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INTRODUCTION |
A large body of evidence supports the view that the rapid turnover
of actin filaments drives actin-based motility processes such as the
forward movement of the leading edge of the lamellipodium of locomoting
cells, the propulsive movement of Listeria monocytogenes, the movement of cortical actin patches in yeast, or the extension of
the growth cone (1). Filaments turn over via a treadmilling mechanism,
whereby the steady growth of barbed ends is fed by the subunits
depolymerizing from the pointed ends (2, 3). While barbed end growth,
which provides the motile force (4), is restricted to specialized
regions of the cell such as the leading edge (5), depolymerization may
occur from all pointed ends in the cell medium. The observed rates of
actin-based movement fall in the range 1-20 µm/min, corresponding to
treadmilling rates of 7-130 subunits/s/filament, i.e. 1-2
orders of magnitude higher than the treadmilling rate measured in
vitro in solutions of pure actin (6).
Actin-binding proteins of the
ADF1/cofilin family have
recently been demonstrated to enhance the treadmilling of F-actin
in vitro by 25-fold and consistently to increase the rate of
Listeria propulsion in platelet extracts (7). We proposed
(7) that these in vitro properties of ADF accounted for the
enhancement of motility of Dictyostelium discoideum (8) due
to ADF overexpression and for its high level of expression in early
development (9). Consistently, ADF was shown to be responsible for the
high rate of filament turnover in yeast (10).
It was initially thought that ADF depolymerized F-actin rapidly due to
a severing activity (11-14). Severing of the filaments was thought to
create a large number of uncapped barbed ends, known to depolymerize
rapidly. It is worth stressing that the above reasoning is incorrect in
the cellular context of the steady state of actin assembly, in which
the concentration of ATP·G-actin always lies between the critical
concentrations of the barbed and pointed ends. In such a situation,
uncapped barbed ends that would putatively be created by severing of
the filaments would immediately grow and not depolymerize. Recent data
show that the ADF-induced enhancement of treadmilling is not due to
severing, but to an increase in the rate of depolymerization from the
pointed ends, which is the rate-limiting step in the treadmilling cycle (7, 15). This function of ADF is mediated by its specific binding to
ADP-bound G- and F-actins, thereby participating in actin filament
assembly while changing the assembly rate parameters in an end-specific
fashion.
A unique feature of all ADFs (except maybe yeast ADF) is their ability
to be activated by rapid dephosphorylation in response to diverse
stimuli that lead to changes in actin dynamics (see Ref. 16 for a
review). The phosphorylated serine (serine 3 in vertebrate ADF and
serine 6 in plant ADF (17)) resides in the N-terminal region of the
molecule. The three-dimensional structure of unphosphorylated ADF from
different sources has been solved using NMR (18) or x-ray diffraction
(19, 20) and shows a remarkable similarity to gelsolin segment-1. An
extensive mutagenesis study, however, demonstrated that yeast cofilin
interacts with actin in a manner quite different from gelsolin
segment-1 (21), consistent with the different functions of these two
proteins. Image reconstruction of frozen hydrated specimens of
filaments decorated with ADF indicates that ADF binds cooperatively to
F-actin while changing the helical periodicity of the filament
(14).
To analyze and simulate the effects of ADF on actin dynamics
quantitatively, it is necessary to have access to the rate parameters of ADF interaction with both G- and F-actins. This rapid kinetic study
is carried out here using human ADF and plant ADF (Arabidopsis thaliana ADF1) on a comparative basis. Indeed,
although structural and functional evidence indicates that all ADFs
have identical regulatory properties (7, 18-21), quantitative
differences may exist between ADFs from different species, which may
introduce important differences in their function. For instance, human
ADF has been reported to cause total depolymerization of F-actin at pH
8.0 (13), whereas plant ADF causes only partial depolymerization at all
pH values (7). The effect of phosphorylation of ADF on its function is
also investigated here using the serine to aspartate mutation of serine
3 in A. thaliana ADF1 to mimic phosphorylation. The binding parameters and the activity of the S6D mutant of
ADF1 have been characterized to determine whether
phosphorylation affects the activity of ADF1 or its binding
to actin.
