Section of Microbiology, Cornell University1, and Agricultural Research Service, USDA2, Ithaca, NY 14853, USA
Author for correspondence: J. B. Russell. Tel: +1 607 255 4508. Fax: +1 607 255 3904. e-mail: jbr8{at}cornell.edu
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ABSTRACT |
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Keywords: protonmotive force, proton conductance, energy spilling, Streptococcus bovis
Abbreviations: 9-AA, 9-aminoacridine; BCECF, 2,7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein; BCECF-AM, acetoxymethyl ester derivative of BCECF; DCCD, dicyclohexylcarbodiimide; TCS, tetrachlorosalicylanilide; TPP, tetraphenylphosphonium
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INTRODUCTION |
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Non-growing Streptococcus bovis ferments sugars at a rate that greatly exceeds its needs for maintenance, lacks alternative respiratory pathways, does not store polymers, and only uses substrate-level phosphorylation to generate ATP (Russell & Strobel, 1990 ). Because non-growth ATP hydrolysis could be strongly inhibited by the F-type ATPase inhibitor DCCD (dicyclohexylcarbodiimide) and was stimulated by protonophores, it appeared that S. bovis was using a proton-pumping F-ATPase to turn over excess ATP. Similar observations have been made in Enterococcus (Harold & Baarda, 1969
) and Lactococcus (Maloney, 1977
).
Previous work indicated that changes in intracellular inorganic phosphate could play a role in regulation of ATP hydrolysis in S. bovis. When energy was in excess, inorganic phosphate decreased, the G0' of ATP hydrolysis was greater, and protonmotive force generated by the F-ATPase was significantly higher (Bond & Russell, 1998
). Gross estimates based on Ohms law indicated that the resistance of the cell membrane to protons declined as protonmotive force increased, allowing faster rates of ATP hydrolysis and proton pumping. However, proton flux rates predicted by these calculations were 2050-fold faster than values previously measured in bacteria (Maloney, 1979
; Rius & Lorén, 1998
; Rius et al., 1994
; Bender et al., 1986
).
The following experiments with S. bovis sought to: (1) generate artificial protonmotive forces similar to those observed in fermenting cells, (2) investigate the relationship between protonmotive force and ion flux, and (3) compare these fluxes to other biological systems.
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METHODS |
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Glucose consumption rate of non-growing cells.
Starved cells (>4 h in stationary phase to deplete intracellular potassium) were washed three times (10000 g, 5 min, 4 °C) in anoxic buffer (100 mM MES/HCl, 0·5 mM MgCl2 and 3 mM cysteine, prepared with double-distilled H2O, pH 6·8), resuspended to a final concentration of 240 µg protein ml-1, gassed with a continuous stream of O2-free N2 in a temperature-controlled chamber (39 °C) and provided with 8 mg glucose ml-1. NaCl or KCl (5 mM), valinomycin (1 µM) or tetrachlorosalicylanilide (TCS; 1 µM) was added where indicated. Glucose consumption by non-growing cells was measured by monitoring bacterial heat production as described previously (Russell & Strobel, 1990 ) with an LKB 2277 Bioactivity monitor equipped with semi-conducting Peltier elements as thermopiles and gold flow cells. The instrument was calibrated with an internal heat source, and the flow cell was sterilized with 95% ethanol and 1 M HCl. The flow cell temperature was set at 39·00 °C. The glucose consumption rate was estimated from the enthalpy (
H) of glucose conversion to lactate [87·6 J (mmol glucose)-1] and the conversion factor 0·278 mW J-1 h-1. As 2 ATP are produced per glucose fermented, non-growing cells must hydrolyse 2 ATP per glucose for glucose consumption to continue.
Intracellular pH measurements using BCECF.
