From the Departments of Physiology and Medicine and the Cardiovascular Research Laboratories, UCLA School of Medicine, Los Angeles, California 90095-1760
Received for publication, June 25, 2000, and in revised form, September 19, 2000
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In a revised topological model of the cardiac
Na+-Ca2+ exchanger, there are nine
transmembrane segments (TMSs) and two possible re-entrant loops
(Nicoll, D. A., Ottolia, M., Lu, Y., Lu, L., and Philipson,
K. D. (1999) J. Biol. Chem. 274, 910-917;
Iwamoto, T., Nakamura, T. Y., Pan, Y., Uehara, A., Imanaga, I.,
and Shigekawa, M. (1999) FEBS Lett. 446, 264-268). The
TMSs form two clusters separated by a large intracellular loop between
TMS5 and TMS6. We have combined cysteine mutagenesis and oxidative
cross-linking to study proximity relationships of TMSs in the
exchanger. Pairs of cysteines were reintroduced into a cysteine-less
exchanger, one in a TMS in the NH2-terminal cluster (TMSs
1-5) and the other in a TMS in the COOH-terminal cluster (TMSs 6-9).
The mutant exchanger proteins were expressed in HEK293 cells, and
disulfide bond formation between introduced cysteines was analyzed by
gel mobility shifts. Western blots showed that S117C/V804C,
A122C/Y892C, A151C/T815C, and A151C/A821C mutant proteins
migrated at 120 kDa under reducing conditions and displayed a partial
mobility shift to 160 kDa under nonreducing conditions. This shift
indicates the formation of a disulfide bond between these paired
cysteine residues. Copper phenanthroline and the cross-linker
N',N'-o-phenylenedimaleimide enhanced the mobility shift to 160 kDa. Our data suggest that TMS7 is
close to TMS3 near the intracellular side of the membrane and is in the
vicinity of TMS2 near the extracellular surface. Also, TMS2 must adjoin
TMS8. This initial packing model of the exchanger brings two
functionally important domains in the exchanger, the Na+-Ca2+ exchanger proteins are present in
a wide variety of tissues (1-3). These polytopic membrane proteins are
electrogenic transporters that can utilize the Na+
electrochemical gradient to exchange three extracellular
Na+ ions for one intracellular Ca2+ ion (4). As
a Ca2+ efflux mechanism, the primary function of the
exchanger is to maintain Ca2+ homeostasis, especially in
excitable cells where rapid and substantial Ca2+ fluxes are
important in signaling pathways. In heart, the
Na+-Ca2+ exchanger is the dominant
Ca2+ efflux mechanism important in beat-to-beat relaxation
(reviewed in Ref. 5).
Based on a combination of hydropathy analysis, cysteine mutagenesis and
sulfhydryl modification, immunolocalization, and functional measurements, the exchanger is modeled to have nine transmembrane segments (TMSs)1 and two
possible re-entrant loops (see Fig. 1) (6-13). The TMSs form two
clusters separated by a large intracellular loop (loop f) between TMSs
5 and 6. The amino-terminal cluster is comprised of TMSs 1-5 with a
possible re-entrant loop between TMSs 2 and 3, whereas the C-terminal
cluster is comprised of TMSs 6-9 with a possible re-entrant loop
between TMSs 7 and 8 (6, 7). Experimental evidence supports the
extracellular localization of the amino terminus and loops c, e, g, and
i and the intracellular localization of loops b, d, f, h and the
carboxyl terminus (6-13). From deletion experiments, it has been
determined that the large cytoplasmic domain (loop f) is not essential
for ion transport (12).
Inspection of the amino acid sequence of the exchanger also reveals
that there are two homologous regions consisting of residues spanning
portions of TMSs 2-3 and TMS7 plus part of loop h, respectively (shading in Fig. 1). These
regions are designated as the 1 and
2
repeats, close to each other.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1 and
2 repeats and are conserved
among all exchangers (14 and reviewed in Ref. 5). Extensive
site-directed mutagenesis studies show that exchanger activity is
highly sensitive to mutagenesis of residues in the
-repeats. Even
conservative mutations alter or eliminate activity (15). The putative
-helices of the
-repeats (TMSs 2, 3, and 7) are amphipathic, and
the hydrophilic faces of these helices may form a portion of the ion
translocation pathway (15).