Kinetics also offer the opportunity to elucidate the mechanism by which
ADF changes the structure of the filament. Kinetic analysis of the
binding of ADF to filaments maintained in different structural states
by drugs (phalloidin), regulatory proteins (gelsolin), or chemical
modification of subdomain 2 provides further insight in the structure
of ADF·F-actin.
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MATERIALS AND METHODS |
Proteins--
Actin was purified from rabbit muscle and isolated
as CaATP·G-actin through Sephadex G-200 chromatography in G buffer (5 mM Tris-Cl, pH 7.5, 0.1 mM CaCl2,
0.2 mM dithiothreitol, 0.2 mM ATP, and 0.01%
NaN3). Procedures to prepare NBD-labeled actin,
MgATP·G-actin, and MgADP·G-actin were as described previously (7).
Subtilisin-cleaved actin (22) was prepared by incubating
CaATP·G-actin (20 µM) in G buffer with subtilisin
Carlsberg (Sigma) at a 1:1500 (w/w) subtilisin/actin ratio for 45 min
at 20 °C. The reaction was stopped by addition of 1 mM
phenylmethylsulfonyl fluoride.
Human ADF and A. thaliana ADF1 were recombinant
proteins expressed in Escherichia coli using the
pET vector and were purified by DEAE-cellulose and SP-Trisacryl
chromatography as described (7). The S6D mutant of ADF1 was
produced and purified in the same way. Gelsolin from human plasma was a
gift from Dr. Yukio Doi. Protein concentrations were determined
spectrophotometrically using extinction coefficients of 0.617 mg
1 cm2 for actin at 290 nm, 0.89 mg
1 cm2 for wild-type and S6D mutant ADFs at
278 nm (7), and 0.64 mg
1 cm2 for human ADF
(13).
Static Fluorescence Measurements--
The change in fluorescence
of NBD-labeled G-actin was used to monitor the binding of ADF to
G-actin under a variety of ionic conditions, as described previously
(7). It was observed that human ADF, in contrast to plant ADF, did not
modify the fluorescence of NBD-labeled actin to an appreciable extent.
Therefore, in most cases, the equilibrium dissociation constant for the
binding of human ADF to G-actin was derived from competition studies,
in which the binding of ADF1 was measured in the presence
of different known amounts of human ADF or by displacing actin-bound
ADF1 by addition of increasing amounts of human ADF.
Fluorescence titration curves were analyzed as follows to derive the
equilibrium dissociation constant of the human ADF·G-actin complex.
The observed fluorescence (F) can be written as follows (Equation 1),
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(Eq. 1)
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where [A], [AX], and [AY]
represent the concentrations of free G-actin,
ADF1·G-actin, and human ADF·G-actin complexes,
respectively, and f0, fx, and
fy represent the specific fluorescence of actin in
the respective species.
Equation l can be written as follows (Equation 2),
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(Eq. 2)
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where Kx and Ky are the
equilibrium dissociation constants for the ADF1·G-actin
(AX) and human ADF·G-actin (AY) complexes,
respectively. Theoretical curves were computer-generated using Equation 2 and calculated values of the concentrations of free ADF1
(X) and free human ADF (Y) as follows. Combining
the laws of mass action and mass conservation leads to Equation 3.
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(Eq. 3)
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The concentration of free human ADF ([Y]) was
calculated as a function of the concentration of free ADF1
([X]) by solving Equation 3. Incremented values of
[X] were computer-generated, from which corresponding
values of [Y] were obtained, for the given values of total
human ADF concentration ([Y0]). Then the total
concentration of ADFl ([X0]) was
calculated as follows (Equation 4).
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(Eq. 4)
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The calculated value of F was plotted as a function
of [X0]. Values of f0,
fx, and fy used in the
calculation of F were experimentally determined. The value
of Kx was first easily derived from analysis of the
titration curve of G-actin by ADF1 in the absence of human
ADF. Its value was used in the calculation of F,
[Y], and [X0]. The value of
Ky was then adjusted manually to obtain the best
global fit to all the titration curves of G-actin by ADF1
at different concentrations [Y0] of human
ADF.
