A Remarkably Stable Phosphorylated Form of
Ca2+-ATPase Prepared from Ca2+-loaded and
Fluorescein Isothiocyanate-labeled Sarcoplasmic Reticulum
Vesicles*
Philippe
Champeil
,
Fernando
Henao§,
Jean-Jacques
Lacapère¶, and
David B.
McIntosh
From the
Unité de Recherche Associée
2096, CNRS et CEA, and the Section de Biophysique des Protéines
et des Membranes, Département de Biologie Cellulaire et
Moléculaire, Commissariat à l'Energie Atomique, Centre
d'Etudes de Saclay, 91191 Gif-sur-Yvette Cedex, France, the
§ Departamento de Bioquímica y Biología
Molecular, Facultad de Ciencias, Universidad de Extremadura, 06080 Badajoz, Spain, ¶ Unité U410, INSERM, Faculté
Xavier Bichat, 16 rue Henri Huchard, 75870 Paris Cedex 18, France, and
the
Department of Chemical Pathology, University of Cape Town
Medical School, Observatory 7925, Cape Town, South Africa
Received for publication, August 2, 2000, and in revised form, October 17, 2000
 |
ABSTRACT |
After the nucleotide binding domain in
sarcoplasmic reticulum Ca2+-ATPase has been
derivatized with fluorescein isothiocyanate at Lys-515, ATPase
phosphorylation in the presence of a calcium gradient, with
Ca2+ on the lumenal side but without Ca2+ on
the cytosolic side, results in the formation of a species that exhibits
exceptionally low probe fluorescence (Pick, U. (1981) FEBS
Lett. 123, 131-136). We show here that, as long as the
free calcium concentration on the cytosolic side is kept in the
nanomolar range, this low fluorescence species is remarkably stable,
even when the calcium gradient is subsequently dissipated by ionophore. This species is a Ca2+-free phosphorylated species. The
kinetics of Ca2+ binding to it indicates that its transport
sites are exposed to the cytosolic side of the membrane and retain a
high affinity for Ca2+. Thus, in the ATPase catalytic
cycle, an intrinsically transient phosphorylated species with transport
sites occupied but not yet occluded must also have been stabilized by
fluorescein isothiocyanate (FITC), possibly mimicking ADP. The low
fluorescence mainly results from a change in FITC absorption. The
Ca2+-free low fluorescence FITC-ATPase species remains
stable after addition of thapsigargin in the absence or presence of
decavanadate, or after solubilization with dodecylmaltoside. The
remarkable stability of this phosphoenzyme species and the changes in
FITC spectroscopic properties are discussed in terms of a putative FITC-mediated link between the nucleotide binding domain and the phosphorylation domain in Ca2+-ATPase, and the possible
formation of a transition state-like conformation with a compact
cytosolic head. These findings might open a path toward structural
characterization of a stable phosphorylated form of
Ca2+-ATPase for the first time, and thus to further
insights into the pump's mechanism.
 |
INTRODUCTION |
The SERCA1a1
Ca2+ pump is a P-type membrane ATPase, whose catalytic
cycle comprises several intrinsically transient auto-phosphorylated forms, the processing of which is tightly coupled to the binding or
dissociation of calcium and hydrogen ions at distant transport sites.
Twenty years ago, Pick and Karlish (1) showed that the use of
fluorescein isothiocyanate (FITC) as a fluorescent covalent label of
Ca2+-ATPase made it possible to monitor conformational
changes of the protein. It was subsequently found that FITC
specifically labels lysine 515 in the ATPase nucleotide binding domain
(2). The fluorescence changes observed upon vanadate or
Ca2+ binding to FITC-ATPase were generally of relatively
small amplitude, but they nevertheless have been widely exploited. Pick
also described the formation under specific conditions of an
FITC-ATPase species with an exceptionally low fluorescence (3).
Surprisingly, however, these latter results were not, to our knowledge,
much exploited or mentioned later.
We report here that this FITC-ATPase species of low fluorescence has
very unusual functional and energetic characteristics; it is a
remarkably stable phosphorylated species, with Ca2+ binding
sites empty, but oriented toward the cytosolic side and of high
affinity. These results are discussed in relation to the putative
mechanism for ion transport by Ca2+-ATPase and in relation
to the environment of FITC in the ATPase nucleotide binding pocket. The
stability of the low fluorescence FITC-ATPase species makes it a good
candidate for helping the long-sought structural characterization of a
phosphorylated form of Ca2+-ATPase, which would provide
significant insight into the conformational flexibility of an ion
transport ATPase during its catalytic cycle.
 |
EXPERIMENTAL PROCEDURES |
In most experiments, the medium contained 100 mM
KCl, 5 mM Mg2+, and 50 mM MOPS-Tris
at pH 7 and 20 °C (buffer A). SR vesicles (prepared as in Ref. 4)
were labeled with FITC (Sigma F 7250) as described previously (5). In
most cases, this incubation was followed by pH neutralization,
centrifugation, and resuspension at 20 mg/ml protein in buffer A to
which 0.25 M sucrose had been added. Certain aliquots of
resuspended labeled vesicles at 20 mg/ml protein were passively loaded
by 1-2 h of equilibration with 5 mM Ca2+ in
buffer A without Mg2+, and frozen without sucrose. Control
vesicles were treated similarly but without FITC.
The procedures used for ordinary fluorescence or stopped-flow
fluorescence measurements, for 45Ca2+ binding
measurements (either at equilibrium or during rapid filtration with
Bio-Logic equipment), and for [32P]EP
measurements (either without acid quenching or after acid quenching),
have already been described (4-8). FITC fluorescence (Spex fluorolog
or PerkinElmer Life Sciences 650-40 instrument) was generally recorded
with excitation and emission wavelengths of 495 and 520 nm,
respectively (2- and 5-nm bandwidths), and plotted as percentage of the
value in the presence of Ca2+ without any correction for
dilution effects (dilution was kept below 1.2% for every addition,
except for that of 10 mM AcP, which resulted in 4%
dilution). The stopped-flow experiments were performed with a Biologic
SFM 3 stopped-flow instrument equipped with a short pathlength optical
cell (1.5-mm cell, FC15), and data points were collected every 2 or 5 ms. The excitation wavelength was 460 nm, and the emission filter was a
broad MTO 531 filter (Massy, France). For the experiment at the final
free concentration of about 30 µM, the low fluorescence
species was first formed by addition of 6 mM EDTA instead
of 2 mM EGTA, and then mixed with 2 mM
Ca2+ plus 4 mM Mg2+. For
phosphorylation experiments performed in the presence of detergent, the
phosphorylation reaction was quenched with 15 mM Pi and 0.5 M perchloric acid (i.e.
4% v/v) instead of our usual 0.12 M perchloric acid, to
precipitate the detergent-solubilized ATPase more easily.
Ca2+ uptake into SR vesicles was measured in different
ways, among which through changes in absorbance of the
calcium-sensitive dye antipyrylazo III (Fluka no. 10795;
e.g. Ref. 9). Absorbance and turbidity were measured either
at a single wavelength or at multiple wavelengths with a diode array HP
8453 spectrophotometer, in a continuously stirred
temperature-controlled cuvette. AcP hydrolysis was deduced from proton
release, using a pH meter (PHM 62, Radiometer) whose analogic output
was amplified (thanks to G. Lecointe in Saclay) to allow acquisition
and digitalization.
