From the Department of Molecular and Cellular
Biology, Faculty of Biotechnology, University of Gdansk, 80-822 Gdansk,
Kladki 24, Poland, the § Department of Molecular Biology,
International Institute of Molecular and Cell Biology UNESCO in Warsaw,
4 Ks. Trojdena Street, 02-109 Warsaw, Poland, and the
¶ Département de Biochimie Médicale, Centre
Medical Universitaire, 1, rue Michel-Servet,
1211 Geneva 4, Switzerland
Received for publication, August 17, 2000, and in revised form, January 11, 2001
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ABSTRACT |
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The ClpX heat shock protein of Escherichia
coli is a member of the universally conserved Hsp100 family of
proteins, and possesses a putative zinc finger motif of the
C4 type. The ClpX is an ATPase which functions both as a
substrate specificity component of the ClpXP protease and as a
molecular chaperone. Using an improved purification procedure we show
that the ClpX protein is a metalloprotein complexed with Zn(II)
cations. Contrary to other Hsp100 family members, ClpXZn(II) exists in
an oligomeric form even in the absence of ATP. We show that the single
ATP-binding site of ClpX is required for a variety of tasks, namely,
the stabilization of the ClpXZn(II) oligomeric structure, binding to
ClpP, and the ClpXP-dependent proteolysis of the The Clp family of ATPases can function both as specificity
components for their corresponding proteolytic subunit partners or
perform certain chaperone functions on their own. For example, they are
able to protect or dissociate other protein aggregates, or dissociate
specific tertiary protein structures (1-9). Both the proteolytic and
chaperone activities of the Clp ATPases depend on ATP hydrolysis. More
recent data demonstrate that chaperone and protease activities can
occur concurrently in the ClpAP complex. For example, the ClpP protein
is able to restore the chaperone activity of a ClpA mutant carrying an
amino acid substitution in its N-terminal ATP-binding site. ClpA(K220V)
is unable to activate RepA on its own, but the presence of ClpP
restores its ability to activate RepA (10).
In the presence of ATP, the ClpA or Hsp104 proteins oligomerize to give
rise to a hexameric ring structure (10-13). Electron microscopic
studies show that the hexameric ClpA or ClpX ring structure binds to
the double ring, barrel-like 7-fold symmetric ClpP component, thus
giving rise to a structure that closely resembles the eukaryotic 26 S
proteosome (14-17). The Clp ATPase activity is induced in the presence
of its corresponding specific protein substrates (4, 18, 19). ClpA,
ClpB, and Hsp104 each possess two ATP-binding sites. It has been shown
by site-directed mutagenesis that the first ATP-binding site of ClpA,
located near the N-terminal end of the protein, is responsible for
hexamer formation and chaperone activity, whereas the second
ATP-binding site is essential for ATP hydrolysis (10, 12, 20).
Interestingly, in the case of yeast Hsp104 the roles are apparently
reversed; the first ATP-binding site is responsible for ATPase activity
and the second site is essential for oligomerization (11). In contrast
to ClpA, ClpB, or Hsp104, the ClpX ATPase possesses only one
ATP-binding site, highly homologous to the one responsible for
hydrolysis (21, 22).
In this paper we investigate the putative role of Zn(II) in ClpX
structure and function. We show that ClpX binds Zn(II) and that such an
effect is important for binding of ATP to ClpX, and its proper
oligomerization. These events influence binding of ClpX to ClpP and,
consequently, the proteolysis reaction catalyzed by the ClpXP protease.
DNA Cloning and Mutagenesis--
Plasmid pBAD24, which contains
the clpX wild type gene, was constructed as follows. The
coding sequence of clpX was amplified from DNA extracted
from Escherichia coli B178 by means of two oligonucleotides
(synthesized by Life Technologies, Inc.) derived from the published
sequence of clpX (21).
The N-terminal primer was 5'-
GGTTCTCATGACAGATAAACGCAAAGATGGC-3', introducing a BspHI
restriction site and the C- terminal primer was
5'-CCAAGCTTCTGCAGTTATTCACCAGATGCCTGTTGCGC-3', carrying a
PstI restriction site. Polymerase chain reaction was
carried out in a reaction volume of 50 µl. The annealing temperature
was 60 °C and 35 cycles were performed. The polymerase chain
reaction product was cloned into the NcoI-PstI
sites of the pBAD24 expression vector.
The sequence of clpX was confirmed by restriction
analysis and automated sequencing using the ABI Prism 310 DNA sequencer (PerkinElmer Life Sciences, Applied Biosystems). The resulting plasmid construct was used to transform E. coli DH5 Oligonucleotide-directed Mutagenesis of clpX--
The
clpX expression phagemid was mobilized with VCSM13
(Stratagene) in strain CJ236 and amino acid substitutions were
generated using the site-directed mutagenesis method of Kunkel et
al. (23). The cysteine to serine substitutions at residues
15, 18, 37, and 40 were introduced using the following oligonucleotides
(synthesized by Interactiva Biotechnologie): C15S,
5'-GCCGCAAAAAGAGGAATACAGCAATTT-3'; C18S, 5'-
GCTTTTGCCGCTAAAAGAGCAATAC-3'; C37S, 5'-ACACATTCGGAGATATACACG-3'; C40S,
5'-CATAAATCAACAGATTCGTCGGAGAT-3'. All residues downstream from codon
C40S were as in wild type ClpX. The DNA sequence was verified by
automated sequencing using the ABI-Prism 310 DNA sequencer (PerkinElmer
Life Sciences Inc., Applied Biosystems).
