Proc. Natl. Acad. Sci. USA Vol. 73, No. 6, pp. 1892-1896, June 1976 Biochemistry
The electrochemical gradient of protons and its relationship to active transport in Escherichia coli membrane vesicles (flow dialysis/membrane potential/energy transduction/lipophilic cations/weak acids)
SOFIA RAMOS, SHIMON SCHULDINER*, AND H. RONALD KABACK The Roche Institute of Molecular Biology, Nutley, New Jersey 07110
Communicated by B. L. Horecker, March 17, 1976
ABSTRACT Membrane vesicles isolated from E. coli generate a trans-membrane proton gradient of 2 pH units under appropriate conditions when assayed by flow dialysis. Using the distribution of weak acids to measure the proton gradient (ApH) and the distribution of the lipophilic cation triphenylmethylphosphonium to measure the electrical potential across the membrane (AI), the vesicles are shown to generate an electrochemical proton gradient (AiH+) of approximately -180 mV at pH 5.5 in the presence of ascorbate and phenazine methosulfate, the major component of which is a ApH of about -110 mV. As external pH is increased, ApH decreases, reaching o at pH 7.5 and above, while AI remains at about -75 mV and internal pH remains at pH 7.5. Moreover, the ability of various electron donors to drive transport is correlated with their ability to generate A4H+. In addition, ApH and Ad can be varied reciprocally in the presence of valinomycin and nigericin. These data and others (manuscript in preparation) provide convincing support for the role of chemiosmotic phenomena in active transport. Despite apparently contradictory initial observations (1-3), an increasing accumulation of experimental evidence (4-6) indicates that chemiosmotic phenomena, as postulated by Mitchell (7-10), play a central role in respiration-linked active transport in Escherichia coli membrane vesicles. It now seems clearly established that membrane vesicles prepared by the techniques developed in this laboratory retain the same orientation as the membrane in the intact cell (see ref. 11 for a summary of the evidence), and that oxidation of electron donors which drive transport in the vesicles results in the generation of a transmembrane electrical potential (interior negative) by means of electrogenic proton extrusion (6, 12-14). The potential is postulated to drive solute accumulation via facilitated diffusion of positively charged substrates such as lysine or via coupled movements of protons with neutral substrates such as lactose or proline (i.e., "symport"). According to the chemiosmotic hypothesis, the total driving force generated by proton extrusion is the electrochemical potential of protons across the membrane (A,4H+) (7-10). This thermodynamic entity is composed of an electrical and a chemical parameter according to the following relationship: = A A =H+
2.3RTA F ApH
[1]
where Ai represents the electrical potential across the membrane, and ApH is the chemical difference in proton concentration across the membrane (2.3RT/F is equal to 58.8 mV at room temperature). Through the use of lipophilic cations and rubidium (in the
presence of valinomycin), a respiration-dependent membrane potential (AI, interior negative) of approximately -75 mV in E. coli membrane vesicles has been documented (6, 13, 14). Moreover it has been shown that the potential causes the appearance of high affinity binding sites for dansyl- and azidophenylgalactosides on the outer surface of the membrane (4, 15) and that the potential is partially dissipated as a result of lactose accumulation (6). Although these findings provide evidence for the chemiosmotic hypothesis, it has also been demonstrated (6, 16) that vesicles are able to accumulate lactose and other substrates to intravesicular concentrations which are 100-fold or greater than those of the external medium. To sustain concentration gradients of this magnitude, a membrane potential of at least -120 mV is required. This observation, in addition so numerous negative attempts to establish the existence of a transmembrane pH gradient (1, 6) has left some doubt as to the quantitative relationship between AILH+ and solute accumulation. [In the apparent absence of a transmembrane pH gradient, it is also not immediately clear how anionic solutes are accumulated, by the chemiosmotic mechanism. suggested for this process (10)]. Recently, Padan et al. (17) have shown that intact E. coli generate a ApH (interior alkaline), and that the magnitude of the ApH is very dependent upon external pH, exhibiting a maximal value of about 2 pH units at pH 6.0 or below. In addition, Rottenberg (18) has utilized acetate to determine ApH in mitochondria, and suggested that this weak acid might be more useful than 5,5'-dimethyloxazolidine-2,4-dione (DMO) (19) in certain systems because it might be less permeant (H. Rottenberg; personal communication). The results presented in this paper were obtained by means of flow dialysis (20), a technique which allows a rapid, continuous determination of changes in the concentration of solutes in the external medium without manipulation of the vesicles. Using this technique, it is demonstrated that E. coli membrane vesicles generate a large proton gradient under appropriate conditions. In addition, we have shown that ApH and AI can be manipulated reciprocally by the ionophores valinomycin and nigericin. The results to be presented, and others (S. Ramos and H. R. Kaback, manuscript in preparation) which will be discussed, leave little doubt as to the primary role of chemiosmotic phenomena in respiration-dependent active transport. METHODS Growth of Cells and Preparation of Membrane Vesicles. E. coli ML 308-225 (i-z-y+a+) was grown on minimal medium A with 1.0% sodium succinate (hexahydrate), and membrane vesicles were prepared as described (21, 22). Vesicles were suspended in 0.1 M potassium phosphate (pH 6.6) and stored in liquid nitrogen. For studies at various pH's, membrane suspensions containing about 4 mg of protein per ml were removed from storage,
Abbreviations: DMO, 5,.5'-dimethyloxazolidine-2,4-dione; TPMP',
triphenylmethylphosphonium (bromide salt); PMS, phenazine methosulfate; CCCP, carbonylcyanide m-chlorophenylhydrazone. * Present address: Department of Molecular Biology, Hadassah Medical School, Hebrew University, Jerusalem, Israel. 1892
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Ramos et al.