Kinetic Measurements--
The kinetics of binding of
ADF1 to G-actin were monitored by the quenching of
NBD-labeled G-actin fluorescence using a stopped-flow apparatus (DX.17
MV, Applied Photophysics Ltd.). NBD-labeled G-actin (in either MgATP-
or MgADP-bound form) in syringe A was mixed with ADF, present at
different concentrations in syringe B. Experiments were carried out at
low ionic strength (G buffer) and at physiological ionic strength (0.1 M KCl and 1 mM MgCl2 added to G
buffer). The change in NBD fluorescence was recorded (
ex = 475 nm). A filter cutting the excitation light (0% transmission at
< 515 nm and 65 and 80% transmission at 530 and 540 nm,
respectively) was placed on the emission beam. Dissociation of
ADF1 from G-actin was similarly monitored by displacing
bound ADF1 from the preformed NBD-labeled ADF1·G-actin complex (syringe A) by an excess of
unlabeled G-actin (syringe B). Dissociation of human ADF from G-actin
was monitored by displacing bound human ADF from the preformed
NBD-labeled ADF·G-actin complex by an excess of ADF1.
The kinetics of binding of plant and human ADFs to F-actin were
monitored in the stopped-flow apparatus using both light scattering and
NBD fluorescence. The changes in light scattering and fluorescence were
recorded using the same solutions of F-actin and ADFs. Light scattering
was monitored at 320 nm, with a 270-µs electronic attenuation filter
to minimize the noise. Superimposable traces recorded from a minimum of
six consecutive shots were averaged and used for kinetic analysis. The
changes in NBD fluorescence were recorded as described above. Because
fluorescence data are less noisy than light scattering, three
superimposable traces coming from consecutive shots were averaged. The
kinetics of dissociation of ADF from F-actin were monitored in a Spex
spectrofluorometer by displacing ADF bound to NBD-labeled F-actin (1-2
µM) by an excess of unlabeled F-actin (10-50
µM).
Sedimentation Assay--
The binding of ADFs and S6D mutant
ADF1 to F-actin and the extent of depolymerization linked
to ADF binding were measured by SDS-polyacrylamide gel electrophoresis
analysis of the pellets and supernatants of samples containing
different amounts of F-actin and ADF (7). Experiments were carried out
at pH 6.5 and 8.2. The steady-state concentration of monomeric actin
and ADF present in the supernatant was estimated by scanning Coomassie
Blue-stained gels using an Arcus II scanner (AGFA Corp., Orangeburg,
NY) and comparing with actin and ADF standards using the National
Institutes of Health Image analysis program.
ATPase Measurements--
Measurements of the steady-state
hydrolysis of ATP resulting from treadmilling of F-actin were carried
out in the presence of different concentrations of wild-type or mutant
ADF proteins as described (7). Actin (15 µM) was
polymerized in the presence of [
-32P]ATP. When steady
state was reached, the solution was split into several samples
supplemented with different amounts of ADF. The hydrolysis of ATP was
monitored at 50-min intervals over a period of 6 h.
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RESULTS |
Equilibrium Binding of Human ADF and Arabidopsis Wild-type and S6D
Mutant ADF1 to G-actin--
A. thaliana
ADF1 causes a 35% quenching of NBD-labeled actin
fluorescence upon binding to G-actin (7). In contrast, the binding of
human ADF very little affected the fluorescence of NBD-labeled G-actin.
Therefore, the equilibrium dissociation constant for human ADF binding
to G-actin was derived from fluorescence titration curves of
NBD-labeled G-actin by Arabidopsis ADF1 in the
presence of human ADF acting as a competitive ligand. Examples are
shown in Fig. 1. Data were fitted as
developed under "Materials and Methods" to derive the values of the
equilibrium dissociation constants. All data obtained under different
ionic and actin-bound nucleotide conditions are summarized in Table
I. The two ADF variants appear to have
very similar affinities for G-actin under a variety of conditions.
