From the Departments of Genetics and Cellular & Molecular Physiology, Yale University School of Medicine, New Haven, Connecticut 06510
Received for publication, March 15, 2001, and in revised form, April 13, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In P2-type ATPases, a stalk
region connects the cytoplasmic part of the molecule, which binds and
hydrolyzes ATP, to the membrane-embedded part through which cations are
pumped. The present study has used cysteine scanning mutagenesis to
examine structure-function relationships within stalk segment 5 (S5) of
the yeast plasma-membrane H+-ATPase. Of 29 Cys mutants that
were made and examined, two (G670C and R682C) were blocked in
biogenesis, presumably due to protein misfolding. In addition, one
mutant (S681C) had very low ATPase activity, and another (F685C)
displayed a 40-fold decrease in sensitivity to orthovanadate,
reflecting a shift in equilibrium from the E2
conformational state toward E1. By far the most striking group of mutants (F666C, L671C, I674C, A677C, I684C, R687C, and Y689C)
were constitutively activated even in the absence of glucose, with
rates of ATP hydrolysis and kinetic properties normally seen only in
glucose-metabolizing cells. Previous work has suggested that activation
of the wild-type H+-ATPase results from kinase-mediated
phosphorylation in the auto-inhibitory C-terminal region of the 100-kDa
polypeptide. The seven residues identified in the present study are
located on one face of the S5 The plasma membrane H+-ATPase of yeast belongs to the
P2 subfamily of cation-transporting ATPases and is
structurally and functionally related to the
Na+,K+-, H+,K+-, and
Ca2+-ATPases of animal cells (1). All of these enzymes have
a 100-kDa catalytic subunit that is anchored in the membrane by four
hydrophobic segments (M1-4) at the N-terminal end of the molecule and
six hydrophobic segments (M5-10) at the C-terminal end. They split ATP
by way of a covalent Recently, the three-dimensional structure of the sarcoplasmic reticulum
Ca2+-ATPase has been solved at 2.6 Å (2),
furnishing a valuable framework for further work on the transport
mechanism of the P2-type ATPases. The cytoplasmic part of
the molecule includes the N- and C-terminal segments, the small
hydrophilic loop between M2 and M3, and the large hydrophilic loop
between M4 and M5 and is organized into three domains: (a) N
(ATP-binding domain), (b) P (phosphorylation), and
(c) A ("actuator" or anchor for domain N). Within the
membrane, the 10 hydrophobic As seen earlier by cryoelectron microscopy of the sarcoplasmic
reticulum enzyme (3) and the plasma membrane H+-ATPase of
Neurospora crassa (4), a stalk-like region connects the
cytoplasmic part of the molecule to the membrane. Not surprisingly, there is growing evidence that the stalk plays a conformationally active role. In a recent study of the yeast plasma membrane
H+-ATPase, our laboratory carried out scanning mutagenesis
to examine structure-function relationships in stalk segment 4 (S4),
which forms a short, relatively well-conserved link between the
phosphorylated aspartate residue (Asp-378) and M4. Kinetic analysis of
mutants along the entire length of S4 revealed 13 consecutive positions at which amino acid substitutions led to a shift in equilibrium from
the E2 state of the ATPase toward the E1 state
(5). Mutagenesis studies of sarcoplasmic reticulum
Ca2+-ATPase have revealed conformationally important
residues in S4 of that enzyme as well (6, 7).
In the work described here, we have turned to stalk segment 5 (S5),
located at the opposite end of the large cytoplasmic loop. In the
sarcoplasmic reticulum Ca2+-ATPase, M5 and S5 form a
continuous Yeast Strains--
Two related strains of Saccharomyces
cerevisiae were used in this study: SY4
(MATa, ura3-52, leu2-3, 112, his4-619, sec6-4ts GAL2, pma1::YIpGAL-PMA1;
Ref. 8) and NY13 (MATa, ura3-52; generously provided by Dr. Peter Novick of the Department of Cell Biology, Yale School of Medicine). In strain SY4, the chromosomal copy
of the PMA1 gene has been placed under control of the
GAL1 promotor by gene disruption (9) using the integrating
plasmid, YIpGAL-PMA1 (8). SY4 also carries the temperature-sensitive sec6-4 mutation, which, upon incubation at 37 °C, blocks
the fusion of secretory vesicles with the plasma membrane (10).