thawed rapidly at 460, diluted at least 10-fold with 0.1 M potassium phosphate buffer at the desired pH, and incubated for 10 min at 25°. The suspension was centrifuged at 40,000 X g for 30 min, and the pellet resuspended and washed once in a similar volume of the same buffer. The final pellet was then resuspended to an appropriate protein concentration in 0.1 M potassium phosphate at the same pH. Transport Assays. Filtration assays (6, 23) were carried out on Millipore Cellotate filters (0.5 ,um pore size). Electron donors and isotopically labeled solutes were used as described. Flow dialysis was performed as described (24) except that the apparatus was modified so that the upper chamber was completely open to the atmosphere, and the reaction mixture was gassed with oxygen. The upper and lower chambers were separated by Spectrapor 1 dialysis tubing (600-8000 molecular weight cut-off; Fisher Scientific), and both chambers were stirred by means of magnetic bars. Membrane vesicles suspended in 0.05 M potassium phosphate at a given pH containing 0.'01 M magnesium sulfate were added to the upper chamber (total volume 0.8 ml), and electron donors, isotopically labeled solutes, and ionophores were added as indicated. Potassium phosphate (0.05 M at the same pH as the buffer in the upper chamber) was pumped from the lower chamber at a rate of 6.0 ml/min with a Pharmacia pump (model P3). Fractions of about 1.7 ml were collected and assayed for radioactivity by liquid scintillation spectrometry. Since phenazine methosulfate (PMS) causes about 5.5% quenching of tritium under the conditions described, control experiments were carried out in the absence of membrane vesicles, and the data were corrected appropriately. Determination of ApH. ApH.was determined by measuring the accumulation of acetate, propionate, butyrate or DMO by flow dialysis unless otherwise stated. Data were quantitated assuming that dialysis rates obtained after addition of nigericin (Fig. 1) represent 0 ApH. Determination of AI. The electrical potential across the membrane (Au') was determined by measuring the accumulation of [3H]triphenylmethylphosphonium (TPMP+) by either filtration (6) or flow dialysis. Calculations. Concentration gradients for solutes taken up by the vesicles were calculated using a value of 2.2 ,ul of intravesicular fluid per mg of membrane protein (25). Internal pH was calculated as described by Waddel and Butler (19), and ApH was determined by difference. The electrical potential (A') was calculated from the Nernst equation (A' = 58.8 log [TPMP+]n/[TPMP+]out) using steady-state concentration values obtained from TPMP+ uptake experiments. The proton electrochemical gradient (AUH+) was calculated by substituting calculated values for A* and ApH into equation (1). Protein Determinations were carried out as described (26). Chemicals. [3H]Triphenylmethylphosphonium bromide was prepared by the Isotope Synthesis Group at Hoffmann-La Roche, Inc. under the direction of Dr. Arnold Liebman as described (6). Other isotopically labeled materials were purchased from New England Nuclear and Amersham-Searle. Valinomycin and carbonylcyanide m-chlorophenylhydrazone (CCCP) were obtained from Calbiochem. Nigericin was the generous gift of Dr. J. Berger of Hoffmann-La Roche, Inc. RESULTS Determination of ApH Fig. 1 depicts a typical flow dialysis experiment carried out as described in Methods. Shortly after [3H]acetate is added to the upper chamber containing membrane vesicles, radioactivity
1893
0 x
.,
0.
15
FRACTION NUMBER
FIG. 1. Ascorbate-PMS-dependent acetate uptake by E. coli ML 308-225 membrane vesicles as determined by flow dialysis. The assay shown was carried out at pH 5.5 as described in Methods with sodium [3H]acetate (685 mCi/mmol) at a final concentration of 18 gM and E. coli ML 308-225 membrane vesicles at a final concentration of 2.5 mg of protein per ml in the upper chamber. As indicated by the arrows, ascorbate and phenazine methosulfate (ASC/PMS), valinomycin (VAL), and nigericin (NIG) were added to the upper chamber at final concentrations of 20 mM and 0.1 mM, 1 ,uM and 1 gM, respectively (closed symbols). Open symbols were obtained from an identical experiment carried out in the absence of ascorbate and PMS. The data have been corrected for a control performed in the absence of membrane vesicles as described in Methods.
appears in the dialysate, increasing linearly for about 2 min, and reaching a maximum which then decreases at a slow and constant rate (open symbols). When ascorbate and PMS are added to the upper chamber (closed symbols), the vesicles accumulate acetate, and its concentration in the dialysate decreases markedly to about 60% of the level observed in the absence of electron donor. Addition of valinomycin, an ionophore which specifically increases the potassium permeability of the membrane (27), causes the vesicles to accumulate more acetate, and its concentration in the dialysate decreases still further to about 50% of the control level. Finally, when nigericin is added, acetate is released from the vesicles, and the external concentration returns to the control level, an observation which is consistent with the notion that nigericin catalyzes an electrically neutral exchange of potassium for protons, and thus collapses ApH (28). Using the equilibrium concentration of acetate observed in the dialysate after addition of ascorbate-PMS (i.e., fraction 25), it can be calculated that the vesicles take up approximately 1.8 nmol of acetate per mg of membrane protein. Making the calculations described in Methods, this value represents an internal pH of 7.5. Since the external pH is 5.5, it is clear that the vesicles generate a ApH (interior alkaline) of approximately 2 units under these conditions. Previous attempts to observe a ApH in this in vitro system with DMO and standard assay techniques have been negative (1, 6). These observations are documented in Table 1 where it it shown that DMO is not accumulated to any extent when uptake is assayed by filtration or rapid centrifugation. With flow dialysis, however, the ApH observed with DMO is similar to that obtained with acetate. Moreover similar ApH values are obtained with propionate and butyrate, whereas filtration gives values which are increasingly lower for propionate and butyrate relative to acetate. Presumably, the low values obtained by filtration and centrifugation are due to the high passive permeability of the vesicle membrane to weak acids
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Proc. Natl. Acad. Sci. USA 73 (1976)
Table 1. Uptake of different weak acids by E. coli ML 308-225 membrane vesicles as determined by various techniques Uptake assayed by:
Acid Acetate Propionate Butyrate DMO
Flow dialysis* (mV) -114 -110 -118 -127
(3.9) (3.2) (4.3) (6.9)
Filtrationt
(mV) -23.4 -3.3 -2 0
(1.1) (0.5) (0.4) (0)
Centrifugationt (mV) -46 (0.5) n.d. n.d. 0 (0)
Values given in parentheses represent uptake of the weak acids in nmol/mg of membrane protein; n.d., not determined. * Flow dialysis was performed as described in Methods and in Fig. 1 using sodium [3H]acetate (685 mCi/mmol), sodium [1-'4C]propionate (53 mCi/mmol), sodium [1-14C]butyrate (25.8 mCi/ mmol), or [2-14C]DMO (11 mCi/mmol) at final concentrations of 37.5, 37.5, 37.5, and 200 gM, respectively. Internal pH was calculated as described by Waddel and Butler (19). The pK values used in the calculations were as follows: acetate, 4.75; propionate, 4.87; butyrate, 4.81; and DMO, 6.3. ApH was calculated and converted to mV as described in Methods. t Filtration assays were performed as described previously (21, 23) and in Methods with the same isotopically labeled weak acids as given above (except that sodium [1,2-14C]acetate at a specific activity of 57 mCi/mmol was used in place of sodium [3H]acetate) at final concentrations of 200 ,uM. A value of zero indicates that the experimental value was insignificantly different from a control sample that has been diluted 20-fold before addition of radioactive solute and filtered and washed (23). Attempts were also made to carry out filtration assays without washing the samples on the filters. Results obtained in this fashion were so variable that the experiments were not reproducible. t Centrifugation assays were carried out in the following manner: aliquots of membrane vesicles (50 ,l containing 0.2-0.4 mg of membrane protein) were diluted to a final volume of 100 Al containing (in final concentrations) 0.05 M potassium phosphate (pH 5.5) and 0.01 M magnesium sulfate. The samples were incubated in an oxygen atmosphere at 250 for about 30 sec, and ascorbate and PMS were added to final concentrations of 20 mM and 0.1 mM, respectively. Immediately thereafter, sodium
[3H]acetate (685 mCi/mmol) or [2-14C]DMO (11 mCi/mmol) were added to final concentrations of 37.5 and 200 AM, respectively. The incubations were continued for 5 min, at which time the samples were transferred as rapidly as possible to centrifuge tubes by means of a Hamilton syringe (5-10 sec). Each sample was then centrifuged in a Beckman Airfuge at about 160,000 x g for about 15 sec (a total of approximately 2 min was required from the time the tubes were placed into the centrifuge until they were removed). The supernatants were discarded, and the pellets were resuspended in 100 ,A water and aliquots (70 Ml) were assayed for radioactivity. Values were corrected for a control sample treated identically in the presence of 1 MM nigericin and 5 AM valinomycin.