View larger version (20K):
[in a new window]
Fig. 1.
Topological model of the
Na+-Ca2+ exchanger. TMSs are represented
by cylinders and are numbered. The -repeat
regions are shaded. All labeled residues within the
transmembrane regions were used in this study. This includes native
cysteines (reintroduced into the cys-less exchanger) and residues
mutated to cysteines. Also shown are Cys-20 and
Cys-792, the two cysteines that form an endogenous disulfide
bond in the wild type protein. The extracellular surface is at the
top. Extramembrane segments are labeled a through
j. CH2O denotes the protein
glycosylation site.
The electrophoretic mobility of the exchanger is different under reducing and nonreducing conditions. The exchanger proteins purified from canine cardiac sarcolemma migrate on SDS-PAGE as two bands with apparent molecular masses of 70 and 120 kDa in the presence of reducing agents (16). The 120-kDa protein species represents the mature protein, whereas the 70-kDa protein is an active proteolytic fragment of variable amount. Under nonreducing conditions, the apparent molecular mass of the 120-kDa protein shifts to 160 kDa (16). It has been shown recently that this mobility shift is due to an intramolecular disulfide bond between the cysteine at position 792 in loop g and the cysteine at either position 14 or 20 near the NH2 terminus of the exchanger (17). Disulfide bond formation apparently induces a significant conformational change in the exchanger protein under SDS-PAGE conditions.
The three-dimensional arrangement of the TMS helices of the
Na+-Ca2+ exchanger is unknown. Understanding
the transport mechanism of the exchanger will require knowledge of the
helix packing. It is not yet possible to crystallize the exchanger
protein because of difficulties in producing a large amount of pure
functional protein as well as difficulty in crystallizing membrane
proteins in general. Alternative approaches to obtain structural
information on membrane proteins are being developed. In this study, we
employed an approach combining cysteine mutagenesis with disulfide
cross-linking (18, 19) to analyze the arrangement of TMSs in the
exchanger. Pairs of cysteines were reintroduced into a cysteine-less
(cys-less) exchanger, and the mutant exchanger proteins were
expressed in HEK cells. Disulfide cross-linking was detected by an
electrophoretic mobility shift assay. Four cross-links have been
identified, which provide initial information on the helix packing of
the exchanger. Our data indicate that the same interface of TMS7 is
close to TMS2 near the extracellular side but is adjacent to TMS3 near the intracellular side of the plasma membrane and that TMS2 adjoins TMS8. This suggests that the functionally important domains, the 1 and
2 repeats, are in the close proximity.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Generation of Mutant Exchangers-- Clone I, a modified version of the cys-less exchanger (6), was used in this study for expression in HEK cells (see "Results"). Single or pairs of cysteines were reintroduced into the cys-less background. Mutations were generated in 300-500-base pair cassettes using the QuickChange site-directed mutagenesis kit (Stratagene) and verified by sequencing. Full-length exchangers were constructed by subcloning the cassettes carrying mutations into the cys-less exchanger.
Expression of Exchanger Proteins in Xenopus Oocytes-- cRNA was prepared using the mMessage mMachine in vitro RNA synthesis kit (Ambion) and injected into Xenopus laevis oocytes. Na+-Ca2+ exchanger activity was measured as Na+ gradient-dependent 45Ca2+ uptake into oocytes as described previously (11).
Na+ Gradient-dependent
45Ca2+ Uptake into Transfected HEK
Cells--
Exchanger cDNAs were subcloned into pcDNA
3.1() vector (Invitrogen) and transfected into HEK293 cells
using LipofectAMINE (Life Technologies, Inc.). 48-72 h
post-transfection, cells were harvested and washed twice with washing
buffer (10 mM MOPS (pH 7.4), 140 mM NaCl).
Cells were then loaded with Na+ by incubation with 10 mM MOPS (pH 7.4), 140 mM NaCl, 1 mM
MgCl2, 0.4 mM ouabain, and 25 µM
nystatin for 10 min at room temperature. Nystatin was removed from the
cells by two washes with washing buffer plus 0.4 mM
ouabain. Uptake was initiated by resuspending the cell pellet in assay
medium: 10 mM MOPS (pH 7.4), 140 mM KCl (or
NaCl as blank), 25 µM CaCl2, 0.4 mM ouabain, and 5 µCi/ml 45Ca2+.
