and used without preincubation. Radiolabeled sugar analog (['4C]thiomethylgalactoside (['4C]TMG), 1.0 Ci/mol; 6-deoxy[3H]glucose, 0.5 Ci/mol; or 2-deoxy['4C]glucose, 1.0 Ci/mol) was added to the cell suspension in a pH stat (pH 7.0, 37°C) to give a final concentration of 4 mM. Samples were taken at the times indicated and filtered through dry, preweighed, 0.2-p.m-pore-size polycarbonate filters without washing. Intracellular and extracellular water volumes and filter and filtrate radioactivities were determined as described above. The amount of intracellular radiolabeled sugar was determined by subtracting the extracellular radioactivity from the total radioactivity on the filter. A control (10 puM gramicidin-treated cells) was used to determine non-PMF uptake or cellular adsorption or both. Uptake of radiolabeled sugar analog after an imposed membrane potential (AO. A membrane potential was imposed by valinomycin treatment of K+-loaded, deenergized cells in low-K+ medium as described by Kashket and Wilson (18) and Maloney (27, 28). Washed cells were suspended in 10 mM K+-FMM or 100 mM taurine-25 mM choline chloride-25 mM KCl (pH 7.2) and preincubated in a pH stat (pH 7.0, 37°C) until acid production ceased. Prior to the [14C]TMG uptake experiments, 2-deoxyglucose was added to the cell suspension to give a final concentration of 1 mM and incubated for 20 min to deplete intracellular PEP. Similarly, prior to the 2-deoxy[14C]glucose uptake experiments, TMG was added to a final concentration of 1 mM and incubated for 20 min. After PEP depletion, the cells were washed once in 100 mM taurine-50 mM choline chloride-0.3 mM KCl (pH 6.5) and then resuspended in the same buffer to give a cell density of 5 mg (dry weight) per ml. The radiolabeled sugar analog 6-deoxy[3H]glucose (0.5 p.Ci/
p.mol), 2-deoxy[14C]glucose (1.0 p.Ci/p.mol), or [14C]TMG (1.0 p.Ci/p.mol) was added to the deenergized, PEP-depleted cells to give a final concentration of 4 mM, and after
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15 min, valinomycin (Sigma) was added to 10 p.M. Uptake measured as described above. Valinomycin was added 10 mM solution in 95% ethanol. Uptake of radiolabeled sugar analog after an imposed transmembrane pH gradient (ApH). A transmembrane pH gradient was imposed by using a modification of the method of Maloney (27). Washed cells were suspended in 50 mM K+-FMM, and endogenous energy reserves were depleted at pH 7.2 in the pH stat and PEP depleted as described above. To 1 ml of the cell suspension (5 mg [dry weight] per ml) 3.5 ml of a 200 mM potassium phosphate buffer (pH 5.0) was added, which rapidly lowered the extracellular pH from 7.2 to 5.2. Immediately, the radiolabeled sugar analog 6deoxy[3H]glucose (0.5 Ci/mol), 2-deoxy[14C]glucose (1.0 Ci/ mol), or ['4C]TMG (1.0 Ci/mol) was added to 4 mM and uptake was monitored as described above. Uptake of radiolabeled sugar analog in exogenously energized cells. Washed cells were suspended in 50 mM K+FMM to 5 mg (dry weight) per ml and energy depleted. Fructose or galactose in 50 mM K+-FMM was then added to the cell suspension in a pH stat to give a final concentration of 10 mM. The fructose-energized cells were allowed to lower the extracellular pH to 5.0, where it was maintained by the addition of 0.1 M KOH. After acid production reached a steady state, 6-deoxy[3H]glucose (0.5 ,uCi/,umol) was added to 4 mM and uptake was measured as described above. The galactose-energized cells were maintained at an extracellular pH of 7.0, and 6-deoxyglucose uptake was measured in the was as a
same way.