(filtration) and to the sensitivity of ApH to oxygen tension (centrifugation). In any case, it should be emphasized that such differences are not observed with TPMP+ or lactose which are accumulated to the same extent when assayed by filtration or flow dialysis. Finally, ApH values obtained by flow dialysis are constant over at least a 100-fold range of weak acid concentrations and over a range of membrane protein concentrations from 1 to 3 mg/ml;. and none of the weak acids utilized is metabolized by the vesiclest. t Membrane vesicles were incubated for 10 min with each of the weak acids as described in Table 1. Aliquots of the reaction mixtures were chromatographed on thin-layer chromatography plates coated with silica gel G and radioautographed as described previously (29), except that the solvent system used was water-saturated ethyl ether/ ammonium hydroxide (7:1, vol/vol).
z0. z
E
w9
EXTERNAL pH
FIG. 2. Effect of external pH on ApH, internal pH, /v, and AjH+. ApH values (0-0) were calculated from flow dialysis experiments carried out with [3H]acetate at the pH values given as described in Fig. 1 and Methods. Internal pH (v-v) was calculated from values for acetate uptake determined at each pH as described in Methods. AdI values (0-*) were calculated from flow dialysis experiments carried out with [3H]TPMP+ (1.33 Ci/mmol) at a final concentration of 24MuM as described in Fig. 1 and Methods, and from filtration assays. The filtration assays were carried out after 5 and 10 min incubations with ascorbate (20 mM) and PMS (0.1 mM) performed as described previously (6) except that [3H]TPMP+ (1.33 Ci/dimol) was used at a final concentration of 24 ,M. Similar results were obtained using flow dialysis and filtration assays. A4H+ values (A-A) were calculated from ApH and A'I as described in Methods.
Effect of external pH on ApH, A', internal pH, and AAH+ It is apparent from Fig. 2 that ApH varies markedly with external pH, as reported by Padan et al. (17) for intact cells. From pH 5.0 to 5.5, ApH remains almost constant at -114 to -113 mV; above pH 5.5, ApH decreases drastically, and is negligible at pH 7.5 and above. Since ascorbate-PMS-dependent [14C]methylamine uptake is not observed at pH 7.5 or above (data not shown), ApH does not become reversed in the vesicles at high external pH as reported for intact cells (17). Significantly, despite marked variation in ApH as a function of external pH, internal pH remains essentially constant at pH 7.5, and AI ranges from a low of about -65 mV at pH 5.0 to a high of about -75 mV at pH 7.0. As a result of these individual variations, AOH+ exhibits a maximum value of -180 mV at pH 5.5 (-110 mV ApH + -70 mV AI) which decreases to a constant value of about -75 mV at pH 7.5 and above (O ApH + -75 mV A'i). Effect of various electron donors on ApH, AI, and Although there is little relationship between the ability of the vesicles to oxidize an electron donor and the ability of that electron donor to drive active transport, a qualitative correlation exists between the ability of various electron donors to drive transport and their ability to generate a AI (interior negative) (6). The data presented in Table 2 corroborate the latter ohservation and demonstrate further that a similar relationship exists with respect to ApH and AAH+. Clearly, ascorbate-PMS and D-lactate produce maximal relative values for each parameter, whereas succinate and especially NADH produce
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Proc. Natl. Acad. Sci. USA 73 (1976)
Table 2. Effect of various electron donors on ApH, A4', and 4LH+
Electron donor
ApH*
(mV) AV
Ascorbate-PMS D-Lactate Succinate NADH
-115 -102 0 0 -59
-74 -70 -64 0 -62
NADH + Q1
4 -189 -172 -64 0 -131
* ApH was calculated from flow dialysis experiments carried out with sodium [6H]acetate (685 mCi/mmol) at a final concentration of i8 AM as described in Methods and in Fig. 1. Ascorbate and PMS, lithium-D-lactate, sodium succinate, NADH (sodium salt), and ubiquinone-1 (Q1) were used at final concentrations of 20 mM and 0.1 mM, 20,mM, 20 mM, 5 mM, and 0.08 mM, respectively. t A'I was determined from filtration assays performed after 5 and 1G min incubations as described previously (6) except that [3H]TPMP+ (1.33 Ci/mmol) was used at a final concentration of 24 AM. Electron donors were used at the concentrations given above, and the data were corrected f6r the amount of TPMP+ taken up in the absence of exogenous electron donors. t AUH + was calculated as described in Methods.
much weaker effects. Moreover, when ubiquinone-1 is added to the vesicles, NADH generates much higher values. This observation is highly significant in that NADH oxidation under these conditions drives transport in the presence of ubiquinone-i, but not in its absence (11). Effect of valinomycin, nigericin, and CCCP on 4pH, AI, and AMH+ As increasing concentrations of valinomycin are added to ves-
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ides at pH 5.5 or 7.5, ascorbate-PMS-dependent TPMP+ uptake decreases to approximately 10% of the control value at 2.5-5 AM valinomycin (Fig. 3A), corresponding to a decrease in A' from about -70 mV to -20 mV (insert). Strikingly, there is an increase in acetate uptake to about 130% of the control value at 1 MM valinomycin [an increase in ApH of about 10 mV (insert)] which remains constant, at higher valinomycin concentrations. The effect of valinomycin on the total driving force at pH 5.5 is relatively small, producing only about a 20% loss at 5 MM valinomycin [from a AMH+ of about -190 mV to -150 mV (insert)]. As shown in Fig. 3B, nigericin induces effects which are opposite to those of valinomycin. Acetate uptake decreases to 0 as the nigericin concentration is increased from 0 to 0.1 AM [from a ApH of -110 mV to 0 (insert)], whereas TPMP+ uptake increases almost 2.5-fold over the same concentration range [from a AI of about -60 mV to -90 mV (insert)]. Despite marked effect on ApH and A+, 0.1 MM nigericin produces only a 45% decrease in the total driving force at pH 5.5 [from a AMH+ of -170 mV to -90 mV (insert)]. It is also apparent that this ionophore has no effect on TPMP+ uptake at pH 7.5, a finding which is consistent with the absence of a ApH at this external pH (see Fig. 2). The proton conductor CCCP inhibits both TPMP+ and acetate uptake at pH 5.5 over a concentration range from 0 to 1 MM CCCP, and diminishes the total driving force at pH 5.5 by approximately 60% (a decrease in AAH+ from -170 mV to about -65 mV at 1 MM CCCP) (Fig. SC). It is also noteworthy that CCCP is significantly more effective at pH 5.5 relative to pH 7.5, and that at pH 5.5, ApH (acetate uptake) is inhibited more markedly than A' (TPMP+ uptake). The extent of inhibition of AAH+ observed at 1 AM CCCP is similar to that observed with many of the vesicular transport systems (30).