Mutagenesis--
Cysteine substitutions were made by polymerase
chain reaction (11) in a 519-base pair BglII-SalI
restriction fragment of the PMA1 gene that had been
subcloned into a modified Bluescript plasmid (Stratagene, La Jolla,
CA). After DNA sequencing, the BglII-SalI restriction fragment
carrying the mutation was moved into plasmid pPMA1.2 (9). The
3.8-kilobase HindIII-SacI fragment, which
contains the entire PMA1 coding region, was then cloned into
yeast expression vector YCp2HSE (9), placing the mutant pma1
allele under the control of two tandemly arranged heat-shock elements.
Plasmids were then transformed into yeast according to the method of
Ito et al. (12).
Isolation of Secretory Vesicles and Quantitation of Expressed
ATPase--
Transformed SY4 cells were grown to mid-exponential phase
(A600 ~ 1) at 23 °C in minimal medium
containing 2% galactose, shifted to medium containing 2% glucose for
3 h, and then heat-shocked at 39 °C for an additional 2 h.
The cells were harvested and washed, and secretory vesicles were
isolated and suspended in 0.8 M sorbitol, 1 mM
EDTA, and 10 mM triethanolamine/acetic acid, pH 7.2 as described previously (13). To determine the level of expressed Pma1
protein, secretory vesicles (5-20 µg) were subjected to
SDS-polyacrylamide gel electrophoresis and immunoblotted. Quantitative
PhosphorImager (Molecular Dynamics) analysis was carried out at two
protein concentrations within the linear range, and the expression
level was calculated from the average of two or more determinations.
Chromosomal Integration of pma1 Mutants--
To test for changes
in glucose regulation of the ATPase, selected mutations were introduced
into the chromosomal copy of the PMA1 gene. For this
purpose, a BglII-SalI fragment carrying the mutation was first subcloned into plasmid pGW201 (generously provided by Dr. David Perlin, Public Health Research Institute, New York, NY;
Ref. 14) to create a mutant pma1 allele with the
URA3 marker at its 3' non-coding end, and the presence of
the mutation was confirmed by DNA sequencing. A 6.1-kilobase
HindIII fragment containing the mutant allele linked to
URA3 was then excised from the plasmid and
transformed into yeast strain NY13 using the Alkali-Cation Yeast
Transformation kit (Bio 101). Finally, the presence of the mutation was
reconfirmed by polymerase chain reaction amplification of chromosomal
DNA and DNA sequence analysis.
Preparation of Plasma Membranes--
Glucose-starved and
glucose-metabolizing cells were prepared according to the protocol of
Serrano (15). Briefly, cells were grown to mid-exponential phase
(A600 = 4-6) in minimal medium containing 4%
glucose at 30 °C, harvested, washed twice with H2O, and
incubated for 1 h in water without glucose (glucose-starved cells); 4% glucose was then added back to an aliquot of the culture for 30 min to obtain glucose-metabolizing cells. In both cases, a
microsomal membrane fraction was prepared as described by Perlin et al. (16), and plasma membranes were isolated from this
fraction by a modified version of the method of Seto-Young et
al. (17). Specifically, the microsomal fraction was suspended in
membrane wash buffer containing 10 mM Tris, pH 7.0, 20%
(v/v) glycerol, 1 mM EGTA, 1 mM EDTA, 1 mM dithiothreitol, and a protease inhibitor mixture
(PIC).1 The suspension (2.0 ml) was applied to a discontinuous gradient of 0.9 ml of 65% (w/v)
sucrose and 2.0 ml of 54% (w/v) sucrose containing 10 mM
Tris, pH 7.0, and 1 mM EDTA. After centrifugation for
2 h at 183,000 × g (SW55 Ti rotor; Beckman)
plasma membranes were collected from the interface, diluted 5-fold with
1 mM EGTA/Tris (pH 7.5)/PIC, and centrifuged at
100,000 × g for 1 h. The plasma membrane pellet
was resuspended in 1 mM EGTA/Tris (pH 7.5)/PIC and stored
at ATP Hydrolysis and Proton Transport--
Unless otherwise noted,
ATP hydrolysis was assayed at 30 °C in 0.5 ml of 50 mM
4-morpholineethanesulfonic acid/Tris, pH 5.7, 5 mM
KN3, 5 mM Na2ATP, 10 mM
MgCl2, and an ATP regenerating system (5 mM
phosphoenolpyruvate and 50 µg/ml pyruvate kinase). The reaction was
terminated after 20-40 min, and the release of inorganic phosphate from ATP was measured by the method of Fiske and Subbarow (18). Specific activity was calculated as the difference between ATP hydrolysis in the presence and absence of 100 µM sodium
orthovanadate, a potent inhibitor of P-type ATPases. IC50
values for vanadate inhibition were determined by measuring ATP
hydrolysis in the presence of increasing concentrations of vanadate.
