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INTRODUCTION |
The mitochondrial enzyme rhodanese (EC 2.8.1.1) catalyzes the
transfer of a sulfur atom from a sulfur donor, e.g.
thiosulfate to an acceptor, e.g. cyanide. In the course of
catalysis, the enzyme cycles between a form containing a transferred
sulfur held as a persulfide (RSSH) on active site cysteine 247 and a
form without the transferred sulfur (1). Rhodanese is a monomeric protein, and its crystal structure is known (2-4). The protein is
folded into two domains that are independently folded but tightly coupled at the interdomain interface. The active site is in the C-terminal domain close to the interdomain interface, and domain separation is associated with inactivation. There are four cysteine residues at sequence positions 63, 247, 254, and 263, and all are
reduced in the native enzyme. The persulfide-containing form of wild
type rhodanese, WT-ES,1 and
rhodanese without the transferred sulfur, WT-E, have virtually identical crystal structures (2, 5, 6). Site-directed mutagenesis shows
that Cys-247 is the only cysteine residue essential for rhodanese
activity (7, 8). Although it is clear from the crystal structure that
none of the cysteine residues in native rhodanese is in a favorable
position to form disulfide bonds, Cys-247 always forms disulfide bonds
with other cysteines in the C-terminal domain during oxidative
inactivation, to which rhodanese is particularly sensitive. These
disulfides can form from the native state during oxidative inactivation
by reagents such as phenylglyoxal in a process that obviously requires
a conformational change. In addition, single site oxidation at the
active site can form a sulfenic acid or higher oxidation states. These
oxidative events are the major reactions that result in incomplete
reactivation of the enzyme after denaturation. The mutagenesis of the
three nonessential cysteine residues as studied here (C63S, C254S, and C263S) to produce the species C3S removes the possibility of
intramolecular disulfide bonds and makes the enzyme an important model
for studies of folding.
Rhodanese has been one of the most studied substrates for assisted
folding by the chaperonin GroEL. Using the wild type enzyme, rhodanese
only binds easily to GroEL when it is presented to the chaperonin after
it has been extensively unfolded. The native enzyme does not interact
with GroEL (9).
The results presented here, based on studies using controlled
proteolysis and mass spectrometry, suggest that the C-terminal domain
of the persulfide-containing C3S (C3S-ES) is considerably less stable
than WT-ES although the specific enzyme activities are the same.
Although it is known that significant binding to GroEL requires forms
of rhodanese that are unfolded further than states with folded domains,
no specific information is known about these less folded forms. Because
it was found that active C3S-ES is less stable than the wild type
enzyme, we compared the properties of the form of C3S lacking
transferred sulfur (C3S-E) and rhodanese (WT-E) with respect to GroEL
binding. The results suggest that active C3S with an unstable
C-terminal domain exists in equilibrium with the conformer(s),
accessible from the native state that can interact with GroEL. This
contrasts with the lack of binding observed with the wild type enzyme.
Thus, we have identified a particular region of C3S that is correlated
with its decreased stability and increased binding to GroEL. These
results present an opportunity for studying the conformational
correlates required for substrate binding to GroEL.
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MATERIALS AND METHODS |
Reagents--
SDS, acrylamide, and bisacrylamide were from
Fisher. Bicinchoninic acid protein assay reagent was purchased
from Pierce. Other chemicals were from Sigma. Sequencing grade,
modified trypsin (reductively methylated) was from Promega (Madison, WI).
Rhodanese Purification--
Recombinant bovine rhodanese was
purified as described previously and stored at
70 °C as a
crystalline suspension in 1.8 M ammonium sulfate containing
1 mM sodium thiosulfate (10). Rhodanese was desalted on a
G-50 column before use. Rhodanese concentration was determined using a
value of A0.1% 280 nm = 1.75 (11).
Preparation of E Rhodanese--
Rhodanese as prepared is in the
sulfur-substituted form. The E form was prepared by treating the enzyme
with a 5-fold molar excess of cyanide (12).
Rhodanese Assay--
Rhodanese activity was measured using a
colorimetric method based on the absorbance at 460 nm of the complex
formed between the reaction product, thiocyanate, and ferric ion
(11).
