From the Department of Chemistry, National
Tsing Hua University, Hsinchu 300, the § Department of
Chemistry, National Changhua University of Education, Changhua 500, and the ¶ Department of Chemistry, National Taiwan University,
Taipei 106, Taiwan
Received for publication, July 6, 2000, and in revised form, October 18, 2000
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ABSTRACT |
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The refolding kinetics of the 140-residue, all
It is still unknown as to how a polypeptide chain folds into a
unique structure out of an astronomical number of energetically possible conformations in a short period of time (1-3).
Characterization of the conformational properties of the early stages
sampled during refolding are crucial to our understanding the rules
determining the folding process (4-8). Much of our knowledge on the
kinetics of refolding/unfolding is derived employing the stopped-flow
optical spectroscopic methods or the quenched-flow hydrogen/deuterium exchange techniques (9-14). By using these techniques, several proteins have been shown to fold via the formation of transient kinetic
intermediate(s) (15-19). Understanding the structural properties of
the intermediates that occur in the folding/unfolding pathway(s) would
explain how the vast conformational space available could be narrowed
down early during folding to yield the native state of the protein
(20-22).
The refolding kinetics of a number of all-helical proteins have been
investigated (23, 24). It is found that helix formation in proteins
occurs within the low millisecond scale of refolding (24). This aspect
is not surprising because formation of a helix involves intrastrand,
local hydrogen bonding interactions. By contrast, very little
information exists on the modes of refolding of all Human acidic fibroblast growth factor
(hFGF-1)1 is a single chain,
heparin-binding protein involved in a variety of important cellular
processes like the proliferation and differentiation of cells (30, 31).
hFGF-1 is a potent mitogen, and due to its involvement in the wound
repair process, it is a potential therapeutic agent (31). hFGF-1 is a
15-kDa, all 1-Anilino-8-naphthalenesulfonic
acid-NH4+ salt was purchased from Sigma.
Ultrapure urea was procured from Merck, and heparin-Sepharose was from
Amersham Pharmacia Biotech. Labeled 15NH4Cl and
urea-d4 were obtained from Cambridge Isotope
Laboratories. All other chemicals used were of high quality analytical
grade. Unless and otherwise mentioned, all solutions were made in 100 mM phosphate buffer (pH 7.0) containing 100 mM
ammonium sulfate. All experiments were performed at 20 °C.
Protein Expression and Purification--
Expression vector for
the truncated form of the human FGF-1 (hFGF-1, residues 15-154) was
constructed and inserted between the NdeI and
BamHI restriction sites in pET20b. The plasmid containing the hFGF-1 insert was transformed into Escherichia coli BL21
(DE3) pLysS. The expressed protein was purified on heparin-Sepharose column using a NaCl gradient (0-1.5 M). The protein was
desalted by ultrafiltration using an Amicon set up. The homogeneity of the protein was assessed using SDS-polyacrylamide gel electrophoresis. The authenticity of the sample was further verified by electron spray-mass analysis. The concentration of the protein was estimated from the extinction coefficient value at 280 nm.
Preparation of Isotope-enriched hFGF-1--
Uniform
15N isotope labeling was achieved using M9 minimal medium
containing 15NH4Cl. To realize maximal
expression yields, the composition of the M9 medium was modified by the
addition of a mixture mixture of vitamins. The expression host strain
E. coli BL21(DE3)pLysS is a vitamin B1-deficient
host, and hence the medium was supplemented with thiamine (vitamin
B1). Protein expression yields were in the range of 25-30
mg/liter of the isotope-enriched medium. The extent of 15N
labeling was verified by electron spray-mass analysis.
Non-fluorescence Measurements--
All fluorescence experiments
were carried out on a Hitachi F-4500 spectrofluorimeter using a 1-cm
quartz cell. The final concentration of the protein used was 100 µg/ml. Intrinsic tryptophan fluorescence was measured using an
excitation wavelength of 280 nm. The tryptophan emission was monitored
between 300 and 450 nm. The excitation and emission slit widths were
set at 2.5 and 10 nm, respectively. The fraction of unfolded species
formed at various concentrations of urea were estimated based on the
308 to 350 nm ratio.
Far-UV CD Measurements--
All far-UV CD experiments were
performed on a Jasco J-720 spectropolarimeter using a quartz cell of
1-cm path length. Each spectra was an average of 5 scans. The
concentration of the protein used was 0.5 mg/ml. Necessary background
corrections were made in all spectra.