Acetylphosphate (AcP) was from Sigma (catalog no. A-0262) and was
freshly prepared at 250 mM. Orthovanadate solutions were prepared freshly as 100 mM alkaline (colorless) solutions
at pH 12, by simple dissolution in water (Sigma S-6508). Decavanadate (yellow) solutions were prepared by titrating and diluting the former
solutions first to pH 2 and then to pH 7 (and 50 mM), or preferably to pH 6 to minimize the presence of other forms of vanadate
(10). The nonionic detergents C12E8 and DM were
obtained from Nikko and Calbiochem (or Anatrace), respectively. Free
Ca2+ concentrations were computed as described previously
(8, 11).
 |
RESULTS |
The Puzzling Properties of an Unusually Stable Low Fluorescence
FITC-ATPase Species Formed from AcP in the Presence of a
Ca2+ Gradient--
Pick (3) reported previously that,
after AcP-dependent Ca2+ uptake by FITC-labeled
SR vesicles and an additional EGTA-induced rise in the Ca2+
gradient across the membrane of these vesicles, an FITC-ATPase species
with an unusually low fluorescence was formed. Trace
A in Fig. 1 shows that this
low fluorescence species can be very stable, much more in fact than
initially reported by Pick. The poorer stability of the low
fluorescence species in Pick's original report appears to be due to
the fact that, in his experiment, EGTA addition did not result in a
free Ca2+ concentration as low as in the experiment
illustrated in our trace A; trace
B in Fig. 1 demonstrates this effect of a higher free
Ca2+. The low fluorescence species was back-converted to a
species of higher but intermediate fluorescence as soon as subsequent addition of Ca2+ raised the external Ca2+, and
the addition of ionophore immediately brought the fluorescence level up
to about 100%. When the ionophore was added before AcP and EGTA, it
completely prevented the appearance of any low fluorescence species
(trace C in Fig. 1). Traces
A--C therefore imply that large drops in the
fluorescence level depend on the presence of a high calcium
concentration on the lumenal side of the vesicles, as concluded
previously by Pick. In agreement with this view, we found that the
half-time for the AcP-induced initial slow drop in fluorescence (in the
absence of any Ca2+-precipitating anion) was similar to
that required for the accumulation of 45Ca2+ in
FITC-labeled SR vesicles (20-30 s, data not shown).

View larger version (26K):
[in this window]
[in a new window]
|
Fig. 1.
The low fluorescence FITC-ATPase species
formed from AcP in the presence of a Ca2+ gradient has
puzzling properties. FITC-labeled SR vesicles were suspended in 2 ml of buffer A, at 20 µg/ml protein. First, as an internal control
for each experiment, 40 µM EGTA followed by 50 µM Ca2+ were added
(upside-down triangles), resulting in
the small well known Ca2+-dependent FITC
fluorescence changes. In the experiment illustrated by trace
A, 10 mM AcP was then added, followed by 2 mM EGTA (resulting in a low free Ca2+
concentration of about 10 nM), 2 mM
Ca2+, and finally 1 µg/ml A23187. In the experiment
illustrated by trace B, 150 µM
Ca2+ was added before AcP (EGTA addition now resulted in a
slightly higher free Ca2+ concentration, about 40 nM). In the experiments illustrated by traces
C and D, similar additions were made, but
ionophore was added either before (C) or after
(D) AcP and EGTA. In the experiment illustrated by
trace E, 6 mM EDTA, followed by 6 mM Mg2+, were added after EGTA; in this case, 1 µg/ml ionomycin was added at the end, instead of A23187
(iono). In the experiment illustrated by trace
F, various aliquots of DM were sequentially added, resulting
in final concentrations of 0.1, 0.2, 0.4, and 1 mg/ml; this experiment
was performed in a medium containing 250 mM NaCl instead of
100 mM KCl, but qualitatively similar results were obtained
in buffer A with C12E8. The first addition of
DM was sufficient to reduce sample turbidity to a minimum. The starting
time for each trace is arbitrary.
|
|
However, much to our surprise, we observed that when the ionophore was
added to the cuvette after the formation of the low fluorescence species, the previous drop in FITC-ATPase fluorescence was
not reversed, at least when the free Ca2+ concentration on
the cytosolic side of the ATPase (i.e. outside the vesicles)
was very low (trace D in Fig. 1). Therefore, the existence of a Ca2+ gradient is absolutely necessary for
the formation of the low fluorescence species, but once this species is
formed, it remains stable even if the gradient collapses.
Trace E in Fig. 1 illustrates another puzzling
feature of this low fluorescence species; although its formation is
known (3) to require Mg2+, a cofactor of
Ca2+-ATPase phosphorylation, once this species was formed,
it no longer required Mg2+ for stability. In fact, it was
even more stable in the absence of Mg2+ than in its
presence, presumably partly due to EDTA-induced Ca2+
chelation and clamping of the free Ca2+ concentration at an
extremely low value. The low fluorescence species also remained stable
after complete detergent-induced solubilization of the vesicles
(trace F), eliminating the possibility that the
resistance to ionophore in panel D could be due
to lack of permeabilization, and revealing an unusual stability for
this detergent-solubilized ATPase in the presence of EGTA. The low fluorescence species remained stable, too, when thapsigargin was added
to it (data not shown,2 but
see Fig. 2 below). Addition of up to 10 mM ATP or ADP, either in the presence or absence of
Mg2+, had no effect either on the low fluorescence species,
including after its formation from partially labeled vesicles (data not shown).

View larger version (27K):
[in this window]
[in a new window]
|
Fig. 2.
The low fluorescence FITC-ATPase species
formed from Pi in the presence of a Ca2+
gradient has properties similar to those of the species formed from
AcP. FITC-labeled SR vesicles that had been passively loaded with
Ca2+ were diluted to 20 µg/ml in buffer A. As an internal
control for each experiment, EGTA was initially added (first
upside-down triangle; its
concentration was 2 mM for traces
A-I). In the experiment illustrated by traces
A-D, Pi was then added at various
concentrations (as indicated), followed by 2 mM
Ca2+ (second upside-down
triangle). In the experiment illustrated by trace
E, 0.25 mM orthovanadate (VO4) was
added instead of Pi. In the other experiments,
Pi was added at 10 mM; in those illustrated by
traces F and G, additions of EGTA,
Ca2+, Pi, and ionophore (2 µM
ionomycin) were made in various orders. In the experiment illustrated
by traces H and I, 1 µg/ml TG, was
added, either after EGTA and before Pi (trace
H), or after formation of the low fluorescence species,
which here had been formed by adding EGTA after Pi
(trace I); in the presence of TG, addition of 2 mM Ca2+ at the end (triangle) was no
longer efficient. In the experiment illustrated by traces
J-N, various concentrations of EGTA were initially added,
resulting in calculated final free Ca2+ concentrations
corresponding to the pCa values indicated.
|
|
The Similar Properties of the Low Fluorescence FITC-ATPase Species
Formed from Inorganic Phosphate in the Presence of a Ca2+
Gradient--
Pick reported that a low fluorescence species could also
be formed after adding both EGTA and Pi (irrespective of
the order) to FITC-labeled vesicles previously loaded passively with
Ca2+, i.e. after phosphorylation from
Pi in the presence of a calcium gradient (3). We fully
confirmed this observation (Fig. 2A). The maximal amplitude
of the observed signal was smaller than that obtained when experiments
were performed with AcP and FITC-labeled vesicles. This smaller
amplitude was presumably due to the fact that some of the vesicles had
lost their impermeability during passive Ca2+ loading (12)
and/or freezing. The Pi-dependence of the formation of the
low fluorescence species revealed a relatively high apparent affinity
for Pi (half-amplitude was obtained for 0.2-0.3
mM Pi, see traces A-D in
Fig. 2), characteristic of gradient-dependent phosphoenzyme
formation (13). Note that orthovanadate (Fig. 2, trace
E) was not able to replace Pi and induced the
same changes with Ca2+-loaded vesicles as those previously
observed with nonloaded vesicles (14).
We found that the low fluorescence species formed from Pi
and Ca2+-loaded vesicles had puzzling properties, too.