Construction and Isolation of C3S--
The rhodanese gene (1.1 kilobases) was cloned between the NcoI and BamH1
sites of Pet-11d. Cysteines (a) 63, (b) 254, and (c) 263 were
sequentially mutated to serines using the QuikChange site-directed
mutagenesis kit (Stratagene, La Jolla, CA). The corresponding mutagenic
oligos were (a) 5'-GGCCTTCAGCCGTGACTCCTCTATCTG-3', (b) 5'-CAGGGCAATGTGGGAGGCGGTGACACC-3', and (c)
5'-ATCGGGCTTGCCTGAGAGGTAAGCAGC-3'. BL21(DE3) competent cells (Novagen,
Madison, WI) were transformed with the mutant DNAs, and the DNA was
purified by miniprep (Qiagen, Valencia, CA). DNA sequencing was
performed by the Center For DNA Technology, University of Texas Health
Science Center, San Antonio, TX. Once the first change was verified,
that DNA was used for the next step of mutagenesis. The T-7 universal
primer was used to sequence into the 5'-end of the rhodanese insert. Likewise, a sequencing primer (5'-ACACAGCCGGAGCCAGATGCAGTA-3' was used
to sequence from the middle of the insert past the C terminus. There
were no unintended changes evident in the primary sequence. C3S was
purified essentially by the procedure used for WT with the exception
that the cells were grown at 25 °C, and the purification steps were
performed at 4 °C.
GroEL and GroES Purification--
GroES7 and
GroEL14 were purified as described previously (13, 14).
Protein concentration was determined by the BCA method (15).
Proteolysis of Rhodanese--
Rhodanese (0.2 mg/ml) in 40 mM NH4HCO3, pH 7.5, was treated
with trypsin (3.75% w/w) at 25 °C for the times indicated under "Results." At each time, 2.5 µl of 200 mM
phenylmethanesulfonyl fluoride and 5 µl of 4× SDS (0.25 M Tris.HCl, pH 6.8, 40% glycerol, 8%
-ME, 8% SDS, 5%
bromphenol blue) sample solution were added to 12.5 µl (2.5 µg) of
the incubating sample, and the resulting solution was boiled for 2 min.
SDS Gel Electrophoresis--
SDS gels shown in Fig. 4,
A and B, were run by the method of Laemmli (16)
using 12.5 and 4% acrylamide in the separating and stacking gels,
respectively. The SDS gels shown in Fig. 4C were run using a
10% Tricine system with a 10% separating gel and a 4% stacking gel
(17). The band intensities were scanned using the program Scion Image
for Windows downloaded from Scion Corp., Frederick, MD.
Mass Spectrometry--
Peptides for mass spectrometry were
prepared from SDS gels by a method developed by Christopher Carroll and
Dr. Susan Weintraub (University of Texas Health Science Center at San
Antonio). Briefly, the gel band of interest was excised from a
Coomassie Brilliant Blue-stained gel. The gel piece was destained with
3 washes using 0.4 ml of 50% acetonitrile in 25 mM
ammonium bicarbonate buffer, pH 8,0. The gel band was then dehydrated
by soaking the piece in 100 µl of 100% acetonitrile until the gel
piece turned opaque. The gel piece was dried in a Speed-Vac and
rehydrated with a volume of trypsin solution (Promega modified
sequencing grade; 10 µg/ml in 25 mM
NH4HCO3 buffer, pH 8.0) sufficient to swell the
gel piece with no excess (10-20 µl). 25 mM
NH4HCO3, 0.02% Zwittergent was added to just
cover the gel piece, and the sample was incubated at 37 °C for
16-24 h. The reacted digestion samples were centrifuged, 10 µl of
5% trifluoroacetic acid were added, and the sample was sonicated and
then centrifuged. The gel pieces were further treated with 2.5%
trifluoroacetic acid, sonicated and centrifuged. The extracted peptides
were pooled. Typically, 1 µl of the digest was used for analysis by
matrix-assisted laser desorption ionization.
Matrix-assisted laser desorption ionization time-of-flight mass spectra
were acquired on an Applied Biosystems Voyager DE-STR using the
matrices sinapinic acid (intact proteins) and
-cyano-4-hydroxycinnamic acid (tryptic digests).
HPLC-electrospray ionization tandem mass spectra were acquired on a
Finnigan LCQ ion trap mass spectrometer. Samples were separated
by on-line HPLC using a Michrom BioResources MAGIC 2002 micro-HPLC
fitted with a PicoFrit capillary column (75-µm inner diameter × 7 cm, 5-µm C18; New Objective). The mobile phase was 0.5% acetic
acid, 0.005% trifluoroacetic acid (A) and 90% acetonitrile, 0.5%
acetic acid, 0.005% trifluoroacetic acid (B), with a flow rate of 0.5 µl/min. Data were acquired in a data-dependent mode, with
an initial survey full mass scan followed by two tandem mass spectra. Data interpretation was accomplished by means of the SEQUEST
software component on the LCQ.