Glucose and analog transmembrane diffusion kinetics. Glutransmembrane diffusion was measured by using a modification of the methods of Winkler and Wilson (47) and Maloney et al. (29). Washed cells were suspended in FMM to give a cell density of 4 mg (dry weight) per ml and preincubated in a pH stat (pH 7.0, 37°C) until acid production had ceased. Radiolabeled glucose or analog ([14C]glucose, 1.0 Ci/mol; 6-deoxy[3H]glucose, 0.5 Ci/mol; or 2-deoxy[14C]glucose, 1.0 Ci/mol) was added to aliquots of the cell suspension to give a series of concentrations between 0.05 and 50 mM. Prior to the 2-deoxy[14C]glucose incubations, the cells were PEP depleted with TMG (see above), and for the [14C]glucose incubations, KF was added (20 mM final concentration) to the deenergized, PEP-depleted cells. Samples were taken at 10-s intervals for 1 min. Each sample was filtered through a 0.2-p.m-pore-size polycarbonate filter and washed rapidly (
Vol. 172, No. 2
0021-9193/90/020556-08$02.00/0
Characterization of Transmembrane Movement of Glucose and Glucose Analogs in Streptococcus mutans Ingbritt STUART G. DASHPER AND ERIC C. REYNOLDS* Biochemistry and Molecular Biology Unit, Faculty of Medicine and Dentistry, University of Melbourne, 711 Elizabeth Street, Melbourne, 3000, Australia Received 1 August 1989/Accepted 23 October 1989
The transmembrane movement of radiolabeled, nonmetabolizable glucose analogs in Streptococcus mutans Ingbritt was studied under conditions of differing transmembrane electrochemical potentials (A*) and pH gradients (ApH). The ApH and A* were determined from the transmembrane equilibration of radiolabeled benzoate and tetraphenylphosphonium ions, respectively. Growth conditions of S. mutans Ingbritt were chosen so that the cells had a low apparent phosphoenolpyruvate (PEP)-dependent glucose:phosphotransferase activity. Cells energized under different conditions produced transmembrane proton potentials ranging from -49 to -103 mV but did not accumulate 6-deoxyglucose intracellularly. An artificial transmembrane proton potential was generated in deenergized cells by creating a A* with a valinomycin-induced K+ diffusion potential and a ApH by rapid acidification of the medium. Artificial transmembrane proton potentials up to -83 mV, although producing proton influx, could not accumulate 6-deoxyglucose in deenergized cells or 2-deoxyglucose or thiomethylgalactoside in deenergized, PEP-depleted cells. The transmembrane diffusion of glucose in PEP-depleted, KF-treated cells did not exhibit saturation kinetics or competitive inhibition by 6-deoxyglucose or 2-deoxyglucose, indicating that diffusion was not facilitated by a membrane carrier. As proton-linked membrane carriers have been shown to facilitate diffusion in the absence of a transmembrane proton potential, the results therefore are not consistent with a proton-linked glucose carrier in S. mutans Ingbritt. This together with the lack of proton-linked transport of the glucose analogs suggests that glucose transmembrane movement in S. mutans Ingbritt is not linked to the transmembrane proton potential.
The oral bacterium Streptococcus mutans and dietary sugar have been implicated in the development of dental caries (7, 26). Fundamental to the existence of S. mutans in dental plaque is its ability to compete effectively for sugar substrates under differing environmental conditions. Competition is at the level of cellular transport. It is well established that S. mutans transports glucose, 2-deoxyglucose, sucrose, lactose, mannose, and fructose by a phosphoenolpyruvate (PEP)-dependent sugar:phosphotransferase system (PTS) (4, 8, 12, 25, 42). Substrate specificity is conferred by the membrane-bound enzymes II, with each protein recognizing a series of structurally related sugars (36, 41). A second glucose transport system in S. mutans has been proposed by Hamilton and St. Martin (9) and Keevil et al. (22) to involve a carrier-mediated proton-glucose symport driven by the transmembrane electrochemical proton potential (Ap) or proton motive force (PMF) (33). Keevil et al. (22) have suggested that sugar transport in S. mutans at the low pH of