Although the formation from Pi of this low fluorescence
species was strictly dependent on lumenal Ca2+ (as deduced
from the fact that preliminary addition of ionophore completely
prevented its appearance), once formed, this low fluorescence species
was again resistant to the subsequent addition of ionophore (Fig. 2,
traces F and G), or even to the
addition of detergent at solubilizing concentrations, as already
described for the species formed from AcP in Fig. 1 (data not shown).
Similarly, although the formation from Pi of this low
fluorescence species was inhibited by thapsigargin, an inhibitor of
Ca2+-dependent changes as well as of
phosphorylation from Pi (15), once formed, this species was
resistant to the addition of thapsigargin (Fig. 2, traces
H and I). Addition of orthovanadate to a
preformed low fluorescence species brought the fluorescence back to a
high level, at a concentration-dependent rate (data not
shown), but decavanadate could bind to this species without
destabilizing it; about the same relative quenching was observed when
decavanadate was added to control FITC-ATPase (16) or to a low
fluorescence TG-stabilized FITC-ATPase species formed from AcP (data
not shown).2
Traces J-N in Fig. 2 show how formation of the
low fluorescence species was influenced by the final free
Ca2+ concentration in the medium. The low fluorescence
species, once formed, was much more stable when the free
Ca2+ concentration dropped to a very low level than when it
was only moderately low, as shown above when the low fluorescence
species was formed after AcP-mediated calcium uptake (cf.
traces A and B in Fig. 1). The
pCa dependence of the amplitude of the
Pi-induced drop revealed a high affinity
(pCa1/2 was about 7, i.e.
Ca1/2 was submicromolar), which in fact was slightly
higher than the overall affinity for Ca2+ binding to SR
ATPase under the same conditions (micromolar Ca1/2; see Ref. 7).
Ca2+ Is Not Bound to the Low Fluorescence FITC-ATPase,
yet This ATPase Species Remains Phosphorylated and Phosphorylation Is
More Stable than without FITC--
Since the existence of a
Ca2+ gradient is required for the initial formation of a
low fluorescence FITC-ATPase species, Pick initially concluded that
this species would contain 2 or at least 1 bound Ca2+
ion(s) (3). If this were the case, as this species, once formed, is
resistant to the subsequent addition of ionophore or detergent, the
putative bound Ca2+ ion(s) should be occluded. We tested
this possibility by loading FITC-labeled SR vesicles with
45Ca2+, creating a low fluorescence species by
adding EGTA, Pi, and then ionophore, and measuring the
amount of 45Ca2+ bound to the ATPase: under
these conditions, however, this residual amount dropped to values
smaller than 0.2 nmol/mg of protein, i.e. to values much
smaller than the ATPase contents in SR vesicles, which typically is
5-7 nmol/mg (Fig. 3A; see
also Ref. 6). The results were similar when the Ca2+-loaded
vesicles were first diluted in EGTA and Pi was added
afterward, or if partially labeled vesicles were used instead of fully
labeled vesicles (data not shown). Therefore, Ca2+ is not
occluded in the low fluorescence FITC-ATPase species; in this ATPase
species, the transport sites are not occupied by Ca2+ at all.

View larger version (32K):
[in this window]
[in a new window]
|
Fig. 3.
The low fluorescence FITC-ATPase species
formed from Pi in the presence of a Ca2+
gradient does not retain Ca2+; the resulting phosphoenzyme
is stabilized by FITC. Panel A, FITC-labeled SR
vesicles (20 mg/ml) were passively loaded with 5 mM
45Ca2+. At time 0, vesicles were diluted 1:20
in buffer A, to which 5 mM EGTA and 10 mM
Pi had been added. After various periods, 40-µl aliquots
were diluted in 4 ml of buffer A, containing either 0.1 mM
Ca2+ (circles) or EGTA and Pi as in
the initial medium (triangles), filtered on an HA Millipore
filter, washed twice with the same medium, and counted
(closed symbols). Alternatively (open
symbols), 0.01 mg/ml ionomycin was added to the vesicles
after 1.5 min, and 40-µl aliquots were again processed after various
periods. Panel B illustrates a control
fluorescence experiment (cf. Fig. 2G).
Panels C and D, at (nominal) time 0 or
10, Ca2+-loaded FITC-labeled vesicles (panel
C) or unlabeled vesicles (panel D) (at
20 mg/ml in a 5 mM Ca2+ medium) were diluted to
1 mg/ml in buffer A supplemented with 1 mM
[32P]Pi and 10 mM EGTA (free
Ca2+ was about 10 nM). Aliquots (40 µl) were
acid-quenched after various periods and filtered (closed
circles). For some of the samples, after 1.5 min we added
one of the following: 0.01 mg/ml ionomycin (open
circles), 8 mM EDTA (squares), 10 µg/ml TG (upside-down triangles), 20 mM cold Pi (rightside-up
triangles), or 10 mM Ca2+
(diamonds). Panels E and F,
same experiment as that illustrated in panels C
and D, but with vesicles made leaky with ionomycin before
phosphorylation (ionophore/protein was 1% w/w). Circles,
control experiments. Squares in panel
E, EDTA was added after 1.5 min.
|
|
Nevertheless, we found that the low fluorescence FITC-ATPase species
was phosphorylated to a high level (Fig. 3C), in fact higher
than that measured for unmodified ATPase under similar conditions (Fig.
3D). This phosphorylation level (close to 4 nmol/mg, i.e. a significant proportion of the above-mentioned ATPase
contents) remained stable, whereas the level of the phosphoenzyme
formed from unmodified vesicles slowly declined with time over minutes, presumably due to the dissipation of the Ca2+ gradient.
Just like the low fluorescence species, the phosphoenzyme formed from
FITC-ATPase and [32P]Pi was resistant to
ionophore-induced (or detergent-induced) collapse of the
Ca2+ gradient, the EDTA-induced removal of
Mg2+, addition of thapsigargin (with or without
decavanadate) and dilution with unlabeled Pi, whereas it
was completely abolished by a rise in the Ca2+
concentration (Fig. 3C and data not shown).2
These properties are at variance with the conventional ones of gradient-dependent phosphoenzyme formed from unmodified
vesicles incubated with [32P]Pi (Fig.
3D), as well as with the properties observed with either FITC-modified or unmodified vesicles previously made leaky to Ca2+ (Fig. 3, E and F).
Ca2+ Binds to the Phosphorylated Low Fluorescence
FITC-ATPase Species from the Cytosolic Side, and Then Gets Transported
into the SR Lumen--
The fact that, in the experiment illustrated in
Fig. 3C, the addition of cold Pi had no effect
on the [32P]EP level of FITC-ATPase implies
that the phosphorylated low fluorescence species is not in
rapid equilibrium with unphosphorylated ATPase and Pi in
the medium. However, when the external Ca2+ concentration
was raised, the low fluorescence FITC-ATPase species was back-converted
to a species with higher fluorescence (Figs. 1 and 2). This implies
that Ca2+ interacts directly with the phosphorylated low
fluorescence species. In view of the fast effect of Ca2+
(Figs. 1, 2, and 3C) compared with its relatively slow
permeation through the membrane (Fig. 3A), this also
suggests that Ca2+ binds to the phosphorylated low
fluorescence species from the external, cytosolic side. This would be
an unconventional conclusion, because phosphoenzyme formation is
generally thought to be associated with the reorientation of
Ca2+ sites toward the lumen, or at least toward the
interior of the membrane for ion occlusion (17, 18). In the next
experiments, we therefore studied in some detail the kinetics of the
events associated with Ca2+ binding to the low fluorescence species.