FIG. 3. Effect of valinomycin (A), nigericin (B), and CCCP (C) on ApH, ASip, and UH+. ApH was determined by flow dialysis in the presence of acetate and ascorbate-PMS at pH 5.5, and the indicated concentrations of valinomycin, nigericin, or CCCP as described in Fig. 1 and ii Methods. Steady-state levels of TPMP+ accumulation (A+) were determined at pH 5.5 (A-A) and pH 7.5 (v-v) in the presence of given concentrations of valinomycin, nigericin, or CCCP by filtration assays as described in Fig. 2. The effect of the inhibitors on the total driving force at pH 5.5 (AiH+, 0-0) was calculated from ApH and AI at each inhibitor concentration. The percentage of the total driving force remaining at each inhibitor concentration is the percentage of AAH+ measured in the absence of inhibitors. The following control values (100%) were obtained in the absence of inhibitors (nmol/mg of membrane protein): acetate uptake 1.6; TPMP+ uptake, 0.80 (at pH 5.5) and 0.91 (at pH 7.5). The mV values plotted in the inserts were calculated from the experimental values presented.
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DISCUSSION These results demonstrate that E. coli ML 308-225 membrane vesicles generate a considerable ApH when assayed under the proper conditions. When considered together with certain previous observations (1, 6) it seems eminently clear that some of the apparent shortcomings of the chemiosmotic hypothesis are resolved. To wit, AIH+ is thermodynamically sufficient to account for the magnitude of solute accumulation by the vesicles at pH 6.6, where most of the transport studies have been carried out. The findings also have an immediate bearing on the question of the polarity of bacterial membrane vesicles prepared by the methods developed in this laboratory (21, 22). Since the AaSH+ established by the vesicles is at least as high as that reported for intact E. colh (17), it seems extremely unlikely that a significant number of the structures can be inverted or seriously damaged. Finally, this work has provided a framework within which to test other more specific aspects of the relationship between A4H+ and active transport. Although space limitations preclude a detailed presentation or discussion of the data (S. Ramos and H. R. Kaback, manuscript in preparation), it should be obvious that valinomycin and nigericin titrations of various transport activities at pH 5.5 and pH 7.5 should yield considerable insight into the coupling between individual components of AAH+ and the accumulation of particular solutes. Such studies have been performed with many vesicular transport systems, and it is clear that the transport systems fall into two groups when assayed at pH 5.5, i.e., those that are driven by ASH+ and those that are driven by ApH. It is also significant that each of the transport systems exhibits considerable activity at pH 7.5 where AAH+ is composed solely of a Ai component, and that valinomycin or nigericin inhibits, stimulates, or produces no effect depending upon the external pH. These effects are particularly striking with respect to the active transport of organic anions. With glucose-6-P, for example, there is dramatic stimulation of accumulation by valinomycin at pH 5.5, and dramatic inhibition at pH 7.5. Nigericin, on the other hand, inhibits glucose-6-P accumulation at pH 5.5, but has no effect at pH 7.5. Thus, it is clear that anion transport is driven by ApH at pH 5.5 or by A*I at pH 7.5.
Proc. Natl. Acad. Sci. USA 73 (1976) 1. Kaback, H. R. (1972) Biochim. Blophys. Acta 265,367-416. 2. Kaback, H. R. & Hong, J.-s. (1973) in CRC Critical Reviews in
3. 4. 5. 6.
Microbiology, eds. Laskin, A. I. & Lechevalier, H. (CRC Press, Ohio), Vol. 2, pp. 333-376. Lombardi, F. J., Reeves, J. P., Short, S. A. & Kaback, H. R. (1974) Ann. N.Y. Acad. Sci. 227,312-327. Kaback, H. R. (1974) Science 186,882-892. Patel, L., Schuldiner, S. & Kaback, H. R. (1975) Proc. Natl. Acad. Sci. USA 72,3387-3391. Schuldiner, S. & Kaback, H. R. (1975) Biochemistry 14,54515461.
7. Mitchell, P. (1967) Nature 191, 144-148. 8. Mitchell, P. (1966) Biol. Rev. Cambridge Philos. Soc. 47,445-502. 9. Mitchell, P. (1973) J. Bioenerg. 4,63-91. 10. Harold, F. M. (1972) Bacteriol. Rev. 36, 172-230. 11. Stroobant, P. & Kaback, H. R. (1975) Proc. Natl. Acad. Sci. USA
72,3970-3974. 12. Reeves, J. P. (1971) Biochem. Biophys. Res. Commun. 45, 931-936. 13. Hirata, H., Altendorf, K. & Harold, F. M. (1973) Proc. Natl. Acad. Sci. USA 70,1804-1808. 14. Altendorf, K., Hirata, H. & Harold, F. M. (1975) J. Biol. Chem.
250, 1405-1412.
15. Schuldiner, S., Rudnick, G., Weil, R. & Kaback, H. R. (1976) Trends in Biochemical Sciences, 1, 41-45.
16. Lombardi, F. J. & Kaback, H. R. (1973) J. Biol. Chem. 247, 7844-7857. 17. Padan, E., Zilberstein, D. & Rottenberg, H. (1976) Eur. J. Biochem., 63,533-541. 18. Rottenberg, H. (1975) J. Bioenerg. 7,61-74. 19. Waddel, W. J. & Butler, T. C. (1959) J. Clin. Invest. 38,720-729. 20. Colowick, S. P. & Womack, F. C. (1969) J. Biol. Chem. 244, 774-777. 21. Kaback, H. R. (1971) in Methods in Enzymology, ed. Jakoby, W. B. (Academic Press, New York, New York), Vol XXII, pp. 99-120. 22. Short, S. A., Kaback, H. R. & Kohn, L. D. (1975) J. Biol. Chem. 250,4291-4296. 23. Kaback, H. R. (1974) in Methods in Enzymology, eds. Fleischer, S. & Packer, L. (Academic Press, New York, New York), Vol. XXXI, pp. 698-709. 24. Schuldiner, S., Weil, R. & Kaback, H. R. (1976) Proc. Natl. Acad. Sci. USA 73, 109-112. 25. Kaback, H. R. & Barnes, E M., Jr. (1971) J. Biol. Chem. 246, 5523-5531. 26. Lowry, 0. H., Rosebrough, N. J., Farr, A. L. & Randall, R. J.
(1951) J. Biol. Chem. 193,265-275.
We are indebted to Dr. Hagai Rottenberg of Tel Aviv University for suggesting the use of acetate, and for helpful discussions. We are also indebted to Dr. Etana Padan of The Hebrew University for allowing us access to her data prior to publication. Finally, we thank Dr. Arnold Liebman and The Isotope Synthesis Group of Hoffmann-La Roche for the preparation of [3H]TPMP+. S. R. is a postdoctoral fellow of the Ministerio de Educacion y Ciencia of Spain.
27. Harold, F. M. (1970) Adv. Microbiol. Physiol. 4,45-104. 28. Harold, F. M., Altendorf, K. H. & Hirata, H. (1974) Ann. N.Y. Acad. Sci. 235, 149-160. 29. Kaback, H. R. & Milner, L. S. (1970) Proc. NatI. Acad. Sci. USA
66, 1008-1012. 30. Kaback, H. R., Reeves, J. P., Short, S. A. & Lombardi, F. J. (1974)
Arch. Biochem. Biophys. 160,215-222.
