Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Venema, K.
Right arrow Articles by Palmgren, M. G.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Venema, K.
Right arrow Articles by Palmgren, M. G.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Volume 270, Number 33, Issue of August 18, pp. 19659-19667, 1995
©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Metabolic Modulation of Transport Coupling Ratio in Yeast Plasma Membrane H-ATPase (*)

(Received for publication, March 29, 1995; and in revised form, June 16, 1995)

Kees Venema (§),  ,  Michael G. Palmgren (¶)

From the Department of Plant Biology, Royal Veterinary and Agricultural University, Thorvaldsensvej 40, DK-1871 Frederiksberg C, Copenhagen, Denmark

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

The plasma membrane proton pump (H-ATPase) of yeast energizes solute uptake by secondary transporters and regulates cytoplasmic pH. The addition of glucose to yeast cells stimulates proton efflux mediated by the H-ATPase. A >50-fold increase in proton extrusion from yeast cells is observed in vivo, whereas the ATPase activity of purified plasma membranes is increased maximally 8-fold after glucose treatment (Serrano, R.(1983) FEBS Lett. 156, 11-14). The low capacity of yeast cells for proton extrusion in the absence of glucose can be explained by the finding that, in H-ATPase isolated from glucose-starved cells, ATP hydrolysis is essentially uncoupled from proton pumping. The number of protons transported per ATP hydrolyzed is significantly increased after glucose activation. We suggest that intrinsic uncoupling is an important mechanism for regulation of pump activity.

INTRODUCTION

The H-ATPase in fungal plasma membranes functions physiologically to hydrolyze ATP and to pump H out of the cell; the resulting electrochemical H gradient provides energy for an array of secondary transport systems (Serrano, 1988). Structurally, the fungal plasma membrane ATPase is a member of the P class of ATPases (Pedersen and Carafoli, 1987), which includes the Na/K-ATPase of animal cell membranes, the H/K-ATPase of gastric mucosa, the Ca-ATPase of sarcoplasmic reticulum, and the plasma membrane CaATPases. Like these enzymes, it contains a major M(r) approx100,000 subunit, which is partly embedded within the membrane bilayer. During the reaction cycle, the major subunit is phosphorylated at a conserved aspartate residue, and also the ATPase activity is highly sensitive to vanadate, which resembles the transition state of phosphate (Cantley et al., 1978).

For all ion-translocating ATPases, an important property is the stoichiometric relationship between ions pumped and ATP molecules split. The number of ions transported per ATP hydrolyzed is the prime determinant of the capacity of these pumps to form a gradient (Läuger, 1991). Two approaches have been used to determine such stoichiometries (Briskin and Hanson, 1992; Sanders, 1990). On the one hand, there are kinetic approaches, in which the addition of ATP leads to detectable transport into the lumen of vesicles or organelles. In the case of the Na/K- and Ca-ATPases, net ion fluxes can be measured using radioisotopes, but in the case of the H-ATPases, such measurements are not possible. Instead, by employing weakly buffered membrane suspensions, the pH change in the external solution measured with a pH electrode or the change in intravesicular acidification measured with DeltapH probes has been taken as a measure for net H fluxes. On the other hand, there is a thermodynamic or electrophysiological approach, in which the free energy for ATP hydrolysis is compared with the free energy available in the steady-state ion gradient produced in vivo or in vitro. As a result of these studies, most investigators favor a stoichiometry of 1 H extruded per 1 ATP split for P-type H-ATPases (Sanders, 1990), although it has been suggested that in Neurospora this ratio is modified to 2 H/1 ATP split, by chronic energy restriction (Warncke and Slayman, 1980).

The results presented in this paper show that the net efflux of H per ATP split by the yeast plasma membrane H-ATPase is a flexible rather than a fixed parameter. Apparently, this ratio can attain at least two values determined by the regulatory state of the pump. Our experimental findings can be explained by assuming that, in one of the regulatory states, H pumping is essentially uncoupled from ATP hydrolysis.

MATERIALS AND METHODS

Yeast Growth and Incubation Conditions

Saccharomyces cerevisiae strain BWG1-7A (MATaade1-100 his4-519 leu2-3, 112 ura3-52) (Guarante et al., 1982) was grown to the stationary phase overnight at 30 °C in medium containing 2% glucose, 1% yeast extract (Difco), and 2% Peptone (Difco). Cells were collected by centrifugation for 10 min at 3000 rpm (Sorvall SS-34 rotor) and washed twice with water. Yeast cells (30-150 mg (fresh weight)/ml) were incubated for 10 min at room temperature with mild agitation in water (glucose-starved cells) or in water supplemented with 2% glucose (glucose-activated cells).

Homogenization and Membrane Preparation

Yeast plasma membranes were purified from glucose-metabolizing and glucose-starved cells by differential and sucrose gradient centrifugation (Villalba et al., 1992). All steps were performed at 4 °C. Cells (1 volume) were homogenized by vortexing with glass beads (2 volumes; 0.5-mm diameter) in medium containing 20% (v/v) glycerol, 10 mM Tris-HCl, pH 7.5, 1 mM EDTA, and 1 mM dithiothreitol (buffer GTED 20) supplied with 1 mM phenylmethylsulfonyl fluoride and 0.1 mg/ml chymostatin. Cell debris and glass beads were removed by centrifugation for 10 min at 3000 rpm (Sorvall SS-34 rotor). The supernatant was diluted two times with buffer GTED 20 and centrifuged for 60 min at 40,000 rpm (Beckman Ti-70 rotor). The pellet was resuspended in 1 ml of buffer GTED 20, layered on top of a 12-ml 43/53% (w/w) sucrose step gradient, and centrifuged for 16 h at 30,000 rpm (Kontron TST 41.14 rotor). Plasma membranes were collected from the 43/53% interface and diluted four times with ice-cold water. After centrifugation at 50,000 rpm (Beckman Ti-70 rotor) for 45 min, the pellet was resuspended in buffer GTED 20 supplemented with 1 mM phenylmethylsulfonyl fluoride and 0.1 mg/ml chymostatin. The plasma membrane fraction was frozen in liquid nitrogen and stored at -80 °C.

Reconstitution of Plasma Membrane H-ATPase

All the following steps were performed at room temperature. Mixed soybean phospholipids (30 mg/ml; L-alpha-phosphatidylcholine, type II-S, Sigma) were dispersed by vortexing under argon for 5 min in 10 mM Mes(^1)-KOH, pH 6.5, 50 mM K(2)SO(4), and 20% (v/v) glycerol. Plasma membranes (106 µg of protein) were mixed with lipids at a lipid/protein mass ratio of 22 in a final volume of 208 µl. The protein/lipid mixture was solubilized by the addition of 12 µl of 1 M octyl glucoside (Sigma), giving an effective mass ratio of 3. The mass ratio is given by the following equation: R = ([detergent] - (critical micelle concentration of detergent))/[phospholipids]. Unsolubilized material was removed by spinning the protein/lipid/detergent mixture at 100,000 g for 10 min in a Beckman Airfuge and discarding the pellet. Disposable syringes (2 ml), fitted with siliconized glass wool at the bottom, were filled with Sephadex G-50 (fine, Pharmacia Biotech Inc.) equilibrated in 10 mM Mes-KOH, pH 6.5, 50 mM K(2)SO(4), and 20% (v/v) glycerol and centrifuged for 5 min at 180 g. Solubilized protein/lipid/detergent mixture (220 µl) was applied to the top of the column and centrifuged again for 7.5 min at 180 g. The volume of the eluate recovered was within 80-100% of the volume applied.

