From the Institut für Biophysik, Johann Wolfgang Goethe-Universität, Theodor-Stern-Kai 7, Haus 74, D-60590 Frankfurt am Main, Germany
Received for publication, December 5, 2002, and in revised form, January 13, 2003
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ABSTRACT |
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Infrared spectroscopy has been used to map substrate-protein interactions: the conformational changes of the sarcoplasmic reticulum Ca2+-ATPase upon nucleotide binding and ATPase phosphorylation were monitored using the substrate ATP and ATP analogues (2'-deoxy-ATP, 3'-deoxy-ATP, and inosine 5'-triphosphate), which were modified at specific functional groups of the substrate. Modifications to the 2'-OH, the 3'-OH, and the amino group of adenine reduce the extent of binding-induced conformational change of the ATPase, with particularly strong effects observed for the latter two. This demonstrates the structural sensitivity of the nucleotide-ATPase complex to individual interactions between nucleotide and ATPase. All groups studied are important for binding and interactions of a given ligand group with the ATPase depend on interactions of other ligand groups.
Phosphorylation of the ATPase was observed for ITP and 2'-deoxy-ATP,
but not for 3'-deoxy-ATP. There is no direct link between the extent of
conformational change upon nucleotide binding and the rate of
phosphorylation showing that the full extent of the ATP-induced
conformational change is not mandatory for phosphorylation. As observed
for the nucleotide-ATPase complex, the conformation of the first
phosphorylated ATPase intermediate E1PCa2 also depends on
the nucleotide, indicating that ATPase states have a less uniform conformation than previously anticipated.
Ligand binding to proteins controls vast numbers of cellular
processes and has attracted great scientific and economic interest. Protein and ligand flexibility are important determinants of the interaction and often lead to ligand binding modes that are not anticipated from structures obtained with other ligands. To these "failure(s) of the rigid receptor hypothesis" (1) is added here an
impressive example: induced-fit binding of nucleotides to the
Ca2+-ATPase. This finding stems from a systematic mapping
of substrate-protein interactions with infrared
(IR)1 spectroscopy. New
approaches like this are welcome in the field of ligand-protein
recognition, since the most informative techniques, NMR and x-ray
crystallography, are laborious and problematic for some systems.
Methods like fluorescence and luminescence that require less
expenditure also provide less molecular information. We expect that
this technology gap will be bridged by IR spectroscopy.
IR spectroscopy, one of the methods of vibrational spectroscopy,
provides direct information on the molecular level, is cost-effective, and can be universally applied from small soluble proteins to large
membrane proteins under near-physiological conditions. Work summarized
in recent reviews (2-5) has shown that the vibrational spectrum
changes characteristically when a ligand binds to a protein. This
provides a direct observation of ligand binding: no marker compound has
to be introduced to report the binding process, as with many other
methods. Previous work has mostly focused on individual interactions
between a ligand and a protein by monitoring the influence of the
protein environment on the vibrational frequency of a particular group
of the ligand, the signal of which is identified in a complex
vibrational spectrum with the help of isotopically labeled ligands
(6-8).
Here we employ a different approach to probe the role of single
functional groups of a ligand in the interaction with a protein: using
IR spectroscopy we monitored the protein conformational change induced
by binding of substrate analogues, which are modified at specific
functional groups of the substrate. This identifies those functional
groups that are important in the interaction with the protein;
structure-interaction relationships are obtained that are similar to
structure-activity relationships in drug development that relate the
chemical structure of compounds to their pharmacological activity.
This work studies the ATP binding site of the sarcoplasmic reticulum
(SR) Ca2+-ATPase (9-12). The SR Ca2+-ATPase,
an intrinsic membrane protein of about 110-kDa molecular mass,
catalyzes Ca2+ transport from the cytoplasm of muscle cells
into SR for relaxing a flexed muscle. The energy required for this
active transport process is provided by hydrolysis of the substrate
ATP, which phosphorylates the ATPase at Asp351. The
specificity of the SR Ca2+-ATPase for nucleotides is not
high and not only ATP, but also some other nucleotides and
non-nucleotide substrates enable Ca2+ uptake (13-18).
The ATPase structure (19) of the Ca2+-loaded state
E1Ca2 shows three cytoplasmic domains, the nucleotide
binding domain (N-domain), the phosphorylation domain (P-domain), and
the actuator domain (A-domain). The structure has been solved with and
without 2',3'-O-(2,4,6-trinitrophenyl)adenosine 5'-monophosphate (TNP-AMP), which binds to the N-domain at considerable distance from the phosphorylation site Asp351 with less
structural effects than ATP (20, 21). ATP is thought to bind to the
surface of the N-domain with the phosphate groups pointing toward the
phosphorylation site Asp351 (21), and most of the residues
associated with the nucleotide binding site are located in the N-domain
(12, 16, 20-24). Closure of the cleft between N- and P-domain is
thought to occur to bring ATP close to Asp351 upon
nucleotide binding (20, 21). In line with that, mutations at
Asp351, Lys352 (25), and Thr353
(26) alter the affinity of ATP to the ATPase.
IR spectroscopy has been used to characterize conformational changes of
several partial reactions of the Ca2+-ATPase pump cycle, as
reviewed in Ref. 2. To detect the small IR absorbance changes generally
associated with protein reactions, the reactions have to be triggered
directly in the IR cuvette for which we use the photolytical release of
nucleotides from photolabile derivatives, i.e.
P3-1-(2-nitrophenyl)ethyl nucleotides (caged
nucleotides) (27). Here we characterize the conformational change
induced by binding of the following ATP analogues: inosine
5'-triphosphate (ITP), 2'-deoxy-ATP, and 3'-deoxy-ATP (Fig.
1). They differ from ATP at individual
functional groups which allows us to investigate the impact of these
groups on the binding-induced conformational change.
Materials
IR samples were prepared as described previously (28).