We initially investigated them by forming a low fluorescence species of
FITC-ATPase (by adding EGTA after AcP-supported Ca2+
uptake) and then monitoring the rate at which subsequent addition of
Ca2+ reversed the previous drop in FITC fluorescence. In
the range of submicromolar Ca2+ concentrations, this
reversal was slow enough to be monitored with a regular fluorometer,
but it accelerated when the free Ca2+ concentration was
raised (Fig. 4A). For
micromolar Ca2+ concentrations, we had to use stopped-flow
detection (Fig. 4B). We found that the rate constant of the
fluorescence rise was about 10 s
1 at 30 µM free Ca2+ (pCa 4.5), and this
was almost the maximal value (the rates measured at 300 µM and 2 mM free Ca2+ were 13 and
15 s
1, respectively). This rate was of the
same order of magnitude as that for Ca2+ binding to
dephosphorylated native ATPase in the absence of nucleotides under
similar conditions (19, 20). The Ca2+ dependence of the
stability of the low fluorescence species was qualitatively similar
after collapsing the calcium gradient by either ionophore or detergent
(data not shown).

View larger version (33K):
[in this window]
[in a new window]
|
Fig. 4.
Ca2+ binding to the low
fluorescence species occurs with fast kinetics and returns the ATPase
to the normal cycle for Ca2+ internalization and
dephosphorylation. Panel A, conventional
fluorescence experiments. The low fluorescence FITC-ATPase species was
formed essentially as in Fig. 1, by sequential addition of 2 mM AcP and 2 mM EGTA (final pCa was
about 8.5, because of the presence of 15 µM
Ca2+). At (nominal) time 0, Ca2+ was then
added, leading to the pCa values indicated. Panel
B, stopped flow fluorescence experiments. The low
fluorescence species, initially formed (at 80 µg protein/ml) by
2.5-min incubation with 2 mM AcP, followed by
Ca2+ chelation, was mixed with various EGTA- or
calcium-containing solutions (final pCa values are
indicated). Panel C, kinetics of
45Ca2+ binding during rapid perfusion.
FITC-ATPase was prepared either in its low fluorescence phosphorylated
state (as shown in panels A and B, but
now at 0.3 mg of protein/ml, triangles), or in its control
nonphosphorylated but Ca2+-deprived state
(circles); control unmodified ATPase was also prepared
(squares). In all cases, 0.3 mg of protein was adsorbed onto
a Millipore HA filter, manually rinsed for a few seconds with 100 µM EGTA, and perfused with 50 µM
45Ca2+ for various periods (see
abscissa). Experiments were repeated in the absence of
membranes for control (diamonds). Panel
D, kinetics of Ca2+-induced dephosphorylation.
32P-labeled phosphoenzyme was prepared as in Fig.
3C. At t = 1.5 min, aliquots were diluted to
2 ml with a solution containing 1 mM EGTA, filtered, rinsed
twice with the same solution and a third time with a solution
containing only 0.1 mM EGTA, and at t = 2 min, the sample was finally perfused for various periods (see
abscissa), with a solution buffered at pCa 4.3 (triangles), pCa 6.4 (circles), or
pCa 8.5 (for control, diamonds). To some of the
samples (closed triangles), 0.01 mg/ml ionomycin
had been added at t = 1.2 min.
|
|
We also directly measured, with rapid filtration equipment at a free
Ca2+ concentration of 50 µM, the kinetics of
45Ca2+ binding to the low fluorescence
phosphorylated species, as well as, in control experiments, the
kinetics of 45Ca2+ binding to either
nonphosphorylated FITC-labeled ATPase or unlabeled SR-ATPase. For short
perfusion periods of up to 1 s, the binding patterns were
essentially similar (Fig. 4C), and biphasic as found previously (21, 22); together with the above stopped-flow fluorescence
data, this confirms that the Ca2+ binding sites on the low
fluorescence species are as accessible from the external compartment of
the vesicles as the binding sites on the control nonphosphorylated
ATPase with or without FITC. However, for longer periods of
45Ca2+ perfusion onto the FITC-ATPase species
of initially low fluorescence, the amount of
45Ca2+ associated with the vesicles appeared to
slowly rise to levels higher than those required for saturation of the
two high affinity binding sites (triangles in Fig.
4C; see below for interpretation).
In the final experiment of this series, we then measured the rate of
Ca2+-induced dephosphorylation of the low fluorescence
species formed from [32P]Pi and
Ca2+-loaded FITC-labeled SR. This rate turned out to be
slower than the rate at which the fluorescence rose and the rate at
which Ca2+ binding took place (open
triangles in Fig. 4D, compare with panels B and C). The slow dephosphorylation was
accelerated (nevertheless remaining slower than Ca2+
binding) when the low fluorescence species was treated with ionophore before Ca2+ was added (closed
triangles in Fig. 4D), while the phosphoenzyme remained stable when no Ca2+ was added, as expected
(diamonds in Fig. 4D). Our interpretation is that
addition of Ca2+ to the Ca2+-free low
fluorescence phosphorylated species allows the ATPase to re-enter the
catalytic cycle and permits ATPase dephosphorylation through the normal
forward pathway, in which dephosphorylation after Ca2+
internalization is slow compared with Ca2+ binding,
especially in the presence of lumenal Ca2+. In
Ca2+-tight vesicles, the internalization of those
45Ca2+ ions that have initially interacted with
the low fluorescence species combines with the passive rebinding of
45Ca2+ (resulting from continuous perfusion) to
newly available dephosphorylated ATPase to explain why bound
45Ca2+ slowly rises to a final level higher
than that required for saturation of the ATPase high affinity binding
sites (as shown by the open triangles in Fig.
4C).
When the Rapid AcP-dependent Turnover of FITC-ATPase
Results in Ca2+ Depletion, a Low Fluorescence Species Forms
Spontaneously; FITC Absorbance Also Changes--
In addition, we found
that the low fluorescence FITC-ATPase species can form spontaneously
after AcP-dependent Ca2+ pumping, before
re-addition of Ca2+ leads to another round of pumping. This
was shown by the fact that in the presence of oxalate and a high enough
concentration of vesicles, i.e. when these vesicles are
capable of depleting the free Ca2+ concentration in the
medium down to submicromolar values, FITC-ATPase was to a large extent
converted spontaneously into a low fluorescence species, after a lag
corresponding to the completion of Ca2+ withdrawal from the
medium (thin top trace in Fig.
5A). Subsequent sequential
Ca2+ additions, which triggered renewed Ca2+
uptake, drove the fluorescence level back toward a higher value until
the added Ca2+ was pumped into the vesicles again. Note
that, in such experiments, the slight increase in turbidity concomitant
with the lumenal precipitation of calcium oxalate (23, 24) may serve as
a marker of the completion of Ca2+ uptake, even in the
absence of any Ca2+-sensitive dye (bottom
thick trace in Fig. 5A). In parallel
experiments, we confirmed that the rates of AcP hydrolysis and
AcP-dependent Ca2+ uptake after formation of
the first microcrystals of calcium oxalate were only marginally slower
for FITC-modified ATPase than for unmodified ATPase (in agreement with
Ref. 5); coupling ratios were also unaltered by FITC modification.
However, FITC-modified ATPase had become completely unable to hydrolyze
ATP, as expected (data not shown).2

View larger version (27K):
[in this window]
[in a new window]
|
Fig. 5.
Spontaneous conversion of FITC-ATPase into a
low fluorescence species, with a different absorption spectrum, after
Ca2+ depletion in the presence of oxalate.