Stopped-flow Fluorescence--
The kinetics of unfolding and
refolding were monitored by fluorescence using an SF-61 stopped-flow
spectrofluorimeter (Hi-Tech Scientific Co., UK). Intrinsic tryptophan
fluorescence measurements were made by exciting the protein at 280 nm
(2.5 nm band pass) via a 0.2-cm path length cell and monitoring the
emission intensity at 350 nm (at 20 °C). For 1:10 mixing ratios in
both unfolding or refolding experiments gave a final protein
concentration of 1 µM hFGF-1 in 100 mM
phosphate buffer (pH 7.0), at 20 °C (containing 100 mM
ammonium sulfate), in different urea concentrations. From 0.2-5
µM protein, no deviations longer than 5% were detected
in the rates or relative amplitudes. The data were fit to a two
exponential equation and analyzed using the RKBIN software provided by
Hi-tech Scientific Co., UK.
For the stopped-flow ANS binding experiments, the stock solution of ANS
(300 µM) was appropriately mixed (in the concentration range between 50 and 250 µM) into the native buffer (100 mM phosphate buffer containing 100 mM ammonium
sulfate (pH 7.0)). 10 µM of equilibrium-unfolded hFGF-1
(in 4 M urea) was diluted 10-fold into 270 µl of the
native buffer containing ANS. This procedure ensured that the
concentration of the protein (1 µM) and urea (0.4 M) remained unchanged during the refolding reaction in the presence of different concentrations of ANS. The binding of ANS was
monitored by setting the excitation and emission wavelengths at 418 and
517 nm, respectively. The bandwidth used in all the measurements was 2 nm.
The protein concentration-dependent ANS binding experiments
were carried out under similar conditions by varying the protein concentration (in the range of 0.5 to 5 µM).
Stopped-flow CD Measurements--
Stopped-flow CD experiments
were performed using SFM-3 Biologic stopped-flow apparatus attached to
a Jasco J-715 spectropolarimeter. The cell path length was 1 mm, and
the dead time of the instrument was estimated to be about 5 ms.
Refolding was initiated by a 10-fold dilution of the denatured hFGF-1
(in 4 M urea) sample with the refolding buffer (100 mM phosphate buffer containing 100 mM ammonium sulfate (pH 7.0), 20 °C). The data presented are an average of 30 scans. The reaction between sodium salt of 2,6-dichlorophenol indophenol and L-ascorbic acid (using a mixing ratio of
1:1) was used to determine the performance limits of the instrument.
The disappearance of the color of 2,6-dichlorophenol indophenol was followed at 524 nm. All solutions contained 0.2 M NaCl to
minimize the ionic strength effects.
The ellipticity changes at 228 nm were analyzed employing an in-built
Biologic software.
Quenched-flow NMR Experiments--
All experiments were carried
out at 20 ± 0.1 °C using a RQF-63 rapid mixing quenched-flow
apparatus (Hi-Tech Scientific Co., UK). Complete denaturation and
exchange of the backbone amide protons with deuterium was achieved by
dissolving 4 M urea into a uniformly 15N
labeled hFGF-1 solution, previously equilibrated for 24 h in D2O at pD 6.6 ± 0.2. Refolding of the denatured
protein was initiated by a 10-fold dilution with 100 mM
phosphate buffer (pH 5.0, containing 100 mM ammonium
sulfate) prepared in H2O. At this pH, negligible labeling
occurred. After variable refolding times ranging from 8 ms to 100 s, the solution was diluted again to 10 times the initial protein
volume with 100 mM phosphate buffer (pH 9.0) (containing 100 nM ammonium sulfate), to initiate labeling of the
deuterated amides in hFGF-1 with protons. After a lapse of 10 ms, the
labeling phase was stopped by a further 3.4 times dilution of the
initial protein volume with 1 M phosphoric acid in water.
It should be mentioned that there were no signs of aggregation during
any of the refolding steps in the quenched-flow experiments.
1H-15N HSQC spectrum of each sample was
recorded on a Bruker DMX 600-MHz NMR spectrometer at 20 °C. A 5-mm
inverse probe with a self-shielded z-gradient was used to
obtain all gradient-enhanced 1H-15N HSQC
spectra. 15N decoupling during acquisition were
accomplished using the GARP sequence. 2048 complex data points were
collected in the 1H dimension of the
1H-15N HSQC experiments. In the 15N
dimension of the spectra, 512 complex data points were obtained. 15N chemical shifts were referenced using the consensus
ratio of 0.0101329118. All spectra were processed on a Silicon Graphics Work station using the UX NMR and Aurelia softwares.
The intensities of the cross-peaks in the HSQC spectra collected at
various refolding times were normalized based on the peak height of the
Ile-70 Two-state Equilibrium Unfolding--
The fluorescence spectrum of
hFGF-1 shows an emission maxima at around 308 nm (Fig.
2, inset A). The fluorescence
of the lone tryptophan residue located at position 121 of the sequence
is completely quenched in the native state of the protein (33, 34). The
quenching effect is attributed to the presence of proximal imidazole
and pyrrole groups in the three-dimensional structure of hFGF-1.