The volume of the vesicles was estimated from the fluorescence of trapped pyranine within reconstituted membrane vesicles. The reconstitution was performed as described above but including 25 mM pyranine in the buffer used for reconstitution and Sephadex G-50 equilibration to ascertain the presence of 25 mM pyranine inside eluted vesicles. In a second gel filtration step, dye trapped in vesicles was separated from external dye. Using an approximation for the surface area of 1 phospholipid molecule of 75 Å^2 (Rossignol et al., 1982) and assuming pure phospholipid vesicles, the mean vesicle radius was estimated.

ATPase Assay

ATPase activity was assayed according to a modified protocol of Baginsky et al.(1967) with 1-3 µg of membrane protein at room temperature. The assay medium (100 µl) contained 20 mM Mes, 20 mM Mops, 50 mM KNO(3) (to inhibit vacuolar ATPase), 5 mM NaN(3) (to inhibit mitochondrial ATPase), 3.5 mM Na(2)MoO(4) (to inhibit acid phosphatase), 1 mM Mg free in solution, and the indicated concentrations of MgATP. The pH was adjusted to the desired pH with N-methyl-D-glucamine. After 20 min, the reaction was stopped by the addition of 500 µl of ice-cold stop solution (10 ml of 102 mM ascorbic acid, 0.3 N HCl, 0.065% sodium dodecyl sulfate mixed with 1 ml of 57 mM NH(4)-heptamolybdate to obtain a bright yellow solution). The tubes were incubated for 10 min on ice to allow formation of the P(i)-molybdate complex. Excess molybdate was complexed by the addition of 450 µl of a solution containing 154 mM NaAsO(2), 68 mM trisodium citrate, and 350 mM acetic acid. After 60 min at room temperature, a stable color had developed, and absorbance at 860 nm was determined.

Concentrations of free Mg and MgATP were calculated using and (Morrison, 1979; Wach et al., 1990):

where K(1) = 14.3 µM (dissociation constant, MgATP), K(2) = 1.44 mM (dissociation constant, MgHATP), and K(H) = 0.107 µM (dissociation constant, HATP).

Measurement of DeltapH with Acridine Orange

H transport was assayed by measuring acridine orange absorbance changes at 495 nm (Palmgren, 1990). Changes in the absorbance of acridine orange observed during the formation of pH gradients are due to accumulation of free dye inside the vesicles and subsequent dimerization leading to a spectral shift of this metachromatic dye (Palmgren, 1991). Reconstituted membrane vesicles (50-100 µl; 0.5 mg of protein/ml) were diluted in 1 ml of 20 mM Mes-KOH, pH 6.0, 40 mM K(2)SO(4), 25 mM KNO(3), 30 µM acridine orange (Sigma), 5 mM NaN(3), 3.5 mM Na(2)MoO(4), and (when indicated) 0.5 µM valinomycin. The reaction was started by the addition of 40 µl of 134 mM MgSO(4) and 124 mM ATP, pH 6.0, to obtain a final concentration of 4 mM MgATP and 1 mM free Mg. The developed pH gradient was dissipated by the addition of 0.5 µM nigericin.

Coupled Assay of ATPase Activity and Proton Transport

ATP hydrolysis and H transport were measured simultaneously with a Shimadzu UV-160A spectrophotometer by coupling ATP hydrolysis to NADH oxidation (measured at 340 nm) (Palmgren, 1990). The cycling time between measurements at 340 and 495 nm, respectively, was 12 s. Assay conditions were as described above for H transport measurements employing acridine orange but including 0.3 mM NADH, 2.4 mM phosphoenolpyruvate (neutralized with KOH), 33 µg/ml pyruvate kinase (Boehringer Mannheim 109 045; solution in glycerol), and 33 µg/ml lactate dehydrogenase (Boehringer Mannheim 127 221; solution in glycerol).

Measurement of DeltapH with Pyranine

Pyranine fluorescence was measured with a Perkin-Elmer LS 50B spectrofluorometer at excitation/emission wavelengths of 460/511 nm. Plasma membrane vesicles were reconstituted as indicated, but with 25 mM pyranine (trisodium salt, pH 7.0; Molecular Probes, Inc.) included in the reconstitution buffer. External probe was separated from vesicles during the gel filtration step. The reaction cuvette was thermostatted at 24 °C. The assay medium (3 ml) contained 20 mM Bis-Tris propane-Mes, pH 7.0, 87.5 mM K(2)SO(4), 50 nM valinomycin, and 10% glycerol. Membrane vesicles (10 µl; 5 µg of protein) were added to the reaction medium and incubated until a stable fluorescence signal was observed. The ATPase was energized by the addition of 120 µl of 126 mM MgS0(4) and 104 mM ATP, pH 7.0, giving rise to a final concentration of 4 mM MgATP and 1 mM free Mg.

At the end of the experiment, internal and external pH values were equilibrated by the addition of 80 nM nigericin, and the fluorescence signal was calibrated with pH in the same cuvette by the addition of aliquots of 0.5 N HCl. A calibration equation was made by fitting the fluorescence versus pH data with a second-order polynome, from which the intravesicular pH during the experiment was calculated. In this way, calibration of pyranine fluorescence with intravesicular pH was achieved.

Passive Proton Fluxes and Determination of Passive Proton Permeability

The net proton flux across the membrane was calculated by derivation of the kinetics of pH gradient dissipation taking into account the buffer strength (B) of pyranine (pK 7.2) and Mes (pK 6.1) according to the following equation: J = B (V/A) dpH/dt = B (r/3) dpH/dt, where J is the net proton flux and V, A, and r are the volume, area, and radius of the vesicle, respectively (Venema et al., 1993). In the absence of a surface potential or diffusion potential, which was ascertained in our experiments by high ionic strength and high concentrations of K at both sides of the membrane in the presence of valinomycin, the net proton permeability is given by Fick's law according to the following equation: P = J DeltaH, where P is the net proton permeability coefficient and DeltaH is the proton gradient between inside and outside of the vesicle (Rossignol et al., 1982).

Measurement of Delta with Oxonol VI

Oxonol VI fluorescence was measured at 614/646-nm excitation/emission wavelengths (Venema et al., 1993). Development of an inside positive membrane potential leads to uptake of the anionic dye into the intravesicular space and to enhanced partitioning into the lipid membrane, giving rise to an augmentation of fluorescence (Apell and Bersch, 1987). The reaction medium contained 20 mM Mes adjusted to pH 6.0 with KOH, 50 mM K(2)SO(4), 10% glycerol, and 50 nM oxonol VI. Reconstituted vesicles (30 µl; 15 µg of membrane protein) were added to the reaction cuvette. The H-ATPase reaction was initiated by the addition of 120 µl of 134 mM MgS0(4) and 124 mM ATP, pH 6.0, giving rise to a final concentration of 4 mM MgATP and 1 mM free Mg.

Protein Estimation

Protein concentration was determined by the method of Bradford(1976) with the Bio-Rad protein assay reagent and bovine serum albumin as the standard.

Gel Electrophoresis and Electrotransfer

Plasma membrane proteins were separated by SDS-polyacrylamide gel electrophoresis on 10% acrylamide using the system of Laemmli(1970). Western blotting with a polyclonal antibody against the C terminus of the yeast plasma membrane H-ATPase (Monk et al., 1991) and a second antibody conjugated to alkaline phosphatase (Promega) was as described previously (Blake et al., 1984).

Proton Extrusion Experiments

Cells were grown to stationary phase overnight in growth medium containing 1% yeast extract, 2% Peptone, and 2% glucose as carbon source. Cells were pelleted and resuspended at a concentration of 1 10^8 cells/ml in 20 mM KCl, and 20 µM antimycin when indicated. After incubation for 30 min, 2% glucose was added. Acidification of the external medium was monitored for 10 min with a pH electrode.