N-terminal Amino Acid Sequence--
C3S was proteolyzed as
before and subjected to 12.5% SDS-PAGE (e.g. Fig. 5). The
protein bands were transferred to Immobilon-PSQ transfer
membrane (Millipore Corp., Bedford, MA) using a Semi-Phor Apparatus
(Hoefer Scientific Instruments, San Francisco). The membrane was
briefly stained with 0.1% Amido Black, 10% methanol and then
destained with 10% acetic acid, 10% methanol. The desired band (20 kDa, Fig. 5, band B shown with an arrow) was cut
and sequenced on a Procise 492-cLC Protein Sequencer (Applied
Biosystems, Foster City, CA).
Acrylamide Quenching of WT-E and C3S-E--
For quenching, a
stock of 1 M acrylamide was made in 50 mM
Tris-Cl, 10 mM KCl, 10 mM MgCl2, pH
7.8 (folding buffer). The absorbance at 295 and 340 nm were measured
for each data point and used to correct the fluorescence for the inner
filter effect. The Stern-Volmer equation,
F0/F = 1 + KSV [Q], was used for analysis. In this
equation, F0 and F are the
fluorescence in the absence and presence of quencher, respectively,
[Q] is the quencher concentration, and KSV is the Stern-Volmer constant, which correlates with the accessibility of the
fluorescence for quenching (19).
Gel Filtration--
Gel filtration was performed to separate
monomeric WT-E and C3S-E and oligomer, formed during
incubation. 150 µl of 3 µM WT-E and C3S-E in 50 mM Tris-Cl, 10 mM KCl, 10 mM
MgCl2, pH 7.8 (folding buffer), were incubated at 25 °C
for 60 min and loaded on a 15-ml (19 × 1 cm) Sephacryl S-100-SR
column (fractionation range 1,000-100,000), equilibrated with 50 mM Tris-Cl, 10 mM KCl, 10 mM
MgCl2, pH 7.8 at 25 °C. 100-µl fractions were
collected and assayed for protein concentration and activity. Two
column volumes of buffer were used for the elution. The same experiment
was done at 25 °C using 0.1 mg/ml rhodanese (33 kDa) and 0.1 mg/ml
bovine serum albumin (66 kDa) for the calibration.
CD Measurements--
CD spectra were scanned at 25 °C from
250 to 200 nm for far UV data using an OLIS DSM 16 UV-visible CD
spectropolarimeter (On Line Instrument Systems, Inc., Bogart,
GA). All samples were in 50 mM
Na2HPO4/NaH2PO4, pH
7.5, at 25 °C. Protein concentration was at 0.1 mg/ml, and the path
length of the cell was 0.1 cm. Data were collected at 0.5-nm intervals.
Appropriate blanks were subtracted from the observed spectra. CD data
were calculated in terms of molar ellipticity [
] (20-22) at each
specified wavelength using a protein molecular weight of 33,000. The
molar ellipticity [
] is expressed as degrees cm2
dmol
1.
Fluorescence Titration of GroEL with WT-E and C3S-E--
The
equilibrium binding of E forms of both WT and C3S rhodanese was studied
using a fluorescence titration method described in the literature (23).
The reaction conditions were similar to the experiments described for
the inactivation studies except that the solutions contained 1 mM ADP (see the legend for Fig. 9). A Fluorolog-3 (Jobin
Yvon-Apex) spectrofluorometer was used. The excitation was at 295 nm
(path length 5 mm, band pass 5 nm), and emission was recorded in the
range of 300-450 nm (path length 2.5 mm, band pass 5 nm). In a 1-ml
cuvette containing either WT or C3S (1 µM), the E form of
rhodanese was formed by adding 5 µM KCN. After
equilibration, 1-5-µl aliquots of a concentrated and
fluorescence-free GroEL were added from a 217 mg/ml stock. The
solutions were equilibrated for another 5 min before the emission spectra were recorded. The titration was continued until the final concentration of GroEL reached about 5 µM. In an
identical experiment the scattering contributions from GroEL alone were
studied by titrating it into the buffer without rhodanese. The overlaid
emission scans, after the subtraction of the GroEL contribution, show
maximum changes at 333 nm for both WT-E and C3S-E. These fluorescence intensity changes (
F) at different GroEL concentrations
could be fitted to a binding model(s) to obtain the equilibrium binding constant, Kd (23). Further details on the models
used and their limitations are presented in the literature (23).