After incubation (typically 1-3 min), the reaction was stopped by
adding 1 ml of ice-cold quenching solution (140 mM KCl, 1 mM EGTA) followed by two additional washes with the
quenching solution. Cell pellets were then dissolved in 1N
NaOH at 80 °C for 20 min. Aliquots of samples were subjected to
scintillation counting and protein assay.
Crude Membrane Vesicle Preparation from Transfected HEK cells and
Cross-linking Procedure--
Transfected cells were washed twice with
washing buffer, resuspended in 20 mM MOPS (pH 7.4), 280 mM NaCl, and homogenized with 10 strokes in a Dounce
homogenizer. After centrifugation at 14,000 × g for 5 min at 4 °C, the pellet was resuspended in washing buffer. The
sample was then passed through a 20-gauge needle 20 times and
centrifuged for 10 min at 4,000 × g at 4 °C to
remove cell debris and nuclei. The supernatant was collected for
cross-linking or stored at 80 °C. Cross-linking was carried out at
20 °C by adding oxidative reagent or thiol-specific homobifunctional cross-linker to the membrane preparation. The final concentrations of
reagents were 3 mM CuSO4, 9 mM
phenanthroline, and 0.5 mM o-PDM or
p-PDM. The reaction was terminated after 20 min by adding
N-ethylmaleimide to a final concentration of 10 mM.
SDS-PAGE and Western Blots--
48-72 h post-transfection, HEK
cells were harvested and washed with washing buffer. Cells were lysed
with radioimmune precipitation buffer (50 mM Tris-HCl (pH
7.4), 150 mM NaCl, 1% sodium deoxycholate, 1% Triton
X-100, 0.1% SDS, 10 mM EDTA) supplemented with 10 mM N-ethylmaleimide. After incubating on ice for
10 min, the samples were centrifuged for 15 min at 11,000 rpm
at 4 °C. Supernatants were subjected to SDS-PAGE. Membrane
vesicles or cross-linked samples were mixed with an equal volume of
radioimmune precipitation buffer (plus 10 mM
N-ethylmaleimide). Electrophoresis was performed on
discontinuous 7% SDS-polyacrylamide gels. For reducing conditions, 2%
-mercaptoethanol was included in the sample buffer. Proteins in
SDS-PAGE gels were transferred to nitrocellulose membrane (Bio-Rad). Blots were probed with exchanger-specific antibodies (C2C12 or R3F1)
(13), and exchanger signals were detected by chemiluminescence (PerkinElmer Life Sciences).
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Expression of the Wild Type and Cys-less Exchangers in HEK
Cells--
We have previously constructed a cys-less exchanger (clone
H) with Na+ gradient-dependent
45Ca2+ uptake activity comparable with that of
the wild type exchanger when expressed in Xenopus oocytes
(6). However, clone H exchanger displayed no activity when expressed in
HEK cells although exchanger protein was synthesized, as revealed by
Western blot (data not shown). To construct a new version of the
cys-less exchanger which would be active in HEK cells, we reintroduced
each of the 15 native cysteines, one at a time, into the H background
and expressed these mutant exchangers in HEK cells. Exchanger activity
was restored to about 30% of the wild type activity when the cysteine
at either position 151 or position 210 was present. In the H exchanger, both cysteines had been mutated to alanine. We tested whether conservative substitution of these cysteines with serine would restore
exchanger activity. Indeed, a cys-less exchanger with a serine at
position 210 (clone I) had 30% of wild type activity when expressed in
HEK cells. Substitution of Cys-151 with serine did not rescue cys-less
exchanger activity. Therefore, we used clone I as the background for
these studies. Fig. 2 compares the activity of clones H and I and the wild type exchanger in transfected cells. It is unknown why the alanine to serine substitution
restored activity.