The hydrophilic compound 2,7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein (BCECF) was purchased as its hydrophobic acetoxymethyl ester derivative (BCECF-AM) (Molecular Probes). S. bovis cells were grown to stationary phase, washed three times in buffer (10 mM K2HPO4, 80 mM KCl, 0·5 mM MgCl2 and 0·5 mM DTT, pH 7·8), concentrated 20-fold and equilibrated/de-energized for 12 h (22 °C) with 0·1 µM valinomycin. After equilibration, cells were centrifuged, resuspended in 0·5 ml of the same buffer, and BCECF-AM was added to a final concentration of 10 µM (from a concentrated stock dissolved in DMSO). BCECF-AM can diffuse into cells and be cleaved by intracellular esterases, and BCECF is trapped inside cells. The fluorescence of BCECF is pH-dependent, and this compound can serve as an indicator of intracellular pH. Cells were incubated with BCECF-AM for 1 h at 39 °C, washed twice with 40 ml of buffer containing 0·1 µM valinomycin, resuspended to a final optical density of approximately 100 (600 nm, 1 cm light path), and stored on ice until use. The presence of valinomycin and excess potassium ions allowed for charge compensation when protons entered cells during the assay (Maloney, 1977 ). If valinomycin was omitted, proton influx rates were significantly inhibited. Cells were diluted 100-fold into 39 °C buffer containing 0·1 µM valinomycin, and equilibrated for 15 s in 3 ml cuvettes in a Bio-Rad VersaFluor fluorimeter (490 nm excitation, 520 nm emission). Extracellular pH was lowered by adding 540 µl 1 M HCl while vigorously stirring the cuvette with a pipette. Measurements could reliably be recorded within 56 s of acid addition. Extracellular pH was determined in the cuvette 30 s after acid addition, with a pH meter. DCCD-treated cells were incubated in 1 mM DCCD for 30 min before being assayed in buffer containing 0·5 mM DCCD. Intracellular pH was calibrated for each batch of cells by adding 1 mM each valinomycin and nigericin, adding HCl, and recording both fluorimeter readings and pH in the cuvette. Cell suspensions were not subjected to more than five readings during calibration, to minimize photobleaching. Calibrations performed after loading and 2 h later gave similar results, indicating that leakage (efflux) of the probe (Molenaar et al., 1991
, 1992
) was not significant.
Measurement of proton flux using 9-aminoacridine (9-AA).
As the neutral form of the fluorescent amine 9-AA can freely cross the cell membrane, the distribution of 9-AA (inside versus outside) is pH-dependent. Because only extracellular 9-AA gives fluorescence, uptake and quenching of 9-AA fluorescence is an indicator of decreased intracellular pH (Casadio et al., 1995 ). Potassium-loaded cells were obtained by harvesting cells during growth, and washing them three times in 4 °C choline buffer (50 mM choline chloride, 10 mM Bistris, 0·5 mM MgCl2, 0·5 mM DTT, prepared with double-distilled H2O, pH 6·8). Cells were concentrated to an OD600 of approximately 100 and kept on ice until use. Warmed buffer (39 °C) containing 0·4 µM 9-AA was placed in pre-warmed acid-washed glass vials, cells were added to a final concentration of 1·5 OD600 units (0·9 mg protein ml-1), and the vial was placed in a Perkin Elmer model 203 fluorescence spectrophotometer (405 nm excitation, 495 nm emission) to obtain a baseline. Unless otherwise stated, 0·1 µM valinomycin was added to establish a membrane potential. Measurements could be obtained within 5 s of closing the instrument door, and quenching of 9-AA was measured over the next 30 s. Rates of proton influx were estimated from the initial rate (first 10% of quenching). Potassium (as KCl) was added to vials to adjust membrane potential, and membrane potentials were calculated using the Nernst equation. Previous work indicated that more than 98% of intracellular potassium in S. bovis is unbound (Strobel & Russell, 1989
). Intracellular [typically 1·5 mmol K+ (g protein)-1, or approx. 350 mM] and extracellular (typically 0·20·4 mM unless otherwise adjusted) potassium concentrations were measured by flame photometry as described below (9-AA did not interfere with flame photometry). DCCD treatment was the same as for BCECF-loaded cells.
Measurement of intracellular potassium efflux using flame photometry.