Proteins--
In all experiments described in this paper highly
purified proteins (95% or greater purity) were used. All steps of
purification were carried out at 0-4 °C. Thirty grams of ClpX
overexpressing E. coli cells were lysed in 170 ml of X
buffer composed of 50 mM Tris (pH 7.8), 200 mM
KCl, 5 mM DTT,1
10% (w/v) sucrose, 300 mM spermidine-HCl, and 1 mg/ml
lysosyme. To ensure complete lysis, following a 45-min incubation at
0 °C, the lysate was transferred to 42 °C for an additional 5 min
and the concentration of salt was increased to 1 M. The
lysate was centrifuged in a Beckman R35 rotor at 20,000 rpm for 30 min
at 0 °C. Proteins in the supernatant (160 ml) were precipitated with ammonium sulfate (0.29 g/ml) and centrifuged at 25,000 rpm for 45 min
at 0 °C using the same rotor. The pellet was resuspended and
dialyzed for 6 h at 0 °C against buffer X1, composed of 50 mM Tris-HCl (pH 7.6), 100 mM KCl, 5 mM DTT, 0.1 mg/ml phenylmethylsulfonyl fluoride, 10% (v/v)
glycerol, and 0.01% Triton X-100. The extract was centrifuged in an
R35 Beckman rotor at 25,000 rpm for 4 min at 0 °C. The supernatant
was applied onto a Q-Sepharose (Amersham Pharmacia Biotech) column
(2.5 × 20 cm) which had previously been equilibrated with buffer
X1. After extensive washing (12 h at 30 ml/min) of the Q-Sepharose
column with buffer X1, the bound proteins were eluted with a salt
gradient (2 × 300 ml) from 100 to 450 mM KCl.
Fractions containing the ClpX protein were pooled (50 ml) and applied
onto a hydroxylapatite column (Bio-Rad; 1 × 10 cm) equilibrated
with buffer X1. The bound proteins were eluted with potassium phosphate
in buffer X1 (2 × 25 ml;) from 0 to 300 mM
KPi (pH 7.2). Pooled fractions containing the ClpX protein
(15 ml) were dialyzed against buffer X2, composed of 50 mM
Tris-HCl (pH 7.6), 100 mM KCl, 10% (v/v) glycerol, 5 mM DTT, and 0.01% Triton X-100 and applied onto a Resource
MonoQ (Amersham Pharmacia Biotech) column (6 ml). A linear gradient of
KCl (2 × 30 ml) from 100 to 400 mM (1 ml/min flow
rate) was used to elute ClpX from the column using an fast protein
liquid chromatography (Amersham Pharmacia Biotech) system. Fractions
containing homogenous ClpX were pooled together (7 ml) and Bradford
Bio-Rad assay was performed to estimate the protein concentration as 5 mg/ml. SDS-PAGE and size exclusion chromatography on Superdex 200 (Amersham Pharmacia Biotech; 1 × 30 cm) and Biosil TSK-3000
(Beckman; 7 × 300 mm) were used to assess protein purity.
The ClpP was purified as described by Wojtkowiak et al.
(24). The GroEL protein was purified as described by Ziemienowicz et al. (25). The wild type Protease Assays--
The standard protease assay reaction
mixture (100 µl) contained 3H-labeled Kinetics of ClpXP-mediated Hydrolysis of ATP-binding Assays--
Reactions were carried out in a volume
of 25 µl. Samples contained 20 µg of ClpX or ClpX ATPase Activity Assay--
ATPase activity was measured
according to the colorimetric method of Lanzetta et al. (28)
in a buffer containing 50 mM Tris/HCl (pH 7.8), 10% (v/v)
glycerol, 10 mM MgCl2, 150 mM KCl,
25 mM NaCl in a final reaction volume of 50 µl. Samples
contained 20 µg of ClpX or ClpX Protein-Protein Interaction Assays--
The sensitive ELISA
assay used for monitoring protein-protein interactions has been
previously described in detail by Wawrzynow et al. (4).
Size Exclusion Chromatography--
The reacting components (100 µl) were incubated at 30 °C for 30 min in buffer B (25 mM Hepes/KOH (pH 7.6), 150 mM KCl, 25 mM NaCl, 5 mM MgCl2, 10% (v/v)
glycerol, and 5 mM DTT) before loading onto a Superdex 200 HR 10/30 sizing column (Amersham Pharmacia Biotech) equilibrated with
the same buffer. The chromatography was carried out at a flow rate of
0.3 ml/min (room temperature) using Gold HPLC system (Beckman) equipped
with a diode array detector. In the case where no ATP (or 0.2 mM) was used in the column buffer, eluting proteins were
monitored using absorption at 280 nm. Fractions were collected and the
presence of protein was visualized following SDS-PAGE and Coomassie
Blue staining. The relative amount of protein was estimated using
densitometry (Bio-Rad). The Superdex 200 column was calibrated with the
following Bio-Rad molecular weight standards: thyroglobulin (670 kDa),
bovine Stoichiometry of Zinc Cations in ClpX--
Increasing
amounts of PMPS (p-hydroxymercuriphenylsulfonic acid), at a
concentration 1 mM, were added to a sample containing 600 µl of 5 µM ClpX. The reactants were mixed and the
absorbance at 500 nm was measured. Before addition of the first aliquot
of PMPS, the spectrophotometer was adjusted to the zero absorbance value. PAR (4-(2-pyridylazo)resorcinol) was present in the cuvette at
0.1 mM throughout the measurements. The absorption
coefficient for the (PAR)2·ZN(II) complex at 500 nm is
Removal of Zn2+ Cations from the ClpXZn(II) Protein
using a Denaturation and Renaturation Procedure--
ClpX was
denatured in the buffer containing 50 mM Tris-HCl (pH 7.8),
10% (v/v) glycerol, 150 mM KCl, 25 mM NaCl, 10 mM DTT, and 8 M urea. In order to remove the
zinc ions the same buffer supplemented with 10 mM EDTA was
exchanged several times. The renaturation was carried out upon slow
dialysis in the same buffer but in the absence of urea and EDTA.