Panel A, buffer A was supplemented with 8 mM oxalate, 40 µM Ca2+, and 0.4 mg/ml SR vesicles that had just been incubated with FITC (2 mg/ml SR
and 16 µM FITC for 60 min). 10 mM AcP was
added to trigger Ca2+ uptake. 100 µM
Ca2+ was subsequently added twice, followed by 1 mM EGTA. FITC fluorescence at this high protein
concentration was recorded (thin bottom
line, plotted after appropriate normalization). In a
parallel measurement, optical densities at 495 nm (top
thick trace) and 545 nm (bottom
trace) were recorded. The 545-nm trace reflects changes in
turbidity mainly due to (initially delayed) precipitation of
Ca2+-oxalate. Panel B, absorption
spectra recorded at various times during such an experiment. As
indicated in panel A, spectra were recorded:
1, in the initial state; 2, after
AcP-dependent withdrawal of Ca2+ from the
medium, and 3, after EGTA addition. At this point, 0.5 mg/ml
C12E8 was added, resulting in
spectrum 4. Readdition of Ca2+ to the
now solubilized sample resulted in spectrum 5 (see Fig. 1F for the related fluorescence recovery).
|
|
Optical density recordings under the exact same conditions allowed us
to conclude that changes in FITC fluorescence were in fact due, at
least in part, to changes in FITC absorbance (thick top trace in Fig. 5A). Using a
diode-array spectrophotometer, it was possible to reveal the changes in
the entire FITC absorption spectrum, as they were large enough to show
up on top of the light scattering by the vesicles (Fig. 5B,
spectra 1-3). These changes were visualized even
more easily after detergent-induced solubilization of the vesicles
(spectra 4 and 5), which was shown
previously (Fig. 1F) to leave the low fluorescence species
stable until Ca2+ was re-added. Similar changes in FITC
absorbance were also seen after Ca2+ depletion in the
presence of 25 mM Pi instead of oxalate, or in
the absence of Ca2+-precipitating anions (data not shown).
FITC is known to have an absorption spectrum (and not only a
fluorescence spectrum) highly sensitive to protonation, polarity, or
interactions (25, 26). FITC spectral absorbance changes for
phosphorylated FITC-ATPase after Ca2+ depletion are
qualitatively similar to those occurring for FITC in either an acidic
or an apolar medium.
Absence of Special Interactions between Chains in the Low
Fluorescence FITC-ATPase--
We asked whether the unusual properties
of the phosphorylated FITC-ATPase species were due to the appearance of
unrecognized ATPase-ATPase interactions. The answer, however, was no:
first, because similar spectroscopic properties were observed with
ATPases that had been labeled with FITC only partially (1 nmol of FITC molecule bound/mg of protein instead of 5-7 nmol/mg) (data not shown);
second, because measurements of the fluorescence polarization of bound
FITC (an index of ATPase-ATPase proximity) showed that most of the
homotransfer-induced depolarization of the bound FITC in FITC-labeled
vesicles was lost upon solubilization at low Ca2+
concentrations, whereas the low fluorescence species remained stable
(data not shown); and third, because size exclusion chromatography experiments indicated that this detergent-solubilized low fluorescence FITC-ATPase species was still essentially monomeric (data not shown).
 |
DISCUSSION |
AcP-dependent Ca2+ uptake followed by
chelation of external Ca2+, as well as phosphorylation from
Pi in the presence of a Ca2+ gradient, both
permit accumulation of a phosphorylated FITC-ATPase species with
unusual properties. The species has an extremely low fluorescein
fluorescence, much lower than any other catalytic intermediate of the
ATPase cycle. Once it has been formed, it no longer depends on the
persistence of a Ca2+ gradient or a membranous state. The
phosphorylated and low fluorescence species has vacant, outwardly
oriented, high affinity Ca2+ binding sites. It is very
stable, as long as the free Ca2+ concentration is kept
close to zero, but re-addition of Ca2+ causes inward
Ca2+ transport followed by dephosphorylation. All these
properties point to a novel species of Ca2+-ATPase, which
appears to be of special interest both for the present functional
description of the mechanism of ion transport and for the future
structural studies of phosphorylated forms of the pump.
Relationship of the Low Fluorescence Species to the Usual Catalytic
Intermediates--
It was suggested that the Ca2+-ATPase
catalytic scheme comprises four major enzyme intermediate species,
namely phosphorylated or nonphosphorylated ATPase with or without bound
Ca2+ (Ref. 27; reviewed in Ref. 28). In terms of a simple
four-species scheme, the Ca2+-free nonphosphorylated form
of ATPase must expose its Ca2+-binding sites toward the
cytosolic side of the SR, whereas the Ca2+-bound
phosphorylated ATPase must expose its Ca2+ binding sites
toward the lumenal side (29). The same rationale is valid for
ATP-supported or AcP-supported activity, since both catalytic cycles
appear to be similar (30-32). In Ca2+-accumulating
vesicles, phosphoenzyme hydrolysis slowing down by the high lumenal
concentration Ca2+ should thus lead to steady-state
accumulation of the Ca2+-bound phosphoenzyme; if excess
EGTA is then added, even more of this phosphoenzyme should form as a
result of the reaction of Pi (derived from AcP hydrolysis)
with residual nonphosphorylated enzyme, by the reverse reaction. The
same Ca2+-bound phosphoenzyme should also be formed from
Pi (again by the reverse reaction) in experiments performed
with passively Ca2+-loaded vesicles like those illustrated
in Fig. 2. However, in the context of such a four-species scheme, a
phosphoenzyme species with outwardly oriented Ca2+ binding
sites cannot be generated upon addition of EGTA to
Ca2+-loaded SR vesicles, and this phosphoenzyme cannot
remain stable after disruption of the initial Ca2+
gradient. Consideration of the more recent hypothesis that during transport, Ca2+ ions move from a first, cytosolically
oriented pair of sites to a second, lumenally oriented pair of sites
(e.g. Ref. 33) does not help much, since in that alternative
view the cytosolically oriented pair of sites is no longer accessible
after ATPase phosphorylation (see Fig. 1 in Ref. 33). In addition, no
evidence for Ca2+ binding to any lumenal site was found in
the atomic structure of Ca2+-ATPase derived from protein
crystallized in the presence of 10 mM Ca2+
(34).
Additional intermediate forms of ATPase have been suggested to
exist within the ATPase catalytic cycle (35-40). As a result, more
elaborate schemes now explicitly include different forms for
Ca2+-bound phosphorylated ATPase (permitting
interconversion between the outside and inside orientations of occupied
Ca2+ sites), different forms for Ca2+-free
nonphosphorylated ATPase (permitting interconversion between the
outside and inside orientations of free Ca2+ sites), and
distinct enzyme forms permitting ion "occlusion" (either for
Ca2+ or for the counter-transported protons) (40, 41); this
is illustrated by the main cycle in Scheme
1. Of course, some of the various
intermediate forms postulated by such schemes may be very transient
during turnover. For instance, with unmodified vesicles, since
dissociation of Ca2+ ions to the outside of the vesicles
was found to become impossible almost concomitantly with ATPase
phosphorylation (17, 18), the early phosphoenzyme form with outwardly
oriented sites in Scheme 1,
Ca2
EP,
must be present in only small amounts. A transient species, however,
can conceivably be made more stable by manipulating experimental
conditions or modifying the enzyme.
We have shown here that when FITC-modified vesicles are suddenly
depleted of external calcium after AcP-dependent loading (Fig. 1), or when Ca2+-loaded FITC-labeled vesicles are
phosphorylated from Pi in the absence of cytosolic
Ca2+ (Fig. 2), a Ca2+-free phosphorylated form
accumulates (Fig. 3), with outwardly oriented (cytosolically oriented)
high affinity sites (Fig. 4). In the context of the above discussion
about the ATPase catalytic scheme, the simplest explanation for these
results is that, in FITC-labeled Ca2+-loaded vesicles, the
early phosphorylated ATPase with outwardly oriented Ca2+
sites,
Ca2
EP-FITC in Scheme 1, is
stabilized and now constitutes a very significant fraction of total
phosphoenzyme, from which Ca2+ can dissociate toward the
cytosolic side. Note that stabilization by FITC of the total level of
EP formed from Pi in the presence of lumenal
Ca2+ is not an interpretation, but a fact (see Fig. 3,
C and D).