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RESULTS |
Expression and Purification of Active, Soluble C3S Requires Lowered
Temperatures--
When expression of C3S was attempted using growth
conditions (37 °C) suitable for wild type rhodanese, very little
enzyme was detected in the extracts. Almost all of the enzyme was
contained in the pellets as demonstrated by SDS gels (data not shown),
and attempts to solubilize and activate this enzyme were unsuccessful. When the cells were grown at 25 °C, a large amount of active enzyme could be recovered in the extracts. Attempts to purify this enzyme using procedures at room temperature as for the wild type enzyme led to
enzyme inactivation, and very little active purified C3S was recovered.
Significant recoveries of C3S were achieved when the purification steps
were carried out at 4 °C. The purified C3S had a specific activity
of 760 IU/mg and was, in this regard, very similar to the wild type protein.
C3S Is Less Stable Than Wild Type Rhodanese--
The growth and
purification conditions suggested that C3S is less stable than the wild
type protein. This expectation is strengthened by the results shown in
Fig. 1, where the effect of urea on the activities of WT and C3S is compared. The activity of C3S is very sensitive to urea, and there is a decrease in activity with even the
smallest addition of urea. The C3S loses half its activity at ~1.75
M urea, whereas WT loses half its activity at ~4.3
M urea.

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Fig. 1.
Activity of C3S-ES ( )
and WT-ES ( ) rhodanese as a function of urea concentration.
Rhodanese was incubated with the desired concentrations of urea for
4 h. The reaction conditions were [protein] = 300 µg/ml,
Na2HPO4/NaH2PO4 = 0.2 M, pH 7.5, and T = 25 °C. Enzyme
activity was measured as described under "Materials and
Methods."
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Comparison of Secondary Structures of C3S and Wild Type
Rhodanese--
Fig. 2 shows the circular
dichroism spectra for C3S and WT rhodanese. As can be observed, there
are only small differences in these spectra. Thus, although there are
significant differences in the apparent stabilities of these proteins,
their secondary structures are identical within experimental error.

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Fig. 2.
Near UV circular dichroism spectra of C3S-E
and WT-E rhodanese. CD spectra in the far UV region were scanned
using a 0.1-cm path length cell at 0.5-nm intervals. The conditions
were [protein] = 100 µg/ml,
Na2HPO4/NaH2PO4 = 0.05 M, pH 7.5, and T = 25 °C. Other details
are under "Materials and Methods."
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Accessibility of Tryptophan Residues in WT-E and C3S-E toward
Collisional Quenching--
Fig. 3 shows
the acrylamide quenching of C3S-E and WT-E. The Stern-Volmer constants
(KSV) are 1.073 ± 0.02 M
1 and 1.481 ± 0.04 M
1, respectively. The higher Stern-Volmer
constant indicates that the tryptophan residues in C3S-E are more
exposed than those in WT-E. There is no significant curvature over the
acrylamide concentration range used, which suggests that there is no
conformational change induced by acrylamide that is associated with
changes in tryptophan exposure. These data are consistent with the view
that C3S-E is less stable than WT-E.

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Fig. 3.
Acrylamide quenching of tryptophan residues
in C3S-E and WT-E. 0.1 µM C3S-ES ( ) and 0.1 µM WT-ES ( ) were incubated in 50 mM
Tris-Cl, 10 mM KCl, 10 mM MgCl2, pH
7.8, containing 0.5 µM KCN for 45 min. Acrylamide
solution in 50 mM Tris-Cl, 10 mM KCl, 10 mM MgCl2, pH 7.8 was added, and tryptophan
fluorescence was measured. The excitation was set at 295 nm, and
emission was at 340 nm. 2-nm band passes were kept for both excitation
and emission.