CD Spectra--
CD spectra measurements were carried out using a
Jasco-J500 CD spectrophotometer in 1-mm cuvettes at 25 °C, in a
buffer containing 20 mM Tris-HCl (pH 7.4), 150 mM KCl, and 25 mM NaCl.
Infrared Spectra--
The IR spectra of ClpX Zn(II), ClpX Atomic Spectroscopy--
This was performed using a AAS 30 Carl
Zeis Jena spectrometer.
ClpX Is a Metalloprotein--
ClpX is a molecular chaperone which
can also work as a specificity factor for the ClpP protease (4, 24). It
contains a single ATP-binding site, a substrate-binding domain called
"sensor and substrate discrimination" or SSD domain (31), and a
putative Zn finger motif, containing four cysteines, of unknown
function. Quantitation of the released Zn(II) ions, upon chelating with PMPS, and by atomic absorption
showed that one Zn(II) ion is bound to
each monomer of ClpX (Table I and Fig.
1). To determine how many Zn(II)
molecules bind to ClpX, we incubated a highly purified ClpX protein
preparation, fully active in the
Using site-directed in vitro mutagenesis procedures, we
replaced the four cysteine residues at positions 15, 18, 37, 40 with serine. As predicted, the resulting mutant, termed ClpX
Quantitative analysis of the specific activity of the ClpX protein in
the The Presence of Zn(II) Affects the Interaction of ClpX with
ClpP--
Fig. 5A shows that,
as judged by the ELISA technique, ClpX Zn(II) strongly interacts with
the ClpP proteolytic subunit only in the presence of ATP. When the ClpX
protein was unfolded by 8 M urea and then folded back in
the absence of Zn(II), it possessed a substantially reduced affinity
for the ClpP protein (Fig. 5B). The subsequent addition of
Zn(II) to such a ClpX preparation restored the proteins ability to form
a stable complex with ClpP. Moreover, only in the presence of Zn(II)
this reaction clearly was ATP-dependent (Fig.
5B).
These ELISA results have been complemented by size exclusion
chromatographic studies. The apparent molecular mass of the ClpP/ClpX Zn(II) heterocomplex is estimated to be 700-800 kDa, assuming an
overall spherical shape for its structure (Fig.
6 and results not shown). Its Stokes
radius is slightly lower than that of the GroEL protein, whose
molecular mass is ~800 kDa (results not shown). A similar high
molecular weight structure was previously described for the ClpP·ClpA
complex (14, 32) and recently for the ClpP·ClpX complex (17). As in
the case of the ClpAP protease, the formation of the oligomeric
ClpP/ClpX Zn(II) structure requires the continuous presence of ATP,
i.e. ATP must be present in both the premixture and mobile
phase buffers (Fig. 6 and results not shown).
Zn(II) and ATP Affect the Proper Oligomerization of ClpX--
The
data presented in Fig. 6 also show that ClpX Zn(II) alone, in the
absence of ATP, or in the presence of low concentrations of ATP (0.2 mM, sufficient for binding of ClpX to ClpP), already behaves as an oligomer. Judging from the shape of its HPLC profile, a
mixture of various ClpX Zn(II) oligomeric forms probably exists. The
HPLC profiles of ClpX Zn(II) were independent of the presence or
absence of exogenous Zn(II) or substitution of ATP by ADP in the
running buffer (results not shown). It turns out that in the absence of
ATP or ADP the Stokes radius of ClpX Zn(II) (Fig. 6A) is
very close to that of the ClpP 14-mer (Fig. 6B), suggesting that ClpXs apparent molecular mass is close to 300 kDa. Therefore, at
least a portion of the ClpX Zn(II) protein presumably exists in an
hexameric form (6 × 46 kDa). A similar oligomeric form has been
reported for the ClpY ATPase (15) and ClpX (17) using an EM approach.
Our EM observations2 also
confirmed these published results. ClpY, an E. coli ATPase, highly homologous to ClpX was recently shown to also be an hexamer (33).
When the ATP concentration was increased to 6 mM, which is
optimal for the ClpXP proteolytic activity (24), an additional oligomerization or stabilization of the specific quaternary structure of ClpX Zn(II) took place (Fig. 6A, and results not shown).
The elution profile of ClpX no longer overlapped that of ClpP,
suggesting that ClpX is found in a form higher than an hexamer under
these conditions. In control experiments we showed that substitution of
ATP by ADP in these experiments also leads to the additional oligomerization of ClpX (result not shown). The Stokes radius of ClpP
did not change at higher concentrations of ATP or ADP (Fig.
6B, and results not shown). We found that the minimal ATP concentration required for the additional oligomerization/stabilization of ClpXs quaternary structure was ~2-3 mM (results not
shown). An analogous ATP-promoted oligomerization has been previously reported for the ClpA and Hsp104 family members. In the absence of nucleotide, ClpA or Hsp104 migrate as monomers, dimers, or trimers
depending on the particular technique used (11, 12, 20).