RESULTS

Effect of Glucose on HExtrusion in Vivo

Yeast cells suspended at high concentrations in unbuffered medium supplemented with 20 mM KCl did not extrude a significant number of H ions (approx0.01 pH unit/min at 10^8 cells/ml) (Fig. 1). The initial pH of the suspension of cells was close to 4. The addition of glucose (100 mM) to the yeast cells caused an extensive acidification of the external medium after a lag phase of <1 min (initial rate of 0.7 pH unit/min at 10^8 cells/ml) (Fig. 1), in accordance with Serrano(1983). The cells were able to reduce external pH to 3, at which point pH stabilized. The subsequent addition of glucose had no effect on pH. The glucose effect was not affected by the inclusion in the medium of antimycin (an inhibitor of electron transport) and therefore was not the result of endogenous respiration producing CO(2). Glucose-induced H secretion was the same whether a fresh overnight culture of cells had been starved for 1 h in H(2)O or grown in the presence of alternative carbon sources (e.g. glycerol). Therefore, the glucose effect in vivo cannot be explained in terms of glycolysis providing extra ATP to be utilized by the pump.

Figure 1: Effect of glucose on in vivo H extrusion from yeast cells. Yeast cells, grown in glucose medium to stationary phase, were washed in water and resuspended in a solution of 20 mM KCl to a final concentration of 10^8 cells/ml. H extrusion was measured by monitoring the pH of the medium with a pH electrode. At the arrow, 2% glucose was added. The initial rate of H extrusion was 0.01-0.02 pH unit/min before the addition of glucose and 0.5-0.7 pH unit/min after the addition of glucose.


Effect of Glucose on Plasma Membrane ATPase Activity

When yeast cells were incubated with glucose, a rapid activation of the plasma membrane H-ATPase measured in purified plasma membranes was observed (Fig. 2), in accordance with Serrano(1983). The activation was rapidly reversed after glucose removal, and it was therefore essential to homogenize the glucose-metabolizing cells without washing. The ATPase activity of the plasma membrane H-ATPase in isolated plasma membranes from yeast grown in glucose was 3-5 times higher (3 µmol/min/mg of protein at pH 6.0) than in plasma membranes isolated from yeast that had been deprived of glucose 10 min prior to homogenization (Fig. 2).

Figure 2: ATP hydrolytic activity as a function of pH of plasma membranes isolated from glucose-starved and glucose-activated cells. Plasma membranes were isolated from glucose-starved and glucose-activated cells as described under ``Materials and Methods.'' ATP hydrolytic activity was measured by measuring the release of inorganic phosphate as described under ``Materials and Methods'' with a MgATP concentration of 4 mM and 1 mM free Mg. The pH was adjusted with N-methyl-D-glucamine. Triangles, glucose-starved; circles, glucose-activated.


The glucose-activated ATPase had a pH optimum around 6, whereas the nonactivated enzyme had a pH optimum around 5.5 (Fig. 2). At all pH values studied, however, the increase in specific activity was never more than 8 times ( Fig. 2and Fig. 4-6) compared with the >50-fold increase in H efflux from whole cells after the addition of glucose in vivo.

Figure 4: Dependence of the rate of ATP hydrolysis on the concentration of MgATP of native and reconstituted plasma membrane ATPases from glucose-activated and glucose-starved yeast cells. Plasma membranes were isolated from glucose-starved and glucose-activated cells as described under ``Materials and Methods.'' ATP hydrolytic activity was measured by measuring the release of inorganic phosphate as described under ``Materials and Methods'' with MgATP concentrations ranging from 0.25 to 7.5 mM and 1 mM free Mg. The data represent the means of four independent repetitions with the same membrane preparation. Open symbols, native ATPase; closed symbols, reconstituted ATPase; triangles, glucose-starved ATPase; circles, glucose-activated ATPase. The data were fitted to the following equation: v/[E](0) = (a[S] + b[S]^2)/(1 + c[S] + d[S]^2) (Koland and Hammes, 1986) with the following values for the constants a, b, c, and d. Glucose-activated native membranes: a = 7.5 µmol/min/mg/mM, b = 2.1 µmol/min/mg/mM^2, c = 5.95/mM, and d = 0.47/mM^2 and r^2 = 0.998; glucose-activated reconstituted membranes: a = 6.5 µmol/min/mg/mM, b = 4.2 µmol/min/mg/mM^2, c = 7.6/mM, and d = 1.2/mM^2 and r^2 = 0.998; glucose-starved native membranes: a = 0.29 µmol/min/mg/mM, b = 0.22 µmol/min/mg/mM^2, c = 0.215/mM, and d = 0.188/mM^2 and r^2 = 0.997; glucose-starved reconstituted membranes: a = 0.043 µmol/min/mg/mM, b = 0.486 µmol/min/mg/mM^2, c = 0.35/mM, and d = 0.39/mM^2 and r^2 = 0.998. In the inset, a logarithmic transformation of the data shows the fit to the Hill equation. Values of 0.64 and 2.2 mM for H and K` were calculated from the slope of the line and the intercept with the yaxis, respectively, for the glucose-activated ATPase and values of 1.64 and 2.9 mM for H and K` for the glucose-starved ATPase.


The promoter of the yeast plasma membrane H-ATPase gene (PMA1) contains recognition sequences for a promoter-binding factor positively regulated by glucose (Capieaux et al., 1989). The glucose-mediated increase in H-ATPase activity, however, was rapid and completed within the 10 min of incubation. This relatively rapid activation suggests that de novo synthesis of H-ATPase does not contribute to the observed increase in activity. This was supported by the protein staining and immunostaining shown in Fig. 3(A-B). The plasma membranes contained a prominent band of M(r) approx105,000 corresponding to the H-ATPase. The intensity of this H-ATPase band was not increased in samples from glucose-activated cells when compared with controls.

Figure 3: SDS-polyacrylamide gel electrophoresis of native and reconstituted plasma membrane vesicles from glucose-activated (GA) and glucose-starved (GS) yeast cells. A, plasma membranes (PM; 12.5 µg of protein) were subjected to SDS-polyacrylamide gel electrophoresis as described under ``Materials and Methods.'' The resulting gels were stained with Coomassie Blue. B, Western blot analysis is shown of plasma membranes (2.5 µg of protein). A polyclonal antibody against the C terminus of the yeast plasma membrane H-ATPase was used. C, plasma membranes (200 µl; 100 µg of protein) in the presence of asolectin and detergent were subjected to centrifugation for 100,000 g in a Beckman Airfuge for 10 min. The pellet was resuspended in the same volume. Equal volumes (25 µl) of total membranes (PM + det), supernatant (sup), and resuspended pellet (pellet) were subjected to SDS-polyacrylamide gel electrophoresis. D, the supernatant from C was passed through a Sephadex G-50 column as described under ``Materials and Methods.'' An aliquot (25 µl) of the eluate (200 µl) was diluted 8-fold and subjected to centrifugation for 100,000 g in a Beckman Airfuge for 30 min (pellet). As a control, another aliquot was diluted 8-fold, after which octyl glucoside was added at the same concentration as during reconstitution to solubilize the vesicles before subjecting it to centrifugation for 100,000 g for 30 min (pellet). Undiluted eluate (eluate; 25 µl) and the resulting pellets (pellet and pellet; resuspended in 25 µl) were subjected to SDS-polyacrylamide gel electrophoresis. Molecular mass standards (in kilodaltons) are shown at the left .


Reconstitution of Plasma Membrane H-ATPase

Isolated yeast plasma membranes do not form vesicles that are sufficiently tight to allow measurements of H pumping. Our next goal was therefore to reconstitute the H-ATPase into liposomes so that its transport properties could be studied. The starting point for this work was the discovery by Perlin et al.(1984) that when isolated plasma membrane vesicles from Neurospora are solubilized with deoxycholate in the presence of asolectin, vesicles are re-formed when detergent is removed by column chromatography. In reconstituted vesicles produced this way, Neurospora H-ATPase constitutes 35% of the protein. In preliminary experiments, we observed that deoxycholate was not so effective for reconstitution of the yeast H-ATPase (data not shown). Octyl glucoside, on the contrary, was found to be superior to deoxycholate and was used in subsequent experiments.