However, unfolding of the protein relieves the quenching effect, and
the fluorescence spectrum of the protein in the unfolded state shows an
emission maxima at around 350 nm (Fig. 2, inset A). Hence,
the refolding/unfolding kinetics of hFGF-1 was monitored by the changes
in the emission intensity at 350 nm.
Fig. 2 shows the urea-induced unfolding curve of hFGF-1 monitored by
fluorescence spectroscopy. The protein is completely unfolded at
concentrations of urea greater than 3.2 M. The urea-induced structural transitions are completely reversible. The concentration of
denaturant (Cm) at which the protein (hFGF-1) is
half unfolded ( Slow Refolding hFGF-1--
Fig. 3
shows the stopped-flow time traces of the refolding of urea-denatured
hFGF-1 (at 25 °C) monitored by the changes in the tryptophan
fluorescence (at 350 nm). Complete signal evolution (at 350 nm) occurs
within a refolding time of 50 s. The fluorescence trace could not
be satisfactorily fit to a single exponential function. A sum of two
exponentials best fits the refolding curve yielding refolding rate
constant values of 0.131 ± 0.02 and 0.023 ± 07 s
hFGF-1 like many other Occurrence of Kinetic Intermediate(s)--
It has been observed
that in many proteins where an intermediate in equilibrium conditions
is not found, the analysis of the refolding kinetics shows the
unambiguous presence of an intermediate. The kinetics of refolding were
examined at 20 °C over a range of urea concentrations (up to 6 M) by monitoring the changes in the tryptophan fluorescence
using the stopped-flow fluorescence technique. Fig.
5 shows the urea dependence of the
natural logarithm of the observed rate constant of folding/unfolding.
Interestingly, the slow phase (minor phase) shows no denaturant
dependence and is consistent with it being a rate-limiting proline
isomerization event. The unfolding rate decreases linearly with the
increase in urea concentration (Fig. 5). However, a prominent curvature could be observed at low concentrations of the denaturant (Fig. 5). In
general, a deviation from linearity ("roll over") is believed to
reflect the accumulation of intermediate species at low denaturant concentrations (36). For a protein folding typically by a two-state mechanism, the Chevron plot (ln k versus
concentration of the denaturant) is expected to be V-shaped, with no
roll over. In summary, these results clearly suggest that the
refolding of hFGF-1 occurs via the formation of transient
intermediate(s).
There is a lot of controversy on the interpretation of the roll over in
the Chevron plot (35-38). Curvatures in Chevron plots, in addition to
accumulation of kinetic intermediates, could also be ascribed to the
movements of the transition state ensemble (36, 39-41). According to
the broad barrier model of protein folding, the folding process is
proposed to occur isoenergetically at the transition state level, and
hence the position of the transition state ensemble is expected to be
sensitive to denaturant conditions (36). Recent studies on the human
spliceosomal protein have demonstrated that the roll over in the
Chevron plot could also be due to the movements of the transition state
along the top of a very broad and flat activation barrier (Hammond
effect, see Refs. 41-44). In general, Hammond effects (during protein
folding/unfolding) are manifested as curvatures in both the limbs of
the Chevron plot (35). In this context, as the roll over is observed in only one of the limbs of the Chevron plot of hFGF-1, it could be
possibly attributed to the accumulation transient intermediate(s) rather than to the movement of the transition state ensemble.
Absence of Hydrophobic Collapse--
ANS is a popular hydrophobic
dye used to probe early events in protein folding (45-47). ANS is
known to bind to transiently exposed hydrophobic patches during folding
of many proteins, leading to a prominent increase in the fluorescence
intensity of the dye. Refolding of hFGF-1 was monitored in the presence
of ANS at 520 nm. No prominent intensity changes were observed within
the dead time of the instrument. Thus, it appears that the refolding
hFGf-1 does not involve early burst phase events such as the
hydrophobic collapse. Interestingly, after 1 s of initiation of
refolding, the ANS fluorescence intensity shows a dramatic increase and
reaches a maximum value at around 2.2 s (Fig.
6). Beyond this refolding time (>2.2 s),
the emission intensity exponentially decreases to reach a steady state
value that is much higher than that of the free ANS (Fig. 6). The
changes in the ANS fluorescence intensity (beyond 2.2 s) could be
best fit to a two exponential term yielding folding rate constant
values of 0.21 ± 0.012 and 0.018 ± 0.008 s
The use of an extrinsic probe such as ANS to monitor the folding
reactions raises the possibility that the binding of the probe to the
protein may perturb the folding reaction (46, 47). We examined this
possibility by studying the refolding of hFGF-1 at varying
concentrations (50-250 µM) of ANS in the refolding buffer. It was found that the hydrophobic dye did not have significant effect(s) on the refolding time constant of the two phases of folding
(data not shown). In addition, the kinetics of refolding of hFGF-1
monitored by changes in the intrinsic tryptophan fluorescence (at 350 nm) was also unaffected by the presence of ANS (up to a concentration
of 250 µM) in the refolding buffer (data not shown). Thus, these results suggest that ANS does not significantly perturb the
refolding kinetics of hFGF-1.