Approximate concentrations of the samples based on 1-µl sample volume were: 1.2 mM Ca2+-ATPase, 0.5 mg/ml
Ca2+ ionophore (A23187), 150 mM
methylimidazole (pH 7.5), 150 mM KCl, 10 mM
CaCl2, 5 mM DTT, and 10 mM caged
nucleotide. ITP samples were also prepared with 10 mM DTT
because of a higher conversion of caged ITP in these samples.
In our control samples the nucleotide binding site was already
saturated with We did not use the physiological co-substrate Mg2+ but 10 mM Ca2+ instead. These conditions were chosen
to block the E1PCa2 Methods
FTIR Measurements--
Time-resolved Fourier transfer IR
measurements of the Ca2+-ATPase reaction were performed at
1 °C as described previously (28, 29). Photolytic release of
nucleotides from their respective caged derivatives was triggered by a
xenon flash tube or a XeCl excimer laser. Spectra were obtained in the
following way: a reference spectrum was recorded with the protein in
the E1Ca2 state. After applying a photolysis flash or a
sequence of flashes, we started to record time-resolved IR spectra with
65-ms time resolution. The number of photolysis flashes needed for
saturating signals was determined as described under "Titration of IR
Signals." Difference spectra were obtained by subtracting the
reference spectrum from a spectrum obtained after nucleotide release.
They reflect ATP binding and ATPase phosphorylation as well as the
photolysis reaction. The spectra were normalized to a standard protein
concentration before averaging spectra from different samples (amide II
absorbance: 0.26) as described (35). A photolysis spectrum was then
subtracted as described in the following paragraph to eliminate the
photolysis band. The resulting spectra are named nucleotide binding
spectra or E1PCa2 formation spectra.
Subtraction of the Photolysis Spectrum--
The spectrum
obtained with the control samples shows only signals caused by the
photolysis of caged AMPPNP. It is named photolysis spectrum
and was used to subtract the photolysis bands from the raw difference
spectra as described (33) using the same time interval for both
spectra. This photolysis spectrum is identical to that of other caged
nucleotides above 1300 cm Nucleotide Binding Spectra--
For the nucleotide binding
spectra time windows after the photolysis flash were evaluated in which
the nucleotide-ATPase complex (E1NTPCa2) accumulates. They
were between 0.46 and 0.90 s for ATP or between 0.46 and 3.24 s for ATP analogues. 23 experiments from 12 samples were averaged for
the ATP binding spectrum (~3 mM released ATP, one flash),
four experiments from four samples for the ITP binding spectrum (~6.6
mM released ITP, three flashes), eight experiments from
four samples for the 2'-deoxy-ATP binding spectrum (~3 mM
released 2'-deoxy-ATP, one flash), and three experiments from three
samples for the 3'-deoxy-ATP binding spectrum (~3 mM released 3'-deoxy-ATP, one flash).
Spectra of E1PCa2 Formation--
The difference
spectra of E1PCa2 formation (E1Ca2 Kinetics of Nucleotide Binding and Phosphorylation Reaction and
Fitting Procedures--
The time constants of nucleotide binding and
ATPase phosphorylation were obtained by fitting the integrated band
intensities of the marker band at 1628 cm Titration of IR Signals--
To obtain saturating signals, we
titrated the amplitude of bands at 1641/1628 cm Absorption Spectra of Nucleotides--
Absorption spectra of 500 mM ATP, 2'- and 3'-deoxy-ATP dissolved in H2O
were measured using two BaF2 windows (5-µm path length) with a Bruker Vector 22 spectrometer equipped with a deuterated triglycine sulfate (DTGS) detector at 20 °C at different pH
values. The population of the C2'-endo and
C3'-endo puckering modes of the three
nucleotides were obtained by calculating the ratio of the areas of
bands fitted to the spectrum near 830 cm Titration of IR Signals with Nucleotide Binding--
We first
established the nucleotide concentration needed to obtain saturating IR
signals. For that we used the difference in amplitude of the band pair
at 1628 and 1641 cm
For these titration experiments, spectra where evaluated in a time slot
where the first phosphoenzyme E1PCa2 accumulates. Strictly
speaking, they have therefore determined the nucleotide concentration
necessary for saturating E1PCa2, not for saturating E1NTPCa2. However, the reactions of nucleotide binding and
of phosphorylation are well separated in time, which ensures that E1NTPCa2 also saturates in the time slot evaluated for the
nucleotide binding spectra: time constants for nucleotide binding
(tn) and phosphorylation
(tp) and the time slot of spectra recording for the nucleotide binding spectra (ts) were: for
ATP, tn Nucleotide Binding Spectra--
Fig.
3A shows IR absorbance changes
induced by nucleotide binding to the Ca2+-ATPase. The
spectra reflect the difference in absorbance between the initial
nucleotide-free state E1Ca2 and the nucleotide-ATPase complex E1NTPCa2. Negative bands are characteristic of
E1Ca2 and positive bands of E1NTPCa2. Groups or
structures not involved in the conformational change do not show up in
the difference spectra.
The difference spectra reflect conformational changes of the protein
backbone in the amide I (1700-1610 cm
The spectrum of ATP binding is in close agreement with the AMPPNP
binding spectrum as noted before (33). The positive signal near 1653 cm
The contour of all nucleotide binding spectra is similar; the main
difference is the amplitude of the signals indicating various extents
of conformational change. For further evaluation we used the MSA
(difference between the absorbance change at 1628 cm Spectra of Phosphoenzyme Formation--
Phosphorylation leads to
the appearance of two bands at 1721 and 1549 cm Phosphorylation Rate--
The rate of phosphorylation was measured
using the marker bands at 1721 and 1549 cm Interaction between ATPase and ATP--
Our results show that
modifications to the amino, 2'-OH, 3'-OH, and
The effects of modifying ATP on nucleotide binding might have several
causes: (i) a direct interaction of the modified group of ATP with the
ATPase, (ii) a direct interaction of the "new" group of the ATP
analogue, and (iii) an indirect effect on the interactions between
protein and ATP via a change in electron density or conformation of ATP.