Potassium-loaded cells were prepared from growing cells as described for 9-AA-based measurements. Choline buffer (1 ml, 39 °C) with added potassium to reduce the membrane potential was placed in pre-warmed acid-washed glass tubes with stir bars. Concentrated cell suspensions were added and allowed to equilibrate for 30 s at 39 °C (final concentration 0·20·4 mg protein ml-1). Valinomycin was added, and cells were centrifuged through silicone oil at various times. Sampling times were from the time of valinomycin addition to the initiation of centrifugation. Samples (100 µl) of supernatant were removed, the tubes were frozen, and once the liquid above the silicone was solid, cell pellets were removed with dog nail-clippers. Supernatants and cell pellets were digested at room temperature for 24 h in 3 M HNO3, and insoluble cell debris was removed by centrifugation (10000 g, 15 min). The potassium concentration was determined by flame photometry (Cole-Parmer 2655-00 Digital Flame Analyzer, Cole-Parmer Instrument Co.).
When the filter method was employed, lower cell densities (0·1 mg protein ml-1) were utilized so cells could be rapidly separated from supernatant. Valinomycin was added to the cell suspensions (1 ml), potassium efflux was terminated with the addition of 2 ml ice-cold 100 mM LiCl, and cell suspensions were immediately filtered through Millipore type HA 0·45 µm filters that had been pre-soaked in 100 mM LiCl. The filter was then rinsed with an additional 2 ml of cold LiCl to remove contaminating potassium from the filter. Filters were digested in 3 M HNO3 for 24 h, and analysed as described. Duplicate samples were centrifuged through silicone oil to obtain supernatant samples for extracellular potassium determination.
Measurement of potassium efflux using CD222.
Potassium-loaded cells were washed and concentrated in choline buffer and placed in pre-warmed, acid-washed glass vials (0·24 mg protein ml-1). CD222 (1 µM), a fluorescent probe sensitive to extracellular potassium (Molecular Probes), was added to the cell suspensions, and preliminary results found CD222 to be sensitive in the range of 0·051·0 mM potassium. After a baseline was obtained (Perkin Elmer model 203 fluorescence spectrophotometer; 365 nm excitation, 465 nm emission), 0·1 µM valinomycin was added, and the increase in fluorescence was monitored. Measurements could be obtained within 5 s of closing the instrument door. Potassium efflux rates were based on the difference between extracellular potassium at time of valinomycin addition and the concentration 10 s later. The system was calibrated by adding potassium (up to 1·5 mM) directly to vials containing cells and using flame photometry to determine the initial extracellular potassium concentrations (typically 0·40·6 mM). Membrane potentials were adjusted by adding extracellular potassium (to 1 mM) or by using cells that had only been allowed to partially deplete their intracellular potassium. Potassium-depleted cells were washed in buffer to remove residual potassium before use.
ATPase activity of inverted vesicles.
Membrane vesicles were prepared as previously described (Russell et al. 1988 ). Briefly, 10 ml of lysozyme/mutanolysin-treated protoplasts were rapidly diluted in 1 litre of low-osmotic-strength buffer, incubated with DNase and RNase, unbroken cells removed by low-speed centrifugation (1000 g, 30 min, 4 °C), and vesicles were harvested by high-speed centrifugation (30000 g, 30 min, 4 °C). This vesicle preparation was washed twice in buffer (50 mM potassium phosphate, pH 7·0, 10 mM MgSO4), concentrated to 24·5 mg protein ml-1, and passed twice through a French pressure cell at 55 MPa. Inverted vesicles were washed twice in buffer (35000 g, 30 min, 4 °C), and concentrated to 14 mg protein ml-1. Some inverted vesicles were assayed for ATPase activity the same day, and the rest were frozen at -80 °C. Freezing for up to 3 d did not significantly affect ATPase activity.
ATPase activity was assayed by a method employing pyruvate kinase as described by Bond & Russell (1996) . The assay contained (per ml): 1020 mg liposome protein, 50 µmol triethanolamine, 2 µmol MgCl2, 0·16 µmol NADH, 0·83 µmol phosphoenolpyruvate, and 5 units each pyruvate kinase and lactate dehydrogenase (pH 7·0, 39 °C). DCCD addition only reduced the ATPase activity 5070%, but pre-incubation of inverted vesicles with DCCD (15 min, 1 mM) eliminated 90% of the activity. The ATPase activity of inverted vesicles and whole cells was normalized using the assumption that 15% of cell protein is associated with the membrane (Maloney, 1987
; Cramer & Knaff, 1991
). Protein was measured by the Lowry method after boiling vesicles in 0·2 M NaOH.