Stabilization of phosphorylated ATPase with Ca2+ sites not
yet occluded might be derived from changes in either the forward or the
reverse rate of occlusion in Scheme 1. Since it is known that this
occlusion reaction is not rate-limiting in the normal cycle, such
FITC-dependent alterations would not greatly reduce the
overall AcP-dependent turnover in leaky vesicles or in the presence of oxalate, as actually observed. However, in tight vesicles with high lumenal Ca2+, the
Ca2
EP-FITC species could accumulate, and
at a low enough external free Ca2+ concentration a
phosphorylated species with unoccupied Ca2+ sites facing
the cytosol,
EP-FITC in Scheme 1, could finally be
formed from the previous one. A scheme similar to Scheme 1 (except for
omission of the occluded states) was in fact proposed previously as an
alternative to an overly simple four-species scheme, to explain
AcP-dependent
45Ca2+-40Ca2+ exchange,
an exchange that is slow but nevertheless measurable in normal SR (42).
Note that, even though the early phosphoenzyme form might be very
transient in the absence of FITC, it implies that phosphorylation
actually precedes ion occlusion itself and does not occur
simultaneously with it. Note also that the absence of Mg2+
at the catalytic site of the "E1P" phosphoenzyme has
also been shown to stabilize a phosphoenzyme form with open
Ca2+ sites, thereby permitting Ca2+ release
toward the cytosolic side (43).
Implications for the Catalytic Cycle of Both FITC-Modified and
Unmodified ATPase: A Role for ADP Dissociation in the Occlusion
Process, and Long Distance Coupling?--
To understand the role of
FITC in the stabilization of the early and open
Ca2
EP-FITC form (with intermediate
fluorescence) that precedes ion occlusion, it is worth mentioning that
previous experiments with a photoactivatable analog of ATP
(TNP-8-azido-ATP) revealed that, when this analog was covalently
tethered to the ATPase active site at Lys-492, most of its slow
hydrolysis was uncoupled from Ca2+ transport, again as if
the transport sites had remained open to the cytosolic medium (44).
This did not occur with the free TNP nucleotide, which exhibited tight
coupling of calcium transport and phosphoenzyme hydrolysis. In the
phosphorylated FITC-labeled ATPase, as in the phosphorylated ATPase
with a tethered nucleotide analog, the nucleotide site remains
permanently occupied, perhaps mimicking partly a state in which ADP has
not yet dissociated itself from the unmodified ATPase (45). It might
thus be speculated that, following phosphorylation, ADP dissociation
from the phosphoenzyme plays a significant role for fast closure of the
cytosolic gate of the ion transport site. With unmodified ATPase, from
which ADP dissociates rapidly, the early phosphoenzyme form with
transport sites still open toward the external side of the SR vesicles
would be very transient, while bound FITC or the tethered analog might stabilize this form by mimicking to some extent bound ADP. To our
knowledge, the impact of ADP dissociation on ion dissociation from
other P-type phosphorylated ATPases has not much been studied. The
possibility we suggest should perhaps be kept in mind as an appealing
speculation, although it is fair to say that it is not immediately
reconciled with the fact that the tight binding of Cr.ATP to
unphosphorylated Ca2+-ATPase favors
Ca2+ occlusion (39, 46).
At this point, it is also worth saying a word about the relative
fluorescence level of the two phosphoenzyme forms with outwardly oriented Ca2+ sites,
Ca2
EP-FITC and
EP-FITC.
The latter obviously has a low fluorescence level. The former, if it
indeed accumulates at steady state in the presence of external
Ca2+, must have intermediate fluorescence (Fig. 1). The
difference between the low and intermediate levels of fluorescence (and
absorbance, see Fig. 5) reflects rearrangement of the FITC environment
upon Ca2+ dissociation or binding, and illustrates the long
distance coupling between the transport sites and the catalytic domain.
Differences in fluorescence level have functional counterparts, since
the Ca2+-free
EP-FITC species is
unusually stable and hardly reacts with water, while rapid re-binding
of Ca2+ allows the enzyme to re-enter the cycle for normal
handling of the phosphoenzyme (Fig. 4). The absolute requirement for
Ca2+ at the transport site for normal phosphoenzyme
processing highlights the extraordinary degree to which partial
reactions at the catalytic site can be coupled to changes at distant
transport sites by long range transmission of information (47).
By Which Molecular Mechanism Does Fluorescein Stabilize the
Phosphorylated Form of FITC-ATPase?--
FITC reacts with lysine 515 at the high affinity nucleotide binding site (2, 5, 48), probably as an
affinity label, mimicking nucleotides. Although under our conditions
FITC derivatization increased 3-4-fold the amount of phosphoenzyme
formed from 1 mM Pi in the presence of a
calcium gradient, and made it unusually stable (Fig. 3, C
and D), FITC seems to have only slight effects on most
individual steps of the cycle, including dephosphorylation and
Ca2+ binding (Refs. 5 and 7; see also Fig. 4B);
Ca2+ occlusion can also occur (although with a modified
rate, as discussed above), since Ca2+ is taken up
efficiently in the presence of AcP. Thus, FITC does not sterically
block any essential conformational change. For understanding the effect
of FITC on the stability of the intermediate and low fluorescence
species,
Ca2
EP-FITC and
EP-FITC, respectively, a clue might come from the
spectroscopic changes experienced by the fluorescein moiety under these conditions.
The absorbance (and therefore fluorescence) of fluorescein is
critically dependent on its protonation state and/or the hydrogen bonding power of the environment (25, 26). Fluorescein has two ionizing
groups, the xanthene phenolic group, 3-OH, with a pKa of about 6.7, and the benzoate carboxyl group,
with a pKa of about 4.5. The dianionic species is
strongly absorbant at 495 nm, whereas the monoanion is much less so.
Therefore, one explanation for the unusually low fluorescence and
absorption of the
EP-FITC species is an increase in
the pKa of the phenolic 3-OH, which would stabilize
the protonated form. This could arise either from an increase in
hydrophobicity around this group or from the close approach of a
negatively charged residue. Alternatively the fluorescence may be
quenched by a salt linkage of the negatively charged 3-O
to a positively charged residue. Lys-515, to which the fluorescein moiety is covalently attached in FITC-modified Ca2+-ATPase,
is located deep in the nucleotide binding pocket, while the rest of
this pocket as well as the region surrounding the phosphorylatable
aspartate contains a large number of charged residues (34). Since the
ATPase turnover probably involves large relative movements of the
various subdomains in the ATPase cytosolic head (34), we suggest that
the large drop in FITC fluorescence and absorbance associated with
formation of an early phosphoenzyme form might reflect the fact that,
at this step, the FITC moiety is brought toward the walls of the
cavity, which could easily result in a charged residue being very close
to the 3-O
of the fluorescein. This could result in salt
type bonding if a positive charge approaches. It is also possible that
if a negative charge approaches and results in protonation (change in
pKa) for the fluorescein 3-OH, the hydroxyl group
could now hydrogen-bond to neighboring residues. In both cases this
could be viewed as a form of cross-linkage between subdomains. Of
course, the carboxyl group of the fluorescein may also contribute
interaction energy. The link resulting from these interactions could be
of moderate strength in the
Ca2
EP-FITC
intermediate, but reorganization of the cytosolic head after the
dissociation of Ca2+ could stabilize it further,
resulting in the unusually stable, low fluorescence, and
Ca2+-free
EP-FITC species.
