From the Institut für Physikalische und
Theoretische Chemie der Universität Erlangen-Nürnberg,
D-91058 Erlangen, Germany, the ¶ Departamento de
Bioquímica y Biología Molecular, Unidad Asociada al
Consejo Superior de Investigaciones Científicas, Universidad
del País Vasco, Aptdo. 644, 48080 Bilbao, Spain, and the
** Centro Nacional de Biotecnología, Consejo Superior de
Investigaciones Científicas, Universidad Autónoma de
Madrid, 28049 Madrid, Spain
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ABSTRACT |
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Changes in the vibrational spectrum of the
chaperonin GroEL in the presence of ADP and ATP have been followed as a
function of time using rapid scan Fourier transform infrared
spectroscopy. The interaction of nucleotides with GroEL was triggered
by the photochemical release of the ligands from their corresponding biologically inactive precursors (caged nucleotides;
P3-1-(2-nitro)phenylethyl nucleotide). Binding
of either ADP or ATP induced the appearance of small differential
signals in the amide I band of the protein, sensitive to protein
secondary structure, suggesting a subtle and localized change in
protein conformation. Moreover, conformational changes associated with
ATP hydrolysis were detected that differed markedly from those observed
upon nucleotide binding. Both, high-amplitude absorbance changes and difference bands attributable to modifications in the interaction between oppositely charged residues were observed during ATP
hydrolysis. Once this process had occurred, the protein relaxed to an
ADP-like conformation. Our results suggest that the secondary structure as well as salt bridges of GroEL are modified during ATP hydrolysis, as
compared with the ATP and ADP bound protein states.
Chaperonin GroEL from Escherichia coli facilitates
protein folding in vivo and in vitro in an
ATP-regulated manner (1, 2). GroEL consists of two heptameric rings
composed of identical subunits that form a double toroid structure.
Each subunit is organized into two major domains, the apical domain
being involved in GroES and substrate binding, and the equatorial one
holding the nucleotide-binding site and most of the intra- and
inter-ring contacts, both linked by a small and flexible intermediate
domain (3).
Numerous ligands have been shown to modulate the conformation of GroEL
(4, 5). Among them, nucleotides can modify the chemical properties,
structure and substrate-binding affinity of the central cavity of the
protein that binds the substrate (1, 6). These modifications depend on
the nature of the bound nucleotide. The cooperative binding of ATP
switches GroEL from a high affinity state for non-folded proteins (T)
to a protein release state (R) which has low affinity for substrate
proteins (7). ATP hydrolysis drives the oligomer cavity through
alternating states with different affinities for unfolded protein, and
it has been recently demonstrated that nucleotide hydrolysis is
required to release GroES, once ATP binds to the adjacent protein ring (8). Electron microscopy
(EM)1 studies have shown that
ATP induces large conformational changes in GroEL (5, 9). Supporting
these findings, proteolysis studies have revealed a conformational
change in the apical GroEL domain in response to nucleotide binding (4,
10). Moreover, ATP hydrolysis has also been correlated with an
increased exposure of hydrophobic patches at the chaperonin surface
(11).
Although the importance of the molecular events associated to the
nucleotide-induced allosteric transition of GroEL has been widely
recognized, they remain as yet mostly unknown (12). In an effort to
obtain such information, we have applied time-resolved infrared
difference spectroscopy combined with the use of caged nucleotides.
This approach allows to monitor subtle changes in the vibrational
spectrum between two protein states whose interconversion can be
triggered in the infrared cuvette. Moreover, time-resolved IR
spectroscopy makes it possible to follow in real-time the formation of
intermediates in the ATPase reaction cycle of GroEL, with the following
advantages. (i) It is a non-invasive technique, i.e. there
is no need to label the protein, and therefore the possible alteration
of the protein structure by the label is avoided. (ii) It allows to
follow, at the same time, the kinetic parameters of overall protein
conformational changes, and modifications of specific protein groups as
a consequence of nucleotide binding and hydrolysis (13, 14). Our
results demonstrate that GroEL undergoes a transient conformational
transition during ATP hydrolysis, that differs from the one observed
upon ATP or ADP binding.