cariogenic plaque would occur predominantly by this PMF-driven transporter, and therefore, this system could be an important virulence determinant. However, the evidence to support the existence of a PMF-driven glucose transporter is equivocal and is based largely on the findings that acid production from glucose by S. mutans is inhibited by certain PMF-dissipating ionophores and that the glucose-PTS activity of toluenized cells is not commensurate with the rate of acid production by whole cells with glucose as the substrate. To interpret these results as evidence for a PMF-driven glucose transporter, it must be assumed that glucose uptake is rate limiting for glycolysis, that the PTS activity of toluenized cells is a quantitative measure of whole-cell *
activity, and that ionophores do not affect the PTS or glycolysis through perturbation of intracellular H+ and K+ concentrations. Transport of a nonmetabolizable glucose analog in S. mutans Ingbritt has been investigated by Keevil et al. (21). They used radiolabeled 6-deoxyglucose, which is an ideal substrate as it cannot be actively translocated by the glucose-PTS. However, the results of this study were again equivocal owing to the low level of accumulation and the high level of cellular adsorption of the radiolabel. Further, the accumulation of 6-deoxyglucose was not correlated with Ap nor was a link between proton and 6-deoxyglucose transmembrane movement demonstrated. In an approach to characterize glucose transport in S. mutans Ingbritt, we attempted to demonstrate a PMF-driven glucose transport system by correlating intracellular accumulation of radiolabeled, nonmetabolizable glucose analogs with Ap. The relative contributions of the PMF-driven system and the PTS were then to be determined under different environmental conditions. However, proton movement across the cell membrane did not result in an intracellular accumulation of 6-deoxyglucose or 2-deoxyglucose. Further, the diffusion of ['4C]glucose in PEP-depleted, metabolically poisoned cells did not exhibit saturation kinetics, indicating the absence of a membrane carrier capable of facilitating diffusion. The results therefore are not consistent with a PMF-linked glucose carrier in S. mutans Ingbritt. MATERIALS AND METHODS Bacterial strains. S. mutans Ingbritt (24) and variant 162 (3) were kindly supplied by B. Krasse (Karolinska Institutets, Stockholm, Sweden) and K. Knox (Institute of Dental Research, Sydney, Australia), respectively. The bacteria were stored as lyophilized cultures in sealed ampoules or in
Corresponding author. 556
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30% glycerol broth at -20°C. The strains were checked regularly according to the criteria of Hardie and Bowden (10). Growth conditions. Bacteria were grown as stationary batch cultures or in continuous culture at 37°C. The growth medium for the stationary batch cultures contained 1% (wt/vol) tryptone, 0.5% (wt/vol) yeast extract, 1.62 mM MgSO4, 0.04 mM FeSO4, 0.06 mM MnCl2, 9.70 mM KH2PO4, 15.31 mM K2HPO4, 35 mM KCl, and 55 mM glucose and was adjusted to pH 7.0. The glucose and the phosphate salts were individually filter sterilized and added to the medium before inoculation. Batch-grown cells were harvested in the mid-exponential phase. Continuous cultures were grown in a BioFlo model C-30 chemostat (New Brunswick Scientific Co., Inc., Edison, N.J.) with a working capacity of 365 ml. The temperature was maintained at 37°C, and the pH was controlled at 7.0 or 5.5 as specified by the automatic addition of 5 M KOH. Cultures were gassed with 5% CO2 in nitrogen. The dilution rate was set at D = 0.1 h-1, which is equivalent to a mean generation time of 6.9 h, and cultures were allowed to equilibrate for 10 generations before sampling. The growth medium was the same as that for the batch cultures except that the glucose was 11 mM, which was limiting. Culture purity was checked regularly by microscopic examination and by the criteria of Hardie and Bowden (10). Chemostatgrown cells were collected from the overflow at 4°C for less than 1 h. Chemostat- or batch-grown cells were centrifuged at 1,000 x g for 