The electrochemical gradient of protons and its relationship to active transport in Escherichia coli membrane vesicles (flow dialysis/membrane potential/energy transduction/lipophilic cations/weak acids)
SOFIA RAMOS, SHIMON SCHULDINER*, AND H. RONALD KABACK The Roche Institute of Molecular Biology, Nutley, New Jersey 07110
Communicated by B. L. Horecker, March 17, 1976
ABSTRACT Membrane vesicles isolated from E. coli generate a trans-membrane proton gradient of 2 pH units under appropriate conditions when assayed by flow dialysis. Using the distribution of weak acids to measure the proton gradient (ApH) and the distribution of the lipophilic cation triphenylmethylphosphonium to measure the electrical potential across the membrane (AI), the vesicles are shown to generate an electrochemical proton gradient (AiH+) of approximately -180 mV at pH 5.5 in the presence of ascorbate and phenazine methosulfate, the major component of which is a ApH of about -110 mV. As external pH is increased, ApH decreases, reaching o at pH 7.5 and above, while AI remains at about -75 mV and internal pH remains at pH 7.5. Moreover, the ability of various electron donors to drive transport is correlated with their ability to generate A4H+. In addition, ApH and Ad can be varied reciprocally in the presence of valinomycin and nigericin. These data and others (manuscript in preparation) provide convincing support for the role of chemiosmotic phenomena in active transport. Despite apparently contradictory initial observations (1-3), an increasing accumulation of experimental evidence (4-6) indicates that chemiosmotic phenomena, as postulated by Mitchell (7-10), play a central role in respiration-linked active transport in Escherichia coli membrane vesicles. It now seems clearly established that membrane vesicles prepared by the techniques developed in this laboratory retain the same orientation as the membrane in the intact cell (see ref. 11 for a summary of the evidence), and that oxidation of electron donors which drive transport in the vesicles results in the generation of a transmembrane electrical potential (interior negative) by means of electrogenic proton extrusion (6, 12-14). The potential is postulated to drive solute accumulation via facilitated diffusion of positively charged substrates such as lysine or via coupled movements of protons with neutral substrates such as lactose or proline (i.e., "symport"). According to the chemiosmotic hypothesis, the total driving force generated by proton extrusion is the electrochemical potential of protons across the membrane (A,4H+) (7-10). This thermodynamic entity is composed of an electrical and a chemical parameter according to the following relationship: = A A =H+
2.3RTA F ApH
[1]
where Ai represents the electrical potential across the membrane, and ApH is the chemical difference in proton concentration across the membrane (2.3RT/F is equal to 58.8 mV at room temperature). Through the use of lipophilic cations and rubidium (in the
presence of valinomycin), a respiration-dependent membrane potential (AI, interior negative) of approximately -75 mV in E. coli membrane vesicles has been documented (6, 13, 14). Moreover it has been shown that the potential causes the appearance of high affinity binding sites for dansyl- and azidophenylgalactosides on the outer surface of the membrane (4, 15) and that the potential is partially dissipated as a result of lactose accumulation (6). Although these findings provide evidence for the chemiosmotic hypothesis, it has also been demonstrated (6, 16) that vesicles are able to accumulate lactose and other substrates to intravesicular concentrations which are 100-fold or greater than those of the external medium. To sustain concentration gradients of this magnitude, a membrane potential of at least -120 mV is required. This observation, in addition so numerous negative attempts to establish the existence of a transmembrane pH gradient (1, 6) has left some doubt as to the quantitative relationship between AILH+ and solute accumulation. [In the apparent absence of a transmembrane pH gradient, it is also not immediately clear how anionic solutes are accumulated, by the chemiosmotic mechanism. suggested for this process (10)]. Recently, Padan et al. (17) have shown that intact E. coli generate a ApH (interior alkaline), and that the magnitude of the ApH is very dependent upon external pH, exhibiting a maximal value of about 2 pH units at pH 6.0 or below. In addition, Rottenberg (18) has utilized acetate to determine ApH in mitochondria, and suggested that this weak acid might be more useful than 5,5'-dimethyloxazolidine-2,4-dione (DMO) (19) in certain systems because it might be less permeant (H. Rottenberg; personal communication). The results presented in this paper were obtained by means of flow dialysis (20), a technique which allows a rapid, continuous determination of changes in the concentration of solutes in the external medium without manipulation of the vesicles. Using this technique, it is demonstrated that E. coli membrane vesicles generate a large proton gradient under appropriate conditions. In addition, we have shown that ApH and AI can be manipulated reciprocally by the ionophores valinomycin and nigericin. The results to be presented, and others (S. Ramos and H. R. Kaback, manuscript in preparation) which will be discussed, leave little doubt as to the primary role of chemiosmotic phenomena in respiration-dependent active transport. METHODS Growth of Cells and Preparation of Membrane Vesicles. E. coli ML 308-225 (i-z-y+a+) was grown on minimal medium A with 1.0% sodium succinate (hexahydrate), and membrane vesicles were prepared as described (21, 22). Vesicles were suspended in 0.1 M potassium phosphate (pH 6.6) and stored in liquid nitrogen. For studies at various pH's, membrane suspensions containing about 4 mg of protein per ml were removed from storage,
Abbreviations: DMO, 5,.5'-dimethyloxazolidine-2,4-dione; TPMP',
triphenylmethylphosphonium (bromide salt); PMS, phenazine methosulfate; CCCP, carbonylcyanide m-chlorophenylhydrazone. * Present address: Department of Molecular Biology, Hadassah Medical School, Hebrew University, Jerusalem, Israel. 1892
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Ramos et al.