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Tryptic Digestion of C3S Leads to a Resistant Fragment Not Observed
with Wild Type Rhodanese--
Figs. 4
and 5 show the comparison between the
time courses of digestion of WT and C3S proteins, respectively, with
trypsin. Panel A shows that the WT enzyme in the presence of
thiosulfate is not susceptible to digestion at the level of trypsin and
the conditions that are used. Panel B shows that C3S, in the
presence of thiosulfate, is not significantly digested by trypsin. WT
enzyme in the absence of thiosulfate is similarly unaffected
(panel C). In contrast to these results, when C3S is treated
with trypsin in the absence of thiosulfate (Fig. 5) there is a
progressive digestion of the parent protein with the appearance of a
band at an apparent molecular weight of 21,000 that remains as an
apparently stable product under these conditions. The amino acid
sequence of the daughter band (Fig. 5, band B) was
determined as discussed under "Materials and Methods." The sequence
corresponded to the N-terminal region of rhodanese, confirming that the
clip occurred in the C-terminal domain. This result is in agreement
with the sequence of C3S bands identified by mass spectra (Fig. 7).

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Fig. 4.
SDS-PAGE of C3S-ES and WT-ES proteolysis
products in the absence and presence of thiosulfate. Proteolysis
of C3S-ES and WT-ES rhodanese were performed as described under
"Materials and Methods." Panel A, digestion of WT-ES in
the presence of 50 mM
Na2S2O3. Lane 1 shows
molecular mass marker with bands at 97.4, 66.2, 45.0, 31.0, and 21.5 kDa, top to bottom. Lanes 2-8
are the products of proteolysis at 0, 5, 10, 15, 20, 25, and 30 min,
respectively. Lane 9 was the trypsin control (24 kDa).
Panel B, digestion of C3S-ES in the presence of 50 mM Na2S2O3. Lane
1 shows molecular mass marker bands, and lanes 2-8
were products of proteolysis at the times described for panel
A. Panel C, digestion of WT rhodanese in the absence of
Na2S2O3. Lane 1 shows
molecular mass marker bands, and lanes 2-8 are the
products of proteolysis at 0, 5, 10, 15, 20, 25, and 30 min,
respectively. Lane 9 was the trypsin control (24 kDa).
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Fig. 5.
C3S-ES in the absence of
Na2S2O3. Lane 1 is the
molecular mass marker with bands at 97.4, 66.2, 45.0, 31.0, 21.5, and
14.4 kDa top to bottom. Lane 2 is the trypsin control. Lanes 3-11 are proteolysis products
at 0, 2.5, 5, 10, 15, 20, 25, 30, and 35 min, respectively. Band
A is the parent band (33 kDa), and band B is the daughter
band.
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Fig. 6 shows the activity of C3S in the
absence of thiosulfate and in the presence and absence of trypsin. In
the absence of trypsin (upper curve, solid
circles), the activity is stable for at least 30 min. However,
with trypsin, the activity falls in a time course that is compatible
with the digestion pattern seen on the SDS gels (lower
curve: solid triangles indicate density of parent band;
solid squares indicate rhodanese activity). This indicates
that C3S is stable in the absence of trypsin and that the activity loss
follows digestion rather than the digestion following inactivation of
the protein to a susceptible form.

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Fig. 6.
Effect of proteolysis on the enzyme activity
of C3S-ES. C3S-ES (200 µg/ml) was proteolyzed with trypsin (7.5 µg/ml) in the presence of 50 mM
Na2S2O3 ( ) and in its
absence ( ). The enzyme activity was measured as a function of time
(see "Materials and Methods"). The data represented by the symbol
are the relative intensities of the major band (33 kDa) from an
identical proteolysis experiment in the absence of
Na2S2O3, shown in Fig. 5
(lanes 3-11).
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Mass Spectrometry Indicates That the C-terminal Domain of C3S Is
Proteolytically Susceptible--
In-gel digestion, peptide extraction,
and mass spectrometry were performed as described under "Materials
and Methods" on the 21-kDa daughter band formed by tryptic digestion
as in Fig. 5. Fig. 7 shows the amino acid
sequences of fragments that are detected from the parent band and the
daughter band after digestion. The total molecular weight of the C3S
daughter fragment from mass spectrometry was determined to be 20,783, which is consistent with the migration of the daughter fragment in the
SDS gels. Fragments were observed from both bands that correspond to
peptides in the N-terminal domain. No fragments were detected in the
daughter band beyond residue Arg-182, although large C-terminal
fragments were detected from the parent band. The single band from WT
with or without thiosulfate displayed the same pattern as the parent band from C3S. It is clear that the stable daughter band that remains
on the gel in Fig. 5, band B, results from cleavages within the C-terminal domain. Thus, the digestion occurs within the C-terminal domain of C3S in the absence of thiosulfate. These results suggest that
the C-terminal domain of C3S is less stable than the corresponding region of WT.