Human ADF, like plant ADF, shows high affinity (108
M
1) for ADP·G-actin and very low affinity
(105 M
1) for ATP·G-actin under
physiological ionic conditions. This result is at variance with
previous reports (12, 13) in which the shift in critical concentration
plots induced by ADF was interpreted as evidence for high affinity
binding of ADF to ATP·G-actin. The reason for this discrepancy will
be clarified under "Discussion."

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Fig. 1.
Fluorescence titration of NBD-labeled G-actin
by plant and human ADFs. a, binding to ATP·G-actin.
Arabidopsis ADF1 at the indicated concentrations
was added to fully NBD-labeled ATP·G-actin (1 µM) in
the absence (closed squares) and presence of 1.1 µM (open squares) or 2.2 µM (open circles) human ADF in G
buffer at pH 7.6. Closed circles represent the
titration curve of ATP·G-actin by human ADF. Continuous
lines were calculated using KD values of
0.07 and 0.05 µM for the complexes of ATP·G-actin with
plant and human ADFs, respectively, in a competitive binding scheme and
normalized specific fluorescence of 1, 0.61, and 1.16 for G-actin,
ADF1·G-actin, and human ADF·G-actin, respectively.
b, binding to ADP·G-actin. Arabidopsis
ADF1 was added to 1 µM NBD-labeled
ADP·G-actin in the absence (closed circles) and
presence of 1 µM (open circles) or
2 µM (open squares) human ADF in G
buffer containing ADP at pH 7.6. The binding of human ADF does not
affect the fluorescence of ADP·G-actin. Curves were calculated using
KD values of 0.03 and 0.02 µM for the
complexes of ADP·G-actin with plant and human ADFs, respectively.
a.u., arbitrary units.
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Table I
Equilibrium parameters for binding of Arabidopsis wild-type and S6D
mutant ADF1 and human ADF to G-actin
Values of KD for complexes of G-actin with
ADF1 and S6D mutant ADF1 were derived from fluorescence
titration curves of NBD-labeled G-actin. Values of
KD for complexes of G-actin with human ADF were
derived from analysis of the competition between ADF1 and human
ADF as described under "Materials and Methods." By convention, a
value of 1.0 is attributed to the specific fluorescence of NBD-labeled
G-actin under each ionic condition.
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The binding of the S6D mutant ADF1 protein was measured in
a similar fashion. S6D mutant ADF1 affected the
fluorescence of NBD-labeled G-actin like the wild-type protein,
indicating that the interfaces of the mutant and wild-type proteins
with G-actin were the same, but showed a 20-fold lower affinity (Table
I).
Kinetics of Binding of ADF to G-actin--
The kinetics of
association of ADF1 with NBD-labeled G-actin were monitored
by fluorescence in the stopped-flow apparatus. At all ionic strengths,
the association of ADF1 with either ATP·G-actin or
ADP·G-actin was extremely rapid and could be monitored by lowering the temperature to 4 °C. At physiological ionic strength and at 4 °C, the time course of the decrease in NBD fluorescence shown in
Fig. 2 reflected the reversible
bimolecular association of ADF1 with actin, with
association and dissociation rate constants of 155 ± 10 µM
1 s
l and 16 ± 10 s
l, respectively. The dissociation rate constant could be
directly determined with a greater accuracy by displacing
ADF1 bound to NBD-labeled ADP·G-actin by an excess of
unlabeled ADP·G actin. The resulting increase in fluorescence is
kinetically limited by the dissociation of ADF1 from
NBD-labeled G-actin. A value of 12 s
l was therefore
obtained for the dissociation rate constant of ADF1.
Similarly, the rate constant for dissociation of human ADF from
ADP·G-actin was derived from the exponential decrease in fluorescence
recorded upon displacing human ADF bound to ADP·G-actin by a large
excess of ADF1. A value of 13 s
1 was obtained
for the dissociation rate constant of human ADF. In conclusion, under
physiological conditions, plant and human ADFs bind ADP·G-actin with
high affinity and in very rapid association/dissociation equilibrium.