It is important to understand the dramatic increase in the ANS
fluorescence (at ~2.2 s) observed during the refolding of the protein. hFGF-1 is a heparin-binding protein, and its cell regulatory properties are known to be strongly dependent on binding to the proteoglycan. Available crystal and solution structures of hFGF-1 reveal that the heparin-binding site is in the segment (consisting of
residues, 110-130) of the protein spanning
Recently, transient aggregates have been shown to form during the
in vitro refolding of proteins from their random coil states (39-41). In this context, it could be argued that the increase in ANS
intensity observed during the refolding (at ~2.2 s) of hFGF-1 could
be due to the higher binding affinity of the hydrophobic dye to the
transiently formed aggregates (formed in the time scale of ~2.2 s).
In the event of formation of transient aggregates (during refolding),
the refolding rate constants of the various phases of folding are
expected to show a linear decrease with the increase in the
concentration of the refolding protein. However, it is found that the
rate constants of both the phases (major and minor) of refolding
(observed by monitoring the changes in the tryptophan fluorescence) do
not significantly change as a function of the protein concentration
(data not shown). These results clearly suggest that refolding of
hFGF-1 does not proceed via the formation of transient protein
aggregates. The observed increase in the ANS emission signal (at ~2.2
s) is probably due to binding of the dye to the heparin binding domain
formed in this time scale (~2.2 s).
Refolding Events Detected by Quenched-flow Hydrogen
Exchange--
Stopped-flow optical techniques only report gross
conformational changes that occur during the refolding of proteins.
They do not provide any information on the structural changes that occur at a residue level. However, the quenched-flow H/D exchange measurements allow the determination of the time scales of formation of
various hydrogen bonds involved in secondary structure during refolding
of the protein (23). It is in this context that we used the
quenched-flow H/D exchange to monitor the chronology of events in the
refolding pathway of hFGF-1.
The 1H-15N HSQC spectra of hFGF-1 has been
completely assigned (32). Hence, it is possible to follow unambiguously
the folding kinetics of 75 well separated, slowly exchanging amide
residues involved in secondary and tertiary structural interactions in the protein (Fig. 7). These amide protons
are distributed throughout the protein molecule. The exchange kinetics
of all residues are adequately described by a single exponential (Fig.
8).
Representative 1H-15N-HSQC spectra of hFGF-1
samples collected after various refolding times are depicted in Fig. 7.
Only a few stable hydrogen bonds were observed to form during the dead
time (~8 ms) of the quenched-flow apparatus. This aspect is
consistent with the conclusions drawn from the stopped-flow
fluorescence experiments. After 3 s, there were only about 20 amide protons whose intensities decrease by 50-60%. This observation
is in marked contrast to the stopped-flow CD data (at 228 nm) where the
ellipticity changes (primarily representing the secondary structural
interactions) are found to be almost complete (~95%) within 3 s
of initiation of folding (Fig. 4). The backbone hydrogen bonds that are
formed and detected by the far-UV CD signal could be unstable due to the local breathing or sliding of one strand relative to the other. The
net consequence of this structural motion(s) is the rapid formation and
disruption of hydrogen bonds. The short lifetime of the hydrogen bonds
probably accounts for their weak protection (against exchange) observed
in the 1H-15N HSQC spectra collected at
refolding times lesser than 5 s (Fig. 7). Interestingly, 50-60%
resonance intensity decrease for half of the amide protons involved in
the secondary structure formation occurred only after 5 s of
refolding. The secondary structure formation in another
If we arbitrarily subdivide the time constants of refolding (
It is interesting to note that the amide protons of some residues,
which are highly protected from deuterium exchange in the native state,
could not be observed even in the 1H-15N HSQC
spectra collected after 8 ms of folding. These include the amide
protons of Leu-13, Tyr-15, and Val-137. This feature could be
rationalized on the basis of the three-dimensional structure of hFGF-1.
hFGF-1 is a
The three-dimensional structure of hFGF-1 could be best described as a
trigonal pyramid with three topological units consisting each of four
antiparallel
Comparison of the time constant of folding of various
In the second phase of folding (average Comparison with the Refolding Kinetics of
Interleukin-1
The refolding kinetics of hFGF-1 and interleukin-1
The results of the present study support the general notion that
folding/unfolding of large proteins (>100 amino acids) is complex and
essentially involves the formation of intermediate(s). Detailed folding
studies using appropriate mutants of hFGF-1 are in progress to
understand the relationship between the conformational stability and
rate of refolding of the protein.