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis profiles of plasma membranes and reconstituted vesicles are compared in Fig. 3. Densitometric scanning showed the 105-kDa H-ATPase band to account for approx20% of the total Coomassie Blue-staining material in the plasma membranes. Octyl glucoside solubilized 40-60% of the total protein and 80-90% of the H-ATPase (Fig. 3C). After treatment with octyl glucoside, the relative amount of the M(r) 105,000 H-ATPase band increased 2-fold to 40% of the solubilized Coomassie Blue-staining material. The SDS-polyacrylamide gel electrophoresis profile remained the same after detergent had been removed by passage of solubilized material through the gel filtration column. After spinning the reconstituted vesicles at 100,000 g in a Beckman Airfuge for 30 min, >50% of the protein was pelleted (Fig. 3D, pellet), suggesting that at least 50% of the H-ATPase was effectively reconstituted. Solubilized H-ATPase was not pelleted under these conditions (Fig. 3D, pellet). As judged from the intensity of the 105-kDa band, the same amount of H-ATPase was present in samples from glucose-activated and glucose-starved cells during all stages of the reconstitution procedure (Fig. 3).

The volume of vesicles was estimated from the fluorescence of trapped pyranine in reconstituted membrane vesicles (Table 1). When this value was compared with the molar amount of lipid present in the samples, it was possible to estimate the mean vesicle size (Table 1). Reconstituted asolectin vesicles had a mean radius of approx70 nm. The volumes of reconstituted plasma membrane vesicles were smaller (mean radius of approx40 nm) and were the same no matter whether the vesicles were derived from glucose-starved or glucose-activated cells.

ATP Hydrolysis by Native and Reconstituted H-ATPases

The dependence of ATP hydrolysis on MgATP concentration was not affected by the reconstitution procedure (Fig. 4). The ATP hydrolysis rate of H-ATPase isolated from glucose-starved cells showed a sigmoidal relationship on MgATP concentration (Fig. 4) that can be fitted by the following equation: v/[E](0) = (a[S] + b[S]^2)/(1 + c[S] + d[S]^2) as described before (Koland and Hammes, 1986). Two interpretations of these results are as follows. 1) The enzyme possesses multiple catalytic sites that interact in a positive cooperative way, and 2) the enzyme can exist in multiple conformational states that catalyze MgATP hydrolysis by parallel pathways (Koland and Hammes, 1986). After activation by glucose, we found that the plot was no longer sigmoidal, but not purely hyperbolic either. However, the data can still be fitted by the same equation as above (Fig. 4). The shape of the curve obtained for glucose-activated ATPase suggests negative cooperativity or the presence of a mixture of enzymes catalyzing the same reaction. It is thus theoretically possible that only part of the ATPase molecules are activated upon addition of glucose, giving rise to a mixture of glucose-activated and glucose-starved ATPases in this preparation. Our data do not permit us to distinguish between the various possibilities. Complex kinetics of the glucose-activated ATPase requiring an equation composed by the sum of two Michaelian terms to fit the experimental data has been described before (Berberián et al., 1993). The data can also be fitted adequately by the Hill equation: v = V(max) S^H/(K` + S^H) (see inset in Fig. 4), classically used to indicate cooperative effects and which can also fit data obeying the equation used by Koland and Hammes (Dixon and Webb, 1979). The difference in shape between the curves for glucose-starved and glucose-activated ATPases is described by changing the value of H from >1 (positive cooperativity) to <1 (negative cooperativity), while the value of K` remains approximately the same.

HPumping by Reconstituted H-ATPase

Reconstitution of the plasma membrane H-ATPase as outlined above allowed direct demonstration of H pumping activity. The MgATP-dependent intravesicular acidification of the plasma membrane asolectin vesicles was followed by the quenching of acridine orange absorbance (Palmgren, 1990). Upon addition of MgATP, a decay in absorbance was observed, which leveled off within 2 min (Fig. 5A). The quenching of acridine orange absorbance was stimulated severalfold by the K ionophore valinomycin (0.5 µM) (Fig. 5A), which dissipates the membrane potential, and was abolished by nigericin (0.1 µg/ml) (Fig. 5A), which catalyzes the electroneutral exchange of H for potassium (Pressman, 1976).

Figure 5: Effect of glucose on H transport and ATP hydrolysis by plasma membrane H-ATPase in reconstituted vesicles. H transport activity and ATPase activity of plasma membrane H-ATPase in reconstituted vesicles derived from glucose-starved and glucose-activated cells were measured simultaneously in the same cuvette as indicated under ``Materials and Methods'' at pH 6.5. Note that only half the amount of protein was used in those assays employing vesicles derived from glucose-activated cells as compared with glucose-starved cells. A, H transport activity expressed as the initial rate of acridine orange absorbance quenching at 495 nm; B, ATP hydrolytic activity as calculated from the coupled NADH oxidation measured at 340 nm. Triangles, membranes from glucose-starved cells (100 µg of membrane protein/ml); circles, membranes from glucose-activated cells (50 µg of membrane protein/ml); open symbols, activity in the absence of valinomycin; closed symbols, activity in the presence of 0.5 µM valinomycin. At the closed arrow, the reaction was started by the addition of MgATP (4 mM MgATP + 1 mM Mg, final pH 6.5). At the open arrow, the pH gradient was collapsed by the addition of 0.5 µM nigericin. Numbers indicate activities in terms of DeltaA/min/ml (H pumping) and nmol of ADP/min/ml (ATPase activity).


ATP hydrolysis by the reconstituted plasma membrane H-ATPase was measured in an assay in which ADP release was coupled to oxidation of NADH. Since the absorbance spectra of acridine orange and NADH are not overlapping, it was possible to measure H pumping (Fig. 6, upperpanel) and ATP hydrolysis (lowerpanel) simultaneously. Neither valinomycin (Fig. 6, lower panel), which facilitates the formation of a pH gradient, nor nigericin (data not shown), which dissipates the pH gradient, affected ATPase activity significantly. The apparent insensitivity of ATP hydrolysis to valinomycin observed here and by others (Dufour et al., 1982; Serrano, 1984) suggests that the reversal potential for ATP hydrolysis has not been reached in these systems. Thus, the maximal rates of H pumping in the system are determined by the passive H permeability of the vesicles, and the quenching of acridine orange absorbance levels off when H influx matches H efflux.

Figure 6: ATP dependence of H transport activity and ATP hydrolytic activity of reconstituted H-ATPase derived from glucose-starved (GS) and glucose-activated (GA) cells. H transport and ATP hydrolysis were measured simultaneously as described under ``Materials and Methods'' with 50 µg of membrane protein/assay, MgATP concentrations ranging from 0 to 6 mM, and 1 mM free Mg, pH 6.0, and in the presence of 0.5 µM valinomycin. Note that H pumping by the glucose-activated ATPase was increased significantly more than ATP hydrolysis. Closed symbols, proton pumping; open symbols, ATPase activity. The experimental data were fitted to the following equation: v/[E](0) = (a[S] + b[S]^2)/(1 + c[S] + d[S]^2) (Koland and Hammes, 1986). For the glucose-activated ATPase, the values were as follows: a = 15 DeltaA/min/mg/mM and b = 5.6 DeltaA/min/mg/mM^2 (H pumping) or a = 5.3 µmol/min/mg/mM and b = 2.0 µmol/min/mg/mM^2 (ATP hydrolysis) and for both H pumping and ATP hydrolysis, c = 2.7/mM and d = 0.44/mM^2. For the glucose-starved ATPase, the values were as follows: a = 0.12 DeltaA/min/mg/mM and b = 0.40 DeltaA/min/mg/mM^2 (H pumping) or a = 0.39 µmol/min/mg/mM and b = 1.31 µmol/min/mg/mM^2 (ATP hydrolysis) and for both H pumping and ATP hydrolysis, c = 0.79/mM and d = 1.12/mM^2.