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Fig. 7.
Sequence of the proteolytic products of C3S
by Mass Spectrometry. The amino acid sequences of C3S parent band
and C3S daughter band after limited proteolysis by trypsin. Sample
preparation, acquisition of mass spectra, and sequence analysis are
discussed under "Materials and Methods." Underlined
portions of the sequences represent fragments detected by mass
spectrometry.
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Binding of Sulfur-loaded Forms of C3S (C3S-ES) and Rhodanese
(WT-ES) to GroEL--
Fig. 8,
panel A, shows the activity of C3S-ES alone and in the
presence of GroEL in folding buffer. The activity of C3S-ES remains
unchanged at least up to 4 h at 25 °C (Fig. 8, panel
A, filled squares). However, the enzyme is stable up to
1 h in the presence of GroEL. After that, there is a small but
significant inactivation (~15%) in the presence of GroEL (Fig. 8,
panel A, solid circles). This loss in activity
may be due to the fact that in dilute solution, less ordered C3S-ES
equilibrated slowly to a population of conformers, some of which can
interact with GroEL, and the unfolding activity of GroEL makes
the protein inactive. In the absence of GroEL, these conformers are in
equilibrium with the native form and show 100% activity in the assay
mixture. Fig. 8, panel B, shows the same experiment with
WT-ES. Both in the absence and presence of GroEL, the enzyme remains
100% active, which indicates that WT-ES is stable in solution and does
not form any conformation that can be recognized by GroEL under the conditions used here (24). These data are consistent with the idea that
the structure of C3S-ES is more labile in solution than WT-ES, so that
it can form more than one kinetically accessible state from the native
enzyme.

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Fig. 8.
Inactivation of C3S-ES and WT-ES in the
presence and absence of GroEL. 0.1 µM C3S-ES
(A) and 0.1 µM WT-ES (B) were
incubated in 50 mM Tris-Cl, 10 mM KCl, 10 mM MgCl2, pH 7.8, in the absence ( ) and
presence ( ) of 0.2 µM GroEL at 25 °C.
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Binding to GroEL of C3S Lacking Transferred Sulfur (C3S-E) and
Rhodanese Lacking Transferred Sulfur (WT-E)--
Fig.
9 shows the same experiment as done in
Fig. 8 with C3S-E and WT-E in folding buffer. Unlike C3S-ES and WT-ES,
the E forms of both the proteins are unstable in solution and lose
activity with time. Fig. 9, panel A, shows that C3S-E
inactivates in solution with a rate constant of 0.08 ± 0.008 min
1 (t1/2 = 8.7 min). But the rate of inactivation becomes more than 2 times faster in the presence of GroEL,
with a rate constant of 0.18 0 ± 0.004 min
1
(t1/2 = 3.85 min). The faster rate of inactivation
in the presence of GroEL suggests that C3S-E may form a less ordered
conformer(s) that can be captured by GroEL. Fig. 9, panel B,
shows that WT-E is also unstable in solution. However, in this case
both in the presence and absence of GroEL it inactivates with a similar
rate as C3S-E, with a rate constant of 0.07 ± 0.003 min
1 (t1/2 = 9.9 min). The observation that the rate of inactivation both in the presence and absence of GroEL
are virtually identical suggests that during inactivation WT-E does not
form any conformation that can be captured by GroEL. The inactivation
of C3S-E and WT-E may be due to intermolecular (for both C3S-E and
WT-E) and intramolecular (for WT-E) disulfide formation. However, the
SDS-PAGE results (without
-ME) of the inactivated C3S-E and WT-E
without GroEL show only a single band corresponding to a 33-kDa
molecular mass (data not shown). Similar experiments with both C3S-E
and WT-E were done where excess cyanide was removed by gel filtration,
and they showed a similar profile. Attempts to isolate oligomeric
species from the above samples by gel permeation chromatography also
yielded only monomers for both C3S-E and WT-E (data not shown). These
data indicate that the change in conformation leading to inactivation
is not due to any intermolecular covalent modification of either C3S-E
or WT-E.

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Fig. 9.
Inactivation of C3S-E and WT-E in the
presence and absence of GroEL. 0.1 µM (A)
C3S-ES and 0.1 µM WT-ES (B) were incubated in
50 mM Tris-Cl, 10 mM KCl, 10 mM
MgCl2, pH 7.8, containing 0.5 µM KCN in the
absence ( ) and presence ( ) of 0.2 µM GroEL at
25 °C.