-sheet, human fibroblast growth factor (hFGF-1) is studied using a
variety of biophysical techniques such as stopped-flow fluorescence,
stopped-flow circular dichroism, and quenched-flow hydrogen exchange in
conjunction with multidimensional NMR spectroscopy. Urea-induced
unfolding of hFGF-1 under equilibrium conditions reveals that the
protein folds via a two-state (native
unfolded) mechanism without
the accumulation of stable intermediates. However, measurement of the
unfolding and refolding rates in various concentrations of urea shows
that the refolding of hFGF-1 proceeds through accumulation of kinetic
intermediates. Results of the quenched-flow hydrogen exchange
experiments reveal that the hydrogen bonds linking the N- and
C-terminal ends are the first to form during the refolding of hFGF-1.
The basic
-trefoil framework is provided by the simultaneous formation of
-strands I, IV, IX, and X. The other
-strands
comprising the
-barrel structure of hFGF-1 are formed relatively
slowly with time constants ranging from 4 to 13 s.
INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS AND DISCUSSION
REFERENCES
-sheet proteins.
As the
-sheet formation involves interstrand interactions between
distant parts of the polypeptide chain, proteins belonging to this
structural class (all
-sheet proteins) are predicted to refold
slowly (25). However, investigation of folding kinetics of several
+
proteins using quenched-flow hydrogen/deuterium (H/D) exchange
revealed that stable
-sheet, and
-helix formation occurs on a
similar time scale (18, 19, 26). Interestingly, folding studies on
interleukin-1
, an all
-sheet protein, showed that stable
-sheet formation only occurs on a time scale greater than 50 s
(25, 27). Thus, it appears that the data base on the refolding kinetics
of all
-sheet proteins needs to be significantly expanded to draw
generalized conclusions on the rates of formation of
-sheets in
proteins (28, 29).
-sheet protein with no disulfide bonds (32, 33). It has
a single tryptophan residue, whose emission properties are known to
describe effectively the conformational changes occurring during the
folding
unfolding transition of the protein (33, 34). Furthermore,
high resolution crystal (31) and NMR structures (32) of hFGF-1 are
available (Fig. 1). These characteristics
render hFGF-1 as an useful model to understand the folding/unfolding
pathways of all
-sheet proteins. In the present study, we
investigate the events in the refolding kinetics of hFGF-1 using a
variety of biophysical techniques such as stopped-flow fluorescence,
stopped-flow CD, and quenched-flow hydrogen exchange. The results
obtained in this study reveal that refolding of the protein occurs very
slowly via the formation of transient intermediate(s). In addition, the
-strands constituting the heparin binding domain appear to form
significantly faster than the other
-strands constituting the
-barrel structure of hFGF-1.
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Fig. 1.
MOLSCRIPT (50) representation of the
structure of hFGF-1. The protein has 12 -strands folded into a
trigonal pyramid structure.
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS AND DISCUSSION
REFERENCES
-methyl protons (
0.2 ppm) in the one-dimensional 1H NMR spectra collected each time prior to recording the
HSQC spectra. The time courses of change in proton occupancies were fitted to a single exponential decay (y = A
exp
kt + C, where A is the
amplitude of the phase, k is the apparent rate constant, and
C is the final amplitude) by the Levenberg-Marquardt nonlinear least squares method, yielding rate constants and phase amplitudes in the kinetic refolding experiment. All data analysis was
performed using Kaleidagraph software (Synergy software).
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS AND DISCUSSION
REFERENCES
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Fig. 2.
Urea-induced unfolding profiles of hFGF-1,
monitored by fluorescence ( ) (308/350 nm) and far-UV CD (
) (228 nm). It could be noticed that the unfolding profiles monitored by
the two spectroscopic techniques are nearly superimposable, implying
that the urea-induced unfolding process of hFGF-1 is cooperative
without the accumulation of stable equilibrium intermediates.
Inset A depicts the intrinsic fluorescence spectra of hFGF-1
in the native and 4 M urea-unfolded states.
Inset B shows the far-UV CD spectra of hFGF-1 in
the native and 4 M urea unfolded states.
Gu = 0) is estimated to be 2.3 M. The m value, which is a measure of the
cooperativity of the unfolding transition, is found to be 1.53 ± 0.4 kcal·mol
1·M
1.