A direct interaction is the most likely cause for the reduced extent of
conformational change observed for the deoxy-ATPs, since (i) no
sterical restrains are expected from the replacement of the hydroxyl
groups by the smaller hydrogen atoms, and (ii) effects on the
equilibrium between sugar conformations of ATP in solution seem to be
less relevant for the sugar conformation of the bound ATP molecule
discussed as follows. 2'- or 3'-H substitution influence the sugar
conformation in solution; according to the absorption spectra of ATP
and 2'- and 3'-deoxy-ATP with deprotonated phosphate groups, the ratios
of C2'-endo and C3'-endo puckering of these three nucleotides are 60:40, 80:20, and 70:30, respectively. This shows that free nucleotides prefer
C2'-endo puckering, particularly
deoxynucleotides. Similar results for ATP and 2'-deoxy-ATP were
obtained before (43-45). NMR investigations demonstrate that
C2'- and C3'-endo types of
conformations are in rapid equilibrium in solution (43), indicating
only a small activation barrier between the conformations. Despite the predominant C2'-endo puckering in solution, the
ATPase seems to choose the C3'-endo conformation
for binding, as determined by NMR (45). Therefore the conformation of
the nucleotide-ATPase complex will not depend on the predominant sugar
puckering in solution and the effects of ribose OH substitution are
best explained by direct interactions of the ribose hydroxyls with the ATPase.
It is less certain whether the reduced extent of conformational change
found with ITP can be explained by a localized interaction between the
amino group and the ATPase. In ITP the carbonyl group replaces the
amino group of ATP, and one of the two endocyclic nitrogen atom is
protonated. These alterations are not localized only on the amino group
but will change the electron density distribution in the entire
six-membered ring and its hydrogen bonding pattern. Therefore the
interaction seems to be located on the six-membered ring of adenine.
Since the most drastic alteration is at the amino group, it is likely,
but not mandatory, that our results reflect a direct interaction of the
ATPase with the amino group.
All functional groups of ATP investigated here are important for
inducing the conformation of the ATP-ATPase complex that is competent
for phosphoryl transfer. This is shown by the dependence of the
phosphorylation rate on the modification of 2'-OH, 3'-OH, and the
adenine amino group. Therefore, interactions distant from the phosphate
groups contribute to approaching or forming the phosphate binding
pocket. Binding of ATP to the ATPase turns out to be an interactive
process where the formation of interactions of a given functional group
of ATP is reinforced by interactions of other groups, which can be at
the opposite end of the ATP molecule.
Distance between
There is no simple link between the extent of conformational change
upon nucleotide binding and the ability to form the phosphoenzyme: (i)
the extent is larger for 2'-deoxy-ATP than for ITP but the apparent
phosphorylation rates are very close, and (ii) the extent is similar
for ITP and 3'-deoxy-ATP, but significant phosphorylation is only
observed for ITP. If the conformational change detected in our spectra
and the distance between Concerted Conformational Change--
The interactions between
nucleotide and protein induce a concerted conformational change upon
nucleotide binding: they join forces to induce strain in the protein.
If one of the interacting groups is modified to become a less effective
binder, the interactions with the respective binding pocket are
impaired, the strain is relieved, and a smaller conformational change
is produced. A weakened interaction therefore affects the
conformational change as a whole instead of producing only local
effects. This concept explains that the modifications of ATP studied
reduce all bands in the amide I region of the difference spectrum. If
an interaction between nucleotide and ATPase had only local effects on
the protein structure, a weakened interaction would selectively reduce
the amplitude of difference bands associated with that conformational
change, but not of all of the bands as observed here. Particularly
interesting is that functional groups of ATP, which interact with
different domains of the protein, produce the same type of
conformational change: the amino function is thought to interact with
the N-domain (19, 24, 46) and the Nucleotide-specific Conformation--
Our results suggest that
nucleotide binding induces a conformation that is characteristic of the
bound nucleotide, as proposed earlier from experiments that did not
monitor the conformation of the nucleotide-ATPase complex directly
(50). In light of the known flexibility of the N-domain (51, 52), this
conformation might represent an average conformation. The (average)
conformation adopted in the ATPase-nucleotide complex seems to be very
sensitive to individual interactions between ATPase and nucleotide,
since the extent of conformational change depends dramatically on the presence of individual functional groups of ATP.
Our finding of a nucleotide specific conformation of the
nucleotide-ATPase complexes is supported by previous reports, in which
different effects of different nucleotides were found on fluorescence
properties (17, 18, 53), partial reaction rates (54-57), protection
against proteolysis (21), effects of aromatic compounds (58),
nucleotide binding properties of mutants (25), and uncoupling (59).
The structures of the nucleotide-ATPase complexes studied differ in two
aspects: (i) the extent of the conformational change induced by
nucleotide binding differs as indicated by the different amplitudes of
the amide I signals, and (ii) structural details of the
nucleotide-ATPase complex differ as shown by the subtle differences of
band positions and spectral shape among the nucleotide binding spectra.
A structure characteristic of the nucleotide is inferred not only for
the nucleotide-ATPase complex but also for E1PCa2 where the
conformation of the phosphoenzyme depends on the nucleotide that was
used for phosphorylation.
The small conformational change upon ITP binding observed here suggests
that soaking E1Ca2 crystals with The Conformational Change Reflected in IR
Spectra--
Conformational changes in two regions of the protein were
proposed to occur upon nucleotide binding (21): (i) movement of the
N-domain toward the P-domain and (ii) movement of the A-domain toward
the P-domain. The latter does not seem to contribute to a large extent
to our spectra for the following reason: Danko et al. (21)
studied protection of the ATPase against proteolytic attack by various
nucleotides. This effect is thought to reflect a movement of the
A-domain. They found no effect for ADP, indicating that ADP does not
promote significant movement of the A-domain. Our IR spectra of
nucleotide binding, however, show that ADP binding induces a
conformational change, the extent of which is two-thirds of that
induced by ATP (28) (Fig. 3B). This shows that the
conformational change of the A-domain contributes not or only to a
small extent to the IR difference spectra.