For determination of Km values, Na2ATP
was varied between 0.15 and 7.5 mM with MgCl2
always in excess of ATP by 5 mM; actual concentrations of
MgATP were calculated by the method of Fabiato and Fabiato (19). To
determine the pH optimum for ATP hydrolysis, the pH of the assay medium
was adjusted to values between 5.2 and 7.5 with Tris base.
ATP-dependent proton transport was determined by measuring
the initial rate of acridine orange fluorescence quenching as described by Ambesi et al. (20). The specific initial rate of
fluorescence quenching for each mutant was adjusted for ATPase
expression and reported as a percentage of the wild-type rate.
Protein Determination--
Protein concentrations were assayed
by the method of Lowry et al. (21) or as modified by Ambesi
et al. (13), with bovine serum albumin as standard.
Selection of Residues for Mutagenesis--
Previous work from our
laboratory has examined structure-function relationships in M5 (Ser-690
through Leu-713) of the yeast Pma1 H+-ATPase (22). In the
study described here, residues from the adjacent S5 region (Ala-661
through Tyr-689) were subjected to cysteine scanning mutagenesis. As
shown in Fig. 1, this region is strongly
conserved throughout the P2-type H+-ATPases of
fungi, higher plants, and protozoa; there is also recognizable homology
with the Na+,K+-,
H+,K+-, and Ca2+-ATPases of animal
cells.
Each of the 29 residues was replaced with Cys, and the mutant alleles
were cloned into expression vector YCp2HSE, transformed into yeast
strain SY4, and expressed under the control of a heat-shock promoter
after turning off the wild-type PMA1 allele (9). Secretory vesicles containing newly synthesized mutant ATPase were then isolated
and analyzed (13).
Expression and ATP Hydrolysis--
As measured by quantitative
immunoblotting (Table I), only two of the
mutant ATPases (G670C and R682C) failed to reach the secretory
vesicles, presumably due to protein misfolding; previous studies have
shown that such mutant forms are typically arrested in the endoplasmic
reticulum (23-25). It is worth noting that Gly-670 is strictly
conserved in all known P2-type H+-ATPases, and
Arg-682 is found in mammalian Na+,K+-,
H+,K+-, and Ca2+-ATPases as well
(Fig. 1).
The remaining 27 mutant H+-ATPases appeared in the
secretory vesicles at 28-109% of the level seen in the wild-type
control. Likewise, 26 of the 27 mutants were able to hydrolyze ATP at
rates ranging from 40% to 151%, after correction for the level of
expression in the secretory vesicles (Table I). Only one mutant (S681C) showed a major reduction in ATP hydrolysis (to 9%), and even here, the
uncorrected ATPase activity was 5-fold greater than the background activity observed in empty-plasmid controls.
Proton Transport--
Because S5 links the large hydrophilic loop
to M5, a membrane segment known to be involved in cation transport (2,
26-30), it was important to test the effects of the Cys substitutions on the ability of the ATPase to pump protons. This was assayed by
fluorescence quenching of the pH-sensitive dye acridine orange (Table
I). In mutants A677C and S681C, ATP-dependent quenching was
clearly above background, but the rates were below the limit at which
they could be measured quantitatively by the acridine orange assay. In
another mutant, Y689C, the rate of proton transport (175%) appeared to
exceed the rate of ATP hydrolysis (107%), but further experiments
showed that this discrepancy could be explained by a shift in the pH
optimum of the mutant enzyme (see below). For all of the other mutants,
the initial rate of ATP-dependent acridine orange quenching
closely paralleled the rate of ATP hydrolysis, indicating that the
substitutions had little or no effect on the coupling between transport
and hydrolysis.