The free energy of unfolding in the absence of the denaturant (
Gu(H2O)) obtained by extrapolation
of
Gu to zero denaturant concentration is
calculated to be 3.46 ± 0.2 kcal·mol
1. To test if the unfolding
transition monitored by fluorescence reflected a total disruption of
the overall structure of the protein, or just a local unfolding, we
analyzed the chemical denaturation of hFGF-1 induced by urea using
far-UV CD spectroscopy. The far-UV CD spectrum of hFGF-1 shows a
prominent positive ellipticity band centered at 228 nm. This CD band
(at 228 nm) is a composite signal and is representative of the
secondary and tertiary structural interactions in the protein (Fig. 2,
inset B). Upon complete unfolding of the protein (at urea
concentrations greater than 3.2 M), the 228 nm far-UV CD
band shows negative ellipticity values (Fig. 2, inset
B). The urea-induced equilibrium unfolding curves followed by
fluorescence and CD are nearly superimposable. Within experimental error(s), the
Gu(H2O) (3.40 ± 0.4 kcal·mole1) estimated from the urea-induced far-UV
ellipticity is identical to that calculated based on fluorescence
spectroscopy. These results strongly suggest that the urea-induced
equilibrium denaturation of hFGF-1 follows a two-state (native
unfolded state) mechanism, without the accumulation of stable intermediate(s).
1 for the faster and slower phases,
respectively (Table I). The first phase
associated with the faster rate accounts for 80-85% of the total
refolding amplitude. The slower phase has an amplitude of about
10-20%. This phase of refolding is probably associated with the
cis-trans-proline isomerization (discussed below). No burst
phase is observed to occur as is evident from the close agreement
between the fluorescence signals at time 0 and that extrapolated from
the equilibrium unfolded hFGF-1 base line. However, we cannot
completely rule out the possibility of occurrence of the burst phase
intermediate(s) because the burst phase changes cannot be traced under
conditions where the intermediate(s) has similar fluorescence as the
unfolded state.
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Fig. 3.
Refolding kinetics of hFGf-1 monitored by
stopped-flow fluorescence (at 350 nm). The refolding trace is
found to best fit with a sum of two exponentials. The refolding rates
(k) and the amplitude (A) of the two phases are
as follows: k1 = 0.131 ± 0.02 s 1; A1 = 81.73 ± 0.34%; k2 = 0.023 ± 0.07 s
1; A2 = 15.2 ± 0.3%.
Rate and time constant of the various phases of refolding of hFGF-1
-barrel proteins shows a far-UV positive
ellipticity band at 228 nm, signifying the secondary and, to a lesser
extent, the tertiary structural interactions in the protein. The far-UV
ellipticity changes (at 228 nm) are complete within a time span of
3 s (Fig. 4). The ellipticity
changes could be best fit to a single exponential equation yielding a
refolding time constant of 1.24 ± 0.65 s
1. The slow minor phase (observed in the
stopped-flow fluorescence experiment) possibly representing the
cis-trans-proline isomerization is not observed in the
stopped-flow CD experiments. In addition, no burst phase ellipticity
change(s) could be detected during the refolding of hFGF-1. Within
experimental error, the ellipticity (at 228 nm) at time 0 is similar to
that obtained from the equilibrium unfolded state base line. It is
interesting to note that the refolding rate constant value of the major
phase estimated using the stopped-flow fluorescence (0.131 ± 0.02 s
1) and stopped-flow CD do not match (Table
I). The fast recovery of the 228 nm ellipticity signal presumably
reflects the rapid acquisition of some portion(s) of the native
structure prior to the burial of the tryptophan in the interior of the
protein. Thus, the results of the stopped-flow fluorescence and
stopped-flow CD experiments strongly suggest the accumulation of
transient intermediate(s) in the kinetic refolding pathway of
hFGF-1.
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Fig. 4.
Stopped-flow kinetics of the refolding of
hFGf-1 monitored by far-UV circular dichroism (at 228 nm). The
refolding trace fits well to a single exponential equation yielding a
time constant of 0.8075 s. The arrow in the figure indicates
the start of the refolding trace.
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Fig. 5.
Urea-dependent folding kinetics
of hFGF-1. The solid circles represent the observed
rate constants for refolding and unfolding obtained from the
stopped-flow fluorescence experiments. The solid line shows
the predicted fit of the kinetic data for a two-state kinetic model.
The small curvature observed at low urea concentrations is
ascribed to the formation of transient kinetic intermediate(s) during
the refolding of hFGF-1.
1, respectively (Table I). The amplitudes of
the fast and slow phase have been estimated to be 90 ± 0.1 and
10 ± 0.2%, respectively.
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Fig. 6.
ANS binding affinity during the refolding of
hFGF-1 monitored by stopped-flow fluorescence at 520 nm. The steep
rise in the ANS fluorescence intensity is attributed to the binding of
the fluorescent dye to the heparin-binding domain formed in the time
scale of about 2.2 s.