15 min at 4°C and washed twice with fermentation minimal medium (FMM) containing 50 mM KCl, 5 mM NaCl, 2 mM MgSO4, 2 mM MnCl2, 8 mM (NH4)2SO4, 1.5 ,uM thiamine, and 8 p,M niacin at pH 7.0. The cells were resuspended in FMM to give a cell density of 1.25 mg (dry weight) of cells per ml. Measurement of glycolytic activity. Glycolytic activity was determined in a pH stat (Radiometer Copenhagen, Copenhagen, Denmark) essentially as described by Marsh et al. (32), using a volumetric standard titrant (0.1 M KOH) and a 10 mM final glucose concentration, which was saturating (39). Before the addition of the prewarmed glucose solution, the cells were depleted of endogenous reserves of carbohydrate by preincubation at pH 7.0 in the pH stat for approximately 22 min. Glycolytic activity was measured as nanomoles of H' neutralized per milligram (dry weight) of cells per minute. The dry weight of cells was determined by filtering 1-ml samples of the cell suspension through dry, preweighed, 0.2-,um-pore-size polycarbonate filters (Nuclepore Corp., Pleasanton, Calif.) supported on Whatman GF/ A filters. The ionophore gramicidin D (Sigma Chemical Co., St. Louis, Mo.) was added as a 10 mM solution in 95% ethanol to give a final concentration of 10 p.M. All controls had equivalent amounts of ethanol added, which had no effect on acid production. PTS assay. The PEP-dependent glucose-PTS was assayed by a modification of the method of Kornberg and Reeves (23). Batch- or chemostat-grown cells were centrifuged at 1,000 x g for 10 min at 4°C. The cell pellet was washed twice in 100 mM potassium phosphate buffer (pH 7.2) containing 5 mM dithiothreitol and then resuspended in the same buffer to a cell density of 10 mg (dry weight) of cells per ml. Tolueneacetone (1:9) was used to permeabilize the cells. The PTS assay reaction mixture contained 100 mM potassium phosphate buffer (pH 7.2), 5 mM MgSO4, 5 mM PEP, 5 mM dithiothreitol, 10 mM KF, 0.3 mM NADH (Sigma), 5 U of
557
L-lactic dehydrogenase (EC 1.1.1.27; Sigma) per ml (700 to 1,400 U/mg of protein), and 0.1 mg (dry weight) of cells. Intracellular water volume determination. Intracellular water volume was measured by a modification of the procedure of Maloney et al. (29) with [1,2-3H]taurine as the extracellular marker (15). Batch-grown cells were washed once in 0.1 M Tris hydrochloride (pH 7.2)-100 mM KCl and resuspended in the same buffer to give a cell density of 4.0 mg (dry weight) of cells per ml. [1,2-3H]taurine (50 p.Ci/pmol) was added to the cell suspension to give a final concentration of 100 ,uM. Samples were filtered through dry, preweighed, 0.2-p.m-pore-size polycarbonate filters (Nuclepore) without washing. Total water on the filter was determined gravimetrically, and radioactivity was determined after filter solubilization (Amersham Australia Pty. Ltd.). Filtrates were also collected, and their radioactivity was measured. The polycarbonate filters retained only approximately 30% of the total water as extracellular water, which allowed an accurate determination of total, extracellular, and intracellular water by the [3H]taurine-gravimetric method. In fact, the [3H] taurine-gravimetric method proved to be slightly more reproducible (coefficient of variation, 4.1%) than the ['4C]taurine-3H20 method (coefficient of variation, 8.8%). The intracellular water volume of S. mutans Ingbritt from four separate determinations was 1.69 ± 0.07 ,ul/mg (dry weight) of cells. This compares well with the value obtained by Noji et al. (34) of 1.60 ± 0.14 p.l/mg (dry weight). Uptake of radiolabeled sugar analog in endogenously energized cells. Washed cells were suspended in 4 ml of either 100 mM Tris-citrate buffer (pH 7.0) or FMM containing 5 mM dithiothreitol to give a cell density of 5 mg (dry weight) per ml and used without preincubation. Radiolabeled sugar analog (['4C]thiomethylgalactoside (['4C]TMG), 1.0 Ci/mol; 6-deoxy[3H]glucose, 0.5 Ci/mol; or 2-deoxy['4C]glucose, 1.0 Ci/mol) was added to the cell suspension in a pH stat (pH 7.0, 37°C) to give a final concentration of 4 mM. Samples were taken at the times indicated and filtered through dry, preweighed, 0.2-p.m-pore-size