H pumping and the hydrolysis of ATP were catalyzed by the same enzyme as based on the following observations. (a) H pumping and ATPase activity exhibited similar dependence on MgATP concentration (Fig. 6), and (b) comparable pH-activity profiles were observed for both H pump activity and ATPase activity in the activated as well as the nonactivated state (data not shown).

Effect of Glucose on HPumping by ATPase

The glucose-activated ATPase had an increased potential for H pumping that exceeded the increase in specific ATPase activity by about an order of magnitude ( Fig. 5and 6). Calculated on an equal protein basis, the rate of H transport exhibited by the glucose-activated ATPase was 20-50 times higher than that of the nonactivated ATPase. Thus, the relative change in H transport observed between glucose-activated and nonactivated ATPases matches the relative changes in H efflux in vivo when yeast cells are challenged with glucose.

Internal pH in liposomes containing H-ATPase from glucose-starved and glucose-activated cells was estimated using the fluorescent pH probe pyranine (Fig. 7). Glucose-activated H-ATPase was able to acidify the interior of the vesicles from pH 7.0 to 6.9, whereas glucose-starved H-ATPase could not produce any detectable acidification of the intravesicular volume.

Figure 7: Internal pH in liposomes containing reconstituted plasma membrane H-ATPase from glucose-activated (GA) and glucose-starved (GS) cells. Internal pH was calculated from the fluorescence of the pH probe pyranine trapped inside reconstituted vesicles as described under ``Materials and Methods.'' The reaction was started by the addition of 4 mM MgATP and 1 mM free Mg, pH 7.0. At the end of the experiment, the pH gradient was abolished by the addition of 80 nM nigericin.


We next employed the Delta probe oxonol VI to study the electrogenic properties of the two regulatory states of the H-ATPase. It appeared that the glucose-activated enzyme readily established a membrane potential both in the absence and presence of K (Fig. 8). In marked contrast, the glucose-starved enzyme did not produce a clear fluorescence signal (Fig. 8). As expected, the stability of the membrane potential was influenced by the presence of K in the intravesicular medium. However, K did not alter the maximal amplitude of the signal produced by glucose-starved and glucose-activated H-ATPases.

Figure 8: Changes in Delta in liposomes containing plasma membrane H-ATPase from glucose-activated (GA) and glucose-starved (GS) cells. Delta was measured by oxonol VI fluorescence quenching as described under ``Materials and Methods.'' Upon development of an inside positive membrane potential by the H-ATPase, the probe will fix to the internal leaflet of the membrane, giving rise to an augmentation of fluorescence. A, membranes were reconstituted as described under ``Materials and Methods'' with 50 mM K(2)SO(4). ox, oxonol. B, membranes were reconstituted in the absence of K. MgSO(4) (1 mM) was included instead of K(2)SO(4) to screen the negative charges of the phospholipids. The membrane potential was abolished by the addition of valinomycin (50 nM; A) or gramicidin (200 nM; B).


Passive Ion Permeability of Reconstituted Vesicles

In principle, H accumulation in membrane vesicles could be stimulated (a) indirectly by preventing the formation of a membrane potential that could otherwise inhibit H influx (e.g. by affecting the permeability of a secondary system), (b) by reducing the passive efflux of H from the vesicles, or (c) by direct stimulation of H transport systems.

Electrical balance between the exterior and interior of the membrane vesicles was obtained during the assay since the potassium ionophore valinomycin was present. In the absence of valinomycin, the passive permeability of the membrane to K was determined according to Venema et al.(1993). Glucose treatment did not change the estimated K permeability (data not shown).

The permeability coefficient for H was determined by analyzing the dissipation of an imposed pH gradient (Fig. 9). Intravesicular pH was determined by the pH probe pyranine. After imposing a pH gradient of 1.1 pH units (pH outside = 7.6; inside = 6.5), intravesicular pH was monitored as a function of time (Fig. 9, left panels). The kinetics of H fluxes could be fitted by a single exponential function (Fig. 9, right panels). First-order kinetics is indicative of emptying a single compartment through a homogeneous barrier. If multilamellar structures had been present, more complex kinetics would have been expected. The permeability coefficients for proteoliposomes derived from glucose-activated cells, glucose-starved cells, and liposomes derived from asolectin were 1.74 ± 0.44 10, 2.20 ± 0.35 10, and 1.20 ± 0.27 10 m s, respectively, confirming that glucose treatment did not alter the passive permeability of the proteoliposomes to H. Taken together, these results suggest that glucose stimulates the active H influx.

Figure 9: Passive H permeabilities of reconstituted plasma membrane vesicles prepared from glucose-starved (GS) and glucose-activated (GA) cells. Plasma membrane vesicles were reconstituted as described under ``Materials and Methods'' in the presence of 2 mM pyranine. Reconstituted vesicles (10 µl; 5 µg of protein) were added to 3 ml of 10 mM Mes adjusted to pH 6.5 with KOH, 50 mM K(2)SO(4), 50 nM valinomycin, and 20% glycerol. The pH of the medium was next raised to pH 7.6 by the addition of 30 µl of 1 MN-methyl-D-glucamine, and the time course of augmentation of pyranine fluorescence at a 460-nm excitation wavelength was followed. The fluorescence signal was calibrated with pH as described under ``Materials and Methods'' (left panels). The proton flux (right panels) and permeability coefficients were calculated as described under ``Material and Methods.'' The kinetics of the proton fluxes were fitted by a logarithmic function (right panels, smooth lines). The estimated permeability coefficients were 1.74 ± 0.44 10 m s for reconstituted glucose-activated membranes, 2.20 ± 0.35 10 m s for reconstituted glucose-starved membranes, and 1.20 ± 0.27 10 m s for reconstituted liposomes.


DISCUSSION

It was shown by Serrano(1983) that ATP hydrolytic activity of the plasma membrane H-ATPase is positively regulated by glucose. We have found that glucose-activated yeast plasma membrane H-ATPase has an increased potential for H pumping that is about an order of magnitude higher than the increase in specific ATPase activity ( Fig. 5and Fig. 6). The glucose-starved H-ATPase was hardly able to establish a membrane potential across the vesicle membrane (Fig. 8), suggesting that in this regulatory state, the H-ATPase is not functioning as an electrogenic pump. The fact that H accumulation is stimulated to a higher degree by glucose than is ATP hydrolysis suggests that H pumping can be regulated independently of ATP hydrolysis. Glucose may alter the H/ATP stoichiometry of the plasma membrane H-ATPase or promote coupling of ATP hydrolysis to H translocation.

It is possible that the absence of a functioning state of the H-ATPase could be due to its relative sensitivity to denaturation by detergent, in the regulatory state induced by glucose starvation. This, however, seems unlikely since the enzyme under the experimental conditions readily hydrolyzes ATP, and the dependence of ATP hydrolysis on MgATP concentration was not altered by the reconstitution procedure (Fig. 3). Another possibility is that the successful reconstitution of enzyme units into sealed vesicles is affected by the same structural change, e.g. regulatory phosphorylation, which could affect enzyme lability or structure. However, we have demonstrated that a large fraction of the detergent-solubilized protein is incorporated into structures that sediment, and we have shown that there is no visible difference in the amount of protein incorporated when starved and glucose-activated membranes are the source or in the amount of ATPase activity recovered. In addition, the volume of vesicles harboring glucose-starved and glucose-activated H-ATPases was the same, indicating that the same amount of sealed vesicle structures is present in both preparations. Therefore, although it remains formally possible that stability of function to the reconstitution procedure may be influenced by the glucose-induced modification, the evidence presented strongly supports a model in which glucose activation modifies the coupling efficiency of the H-ATPase.