Interestingly, increasing the ATP concentration from 0.2 to 6 mM did not change the Stokes radius of the ClpP·ClpX
Zn(II) complex (Fig. 6, C and D). Assuming that
all these structures possess a spherical shape, the estimated Stokes
radius of the ClpP/ClpX hetero-oligomer suggests that two hexameric
ClpX Zn(II) molecules are bound to the two ClpP heptamers. These
experiments also suggest that at low concentrations of ATP (0.2 mM) the presence of ClpP helps the ClpX Zn(II) oligomers
either to oligomerize further or to bind to opposite sides of the
barrel-shaped, 7-fold symmetric ClpP structure, as recently shown by
electron microscopy (17).
The release of Zn(II) from the ClpX complex partially inhibited the
oligomerization of ClpX when tested at either high (6 mM)
or low (0.2 mM) concentrations of ATP (results not shown), suggesting that not only ATP but also Zn(II) are required for ClpXs
proper oligomerization. Interestingly the ClpX Zn(II) Is Required for Binding of ATP to ClpX--
Previously we
have shown that ClpX possesses an ATPase activity, stimulatable by its
various protein substrates (4). As shown in Table
II, the ClpX Previous studies have established that zinc fingers, and other
metal-binding protein domains, are involved in protein/DNA interactions, protein folding, as well as protein-protein interactions (for a review, see Ref. 34). We had earlier shown that another molecular chaperone, DnaJ, is also a metalloprotein. In that case, the
binding of two Zn(II) metal ions per DnaJ monomer stabilized the
protein and affected its affinity for its various protein substrates
(30). The C4HC4H motif of the DnaJ protein
could be a novel metal binding motif that differs from those of the established major classes, namely the TFIIIA/Krupple group
(C2H2), steroid receptor (C8),
retroviral gag protein (C2HC), fungal protein such as Gal4
(C6), and bmi-1, the nuclear regulator of the
myc oncogene (C3HC4) (for
reviews, see Refs. 34 and 35). In the case of ClpX the putative Zn
binding motif probably belongs to the C4 family and
contains the following sequence:
CX2CX18CX2C. However, the data
presented in this paper only demonstrate that at least one of the four
cysteine residues is required for Zn(II) binding. More detailed
analysis of the ClpX zinc-binding motif is necessary to really prove
that ClpX contains a C4 binding motif.
In this work we have shown that dissociation of Zn(II) from the ClpX
complex affects all known activities of the ClpX protein, i.e. binding of ATP, protein oligomerization, binding to
ClpP, and ClpXP-dependent proteolysis. Although it is
difficult to definitely conclude which effect is the primary one, it is
very likely that the inability of ClpX lacking Zn(II) to bind ATP, is
responsible for eliminating all of its known activities.
The Zn(II)-dependent binding of nucleotides to proteins was
previously shown to occur in the case of the E. coli MukB
and SlyD proteins. The homodimeric MukB, which is required for the correct partitioning of the bacterial chromosome into daughter cells,
binds GTP or ATP in the presence of Zn(II) but not of Mg(II) (36). In
the case of SlyD, the peptidyl prolyl cis-trans isomerase, the binding
of nucleotide was also Zn(II)-dependent (37). It is
possible that the positively charged Zn(II) is required for the
stabilization of the complex between protein and the negatively charged
phosphate groups of ATP.
We have shown that ClpX Zn(II) is present in an oligomeric state even
in the absence of ATP. Such an oligomeric structure in the absence of
ATP was also observed for ClpY(HslU), a protein highly homologous to
ClpX (15). Interestingly, in the case of ClpX, the tendency for
oligomerization decreases in the absence of Zn(II). In the case of the
mutant ClpX The Zn(II)-dependent oligomerization of ClpX in the
absence of exogenous ATP is not due to the presence of tight ClpX·ATP or ADP complexes in the purified preparations of ClpX protein. Preliminary experiments show that the oligomerization of ClpX is
reduced in the case of ClpX point mutants located in the ATP-binding site. In this case, the ClpX mutant protein does not bind ATP but still
interacts with Zn(II).3 The
results presented in this paper suggest that two factors influence the
oligomerization of ClpX, namely Zn(II) and ATP. It is possible that in
the absence of ATP, the ClpX Zn(II) protein possesses a tendency to
form different oligomeric structures and that binding of ATP stabilizes
one of these structures, very likely the hexameric ring.
Here we have also shown that at physiological concentrations of ATP
(2-6 mM), at which the single ClpX ATP-binding site should be saturated, even further protein oligomerization can take place. Such
phase transition at high ATP concentrations suggests that ClpX can
adopt an unique quaternary structure in the absence of ClpP. One
possibility, suggested by EM studies (17), is that under such
conditions a double hexameric ring sandwich is formed. Interestingly,
such a ClpX quaternary structure is formed also in the presence of ADP,
suggesting that nucleotide binding but not hydrolysis is required for
this event. Moreover, this finding suggests that following ATP
hydrolysis this putative ClpX structure can persist.
Finally, we have shown that low concentrations of ATP (0.2 mM), under which ClpX alone is not present in the putative
double hexameric ring state, are, nevertheless, sufficient for
ClpP·ClpX complex formation. Surprisingly, the Stokes radius of the
ClpP·ClpX complex formed at 0.2 mM ATP is similar to that
formed at 6 mM ATP, suggesting that both complexes contain
the same number of ClpX subunits. Estimation of the molecular mass of
the ClpP·ClpX heterocomplex (700-800 kDa) leads to the conclusion,
assuming an overall spherical shape for this structure, that ClpX is
present in a double hexameric ring state in this complex. This suggests that at 0.2 mM ATP the presence of ClpP helps the ClpX
protein to oligomerize further or that two hexameric ClpX structures
can bind simultaneously to opposite sides of the ClpP barrel-like structure (17).