polycarbonate filters without washing. Intracellular and extracellular water volumes and filter and filtrate radioactivities were determined as described above. The amount of intracellular radiolabeled sugar was determined by subtracting the extracellular radioactivity from the total radioactivity on the filter. A control (10 puM gramicidin-treated cells) was used to determine non-PMF uptake or cellular adsorption or both. Uptake of radiolabeled sugar analog after an imposed membrane potential (AO. A membrane potential was imposed by valinomycin treatment of K+-loaded, deenergized cells in low-K+ medium as described by Kashket and Wilson (18) and Maloney (27, 28). Washed cells were suspended in 10 mM K+-FMM or 100 mM taurine-25 mM choline chloride-25 mM KCl (pH 7.2) and preincubated in a pH stat (pH 7.0, 37°C) until acid production ceased. Prior to the [14C]TMG uptake experiments, 2-deoxyglucose was added to the cell suspension to give a final concentration of 1 mM and incubated for 20 min to deplete intracellular PEP. Similarly, prior to the 2-deoxy[14C]glucose uptake experiments, TMG was added to a final concentration of 1 mM and incubated for 20 min. After PEP depletion, the cells were washed once in 100 mM taurine-50 mM choline chloride-0.3 mM KCl (pH 6.5) and then resuspended in the same buffer to give a cell density of 5 mg (dry weight) per ml. The radiolabeled sugar analog 6-deoxy[3H]glucose (0.5 p.Ci/
p.mol), 2-deoxy[14C]glucose (1.0 p.Ci/p.mol), or [14C]TMG (1.0 p.Ci/p.mol) was added to the deenergized, PEP-depleted cells to give a final concentration of 4 mM, and after
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15 min, valinomycin (Sigma) was added to 10 p.M. Uptake measured as described above. Valinomycin was added 10 mM solution in 95% ethanol. Uptake of radiolabeled sugar analog after an imposed transmembrane pH gradient (ApH). A transmembrane pH gradient was imposed by using a modification of the method of Maloney (27). Washed cells were suspended in 50 mM K+-FMM, and endogenous energy reserves were depleted at pH 7.2 in the pH stat and PEP depleted as described above. To 1 ml of the cell suspension (5 mg [dry weight] per ml) 3.5 ml of a 200 mM potassium phosphate buffer (pH 5.0) was added, which rapidly lowered the extracellular pH from 7.2 to 5.2. Immediately, the radiolabeled sugar analog 6deoxy[3H]glucose (0.5 Ci/mol), 2-deoxy[14C]glucose (1.0 Ci/ mol), or ['4C]TMG (1.0 Ci/mol) was added to 4 mM and uptake was monitored as described above. Uptake of radiolabeled sugar analog in exogenously energized cells. Washed cells were suspended in 50 mM K+FMM to 5 mg (dry weight) per ml and energy depleted. Fructose or galactose in 50 mM K+-FMM was then added to the cell suspension in a pH stat to give a final concentration of 10 mM. The fructose-energized cells were allowed to lower the extracellular pH to 5.0, where it was maintained by the addition of 0.1 M KOH. After acid production reached a steady state, 6-deoxy[3H]glucose (0.5 ,uCi/,umol) was added to 4 mM and uptake was measured as described above. The galactose-energized cells were maintained at an extracellular pH of 7.0, and 6-deoxyglucose uptake was measured in the was as a
same way.
Glucose and analog transmembrane diffusion kinetics. Glutransmembrane diffusion was measured by using a modification of the methods of Winkler and Wilson (47) and Maloney et al. (29). Washed cells were suspended in FMM to give a cell density of 4 mg (dry weight) per ml and preincubated in a pH stat (pH 7.0, 37°C) until acid production had ceased. Radiolabeled glucose or analog ([14C]glucose, 1.0 Ci/mol; 6-deoxy[3H]glucose, 0.5 Ci/mol; or 2-deoxy[14C]glucose, 1.0 Ci/mol) was added to aliquots of the cell suspension to give a series of concentrations between 0.05 and 50 mM. Prior to the 2-deoxy[14C]glucose incubations, the cells were PEP depleted with TMG (see above), and for the [14C]glucose incubations, KF was added (20 mM final concentration) to the deenergized, PEP-depleted cells. Samples were taken at 10-s intervals for 1 min. Each sample was filtered through a 0.2-p.m-pore-size polycarbonate filter and washed rapidly (