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Reactivation of Inactivated C3S-E and WT-E by the GroEL Chaperonin
System--
C3S-E and WT-E inactivated in the absence or in the
presence of GroEL, as in Fig. 9, were incubated further to get maximum inactivation, and then their reactivation was studied. Table
I (first line) shows that all the
incubated samples were almost inactive. When both thiosulfate (50 mM) and
-ME (0.2 M) were added to these
inactivated samples, only a small amount of reactivation (~10-20%)
was observed from all samples. The small reactivation in the presence
of high concentrations of reductant also shows that disulfide bond
formation was not the predominant factor for inactivation. When
GroEL/GroES/ATP were added to those samples, significant reactivation
was observed with all the samples. GroEL could capture inactivated
C3S-E and reactivate to 100% in the presence of GroES and ATP (Table
I). This is the highest reactivation of inactivated rhodanese that has
been observed with this enzyme. Reactivation to ~70.4% was also
observed with C3S-E, where the inactivation was done in the presence of
GroEL. Similar results were obtained with WT-E, where 72% reactivation
was noted. The complete reactivation of C3S-E indicates that it can
form a conformer in solution that is capable of more productive binding
to GroEL. This very high reactivation clearly shows that inactivation
is primarily due to non-covalent conformational change in the
protein.
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Table I
Chaperonin-assisted reactivation of WT-E and C3S-E after inactivation
in solution
0.1 µM rhodanese and C3S were taken in 50 mM
Tris-Cl, 10 mM KCl, 10 mM MgCl2, pH
7.8, with and without 0.2 µM GroEL14. The samples
were incubated in the presence of 0.5 µM KCN for 45 min
at 25 °C, and the activities were measured. 50 mM
thiosulfate and 0.2 M -ME were added and incubated at
25 °C for 30 min, and the activities were measured. GroEL and GroES
were added so that each sample contained 0.2 µM
GroEL14 and 0.5 µM GroES7. Finally, 5 mM ATP was added and incubated at 25 °C for 120 min. The
protein activity was calculated, taking native protein in the same
buffer containing all the components described above except KCN, as
100%. % activities after the noted treatments.
|
|
GroEL binding of the E forms of rhodanese was investigated using
titration of GroEL into a fixed amount WT-E and C3S-E using fluorescence ("Materials and Methods"). The aim was to understand if the better reactivation of C3S-E relative to WT-E could be due to
the increase in the population of specific sites that have higher
affinity for the GroEL protein. The decrease in
F upon the increase in GroEL concentrations follows a binding curve in the
case of WT-E (data not shown) with a binding constant
(Kd) of 0.20 ± 0.01 µM. The
results for C3S did not follow a binding curve in the GroEL
concentrations studied (data not shown). Therefore, contrary to the
better inactivation of C3S-E than WT-E in the presence of GroEL (Fig.
9), the fluorescence titration apparently showed the reverse. However,
it must be noted that the fluorescence changes were not significant and
probably are not better than the results obtained from activity studies
(Fig. 9).
 |
DISCUSSION |
One of the major side reactions that limits the folding of
denatured rhodanese is the formation of intramolecular disulfide bonds
involving the active site cysteine residue and the nonessential sulfhydryl groups (25, 26). According to the x-ray
structure, the cysteines are not close enough to form these disulfides.
This implies flexibility in solution that would permit the observed disulfide formation. We find in this study that the removal of the
three nonessential cysteine residues (cysteines 63, 254, and 263) to
form C3S makes rhodanese less stable in its C-terminal domain, although
the specific activities at 25 °C and the average secondary
structures reported by CD are the same for the mutant and WT. In
addition, C3S is protected against proteolysis by the presence of the
substrate, thiosulfate.
The x-ray structure of WT rhodanese permits us to speculate as to the
relation between the mutations prepared here and the consequences that
are observed. Rhodanese is folded into two separate globular domains
(N-terminal domain consisting of residues 1-142 and C-terminal domain
consisting of residues 159-293) that are tightly coupled at an
interdomain interface by numerous hydrophobic interactions. The domains
are covalently connected by a tether consisting of a long loop of
residues 143-158 that interacts with the N-terminal domain. The two
domains are of nearly equal size, and they have very similar
conformations. Thus, it is reasonable that the N-terminal domain is
more stable than the C-terminal domain due to the additional
interactions with the tether. The active site Cys-247 that holds the
sulfur transferred from thiosulfate is in the interdomain region, and
it has been shown that the activity of the enzyme is sensitive to
conditions that disrupt the interdomain interactions. Numerous attempts
at preparing individual domains have been unsuccessful, presumably due
to the extensive hydrophobic surfaces that would be exposed on the
individual domains.