The Low Fluorescence Species: A Supercompact Form of the ATPase
Cytosolic Head, Almost a Transition-like State? Potential Value for
Future Structural Studies--
Along this line, it is particularly
appealing to further speculate that the above-mentioned
"cross-link" could occur between the nucleotide binding domain and
the phosphorylation domain. We know that, at some stage during the
catalytic cycle, the nucleotide domain and the phosphorylation domain
must be able to come close together, to make phosphoryl transfer
possible. The early phosphoenzyme form that is stabilized by FITC is as
close as possible to that state. A further possibility that we may
consider is that the low fluorescence is in fact caused by the
interaction of the FITC moiety with the phosphoryl group itself. It
could help to fix domain N and domain P (34) together and
simultaneously alter the reactivity of the phosphoryl group. The low
fluorescence species might therefore be a compact phosphoenzyme form,
with the cytosolic head domains tightly associated.
Independently of the precise assignment of the residues interacting
with FITC in the low fluorescence species, the structure of the low
fluorescence species might mimic not only that of an early
phosphoenzyme form, but also that of the transition state for
phosphoryl transfer. Indeed, the phosphorylation event for unmodified
ATPase can be broken into several substeps, as illustrated in Scheme
2.
Phosphorylation is thought to be preceded by a rate-limiting
nucleotide-induced conformational change to a species,
Ca2
aE·ATP, from which
phosphorylation is very rapid (49-51). In Scheme 2, the transition
state for phosphoryl tranfer is placed in square brackets to indicate
its transient existence. Jencks and co-workers (50, 52) have found that
Ca2+ dissociates toward the cytosolic side of the membrane
from both
Ca2
E·ATP and
Ca2
aE·ATP. The low
fluorescence FITC-ATPase species, or more likely the
Ca2+-bound species with intermediate fluorescence from
which it is immediately derived, might to some extent mimic this
transition state for phosphoryl transfer. If this idea is correct, the
ATPase cytosolic head can again be expected to be in a very compact
state, with restricted access of water or other phosphoryl acceptors to
the active site.
At any rate, the low fluorescence phosphorylated form of
FITC-ATPase can probably be safely classified as an
"E1P-like" form, on the basis of its being derived from
an early phosphoenzyme in the cycle and of its poor reactivity to
water. The structure of its head region is likely to be substantially
different from that of either the Ca2·E1 or
the E2·VO4 states; the structure of the former
state, deduced from Ca2+-ATPase three-dimensional crystals
grown in the presence of Ca2+, shows a widely open head
region, with the nucleotide binding domain and the phosphorylation
domain separated by ~25 Å (34), whereas two-dimensional crystals of
the E2·VO4 species show a more closed head
region in which full interaction of the nucleotide binding and
phosphorylation domains is nevertheless hindered by decavanadate
binding at the interface between domains (34, 53). The structure of the
low fluorescence species could provide critical new information on head
domain interactions in a closed state and, by comparison with the other
structures, on the extent and nature of domain movements.
In this direction, the fact that the low fluorescence species remained
stable in a detergent-solubilized state (Fig. 1) might be of future
value for three-dimensional crystallization attempts (e.g.
as in Ref. 34 or 54). In addition, we have now found that it is
possible to grow two-dimensional crystals from the low fluorescence and
phosphorylated ATPase in the presence of thapsigargin and decavanadate
(work in progress),2 i.e. under conditions
previously shown to induce the formation of two-dimensional arrays of
unphosphorylated ATPase (53, 55-57). The present
demonstration of a stable, phosphorylated form of FITC-ATPase might
thus provide a starting point toward the future crystallization and
structural analysis for the first time of a phosphorylated form of the pump.
 |
ACKNOWLEDGEMENTS |
We are very grateful to
Stéphane Orlowski for participating to initial experiments, to
Gérard Lecointe for building an amplifier for our pH meter
analogic output, to Franck Delavoie for generating many two-dimensional
FITC-ATPase arrays and analyzing them by electron microscopy, to Marc
le Maire for helping characterize by high pressure liquid
chromatography the detergent-solubilized low fluorescence species and
discussing our results, to Carlos Guttiérrez-Merino for
specifically discussing the spectroscopic properties of FITC in the low
fluorescence ATPase species, to Jesper Møller and Jens P. Andersen for
discussion on various occasions, and to Mathilde Dreyfus for help with copyediting.
 |
FOOTNOTES |
*
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.
Published, JBC Papers in Press, November 6, 2000, DOI 10.1074/jbc.M006980200
2
Further details were submitted with the
manuscript, but were withdrawn for the sake of conciseness, at the
request of the Editor. They are available from the authors.
 |
ABBREVIATIONS |
The abbreviations used are:
SERCA1a, sarcoplasmic or endoplasmic reticulum ATPase, type 1a;
SR, sarcoplasmic
reticulum;
C12E8, octaethylene glycol
monododecyl ether;
DM,
-D-dodecyl maltoside;
MOPS, 4-morpholinepropanesulfonic acid;
A23187, calcimycin;
Pi, inorganic phosphate;
FITC, fluorescein 5'-isothiocyanate;
AcP, acetylphosphate;
VO4, vanadate;
TG, thapsigargin;
E1P, E2P, names given to the postulated
different conformations of phosphorylated ATPase.
 |
REFERENCES |
1.
|
Pick, U.,
and Karlish, S. J. D.
(1980)
Biochim. Biophys. Acta
626,
255-261[Medline]
[Order article via Infotrieve]
|
2.
|
Mitchinson, C.,
Wilderspin, A. F.,
Trinniman, B. J.,
and Green, N. M.
(1986)
FEBS Lett.
146,
87-92[CrossRef]
|
3.
|
Pick, U.
(1981)
FEBS Lett.
123,
131-136[CrossRef][Medline]
[Order article via Infotrieve]
|
4.
|
Champeil, P.,
Guillain, F.,
Vénien, F.,
and Gingold, M. P.
(1985)
Biochemistry
24,
69-81[Medline]
[Order article via Infotrieve]
|
5.
|
Champeil, P.,
Riollet, S.,
Orlowski, S.,
Guillain, F.,
Seebregts, C. J.,
and McIntosh, D. B.
(1988)
J. Biol. Chem.
263,
12288-12294[Abstract/Free Full Text]
|
6.
|
Orlowski, S.,
and Champeil, P.
(1991)
Biochemistry
30,
352-361[Medline]
[Order article via Infotrieve]
|
7.
|
Orlowski, S.,
and Champeil, P.
(1993)
FEBS Lett.
328,
296-300[CrossRef][Medline]
[Order article via Infotrieve]
|
8.
|
Champeil, P.,
Henao, F.,
and de Foresta, B.
(1997)
Biochemistry
36,
12383-12393[CrossRef][Medline]
[Order article via Infotrieve]
|
9.
|
Riollet, S.,
and Champeil, P.
(1987)
Anal. Biochem.
162,
160-162[Medline]
[Order article via Infotrieve]
|
10.
|
Clark, R. J. H.
(1973)
in
Comprehensive Inorganic Chemistry
(Bailar, J. C.
, Emeleus, H. J.
, Nyholm, R.
, and Trotman-Dickenson, A. F., eds), Vol. 3
, pp. 491-551, Pergamon Press, New York
|
11.
|
Tsien, R. T.,
and Pozzan, T.
(1989)
Methods Enzymol.
172,
230-262[Medline]
[Order article via Infotrieve]
|
12.
|
Dupont, Y.
(1978)
Biochem. Biophys. Res. Commun.
82,
893-900[Medline]
[Order article via Infotrieve]
|
13.
|
Beil, F. U.,
von Chak, D.,
and Hasselbach, W.
(1977)
Eur. J. Biochem.
81,
151-164[Abstract]
|
14.
|
Pick, U.,
and Karlish, S. J. D.