Instead, it is likely that the anticipated hinge movement of the N- and
P-domain upon ATP binding causes the amide I signals. The hinge
movement will, however, not directly reflect in our spectra,
because highly mobile structural elements give broad IR bands before
and after the conformational change, which largely cancel in the
difference spectrum. Therefore IR spectroscopy will largely miss a
conformational change in the mobile hinge region itself. In
line with this, only small bands in a limited spectral region (1660 to
1680 cm
These conformational changes in well structured regions might report
the hinge movement indirectly, since a conformational change in the
hinge region will also affect the connecting stretches. These stretches
become more ordered the more they are incorporated into the domains and
therefore give rise to distinct bands in the amide I region; a hinge
movement will alter the relative orientation of the connecting amide
groups and their hydrogen bonding and therefore affect their amide I
signals. From these considerations we think that our spectra detect the
hinge movement indirectly because it is reported by
structured backbone stretches that link hinge and domains.
The Hinge Movement between N- and P-domain--
The hinge movement
upon nucleotide binding seems, however, to be less pronounced than
anticipated in the structural models (61, 62). Fluorescence energy
transfer experiments show that distances between fluorescence labels in
the N- and the P-domain do not change between E1Ca2 or an
E2 conformation, as reviewed in Ref. 63. Of particular interest is the
unchanged distance of two pairs of residues for which a change in
distance is expected from the two x-ray structures (19, 46). The
distance between Cys344 and Lys515 increases
from 46 Å in E2 to 50 Å in E1Ca2, and that between Cys344
and Glu439 changes from 38 Å in E2 to 45 Å in E1Ca2.
These distance changes should result in decreases in fluorescence
energy transfer by 33 and 41%, respectively, which are not observed
(64, 65).
The hinge movement can bring the N- and P-domain close together in the
E1Ca2 state, since they can be cross-linked with
glutaraldehyde (66). The cross-linked cleft of E1Ca2
resembles that of the E2 structure, since the cross-linked residues are
only 5 Å apart in the E2 structure but 21 Å in the E1Ca2
structure. In line with these experiments, closure of the hinge could
be modeled with the N- and P-domain structures of E1Ca2
without steric clashes, and this brings the two cross-linked residues
as close as 4 Å (62). The mobility of the N-domain (51, 52) implies
that it is likely to move rather independently from the rest of the protein and that the hinge angle might depend less on the E2 and E1Ca2 state than expected from the crystal structures. The
more closed conformation in E2 and the open cleft between the N- and P-domain in E1Ca2 of the crystal structures therefore most
likely do not represent the average conformation of these states in
solution. They are probably adopted in the crystals because of crystal
contacts that are made possible by the mobility of the N-domain in both states (51, 52). In solution the average position of the N-domain will
be probably in between those observed in the two crystal structures.
Therefore it is plausible to assume that the cleft is less open for
E1Ca2 in solution than in the E1Ca2 crystal
structure, and that upon nucleotide binding the hinge movement
between the N- and P-domain will be smaller than anticipated from the
crystal structure.
Our results demonstrate that IR spectroscopy can be used to map
ligand-protein interactions and may become an important tool for
research as well as for drug and herbicide optimization. In the
particular case of ATP binding to the SR Ca2+-ATPase,
modifications to the 2'-OH, 3'-OH, and amino group of ATP reduce the
induced-fit movement of the Ca2+-ATPase, with the
six-membered ring of adenine and the 3'-OH of ribose exerting key
interactions. Nucleotide binding seems to be a flexible and interactive
process: the conformation of the complex is characteristic of the bound
nucleotide, and the interactions to a given ligand group depend on
interactions of other ligand groups. This finding may also shed new
light on the ongoing controversy on the number of nucleotide binding
sites. Many of these studies have been conducted with ATP analogues. If
binding of an ATP analogue induces a conformation that is
characteristic of only that analogue, results with different analogues
are not necessarily comparable and do not necessarily reflect the
effects of ATP binding. Therefore we propose that some of the
conflicting results can be explained by the different conformations of
the complexes obtained with different analogues.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Structures of ATP and ATP
analogues.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
,
-imidoadenosine 5'-triphosphate (AMPPNP) at the
beginning of the experiment. The composition of these samples was the
same as of that with caged AMPPNP (10 mM) except for the presence of AMPPNP (5 mM). Photolysis of caged AMPPNP in
these samples did not lead to further nucleotide binding and
conformational changes, they only revealed the effects of caged
compound photolysis.
E2P transition to achieve a maximum
level of E1PCa2 in the steady state after nucleotide
release while slowing down the phosphorylation reaction (29).
Replacement of Mg2+ at the catalytic site by
Ca2+ decreases the rate of phosphorylation by 1 order of
magnitude (30-32), enabling a longer observation time for
E1ATPCa2 and therefore a better signal to noise ratio of
the ATP binding spectrum. Ca2+ instead of Mg2+
has been used by us before (29, 33, 34) and gave very similar spectra
for the E1Ca2
E1PCa2 (34) and the
E1Ca2
E2P reaction (compare Fig. 6A of Ref.
34 and Fig. 4a of Ref. 29).
1, i.e. outside the
region of phosphate absorption.
E1PCa2) in Fig. 4 were obtained by subtracting the
reference spectrum (E1Ca2) from the spectrum of the protein
in the E1PCa2 state obtained in the time window between 4.5 and 28.1 s. A photolysis spectrum averaged in the same time window
was subtracted using the same subtraction factors as for the respective
nucleotide binding spectra.