O
replication protein. Release of Zn(II) from ClpX protein affects the
ability of ClpX to bind ATP. ClpX, free of Zn(II), cannot oligomerize,
bind to ClpP, or participate in ClpXP-dependent
proteolysis. We also show that ClpX
Cys, a mutant protein whose four
cysteine residues at the putative zinc finger motif have been replaced
by serine, behaves in similar fashion as wild type ClpX protein whose
Zn(II) has been released either by denaturation and renaturation, or
chemically by p-hydroxymercuriphenylsulfonic acid.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
(F-,
dlacZ
M15,
lacZYA-argF, U169, deoR, recA1, end A1, phoA, hsdR17,
supE44,
- thi-1, gyrA96, relA1) and CJ236
(dut1, ung1, thi1, relA1/pCJ105, Cmr)
strains for overproduction and mutagenesis, respectively.
O protein was purified as
described by Roberts and McMacken (26).
O protein (0.5 µg, 40,000 cpm), ClpX (0.2 µg), ClpP (2 µg) in 20 mM
Hepes/KOH (pH 7.2), 10 mM MgCl2, 10 mM ATP, 0.5% Brij 58, 4 mg/ml bovine serum albumin. The
reaction was assembled on ice, then transferred to 30 °C for the
desired time. The reaction was stopped by the addition of ice-cold
trichloroacetic acid (final concentration 10% (v/v)). After
centrifugation (10 min, 5,000 × g, at 4 °C) the
radioactivity present in the soluble fraction was estimated following
the addition of toluene/Triton X-100 scintillation fluid.
O
Protein--
Proteolysis was carried out in a buffer containing 20 mM Hepes/KOH (pH 7.2), 10 mM MgCl2,
10 mM ATP, 0.5% Brij 58. Each assay included 5 µg of
ClpP, 10 µg of ClpXZn(II), or ClpXZn(
), or ClpX
Cys, and 5 µg
of the
O substrate. The reaction mixtures were incubated at
30 °C, and at the desired times, 25-µl portions were withdrawn and
processed by 12.5% SDS-PAGE. The relative amount of non-hydrolyzed protein was estimated using densitometry (Bio-Rad). The data are presented in Table II.
Cys protein and
1.5 mM [
-32P]ATP in a buffer composed of
0.5 mM magnesium acetate, 15% (v/v) glycerol, 0.01%
Triton X-100, and 50 mM Tris-HCl (pH 7.8). After 15 min of
incubation at 0 °C, samples were filtrated through nitrocellulose filters, and washed with 500 µl of the above buffer. Radioactivity of
bound ATP was quantified by liquid scintillation counting (27).
Cys protein and 4 mM
ATP. The reactions were stopped by addition of 800 µl of malachite
green (0.034%) and ammonium molybdate (10.5 g/liter in 1 N
HCl) mixture and 100 µl of 34% citric acid. Absorption at 660 nm was
measured after 30 min of incubation at room temperature. Results were
compared with the calibration curve prepared for the phosphate salt
(28).
-globulin (158 kDa), chicken ovalbumin (44 kDa), equine
myoglobin (17.5 kDa). Purified GroEL was also used for the estimation
of the molecular weight of ClpP·ClpX complex.
= 6.6 × 104 M
1
cm
1 (29).
Cys,
and ClpX Zn(
) were collected as described previously (30).
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
O proteolysis assay (24), with
PMPS, known to release Zn(II) from zinc-binding proteins (30). The
formation of a mercaptide bond between the free Cys residue and PMPS
can be monitored by absorbance at 250 nm. The addition of the high
affinity metallochrome indicator PAR, which changes color (absorbance
at 500 nm) after the formation of the PAR·Zn(II) complex, allowed the
monitoring of the amount of Zn(II) released from ClpX after PMPS
treatment (Fig. 1). Control experiments, using atomic absorption, show
that treatment of ClpX with PMPS (see "Materials and Methods" for
details) leads to the complete removal of Zn(II) from the ClpX protein
(Table I).
The molecular ratio of zinc ligand complexed with ClpX protein
determined by atomic absorbance
View larger version (15K):
[in a new window]
Fig. 1.
Release of Zn(II) from ClpX following
titration with the mercurial reagent PMPS. An increasing amount of
PMPS was added to the ClpX protein (filled squares) or
ClpX Cys (open circles) both at 5 µM. The
reactants were mixed and the absorption was measured at 500 nm in the
presence of 0.1 mM PAR.
Cys, does not
bind Zn(II) (Fig. 1 and Table I). Release of Zn(II) ions from ClpX by
either PMPS or a denaturation and renaturation procedure (see
"Materials and Methods") does not lead to major changes in the
secondary structure of the ClpX protein, as judged from the CD spectra
(Fig. 2). Also, in the case of the
ClpX
Cys mutant the ClpX secondary structure is largely unaffected
(Fig. 2), suggesting that both wild type and mutant proteins fold in an
overall similar fashion. If the zinc-binding domain is found
predominantly in a
-sheet or loop structure, it could be unfolded in
the unliganded state with little effect on the CD spectrum of the
full-length protein. The application of a more sensitive approach,
namely infrared absorbance spectroscopy, suggests that the differences in secondary structure between ClpX, ClpX treated with PMPS, and ClpX
Cys mutant are indeed minor (Fig.