Of the nonessential sulfhydryl groups, Cys-254 has been shown to be the
most influential at affecting the structure and function of rhodanese
(3, 4, 6). This is reasonable since Cys-63 is in the N-terminal domain,
and the main effects in C3S are in the C-terminal domain. Cys-263 has
been shown to have smaller effects, which is reasonable since it is
close to the surface of the protein. Helices from each of the domains
(D' helix in the C-terminal domain, residues 251-264, and the D helix
in the N-terminal domain, residues 107-119) cross each other at
Cys-254, which appears to interact with Trp-113 and Pro-109 from the
N-terminal domain. A number of residues from the D and D' helices
interact hydrophobically to stabilize the interdomain interface. Thus, the mutation at Cys-254 can affect the interdomain surfaces or their
interactions. Changes in this contact region can affect the domain
structure and permit the proteolytic clip that occurs within the
C-terminal domain to leave a stable fragment that ends at Arg-182.
The proteolysis is interesting. Arg-182, the final cleavage point, is
protected from proteolytic access by a long turn in the structure
consisting of residues 189-211 (4). Movement of this turn would permit
proteolytic access, and the unstructured nature of the turn would make
changes in its orientation very difficult to detect by CD.
Interestingly, Arg-186 separates this loop from bond 182-183. Arg-186
is the cationic binding site for the substrate thiosulfate. Thus,
thiosulfate binding can easily be envisioned as stabilizing the
structure or covering Arg-182, thus preventing proteolytic access.
GroEL does not bind native rhodanese in the form containing transferred
sulfur, as studied by activity and direct measurement of complex
formation (27, 28). It has been established that for productive binding
to GroEL and its reactivation in the presence of GroES/ATP, rhodanese
domains must be unfolded (28). The rhodanese form containing
transferred sulfur, WT-ES, does not bind to GroEL, as there is no loss
in activity in the presence of GroEL. The sulfur-loaded form of mutant
rhodanese, C3S-ES, shows the same specific activity as WT-ES and
remains fully active in solution. However, it shows slow but
significant inactivation in the presence of GroEL. These data are
consistent with the view that C3S-ES has a less stable structure than
WT-ES and, thus, is able to exist in an equilibrium with an
intermediate(s) that can bind to GroEL. Similar observations have been
made with dihydrofolate reductase in the absence of substrates (29, 30)
and pre-
-lactamase (31), where starting with the native protein,
stable complexes can be formed with GroEL. Mitochondrial aspartate
aminotransferase forms a modified but catalytically active conformation
that can be bound to GroEL (18). Although the sulfur-loaded forms of both rhodanese and C3S are stable in solution, the forms lacking transferred sulfur inactivate readily. The inactivation may be due to
the formation of inter- and intramolecular disulfide-bonded species.
The addition of high concentrations of reductant shows insignificant
reactivation. Only monomeric species are detected on non-reducing
SDS-PAGE and on gel permeation chromatography. All these data rule out
the possibility of disulfide-linked species as a cause of inactivation
of C3S-E. C3S-E inactivates at a faster rate when co-incubated with
GroEL. The faster rate of inactivation may be due to the shift of
equilibrium between the native state and the conformer(s) recognizable
by GroEL. The addition of GroEL/GroES/ATP to C3S-E after complete
inactivation generates 100% active protein. WT-E does not show any
additional inactivation in the presence of GroEL, which indicates that
GroEL cannot bind the intermediates formed during inactivation. The
addition of GroEL/GroES/ATP to inactivated samples leads to a maximum
of ~72% reactivation with WT. The incomplete reactivation may be due
to the fact that either GroEL cannot capture all the conformers formed
or cannot reactivate all of those to active protein. The greater
accessibility of the tryptophan residues in C3S-E suggests that that
they are more exposed, and it is consistent with the protein having a
more labile structure than WT-E. This loss in compactness in the
structure of C3S is reflected in more productive binding to GroEL. All
these data suggest that C3S has less compact structure in its
C-terminal domain, and it is this domain that preferentially binds to
GroEL. Importantly, the differential stability of C3S is induced by the binding of the substrate, thiosulfate. These studies will provide a
means to identify and characterize the binding determinants on protein
targets for GroEL.