-strands IX and X
(31-33). Recent studies indicate that several polysulfonated compounds
could nonspecifically bind to the positively charged heparin-binding
site (33). In this background, the drastic increase in the ANS
fluorescence intensity observed at around 2 s could be attributed
to the transient binding of ANS to the solvent-exposed, heparin binding
domain (residues spanning
-strands IX and X) formed in this time
scale (~2.2 s). This explanation is consistent with the average time
constant of refolding of the
-strands IX and X (constituting the
polyanion binding site), estimated from the quenched-flow H/D exchange
experiments (see below). The heparin-binding site is densely populated
by cationic residues and is interspersed with many nonpolar residues
such as Asn-109, Tyr-111, Ala-111, Trp-121, and Val-123. It appears
that the strong ANS binding affinity of the folding species realized at
~2.2 s is mediated by both the charge and hydrophobic interactions
with the fluorescent dye. However, in the subsequent stages of folding
of the protein (>2.2 s), the nonpolar groups in the heparin-binding
site appear to be sequestered into the interior of the protein, leading
to a decrease in the binding affinity of ANS to the protein. This
aspect is exemplified by the exponential decrease in the ANS emission intensity beyond 2.2 s (Fig. 6).
View larger version (28K):
[in a new window]
Fig. 7.
1H-15N HSQC spectra
of hFGF-1 samples prepared by quenched-flow hydrogen exchange
experiments at various refolding time periods.
View larger version (26K):
[in a new window]
Fig. 8.
Comparison of the observed time courses of
selected residues as a function of length of the refolding time.
All residues were fit to a first order exponential decay. A
depicts the percentage of proton occupancy of residues located in the
various -strands. The percentage of proton occupancy of residues
located in the unstructured region is represented in
B.
-barrel
protein interleukin-1
, monitored by quenched-flow H/D exchange, also
revealed a similar trend (26).
) of
various residues estimated by the quenched-flow H/D exchange technique
into three classes, as fast (1 <
< 3 s), medium
(5 <
< 10 s), and slow (
> 20 s)
folding, most of the residues in
-strands I, IV, IX, and X fit into
the fast folding category (Fig. 9). The
slow folding residues are mostly confined to the unstructured loop
regions inter-spread between the various
-strands. There are 11 residues whose refolding time constant fits into the medium folding
set. These residues are mostly concentrated in
-strands III, V, and
VIII (Fig. 9).
View larger version (34K):
[in a new window]
Fig. 9.
A plot of the time constant of refolding
versus the amino acid residue number
(A). It could be observed that most of the
residues located in -strands I, IV, IX, and X have low refolding
time constant values. The y axis scale is broken to depict
clearly the differences in the refolding time constant values of the
fast folding residues. B, average time constant of various
-strands in hFGF-1. The y axis scale is broken to
indicate clearly the trends in the average time constant values of
various
-strands in the protein.
-barrel protein and the N and C termini of the molecule
are strongly bridged by three hydrogen bonds between, Leu-13-NH and
Leu-135-CO, Tyr-15-NH and Leu-133-CO, Val-137-NH and Pro-11-CO (31,
32). The absence of the 1H-15N cross-peaks
representing the amide protons of Leu-13, Tyr-15, and Val-137 in the
first 1H-15N HSQC spectrum obtained following
the dead-time of the quenched-flow apparatus (<8 ms) strongly suggests
that the formation of hydrogen bonds linking the extreme ends (N and C
termini) of the
-barrel is probably the first event in the refolding
pathway of hFGF-1 (Fig. 10).
View larger version (26K):
[in a new window]
Fig. 10.
Schematic representation
of the initial events in the refolding of hFGF-1.
A, hFGF-1 in the 4 M urea-unfolded state.
B, the bridging of the N- (indicated in yellow)
and C-terminal ends (indicated in green) of the hFGF-1
molecule through backbone hydrogen bonds (indicated by green
dotted lines) between Leu-13-NH and Leu-135-CO, Tyr-15-NH and
Leu-133-CO, and Val-137-NH and Pro-11-CO appears to be the first
detectable event in the refolding of hFGF-1. The N- and C-terminally
linked topology probably provides the structural mold for subsequent
events. C among the various -strands in hFGF-1,
-strands, I, IV, IX, and X form (indicated in yellow) in
similar time scales (~2-2.5 s) and appear to provide the basic
-trefoil framework. Subsequently, D, the trigonal pyramid
structure (as seen in the native state) is built by progressive
layering of the various other
-strands (indicated in
blue) (in the order of strand V, strand XI (
= 5-7
s) > strand XII, strand VIII, strand III, strand II, strand VI,
and strand VII (
= 10-13 s)).
-sheets (31, 32). The
-strands comprising the
structural units A are
-strands I-III and XII. The structural units
B and C consist of
-strands, IV-VII and
-strands VIII-XI, respectively.