Kinetic Properties--
The mutant ATPases were next assayed for
MgATP dependence, vanadate sensitivity, and the effect of pH on the
rate of ATP hydrolysis, parameters that can provide a useful clue to
changes in reaction mechanism. As summarized in Table
II, there was one mutant (F685C) that
displayed an increase in the IC50 for vanadate (to 42 µM) and a decrease in the apparent Km
for MgATP (to 0.3 mM); F685C also showed a measurable
alkaline shift in pH optimum (to pH 6.0). As pointed out previously,
coordinated changes of this kind can readily be accounted for by a
shift in conformational equilibrium from the E2 state (high
affinity for vanadate; low affinity for MgATP) to the E1
state (low affinity for vanadate; high affinity for ATP) (5, 20, 22).
Similar mutants have been described for mammalian
Na+,K+-ATPase by Daly et al. (31)
and Boxenbaum et al. (32).
More striking were seven mutants (F666C, L671C, I674C, A677C, I684C,
R687C, and Y689C) with a 2- to 3-fold decrease in the apparent
Km for MgATP (to 0.2-0.3 mM) and a rise
in pH optimum (to pH 6.0-6.3) but no change (or even a slight
decrease) in the IC50 for vanadate (Table II). This
constellation of kinetic changes is reminiscent of that seen when
wild-type yeast cells, previously starved of glucose, are returned to a
glucose-containing medium. Under such conditions, the
H+-ATPase undergoes a rapid 10-fold increase in activity,
along with a lowering of the Km for MgATP and an
alkaline shift of pH optimum (33). There is evidence that glucose
activation results from kinase-mediated phosphorylation of the ATPase,
very likely in the C-terminal region of the polypeptide (see
"Discussion"). Thus, it seemed possible that the seven
mutations listed above may have somehow shifted the ATPase into a
constitutively activated state.
Glucose Activation--
To test this idea, the seven mutations
highlighted in Table I were integrated into the chromosomal copy of the
PMA1 gene to see whether the H+-ATPase was still
capable of glucose activation at the plasma membrane. For the
experiments to be described, wild-type and mutant cells were grown to
mid-exponential phase in glucose medium and incubated under two
conditions: (a) 90 min of starvation in glucose-free medium,
and (b) 60 min of starvation followed by a 30-min incubation in medium containing 4% glucose. Plasma membranes were then isolated and assayed.
Based on quantitative immunoblotting, wild-type and mutant plasma
membranes contained roughly equal amounts of Pma1 ATPase, and there was
little if any change in the amount when starved cells were incubated
briefly with glucose (Table III).
Activity measurements revealed a striking difference between the
wild-type and mutant enzymes, however. Across a wide range of pH
values, the wild-type ATPase had relatively low activity in starved
cells but was conspicuously stimulated when the cells were treated with glucose (Fig. 2); the increase in
activity ranged from 4.4-fold at pH 5.7 to 9.0-fold at pH 6.25 (Table
III). The seven mutant ATPases, on the other hand, were already
activated in starved cells and changed very little upon glucose
treatment (Fig. 2; Table III).
Glucose also had a less pronounced effect on the kinetic properties of
the mutant ATPases (Table IV). Whereas
the Km of the wild-type enzyme for MgATP fell 8-fold
(from 3.1 mM to 0.4 mM) upon incubation with
glucose, the Km values of the mutant enzymes were
already reduced in starved cells (0.2-1.2 mM). Likewise,
although the pH optimum of the wild-type ATPase displayed a small but
reproducible alkaline shift (from pH 5.7 to pH 6.1) after glucose
treatment, the mutants had pH optima between pH 6.0 and pH 6.3, even
under starvation conditions. Taken together, the kinetic findings
reinforce the notion that single amino acid substitutions in S5 have
led to a constitutive activation of the Pma1 ATPase.
The original goal of this study was to identify amino acid
residues in stalk segment 5 that play an important role in the reaction
cycle of the yeast Pma1 H+-ATPase. The strategy, as
described previously (5, 20), was to screen for mutations that increase
the IC50 for vanadate by shifting the conformational
equilibrium away from the E2 state (to which vanadate
binds) toward E1; typically, such mutants also display a
reduced Km for MgATP and an alkaline change in pH
optimum. Unlike stalk segment 4, where mutations of this type occurred
at 13 consecutive positions, only 1 vanadate-resistant mutant (F685C)
was seen in S5. At the corresponding position of the sarcoplasmic
reticulum Ca2+-ATPase (Tyr-754), Sorensen and Andersen (34)
have also observed kinetic changes upon substitution with Ala, but in
this case, there was an increase in apparent affinity for vanadate,
reflecting an elevated dephosphorylation rate of E2P to
E2.