3). Thus, CD and IR spectroscopy cannot
detect the local conformational changes of ClpX, which should occur
following the release of Zn(II) from the ClpX protein. However, the
release of Zn(II) from ClpX partially inhibits the ATP-dependent hydrolysis of the
O protein substrate by
the ClpXP protease (Fig. 4). This partial
activity of the PMPS-treated ClpX could be due to replacement of the
zinc ligand with PMPS, which forms a tight complex with cysteine or/and
the formation of S-S bridges between the free Cys residues, which in
turn could stabilize the ClpX Zn(
) apo-protein structure, and thus
partially compensate for the loss of Zn(II). Consistent with this
interpretation, for the ClpX
Cys mutant protein the
O proteolysis
reaction is completely blocked (Fig. 4). Since the four cysteines are
substituted with serines, the stabilizing effect of PMPS or potential
disulfide bond formation should not occur. In a control experiment we
showed that the ClpXP proteolytic activity is partially inhibited by the presence of increasing concentrations of different divalent cations. It turns out that neither Mg(II) nor Co(II) ions exert an
inhibitory effect on ClpXP proteolytic activity, whereas either Ca(II)
or Zn(II) ions present in millimolar concentrations partially inhibit
ClpXP proteolytic activity (results not shown). Partial inhibition of
ClpXPs proteolytic activity in the presence of a high concentration of
Zn(II) (1 mol of ClpX to 100 or 1000 mol of Zn(II)) is probably due to
the partial ClpX precipitation under these conditions (results not
shown).
View larger version (15K):
[in a new window]
Fig. 2.
Far ultraviolet CD spectra of ClpX Zn(II),
ClpX Cys, and ClpXZn(
) proteins. Far
UV-CD spectra of 5 µM ClpX (solid line), 5 µM ClpX
Cys (dotted line), and 5 µMClpXZn(-) (dashed line) in 20 mM Tris/HCl
(pH 7.4), 150 mM KCl, 25 mM NaCl was obtained
with a Jasco-J500 CD spectrophotometer, using 1-mm thermoregulated
cuvettes at 25 °C.
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Fig. 3.
The original infrared absorbance spectrum
collected for ClpX Zn(II) (a), and the difference
spectra between ClpX Zn(II)/ClpX Zn( ) (b), or ClpX
Zn(II)/(ClpX
Cys (c).
Arrows indicate the wavenumber characteristic for the
-structure.
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Fig. 4.
Kinetics of ClpXP-mediated hydrolysis of
O protein. Proteolysis was carried out in a
buffer containing 20 mM Hepes/KOH (pH 7.2), 10 mM MgCl2, 10 mM ATP, 0.5% Brij 58. Each assay included 5 µg of ClpP, 10 µg of ClpX Zn(II)
(filled squares), ClpX Zn(
) (filled circles),
or ClpX
Cys (filled triangles) and 5 µg of the
O
substrate. The reaction mixtures were incubated at 30 °C, and at the
indicated times, 25-µl portions were withdrawn and processed by
12.5% SDS-PAGE. The amount of
O which was left after hydrolysis was
analyzed densitometrically.
O proteolytic assay shows that during the various steps of ClpX
purification there is a partial loss of ClpX activity (result not
shown). This result could be due to a partial loss of the Zn(II)
cation. To bypass this potential problem, we elaborated a new method
for ClpX purification which ended up with fully active ClpX protein
complexed with Zn(II) (see "Materials and Methods" for details).
The identity of ClpX was verified by N-terminal sequencing analysis and
Western blot analysis (results not shown). Using this new purification
procedure, we were able to obtain ~35 mg of 98% pure ClpX Zn(II)
protein starting from 30 g of ClpX overproducing bacteria.
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Fig. 5.
Binding of ClpX to the ClpP protein.
Panel A, ClpP or bovine serum albumin (50 µl of 0.01 mg/ml) in PBS buffer were first fixed onto ELISA plate wells as
previously described (4) and incubated for 1 h at room
temperature. Increasing amounts of the ClpX Zn(II) wild type protein in
buffer B (25 mM HEPES/KOH (pH 7.6), 150 mM KCl,
25 mM NaCl, 5 mM MgCl2, 1 mM DTT, 2.5% (v/v) glycerol, and 0.05% Triton X-100) were
then added in the presence (open squares) or absence
(filled squares) of 1 mM ATP. In a control
experiment, increasing amounts of ClpX Zn(II) (in the presence of 1 mM ATP) were added to the wells with bovine serum albumin
(filled circles). After incubation for 30 min at room
temperature, the ELISA plates were washed once with buffer B and three
times with PBS, both supplemented with bovine serum albumin. Following
this, 100 ml of a 1:10,000 dilution of anti-ClpX serum was added and
the plates were incubated for 2 h at room temperature. The amount
of ClpX protein retained was detected using the TMB peroxidase EIA
substrate kit (Bio-Rad). The absorbance at 490 nm was determined using
a microplate reader (Cambridge Technology). Panel B, the
same conditions as those for Panel A were used with the
exception that appropriate amounts of CpX Zn( ) apoprotein, with or
without subsequent addition of 5 µM ZnCl2
were added to the ELISA plates. Open squares and
filled squares represent the complex formation between the
CpX Zn(
) apoprotein and ClpP with and without ATP, respectively.
Filled circles and open circles show the protein
complex in the presence of 5 µM ZnCl2 and in
the absence and presence of ATP, respectively.
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Fig. 6.
Detection of a ClpP·ClpX complex using size
exclusion chromatography. Panel A represents ClpX
Zn(II) alone (40 µg) chromatographed on Superdex-200 column in the
presence of 6 mM ATP (filled symbols) or 0.2 mM ATP (open symbols). Panel B shows
that ClpP (80 µg) does not oligomerize further in the presence of
nucleotide (tested in the range 0.2-6 mM ATP).