Sample Preparation--
GroEL was overexpressed from E. coli and purified as described previously (15). The protein was
concentrated with microconcentration filters (Centricon-50 (Amicon)),
and the buffer was exchanged by repeated concentration and dilution
steps with buffer 100 mM imidazole, 100 mM
NaCl, 30 mM MgCl2 prepared in H2O
or D2O, pH or pD 7.0. Infrared samples were prepared by
drying 1 µl of caged nucleotide (20 mM) and 1 µl of 40 mM KCl onto a CaF2 window with a trough of
8-µm depth and 8-mm diameter. For the samples containing DTT, an
additional 1 µl of 20 mM DTT was dried onto the window. They were rehydrated with the same volume of GroEL dissolved in the
above buffer and the samples were sealed with a second flat window and
thermostated at 25 °C during the experiment. Protein subunit
concentration was 0.8 mM, as determined by the
bicinchoninic acid assay (Sigma). Control experiments were measured on
identical samples, prepared without GroEL.
Infrared Spectroscopy--
Infrared measurements were carried
out on a modified Bruker IFS 66 spectrophotometer equipped with a
HgCdTe detector. Data were acquired with double sided interferograms in
a forward-backward mode at a spectral resolution of 4 cm Kinetic Analysis--
To fit the IR absorbance changes to a
kinetic model, we used integrated band intensities. Integration was
performed with respect to a baseline that was drawn between two limits
at each side of the band. To avoid the overlapping effect expected in a
complex difference profile, in some cases the integration boundaries
were chosen so that the resulting area was characteristic of a single band. To determine the rate constants, several bands with high signal-to-noise ratio were selected. For the kinetic analysis the time
slots of spectra recording were represented by their average times.
ATPase Hydrolysis--
The ATPase activity of GroEL was assayed
at 25 °C, using malachite green to measure the amount of inorganic
phosphate released upon ATP hydrolysis as previously reported (16). The
reaction was started by adding GroEL (final oligomer concentration 0.49 µM) to the assay solution containing 50 mM
imidazole, 40 mM KCl, 30 mM Mg2Cl,
100 mM NaCl, pH 7.0, and 1.5 mM free or caged
ATP. Aliquots of the reaction mixture were diluted with 3 volumes of 1 M HClO4, and centrifuged (30 min, 14,000 × g). The supernatant was mixed with the malachite green
reagent and allowed to stand for 2 h at room temperature for the
color to develop. Control experiments were also performed to account
for the spontaneous hydrolysis of the nucleotide, which was found to be
negligible under our experimental conditions.
Nucleotide Binding Assay--
150 µM free and
caged nucleotides in the above buffer were centrifuged in the presence
of the same concentration of GroEL subunits, using microconcentration
filters (Microcon-30 (Amicon)). The filtration membrane retained all
protein material and bound nucleotide while allowed unbound nucleotide
to pass through. Nucleotide concentration in the filtrate was estimated
from the absorbance at 260 nm, using an extinction coefficient of
15,400 M Electron Microscopy and Image Processing--
Samples containing
GroEL and GroES (1:2 molar ratio) in 50 mM imidazole, 40 mM KCl, 30 mM Mg2Cl, 100 mM NaCl, pH 7.0, were incubated with 3 mM caged
nucleotides. After 30 min at 25 °C, they were negatively stained
with 1% uranyl acetate on thin coated collodion grids previously glow
discharged for 15 s. Transmission electron microscopy (JEOL
1200EX-II electron microscope operated at 120 KV) and image processing
were performed as previously reported (5).
Binding and Hydrolysis of Caged Nucleotides by GroEL--
Prior to
the spectroscopic characterization of the interaction of nucleotides
with GroEL, it is necessary to ensure that their caged analogs are not
bound or hydrolyzed by the protein. ATPase activity measurements
clearly indicate that caged ATP is not modified by GroEL, and that
hydrolysis only occurs after ATP release from the cage (Fig.