|
Mobility Shift of Mutant Exchangers with Substituted Cysteine
Pairs--
We have previously identified an intramolecular disulfide
bond between cysteines 20 and 792 in the exchanger that induces a
mobility shift on SDS-PAGE under nonreducing conditions (17). These two
cysteines are located in extracellular segments connecting TMSs in the
NH2-terminal and COOH-terminal TMS clusters, respectively. We reasoned that other cross-links between substituted cysteines in
TMSs in the NH2-terminal cluster (TMSs 1-5) to those in
the COOH-terminal cluster (TMSs 6-9) might also give rise to an
electrophoretic mobility shift. To test this possibility, we
reintroduced the native cysteine residue Cys-151 in TMS3 into the
cys-less background. This cysteine was then paired with a series of
cysteines (native or substituted) in the TMSs in the COOH-terminal
cluster (Table I). The mutant exchangers
carrying cysteine pairs were expressed in HEK cells and analyzed by
Western blot under reducing and nonreducing conditions (Fig.
3). In mutants A151C/T815C and
A151C/A821C, the majority of the protein migrated as the 120-kDa band,
whereas a fraction (10-15% of total exchanger protein) had an
apparent molecular mass of 160 kDa under nonreducing conditions. The
two mutant proteins migrated as a 120-kDa band when the reducing
reagent -mercaptoethanol was present. These results suggest that
Cys-151 can form disulfide bonds with either Cys-815 or Cys-821 to give rise to a 160-kDa species as previously observed with mutants carrying
Cys-20 and Cys-792. This partial disulfide bond formation between
Cys-151 and Cys-815 or Cys-821 also suggests that TMS3 and TMS7 are
within close proximity.
|
|
Screening of Mutant Exchangers for Mobility Shifts-- To further analyze proximities between different TMSs, we constructed a series of cysteine substitutions at sites modeled to be in the TMSs. Mutants carrying a single cysteine substitution in the cys-less background were first assayed for exchanger activity in oocytes using Na+ gradient-dependent 45Ca2+ uptake (Refs. 6, 11, and 15 and data not shown). Only active cysteine substitutions were then paired, and mutant exchangers carrying substituted cysteine pairs were expressed in HEK cells and analyzed for electrophoretic mobility on SDS-PAGE. Cysteine substitutions were tested in each of the TMSs. Pairings were based on the current topological model such that paired cysteines were modeled to reside within the TMSs near the same side of the plasma membrane. Table I summarizes all the cysteine pair mutants that have been tested for activity and mobility shift. Among the 66 cysteine pairs, four active mutants displayed mobility shifts (Table I) and were further analyzed using cross-linking techniques.
Cross-linking of Exchanger Mutants--
The proximity
relationships between selected cysteine residues were further analyzed
in cross-linking experiments using the oxidative reagent CuPhe or the
homobifunctional thiol-specific linkers o-PDM and
p-PDM. CuPhe catalyzes oxidation of adjacent thiol groups to
promote the disulfide bond formation between cysteine residues.
o-PDM and p-PDM are noncleavable, rigid
homobifunctional reagents in which the maleimido groups are coupled to
a benzene ring in the ortho or para position at
fixed distances of 6 or 10 Å, respectively (19). Membrane vesicles
prepared from transfected cells expressing mutant exchanger were
subjected to cross-linking, SDS-PAGE, and immunoblot analysis. For all
cysteine pairs that showed mobility shifts (151/815, 151/821, 117/804,
and 122/892), treatment with CuPhe and o-PDM enhanced the
conversion of the 120-kDa band to the 160-kDa band under nonreducing
conditions (Fig. 4 and data not shown).
Total exchanger signal (120- + 160-kDa bands) was reduced in samples
from mutant A151C/T815C upon treatment with CuPhe or o-PDM
(not shown). Apparently this mutant protein was prone to aggregation in
the presence of cross-linking reagents. In samples treated with
p-PDM, only a small increase of the 160-kDa species was
observed (Fig. 4). The data are quantified in Table II. No 160-kDa band was ever seen with
the cys-less exchanger (Fig. 4). Under reducing conditions, the
presence of -mercaptoethanol converted the 160-kDa band to the
120-kDa band in untreated and CuPhe-treated samples. A fraction of the
160-kDa exchanger protein was resistant to
-mercaptoethanol in
o-PDM- or p-PDM-treated samples (Fig. 4). In some
samples, an additional band with an apparent molecular mass of 140 kDa
was present (Fig. 4). The origin of this band is unknown, and it has
been observed in our previous studies (16, 17). Although the exact
position and intensity of this band vary in different sample
preparations, the band was not affected by the presence or absence of
reducing reagents as also noted previously (17) or by cross-linking
reagents. Taken together, these results indicate that cysteine pairs
within each of the three mutants (A151C/A821C, S117C/V804C,
A122C/Y892C) are close to each other (within a distance of 6 Å).