1 for nucleotide
binding and two marker bands at 1721 and 1549 cm
1 for
phosphorylation (29, 35) with the second (the third for ITP results)
and first order exponential decay equations, respectively (Origin 5.0),
and averaging the resulting time constants. 23 experiments were
averaged for ATP, 8 for 2'-deoxy-ATP and 11 for ITP.
1 by
repeating an experiment on the same sample consisting of a reference
spectrum, a photolysis flash, and a spectrum of 300 scans in a time
interval when the first phosphoenzyme E1PCa2 had formed.
The number of flashes needed for saturating signals was used in further experiments.
1 for
C2'-endo puckering and near 814 cm
1 for C3'-endo puckering
(36-38).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1 in the amide I region of the IR
spectrum, which is sensitive to conformational changes.
This difference is termed maximum signal amplitude (MSA). Fig.
2 shows the result of titrations with
ATP, ITP, 2'-deoxy-ATP, and 3'-deoxy-ATP applying a total of eight flashes, which released ~9.4 mM nucleotide. According to
Fig. 2, the binding-induced amplitude difference MSA reached saturating values with the first flash for ATP, 2'- and 3'-deoxy-ATP, and with the
third flash for ITP. From the photolysis efficiency of 30% we then
calculated the saturating nucleotide concentration assuming 1 µl
of sample volume: 3 mM (one flash) for ATP and 2'- and
3'-deoxy-ATP and 6.6 mM (three flashes) for ITP.
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Fig. 2.
Titration of IR signals of E1PCa2
formation (1 °C, pH 7.5). MSA is the difference between the
absorbance change at 1628 cm 1 and that at 1641 cm
1. Data points are connected by lines to guide the eye
of the reader. Approximate concentrations of the samples were: 1.2 mM Ca2+-ATPase, 0.5 mg/ml Ca2+
ionophore (A23187), 150 mM methylimidazole, 150 mM KCl, 10 mM CaCl2, 10 mM DTT, and 10 mM caged nucleotide.
0.11 s, tp = 2.0 s, ts = 0.46-0.90 s; for 2'-deoxy-ATP, tn
0.25 s, tp = 6.7 s, ts = 0.46-3.24
s; for 3'-deoxy-ATP, tn
0.49 s,
tp > 100 s, ts = 0.46-3.24 s; and for ITP, tn
0.23 s, tp = 8.66 s, ts = 0.46-3.24 s (see "Methods"). The
starting time of 0.46 s for spectra averaging might seem to be
early for ITP and 2'- and 3'-deoxy-ATP, since it is close to the time
constant for nucleotide binding. However, increasing the starting time
to 1.5 s for 3'-deoxy-ATP and 0.8 s for ITP and 2'-deoxy-ATP
did not change the maximum signal amplitude MSA by more than 3%.
Therefore we kept the time slot for averaging consistent for the three
ATP analogues and as large as possible for an optimum signal to noise ratio.
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Fig. 3.
A, nucleotide binding spectra
(E1Ca2 E1NTPCa2) of ATP, 2'-deoxy-ATP,
3'-deoxy-ATP, and ITP binding to the Ca2+-ATPase (1 °C
and pH 7.5). The labels indicate the band positions of the ATP binding
spectrum. B, MSA values of the nucleotide binding spectra
with S.D. bars. MSA is the difference between the absorbance
change at 1628 cm
1 and that at 1641 cm
1 in
the nucleotide binding spectra. In the samples there were: 1.2 mM Ca2+-ATPase, 0.5 mg/ml Ca2+
ionophore (A23187), 150 mM methylimidazole, 150 mM KCl, 10 mM CaCl2, 5 mM DTT (10 mM DTT for ITP samples), and 10 mM caged nucleotide.
1) and amide II
(1580-1500 cm
1) region of the spectra. In addition,
environmental and structural changes of side chains and nucleotide
contribute in the whole mid-IR region shown. We will focus here on the
binding-induced absorbance changes in the amide I region.
1 is characteristic of an
-helical structure, the
signals near 1693, 1641, and 1628 cm
1 of
-sheets. Turn
structures likely contribute to the signals near 1665 cm
1. The spectrum indicates that
-helices,
-sheets,
and turns are affected by ATP binding in line with previous findings
(33).
1 and
that at 1641 cm
1). As shown in Fig. 3B, the
largest binding-induced signals were obtained with ATP and AMPPNP (28)
(MSA
3 × 10
3), medium size signals
(MSA
2 × 10
3) with ADP (28), and
2'-deoxy-ATP, and the smallest signals (MSA
1 × 10
3) with ITP and 3'-deoxy-ATP. The different amplitudes
of the nucleotide binding spectra cannot be explained by incomplete
binding to the ATPase, since we have verified that saturating signals
have been obtained (see above). MSA values shown in Fig. 3 for
nucleotide binding spectra differ slightly from MSA values shown in
Fig. 2, for which a time slot was evaluated in which
E1PCa2, not E1NTPCa2, accumulated, because of
conformational changes accompanying the phosphorylation reaction.
1, which
serve here as marker bands for the first phosphorylated intermediate
E1PCa2 (29, 35). ATP, 2'-deoxy-ATP, and ITP, but not
3'-deoxy-ATP, phosphorylate the ATPase at a rate that is sufficiently
high to observe accumulation of the E1PCa2 state. Fig.
4 shows spectra of E1PCa2
formation from E1Ca2, i.e. the absorbance of
E1PCa2 minus the absorbance of E1Ca2 (see
"Methods"). As found for nucleotide binding, the shape of the
E1PCa2 formation spectra is similar for the analogues but
the amplitude is different. In contrast, the same amplitude
is observed for the band at 1721 cm
1, which has been
tentatively assigned to the C=O group of Asp351 formed upon
phosphorylation (34, 35). This local probe of the phosphorylation
reaction shows that E1PCa2 accumulates to the same extent
with ITP and 2'-deoxy-ATP as with ATP. The smaller signals obtained
with the analogues can therefore not be explained by incomplete
phosphorylation. Instead they are due to a smaller extent of
conformational change showing that the conformation of
E1PCa2 depends on the nucleotide used for phosphorylation
of the ATPase.