1). Furthermore, they also demonstrate
that the presence of the free cage does not significantly affect the
ATPase activity of the protein. However, they do not rule out the
possible interaction between the caged ligands and the protein. In
order to test whether binding of caged ADP and ATP to GroEL was taking place before nucleotide release from the cage, two types of experiments were performed. First, filtration experiments pointed out that, unlike
free nucleotides which can bind tightly to half of the 14 available
sites in GroEL (Kd<15 µM; 17), none
of the caged nucleotides were able to bind to the protein, since all the initially added caged ligands appeared in the filtrate (data not
shown). Second, electron microscopy clearly showed that neither caged
ATP nor caged ADP were able to induce the formation of GroES·GroEL asymmetric complexes, in contrast to the free nucleotides (5) (data not
shown). Therefore, we conclude that caged nucleotides do not bind to
GroEL.
ATP-induced Conformational Changes--
The interaction between
ATP and GroEL is accomplished by triggering the release of the ligand
from an inactive (caged) photolabile analog with a UV flash, which does
not disturb the sample (18). The spectrum recorded before illuminating
the sample is the reference, and those measured after the flash are the
samples. The difference spectra show absorbance changes that reflect
conformational alterations of functionally important regions of the
protein, while contributions from the rest of the molecule are
canceled. Positive signals in the difference spectra refer to the
nucleotide-bound GroEL conformation, whereas negative signals are
characteristic of the free, unliganded protein.
The difference IR spectrum recorded 3.6 s after release of ATP to
the medium displays sharp differential features, indicating a
nucleotide-induced conformational transition between distinct protein
conformations (Fig. 2). These signals can
essentially be attributed to molecular processes taking place during
caged ATP photolysis, ATP binding and hydrolysis, and to protein
conformational changes associated with its ATPase activity. The
assignment of the experimental absorbance changes to any of these
processes requires investigation of the photolysis of caged ATP in the
absence of protein (Fig. 2, dashed lines). The bands at 1692 cm
The strongest difference bands in H2O are two positive
signals at 1654 and 1572 cm
A possible interpretation of the 1568 cm
Alternatively, if a modification of the interaction between oppositely
charged residues were responsible for the 1568 cm
To distinguish the possible different conformational states generated
as a consequence of the interaction of ATP with GroEL, we have analyzed
the time dependence of the absorbance changes in the infrared
difference spectrum of the protein (Fig.
3). Release of ATP from the cage occurs
within the first 33 ms after the flash, as judged by the time course of
the intensity of the photolysis bands (see the negative band at 1526 cm ADP-induced Conformational Changes--
A comparison of the
difference spectra recorded 3.6 s after the photolysis flash, in
the presence (solid lines) and absence (dashed
lines) of GroEL, reveals that ADP binding to the protein induces
the appearance of absorbance changes in the 1700-1610 cm There are a number of experimental evidences indicating that the
conformational rearrangements induced in GroEL by the physiologically relevant nucleotides might have functional implications. EM studies have shown that the apical domains of GroEL move differently in the
presence of ATP or ADP (5, 9). Moreover, only in the presence of ATP
and during its hydrolysis does GroEL seem to transiently expose
additional hydrophobic residues as judged from
bis-1-anilino-8-naphthalenesulfonate binding experiments (11).
Functional studies with a disulfide GroEL mutant have also shown that
nucleotide hydrolysis is coupled to a spatial rearrangement of the
protein particle which, in turn, promotes release of the substrate
(25). So far, these experimental evidences have been analyzed in terms
of changes in the tertiary and/or quaternary structures of the protein,
paying little attention to its secondary structure. This is mainly due
to limitations of the spectroscopic techniques to monitor local rather
than overall conformational changes (26). In this context,
"reaction-induced infrared difference spectroscopy" provides an
appropriate tool to analyze the small and localized structural changes
associated with binding to and/or hydrolysis of different nucleotides
by GroEL.