An alternative artifact is that the acridine orange signal is not linearly related to changes in the rate of H pumping. This is less likely since acridine orange absorbance changes were closely related to changes in ATPase activity when ATP concentration (Fig. 6) and pH (data not shown) were altered. In addition, by employing two fluorescent probes (pyranine and oxonol VI) that report intravesicular pH (Fig. 7) and Delta (Fig. 8), respectively, the discrepancy between activation of H pumping and ATP hydrolysis was confirmed.

The maximal gradient in our system is probably determined by the high H leakiness of the liposomes both in the presence and absence of protein (Fig. 9). The maximal pH gradient produced by the glucose-activated ATPase amounted to 0.1 pH unit (measured at pH 7.0 in the extravesicular medium) when using pyranine as a probe to report intravesicular pH (Fig. 7), and 1 pH unit (pH 6.0 in the extravesicular medium; pH 5.0 inside the vesicles) (data not shown) when measured by employing acridine orange and using the pH jump method introduced by Dufour et al.(1982). This discrepancy is probably caused by the different mechanism by which these probes report pH gradients. Pyranine, trapped inside the lumen of vesicles, reports the mean internal pH of all vesicles. Acridine orange, on the contrary, only accumulates inside vesicles that harbor functional H-ATPase. Thus, it seems likely that a population of vesicles does not contain any H-ATPase at all. Using pure Neurospora H-ATPase protein, a different reconstitution procedure, and 100 times more asolectin relative to protein than in the present study, Goormaghtigh et al. (1986) found that <0.5% of liposomes contained H-ATPase.

Most authors have suggested a coupling ratio of 1 H transported per ATP hydrolyzed for plasma membrane H-ATPase from yeast (Serrano, 1984), Neurospora (Warncke and Slayman, 1980; Perlin et al., 1986), algae (Blatt et al., 1990), and higher plants (Brauer et al., 1989; Briskin and ReynoldsNiesman, 1991). The limits for the H/ATP stoichiometry of the pump are set by the free energy supplied by the chemical reaction per turnover (Läuger, 1991). If the pump is tightly coupled and if leakage pathways are negligible, the system reaches equilibrium when the electrochemical gradient counterbalances the chemical driving force (DeltaG). If the pump translocates n ions/cycle, this equilibrium condition is given by the following equation: DeltaG = n(RT 2.3 DeltapH + zFV) (where z is the valency of the ion, F is the Faraday constant, and V is the membrane potential). Assuming that the free energy for ATP hydrolysis under physiological conditions is 40 kJ/mol, the maximal pH gradients that can be created (for V = 0) would be 6.8, 3.4, and 0.68 for n = 1, 2, and 10, respectively. In vivo, glucose-metabolizing cells can sustain pH gradients of at least 4 pH units (Serrano, 1984). The size of the electrical gradient produced by the plasma membrane H-ATPase (membrane potentials of up to 300 mV are generated by the Neurospora H-ATPase (Gradmann et al., 1978)) makes it a potent electrogenic transport protein. With a ratio of 5-10 H pumped per ATP consumed, the maximal capacity for formation of electrochemical gradients would be far below these values. It is therefore reasonable to suggest a 1 H/ATP stoichiometry for the ATPase under conditions where it generates maximal pH gradients and membrane potentials. The initial rates of H translocation observed by us suggest that the glucose-activated H-ATPase translocates more H per ATP consumed than the enzyme isolated from glucose-deprived cells (Fig. 5). Assuming that the activated H-ATPase operates with a stoichiometry of 1 H/ATP, our results immediately suggest that the stoichiometry of the nonactivated yeast ATPase is <1 H/ATP (e.g. 0.1), i.e. net transport of H is essentially uncoupled from the splitting of ATP. In future studies, the actual H/ATP stoichiometry of the purified yeast plasma membrane H-ATPase before and after glucose activation needs to be determined, e.g. by optimizing the reconstitution procedure, by a thermodynamic approach using the patch-clamp method (Davies et al., 1994), or after reconstitution of the ATPase into planar lipid bilayer membranes (Ziegler et al., 1993).

Intrinsic uncoupling (defined here as ATP hydrolysis without net translocation of the full potential complement of H) has previously been suggested to play a role in the regulation of a variety of ion pumps such as bacteriorhodopsin (Westerhoff and Dancsházy, 1984; Caplan, 1988), vacuolar H-ATPase (Davies et al., 1994; Kibak et al., 1993; Tu et al., 1987; Yoshinori and Nelson, 1988), cytochrome oxidase (Blair et al., 1986), F(0)F(1)-ATPase (Krenn et al., 1993; Muller, 1993; Pietrobon et al., 1986; van Walraven et al., 1990), and sarcoplasmic reticulum Ca-ATPase (Caplan, 1988; Inesi and de Meis, 1989; Meltzer and Berman, 1984; Navarro and Essig, 1984; Soler et al., 1990). Intrinsic uncoupling has been suggested to play a role in providing a ``safety valve'' for the formation of gradients (Caplan, 1988) or in matching the pump to the load for optimization purposes (Stucki, 1980).

The mechanistic implications of H-ATPase uncoupling remain to be explored. Intrinsic uncoupling of H-ATPase may arise in at least two ways (Läuger, 1991). First, kinetic studies on members of the P class of ATPases suggest that hydrolysis of the aspartylphosphoryl (E-P) intermediate is closely associated with the simultaneous translocation of the transported ion(s). The phosphorylated state, however, may spontaneously dephosphorylate without ion translocation (slippage). However, since the nonactivated ATPase is not deficient in H transport and is able to build up a H gradient (Fig. 5A), slippage would have to be partial. Second, glucose activation may result in a decreased H permeability intrinsic to the yeast plasma membrane H-ATPase. Intrinsic H transport in the reverse direction by a process that is not linked to ATP synthesis may take place without conformational change of the protein (tunneling) or may occur by a carrier-like operation mode of the pump involving conformational changes. At least tunneling may be specific for active ATPases (Fröhlich, 1988), which could explain why the apparent passive permeability of vesicles, measured in absence of ATP, is not affected by glucose regulation of the H-ATPase (Fig. 9). It is thus possible that the intrinsic pathway is rendered more permeable under conditions of pump turnover, allowing a higher leakage of H.

Removal of the last 11 amino acids from the yeast H-ATPase (Glu stop) produces an enzyme in glucose-starved cells with kinetic parameters similar to those of the glucose-activated wild-type H-ATPase (Portillo et al., 1989). The truncated H-ATPase is not activated further in glucose-metabolizing cells. The same phenotype is exhibited by a mutation (Ala Val) affecting a residue in the nucleotide-binding site that is located in the large central cytoplasmic domain (Cid and Serrano, 1988). Therefore, the C terminus seems to interact with this site. Glucose-activated H-ATPase is phosphorylated at a residue not phosphorylated in glucose-starved cells (Chang and Slayman, 1991). A double mutation at the C terminus destroying putative phosphorylation sites (Ser Ala,Thr Ala) locks the enzyme in the inhibited state. This double mutation results in almost no activation of the H-ATPase by glucose and no growth of yeast in glucose medium (Portillo et al., 1991), suggesting that kinase-mediated phosphorylation of amino acids at the C terminus is part of the glucose response. A Tyr Gly mutant at the top of transmembrane segment M5 of the Ca-ATPase of sarcoplasmic reticulum is uncoupled in the sense that it catalyzes a high rate of Ca-activated ATP hydrolysis without net accumulation of Ca in membrane vesicles (Andersen, 1995). It has been suggested that the side chain of Tyr might play a critical role in the gating mechanism normally preventing the occluded calcium ions from dissociating to the cytoplasmic site upon dephosphorylation (Andersen, 1995). In analogy, one could speculate that the C terminus of the yeast H-ATPase stabilizes a conformation of the enzyme that is unable to effectively occlude H.