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[in a new window]
Fig. 4.
Spectra of E1PCa2 formation
(E1Ca2 E1PCa2)
obtained with ATP, 2'-deoxy-ATP, and ITP. Conditions are the same
as described in the legend to Fig. 3.
1 (29, 35). The
kinetics of the two bands is shown in Fig. 5. Phosphorylation of the ATPase by
2'-deoxy-ATP or ITP is slower (0.15 ± 0.01 s
1 and
0.12 ± 0.01 s
1, respectively) than by ATP
(0.51 ± 0.03 s
1). Slower phosphorylation of the
ATPase with ITP and 2'-deoxy-ATP compared with ATP has been observed
(39, 40), but without specifying the rate for ATP. Previous findings
revealed a low phosphoenzyme concentration for both 2'- and
3'-deoxy-ATP (40), which we found only for 3'-deoxy-ATP. This is likely
due to the different buffers used, since nucleotide binding (41) and
the associated conformational change (28) depend on the composition of
the medium.
View larger version (15K):
[in a new window]
Fig. 5.
Kinetics of phosphoenzyme
(E1PCa2) formation with ATP, 2'-deoxy-ATP, and ITP at 1721 and 1549 cm 1. Conditions are
the same as described in the legend to Fig. 3.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-phosphate groups of
ATP affect the binding-induced conformational change of the ATPase.
These groups are therefore important for the interaction between ATP
and the ATPase. 3'-OH and the region near the amino group have the most
significant influence on the induced-fit movement of the ATPase, since
modification to either of these groups reduces the extent of backbone
conformational change seen by IR spectroscopy to only one-third of that
obtained with ATP. They are therefore important groups of ATP that
anchor ATP to the ATPase. The importance of the functional groups of ATP investigated here for several partial steps of the ATPase reactions
cycle has been shown before (39, 40, 42). The new finding here is that
modifications of the ATP molecule have a direct effect on the structure
of the nucleotide-ATPase complex. This is valid not only for side chain
orientation as often found (1) but also for backbone conformation: with
some ATP analogues the binding-induced conformational change of the
backbone seen by the IR spectroscopy was found to be only one-third of
that for ATP.
-Phosphate and Asp351--
Our
data show that phosphorylation does not strictly depend on the full
extent of the conformational change achieved by ATP binding. This is in
line with the observation that pseudo substrates like acetyl phosphate
are able to produce the same kind of phosphoenzyme as ATP (42),
although they are not expected to induce the same conformational change
as ATP, because their structures are even more different from ATP than
those of the nucleotides investigated here.
-phosphate and phosphorylation site
Asp351 were correlated, a small conformational change would
place the
-phosphate further away from Asp351 than a
larger one and result in slower phosphorylation. Therefore the
conformational change seen in our spectra seems to be not or only
weakly correlated with the distance between
-phosphate and
Asp351. This finding is in line with models where the
-phosphate in the nucleotide-ATPase complex is still some distance
away from the phosphorylation site, as proposed by Hua et
al. (46) for the Ca2+-ATPase and Ettrich et
al. (47) for the Na+/K+-ATPase. Then, the
-phosphate arrives at the catalytic site only after nucleotide
binding, which could take place during the conformational change after
nucleotide binding that has been identified as the rate-limiting step
for phosphorylation (48). A
-phosphate in some distance to the
phosphorylation site provides a possibility of binding a regulatory ATP
molecule to the phosphoenzyme at the same site (49).
-phosphate with the P-domain (25, 26, 46). Despite that, the absence of the
-phosphate in ADP (28) or
of the adenine amino group in ITP both reduce the amplitude of the same
bands. This shows that the concerted conformational change detected
here is caused by interactions of the nucleotide with different protein
domains: the N- and the P-domain.
,
-iminoinosine 5'-triphosphate (IMPPNP) may not disrupt the crystals as ATP does (19)
because of the relatively small conformational change seen here for
ITP. This may therefore enable the investigation of nucleotide binding
at atomic resolution. The conformational change induced by ATP binding
may then be extrapolated from the conformational change seen upon
IMPPNP binding.
1) can be assigned to mobile structures, since
they exchange their amide proton upon
1H2O/2H2O exchange
(33). Instead the nucleotide binding bands in the amide I region are
caused by backbone stretches within well defined and stable structures,
since they are hardly affected by
1H2O/2H2O exchange
(33). In line with this finding of conformational changes in well
structured regions, NMR spectroscopy has detected changes of backbone
conformation in the N-domain upon AMP binding (60).
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ACKNOWLEDGEMENTS |
---|
We thank W. Mäntele for continuous support, W. Hasselbach (Max-Planck-Institut, Heidelberg, Germany) for the gift of Ca2+-ATPase and J. E. T. Corrie (National Institute for Medical Research, London) and F. von Germar for the preparation of caged compounds. We are grateful to C. Toyoshima for sharing unpublished results with us.
![]() |
FOOTNOTES |
---|
* This work was supported by Deutsche Forschungsgemeinschaft Grant Ba1887/2-1.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: Dept. of Biochemistry
and Biophysics, Arrhenius Laboratories for Natural Sciences, Stockholm
University, S-106 91 Stockholm, Sweden. Tel.: 46-8-16-2452; Fax:
46-8-15-5597; E-mail: Andreas.Barth@dbb.su.se.