Before discussing the differences between the GroEL conformational
states generated in the presence of ATP or ADP, we shall analyze the
time dependence of the ATP-induced absorbance changes. This might help
to distinguish the consequences of nucleotide binding from its
subsequent hydrolysis on GroEL conformation. As shown by the intensity
of the photolysis bands, nucleotide release to the medium occurs within
the first 33 ms after the flash. Under the experimental conditions used
in this work (high protein and nucleotide concentration, 25 °C) ATP
binding to GroEL is strongly favored. This is supported by the fact
that the ADP-induced absorbance changes reach their maximum amplitude
0.42 s after nucleotide release from the cage. Considering that
the affinity of GroEL for both nucleotides is similar, it is reasonable
to assume that the spectra recorded in the presence of ATP before this
time would correspond to the ATP-bound GroEL state, without contributions from ATP hydrolysis. The corresponding difference spectra
indicate that ATP binding to GroEL induces a small rearrangement of its
secondary structure elements which, in contrast to what is found during
nucleotide hydrolysis, does not affect basic and acidic amino acid side
chains. Although the rate constant of the initial event compares well
with the one described for ATP binding to pyrene-labeled GroEL (17,
27), it should be mentioned that under our experimental conditions
overlapping of the binding and hydrolysis steps might obscure the
kinetic analysis of the binding process.
During the subsequent ATP hydrolysis step, the protein experiences a
more pronounced conformational change, as evidenced by a 3-fold
increase in the intensity of differential bands attributable to the
polypeptide backbone. Moreover, our results also show the appearance of
specific signals tentatively assigned to acidic and basic residues
changing their chemical environment, indicating that ATP hydrolysis is
probably coupled to modifications of salt bridges of the protein. The
transient character of this conformational state is demonstrated by the
recovery of the absorbance changes characteristic of the ADP-bound
state after nucleotide hydrolysis. There are several salt bridges
involved in intra- and intersubunits contacts within the same protein
ring, as well as in inter-ring interactions. It has been shown that the
intrasubunit salt bridge between Glu-409 and Arg-501, at the hinge
between the equatorial and the apical domain and close to the
ATP-binding site, becomes significantly weaker during the ATP-induced
allosteric transitions of GroEL (28). The ion pair formed by
Glu-434 and Lys-105, involved in intra-ring communication, is also
believed to change in the presence of ATP, since the nucleotide greatly
reduces the electron density at this contact site (9). Salt bridges
also provide numerous contact sites between neighboring subunits. Based
on EM and modeling studies, it has been proposed that during the ATP-induced transition from the T to the R state, several salt bridges
between subunit apical domains are modified (29). A conclusive
assignment of the experimentally observed difference signals to
specific amino acids will require the characterization of selected
mutants. The time constant of the slow process is twice lower than the
one estimated from ATPase activity and fluorescence measurements (17,
30). This discrepancy might well be caused by the higher protein and
ATP concentrations used in this work, as it has been shown for the
Ca2+-ATPase from sarcoplasmic reticulum (14).
The difference between the ADP- and ATP-bound states of GroEL is
difficult to ascertain, due to the small amplitude of the absorbance
changes ( The role of ATP hydrolysis in GroEL function still remains unclear. On
one hand, kinetic and structural data indicate that ATP binding is
responsible for the conformational changes in GroEL that release the
bound substrate, while nucleotide hydrolysis simply resets the system.
On the other hand, EM (5, 9) and biochemical data (11) indicate that
the ADP-bound state is structurally different from the unliganded and
ATP-hydrolyzing state. Recent experiments have shown that ATP
hydrolysis in the cis ring weakens the binding of GroEL and
primes it for release, after ATP binding to the trans ring
(8). In this context, our results suggest that during ATP hydrolysis,
GroEL undergoes a unique conformational transition, different from
those caused by ATP or ADP-binding, which involves changes in
solvent-exposed secondary structure elements and modifications of salt
bridges of the protein.
INTRODUCTION
Top
Abstract
Introduction
References
MATERIALS AND METHODS
1. One interferometer cycle corresponding to two
spectra needed 65 ms for completion. After recording a blank difference
spectrum to control for the signal-to-noise ratio and base-line
stability, the experiment started by measuring a reference spectrum
coadded from 300 scans which characterized the unperturbed sample.