thawed rapidly at 460, diluted at least 10-fold with 0.1 M potassium phosphate buffer at the desired pH, and incubated for 10 min at 25°. The suspension was centrifuged at 40,000 X g for 30 min, and the pellet resuspended and washed once in a similar volume of the same buffer. The final pellet was then resuspended to an appropriate protein concentration in 0.1 M potassium phosphate at the same pH. Transport Assays. Filtration assays (6, 23) were carried out on Millipore Cellotate filters (0.5 ,um pore size). Electron donors and isotopically labeled solutes were used as described. Flow dialysis was performed as described (24) except that the apparatus was modified so that the upper chamber was completely open to the atmosphere, and the reaction mixture was gassed with oxygen. The upper and lower chambers were separated by Spectrapor 1 dialysis tubing (600-8000 molecular weight cut-off; Fisher Scientific), and both chambers were stirred by means of magnetic bars. Membrane vesicles suspended in 0.05 M potassium phosphate at a given pH containing 0.'01 M magnesium sulfate were added to the upper chamber (total volume 0.8 ml), and electron donors, isotopically labeled solutes, and ionophores were added as indicated. Potassium phosphate (0.05 M at the same pH as the buffer in the upper chamber) was pumped from the lower chamber at a rate of 6.0 ml/min with a Pharmacia pump (model P3). Fractions of about 1.7 ml were collected and assayed for radioactivity by liquid scintillation spectrometry. Since phenazine methosulfate (PMS) causes about 5.5% quenching of tritium under the conditions described, control experiments were carried out in the absence of membrane vesicles, and the data were corrected appropriately. Determination of ApH. ApH.was determined by measuring the accumulation of acetate, propionate, butyrate or DMO by flow dialysis unless otherwise stated. Data were quantitated assuming that dialysis rates obtained after addition of nigericin (Fig. 1) represent 0 ApH. Determination of AI. The electrical potential across the membrane (Au') was determined by measuring the accumulation of [3H]triphenylmethylphosphonium (TPMP+) by either filtration (6) or flow dialysis. Calculations. Concentration gradients for solutes taken up by the vesicles were calculated using a value of 2.2 ,ul of intravesicular fluid per mg of membrane protein (25). Internal pH was calculated as described by Waddel and Butler (19), and ApH was determined by difference. The electrical potential (A') was calculated from the Nernst equation (A' = 58.8 log [TPMP+]n/[TPMP+]out) using steady-state concentration values obtained from TPMP+ uptake experiments. The proton electrochemical gradient (AUH+) was calculated by substituting calculated values for A* and ApH into equation (1). Protein Determinations were carried out as described (26). Chemicals. [3H]Triphenylmethylphosphonium bromide was prepared by the Isotope Synthesis Group at Hoffmann-La Roche, Inc. under the direction of Dr. Arnold Liebman as described (6). Other isotopically labeled materials were purchased from New England Nuclear and Amersham-Searle. Valinomycin and carbonylcyanide m-chlorophenylhydrazone (CCCP) were obtained from Calbiochem. Nigericin was the generous gift of Dr. J. Berger of Hoffmann-La Roche, Inc. RESULTS Determination of ApH Fig. 1 depicts a typical flow dialysis experiment carried out as described in Methods. Shortly after [3H]acetate is added to the upper chamber containing membrane vesicles, radioactivity
1893
0 x
.,
0.
15
FRACTION NUMBER
FIG. 1. Ascorbate-PMS-dependent acetate uptake by E. coli ML 308-225 membrane vesicles as determined by flow dialysis. The assay shown was carried out at pH 5.5 as described in Methods with sodium [3H]acetate (685 mCi/mmol) at a final concentration of 18 gM and E. coli ML 308-225 membrane vesicles at a final concentration of 2.5 mg of protein per ml in the upper chamber. As indicated by the arrows, ascorbate and phenazine methosulfate (ASC/PMS), valinomycin (VAL), and nigericin (NIG) were added to the upper chamber at final concentrations of 20 mM and 0.1 mM, 1 ,uM and 1 gM, respectively (closed symbols). Open symbols were obtained from an identical experiment carried out in the absence of ascorbate and PMS. The data have been corrected for a control performed in the absence of membrane vesicles as described in Methods.
appears in the dialysate, increasing linearly for about 2 min, and reaching a maximum which then decreases at a slow and constant rate (open symbols). When ascorbate and PMS are added to the upper chamber (closed symbols), the vesicles accumulate acetate, and its concentration in the dialysate decreases markedly to about 60% of the level observed in the absence of electron donor. Addition of valinomycin, an ionophore which specifically increases the potassium permeability of the membrane (27), causes the vesicles to accumulate more acetate, and its concentration in the dialysate decreases still further to about 50% of the control level. Finally, when nigericin is added, acetate is released from the vesicles, and the external concentration returns to the control level, an observation which is consistent with the notion that nigericin catalyzes an electrically neutral exchange of potassium for protons, and thus collapses ApH (28). Using the equilibrium concentration of acetate observed in the dialysate after addition of ascorbate-PMS (i.e., fraction 25), it can be calculated that the vesicles take up approximately 1.8 nmol of acetate per mg of membrane protein. Making the calculations described in Methods, this value represents an internal pH of 7.5. Since the external pH is 5.5, it is clear that the vesicles generate a ApH (interior alkaline) of approximately 2 units under these conditions. Previous attempts to observe a ApH in this in vitro system with DMO and standard assay techniques have been negative (1, 6). These observations are documented in Table 1 where it it shown that DMO is not accumulated to any extent when uptake is assayed by filtration or rapid centrifugation. With flow dialysis, however, the ApH observed with DMO is similar to that obtained with acetate. Moreover similar ApH values are obtained with propionate and butyrate, whereas filtration gives values which are increasingly lower for propionate and butyrate relative to acetate. Presumably, the low values obtained by filtration and centrifugation are due to the high passive permeability of the vesicle membrane to weak acids
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Proc. Natl. Acad. Sci. USA 73 (1976)
Table 1. Uptake of different weak acids by E. coli ML 308-225 membrane vesicles as determined by various techniques Uptake assayed by:
Acid Acetate Propionate Butyrate DMO
Flow dialysis* (mV) -114 -110 -118 -127
(3.9) (3.2) (4.3) (6.9)
Filtrationt
(mV) -23.4 -3.3 -2 0
(1.1) (0.5) (0.4) (0)
Centrifugationt (mV) -46 (0.5) n.d. n.d. 0 (0)
Values given in parentheses represent uptake of the weak acids in nmol/mg of membrane protein; n.d., not determined. * Flow dialysis was performed as described in Methods and in Fig. 1 using sodium [3H]acetate (685 mCi/mmol), sodium [1-'4C]propionate (53 mCi/mmol), sodium [1-14C]butyrate (25.8 mCi/ mmol), or [2-14C]DMO (11 mCi/mmol) at final concentrations of 37.5, 37.5, 37.5, and 200 gM, respectively. Internal pH was calculated as described by Waddel and Butler (19). The pK values used in the calculations were as follows: acetate, 4.75; propionate, 4.87; butyrate, 4.81; and DMO, 6.3. ApH was calculated and converted to mV as described in Methods. t Filtration assays were performed as described previously (21, 23) and in Methods with the same isotopically labeled weak acids as given above (except that sodium [1,2-14C]acetate at a specific activity of 57 mCi/mmol was used in place of sodium [3H]acetate) at final concentrations of 200 ,uM. A value of zero indicates that the experimental value was insignificantly different from a control sample that has been diluted 20-fold before addition of radioactive solute and filtered and washed (23). Attempts were also made to carry out filtration assays without washing the samples on the filters. Results obtained in this fashion were so variable that the experiments were not reproducible. t Centrifugation assays were carried out in the following manner: aliquots of membrane vesicles (50 ,l containing 0.2-0.4 mg of membrane protein) were diluted to a final volume of 100 Al containing (in final concentrations) 0.05 M potassium phosphate (pH 5.5) and 0.01 M magnesium sulfate. The samples were incubated in an oxygen atmosphere at 250 for about 30 sec, and ascorbate and PMS were added to final concentrations of 20 mM and 0.1 mM, respectively. Immediately thereafter, sodium
[3H]acetate (685 mCi/mmol) or [2-14C]DMO (11 mCi/mmol) were added to final concentrations of 37.5 and 200 AM, respectively. The incubations were continued for 5 min, at which time the samples were transferred as rapidly as possible to centrifuge tubes by means of a Hamilton syringe (5-10 sec). Each sample was then centrifuged in a Beckman Airfuge at about 160,000 x g for about 15 sec (a total of approximately 2 min was required from the time the tubes were placed into the centrifuge until they were removed). The supernatants were discarded, and the pellets were resuspended in 100 ,A water and aliquots (70 Ml) were assayed for radioactivity. Values were corrected for a control sample treated identically in the presence of 1 MM nigericin and 5 AM valinomycin.