Panels C and D represent the physical presence of
ClpX Zn(II) and ClpP in the ClpP·ClpX complex, respectively. ClpX
Zn(II) protein (40 µg) was preincubated with ClpP (160 µg) for 30 min at 30 °C in the presence of 6 mM ATP (filled
symbols) or 0.2 mM ATP (open symbols) and
injected onto a Superdex-200 column equilibrated with buffer B. After
chromatography on Superdex 200 column, 30-µl aliquots from each 1-min
fractions were loaded onto an SDS-polyacrylamide gel. The stained bands
were analyzed densitometrically using a Bio-Rad densitometer. The GroEL
(800 kDa), thyroglobulin (670 kDa), ClpP (300 kDa), bovine -globulin
(158 kDa), chicken ovalbumin (44 kDa), and equine myoglobin (17.5 kDa)
proteins served as molecular mass standards.
Cys mutant did not
form an oligomeric structure (Fig. 7).
Assuming a spherical shape of ClpX, the ClpX
Cys mutant, when
chromatographed at room temperature, behaved like a monomer (Fig. 7).
We conclude that Zn(II) and ATP (or ADP) are required for the formation
and/or stabilization of the oligomeric state of the ClpX chaperone, and that the oligomeric form is required for binding of ClpX Zn(II) to the
ClpP protease.
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Fig. 7.
Size exclusion chromatography of
ClpX Cys and ClpX Zn(II) proteins. The
ClpX
Cys (lane a) or ClpX Zn(II) (lane b)
proteins (40 µg each) were preincubated for 30 min at 25 °C in the
presence of 0.2 mM ATP and injected onto a Superdex 200 column equilibrated with buffer B supplemented with 0.2 mM
ATP (see "Materials and Methods" for details). Chromatography was
performed as described under "Materials and Methods."
Cys mutant does not
possess any detectable ATPase activity. Also, when Zn(II) is released
from ClpX by PMPS treatment, ATP hydrolysis is strongly inhibited
(Table II). This effect is most likely caused by the lack of binding of
ATP to either ClpX
Cys or ClpX Zn(
). As shown in Table II, the
ClpX
Cys mutant or ClpX wild type free of Zn(II), ClpX Zn(
) do not
bind ATP. We conclude that in the absence of zinc, the ClpX protein is
not able to bind ATP, and as a consequence not able to oligomerize or
bind to ClpP.
Binding and hydrolysis of ATP by ClpX Zn(II), ClpXCys, and ClpX
Zn(
) proteins
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
Cys, in which binding of Zn(II) is completely blocked,
the protein behaves exclusively as a monomer. The involvement of Zn(II)
in ClpX oligomerization could be due to direct protein-protein
interaction, or to Zn(II)-dependent exposure of hydrophobic
patches on the ClpX protein surface. It has been shown before for the
chaperone GroEL that the presence of Zn(II) increases the amount of
hydrophobic surface exposed (38). It has also been postulated that
zinc-binding finger motifs are directly involved in protein-protein
interactions (39).
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FOOTNOTES |
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* This work was supported by Polish State Committee for Scientific Research Grant 6P04A04219, Foundation for Polish Science Grant 15/2000, UNESCO grant, and Swiss National Foundation Grant 3147283-96.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. E-mail:
zylicz@iimcb.gov.pl.
Published, JBC Papers in Press, March 13, 2001, DOI 10.1074/jbc.M007507200
2 B. Banecki, A. Wawrzynow, J. Puzewicz, C. Georgopoulos, and M. Zylicz, unpublished observations.
3 J. Puzewicz, B. Banecki, and M. Zylicz, unpublished results.
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ABBREVIATIONS |
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The abbreviations used are: DTT, dithiothreitol; PAGE, polyacrylamide gel electrophoresis; ELISA, enzyme-linked immunosorbent assay; PMPS, p-hydroxymercuriphenylsulfonic acid; PAR, 4-(2-pyridylazo)resorcinol; HPLC, high performance liquid chromatography.
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REFERENCES |
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---|
1. | Squires, C., and Squires, C. L. (1992) J. Bacteriol. 174, 1081-1085[Medline] [Order article via Infotrieve] |
2. | Parsell, D. A., Kowal, A. S., Singer, M. A., and Lindquist, S. (1994) Nature 372, 475-478[CrossRef][Medline] [Order article via Infotrieve] |
3. |
Wickner, S.,
Gottesman, S.,
Skowyra, D.,
Hoskins, J.,
McKenney, K.,
and Maurizi, M. R.
(1994)
Proc. Natl. Acad. Sci. U. S. A.
91,
12218-12222 |
4. | Wawrzynow, A., Wojtkowiak, D., Marszalek, J., Banecki, B., Jonsen, M., Graves, B., Georgopoulos, C., and Zylicz, M. (1995) EMBO J. 14, 1867-1877[Abstract] |
5. | Levchenko, I., Luo, L., and Baker, T. A. (1995) Genes Dev. 9, 2399-2408[Abstract] |
6. | Kruklitis, R., Welty, D. J., and Nakai, H. (1996) EMBO J. 15, 935-944[Abstract] |
7. |
Pak, M.,
and Wickner, S.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
4901-4906 |
8. | Glover, J. R., and Lindquist, S. (1998) Cell 94, 73-82[Medline] [Order article via Infotrieve] |
9. |
Jones, J. M.,
Welty, D. J.,
and Nakai, H.
(1998)
J. Biol. Chem.
273,
459-465 |
10. |
Pak, M.,
Hoskins, J. R.,
Singh, S. K.,
Maurizi, M. R.,
and Wickner, S.