|
|
In other studies using similar approaches, it has been demonstrated that ligand binding leads to an altered cross-linking pattern in lactose permease and P-glycoprotein (19, 20). We tested whether the presence or absence of Na+ and/or Ca2+, the two ligands for the exchanger, has any effect on cross-linking. Membrane vesicles were prepared under different conditions in which buffers containing Na+ only, Na+ plus Ca2+, K+ only, or K+ plus Ca2+ were used. No significant difference in cross-linking with different vesicle preparations was observed by Western blot analysis (data not shown).
Effect of Cysteine Disulfide Bond Formation on Exchanger
Activity--
Each individual cysteine substitution mutant has been
tested in Xenopus oocytes and showed Na+
gradient-dependent 45Ca2+ uptake
activity. To study the effects of disulfide bond formation on exchanger
activity, HEK cells transfected with exchanger constructs carrying
selected cysteine pair mutants were assayed for exchanger activity. As
shown in Fig. 5, A151C/T815C and
A151C/A821C mutants have an increased exchanger activity, whereas
activity in other mutants maintained the same level as that of the
cys-less exchanger, including mutants S117C/V804C and A122C/Y892C,
which displayed mobility shifts. Perhaps the disulfide bond between
Cys-151 and Cys-815 or Cys-821 facilitates a more active conformation
of the exchanger or enhances the cell surface expression of the
exchanger protein. In contrast, mutant S117C/K909C was inactive (Fig.
5) although Western blot experiments showed that it displayed partial mobility shift under nonreducing conditions (Table I). This inhibition could not be reversed by treatment with -mercaptoethanol.
Disulfide bond formation in mutant S117C/K909C may disrupt trafficking
of exchanger to the plasma membrane or constrain conformational
flexibility for active ion transport. As with the data shown in Fig. 2,
empty vector transfected cells showed no significant
Na+-Ca2+ exchange activity.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
A number of membrane proteins have been reported to have a retarded electrophoretic mobility in SDS-PAGE under nonreducing conditions (20, 21). This is in contrast to what has been observed for many soluble proteins in which preserving intramolecular disulfide bonds during SDS-PAGE generally leads to an increased mobility. It has been further noted that for the mobility shift of membrane proteins to occur in the absence of reducing agents, residues involved in disulfide bond formation or cross-linking must be distantly spaced (20). We have reported that purified cardiac Na+-Ca2+ exchanger proteins migrate differently in SDS-PAGE depending on the redox environment: a 120-kDa band in the presence and a 160-kDa band in the absence of dithiothreitol (16). By systematically removing each of the 15 endogenous cysteine residues, Santacruz-Toloza et al. (17) determined that the mobility shift to 160 kDa under nonreducing conditions is primarily due to the presence of an intramolecular disulfide bond between residues Cys-20 and Cys-792. Residues Cys-20 and Cys-792 are located in extracellular segments connecting TMSs in the NH2-terminal and COOH-terminal clusters, respectively. Between the two clusters is a large intracellular loop (loop f about 550 amino acid residues in length) that is capable of undergoing dramatic conformational changes (22). The conformation of loop f may contribute to the variation in apparent molecular mass of the exchanger. Under reducing conditions, when there is no disulfide bond connecting the two TMS clusters, loop f may form a more compact structure, and the exchanger migrates as a 120-kDa band. When a disulfide bond between the two TMS clusters brings helices into a more tightly packed bundle, loop f may extend further from the membrane. This "parachute" effect may result in the mobility shift to 160 kDa under nonreducing conditions.