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Fig. 2.
Kinetics of binding of ADF to
ADP·G-actin. NBD-labeled ADP·G-actin (1 µM) was
rapidly mixed in the stopped-flow apparatus with ADF1 at 0, 0.37, 1, 1.5, 2, and 3 µM (top to
bottom curves). Noisy curves represent the data
coming from six averaged traces at each concentration. The dead time
was 2 ms. Conditions were physiological ionic strength buffer at pH 7.6 and 4 °C. Solid lines are kinetic curves
calculated using KINSIM (44) within a reversible bimolecular reaction
scheme, with k+ = 155 µM 1·s 1,
k = 16 s l, and 25% quenching of
NBD-labeled actin fluorescence upon binding ADF1.
a.u., arbitrary units.
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Kinetics of Binding of ADF to F-actin--
The kinetics of
association of plant and human ADFs with F-actin were monitored in the
stopped-flow apparatus by both NBD fluorescence and light scattering in
ATP-containing F buffer (G buffer supplemented with 0.1 M
KCl and 1 mM MgCl2) at 20 °C and pH 7.6. While the fluorescence of NBD-labeled G-actin was unaffected by human
ADF, the fluorescence of NBD-labeled F-actin was quenched by human ADF,
as previously observed with Arabidopsis ADF1 in static measurements (7). Fig. 3
(a and b) shows that the quenching of
fluorescence temporally correlates with an increase in light scattering, which reflects the 42% increase in mass per unit length of
the filament associated with ADF (18 kDa) binding to each F-actin subunit (42 kDa). The initial increase in light scattering was followed
by a slower decrease (Fig. 3c), which, as previously observed with ADF1 (7), reflects the partial
depolymerization of ADF·F-actin, and relaxation toward a new steady
state in which ADF is bound in part to ADP·G-actin and in part to
F-actin. Therefore, the amount of ADF·F-actin at the end of the rapid
phase is greater than at the end of the slow phase. The extent of
fluorescence decrease reached at the end of the rapid phase (Fig.
3d) truly represents the binding of ADF to F-actin before
the partial depolymerization of ADF·F-actin into ADF·ADP·G-actin
has occurred. This curve therefore does not exhibit the strong
sigmoidal shape displayed by curves derived from sedimentation data,
when the amount of ADF·F-actin was plotted versus the
concentration of ADF in the supernatant, misleadingly assumed to
represent the concentration of free ADF (14). In view of the present
and previous (7) data, this strong sigmoidal behavior results from the
binding of ADF to both G- and F-actins at steady state. The extent of
depolymerization was greater at pH 8 than at pH 6.8 (7); otherwise, the
overshoot kinetic curves observed in light scattering were similar at
all pH values.

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Fig. 3.
Kinetics of binding of ADF to F-actin.
NBD-labeled F-actin (4 µM) was rapidly mixed with human
ADF at the indicated concentrations. Conditions were physiological
ionic conditions and ATP-containing F buffer at pH 7.6 and 20 °C.
a, rapid quenching of NBD fluorescence; b,
temporally correlated rapid increase in light scattering; c,
evolution of light scattering (same experiment as in b) on a
longer time base, showing that the rapid increase in light scattering
is followed by a slow decrease corresponding to the depolymerization
process; d, amplitude data from a. The relative
extent of fluorescence change at the end of the rapid phase
( F/ Fmax) was measured at
different total concentrations of ADF.
[ADF0]/[F0] represents the ratio of the
total concentrations of ADF and F-actin. The dashed
line represents the curve expected for michaelian binding of
ADF to F-actin with an equilibrium dissociation constant of 0.3 µM.
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Fig. 4a shows that human ADF
bound more slowly to F-actin than Arabidopsis
ADF1. The extent of fluorescence quenching was also
slightly lower for human ADF. For the two ADF species, the time courses
of the increase in light scattering and of quenching of NBD-labeled
F-actin fluorescence were not consistent with a simple bimolecular
reaction, but showed an acceleration following a lag phase, indicating
that ADF bound to F-actin in a kinetically cooperative fashion (Fig.
