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 PontDagger

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



    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

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 beta -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.



    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

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 beta -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 right-arrow 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 alpha 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 beta 1-subunits, which are only 30% identical. Assembly of the alpha - and beta -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 alpha -subunits can form functional complexes with both beta -subunits, they have a preference for their own beta -subunit (15). This preference is probably determined by the C-terminal half of the extracellular loop between transmembrane segments seven and eight of the alpha -subunit where the alpha beta 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 beta -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.


    EXPERIMENTAL PROCEDURES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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Expression Constructs-- The rat gastric H+,K+-ATPase alpha - and beta -subunits and the rat Na+,K+-ATPase alpha 1- and beta 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 alpha -subunits. These restriction sites were chosen in the intracellular domain close to putative transmembrane helices. A NarI restriction site was already present in both alpha -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 alpha - and beta -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 [gamma -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 [gamma -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 alpha - and beta -subunits and the rat and sheep cDNA clones of the Na+,K+-ATPase alpha 1- and beta 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. [gamma -32P]ATP (3000 Ci·mmol-1) was purchased from Amersham Pharmacia Biotech (Buckinghamshire, United Kingdom).


    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Seven chimeric constructs were produced after introduction of six unique restriction sites in both the cDNAs of the rat gastric H+,K+-ATPase alpha -subunit and the rat Na+,K+-ATPase alpha 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 alpha -subunits were introduced in the genome of a baculovirus together with the H+,K+-ATPase beta -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 alpha -subunit), HK9 (25) (directed against the N terminus of the H+,K+-ATPase alpha -subunit), L16 (26) (directed against M6 of Na+,K+-ATPase), 2G11 (27) (directed against the H+,K+-ATPase beta -subunit), and G34 (28) (directed against the Na+,K+-ATPase beta -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).

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.

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. black-square, H+,K+-ATPase; open circle , Na+,K+-ATPase; black-triangle, HNN6; down-triangle, HN16. The values presented are the mean of three experiments. The maximal ATPase activity was set at 100%.

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. black-square, H+,K+-ATPase; open circle , Na+,K+-ATPase; black-triangle, HNN6; down-triangle, HN16. The values presented are the mean of three experiments. The maximal ATPase activity was set at 100%.

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%. black-square, H+,K+-ATPase; open circle , Na+,K+-ATPase; black-triangle, HNN6; down-triangle, 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. black-square, H+,K+-ATPase; open circle , Na+,K+-ATPase; black-triangle, HNN6; down-triangle, HN16. The values presented are the mean of three experiments. The ATPase activity in the absence of vanadate was set at 100%.

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 [gamma -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.

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 [gamma -32P]ATP, the phosphorylation level was determined and corrected for that of mock-infected cells. black-square, H+,K+-ATPase; , HN56; diamond , HNn6; open circle , Na+,K+-ATPase; black-triangle, HNN6; down-triangle, HN16. The values are representative of two enzyme preparations. The maximal phosphorylation level was set at 100%.



    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

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 alpha -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 alpha -subunit, together with the beta -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 alpha - and beta -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 beta -subunits for optimal activity (15). When the beta -subunits were exchanged, the enzyme activity decreased and the apparent K+ affinity of the (hybrid) ATPases was the highest when the beta -subunit originated from Na+,K+-ATPase. It has been demonstrated that the binding region for the beta -subunit is located in the C-terminal region of both the Na+,K+-ATPase and H+,K+-ATPase alpha -subunits (17, 18, 36, 37). Hence, an optimal assembly between the chimeric alpha -subunits and the H+,K+-ATPase beta -subunit was ensured through the presence of the C-terminal 187 amino acids of the H+,K+-ATPase alpha -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 alpha 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 beta -subunit binding domain, and in contrast to our study, Canfield and Levenson (32) used the beta -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 beta -subunit (18, 43). Disruption of the beta -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 beta -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 alpha -beta 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 alpha - and beta -subunits and the rat and sheep cDNA clones of the Na+,K+-ATPase alpha 1- and beta 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.

Dagger 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
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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]


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