By introducing paired cysteine residues back into the cys-less background, one in each half of the protein, we show here that a number of exchanger mutants also displayed a mobility shift under nonreducing conditions (Fig. 3). This implies that spontaneous disulfide bond formation occurs between the introduced paired cysteines. Treatment with CuPhe significantly enhanced the mobility shift. Thus, the mobility shift provides a useful tool to study proximity relationships of transmembrane helices in the exchanger.
We screened a total of 66 exchanger mutants with double cysteines
(Table I). Four pairs induced mobility shifts that were enhanced with
cross-linking reagents (Figs. 3 and 4). Based on these data, we propose
an initial packing model for TMSs 2, 3, 7, and 8 of the exchanger (Fig.
6). Disulfide bond formation between S117C and V804C indicates that TMS7 is in close proximity to TMS2 at
the extracellular side of the membrane. At the intracellular side of
the membrane, TMS7 is in the vicinity of TMS3 as indicated by
cross-linking of A151C/A821C. Furthermore, residues Cys-804 and Cys-821
in TMS7, which are adjacent to TMS2 and TMS3, are located on the same
surface of TMS7 (Fig. 6) according to helical wheel modeling. This
suggests that the same surface of TMS7 interacts with TMS2 at the
extracellular and TMS3 at the intracellular side of the membrane.
Possibly TMS7 tilts in the membrane between TMSs 2 and 3 or has a bend.
A proline residue at position 813 in the center of TMS7 may facilitate
a nonhelical structure and/or a bend. On this surface of TMS7,
mutations S811T (15), S818A (15), and S818C (data not shown) inhibited
exchanger activity when the exchangers were expressed in oocytes.
|
Our model (Fig. 6) places TMS2 between TMSs 7 and 8. This requires a reasonable length for the loop connecting TMSs 7 and 8. Significantly, loop h connecting TMSs 7 and 8 is modeled to be 49 amino acid residues in length and includes a speculative P-loop-like region (Ref. 6 and Fig. 1). Also, there is experimental evidence that loop c connecting TMSs 2 and 3 re-enters the membrane from the extracellular side of the membrane (7). Thus, the two proposed re-entrant loops may be constrained to be in close proximity.
It has been suggested previously that the 1 and
2 repeats,
comprising parts of TMSs 2, 3, and 7 and loop h, form a portion of the
ion translocation pathway (14, 15). Significantly, our packing model
now indicates that the
-repeats are in close proximity. Within the
-repeat regions, we have previously identified mutations in TMSs 2, 3, and 7, which either abolish or lead to a decreased exchanger
activity, including mutations at Ser-109, Ser-110, Glu-113, and Glu-120
(TMS2); Ser-139 and Asn-143 (TMS3); Ser-811, Asp-814, and
Ser-818 (TMS7) (Ref. 15 and Qiu et
al.2). Strikingly, these
residues line the helical surfaces that face one another according to
our initial helix packing model. Therefore, our data provide structural
evidence for the earlier suggestion that hydrophilic faces of
amphipathic TMSs 2, 3, and 7 form a portion of the ion translocation pathway.
Also, our data support recent topology models (6, 7) that suggest that
residues 804 and 821 of TMS7 must be near the extracellular and
intracellular surfaces, respectively. In initial models (23), this
would not be possible as the orientation of TMS7 (TMS8 of earlier
models) would be reversed. This information will be helpful in
designing future experiments to further elucidate helical interactions
within the exchanger. More importantly, accumulation of structural
information will greatly facilitate understanding of the mechanism of
ion translocation.
![]() |
ACKNOWLEDGEMENTS |
---|
We are grateful to Dr. H. R. Kaback for encouraging this project and to Dr. Beate Quednau for commenting on the manuscript.
![]() |
FOOTNOTES |
---|
* This work was supported by National Institutes of Health Research Grant HL49101 (to K. D. P.), by a grant from the American Heart Association, Western States Affiliate (to Z. Q.), and by the Laubisch Fund.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: Cardiovascular
Research Laboratories, MRL 3-645, UCLA School of Medicine, Los Angeles, CA 90095-1760. Tel.:310-825-7679; Fax: 310-206-5777; E-mail:
Kphilipson@mednet.ucla.edu.