(1999)
J. Biol. Chem.
274,
19316-19322 |
11. |
Parsell, D. A.,
Kowal, A. S.,
and Lindquist, S.
(1994)
J. Biol. Chem.
269,
4480-4487 |
12. |
Singh, S. K.,
and Maurizi, M. R.
(1994)
J. Biol. Chem.
269,
29537-29545 |
13. | Seol, J. H., Woo, K. M., Kang, M. S., Ha, D. B., and Chung, C. H. (1995) Biochem. Biophys. Res. Commun. 217, 41-51[CrossRef][Medline] [Order article via Infotrieve] |
14. | Kessel, M., Maurizi, M. R., Kim, B., Kocsis, E., Trus, B., Singh, S. K., and Steven, A. C. (1995) J. Mol. Biol. 250, 587-594[CrossRef][Medline] [Order article via Infotrieve] |
15. | Kessel, M., Wu, W., Gottesman, S., Kocsis, E., Steven, A. C., and Maurizi, M. R. (1996) FEBS Lett. 398, 274-278[CrossRef][Medline] [Order article via Infotrieve] |
16. | Wang, J., Hartling, J. A., and Flanagan, J. M. (1997) Cell 91, 447-456[Medline] [Order article via Infotrieve] |
17. |
Grimaud, R.,
Kessel, M.,
Beuron, F.,
Steven, A. C.,
and Maurizi, M. R.
(1998)
J. Biol. Chem.
273,
12476-12481 |
18. |
Thompson, M. W.,
and Maurizi, M. R.
(1994)
J. Biol. Chem.
269,
18201-18208 |
19. |
Woo, K. M.,
Kim, K. I.,
Goldberg, A. L.,
Ha, D. B.,
and Chung, C. H.
(1992)
J. Biol. Chem.
267,
20429-20434 |
20. |
Seol, J. H.,
Baek, S. H.,
Kang, M. S.,
Ha, D. B.,
and Chung, C. H.
(1995)
J. Biol. Chem.
270,
8087-8092 |
21. |
Gottesman, S.,
Clark, W. P.,
de Crecy-Lagard, V.,
and Maurizi, M. R.
(1993)
J. Biol. Chem.
268,
22618-22626 |
22. | Schirmer, E. C., Glover, J. R., Singer, M. A., and Lindquist, S. (1996) Trends Biochem. Sci. 21, 289-296[CrossRef][Medline] [Order article via Infotrieve] |
23. | Kunkel, T. A., Roberts, J. D., and Zakour, R. A. (1987) Methods Enzymol. 154, 370-382 |
24. |
Wojtkowiak, D.,
Georgopoulos, C.,
and Zylicz, M.
(1993)
J. Biol. Chem.
268,
22609-22617 |
25. |
Ziemienowicz, A.,
Skowyra, D.,
Zeilstra-Ryalls, J.,
Fayet, O.,
Georgopoulos, C.,
and Zylicz, M.
(1993)
J. Biol. Chem.
268,
25425-25431 |
26. | Roberts, J. D., and McMacken, R. (1983) Nucleic Acids Res. 11, 7435-7452[Abstract] |
27. |
Hupp, T. R.,
and Kaguni, J. M.
(1993)
J. Biol. Chem.
268,
13128-13136 |
28. | Lanzetta, P. A., Alvarez, L. J., Reinach, P. S., and Candia, O. A. (1979) Anal. Biochem. 100, 95-97[Medline] [Order article via Infotrieve] |
29. |
Hunt, J. B.,
Neece, S. H.,
Schachman, H. K.,
and Ginsburg, A.
(1984)
J. Biol. Chem.
259,
14793-14803 |
30. |
Banecki, B.,
Liberek, K.,
Wall, D.,
Wawrzynow, A.,
Georgopoulos, C.,
Bertoli, E.,
Tanfani, F.,
and Zylicz, M.
(1996)
J. Biol. Chem.
271,
14840-14848 |
31. |
Smith, C. K.,
Baker, T. A.,
and Sauer, R. T.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
6678-6682 |
32. | Maurizi, M. R. (1992) Experientia 48, 178-201[Medline] [Order article via Infotrieve] |
33. | Bochtler, M., Hartman, C., Song, H. K., Bourenkov, G. P., and Huber, R. (2000) Nature 403, 800-805[CrossRef][Medline] [Order article via Infotrieve] |
34. |
Berg, J. M.
(1990)
J. Biol. Chem.
265,
6513-6516 |
35. | Haupt, Y., Alexander, W. S., Barri, G., Klinken, S. P., and Adams, J. M. (1991) Cell 65, 753-763[Medline] [Order article via Infotrieve] |
36. | Niki, H., Imamura, R., Kitaoka, M., Yamanaka, K., Ogura, T., and Hiraga, S. (1992) EMBO J. 11, 5101-5109[Abstract] |
37. | Mitterauer, T., Nanoff, C., Ahorn, H., Freissmuth, M., and Hohenegger, M. (1999) Biochem. J. 342, 33-39[CrossRef][Medline] [Order article via Infotrieve] |
38. |
Brazil, B. T.,
Ybarra, J.,
and Horowitz, P. M.
(1998)
J. Biol. Chem.
273,
3257-3263 |
39. |
Kuroda, S.,
Tokunaga, C.,
Kiyohara, Y.,
Higuchi, O.,
Konishi, H.,
Mizuno, K.,
Gill, G. N.,
and Kikkawa, U.
(1996)
J. Biol. Chem.
271,
31029-31032 |