(filtration) and to the sensitivity of ApH to oxygen tension (centrifugation). In any case, it should be emphasized that such differences are not observed with TPMP+ or lactose which are accumulated to the same extent when assayed by filtration or flow dialysis. Finally, ApH values obtained by flow dialysis are constant over at least a 100-fold range of weak acid concentrations and over a range of membrane protein concentrations from 1 to 3 mg/ml;. and none of the weak acids utilized is metabolized by the vesiclest. t Membrane vesicles were incubated for 10 min with each of the weak acids as described in Table 1. Aliquots of the reaction mixtures were chromatographed on thin-layer chromatography plates coated with silica gel G and radioautographed as described previously (29), except that the solvent system used was water-saturated ethyl ether/ ammonium hydroxide (7:1, vol/vol).
z0. z
E
w9
EXTERNAL pH
FIG. 2. Effect of external pH on ApH, internal pH, /v, and AjH+. ApH values (0-0) were calculated from flow dialysis experiments carried out with [3H]acetate at the pH values given as described in Fig. 1 and Methods. Internal pH (v-v) was calculated from values for acetate uptake determined at each pH as described in Methods. AdI values (0-*) were calculated from flow dialysis experiments carried out with [3H]TPMP+ (1.33 Ci/mmol) at a final concentration of 24MuM as described in Fig. 1 and Methods, and from filtration assays. The filtration assays were carried out after 5 and 10 min incubations with ascorbate (20 mM) and PMS (0.1 mM) performed as described previously (6) except that [3H]TPMP+ (1.33 Ci/dimol) was used at a final concentration of 24 ,M. Similar results were obtained using flow dialysis and filtration assays. A4H+ values (A-A) were calculated from ApH and A'I as described in Methods.
Effect of external pH on ApH, A', internal pH, and AAH+ It is apparent from Fig. 2 that ApH varies markedly with external pH, as reported by Padan et al. (17) for intact cells. From pH 5.0 to 5.5, ApH remains almost constant at -114 to -113 mV; above pH 5.5, ApH decreases drastically, and is negligible at pH 7.5 and above. Since ascorbate-PMS-dependent [14C]methylamine uptake is not observed at pH 7.5 or above (data not shown), ApH does not become reversed in the vesicles at high external pH as reported for intact cells (17). Significantly, despite marked variation in ApH as a function of external pH, internal pH remains essentially constant at pH 7.5, and AI ranges from a low of about -65 mV at pH 5.0 to a high of about -75 mV at pH 7.0. As a result of these individual variations, AOH+ exhibits a maximum value of -180 mV at pH 5.5 (-110 mV ApH + -70 mV AI) which decreases to a constant value of about -75 mV at pH 7.5 and above (O ApH + -75 mV A'i). Effect of various electron donors on ApH, AI, and Although there is little relationship between the ability of the vesicles to oxidize an electron donor and the ability of that electron donor to drive active transport, a qualitative correlation exists between the ability of various electron donors to drive transport and their ability to generate a AI (interior negative) (6). The data presented in Table 2 corroborate the latter ohservation and demonstrate further that a similar relationship exists with respect to ApH and AAH+. Clearly, ascorbate-PMS and D-lactate produce maximal relative values for each parameter, whereas succinate and especially NADH produce
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Proc. Natl. Acad. Sci. USA 73 (1976)
Table 2. Effect of various electron donors on ApH, A4', and 4LH+
Electron donor
ApH*
(mV) AV
Ascorbate-PMS D-Lactate Succinate NADH
-115 -102 0 0 -59
-74 -70 -64 0 -62
NADH + Q1
4 -189 -172 -64 0 -131
* ApH was calculated from flow dialysis experiments carried out with sodium [6H]acetate (685 mCi/mmol) at a final concentration of i8 AM as described in Methods and in Fig. 1. Ascorbate and PMS, lithium-D-lactate, sodium succinate, NADH (sodium salt), and ubiquinone-1 (Q1) were used at final concentrations of 20 mM and 0.1 mM, 20,mM, 20 mM, 5 mM, and 0.08 mM, respectively. t A'I was determined from filtration assays performed after 5 and 1G min incubations as described previously (6) except that [3H]TPMP+ (1.33 Ci/mmol) was used at a final concentration of 24 AM. Electron donors were used at the concentrations given above, and the data were corrected f6r the amount of TPMP+ taken up in the absence of exogenous electron donors. t AUH + was calculated as described in Methods.
much weaker effects. Moreover, when ubiquinone-1 is added to the vesicles, NADH generates much higher values. This observation is highly significant in that NADH oxidation under these conditions drives transport in the presence of ubiquinone-i, but not in its absence (11). Effect of valinomycin, nigericin, and CCCP on 4pH, AI, and AMH+ As increasing concentrations of valinomycin are added to ves-
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ides at pH 5.5 or 7.5, ascorbate-PMS-dependent TPMP+ uptake decreases to approximately 10% of the control value at 2.5-5 AM valinomycin (Fig. 3A), corresponding to a decrease in A' from about -70 mV to -20 mV (insert). Strikingly, there is an increase in acetate uptake to about 130% of the control value at 1 MM valinomycin [an increase in ApH of about 10 mV (insert)] which remains constant, at higher valinomycin concentrations. The effect of valinomycin on the total driving force at pH 5.5 is relatively small, producing only about a 20% loss at 5 MM valinomycin [from a AMH+ of about -190 mV to -150 mV (insert)]. As shown in Fig. 3B, nigericin induces effects which are opposite to those of valinomycin. Acetate uptake decreases to 0 as the nigericin concentration is increased from 0 to 0.1 AM [from a ApH of -110 mV to 0 (insert)], whereas TPMP+ uptake increases almost 2.5-fold over the same concentration range [from a AI of about -60 mV to -90 mV (insert)]. Despite marked effect on ApH and A+, 0.1 MM nigericin produces only a 45% decrease in the total driving force at pH 5.5 [from a AMH+ of -170 mV to -90 mV (insert)]. It is also apparent that this ionophore has no effect on TPMP+ uptake at pH 7.5, a finding which is consistent with the absence of a ApH at this external pH (see Fig. 2). The proton conductor CCCP inhibits both TPMP+ and acetate uptake at pH 5.5 over a concentration range from 0 to 1 MM CCCP, and diminishes the total driving force at pH 5.5 by approximately 60% (a decrease in AAH+ from -170 mV to about -65 mV at 1 MM CCCP) (Fig. SC). It is also noteworthy that CCCP is significantly more effective at pH 5.5 relative to pH 7.5, and that at pH 5.5, ApH (acetate uptake) is inhibited more markedly than A' (TPMP+ uptake). The extent of inhibition of AAH+ observed at 1 AM CCCP is similar to that observed with many of the vesicular transport systems (30).