4b). This behavior suggests that the binding of the first
ADF molecules to a bare filament is accompanied by a large cooperative
structural change of this filament, allowing the faster subsequent
binding of ADF.

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Fig. 4.
ADF binds to F-actin with a strong kinetic
cooperativity. a, comparison of plant and human ADF
binding kinetics. NBD-labeled F-actin (2.5 µM) was mixed
with Arabidopsis ADF1 (thick
lines) or human ADF (thin lines) at
2.5, 5, 7.5, 10, 12.5, 15, and 20 µM (top to
bottom curves). Note the difference in the
association rates and extent of fluorescence quenching displayed by the
two ADFs. Conditions were as described for Fig. 3. b, fit of
the proposed model to the kinetic data for ADF1 binding to
F-actin (from a) within the scheme given under
"Results," using the following values of the rate parameters:
n = 2, kn = 0.037 µM 2·s 1, and
ke = 130 µM 1·s 1. a.u.,
arbitrary units.
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To get more insight into the mechanism by which ADF modifies the
structure of F-actin, attempts were made to elaborate a model quantitatively accounting for the time courses of ADF binding to
F-actin shown in Fig. 4b. A simple model in which the
binding site consisted of two neighboring F-actin subunits
(F2), binding two consecutive ADF molecules with very
different rate constants, was first envisaged. It was possible to find
sets of rate parameters that generated calculated curves able to fit
some of the experimental curves, but this model failed to provide a
satisfactory global fit to a series of time courses covering a broad
range of ADF concentrations. The curves calculated using this model did
not reconstitute the highly cooperative concentration dependence
exhibited by the experimental kinetic curves.
A satisfactory fit to the complete series of kinetic curves was
provided by a model within which ADF binds to a bare filament in two
consecutive steps. In an initial slow "nucleation" step, the
cooperative association of ADF with a critical number (n) of
neighboring F-actin subunits (forming a "nucleus") changes locally
the structure of the filament, thus allowing faster subsequent binding
of ADF along the filament in a "zipper-like" elongation process.
This model is described by the following scheme: F + n ADF
Fn (nucleation) and Fn + ADF
Fn + F·ADF (elongation), leading to Equations 5 and 6,
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(Eq. 5)
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(Eq. 6)
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with [F0] = [F] + [F·ADF] and
[ADF0] = [ADF] + [F·ADF], where [F0]
and [ADF0] represent the total concentrations of F-actin subunits and ADF, respectively, and [Fn] is the concentration of "nucleating" sites on F-actin. For simplicity, it is assumed that [Fn]
[F0] and that the reactions are
irreversible. The change in fluorescence is proportional to the
concentration of ADF·F-actin. In Equation 6, the factor
[F]/[F0] is included to express the fact that the
concentration of nuclei ([Fn]) decreases as stretches of
F-actin are entirely decorated by ADF in the progress of the binding
reaction.
The kinetics of ADF binding to F-actin were simulated within the above
scheme by integration of Equations 5 and 6 using Excel (Microsoft). A
good fit to the whole series of curves could be obtained only for
n = 2. Higher values of n were
unsatisfactory, independent of the values of kn and
ke. Calculated curves (Fig. 4b)
represented the best fit to the data, using the experimental values of
F0 and ADF0 and values of 0.037 and 130 µM
1·s
1 for
kn and ke, respectively.
The kinetic cooperativity in the binding of ADF to F-actin results in
equilibrium binding curves (expressed as a function of total ligand)
that differ, in the 50-90% saturation range, from the behavior
expected for michaelian binding. The data points above 50% saturation
(Fig. 3d) fall above the dashed line
representing michaelian binding of ADF to F-actin with an equilibrium
dissociation constant of 0.3 µM (7), consistent with
cooperative binding.
The rate of ADF binding to F-actin was pH-dependent.