Photolysis of caged nucleotides was triggered with a Xenon flash tube
with high UV output. The voltage of the flash power supply was adjusted to release approximately 1.5 mM nucleotide/flash. After the
flash, 20 spectra at 2 scans each, 20 spectra at 4 scans each, 10 spectra at 10 scans each, 20 spectra at 20 scans each, and 20 spectra at 100 scans each were recorded. To additionally improve the
signal-to-noise ratio, signals obtained from different samples were
averaged after normalizing the solvent-subtracted reference spectra of
the protein to the same absorbance value in the amide I (0.4 absorbance
unit in D2O) or amide II (0.06 absorbance unit in
H2O) bands. The difference spectra were not subjected to
smoothing or other resolution enhancement procedures, such as
deconvolution or derivation. The noise level in these spectra, as
confirmed from control experiments, was estimated to be around 5 × 10
5 absorbance unit at frequencies above 1750 cm
1, where no signals appear, and approximately
10
4 absorbance unit at around 1650 in the H2O
samples due to the strong absorbance from H-O-H bending modes. These
noise levels confirmed the reliability of the experimentally observed
weak signals.
1 cm
1. Control
experiments without nucleotide were carried out to subtract background
contributions to this value.
RESULTS
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Fig. 1.
ATPase activity of GroEL in the presence of
free and caged ATP. ATP hydrolysis by GroEL (0.49 µM
oligomer concentration) in the presence of 1.5 mM
nucleotide, 50 mM imidazole, 40 mM KCl, 30 mM Mg2Cl, 100 mM NaCl, pH 7.0, at
25 °C. ATP ( ), caged-ATP (
), and caged-ATP after releasing the
nucleotide form the cage (
).
1 (H2O), 1686 cm
1
(D2O), and those appearing at 1526 cm
1 and
below 1350 cm
1 in both media, contain contributions from
the photolysis reaction. From previous studies, it is known that ATP
hydrolysis gives rise to difference absorbance signals below 1300 cm
1 (19), thus leaving the 1800-1300 cm
1
spectral region free from unwanted signals overlapping with the protein
bands. Therefore, the absorbance changes that will be considered below,
which are observed upon nucleotide release in the presence of GroEL,
can be assigned to protein conformational changes induced by ATP
binding and its subsequent hydrolysis.
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Fig. 2.
Infrared absorbance changes of GroEL in
the presence of ATP. IR difference spectra obtained 3.6 s
after release of approximately 1.5 mM ATP in samples
containing GroEL, 0.8 mM protein subunit ( ), and no
protein-photolysis reaction (· · ·). The spectra recorded at
25 °C in H2O and D2O buffers are the average
of 19 and 11 different experiments, respectively. Insets,
difference spectra recorded under the same experimental conditions but
in the presence of 20 mM DTT.
1. Less intense differential
bands appear at 1692(+)/1673(
), 1641(
)/1629(+), 1556(+)/1546(
),
and 1511(+)/1508(
) cm
1. The effect of deuteration helps
to distinguish bands caused by amide modes of the polypeptide backbone
from bands due to amino acid side chains. The amide I modes (1700-1610
cm
1; mainly due to peptide C=O groups) shift up to 10 cm
1, while the amide II band (mainly due to
amide-backbone NH groups) shifts from 1550 cm
1 to 1460 cm
1 (20-22). In contrast, amino acid side chains
absorbing in this spectral region, i.e. those from Asn, Gln,
Lys, Arg, show larger shifts (23, 24). Deuteration of the sample
induces the following noticeable changes in the difference spectrum of
the protein. 1) The absorbance changes at 1692, 1654, and 1629 cm
1 are downshifted to 1686, 1648, and 1607 cm
1, respectively. 2) The intensity of the differential
signal observed in H2O at 1556(+)/1546(
) is strongly
reduced. 3) The signal at 1511(+)/1508(
) cm
1 almost
completely disappears. The small shift observed for the 1692 and 1654 cm
1 bands suggests that they are due to changes in the
amide I absorbance of the polypeptide backbone. The existence of an
ATP-induced conformational change is also indicated by the
1556(+)/1546(
) differential feature, which can be assigned to the
amide II mode of the protein, because it is virtually not detected upon
deuteration. Instead, a band is observed at 1469 cm
1,
which may represent the amide II' mode.
