(Received for publication, March 29, 1995; and in revised form, June 16, 1995)
From the
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.
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 Ca
ATPases. Like these enzymes, it contains a
major M
100,000 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
pH 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.
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 (Rossignol et al., 1982) and assuming pure
phospholipid vesicles, the mean vesicle radius was estimated.
Concentrations of free Mg and MgATP
were calculated using and (Morrison,
1979; Wach et al.,
1990):
where K = 14.3 µM (dissociation constant, MgATP
), K
= 1.44 mM (dissociation
constant, MgHATP
), and K
= 0.107 µM (dissociation constant,
HATP
).
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.
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
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.
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]
= (a[S] + b[S]
)/(1
+ c[S] + d[S]
) (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
, c =
5.95/mM, and d = 0.47/mM
and r
= 0.998; glucose-activated
reconstituted membranes: a = 6.5
µmol/min/mg/mM, b = 4.2
µmol/min/mg/mM
, c =
7.6/mM, and d = 1.2/mM
and r
= 0.998; glucose-starved native
membranes: a = 0.29 µmol/min/mg/mM, b = 0.22 µmol/min/mg/mM
, c = 0.215/mM, and d =
0.188/mM
and r
=
0.997; glucose-starved reconstituted membranes: a =
0.043 µmol/min/mg/mM, b = 0.486
µmol/min/mg/mM
, c =
0.35/mM, and d = 0.39/mM
and r
= 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
105,000 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
.
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
20% 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
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
70 nm. The volumes of reconstituted plasma membrane vesicles were
smaller (mean radius of
40 nm) and were the same no matter whether
the vesicles were derived from glucose-starved or glucose-activated
cells.
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
A
/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]
= (a[S] + b[S]
)/(1
+ c[S] + d[S]
) (Koland and Hammes, 1986). For the
glucose-activated ATPase, the values were as follows: a = 15
A
/min/mg/mM and b = 5.6
A
/min/mg/mM
(H
pumping) or a = 5.3
µmol/min/mg/mM and b = 2.0
µmol/min/mg/mM
(ATP hydrolysis) and for both
H
pumping and ATP hydrolysis, c =
2.7/mM and d = 0.44/mM
.
For the glucose-starved ATPase, the values were as follows: a = 0.12
A
/min/mg/mM and b = 0.40
A
/min/mg/mM
(H
pumping) or a = 0.39
µmol/min/mg/mM and b = 1.31
µmol/min/mg/mM
(ATP hydrolysis) and for both
H
pumping and ATP hydrolysis, c =
0.79/mM and d =
1.12/mM
.
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).
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 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 in liposomes
containing plasma membrane H
-ATPase from
glucose-activated (GA) and glucose-starved (GS)
cells.
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
SO
. ox, oxonol. B,
membranes were reconstituted in the absence of K
.
MgSO
(1 mM) was included instead of
K
SO
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).
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
SO
, 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.
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
(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 (
G). If the
pump translocates n ions/cycle, this equilibrium condition is
given by the following equation:
G = n(RT
2.3
pH + 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
F
-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.