A closer correspondence between the two ATPases was evident at Arg-682
(Arg-751 in the SERCA Ca2+-ATPase), a residue found
in all known P-type ATPases. Here, substitution by Cys in the
H+-ATPase or by Ala, Ile, or Glu in the
Ca2+-ATPase virtually abolished expression, suggesting that
the conserved Arg plays an essential role in protein folding. Indeed,
in the recently published crystal structure of the
Ca2+-ATPase (2), Arg-751 interacts with several residues in
the cytoplasmically exposed M6-M7 loop; it is easy to imagine that these interactions may serve to stabilize the protein.
By far the most significant finding of the present study was that seven
periodically spaced residues in S5 play a role in the regulation of
yeast H+-ATPase by glucose. Rapid, reversible
down-regulation of ATPase activity during glucose starvation was first
observed by Serrano (33), who found it to be associated with a
reduction in the apparent affinity for MgATP and a shift in pH optimum
from pH 6.0 to pH 5.7. Since then, evidence has mounted that
down-regulation is mediated by the C terminus of the ATPase,
functioning as an auto-inhibitory domain. Consistent with this idea,
the yeast ATPase can be activated by deleting 11 or 18 amino acid
residues from the C terminus (35, 36). Although the mechanism is not
yet fully understood in molecular terms, glucose has been shown to trigger kinase-mediated phosphorylation of the ATPase on one or more
Ser/Thr residues (37), and site-directed mutagenesis has identified two
potential phosphorylation sites in the C-terminal region (Ser-899 and
Thr-912) at which amino acid substitutions modify glucose regulation
(38). A recent study by Goossens et al. (39) provides
evidence that a novel protein kinase known as Ptk2 acts on the first of
the two sites, whereas the second is a potential phosphorylation site
for calmodulin-dependent protein kinase II. Based on all of
these findings, it is attractive to think that glucose induces the
phosphorylation of Ser-899 and Thr-912, somehow displacing the C
terminus and releasing the ATPase from auto-inhibition. A more detailed
discussion of this hypothesis can be found in a recent review by
Portillo (40).
If the C terminus can indeed inhibit the activity of the yeast
H+-ATPase, one would like to know how it does so. A
previous study by Eraso and Portillo (41) took a useful step toward
answering this question by isolating intragenic suppressors of the
double mutant S911A/T912A, which exhibits little or no glucose
activation and cannot grow on glucose medium. Fourteen second-site
suppressor mutations were isolated by their ability to restore growth
on glucose. Of them, four mapped in the C-terminal region, three mapped
in S2, two mapped in S4, four mapped in the presumed ATP-binding domain
of the large cytoplasmic loop, and two mapped toward the end of the
cytoplasmic loop in the region that we have defined as S5. Direct
assays of isolated plasma membranes showed that all of the suppressor
mutations produced significant increases in ATPase activity in
glucose-starved cells. However, the kinetic properties of the
mutant enzymes varied, with some exhibiting changes in
Km and pH optimum characteristic of the activated state, and others not doing so. Most relevant to the present study are
the two suppressor mutations mapping in S5. One, G670S, led to a 7-fold
increase in the rate of ATP hydrolysis in starved cells, along with a
significant drop in Km (from 4.0 to 1.2 mM), a rise in pH optimum (from pH 5.5 to pH 6.3), and a
fall in the IC50 for vanadate (from 10.0 to 0.5 µM); it therefore met the criteria for constitutive
activation as described above. The other suppressor mutant isolated by
Eraso and Portillo (41), P669L, displayed a 10-fold elevation in the
rate of hydrolysis in starved cells, accompanied by a smaller rise in
pH optimum (to pH 6.0) and a fall in the IC50 (to 0.9 µM) but not by a detectable change in
Km; thus, it seems to be an intermediate case. Of
the related mutations that were constructed in the present study, G670C
blocked biogenesis and could not be characterized, whereas P669C had
reasonable specific activity, a slightly lowered Km
and IC50, and a slightly elevated pH optimum (see Table
II).
At the structural level, it is presumably significant that the seven
constitutively activated mutants described in the present study map to
one face of an -helix, consistent with the
idea that mutations along this face serve to release the
auto-inhibition.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-aspartyl phosphate intermediate and alternate
between two major conformational states (E1 and
E2) during the reaction cycle.
-helices assemble together in a complex
way, with 4 of the inner helices (M4, M5, M6, and M8) providing the
actual Ca2+-liganding residues.
-helix, 60-Å long, stretching from the membrane into the
cytoplasm and seeming to serve as a "center mast" around which the
rest of the molecule is organized (2). To discover whether S5 of the
yeast H+-ATPase contains conformationally active residues,
we performed cysteine scanning mutagenesis along the entire segment,
from Ala-661 to Tyr-689, followed by kinetic analysis of each of the
mutants. Based on the results, only one residue influences the
equilibrium between E1 and E2 conformations. By
contrast, there are seven residues at which mutations alter the
regulation of the ATPase by glucose, leading to a constitutively
activated form of the enzyme.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
70 °C. All procedures were carried out at 4 °C.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (18K):
[in a new window]
Fig. 1.
Sequence alignment of stalk segment 5. The 29-amino acid stretch of stalk segment 5 has been aligned for nine
representative P2-type ATPases. Swiss Protein Database
sequence accession numbers are (from top to
bottom) P05030, P07038, P09627, P54210, P20649, Q00804,
P04191, P04074, and P09626. Residues identical to the yeast Pma1
sequence are indicated by a period.
Effect of mutations in stalk segment 5 on H+-ATPase expression
and activity in secretory vesicles
Kinetic properties of mutant H+-ATPases in secretory vesicles
H+-ATPase activity in plasma membranes isolated from
glucose-starved and glucose-metabolizing cells
View larger version (21K):
[in a new window]
Fig. 2.
Effect of glucose on ATPase activity as a
function of pH. Plasma membranes from yeast cells expressing
wild-type (top) or Y689C (bottom) ATPase were
isolated from glucose-starved cells ( ) or glucose-metabolizing cells
(
). ATPase activity was determined as a function of pH as described
under "Experimental Procedures."
Kinetic properties of mutant ATPase in the plasma membrane
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-helix, along with the P669L and G670S mutations of
Eraso and Portillo (Ref. 41; Fig.
3). Also on the same face are
F685C (a mutation affecting the equilibrium between E1 and
E2 conformations; see above) and R682C (one of two
mutations leading to arrest in the endoplasmic reticulum). Although it
is tempting to think that this face of the S5
-helix may interact
directly with the C terminus to bring about auto-inhibition, the
interaction may equally well be indirect; high-resolution structures of
the ATPase in the glucose-starved and glucose-metabolizing states will
be required to settle this question.
View larger version (40K):
[in a new window]
Fig. 3.
Helical wheel analysis of S5. Residues
at which mutations lead to constitutive activation are highlighted by a
shaded box (this study) or an empty box
(41). Residues at which mutations cause a block in biogenesis
(R682C) or a shift in the E1-E2 equilibrium
(F685C) are marked with an asterisk.
In the meantime, it is interesting to consider glucose regulation of
the yeast H+-ATPase in a more general framework. Although
N- and C-terminal sequences have been poorly conserved among
P2-type ATPases, both termini have taken on regulatory
roles in various members of the group. For example, the C terminus of
the plant plasma membrane H+-ATPase, which is considerably
longer than that of the yeast enzyme, has been well documented to
function as an auto-inhibitory domain, and inhibition is reversed by
the binding of 14-3-3 proteins to the C terminus (reviewed by Morsomme
and Boutry (42)). Once again, mutations elsewhere in the ATPase can
also reverse the inhibition (43, 44). The plasma membrane
Ca2+-ATPase of animal cells is also regulated by means of
the C terminus, where calmodulin binding stimulates the rate of ATP
hydrolysis and improves the affinities for Ca2+ and ATP
(reviewed by Carafoli and Brini (45)). On the other hand, a recently
described plant endomembrane Ca2+-ATPase is thought to be
activated by the binding of calmodulin to its very long N terminus
(46). Thus, as one might have predicted, the P2-type
ATPases have evolved a variety of regulatory mechanisms, each involving
its own auxiliary molecules and binding sites. Given the high degree of
interest in this area, the next few years are likely to see continued
progress toward understanding the molecular mechanisms of regulation.
![]() |
ACKNOWLEDGEMENTS |
---|
We are grateful to Brett Mason, Valery Petrov, and Anthony Ambesi for helpful discussions.
![]() |
FOOTNOTES |
---|
* This work was supported by National Institutes of Health Grant GM-15761 and by a Fogarty Postdoctoral Fellowship (to M. M.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed: Depts. of Genetics and
Cellular & Molecular Physiology, Yale University School of Medicine,
333 Cedar St., New Haven, CT 06510. Tel.: 203-785-2690; Fax:
203-737-1771.
§ Present address: Universidad Nacional Autonoma de Mexico, Departamento de Bioquimica, Facultad de Medicina, Mexico, D.F., 04510 Mexico.
Published, JBC Papers in Press, April 16, 2001, DOI 10.1074/jbc.M102332200
![]() |
ABBREVIATIONS |
---|
The abbreviation used is: PIC, protease inhibitor mixture (1 µg/ml leupeptin, 1 µg/ml pepstatin, 1 µg/ml aprotinin, and 2 µg/ml chymostatin).
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Lutsenko, S., and Kaplan, J. H. (1995) Biochemistry 34, 15607-15613[Medline] [Order article via Infotrieve] |
2. | Toyoshima, C., Nakasako, M., Nomura, H., and Ogawa, H. (2000) Nature 405, 647-655[CrossRef][Medline] [Order article via Infotrieve] |
3. | Zhang, P., Toyoshima, C., Yonekura, K., Green, N. M., and Stokes, D. L. (1998) Nature 392, 835-839[CrossRef][Medline] [Order article via Infotrieve] |
4. | Auer, M., Scarborough, G. A., and Kuhlbrandt, W. (1998) Nature 392, 840-843[CrossRef][Medline] [Order article via Infotrieve] |
5. |
Ambesi, A.,
Miranda, M.,
Allen, K. E.,
and Slayman, C. W.
(2000)
J. Biol. Chem.
275,
20545-20550 |
6. |
Zhang, Z.,
Sumbilla, C.,
Lewis, D.,
Summers, S.,
Klein, M. G.,
and Inesi, G.
(1995)
J. Biol. Chem.
270,
16283-16290 |
7. | Garnett, C., Sumbilla, C., Belda, F. F., Chen, L., and Inesi, G. (1996) Biochemistry 35, 11019-11025[CrossRef][Medline] [Order article via Infotrieve] |
8. |
Nakamoto, R. K.,
Rao, R.,
and Slayman, C. W.
(1991)
J. Biol. Chem.
266,
7940-7949 |
9. | Cid, A., Perona, R., and Serrano, R. (1987) Curr. Genet. 12, 105-111[Medline] [Order article via Infotrieve] |
10. | Novick, P., Field, C., and Schekman, R. (1980) Cell 21, 205-215[Medline] [Order article via Infotrieve] |
11. | Sarkar, G., and Sommer, S. S. (1990) BioTechniques 8, 404-407[Medline] [Order article via Infotrieve] |
12. | Ito, H., Fukuda, Y., Murata, K., and Kimura, A. (1983) J. Bacteriol. 153, 163-168[Medline] [Order article via Infotrieve] |
13. | Ambesi, A., Allen, K. E., and Slayman, C. W. (1997) Anal. Biochem. 251, 127-129[CrossRef][Medline] [Order article via Infotrieve] |
14. |
Wang, G.,
Tamas, M.,
Hall, M. J.,
Pascual-Ahuir, A.,
and Perlin, D. S.
(1996)
J. Biol. Chem.
271,
25438-25445 |
15. | Serrano, R. (1988) Biochim. Biophys. Acta 947, 1-28[Medline] [Order article via Infotrieve] |
16. |
Perlin, D. S.,
Harris, S. L.,
Seto-Young, D.,
and Haber, J. E.
(1989)
J. Biol. Chem.
264,
21857-21864 |
17. | Seto-Young, D., Monk, B. C., and Perlin, D. (1992) Biochim. Biophys. Acta 1102, 213-219[Medline] [Order article via Infotrieve] |
18. |
Fiske, C. H.,
and Subbarow, Y.
(1925)
J. Biol. Chem.
66,
375-400 |
19. | Fabiato, A., and Fabiato, F. (1979) J. Physiol. Paris 75, 463-505[Medline] [Order article via Infotrieve] |
20. |
Ambesi, A.,
Pan, R. L.,
and Slayman, C. W.
(1996)
J. Biol. Chem.
271,
22999-23005 |
21. |
Lowry, O. H.,
Rosebrough, N. J.,
Farr, A. L.,
and Randall, R. J.
(1951)
J. Biol. Chem.
193,
265-275 |
22. |
Dutra, M. B.,
Ambesi, A.,
and Slayman, C. W.
(1998)
J. Biol. Chem.
273,
17411-17417 |
23. |
Nakamoto, R. K.,
Verjovski-Almeida, S.,
Allen, K. E.,
Ambesi, A.,
Rao, R.,
and Slayman, C. W.
(1998)
J. Biol. Chem.
273,
7338-7344 |
24. |
DeWitt, N. D.,
Tourinho dos Santos, C. F.,
Allen, K. E.,
and Slayman, C. W.
(1998)
J. Biol. Chem.
273,
21744-21751 |
25. |
Sen Gupta, S.,
DeWitt, N. D.,
Allen, K. E.,
and Slayman, C. W.
(1998)
J. Biol. Chem.
273,
34328-34334 |
26. | Clarke, D. M., Loo, T. W., Inesi, G., and MacLennan, D. H. (1989) Nature 339, 476-478[CrossRef][Medline] [Order article via Infotrieve] |
27. |
Clarke, D. M.,
Loo, T. W.,
and MacLennan, D. H.
(1990)
J. Biol. Chem.
265,
6262-6267 |
28. |
Arguello, J. M.,
Peluffo, R. D.,
Feng, J.,
Lingrel, J. B.,
and Berlin, J. R.
(1996)
J. Biol. Chem.
271,
24610-24616 |
29. | Nielsen, L. M., Pedersen, P. A., Karlish, S. J. D., and Jorgensen, P. L. (1998) Biochemistry 37, 1961-1968[CrossRef][Medline] [Order article via Infotrieve] |
30. |
Petrov, V. V.,
Padmanabha, K. P.,
Nakamoto, R. K.,
Allen, K. E.,
and Slayman, C. W.
(2000)
J. Biol. Chem.
275,
15709-15716 |
31. |
Daly, S. E.,
Lane, L. K.,
and Blostein, R.
(1996)
J. Biol. Chem.
271,
23683-23689 |
32. |
Boxenbaum, N.,
Daly, S. E.,
Javaid, Z. Z.,
Lane, L. K.,
and Blostein, R.
(1998)
J. Biol. Chem.
273,
23086-23092 |
33. | Serrano, R. (1983) FEBS Lett. 156, 11-14[CrossRef][Medline] [Order article via Infotrieve] |
34. |
Sorensen, T. L.-M.,
and Andersen, J. P.
(2000)
J. Biol. Chem.
275,
28954-28961 |
35. | Portillo, F., de Larrinoa, I. F., and Serrano, R. (1989) FEBS Lett. 247, 381-385[CrossRef][Medline] [Order article via Infotrieve] |
36. | Mason, A. B., Kardos, T. B., and Monk, B. C. (1998) Biochim. Biophys. Acta 1372, 261-271[Medline] [Order article via Infotrieve] |
37. | Chang, A., and Slayman, C. W. (1991) J. Cell Biol. 115, 289-295[Abstract] |
38. | Portillo, F., Eraso, P., and Serrano, R. (1991) FEBS Lett. 287, 71-74[CrossRef][Medline] [Order article via Infotrieve] |
39. |
Goossens, A.,
De la Fuente, N.,
Forment, J.,
Serrano, R.,
and Portillo, F.
(2000)
Mol. Cell. Biol.
20,
7654-7661 |
40. | Portillo, F. (2000) Biochim. Biophys. Acta 1469, 31-42[Medline] [Order article via Infotrieve] |
41. |
Eraso, P.,
and Portillo, F.
(1994)
J. Biol. Chem.
269,
10393-10399 |
42. | Morsomme, P., and Boutry, M. (2000) Biochim. Biophys. Acta 1465, 1-16[Medline] [Order article via Infotrieve] |
43. | Morsomme, P., de Kerchove d'Exaerde, A., De Meester, S., Thines, D., Goffeau, A., and Boutry, M. (1996) EMBO J. 15, 5513-5526[Abstract] |
44. |
Morsomme, P.,
Dambly, S.,
Maudoux, O.,
and Boutry, M.
(1998)
J. Biol. Chem.
273,
34837-34842 |
45. | Carafoli, E., and Brini, M. (2000) Curr. Opin. Chem. Biol. 4, 152-161[CrossRef][Medline] [Order article via Infotrieve] |
46. |
Curran, A.,
Hwang, I.,
Corbin, J.,
Martinez, S.,
Rayle, D.,
Sze, H.,
and Harper, J. F.
(2000)
J. Biol. Chem.
275,
30301-30308 |