(1982)
J. Biol. Chem.
257,
6120-6126[Free Full Text]
|
15.
|
Sagara, Y.,
and Inesi, G.
(1991)
J. Biol. Chem.
266,
13503-13506[Abstract/Free Full Text]
|
16.
|
Highsmith, S.,
Barker, D.,
and Scales, D. J.
(1985)
Biochim. Biophys. Acta
817,
123-133[Medline]
[Order article via Infotrieve]
|
17.
|
Verjovski-Almeida, S.,
and Inesi, G.
(1978)
J. Biol. Chem.
254,
18-21[Medline]
[Order article via Infotrieve]
|
18.
|
Dupont, Y.
(1980)
Eur. J. Biochem.
109,
231-238[Abstract]
|
19.
|
Dupont, Y.
(1984)
Anal. Biochem.
142,
504-510[Medline]
[Order article via Infotrieve]
|
20.
|
Petithory, J. R.,
and Jencks, W. P.
(1988)
Biochemistry
27,
8626-8635[Medline]
[Order article via Infotrieve]
|
21.
|
Champeil, P.,
Gingold, M. P.,
Guillain, F.,
and Inesi, G.
(1983)
J. Biol. Chem.
258,
4453-4458[Abstract/Free Full Text]
|
22.
|
Mintz, E.,
Mata, A. M.,
Forge, V.,
Passafiume, M.,
and Guillain, F.
(1995)
J. Biol. Chem.
270,
27160-27164[Abstract/Free Full Text]
|
23.
|
Feher, J. J.,
and Briggs, F. N.
(1980)
Cell Calcium
1,
105-118
|
24.
|
Madeira, V. M. C.
(1984)
Biochim. Biophys. Acta
769,
284-290[Medline]
[Order article via Infotrieve]
|
25.
|
Klonis, N.,
and Sawyer, W. H.
(1996)
J. Fluorescence
6,
147-157
|
26.
|
Klonis, N.,
Clayton, A. H. A.,
Voss, E. W.,
and Sawyer, W. H.
(1998)
Photochem. Photobiol.
67,
500-510[CrossRef][Medline]
[Order article via Infotrieve]
|
27.
|
Makinose, M.
(1973)
FEBS Lett.
37,
140-143[CrossRef][Medline]
[Order article via Infotrieve]
|
28.
|
Champeil, P.
(1996)
in
Biomembranes
(Lee, A. G., ed), Vol. 5
, pp. 43-76, JAI Press, Greenwich, CT
|
29.
|
Jencks, W. P.
(1989)
J. Biol. Chem.
264,
18855-18858[Free Full Text]
|
30.
|
Friedman, Z.,
and Makinose, M.
(1970)
FEBS Lett.
11,
69-72[CrossRef][Medline]
[Order article via Infotrieve]
|
31.
|
Pucell, A.,
and Martonosi, A.
(1971)
J. Biol. Chem.
246,
3389-3397[Abstract/Free Full Text]
|
32.
|
Bodley, A. L.,
and Jencks, W. P.
(1987)
J. Biol. Chem.
262,
13997-14004[Abstract/Free Full Text]
|
33.
|
Jencks, W. P.,
Yang, T.,
Peisach, D.,
and Myung, J.
(1993)
Biochemistry
32,
7030-7034[Medline]
[Order article via Infotrieve]
|
34.
|
Toyoshima, C.,
Nakasako, M.,
Nomura, H.,
and Ogawa, H.
(2000)
Nature
405,
645-655
|
35.
|
de Meis, L.,
and Vianna, A. L.
(1979)
Annu. Rev. Biochem.
48,
275-292[CrossRef][Medline]
[Order article via Infotrieve]
|
36.
|
Chiesi, M.,
and Inesi, G.
(1980)
Biochemistry
19,
2912-2918[Medline]
[Order article via Infotrieve]
|
37.
|
Yamaguchi, M.,
and Kanazawa, T.
(1984)
J. Biol. Chem.
259,
9526-9531[Abstract/Free Full Text]
|
38.
|
Lévy, D.,
Seigneuret, M.,
Bluzat, A.,
and Rigaud, J. L.
(1990)
J. Biol. Chem.
265,
19524-19534[Abstract/Free Full Text]
|
39.
|
Vilsen, B.,
and Andersen, J. P.
(1986)
Biochim. Biophys. Acta
855,
429-431[Medline]
[Order article via Infotrieve]
|
40.
|
McIntosh, D. B.,
Ross, D. C.,
Champeil, P.,
and Guillain, F.
(1991)
Proc. Natl. Acad. Sci. U. S. A.
88,
6437-6441[Abstract]
|
41.
|
Andersen, J. P.
(1995)
BioSci. Rep.
15,
243-261[Medline]
[Order article via Infotrieve]
|
42.
|
Takakuwa, Y.,
and Kanazawa, T.
(1984)
J. Biochem. (Tokyo)
95,
543-550[Abstract]
|
43.
|
Chiesi, M.,
and Wen, Y. S.
(1983)
J. Biol. Chem.
258,
6078-6085[Abstract/Free Full Text]
|
44.
|
McIntosh, D. B.,
and Woolley, D. G.
(1994)
J. Biol. Chem.
269,
21587-21595[Abstract/Free Full Text]
|
45.
|
Inesi, G.,
and de Meis, L.
(1989)
J. Biol. Chem.
264,
5929-5936[Abstract/Free Full Text]
|
46.
|
Vilsen, B.,
and Andersen, J. P.
(1992)
J. Biol. Chem.
267,
3539-3550[Abstract/Free Full Text]
|
47.
|
Inesi, G.,
Lewis, D.,
Nikic, D.,
Hussain, A.,
and Kirtley, M. E.
(1992)
Adv. Enzymol. Relat. Areas Mol. Biol.
65,
185-215[Medline]
[Order article via Infotrieve]
|
48.
|
Pick, U.,
and Bassilian, S.
(1981)
FEBS Lett.
123,
127-130[CrossRef][Medline]
[Order article via Infotrieve]
|
49.
|
Coan, C. R.,
Verjovski-Almeida, S.,
and Inesi, G.
(1979)
J. Biol. Chem.
254,
2968-2974[Medline]
[Order article via Infotrieve]
|
50.
|
Petithory, J. R.,
and Jencks, W. P.
(1986)
Biochemistry
25,
4493-4497[Medline]
[Order article via Infotrieve]
|
51.
|
Obara, M.,
Suzuki, H.,
and Kanazawa, T.
(1988)
J. Biol. Chem.
263,
3690-3697[Abstract/Free Full Text]
|
52.
|
Stahl, N.,
and Jencks, W. P.
(1987)
Biochemistry
26,
7654-7667[Medline]
[Order article via Infotrieve]
|
53.
|
Zhang, P.,
Toyoshima, C,
Yonekura, K.,
Green, N. M.,
and Stokes, D. L.
(1998)
Nature
392,
835-839[CrossRef][Medline]
[Order article via Infotrieve]
|
54.
|
Lacapère, J. J.,
Stokes, D. L.,
Olofsson, A.,
and Rigaud, J. L.
(1998)
Biophys. J.
75,
1319-1329[Abstract/Free Full Text]
|
55.
|
Sagara, Y.,
Wade, J. B.,
and Inesi, G.
(1992)
J. Biol. Chem.
267,
1286-1292[Abstract/Free Full Text]
|
56.
|
Stokes, D. L.,
and Lacapère, J. J.
(1994)
J. Biol. Chem.
269,
11606-11613[Abstract/Free Full Text]
|
57.
|
Dux,
and Martonosi, A.
(1983)
J. Biol. Chem.
258,
2599-2603[Free Full Text]
|
Copyright © 2001 by The American Society for Biochemistry and Molecular Biology, Inc.