Jencks(1980) has defined certain rules for the reaction cycle that need to be obeyed for coupling in ion pumps. The main concept that emerges is that ATP hydrolysis does not occur without ion transport, and no reverse flux of ions occurs without ATP synthesis. Thus, existing models for ion pumps, which are generally based on mechanisms having integral stoichiometry, cannot account for our experimental findings of variable coupling. Since uncoupling would theoretically result in futile cycling of ATP and is typically induced only under in vitro conditions, variable stoichiometry has remained a controversial concept. In this paper, we have demonstrated a change in coupling ratio of an ion pump induced by a metabolite under in vivo conditions. This points to a physiological role for uncoupling as a mechanism for regulation of pump activity. The tightly coupled high activity state induced by glucose may be essential for the formation of the very steep H gradients required for efficient solute uptake. Regulated uncoupling may be advantageous when taking into consideration that, with a stoichiometry of 1 H/ATP, yeast H-ATPase is physiologically irreversible (Serrano, 1984). Uncoupling intrinsic to the pump would allow for regulation of the magnitude of the steady-state electrochemical gradient. Still, the partially uncoupled low activity state of the ATPase may be sufficient to maintain H gradients required for normal growth. It seems clear, however, that extensive kinetic controls must operate to avoid undesired H leakage or futile consumption of ATP.

FOOTNOTES

*
This work was supported by the Danish Natural Science Research Council, the NOVO Nordisk Fonden, and the European Communities' BIOTECH Programme as part of the Project of Technological Priority 1993-1996. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
A European Molecular Biology Organization short term fellow.

To whom correspondence should be addressed. Tel.: 45-3528-3338; Fax: 45-3528-3333.

(^1)
The abbreviations used are: Mes, 4-morpholineethanesulfonic acid; Mops, 3-(N-morpholino)propanesulfonic acid.

ACKNOWLEDGEMENTS

We are grateful to Frank C. Lanfermeijer and Morten Kielland-Brandt for valuable discussions and critical reading of the manuscript.

REFERENCES

  1. Andersen, J. P. (1995) J. Biol. Chem. 270,908-914 [Abstract/Free Full Text]
  2. Apell, H. J., and Bersch, B. (1987) Biochim. Biophys. Acta 903,480-494 [Medline] [Order article via Infotrieve]
  3. Baginsky, E. S., Foa, P. P., and Zak, B. (1967) Clin. Chem. 13,326-332 [Abstract]
  4. Berberián, G., Helguera, G., and Beaugé, L. (1993) Biochim. Biophys. Acta 1153,283-288 [Medline] [Order article via Infotrieve]
  5. Blair, D. F., Gelles, J., and Chan, S. I. (1986) Biophys. J. 50,713-733 [Medline] [Order article via Infotrieve]
  6. Blake, M. S., Johnston, K. H., Russell-Jones, G. J., and Gotschlich, E. C. (1984) Anal. Biochem. 128,302-311
  7. Blatt, M. R., Beilby, M. J., and Tester, M. (1990) J. Membr. Biol. 114,205-223 [CrossRef][Medline] [Order article via Infotrieve]
  8. Bradford, M. M. (1976) Anal. Biochem. 72,248-254 [CrossRef][Medline] [Order article via Infotrieve]
  9. Brauer, D., Hsu, A.-F., and Thomas, C. E. (1989) Plant Physiol. 89,464-471 [Abstract/Free Full Text]
  10. Briskin, D. P., and Hanson, J. B. (1992) J. Exp. Bot. 43,269-289 [Abstract/Free Full Text]
  11. Briskin, D. P., and Reynolds-Niesman, I. (1991) Plant Physiol. 95,242-250 [Abstract/Free Full Text]
  12. Cantley, L. C., Jr., Cantley, L. G., and Josephson, L. (1978) J. Biol. Chem. 253,7361-7368 [Free Full Text]
  13. Capieaux, E., Vignais, M.-L., Sentenac, A., and Goffeau, A. (1989) J. Biol. Chem. 264,7437-7446 [Abstract/Free Full Text]
  14. Caplan, S. R. (1988) in The Ion Pumps: Structure, Function, and Regulation (Stein, W., ed) pp. 377-386, Alan R. Liss, Inc., New York
  15. Chang, A., and Slayman, C. W. (1991) J. Cell Biol. 115,289-295 [Abstract/Free Full Text]
  16. Cid, A., and Serrano, R. (1988) J. Biol. Chem. 263,14134-14139 [Abstract/Free Full Text]
  17. Davies, J. M., Hunt, I., and Sanders, D. (1994) Proc. Natl. Acad. Sci. U. S. A. 91,8547-8551 [Abstract/Free Full Text]
  18. Dixon, M., and Webb, E. C. (1979) Enzymes , Longman Group UK Ltd., London
  19. Dufour, J.-P., Goffeau, A., and Tsong, T. Y. (1982) J. Biol. Chem. 257,9365-9371 [Free Full Text]
  20. Fröhlich, O. (1988) J. Membr. Biol. 101,189-198 [CrossRef][Medline] [Order article via Infotrieve]
  21. Goormaghtigh, E., Chadwick, C., and Scarborough, G. A. (1986) J. Biol. Chem. 261,7466-7471 [Abstract/Free Full Text]
  22. Gradmann, D., Hansen, U.-P., Long, W. S., Slayman, C. L., and Warncke, J. (1978) J. Membr. Biol. 39,333-367 [CrossRef][Medline] [Order article via Infotrieve]
  23. Guarante, L., Yocum, R. R., and Gifford, P. (1982) Proc. Natl. Acad. Sci. U. S. A. 79,7410-7414 [Abstract/Free Full Text]
  24. Inesi, G., and de Meis, L. (1989) J. Biol. Chem. 264,5929-5936 [Abstract/Free Full Text]
  25. Jencks, W. P. (1980) Adv. Enzymol. Relat. Areas Mol. Biol. 51,75-106 [Medline] [Order article via Infotrieve]
  26. Kibak, H., Van Eeckhout, D., Cutler, T., Taiz, S. L., and Taiz, L. (1993) J. Biol. Chem. 268,23325-23333 [Abstract/Free Full Text]
  27. Koland, J. G., and Hammes, G. G. (1986) J. Biol. Chem. 261,5936-5942 [Abstract/Free Full Text]
  28. Krenn, B. A., van Walraven, H. S., Scholts, M. J. C., and Kraayenhof, R. (1993) Biochem. J. 294,705-709
  29. Laemmli, U. K. (1970) Nature 227,680-685 [CrossRef][Medline] [Order article via Infotrieve]
  30. L ä uger, P. (1991) Electrogenic Ion Pumps , Sinauer Associates, Inc., Sunderland, MA
  31. Meltzer, S., and Berman, M. C. (1984) J. Biol. Chem. 259,4244-4253 [Abstract/Free Full Text]
  32. Monk, B. C., Montesinos, C., Ferguson, C., Leonard, K., and Serrano, R. (1991) J. Biol. Chem. 266,18097-18103 [Abstract/Free Full Text]
  33. Morrison, J. F. (1979) Methods Enzymol. 63,257-294 [Medline] [Order article via Infotrieve]
  34. Muller, A. W. J. (1993) Physiol. Chem. Phys. Med. NMR 25,95-111
  35. Navarro, J., and Essig, A. (1984) Biophys. J. 46,709-717 [Medline] [Order article via Infotrieve]
  36. Palmgren, M. G. (1990) Plant Physiol. (Bethesda) 94,882-886 [Abstract/Free Full Text]
  37. Palmgren, M. G. (1991) Anal. Biochem. 192,316-321 [CrossRef][Medline] [Order article via Infotrieve]
  38. Pedersen, P. L., and Carafoli, E. (1987) Trends Biochem. Sci. 12,146-150 [CrossRef]
  39. Perlin, D. S., Kasamo, K., Brooker, R. J., and Slayman, C. W. (1984) J. Biol. Chem. 259,7884-7892 [Abstract/Free Full Text]
  40. Perlin, D. S., San Fransisco, M. J. D., Slayman, C. W., and Rosen, B. P. (1986) Arch. Biochem. Biophys. 248,53-61 [CrossRef][Medline] [Order article via Infotrieve]
  41. Pietrobon, D., Zoratti, M., Azzone, G. F., and Caplan, S. R. (1986) Biochemistry 25,767-775 [CrossRef][Medline] [Order article via Infotrieve]
  42. Portillo, F., Larrinoa, I. F., and Serrano, R. (1989) FEBS Lett. 247,381-385 [CrossRef][Medline] [Order article via Infotrieve]
  43. Portillo, F., Eraso, P., and Serrano, R. (1991) FEBS Lett. 287,71-74 [CrossRef][Medline] [Order article via Infotrieve]
  44. Pressman, B. C. (1976) Annu. Rev. Biochem. 45,501-530 [CrossRef][Medline] [Order article via Infotrieve]
  45. Rossignol, M., Thomas, P., and Grignon, C. (1982) Biochim. Biophys. Acta 684,195-199 [Medline] [Order article via Infotrieve]
  46. Sanders, D. (1990) Annu. Rev. Plant Physiol. Plant Mol. Biol. 41, 77-107, and references therein [CrossRef]
  47. Serrano, R. (1983) FEBS Lett. 156,11-14 [CrossRef][Medline] [Order article via Infotrieve]
  48. Serrano, R. (1984) Curr. Top. Cell. Regul. 23,87-126 [Medline] [Order article via Infotrieve]
  49. Serrano, R. (1988) Biochim. Biophys. Acta 947,1-28 [Medline] [Order article via Infotrieve]
  50. Soler, F., Teruel, J. A., Fernández-Belda, F., and Gómez-Fernández, J. C. (1990) Eur. J. Biochem. 192,347-354 [Medline] [Order article via Infotrieve]
  51. Stucki, J. W. (1980) Eur. J. Biochem. 109,269-283 [Medline] [Order article via Infotrieve]
  52. Tu, S.-I., Nagahashi, G., and Brouillette, J. N. (1987) Arch. Biochem. Biophys. 256,625-637 [CrossRef][Medline] [Order article via Infotrieve]
  53. van Walraven, H. S., Scholts, M. J. C., Koppenaal, F., Bakels, R. H. A., and Krab, K. (1990) Biochim. Biophys. Acta 1015,425-434 [CrossRef]
  54. Venema, K., Gibrat, R., Grouzis, J.-P., and Grignon, C. (1993) Biochim. Biophys. Acta 1146,87-96 [Medline] [Order article via Infotrieve]
  55. Villalba, J. M., Palmgren, M. G., Berberián, G. E., Ferguson, C., and Serrano, R. (1992) J. Biol. Chem. 267,12341-12349 [Abstract/Free Full Text]
  56. Wach, A., Ahlers, J., and Gräber, P. (1990) Eur. J. Biochem. 189,675-682 [Medline] [Order article via Infotrieve]
  57. Warncke, J., and Slayman, C. L. (1980) Biochim. Biophys. Acta 591,224-233 [Medline] [Order article via Infotrieve]
  58. Westerhoff, H. V., and Dancsházy, Z. (1984) Trends Biochem. Sci. 7,112-117
  59. Yoshinori, M., and Nelson, N. (1988) in The Ion Pumps: Structure, Function, and Regulation (Stein, W., ed) pp. 387-394, Alan R. Liss, Inc., New York
  60. Ziegler, W., Slayman, C. L., and Cartwright, C. P. (1993) Gen. Physiol. Biophys. 12,429-443 [Medline] [Order article via Infotrieve]
©1995 by The American Society for Biochemistry and Molecular Biology, Inc.


Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Biol. Chem.Home page
M. J. Buch-Pedersen, E. L. Rudashevskaya, T. S. Berner, K. Venema, and M. G. Palmgren
Potassium as an Intrinsic Uncoupler of the Plasma Membrane H+-ATPase
J. Biol. Chem., December 15, 2006; 281(50): 38285 - 38292.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
A. S. Fraysse, A. L. B. Moller, L. R. Poulsen, B. Wollenweber, M. J. Buch-Pedersen, and M. G. Palmgren
A Systematic Mutagenesis Study of Ile-282 in Transmembrane Segment M4 of the Plasma Membrane H+-ATPase
J. Biol. Chem., June 10, 2005; 280(23): 21785 - 21790.
[Abstract] [Full Text] [PDF]

Home page
 Physiol. Rev.Home page
C. A. Wagner, K. E. Finberg, S. Breton, V. Marshansky, D. Brown, and J. P. Geibel
Renal Vacuolar H+-ATPase
Physiol Rev, October 1, 2004; 84(4): 1263 - 1314.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
M. J. Buch-Pedersen and M. G. Palmgren
Conserved Asp684 in Transmembrane Segment M6 of the Plant Plasma Membrane P-type Proton Pump AHA2 Is a Molecular Determinant of Proton Translocation
J. Biol. Chem., May 9, 2003; 278(20): 17845 - 17851.
[Abstract] [Full Text] [PDF]

Home page
 Mol. Cell. Biol.Home page
A. Goossens, N. de la Fuente, J. Forment, R. Serrano, and F. Portillo
Regulation of Yeast H+-ATPase by Protein Kinases Belonging to a Family Dedicated to Activation of Plasma Membrane Transporters
Mol. Cell. Biol., October 15, 2000; 20(20): 7654 - 7661.
[Abstract] [Full Text]

Home page
 Mol. Cell. Biol.Home page
J. M. Mulet, M. P. Leube, S. J. Kron, G. Rios, G. R. Fink, and R. Serrano
A Novel Mechanism of Ion Homeostasis and Salt Tolerance in Yeast: the Hal4 and Hal5 Protein Kinases Modulate the Trk1-Trk2 Potassium Transporter
Mol. Cell. Biol., May 1, 1999; 19(5): 3328 - 3337.
[Abstract] [Full Text] [PDF]

Home page
 J. Bacteriol.Home page
A. Hernández, D. T. Cooke, and D. T. Clarkson
Effects of Abnormal-Sterol Accumulation on Ustilago maydis Plasma Membrane H+-ATPase Stoichiometry and Polypeptide Pattern
J. Bacteriol., January 15, 1998; 180(2): 412 - 415.
[Abstract] [Full Text]

Home page
 J. Biol. Chem.Home page
M. J. Buch-Pedersen, K. Venema, R. Serrano, and M. G. Palmgren
Abolishment of Proton Pumping and Accumulation in the E1P Conformational State of a Plant Plasma Membrane H+-ATPase by Substitution of a Conserved Aspartyl Residue in Transmembrane Segment 6
J. Biol. Chem., December 8, 2000; 275(50): 39167 - 39173.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Venema, K.
Right arrow Articles by Palmgren, M. G.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Venema, K.
Right arrow Articles by Palmgren, M. G.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 1995 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement