Chimeras of X+,K+-ATPases
THE M1-M6 REGION OF Na+,K+-ATPase
IS REQUIRED FOR Na+-ACTIVATED ATPase ACTIVITY, WHEREAS THE
M7-M10 REGION OF H+,K+-ATPase IS INVOLVED IN
K+ DE-OCCLUSION*
Jan B.
Koenderink,
Herman G. P.
Swarts,
H. Christiaan
Stronks,
Harm P. H.
Hermsen,
Peter H. G. M.
Willems, and
Jan Joep
H. H. M.
De Pont
From the Department of Biochemistry, Institute of Cellular
Signalling, University of Nijmegen, P. O. Box 9101, 6500 HB
Nijmegen, The Netherlands
Received for publication, November 30, 2000, and in revised form, January 4, 2001
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ABSTRACT |
In this study we reveal regions of
Na+,K+-ATPase and
H+,K+-ATPase that are involved in cation
selectivity. A chimeric enzyme in which transmembrane hairpin M5-M6 of
H+,K+-ATPase was replaced by that of
Na+,K+-ATPase was phosphorylated in the absence
of Na+ and showed no K+-dependent
reactions. Next, the part originating from
Na+,K+-ATPase was gradually increased in the
N-terminal direction. We demonstrate that chimera HN16, containing the
transmembrane segments one to six and intermediate loops of
Na+,K+-ATPase, harbors the amino acids
responsible for Na+ specificity. Compared with
Na+,K+-ATPase, this chimera displayed a similar
apparent Na+ affinity, a lower apparent K+
affinity, a higher apparent ATP affinity, and a lower apparent vanadate
affinity in the ATPase reaction. This indicates that the
E2K form of this chimera is less stable
than that of Na+,K+-ATPase, suggesting that it,
like H+,K+-ATPase, de-occludes K+
ions very rapidly. Comparison of the structures of these chimeras with
those of the parent enzymes suggests that the C-terminal 187 amino
acids and the
-subunit are involved in K+ occlusion.
Accordingly, chimera HN16 is not only a chimeric enzyme in structure,
but also in function. On one hand it possesses the Na+-stimulated ATPase reaction of
Na+,K+-ATPase, while on the other hand it has
the K+ occlusion properties of
H+,K+-ATPase.
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INTRODUCTION |
Enzymes that belong to the family of P-type ATPases can facilitate
active transport of cations across the plasma membrane (1).
Na+,K+-ATPase and gastric
H+,K+-ATPase are two particular members of this
family, since they possess a
-subunit and couple ATP hydrolysis to
counter-transport of cations. The latter process can be described by
the Albers-Post scheme (2, 3). Although
Na+,K+-ATPase and
H+,K+-ATPase have many structural and
functional similarities, they differ in cation specificity. Both enzyme
activities need K+, but the
Na+,K+-ATPase activity is stimulated by
Na+ and that of H+,K+-ATPase by
H+. A major difference between the two enzymes is that the
occlusion of K+ can easily be measured in
Na+,K+-ATPase (4), whereas in
H+,K+-ATPase one needs very special conditions
to measure this (5). This difference is due to the much faster rate of
the E2K
E1K conversion in the latter enzyme (6, 38). It has been shown that four
negatively charged residues present in transmembrane segments 4-6 play
a role in cation-activated reactions of both ATPases (7-13). With the
exception of a single residue in
M61 (Asp804 in
Na+,K+-ATPase; Glu820 in
H+,K+-ATPase), these amino acid residues are
similar in both ATPases. Their analogous residues in SERCA1a
Ca2+-ATPase also play a role in Ca2+ binding as
revealed by the recently published crystal structure (14).
The catalytic
1-subunit of
Na+,K+-ATPase and that of gastric
H+,K+-ATPase share a high degree of identity
(63%) in contrast to their heavily glycosylated
1-subunits, which are only 30% identical. Assembly of
the
- and
-subunits is essential for enzyme activity (15) and
occurs before the subunits are transported from the endoplasmic
reticulum to the plasma membrane (16). Although the
-subunits can
form functional complexes with both
-subunits, they have a
preference for their own
-subunit (15). This preference is probably
determined by the C-terminal half of the extracellular loop between
transmembrane segments seven and eight of the
-subunit where the

interaction occurs (17, 18).
The similarity between the catalytic subunits of
Na+,K+-ATPase and
H+,K+-ATPase made it possible to prepare
chimeras and to test their catalytic properties. Because of the
postulated role of the M5-M6 hairpin in cation binding, we first
prepared a chimeric enzyme from gastric
H+,K+-ATPase in which only this hairpin was
replaced by that of Na+,K+-ATPase. ATP could
phosphorylate this chimeric enzyme, but no K+-stimulated
ATPase activity could be measured (19). Moreover, Na+ ions
did not stimulate the phosphorylation reaction. We next prepared
chimeras in which the part originating from
Na+,K+-ATPase was gradually increased into the
N-terminal direction. In all these chimeras the C-terminal part,
including the last four transmembrane segments and the
-subunit,
originated from the gastric H+,K+-ATPase. In
this study we show that only chimeras that contain all six N-terminal
transmembrane domains and their intervening loops display
Na+ activation of the ATPase activity and the ATP
phosphorylation reaction. Furthermore, we provide evidence for a role
of the C-terminal region in K+ de-occlusion.
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EXPERIMENTAL PROCEDURES |
Expression Constructs--
The rat gastric
H+,K+-ATPase
- and
-subunits and the rat
Na+,K+-ATPase
1- and
1-subunits were cloned into the pFastbacdual vector
(Life Technologies, Inc., Breda, The Netherlands) as described previously (15). We used the Altered Sites II in vitro
mutagenesis system (Promega, Madison, WI) to introduce silent mutations
to generate MunI, DraI, PvuI,
NheI, VspI, and SstII restriction
sites in the cDNA of the H+,K+-ATPase and
Na+,K+-ATPase
-subunits. These restriction
sites were chosen in the intracellular domain close to putative
transmembrane helices. A NarI restriction site was already
present in both
-subunits. Thereafter, the
VspI-SstII fragment, the
NarI-SstII fragment, the
NheI-SstII fragment, the
PvuI-SstII fragment, the
DraI-SstII fragment, the
MunI-SstII fragment, and the N terminus until
SstII were replaced by the similar fragments of
Na+,K+-ATPase, resulting in the chimeras HN56,
HNn6, HN46, HN36, HN26, HN16, and HNN6, respectively (Fig. 1). The
sequence of all mutants was verified.
Production of Recombinant Viruses--
Competent DH10bac
Escherichia coli cells (Life Technologies, Inc., Breda, The
Netherlands) harboring the baculovirus genome (bacmid) and a
transposition helper vector were transformed with the pFastbacdual
transfer vector containing different cDNAs. Upon transition between
the Tn7 sites in the transfer vector and the bacmid, recombinant
bacmids were selected and isolated (20). Subsequently, Sf9
insect cells were transfected with recombinant bacmids using
Cellfectin reagent (Life Technologies, Inc.). After 3 days, the
recombinant baculoviruses were harvested and used to infect Sf9
cells at a multiplicity of infection of 0.1. Four days after infection,
the amplified viruses were harvested. As a mock a baculovirus not
expressing the
- and
-subunit of
H+,K+-ATPase or
Na+,K+-ATPase was prepared.
Preparation of Membranes--
Sf9 cells were grown at
27 °C in 100-ml spinner flask cultures (21). For production of the
ATPases subunits, 1.0-1.5·106
cells·ml
1 were infected at a multiplicity
of infection of 1-3 in Xpress medium (BioWittaker,
Walkersville, MD) containing 1% (v/v) ethanol (22) and incubated for 3 days. The Sf9 cells were harvested by centrifugation at
2,000 × g for 5 min and resuspended at 0 °C in 0.25 M sucrose, 2 mM EDTA, and 25 mM
Hepes/Tris (pH 7.0). The membranes were sonicated twice for 30 s
at 60 watts (Branson Power Co., Danbury, CT), after which the disrupted
cells were centrifuged at 10,000 × g for 30 min. The
supernatant was recentrifuged at 100,000 × g for 60 min, and the pelleted membranes were resuspended in the above mentioned
buffer and stored at
20 °C.
Protein Determination--
Protein was determined with the
modified Lowry method described by Peterson (23) using bovine serum
albumin as a standard.
ATPase Activity Assay--
The ATPase activity was determined
with a radiochemical method (10). For this purpose Sf9 membranes
were added to 100 µl of medium, which contained under standard
conditions 50 mM Tris-acetic acid (pH 7.0), 0.2 mM EDTA, 0.1 mM EGTA, 1 mM
Tris-N3, 1.3 mM MgCl2, 10 mM KCl, 100 mM NaCl, and 100 µM
[
-32P]ATP. Other conditions are indicated in the
legends of the Figs. 3-6. After incubation at 37 °C, the
reaction was stopped by the addition of 500 µl 10% (w/v) charcoal in
6% (w/v) trichloroacetic acid, and after incubation at 0 °C, the
mixture was centrifuged for 30 s (10,000 × g). To
200 µl of the clear supernatant, containing the liberated inorganic
phosphate (32Pi), 4 ml of OptiFluor
(Canberra Packard, Tilburg, The Netherlands) was added, and the mixture
was analyzed by liquid scintillation analysis. Blanks were prepared by
incubating in the absence of enzyme. The specific activity is presented
as the difference between that of the expressed enzyme and the mock.
ATP Phosphorylation Assay--
ATP phosphorylation was
determined as described previously (15). Sf9 membranes
were incubated at 21 °C in 50 mM Tris-acetic acid (pH
6.0), 0.2 mM EDTA, 1.2 mM MgCl2 and
0-300 mM NaCl in a volume of 50 µl. After 30-60 min
preincubation 10 µl of 0.6 µM
[
-32P]ATP was added and incubated for 10 s at
21 °C. The reaction was stopped by adding 5% (w/v) trichloroacetic
acid in 0.1 M phosphoric acid, and the phosphorylated
protein was collected by filtration over a 0.8-µm membrane filter
(Schleicher and Schuell, Dassel, Germany). After repeated washing, the
filters were analyzed by liquid scintillation analysis. The specific
phosphorylation level is presented as the phosphorylation level
obtained with the expressed enzyme minus that of the mock.
Calculations--
The K0.5 value is
defined as the concentration of effector giving the half-maximal
activation and the IC50 as the value giving 50% inhibition
of the maximal activation. Data are presented as mean values with S.E.
of the mean. Differences were tested for significance by means of the
Student's t test.
Materials--
The rat cDNA clones of the
H+,K+-ATPase
- and
-subunits and the rat
and sheep cDNA clones of the Na+,K+-ATPase
1- and
1-subunits were provided by Drs.
G. E. Shull and J. B Lingrel, respectively. All enzymes used for
DNA cloning were purchased from Life Technologies, Inc.
[
-32P]ATP (3000 Ci·mmol
1)
was purchased from Amersham Pharmacia Biotech (Buckinghamshire, United Kingdom).
 |
RESULTS |
Seven chimeric constructs were produced after introduction of six
unique restriction sites in both the cDNAs of the rat gastric H+,K+-ATPase
-subunit and the rat
Na+,K+-ATPase
1-subunit. First,
the transmembrane hairpin M5-M6 of H+,K+-ATPase
was exchanged by the similar region of
Na+,K+-ATPase, generating the chimeric ATPase
HN56. Next, the part originating from
Na+,K+-ATPase was progressively increased in
the N-terminal direction, generating the chimeric ATPases HNn6, HN46,
HN36, HN26, HN16, and HNN6 (Fig. 1).
These chimeric
-subunits were introduced in the genome of a
baculovirus together with the H+,K+-ATPase
-subunit. As controls the wild-type
H+,K+-ATPase and
Na+,K+-ATPase (HK and NaK) were also expressed.
The recombinant baculoviruses were used to infect Sf9 insect
cells, and the membrane fractions of these cells expressing the ATPase
proteins were isolated. Western blot analysis, using the antibodies HKB
(24) (directed against the large intracellular loop of the
H+,K+-ATPase
-subunit), HK9 (25) (directed
against the N terminus of the H+,K+-ATPase
-subunit), L16 (26) (directed against M6 of
Na+,K+-ATPase), 2G11 (27) (directed against the
H+,K+-ATPase
-subunit), and G34 (28)
(directed against the Na+,K+-ATPase
-subunit), was used to check the produced proteins. This analysis
revealed similar expression levels for the wild-type H+,K+-ATPase,
Na+,K+-ATPase, and the chimeras HNn6, HN46,
HN36, HN26, HN16, and HNN6. The expression level of chimera HN56,
however, was slightly lower than that of the wild-type
H+,K+-ATPase (data not shown).

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Fig. 1.
Schematic representation of chimeras and
wild-type enzymes. The open lines represent
H+,K+-ATPase sequences, and the black
lines represent Na+,K+-ATPase sequences.
HK, H+,K+-ATPase; NaK,
Na+,K+-ATPase; HN56,
H+,K+-ATPase with amino acids
Leu776-Arg846 replaced by those of
Na+,K+-ATPase
(Leu762-Arg832); HNn6,
H+,K+-ATPase with amino acids
Ala519-Arg846 replaced by those of
Na+,K+-ATPase
(Ala505-Arg832); HN46,
H+,K+-ATPase with amino acids
Leu346-Arg846 replaced by those of
Na+,K+-ATPase
(Leu332-Arg832); HN36,
H+,K+-ATPase with amino acids
Ile293-Arg846 replaced by those of
Na+,K+-ATPase
(Ile279-Arg832); HN26,
H+,K+-ATPase with amino acids
Lys171-Arg846 replaced by those of
Na+,K+-ATPase
(Lys157-Arg832); HN16,
H+,K+-ATPase with amino acids
Leu105-Arg846 replaced by those of
Na+,K+-ATPase
(Leu91-Arg832); HNN6,
H+,K+-ATPase with amino acids
Met1-Arg846 replaced by those of
Na+,K+-ATPase
(Met 5-Arg832).
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Both Na+,K+-ATPase and
H+,K+-ATPase hydrolyze ATP to actively
transport cations across the plasma membrane. This ATPase activity was
measured at 100 µM ATP in the absence of K+
and Na+, in the presence of 10 mM
K+, and in the combined presence of K+ (10 mM) and Na+ (100 mM). The ATPase
activity of the wild-type H+,K+-ATPase was
stimulated by K+, but inhibited when Na+ was
additionally present (Fig. 2). The
chimeras HN56, HNn6, HN46, HN36, and HN26 did not possess significant
ATPase activity above that of the mock-infected cells, either in the
absence or the presence of K+. Addition of Na+
in the presence of K+ also had no effect on the activity of
the latter chimeras. In contrast to
H+,K+-ATPase, the ATPase activities of the
chimeras HN16 and HNN6 were only slightly stimulated by the addition of
K+, whereas similarly to that of the wild-type
Na+,K+-ATPase, the activities were strongly
stimulated by the combined presence of Na+ and
K+. The ATPase activity levels, expressed per milligram of
protein, of H+,K+-ATPase,
Na+,K+-ATPase, HNN6, and HN16 at 100 µM ATP were rather similar.

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Fig. 2.
ATPase activity of chimeras and wild-type
enzymes. The assay was performed at 37 °C in the presence of
0.2 mM EDTA, 0.1 mM EGTA, 1.3 mM
MgCl2, 50 mM Tris-acetic acid (pH 7.0), and 100 µM ATP. Depending on the conditions described in the
figure, 10 mM KCl and/or 100 mM NaCl was
included in the incubation medium. The ATPase activity determined was
corrected for that of mock-infected cells. The values presented are the
mean ± S.E. of four enzyme preparations.
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We compared the Na+ dependence of the overall ATPase
activity of Na+,K+-ATPase,
H+,K+-ATPase, and the chimeras HNN6 and HN16 in
the presence of 10 mM K+ at 100 µM ATP (Fig. 3). The
H+,K+-ATPase activity was not stimulated by the
addition of Na+. Increasing concentrations of
Na+, however, gradually inhibited the activity of this
enzyme (apparent IC50 = 66 mM).
Na+,K+-ATPase activity was stimulated by the
addition of Na+ (apparent K0.5 = 4.7 mM). Na+ also raised the
K+-stimulated ATPase activity of the chimeras HNN6 and HN16
with similar apparent K0.5 values (8.5 and 6.1 mM, respectively). At very high Na+
concentrations the K+-stimulated ATPase activities of
Na+,K+-ATPase and the chimeras HN16 and HNN6
were also inhibited. Thus the Na+ activation curves of HN16
and HNN6 are rather similar to that of
Na+,K+-ATPase.

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Fig. 3.
Effect of Na+ on the ATPase
activity of chimeras and wild-type enzymes. The assay was
performed at 37 °C in the presence of 0.2 mM EDTA, 0.1 mM EGTA, 1.3 mM MgCl2, 10 mM KCl, 50 mM Tris-acetic acid (pH 7.0), 100 µM ATP, and different concentrations of NaCl. The ATPase
activity determined was corrected for that of mock-infected cells. ,
H+,K+-ATPase; ,
Na+,K+-ATPase; , HNN6; , HN16. The values
presented are the mean of three experiments. The maximal ATPase
activity was set at 100%.
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To compare the K+ activation characteristics of the two
ATPases and the active chimeras, ATPase activity measurements were carried out with 100 µM ATP using 30 mM
Na+, which is almost optimal for
Na+,K+-ATPase, whereas the
H+,K+-ATPase activity is still 70% of the
maximal activity. All ATPases showed a biphasic K+
activation curve (Fig. 4). The increasing
parts of these curves are due to K+ activation of the
dephosphorylation step. The decreasing parts are due to inhibition by
K+ of the E2 to
E1 transition. The K+ inhibition of
Na+,K+-ATPase is enhanced by using lower than
optimal ATP concentrations. In the absence of K+ and the
presence of 30 mM Na+, the
H+,K+-ATPase activity was already 41% of the
maximal ATPase activity, which is likely to be due to a
K+-like effect of Na+ on the dephosphorylation
reaction (29). The apparent K0.5 value of the
increasing part of this curve was 0.2 mM K+.
The K+-activated ATPase curve of
Na+,K+-ATPase (apparent
K0.5 = 0.4 mM) was shifted to the
right compared with the curve of H+,K+-ATPase.
The apparent K+ sensitivities of the chimeras HNN6 and HN16
were considerably lower (1.3 and 1.6 mM, respectively).

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Fig. 4.
Effect of K+ on the ATPase
activity of chimeras and wild-type enzymes. The assay was
performed at 37 °C in the presence of 0.2 mM EDTA, 0.1 mM EGTA, 1.3 mM MgCl2, 30 mM NaCl, 50 mM Tris-acetic acid (pH 7.0), 100 µM ATP, and different concentrations of added KCl. The
ATPase activity determined was corrected for that of mock-infected
cells. , H+,K+-ATPase; ,
Na+,K+-ATPase; , HNN6; , HN16. The values
presented are the mean of three experiments. The maximal ATPase
activity was set at 100%.
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To determine whether the ATP affinity of these chimeras was changed
compared with that of the wild-type enzymes, we measured the ATPase
activity at 10 mM K+, 30 mM
Na+, and varying concentrations of ATP (1-3,000
µM) (Fig. 5). The apparent
ATP affinity of H+,K+-ATPase (38 µM) was higher than that of
Na+,K+-ATPase (113 µM), and the
affinities of the chimeras HNN6 and HN16 were even higher (13 and 8 µM, respectively). Since it is known that ATP drives the
enzyme from the E2K to the
E1 conformation, this suggests that the
E2K conformers of HNN6 and HN16 are less stable
than those of Na+,K+-ATPase. This was confirmed
by studies with vanadate. This compound reacts with the
E2K conformation of the enzyme and forms a
stable intermediate that inhibits enzyme activity. The longer the
enzyme is in this conformation during the reaction cycle the lower the vanadate concentration needed for 50% inhibition (30). Fig. 6 shows that the chimeras, which have a
high apparent affinity for ATP, have a very low affinity for vanadate,
suggesting that the observed high ATP affinity is due to a preference
of the chimeras HNN6 and HN16 for the E1
conformation.

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Fig. 5.
Effect of ATP on the ATPase activity of
chimeras and wild-type enzymes. The assay was performed at
37 °C in the presence of 0.2 mM EDTA, 0.1 mM
EGTA, 1.3 mM MgCl2, 10 mM KCl, 30 mM NaCl, 50 mM Tris-acetic acid (pH 7.0), and
different concentrations of ATP. The ATPase activity determined was
corrected for that of mock-infected cells, and the maximal ATPase
activity was set at 100%. , H+,K+-ATPase;
, Na+,K+-ATPase; , HNN6; , HN16. The
values presented are the mean of three experiments.
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Fig. 6.
Effect of vanadate on the ATPase activity of
chimeras and wild-type enzymes. The assay was performed at
37 °C in the presence of 0.2 mM EDTA, 0.1 mM
EGTA, 1.3 mM MgCl2, 10 mM KCl, 30 mM NaCl, 50 mM Tris-acetic acid (pH 7.0), 100 µM ATP, and different concentrations of vanadate. The
ATPase activity determined was corrected for that of mock-infected
cells. , H+,K+-ATPase; ,
Na+,K+-ATPase; , HNN6; , HN16. The values
presented are the mean of three experiments. The ATPase activity in the
absence of vanadate was set at 100%.
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A characteristic property of P-type ATPases is the formation of an
acid-stable phosphorylated intermediate during the catalytic cycle. We
measured the phosphorylation capacity of the wild-type enzymes and the
chimeras with 0.1 µM ATP in the absence of
K+, with and without 100 mM Na+ at
21 °C and pH 6.0 (Fig. 7). The ATP
concentration used was approximately four and eight times the
K0.5 for ATP in the phosphorylation reaction of
H+,K+-ATPase and
Na+,K+-ATPase, respectively (15). The chimeras
HN56 and HNn6 that showed no K+-stimulated ATPase reaction
could be phosphorylated. The chimera HNn6 and
H+,K+-ATPase were both phosphorylated to a
level of 6.2 pmol of EP mg
1
protein, whereas the phosphorylation level of chimera HN56 was only 2.6 pmol EP mg
1 protein. The addition
of 100 mM Na+ decreased the phosphorylation
level of these enzymes. The amounts of phosphoenzyme of HN46, HN36, and
HN26 were not significantly different from that of the mock-infected
cells. Chimeras HN16 and HNN6 were phosphorylated to a level of about 2 pmol EP mg
1 protein. The addition
of 100 mM Na+ increased this phosphorylation
level to 3.1 and 3.6 pmol EP mg
1
protein, respectively. Phosphorylation of the wild-type
Na+,K+-ATPase did hardly occur in the absence
of added Na+ and was stimulated by this cation up to a
maximal level of 1.6 pmol EP mg
1
protein.

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Fig. 7.
ATP phosphorylation level of chimeras and
wild-type enzymes. Membranes were preincubated for at least 10 min
at 21 °C in the presence of 50 mM Tris-acetic acid (pH
6.0), 1.2 mM MgCl2, 0.2 mM EDTA,
and with or without 100 mM NaCl. After phosphorylation for
10 s at 21 °C with 0.1 µM
[ -32P]ATP, the phosphorylation level was determined
and corrected for that of mock-infected cells. The values presented are
the mean ± S.E. of four enzyme preparations.
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Na+ stimulated the formation of a phosphorylated
intermediate of Na+,K+-ATPase as well as of the
chimeras HNN6 and HN16, whereas it decreased the phosphorylation level
of H+,K+-ATPase and the chimeras HNn6 and HN56.
In Fig. 8 we used different concentrations of Na+ to investigate the cation dependence
of the phosphorylation process. The phosphorylation levels of
H+,K+-ATPase and chimeras HN56 and HNn6 were
dose-dependently decreased by Na+ (Fig.
8A), which is probably due to K+-like effects of
Na+ on the dephosphorylation reaction (29). The
phosphorylation levels of the wild-type
Na+,K+-ATPase and the chimeras HN16 and HNN6
were dose-dependently increased by Na+. The
phosphorylation level of the chimeras HNN6 and HN16 at 20 mM Na+ was almost three times the level
measured in the absence of added Na+ and was higher than
that measured at a Na+ concentration of 100 mM
(Fig. 7). The phosphorylation level of the wild-type
Na+,K+-ATPase was gradually stimulated by
Na+, until it reached a maximum at 100 mM
Na+ (Fig. 8B) (10). The different kinetics of
these activation curves makes comparison difficult. It is likely that
the difference is due to the fact that the chimeras are in the
E1 form, whereas wild-type
Na+,K+-ATPase is in the
E2 conformation. Part of the added
Na+ is therefore needed to drive the latter enzyme in the
E1 form before it can be phosphorylated.

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Fig. 8.
Effect of Na+ on the ATP
phosphorylation level of chimeras and wild-type enzymes. Membranes
were preincubated for at least 10 min at 21 °C in the presence of 50 mM Tris-acetic acid (pH 6.0), 1.2 mM
MgCl2, 0.2 mM EDTA, and different
concentrations of NaCl. After phosphorylation for 10 s at 21 °C
with 0.1 µM [ -32P]ATP, the
phosphorylation level was determined and corrected for that of
mock-infected cells. , H+,K+-ATPase; ,
HN56; , HNn6; , Na+,K+-ATPase; , HNN6;
, HN16. The values are representative of two enzyme
preparations. The maximal phosphorylation level was set at 100%.
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 |
DISCUSSION |
The first high resolution structure of a P-type ATPase
(Ca2+-ATPase) has been elucidated recently (14). This huge
step forward does, however, not answer all questions regarding the
structure-function relationship of P-type ATPases. The cation-binding
residues present in the occlusion pocket might not be fully responsible
for the cation specificity of P-type ATPases. Indeed, many reports
demonstrate that regions outside the proposed occlusion pocket
influence the cation selectivity (31-34). In the present study we shed
some light on the regions involved in the cation specificity of
Na+,K+-ATPase and
H+,K+-ATPase. We demonstrate that only chimeras
containing transmembrane segments one to six and the intervening
regions of the Na+,K+-ATPase
-subunit harbor
the amino acids responsible for the Na+ stimulation of the
ATPase activity in the presence of K+. On the other hand
the C-terminal part of the
-subunit, together with the
-subunit,
most likely determines the difference in K+ de-occlusion
properties between both parental ATPases.
The crystal structure of SERCA1a Ca2+-ATPase revealed all
residues that coordinate the Ca2+ atoms (14). According to
this model, the complete cation binding pocket of
H+,K+-ATPase was replaced by that of
Na+,K+-ATPase when both transmembrane hairpins
M3-M4 and M5-M6 were exchanged. This chimera (HN34/56), however,
possessed a (H+,)K+-ATPase activity that could
not be stimulated by Na+ (19). Therefore, it is not likely
that only these residues are responsible for the Na+
activation of Na+,K+-ATPase. Because of the
postulated role of the M5-M6 hairpin in cation binding (35), we first
prepared a chimeric H+,K+-ATPase in which only
this hairpin was replaced by that of
Na+,K+-ATPase (HN56). We demonstrated that ATP
could phosphorylate this chimera, but K+ did not stimulate
the dephosphorylation reaction (19). In the present study this chimera
was used as the starting enzyme: the part originating from
Na+,K+-ATPase was gradually increased in the
N-terminal direction. Only when the chimera was extended to include the
M1-M2 hairpin of Na+,K+-ATPase was
Na+ activation observed.
Assembly of
- and
-subunits is a crucial step in the formation of
active X+,K+-ATPases. We previously
demonstrated that Na+,K+-ATPase and
H+,K+-ATPase require their own
-subunits for
optimal activity (15). When the
-subunits were exchanged, the enzyme
activity decreased and the apparent K+ affinity of the
(hybrid) ATPases was the highest when the
-subunit originated from
Na+,K+-ATPase. It has been demonstrated that
the binding region for the
-subunit is located in the C-terminal
region of both the Na+,K+-ATPase and
H+,K+-ATPase
-subunits (17, 18, 36, 37).
Hence, an optimal assembly between the chimeric
-subunits and the
H+,K+-ATPase
-subunit was ensured through
the presence of the C-terminal 187 amino acids of the
H+,K+-ATPase
-subunit in all chimeras
described in this paper.
Na+,K+-ATPase and
H+,K+-ATPase need hydrolysis of ATP to
transport cations across the membrane. Only two of the seven chimeras produced possessed this ATP hydrolyzing activity. The formation of an
acid-stable phosphorylated intermediate during the catalytic cycle is a
characteristic property of P-type ATPases.
X+,K+-ATPases react with K+ ions
and are dephosphorylated subsequently (35). When K+ ions
are absent during incubation, the enzymes presumably accumulate in the
phosphorylated state. Four of the chimeras were significantly phosphorylated compared with the background. Chimeras HN56 and HNn6,
however, did not show a K+-stimulated dephosphorylation
reaction (not shown), and they did not possess
K+-stimulated ATPase activity. Moreover, Na+
did not increase the phosphorylation levels of HN56 and HNn6, indicating that these chimeras most likely do not possess the amino
acids that specify the selectivity for Na+. The chimeras
HN46, HN36, and HN26 showed no phosphorylation or ATPase activity at
all. These chimeras contained the complete intracellular domain between
M4 and M5 that, according to the Toyoshima model (14), includes the
nucleotide binding (N) and phosphorylation (P) domains of
Na+,K+-ATPase. In these chimeras either the
complete A-domain (HN46, HN36) or part of it (HN26) originated from
H+,K+-ATPase. It seems likely that certain
structural changes within the ATPases can impede the enzyme activity.
In that sense, it is remarkable that chimera HN34/56 that has similar
transmembrane segments as HN36, but has intracellular domains that only
originate from H+,K+-ATPase, has
K+-stimulated ATPase activity (19).
When chimera HN26 was extended in the N-terminal direction with
transmembrane segments one and two of
Na+,K+-ATPase, Na+-stimulated
K+-ATPase activity became apparent. This activity was
independent of the origin of the N-terminal intracellular part. Both
chimeras (HN16 and HNN6) also possessed Na+-stimulated
phosphorylation capacity. In addition, (i) these chimeras were already
partially phosphorylated in the absence of Na+, and (ii)
their apparent affinity for Na+ in the phosphorylation
process was higher than that of the wild-type enzyme. Both observations
probably reflect a high preference of these chimeras for the
E1 conformation. This effect has been observed previously (10) for the Na+,K+-ATPase mutant
D804A. It might be that the presence of the chimera in the
E1 conformation is sufficient for activating the
phosphorylation process and thus initiates the ATPase reaction.
The finding that the chimeras HN16 and HNN6 are
Na+-sensitive suggests that the first transmembrane hairpin
is involved in Na+ selectivity. It is tempting to speculate
how this fits with the Toyoshima model for Ca2+-ATPase (14)
that shows that only amino acids from M4, M5, M6, and M8 are directly
involved in Ca2+ binding. It is known that, whereas
Ca2+-ATPase binds and transports two Ca2+ ions,
Na+,K+-ATPase transports three Na+
ions, and binding of all three ions is needed for ATPase activity. It
could theoretically be that one of the Na+ ions binds to a
region of the protein that includes the M1-M2 hairpin. Alternatively,
the interaction of the M1-M2 hairpin with the M3-M4 and M5-M6 hairpins
through the hydrogen bond network might be necessary to obtain a
functional Na+-binding pocket.
In the past decade, it has been demonstrated that the predicted
transmembrane segments 4-6 are involved in cation occlusion (35).
Recently, Mense et al. (34) observed that when three residues present in the fourth transmembrane segment of
Na+,K+-ATPase were replaced by those of
H+,K+-ATPase, the enzyme showed partial
K+-stimulated ATPase activity in the absence of
Na+. The authors suggested that these three residues are
involved in the Na+/H+ selectivity of
Na+,K+-ATPase. We, however, witnessed that the
K+-stimulated ATPase activity of a chimera containing
transmembrane hairpin M3-M4 of Na+,K+-ATPase
(HN34) was not stimulated by Na+ (19). Canfield and
Levenson (32) replaced parts of the rat
1-subunit of
Na+,K+-ATPase by those of rat gastric
H+,K+-ATPase. These constructs were transfected
into ouabain-sensitive human HEK 293 cells. By measuring the ability to
transfer ouabain resistance, they demonstrated that four discrete
regions of Na+,K+-ATPase could not be exchanged
by H+,K+-ATPase without loss of
function. These regions are Leu63-Ile117,
Ala320-Lys413,
Val736-Gln861, and
Val898-Ile953. Blostein et al. (33)
suggested that both the N-terminal half of the intracellular M4-M5 loop
and the adjacent transmembrane helice(s) of
Na+,K+-ATPase and
H+,K+-ATPase play a role in cation selectivity.
Chimera HN16 includes all these fragments, except
Val898-Ile953. This region, however, contains
the
-subunit binding domain, and in contrast to our study, Canfield
and Levenson (32) used the
-subunit of
Na+,K+-ATPase.
The E2K to E1K reaction
of H+,K+-ATPase is more than 2 orders of
magnitude faster than that of the Na+,K+-ATPase
transition (6, 38). Moreover, the E2K conformer
of the Na+,K+-ATPase is 3 orders of magnitude
more stable than E1K, while the E1K and E2K conformations
of the H+,K+-ATPase are energetically nearly
equivalent (6, 38). This explains the difficulty of measuring
K+ occlusion in H+,K+-ATPase (5).
Chimeras HNN6 and HN16 have a high apparent affinity for ATP and a low
affinity for vanadate compared with
Na+,K+-ATPase. This is attributable to the
relative rates of interconversion of the
E1/E2 enzyme
conformations, which yield an equilibrium that is probably more close
to that of H+,K+-ATPase than to that of
Na+,K+-ATPase. Consequently, the chimeras HNN6
and HN16 most likely possess the K+ de-occlusion properties
of the H+,K+-enzyme and not those of
Na+,K+-ATPase. This phenomenon seems to be more
prominent in HN16 than in HNN6. The N terminus, however, is physically
on the other side of the molecule (according to the
Ca2+-ATPase structure (14)) and therefore probably changes
the relative rates of the interconversion of the
E1/E2 conformation by
itself. The different nonsaturating ATP concentrations for
Na+,K+-ATPase and chimeras HNN6 and HN16
probably resulted in different apparent K+ affinities (Fig.
4). When the ATP concentration used is far below its saturating
concentration (Na+,K+-ATPase), then this
results in an apparently higher affinity for K+.
Several studies imply an important role for the M5-M6 hairpin in
occlusion of cations and suggest that this hairpin is stabilized by
interactions with other fragments, such as the M7-M8 hairpin (39-42).
The M7-M8 loop is most likely involved in interactions with the
-subunit (18, 43). Disruption of the
-subunit, by S-S bridge
reduction results in a loss of ATPase activity (44, 45), which has been
shown for Na+,K+-ATPase to lead to a loss of
K+ occlusion (45). Furthermore, it was demonstrated that in
hybrid X+,K+-ATPases, in which the
-subunits
were exchanged, the K+ affinity was modified (15). Ishii
et al. (46) showed that the C-terminal 102 amino acids of
Na+,K+-ATPase are sufficient to shift the
K+ sensitivity for activation of the
Ca2+-ATPase. Geering (47) recently speculated that the
K+-transport function, common to all oligomeric
P2-type ATPases, necessitates a particular amino acid
composition of the C-terminal transmembrane pairs. This specific
arrangement is not compatible with membrane insertion mediated only by
intramolecular interactions and has required, during evolution, the
association of a helper protein that assists the correct packing of
K+-transporting P-type ATPases. Or et al. (48)
previously suggested that the M7-M8 loop interacts with M4-M6
containing the cation sites so that disruption of the
-
interaction alters the disposition of this loop and inactivates cation
occlusion. Our results strengthen this hypothesis, that is we provide
evidence that the C-terminal regions of
Na+,K+-ATPase and
H+,K+-ATPase are likely to be involved in
K+ de-occlusion.
In this study we revealed that the
Na+,K+-ATPase section M1-M6, with a surprising
role for the first transmembrane hairpin, is involved in
Na+ activation. On the other hand the C-terminal 187 amino
acids may play a role in K+ occlusion. Moreover, HNN6 and
HN16 are not only chimeric enzymes in structure, but also in function.
On one hand they possess the Na+-stimulated ATPase reaction
of Na+,K+-ATPase, while on the other hand they
have the K+ de-occlusion properties of
H+,K+-ATPase.
 |
ACKNOWLEDGEMENTS |
We are thankful to Dr. K. Fendler for useful
discussions. We also thank Drs. M. Caplan, J. Forte, W. J. Ball,
Jr., and J. V. Møller for generously providing the various
antibodies. Furthermore, we thank Drs. G. E. Shull and J. B
Lingrel who provided us with the rat cDNA clones of the
H+,K+-ATPase
- and
-subunits and the rat
and sheep cDNA clones of the Na+,K+-ATPase
1- and
1-subunits, respectively.
 |
FOOTNOTES |
*
This work was supported by the Netherlands Foundation for
Scientific Research (NWO-ALW) through Grant 805-05.041.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. Tel.: 31-24-3614260;
Fax: 31-24-3616413; E-mail: j.depont@bioch.kun.nl.
Published, JBC Papers in Press, January 16, 2001, DOI 10.1074/jbc.M010804200
 |
ABBREVIATIONS |
The abbreviations used are:
M, transmembrane segment;
X+, Na+ or
H+;
Sf, Spodoptera frugiperda.
 |
REFERENCES |
1.
|
Pedersen, P. L.,
and Carafoli, E.
(1987)
Trends Biochem. Sci.
12,
146-150[CrossRef]
|
2.
|
Albers, R. W.
(1967)
Annu. Rev. Biochem.
36,
727-756
|
3.
|
Post, R. L.,
Kume, S.,
Tobin, T.,
Orcutt, B.,
and Sen, A. K.
(1969)
J. Gen. Physiol.
54,
306s-326s
|
4.
|
Beaugé, L. A.,
and Glynn, I. M.
(1979)
Nature
280,
510-512[Medline]
[Order article via Infotrieve]
|
5.
|
Rabon, E. C.,
Smillie, K.,
Seru, V.,
and Rabon, R.
(1993)
J. Biol. Chem.
268,
8012-8018[Abstract/Free Full Text]
|
6.
|
Faller, L. D.,
Diaz, R. A.,
Scheiner-Bobis, G.,
and Farley, R. A.
(1991)
Biochemistry
30,
3503-3510[Medline]
[Order article via Infotrieve]
|
7.
|
Pedersen, P. A.,
Nielsen, J. M.,
Rasmussen, J. H.,
and Jørgensen, P. L.
(1998)
Biochemistry
37,
17818-17827[CrossRef][Medline]
[Order article via Infotrieve]
|
8.
|
Price, E. M.,
Rice, D. A.,
and Lingrel, J. B
(1989)
J. Biol. Chem.
264,
21902-21906[Abstract/Free Full Text]
|
9.
|
Kuntzweiler, T. A.,
Arguello, J. M.,
and Lingrel, J. B
(1996)
J. Biol. Chem.
271,
29682-29687[Abstract/Free Full Text]
|
10.
|
Koenderink, J. B.,
Swarts, H. G. P.,
Hermsen, H. P. H.,
Willems, P. H. G. M.,
and De Pont, J. J. H. H. M.
(2000)
Biochemistry
39,
9959-9966[CrossRef][Medline]
[Order article via Infotrieve]
|
11.
|
Hermsen, H. P. H.,
Swarts, H. G. P.,
Koenderink, J. B.,
and De Pont, J. J. H. H. M.
(1998)
Biochem. J.
331,
465-472[Medline]
[Order article via Infotrieve]
|
12.
|
Swarts, H. G. P.,
Klaassen, C. H. W.,
De Boer, M.,
Fransen, J. A. M.,
and De Pont, J. J. H. H. M.
(1996)
J. Biol. Chem.
271,
29764-29772[Abstract/Free Full Text]
|
13.
|
Asano, S.,
Furumoto, R.,
Tega, Y.,
Matsuda, S.,
and Takeguchi, N.
(2000)
J. Biochem. (Tokyo)
127,
993-1000[Abstract]
|
14.
|
Toyoshima, C.,
Nakasako, M.,
Nomura, H.,
and Ogawa, H.
(2000)
Nature
405,
647-651[CrossRef][Medline]
[Order article via Infotrieve]
|
15.
|
Koenderink, J. B.,
Swarts, H. G. P.,
Hermsen, H. P. H.,
and De Pont, J. J. H. H. M.
(1999)
J. Biol. Chem.
274,
11604-11610[Abstract/Free Full Text]
|
16.
|
Geering, K.
(1991)
FEBS Lett.
285,
189-193[CrossRef][Medline]
[Order article via Infotrieve]
|
17.
|
Wang, S. G.,
Eakle, K. A.,
Levenson, R.,
and Farley, R. A.
(1997)
Am. J. Physiol.
272,
C923-C930[Abstract/Free Full Text]
|
18.
|
Melle-Milovanovic, D.,
Milovanovic, M.,
Nagpal, S.,
Sachs, G.,
and Shin, J. M.
(1998)
J. Biol. Chem.
273,
11075-11081[Abstract/Free Full Text]
|
19.
|
Koenderink, J. B.,
Hermsen, H. P. H.,
Swarts, H. G. P.,
Willems, P. H. G. M.,
and De Pont, J. J. H. H. M.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
11209-11214[Abstract/Free Full Text]
|
20.
|
Luckow, V. A.,
Lee, S. C.,
Barry, G. F.,
and Olins, P. O.
(1993)
J. Virol.
67,
4566-4579[Abstract]
|
21.
|
Klaassen, C. H. W.,
Van Uem, T. J. F.,
De Moel, M. P.,
De Caluwé, G. L. J.,
Swarts, H. G. P.,
and De Pont, J. J. H. H. M.
(1993)
FEBS Lett.
329,
277-282[CrossRef][Medline]
[Order article via Infotrieve]
|
22.
|
Klaassen, C. H. W.,
Swarts, H. G. P.,
and De Pont, J. J. H. H. M.
(1995)
Biochem. Biophys. Res. Commun.
210,
907-913[CrossRef][Medline]
[Order article via Infotrieve]
|
23.
|
Peterson, G. L.
(1983)
Methods Enzymol.
91,
95-106[Medline]
[Order article via Infotrieve]
|
24.
|
Gottardi, C. J.,
and Caplan, M. J.
(1993)
J. Biol. Chem.
268,
14342-14347[Abstract/Free Full Text]
|
25.
|
Gottardi, C. J.,
and Caplan, M. J.
(1993)
J. Cell Biol.
121,
283-293[Abstract]
|
26.
|
Ning, G.,
Maunsbach, A. B.,
Lee, Y. J.,
and Møller, J. V.
(1993)
FEBS Lett.
336,
521-524[CrossRef][Medline]
[Order article via Infotrieve]
|
27.
|
Chow, D. C.,
and Forte, J. G.
(1993)
Am. J. Physiol.
265,
C1562-C1570[Abstract/Free Full Text]
|
28.
|
Peters, W. H. M.,
Ederveen, A. G. H.,
Salden, M. H. L.,
De Pont, J. J. H. H. M.,
and Bonting, S. L.
(1984)
J. Bioenerg. Biomembr.
16,
223-231[Medline]
[Order article via Infotrieve]
|
29.
|
Swarts, H. G. P.,
Klaassen, C. H. W.,
Schuurmans Stekhoven, F. M. A. H.,
and De Pont, J. J. H. H. M.
(1995)
J. Biol. Chem.
270,
7890-7895[Abstract/Free Full Text]
|
30.
|
Cantley, L. C.,
Cantley, L. G.,
and Josephson, L.
(1978)
J. Biol. Chem.
253,
7361-7368[Medline]
[Order article via Infotrieve]
|
31.
|
Schneider, H.,
and Scheiner-Bobis, G.
(1997)
J. Biol. Chem.
272,
16158-16165[Abstract/Free Full Text]
|
32.
|
Canfield, V. A.,
and Levenson, R.
(1998)
Biochemistry
37,
7509-7516[CrossRef][Medline]
[Order article via Infotrieve]
|
33.
|
Blostein, R.,
Dunbar, L.,
Mense, M.,
Scanzano, R.,
Wilczynska, A.,
and Caplan, M. J.
(1999)
J. Biol. Chem.
274,
18374-18381[Abstract/Free Full Text]
|
34.
|
Mense, M.,
Dunbar, L. A.,
Blostein, R.,
and Caplan, M. J.
(2000)
J. Biol. Chem.
275,
1749-1756[Abstract/Free Full Text]
|
35.
|
Møller, J. V.,
Juul, B.,
and Le Maire, M.
(1996)
Biochim. Biophys. Acta
1286,
1-51[Medline]
[Order article via Infotrieve]
|
36.
|
Lemas, M. V.,
Takeyasu, K.,
and Fambrough, D. M.
(1992)
J. Biol. Chem.
267,
20987-20991[Abstract/Free Full Text]
|
37.
|
Colonna, T. E.,
Huynh, L.,
and Fambrough, D. M.
(1997)
J. Biol. Chem.
272,
12366-12372[Abstract/Free Full Text]
|
38.
|
Rabon, E. C.,
Bassilian, S.,
Sachs, G.,
and Karlish, S. J. D.
(1990)
J. Biol. Chem.
265,
19594-19599[Abstract/Free Full Text]
|
39.
|
Lutsenko, S.,
Anderko, R.,
and Kaplan, J. H.
(1995)
Proc. Natl. Acad. Sci. U. S. A.
92,
7936-7940[Abstract]
|
40.
|
Lutsenko, S.,
Daoud, S.,
and Kaplan, J. H.
(1997)
J. Biol. Chem.
272,
5249-5255[Abstract/Free Full Text]
|
41.
|
Shainskaya, A.,
Nesaty, V.,
and Karlish, S. J. D.
(1998)
J. Biol. Chem.
273,
7311-7319[Abstract/Free Full Text]
|
42.
|
Shainskaya, A.,
Schneeberger, A.,
Apell, H. J.,
and Karlish, S. J.
(2000)
J. Biol. Chem.
275,
2019-2028[Abstract/Free Full Text]
|
43.
|
Lemas, M. V.,
Hamrick, M.,
Takeyasu, K.,
and Fambrough, D. M.
(1994)
J. Biol. Chem.
269,
8255-8259[Abstract/Free Full Text]
|
44.
|
Chow, D. C.,
Browning, C. M.,
and Forte, J. G.
(1992)
Am. J. Physiol.
263,
C39-C46[Abstract/Free Full Text]
|
45.
|
Lutsenko, S.,
and Kaplan, J. H.
(1993)
Biochemistry
32,
6737-6743[Medline]
[Order article via Infotrieve]
|
46.
|
Ishii, T.,
Hata, F.,
Lemas, M. V.,
Fambrough, D. M.,
and Takeyasu, K.
(1997)
Biochemistry
36,
442-451[CrossRef][Medline]
[Order article via Infotrieve]
|
47.
|
Geering, K.
(2000)
J. Membr. Biol.
174,
181-190[CrossRef][Medline]
[Order article via Infotrieve]
|
48.
|
Or, E.,
Goldshleger, R.,
Shainskaya, A.,
and Karlish, S. J. D.
(1998)
Biochemistry
37,
8197-8207[CrossRef][Medline]
[Order article via Infotrieve]
|
Copyright © 2001 by The American Society for Biochemistry and Molecular Biology, Inc.
Copyright © 2001 by the American Society for Biochemistry and Molecular Biology.