1 band is the
appearance of carboxylate group(s) upon ATP binding and/or hydrolysis. Indeed, a band at this position has been assigned to -COO
groups of Glu or Asp residues and it has been shown to be influenced very little, as compared with other side chain vibrations, by deuteration (23, 24). This could be achieved by either a change in the
ionization state of a Glu or Asp residue(s), or a modification of the
interaction between oppositely charged residues, i.e. salt bridges. If the former hypothesis would hold, the appearance of this
positive band should be accompanied by a negative counterpart at around
1700-1730 cm
1, that would indicate the disappearance of
a -COOH group(s). Although a signal at this position is not observed in
the difference IR spectra, we have carried out the same experiments in
the presence of DTT to eliminate the photolysis signal that appears at
1692 cm
1 (H2O) and 1686 cm
1
(D2O), that may partially overlap with a COOH vibration.
The results (Fig. 2, insets) demonstrate the presence of a
weak negative signal at 1700 cm
1 in H2O that
is shifted upon deuteration to 1695 cm
1. The position of
this differential feature, downshifted from the characteristic
frequency of the -COOH groups, makes its assignment to a carboxylic
group unlikely. It is important to note that the positions and
amplitudes of all the differential signals mentioned above are
maintained, indicating that the presence of DTT does not modify the
structural transition brought about by the interaction of ATP with GroEL.
1
positive signal, "marker" differential bands for Arg and/or Lys residues should also be observed in the infrared difference spectra. These bands appear in the 1700-1500 cm
1 spectral region,
where they can overlap with vibrations from the amide I and amide II
modes of the protein. Based on studies of model compounds (23, 24), the
protonated side chain of arginine residues gives rise to two signals at
around 1673 and 1633 cm
1 in H2O that shift to
1607 and 1586 cm
1 in D2O, while the
NH3+ modes of Lys at around 1513 cm
1 disappear upon deuteration. Due to the possible
overlap with protein backbone modes, the assignment of the differential
features at around 1673 cm
1 to Arg residues is not
straightforward. However, the putative assignment of the 1629 cm
1 signal to these residues is supported by the fact
that it is virtually abolished upon deuteration. Furthermore, the
absorbance change at 1607 cm
1 in D2O may be
attributed to Arg side chains, and possible contributions from protein
backbone signals are not likely. These results, together with the
disappearance of the 1511(+)/1508(
) cm
1 feature upon
deuteration, suggest that both Arg and Lys residues could experience a
change in their chemical environments during the interaction of ATP
with GroEL. The alternative assignment of the 1511(+)/1508(
)
cm
1 spectral feature, observed only in H2O,
to Tyr residues can be reasonably discarded because if this were the
case neither the position nor the intensity of these vibrations should
be significantly affected by deuteration (23, 24).
1). The corresponding infrared difference spectrum
shows a positive differential signal at 1648 cm
1,
attributable to the protein, which is maintained in the spectra recorded during the first 0.23 s after the flash. At longer time intervals, up to 7 s, the intensity of this and other positive (1607 and 1568 cm
1) and negative (1673 and around 1620 cm
1) differential features increases. The amplitude of
these absorbance changes is maintained during approximately 60 s,
a time interval that most likely corresponds to a multiple turnover
reaction where the enzyme is in the steady-state (note that the
[ATP]/[GroEL subunit] is approximately 2). This interpretation
would be in accordance with kinetic studies using pyrene-labeled GroEL
(17). Afterward, a slow structural change drives GroEL into a
conformation analogous to that exhibited in the presence of ADP (see
below). As compared with the IR difference spectra obtained within the 10-60-s interval, the spectrum of this conformation lacks the 1568 cm
1 signal and shows a positive signal at 1627 cm
1, both characteristic of the ADP-bound state (see Fig.
3, top trace, for a comparison). The analysis of the time
course of the absorbance changes after ATP release is shown in Fig.
4. The selected signals are tentatively
assigned to alterations of the amide I mode of the polypeptide backbone
(1648 cm
1), of acidic amino acid side chains (1568 cm
1), and of phosphate vibrations (1273 cm
1). To properly analyze the time course of these
signals, two time constants were required. Their values for the fast
phase are 0.4, 0.55, and 0.8 s for the 1273, 1648, and 1568 cm
1 bands, respectively, while those of the slow phase
are 3.3 ± 0.3 s for all bands.