Other assays.
Membrane potential was estimated from the uptake of [3H]TPP+. Cells were incubated with [3H]TPP+ (10 nM, 851 GBq mmol-1; Amersham) for 2 min (or shorter intervals, if indicated). The cells were centrifuged through silicone oil, and [3H]TPP+ in pellets and supernatants was measured by scintillation counting. Cells were de-energized for 10 min with 10 µM valinomycin and 10 µM nigericin to correct for nonspecific [3H]TPP+ binding, and separate determinations of nonspecific binding were made for each potassium concentration or medium condition (Lolkema et al., 1982 ). [14C]Benzoate (1 µM final concentration, 0·81 GBq mmol-1; Amersham) was used in a similar fashion to determine intracellular pH of glycolysing cells. The intracellular volume of whole cells was assumed to be 4·3 µl (mg protein)-1 (Strobel & Russell, 1989
). Cellular ATP was assayed by extracting cells with perchloric acid, neutralizing with KOH and K2CO3, and measuring luminescence using a luciferinluciferase assay kit (Sigma) as previously described (Russell & Strobel, 1990
).
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RESULTS |
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DISCUSSION |
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Otto (1984) noted that L. lactis had a potential futile cycle of glycolytic enzymes, but this cycle only accounted for 1·5% of the glycolytic rate. Harold & Baarda (1969)
, Maloney (1977)
and Maloney & Hanson (1982)
found that ATP turnover in enterococci and lactococci could be strongly inhibited by DCCD, an inhibitor of membrane F-ATPases, and Maloney (1977)
noted that this ATP consumption could also be stimulated by a protonophore. Previous work indicated that ATP dissipation in S. bovis cells was stimulated by TCS (a protonophore) and inhibited by DCCD (Russell & Strobel, 1990
), and similar results were noted in our initial experiments (Figs 1
and 2
). Inverted S. bovis membrane vesicles hydrolysed ATP at a rapid rate [equivalent to 13 mmol H+ (g protein)-1 h-1], and virtually all of this activity was DCCD sensitive. These observations supported the idea that that proton-pumping, membrane-bound ATPase activity was the dominant mechanism of non-growth ATP hydrolysis in S. bovis.
Because ATP hydrolysis is the energy source for protonmotive force generation in streptococci (Harold & Papineau, 1972 ), ATP hydrolysis is inhibited as the two forces approach equilibrium (
p=
G0 of ATP hydrolysis/n, where n is the ratio of protons pumped/ATP) (Hirata et al., 1986
; Van Walraven et al., 1996
). In order for ATP hydrolysis to continue, protonmotive force must continually be dissipated via proton conductance across the cell membrane. Reported values of passive proton conductance in Gram-positive species are less than 1 mmol H+ (g protein)-1 h-1 (Maloney, 1979
; Rius & Lorén, 1998
; Rius et al., 1994
; Bender et al., 1986
). However, S. bovis had an ATP hydrolysis rate of 39 mmol ATP (g protein)-1 h-1, and based on a ratio of protons pumped/ATP of 4, the proton conductance of S. bovis was 156 mmol H+ (g protein)-1 h-1, a value 150-fold greater than previously measured values. The protonmotive force of glycolysing S. bovis cells was approximately -150 mV, but proton conductance values in other bacteria were estimated at much lower driving forces (approx. -6 mV). Ohms law states that current (proton flux) increases as a linear function of driving force (mV), but even this correction could not explain the estimated membrane conductance of S. bovis cells.
Rapid pH shifts are a straightforward method for generating large protonmotive forces, and a variety of workers have used this technique to estimate proton flux in liposomes (Deamer & Nichols, 1989 ). Converting rates of pH change to rates of proton flux is dependent on an estimate of cellular buffering capacity, and while literature values for lactic acid bacteria average approximately 200 nmol H+ (mg protein)-1 per pH unit over an internal pH of 7 to 8, values vary from 50 to at least 350 nmol H+ (mg protein)-1 per pH unit (Maloney, 1979
; Rius et al., 1994
; Rius & Lorén, 1998
). When S. bovis cells were subjected to a pH gradient of 2·75 pH units (-170 mV), the internal pH declined at a rate of 0·15 pH units s-1, and based on a buffering capacity estimate of 200 nmol H+ (mg protein)-1 per pH unit, proton conductance was 108 mmol H+ (g protein)-1 h-1. As the magnitude of the pH gradient was decreased, the initial rate of proton conductance declined in a non-ohmic fashion, to an estimated rate of 10 mmol H+ (g protein)-1 h-1 at a
pH of 0·75 (-46 mV). These rates were much faster than previously measured values of proton conductance in similar bacteria.
Electrogenic potassium diffusion is also a common method of generating large membrane potentials, because large potassium efflux can only occur if other ions (e.g. protons) enter the cell. For instance, when S. bovis cells were loaded with 300 mM potassium, each cell had approximately 12x107 internal potassium ions, and over 98% of this potassium was free (Strobel & Russell, 1989 ). Based on a membrane capacitance of 1 µF cm-2, an efflux of only 30000 potassium ions per cell (<0·025% of the total potassium) would have generated a membrane potential of -150 mV (Maloney, 1987
; Cramer & Knaff, 1991
). Because valinomycin-treated cells lost more than 50% of their intracellular potassium in a very short time, it appeared that influx of another ion was allowing potassium loss. 9-AA measurements indicated that valinomycin-treated cells took up protons at a rapid rate, but these measurements were not quantitative.
Maloney (1977) used extracellular pH measurements to estimate proton fluxes in dense suspensions of L. lactis cells, and noted that potassium efflux via valinomycin was balanced by a stoichiometric (1:1) influx of protons. The idea that potassium efflux was coupled with proton influx was also supported by the observation that TCS greatly increased the rate of potassium efflux and proton influx. Because membrane potentials were always below the threshold required for proton influx via the ATPase (as evidenced by the lack of an effect of DCCD and the data of Maloney, 1977
), potassium movements provided an estimate of proton conductance across the cell membrane. Regardless of the method used, potassium efflux measurements demonstrated that proton influx was a non-ohmic function of imposed membrane potential, and rates were always in excess of previously reported values.
Proton conductance is often expressed in terms of membrane surface area. S. bovis has a mean cell composition of 50% protein and 10% lipid by weight (Russell & Robinson, 1984 ), and Brookes et al. (1997)
estimated that lipid bilayers have 2200 cm2 of surface area per mg lipid. Based on this relationship [440 cm2 (mg protein)-1] a proton leak rate of 100 mmol H+ (g protein)-1 h-1 of S. bovis corresponds to 6·2x10-11 mol H+ (cm membrane)-2 s-1. Respiring liver mitochondria have proton leak rates that range from 0·8 to 3·6x10-11 mol H+ (cm membrane)-2 s-1 and these leak rates are also non-ohmic (Krishnamoorthy & Hinkle, 1984
; Brookes et al., 1997
; Brand et al., 1994
).
Other lactic acid bacteria had non-growth ATP hydrolysis at rates comparable to those estimated for S. bovis (Rosenberger & Elsden, 1960 ; Fordyce et al., 1984
), but direct measurements of proton conductance could not explain these high rates of ATP turnover (Maloney, 1979
; Rius & Lorén, 1998
; Rius et al., 1994
; Bender et al., 1986
). These discrepancies can be explained by the non-ohmic relationship between proton conductance and driving force. When the driving force was low (<40 mV), the proton conductance of S. bovis was similar to the values previously reported. However, if the driving force was similar to ones found in glycolysing cells (160 mV), the proton conductance was at least 10-fold higher. Further work is needed to define the nature of this proton-conducting pathway.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Received 23 August 1999;
revised 11 November 1999;
accepted 2 December 1999.
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