Published, JBC Papers in Press, January 21, 2003, DOI 10.1074/jbc.M212403200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
IR, infrared;
SR, sarcoplasmic reticulum;
N-domain, the nucleotide binding domain;
P-domain, the phosphorylation domain;
A-domain, the actuator domain;
TNP-AMP, 2',3'-O-(2,4,6-trinitrophenyl)adenosine
5'-monophosphate;
caged nucleotide, P3-1-(2-nitrophenyl)ethyl nucleotides;
A23187, calcium ionophore;
AMPPNP, ,
-imidoadenosine 5'-triphosphate;
E1Ca2, the nucleotide-free ATPase;
E1NTPCa2, the nucleotide-ATPase complex;
DTGS, deuterated triglycine sulfate;
MSA, maximum signal amplitude;
IMPPNP,
,
-iminoinosine
5'-triphosphate.
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REFERENCES |
---|
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---|
1. | Davis, A. M., and Teague, S. J. (1999) Angew. Chem. Int. Ed. 38, 736-749[CrossRef] |
2. | Barth, A., and Zscherp, C. (2000) FEBS Lett. 477, 151-156[CrossRef][Medline] [Order article via Infotrieve] |
3. | Wharton, C. W. (2000) Nat. Prod. Rep. 17, 447-453[CrossRef][Medline] [Order article via Infotrieve] |
4. | Carey, P. R., and Tonge, P. J. (1995) Acc. Chem. Res. 28, 8-13 |
5. | Deng, H., and Callender, R. (1999) Methods Enzymol. 308, 176-201[Medline] [Order article via Infotrieve] |
6. | Belasco, J. G., and Knowles, J. R. (1980) Biochemistry 19, 472-477[Medline] [Order article via Infotrieve] |
7. | Cepus, V., Scheidig, A. J., Goody, R. S., and Gerwert, K. (1998) Biochemistry 37, 10263-10271[CrossRef][Medline] [Order article via Infotrieve] |
8. | Baenziger, J. E., Miller, K. W., and Rothschild, K. J. (1993) Biochemistry 32, 5448-5454[Medline] [Order article via Infotrieve] |
9. | Hasselbach, W., and Makinose, M. (1961) Biochem. Z. 333, 518-528 |
10. | Hasselbach, W. (1974) in The Enzymes (Boyer, P. D., ed), 3rd Ed. , pp. 431-467, Academic Press, New York |
11. | Andersen, J. P. (1989) Biochim. Biophys. Acta 988, 47-72[Medline] [Order article via Infotrieve] |
12. | Lee, A., and East, J. (2001) Biochem. J. 356, 665-683[CrossRef][Medline] [Order article via Infotrieve] |
13. | Hasselbach, W. (1979) Top. Curr. Chem. 78, 1-56[Medline] [Order article via Infotrieve] ] |
14. | Hasselbach, W. (1981) in Membrane Transport (Bonting, S. L. , and De Pont, J. J. H. H. M., eds) , pp. 183-208, Elsevier Science Publishers B. V., Amsterdam |
15. | Inesi, G., and De Meis, L. (1985) in The Enzymes of Biological Membranes (Martonosi, A., ed), 2nd Ed., Vol. 3 , pp. 157-191, Plenum Press, New York |
16. | McIntosh, D. B. (1998) Adv. Mol. Cell Biol. 23A, 33-99 |
17. |
Lacapere, J.-J.,
Bennett, N.,
Dupont, Y.,
and Guillain, F.
(1990)
J. Biol. Chem.
265,
348-353 |
18. | Wakabayashi, S., and Shigekawa, M. (1990) Biochemistry 29, 7309-7318[Medline] [Order article via Infotrieve] |
19. | Toyoshima, C., Nakasako, M., Nomura, H., and Ogawa, H. (2000) Nature 405, 647-655[CrossRef][Medline] [Order article via Infotrieve] |
20. | MacLennan, D. H., and Green, N. M. (2000) Nature 405, 633-634[CrossRef][Medline] [Order article via Infotrieve] |
21. | Danko, S., Yamasaki, K., Daiho, T., Suzuki, H., and Toyoshima, C. (2001) FEBS Lett. 505, 129-135[CrossRef][Medline] [Order article via Infotrieve] |
22. | Andersen, J. P., and Vilsen, B. (1992) Acta Physiol. Scand. 146, 151-159 |
23. | Mcalennan, D. H., Clarke, D. M., Loo, T. W., and Skerjanc, I. S. (1992) Acta Physiol. Scand. 146, 141-150[Medline] [Order article via Infotrieve] |
24. |
McIntosh, D. B.,
Woolley, D. G.,
Vilsen, B.,
and Andersen, J. P.
(1996)
J. Biol. Chem.
271,
25778-25789 |
25. |
McIntosh, D.-B.,
Woolley, D.-G.,
MacLenna, D.-H.,
Vilsen, B.,
and Andersen, J.-P.
(1999)
J. Biol. Chem.
274,
25227-25236 |
26. |
Clausen, J. D.,
McIntosh, D. B.,
Wooley, D. G.,
and Andersen, J. P.
(2001)
J. Biol. Chem.
276,
35741-35750 |
27. | Kaplan, J. H., Forbush, B., and Hoffman, J. F. (1978) Biochemistry 17, 1929-1935[Medline] [Order article via Infotrieve] |
28. | Liu, M., and Barth, A. (2002) Biospectroscopy 67, 267-270[Medline] [Order article via Infotrieve] |
29. |
Barth, A.,
von Germar, F.,
Kreutz, W.,
and Mäntele, W.
(1996)
J. Biol. Chem.
271,
30637-30646 |
30. | Suzuki, H., Nakamura, S., and Kanazawa, T. (1994) Biochemistry 33, 8240-8246[Medline] [Order article via Infotrieve] |
31. |
Shigekawa, M.,
Wakabayashi, S.,
and Nakamura, H.
(1983)
J. Biol. Chem.
258,
8698-8707 |
32. |
Lacapere, J.-J.,
and Guillain, F.