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Fig. 3.
Time dependence of the IR difference spectrum
of GroEL in the presence of ATP. Measurements were performed after
release of 1.5 mM ATP in a sample containing 0.8 mM GroEL subunit, at 25 °C in D2O buffer.
Spectra were recorded at the times indicated below each
trace after the photolysis flash. Average of 11 different experiments.
The spectrum labeled ADP (top, thick trace) was recorded
15 s after ADP release in the presence of GroEL.
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Fig. 4.
Kinetics of selected infrared absorbance
changes of GroEL in the presence of ATP, measured in D2O
buffer. Labels refer to the wave number of absorption in
cm 1. Different constants were added to each kinetic trace
to allow for a clear and simultaneous representation.
, 1648 cm
1;
, 1568 cm
1;
, 1273 cm
1.
1 spectral region (amide I band) that cannot be
attributed to the photolysis reaction (Fig.
5). These absorbance changes are located at 1647 and 1628 cm
1 in both H2O and
D2O. The possible contribution of amino acid side chains to
these differential signals can be better considered by analyzing the
effect of deuteration on the difference spectrum. As stated before, the
absence of significant shifts in band position upon deuteration allows
assignment of the observed major absorbance changes to peptide C=0
groups. It is important to note that none of these bands shows the
kinetic behavior observed in the ATP-containing samples (data not
shown), and that they reach their maximum amplitude 0.42 s after
nucleotide release to the medium. More significantly, the ADP-induced
difference spectra do not display absorbance changes attributable to
basic and acidic amino acid side chains, as observed in the presence of
ATP.
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Fig. 5.
Effect of ADP on the infrared difference
spectrum of GroEL. Absorbance changes after ADP release in samples
with ( ) and without (· · ·) GroEL. IR difference spectra
recorded 3.6 s after the release of approximately 1.5 mM ADP. Protein subunit concentration was 0.8 mM. Samples were prepared in H2O and
D2O buffers, and measured at 25 °C. Average of 10 (H2O) and six (D2O) experiments.
DISCUSSION
1% of the amide I band intensity). This, nevertheless,
suggests that the conformational change induced in GroEL upon
nucleotide binding is localized, and it might reflect modifications
within existing secondary structure elements rather than a net change
of secondary structure (14). However, the conformation of the active,
ATP-hydrolyzing state of GroEL is clearly distinct from those of the
nucleotide-bound states. The amplitude of the major absorbance changes
within the amide I increases 3 and 4 times as compared with those
observed upon ATP and ADP binding, respectively. The position of the
strongest absorbance change in H2O (1654 cm
1)
and D2O (1648 cm
1) has been described for
solvent-exposed
-helical structures (31, 32). Less intense
differential signals are characteristic of turns (1686, 1673 cm
1 in D2O) and
-structures (negative
band, at approximately 1626 cm
1 in
D2O). The fact that a differential signal attributable to the amide II mode of the protein also appears during ATP hydrolysis supports the existence of this transient GroEL conformation.
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ACKNOWLEDGEMENT |
---|
We thank Dr. F. M. Goñi for critically reading the manuscript.
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FOOTNOTES |
---|
* This work was supported in part by University of the Basque Country Grant EB200/96 (to A. M.), CICYT Grants BIO97-0820 and PB 97-1225 (to A. M. and J. M. V.), and Acción Concertada Hispano-Alemana.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.
§ Supported by the Deutsche Forschungsgemeinschaft.
Recipient of a fellowship from the University of the Basque Country.
Recipient of a fellowship from the Comunidad Autónoma de Madrid.
§§ Supported by the Deutsche Forschungsgemeinschaft. Present address: Institut für Biophysik, Johan Wolfgang Goethe Universität, Theodor Stern Kai 7, D-60590 Frankfurt.
¶¶ To whom correspondence should be addressed. Tel.: 34-94-6012624; Fax: 34-94-4648500; E-mail: gbpmuvia{at}lg.ehu.es.
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ABBREVIATIONS |
---|
The abbreviations used are: EM, electron microscopy; DTT, dithiothreitol; IR, infrared.
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REFERENCES |
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