-strands
comprising the
-barrel structure of hFGF-1 shows that there is a
clear pattern in the rates of the formation of the 12
-strands. The
average time constants of the
-strands in the protein are as follows
(Fig. 9):
-strand I,
= 1.89 ± 0.24 s;
-strand
II,
= 11.52 ± 0.25 s;
-strand III,
= 8.50 ± 0.31 s;
-strand IV,
= 2.44 ± 0.1 s;
-strand V,
= 4.39 ± 0.25 s;
-strand VI,
= 11.78 ± 0.41 s;
-strand VII,
= 13.33 ± 0.15 s;
-strand VIII,
= 9.50 ± 0.30 s;
-strand IX,
= 2.33 ± 0.34 s;
-strand X,
= 2.5 ± 0.21 s;
-strand XI,
= 6.32 ± 0.34 s; and
-strand XII,
= 11.69 ± 0.41 s. The hFGF-1 molecule with the N-
and C-terminal ends bridged together appears to constitute the basic
mold for subsequent folding events (Fig. 10). The
-strands in the
three topological (A, B, and C) units comprising the trigonal pyramid structure (of hFGF-1) appear to form in a phased manner.
-Strands I,
IV, IX, and X appear to form simultaneously to establish the first
layer of the trigonal pyramid structure (Figs. 9 and 10). The drastic
increase in the ANS fluorescence intensity observed after 2.2 s
could be attributed to the transient binding of the polysulfonated dye
to the heparin binding domain (comprising of the residues in
-strands IX and X) formed in this time scale (
~2.2 ms). The
simultaneous formation of
-strands I, IV, IX, and X could be
ascribed to the hydrogen bonds linking these
-strands (strands I,
IV, IX, and X) with each other in the native state of the protein
(Figs. 9 and 10).
= 4 to 6.5 s),
-strands V and XI are formed (Fig. 9B). The formation of
-strands, II, III, VI
VIII, and XII occurs simultaneously in the
final phase (average
= 9-12 s) of refolding of hFGF-1. Thus,
it appears that the protein regains most of its native interactions
within a time span of about 20 s. The slow phase of folding (
>20 s) probably pertains to conformational reorganization events
arising out of cis-trans-proline isomerization.
--
The three-dimensional structures of hFGF-1 and
interleukin-1
exhibit strong structural homology. Although both
proteins share only 10-15% sequence similarity, their backbone in 9 of the 12
-sheet strands could be superimposed with a root mean
square deviation of 0.5 Å (48). The folding pathway of
interleukin-1
has been investigated using a variety of spectroscopic
techniques including quenched-flow hydrogen exchange experiments (26,
49). It is informative to compare the events in the refolding kinetics of hFGF-1 and interleukin-1
. Striking similarities could be observed in the refolding kinetics of hFGF-1 and interleukin-1
. Both the proteins fold very slowly, and complete folding of these proteins occurs on a time scale greater than 50 s (26). As observed in hFGF-1, the formation of unstable
-sheets in interleukin-1
is shown to occur rapidly, but stabilization of the three-dimensional structure with the establishment of native hydrogen bonds begins only
after 1 s of initiation of refolding. Additionally, the refolding of both hFGF-1 and interleukin-1
involves the formation of kinetic intermediate(s).
also show
significant difference(s). Unlike hFGF-1, where no burst phase hydrophobic collapse has been observed, the refolding pathway of
interleukin-1
involves clustering of the hydrophobic residues as
evidenced by the strong ANS binding in the burst phase of refolding. In
hFGF-1 the formation of hydrogen bonds between the N- and C-terminal ends is the first detectable event. Thus, comparison of the folding kinetics of these two structurally homologous proteins reveals that
although the time scale for refolding of the proteins is dependent on
their overall structural architecture, the individual events in the
refolding pathway appear to be governed by the unique local and long
range interactions present in the protein in context.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Prof. Ing-Ming Chiu, Ohio State University, for providing us the hFGF-1 clone. We also thank the anonymous referee for valuable comments on an earlier version of the manuscript.
![]() |
FOOTNOTES |
---|
* This work was supported by research grants from National Science Council, Taiwan, Dr. C. S. Tsong Memorial Medical Research Foundation (to C. Y.), and the National Health Research Institute, Taiwan (to D.-H. C).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. Tel.: 886 35 715131 (ext. 5605); Fax: 886 35 711082; E-mail:
cyu@mx.nthu.edu.tw.
Published, JBC Papers in Press, October 18, 2000, DOI 10.1074/jbc.M005921200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: hFGF, human fibroblast growth factor; ANS, 1-anilino-8-naphthalene sulfonate; H/D, hydrogen/deuterium.
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