Published, JBC Papers in Press, October 16, 2000, DOI 10.1074/jbc.M005571200
2 Z. Qiu, D. A. Nicoll, and K. D. Philipson, unpublished results.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: TMS, transmembrane segment; PAGE, polyacrylamide gel electrophoresis; MOPS, 4-morpholinepropanesulfonic acid; o-PDM, N',N'-o-phenylenedimaleimide; p-PDM, N',N'-p-phenylenedimaleimide; CuPhe, copper phenanthroline.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. |
Kofuji, P.,
Lederer, W.,
and Schulze, D. H.
(1992)
Am. J. Physiol.
263,
C1241-C1249 |
2. |
Lee, S. L., Yu, A.,
and Lytton, J.
(1994)
J. Biol. Chem.
269,
14849-14852 |
3. |
Quednau, B. D.,
Nicoll, D. A.,
and Philipson, K. D.
(1997)
Am. J. Physiol.
272,
C1250-C1261 |
4. |
Reeves, J. P.,
and Hale, C. C.
(1984)
J. Biol. Chem.
259,
7733-7739 |
5. | Philipson, K. D., and Nicoll, D. A. (2000) Annu. Rev. Physiol. 62, 113-133 |
6. |
Nicoll, D. A.,
Ottolia, M.,
Lu, Y.,
Lu, L.,
and Philipson, K. D.
(1999)
J. Biol. Chem.
274,
910-917 |
7. | Iwamoto, T., Nakamura, T. Y., Pan, Y., Uehara, A., Imanaga, I., and Shigekawa, M. (1999) FEBS Lett. 446, 264-268[CrossRef][Medline] [Order article via Infotrieve] |
8. | Hryshko, L. V., Nicoll, D. A., Weiss, J. N., and Philipson, K. D. (1993) Biochim. Biophys. Acta 1151, 35-42[Medline] [Order article via Infotrieve] |
9. | Sahin-Toth, M., Nicoll, D. A., Frank, J. S., Philipson, K. D., and Friedlander, M. (1995) Biochem. Biophys. Res. Commun. 212, 968-974[CrossRef][Medline] [Order article via Infotrieve] |
10. | Cook, O., Low, W., and Rahamimoff, H. (1998) Biochim. Biophys. Acta 22, 40-52 |
11. |
Doering, A. E.,
Nicoll, D. A.,
Lu, Y.,
Lu, L.,
Weiss, J. N.,
and Philipson, K. D.
(1998)
J. Biol. Chem.
273,
778-783 |
12. | Matsuoka, S., Nicoll, D. A., Reilly, R. F., Hilgemann, D. W., and Philipson, K. D. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 3870-3874[Abstract] |
13. |
Porzig, H.,
Li, Z.,
Nicoll, D. A.,
and Philipson, K. D.
(1993)
Am. J. Physiol.
265,
C748-C756 |
14. |
Schwarz, E. M.,
and Benzer, S.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
10249-10254 |
15. |
Nicoll, D. A.,
Hryshko, L. V.,
Matsuoka, S.,
Frank, J. S.,
and Philipson, K. D.
(1996)
J. Biol. Chem.
271,
13385-13391 |
16. | Philipson, K. D., Longoni, S., and Ward, R. (1988) Biochim. Biophys. Acta 945, 298-306[Medline] [Order article via Infotrieve] |
17. |
Santacruz-Toloza, L. K.,
Ottolia, M.,
Nicoll, D. A.,
and Philipson, K. D.
(2000)
J. Biol. Chem.
275,
182-188 |
18. | Pakula, A. A., and Simon, M. I. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 4144-4148[Abstract] |
19. | Wu, J., and Kaback, H. R. (1997) J. Mol. Biol. 270, 285-293[CrossRef][Medline] [Order article via Infotrieve] |
20. |
Loo, T. W.,
and Clarke, D. M.
(2000)
J. Biol. Chem
275,
5253-5256 |
21. |
Rice, W. J.,
Green, N. M.,
and MacLennan, D. H.
(1997)
J. Biol. Chem.
272,
31412-31419 |
22. |
Levitsky, D. O.,
Nicoll, D. A.,
and Philipson, K. D.
(1994)
J. Biol. Chem.
269,
22847-22852 |
23. | Nicoll, D. A., Longoni, S., and Philipson, K. D. (1990) Science 250, 562-656[Medline] [Order article via Infotrieve] |