FIG. 3. Effect of valinomycin (A), nigericin (B), and CCCP (C) on ApH, ASip, and UH+. ApH was determined by flow dialysis in the presence of acetate and ascorbate-PMS at pH 5.5, and the indicated concentrations of valinomycin, nigericin, or CCCP as described in Fig. 1 and ii Methods. Steady-state levels of TPMP+ accumulation (A+) were determined at pH 5.5 (A-A) and pH 7.5 (v-v) in the presence of given concentrations of valinomycin, nigericin, or CCCP by filtration assays as described in Fig. 2. The effect of the inhibitors on the total driving force at pH 5.5 (AiH+, 0-0) was calculated from ApH and AI at each inhibitor concentration. The percentage of the total driving force remaining at each inhibitor concentration is the percentage of AAH+ measured in the absence of inhibitors. The following control values (100%) were obtained in the absence of inhibitors (nmol/mg of membrane protein): acetate uptake 1.6; TPMP+ uptake, 0.80 (at pH 5.5) and 0.91 (at pH 7.5). The mV values plotted in the inserts were calculated from the experimental values presented.
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DISCUSSION These results demonstrate that E. coli ML 308-225 membrane vesicles generate a considerable ApH when assayed under the proper conditions. When considered together with certain previous observations (1, 6) it seems eminently clear that some of the apparent shortcomings of the chemiosmotic hypothesis are resolved. To wit, AIH+ is thermodynamically sufficient to account for the magnitude of solute accumulation by the vesicles at pH 6.6, where most of the transport studies have been carried out. The findings also have an immediate bearing on the question of the polarity of bacterial membrane vesicles prepared by the methods developed in this laboratory (21, 22). Since the AaSH+ established by the vesicles is at least as high as that reported for intact E. colh (17), it seems extremely unlikely that a significant number of the structures can be inverted or seriously damaged. Finally, this work has provided a framework within which to test other more specific aspects of the relationship between A4H+ and active transport. Although space limitations preclude a detailed presentation or discussion of the data (S. Ramos and H. R. Kaback, manuscript in preparation), it should be obvious that valinomycin and nigericin titrations of various transport activities at pH 5.5 and pH 7.5 should yield considerable insight into the coupling between individual components of AAH+ and the accumulation of particular solutes. Such studies have been performed with many vesicular transport systems, and it is clear that the transport systems fall into two groups when assayed at pH 5.5, i.e., those that are driven by ASH+ and those that are driven by ApH. It is also significant that each of the transport systems exhibits considerable activity at pH 7.5 where AAH+ is composed solely of a Ai component, and that valinomycin or nigericin inhibits, stimulates, or produces no effect depending upon the external pH. These effects are particularly striking with respect to the active transport of organic anions. With glucose-6-P, for example, there is dramatic stimulation of accumulation by valinomycin at pH 5.5, and dramatic inhibition at pH 7.5. Nigericin, on the other hand, inhibits glucose-6-P accumulation at pH 5.5, but has no effect at pH 7.5. Thus, it is clear that anion transport is driven by ApH at pH 5.5 or by A*I at pH 7.5.
Proc. Natl. Acad. Sci. USA 73 (1976) 1. Kaback, H. R. (1972) Biochim. Blophys. Acta 265,367-416. 2. Kaback, H. R. & Hong, J.-s. (1973) in CRC Critical Reviews in
3. 4. 5. 6.
Microbiology, eds. Laskin, A. I. & Lechevalier, H. (CRC Press, Ohio), Vol. 2, pp. 333-376. Lombardi, F. J., Reeves, J. P., Short, S. A. & Kaback, H. R. (1974) Ann. N.Y. Acad. Sci. 227,312-327. Kaback, H. R. (1974) Science 186,882-892. Patel, L., Schuldiner, S. & Kaback, H. R. (1975) Proc. Natl. Acad. Sci. USA 72,3387-3391. Schuldiner, S. & Kaback, H. R. (1975) Biochemistry 14,54515461.
7. Mitchell, P. (1967) Nature 191, 144-148. 8. Mitchell, P. (1966) Biol. Rev. Cambridge Philos. Soc. 47,445-502. 9. Mitchell, P. (1973) J. Bioenerg. 4,63-91. 10. Harold, F. M. (1972) Bacteriol. Rev. 36, 172-230. 11. Stroobant, P. & Kaback, H. R. (1975) Proc. Natl. Acad. Sci. USA
72,3970-3974. 12. Reeves, J. P. (1971) Biochem. Biophys. Res. Commun. 45, 931-936. 13. Hirata, H., Altendorf, K. & Harold, F. M. (1973) Proc. Natl. Acad. Sci. USA 70,1804-1808. 14. Altendorf, K., Hirata, H. & Harold, F. M. (1975) J. Biol. Chem.
250, 1405-1412.
15. Schuldiner, S., Rudnick, G., Weil, R. & Kaback, H. R. (1976) Trends in Biochemical Sciences, 1, 41-45.
16. Lombardi, F. J. & Kaback, H. R. (1973) J. Biol. Chem. 247, 7844-7857. 17. Padan, E., Zilberstein, D. & Rottenberg, H. (1976) Eur. J. Biochem., 63,533-541. 18. Rottenberg, H. (1975) J. Bioenerg. 7,61-74. 19. Waddel, W. J. & Butler, T. C. (1959) J. Clin. Invest. 38,720-729. 20. Colowick, S. P. & Womack, F. C. (1969) J. Biol. Chem. 244, 774-777. 21. Kaback, H. R. (1971) in Methods in Enzymology, ed. Jakoby, W. B. (Academic Press, New York, New York), Vol XXII, pp. 99-120. 22. Short, S. A., Kaback, H. R. & Kohn, L. D. (1975) J. Biol. Chem. 250,4291-4296. 23. Kaback, H. R. (1974) in Methods in Enzymology, eds. Fleischer, S. & Packer, L. (Academic Press, New York, New York), Vol. XXXI, pp. 698-709. 24. Schuldiner, S., Weil, R. & Kaback, H. R. (1976) Proc. Natl. Acad. Sci. USA 73, 109-112. 25. Kaback, H. R. & Barnes, E M., Jr. (1971) J. Biol. Chem. 246, 5523-5531. 26. Lowry, 0. H., Rosebrough, N. J., Farr, A. L. & Randall, R. J.
(1951) J. Biol. Chem. 193,265-275.
We are indebted to Dr. Hagai Rottenberg of Tel Aviv University for suggesting the use of acetate, and for helpful discussions. We are also indebted to Dr. Etana Padan of The Hebrew University for allowing us access to her data prior to publication. Finally, we thank Dr. Arnold Liebman and The Isotope Synthesis Group of Hoffmann-La Roche for the preparation of [3H]TPMP+. S. R. is a postdoctoral fellow of the Ministerio de Educacion y Ciencia of Spain.
27. Harold, F. M. (1970) Adv. Microbiol. Physiol. 4,45-104. 28. Harold, F. M., Altendorf, K. H. & Hirata, H. (1974) Ann. N.Y. Acad. Sci. 235, 149-160. 29. Kaback, H. R. & Milner, L. S. (1970) Proc. NatI. Acad. Sci. USA
66, 1008-1012. 30. Kaback, H. R., Reeves, J. P., Short, S. A. & Lombardi, F. J. (1974)
Arch. Biochem. Biophys. 160,215-222.