(1990)
J. Biol. Chem.
265,
8583-8589 |
33. |
Von Germar, F.,
Barth, A.,
and Mäntele, W.
(2000)
Biophys. J.
78,
1531-1540 |
34. | Barth, A., Kreutz, W., and Mäntele, W. (1994) Biochim. Biophys. Acta 1194, 75-91[Medline] [Order article via Infotrieve] |
35. |
Barth, A.,
and Mäntele, W.
(1998)
Biophys. J.
75,
538-544 |
36. | Wartell, R. M., and Harrell, J. T. (1986) Biochemistry 25, 2664-2671[Medline] [Order article via Infotrieve] |
37. | Taillandier, E., Ridoux, J. P., Liquier, J., Leupin, W., Denny, W. A., Wang, Y., Thomas, G. A., and Peticolas, W. L. (1987) Biochemistry 26, 3361-3368[Medline] [Order article via Infotrieve] |
38. | Ouali, M., Letellier, R., Sun, J. S., Akhebat, A., Adnet, F., Liquier, J., and Taillandier, E. (1993) J. Am. Chem. Soc. 115, 4264-4270 |
39. |
de Meis, L.,
and de Mello, M.-C.-F.
(1973)
J. Biol. Chem.
248,
3691-3701 |
40. |
Coan, C.,
Amaral, J. A.,
and Verjovski-Almeida, S.
(1993)
J. Biol. Chem.
268,
6917-6924 |
41. | Hasselbach, W., and The, R. (1975) Eur. J. Biochem. 53, 105-113 |
42. |
Bodley, A. L.,
and Jencks, W. P.
(1987)
J. Biol. Chem.
262,
13997-14004 |
43. | Schleich, T., Blackburn, B. J., Lapper, R. D., and Smith, I. C. P. (1972) Biochemistry 11, 137-145[Medline] [Order article via Infotrieve] |
44. | Davies, D. B., and Danyluk, S. S. (1974) Biochemistry 13, 4417-4434[Medline] [Order article via Infotrieve] |
45. | Clore, G. M., Gronenborn, A. M., Mitchinson, C., and Green, N. M. (1982) Eur. J. Biochem. 128, 113-117[Abstract] |
46. | Hua, S., Inesi, G., Nomura, H., and Toyoshima, C. (2002) Biochemistry 41, 11405-11410[CrossRef][Medline] [Order article via Infotrieve] |
47. | Ettrich, R., Melichercik, M., Teisinger, J., Ettrichova, O., Krumscheid, R., Hofbauerova, K., Kvasnicka, P., Schoner, W., and Amler, E. (2001) J. Mol. Model 7, 184-192 |
48. | Petithory, J. R., and Jencks, W. P. (1986) Biochemistry 25, 4493-4497[Medline] [Order article via Infotrieve] |
49. |
Bishop, J. E.,
Al-Shawi, M. K.,
and Inesi, G.
(1987)
J. Biol. Chem.
262,
4658-4663 |
50. | Rubtsov, A. M., Quinn, P. J., and Boldyrev, A. A. (1988) FEBS Lett. 238, 240-244[CrossRef][Medline] [Order article via Infotrieve] |
51. | Jona, I., Matko, J., and Martonosi, A. (1990) Biochim. Biophys. Acta 1028, 183-199[Medline] [Order article via Infotrieve] |
52. | Huang, S., and Squier, T. C. (1998) Biochemistry 37, 18064-18073[CrossRef][Medline] [Order article via Infotrieve] |
53. | Kubo, K., Suzuki, H., and Kanazawa, T. (1990) Biochim. Biophys. Acta 1040, 251-259[Medline] [Order article via Infotrieve] |
54. |
Champeil, P.,
Riollet, S.,
Orlowski, S.,
Guillain, F.,
Seebregts, C. J.,
and McIntosh, D. B.
(1988)
J. Biol. Chem.
263,
12288-12294 |
55. | Hobbs, A. S., Albers, R. W., Froehlich, J. P., and Heller, P. F. (1985) J. Biol. Chem. 260, 2035-2057[Abstract] |
56. |
Ferreira, S. T.,
and Verjovski-Almeida, S.
(1988)
J. Biol. Chem.
263,
9973-9980 |
57. | Scofano, H. M., Vieyra, A., and De Meis, L. (1979) J. Biol. Chem. 254, 10227-10231[Abstract] |
58. |
Petretski, J. H.,
Wolosker, H.,
and De Meis, L.
(1989)
J. Biol. Chem.
264,
20339-20343 |
59. |
Fortea, M. I.,
Soler, F.,
and Fernandez-Belda, F.
(2000)
J. Biol. Chem.
275,
12521-12529 |
60. | Abu-Abed, M., Mal, T.-K., Kainosho, M., Maclennan, D.-H., and Ikura, M. (2002) Biochemistry 41, 1156-1164[CrossRef][Medline] [Order article via Infotrieve] |
61. | Toyoshima, C., and Nomura, H. (2002) Nature 418, 605-611[CrossRef][Medline] [Order article via Infotrieve] |
62. | Xu, C., Rice, W. J., He, W., and Stokes, D. L. (2002) J. Mol. Biol. 316, 201-211[CrossRef][Medline] [Order article via Infotrieve] |
63. | Bigelow, D. J., and Inesi, G. (1992) Biochim. Biophys. Acta 1113, 323-338[Medline] [Order article via Infotrieve] |
64. | Stefanova, H. I., Mata, A. M., Gore, M. G., East, J. M., and Lee, A. G. (1993) Biochemistry 32, 6095-6103[Medline] [Order article via Infotrieve] |
65. | Mata, A. M., Stefanova, H. I., Gore, M. G., Khan, Y. M., East, J. M., and Lee, A. G. (1993) Biochim. Biophys. Acta 1147, 6-12[Medline] [Order article via Infotrieve] |
66. |
McIntosh, D. B.
(1992)
J. Biol. Chem.
267,
22328-22335 |