Kinetic cooperativity was observed at all pH values; however, the
binding was ~4-fold slower at pH 8.2 than at pH 7.2.
The kinetics of ADF binding to Ca·F-actin or Mg·F-actin were
similarly cooperative and not appreciably different in rate. The only
difference was that the quenching of fluorescence of NBD-labeled
F-actin was 11.5% greater when Mg2+ was bound than when
Ca2+ was bound to actin.
Internal cooperativity in the structure of F-actin has already been
observed (23-27). Both the structure and the flexural rigidity of the
actin filament are affected by the nature of the bound nucleotide or
ligands like phalloidin (28-30). A cooperative change in structure
propagating over long distances along the filament has been observed
when filaments are capped by gelsolin or assembled from actin
proteolytically cleaved in subdomain 2 (27, 28). This structural change
is visualized by a bridge of density connecting the two strands of the
long pitch helix. The fact that ADF does not bind to filaments
stabilized by phalloidin or BeF3
(7)
and promotes a structural change in the filament when it binds
ADP·F-actin prompted us to examine the kinetics of ADF binding to
filaments capped by gelsolin or assembled from subtilisin-cleaved actin. Fig. 5a shows that the
rate of ADF binding to capped filaments increased with the
gelsolin/actin ratio. The effect of gelsolin could be detected at a
1:200 ratio to actin. At a gelsolin/actin ratio of 1:20, the lag time
disappeared, and ADF bound to capped filaments with a rapid initial
rate. At gelsolin/actin ratios of 1:20 and 1:10, the quenching of
NBD-labeled F-actin fluorescence due to ADF binding was reduced by 10 and 20%, respectively, indicating that ADF did not bind to the two
terminal F-actin subunits interacting with gelsolin at the barbed end,
which both exhibited the fluorescence of F-actin. These results suggest
that, in agreement with Egelman and coworkers (27), the structure of
the filaments is affected by gelsolin in such a way that ADF binding is
facilitated and does not require a nucleation step. However, our data
indicate that the structural change of the filament induced by gelsolin propagates over 10-20 rather than several hundred subunits.
Similarly, when loop 45-52 in actin subdomain 2 was cleaved by
subtilisin, ADF bound much faster to F-actin, with a marked reduction
of the lag time (Fig. 5b).

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Fig. 5.
ADF binds faster to gelsolin-capped or to
subtilisin-cleaved F-actin. a, ADF1 (5 µM) was mixed in the stopped-flow apparatus with
NBD-labeled F-actin (2.5 µM) preincubated overnight in
the presence of gelsolin at the indicated gelsolin/actin ratios.
Conditions were ATP-containing F buffer at pH 7.6 and 20 °C.
t1/2 values were 400, 300, 200, and 150 ms at
gelsolin/actin ratios of 0, 1:200, 1:20, and 1:10, respectively.
b, ADF1 (6 µM) was mixed in the
stopped-flow apparatus with 4 µM standard F-actin
(thin curve) or subtilisin-cleaved F-actin
(thick curve). Conditions were as described for
a. t1/2 values were 350 and 80 ms for
control and subtilisin-cleaved actin, respectively. a.u.,
arbitrary units.
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Phalloidin inhibited ADF binding to F-actin cooperatively in a
substoichiometric fashion (Fig. 6). Over
90% of the F-actin subunits did not bind ADF at a phalloidin/actin
ratio of 1:3, indicating that binding of phalloidin to one subunit
cooperatively modifies the local structure of the filament, making a
sequence of three subunits inaccessible to ADF. The binding of ADF to
accessible sites was also much slower than to bare filaments.

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Fig. 6.
Phalloidin inhibits ADF binding to F-actin in
a substoichiometric cooperative fashion. ADF1 (5 µM) was mixed in the stopped-flow apparatus with 2.5 µM NBD-labeled F-actin preincubated overnight in the
presence of phalloidin at the indicated phalloidin/actin ratios.
Inset, dependence of the extent of fluorescence quenching on
the phalloidin/actin ratio (r). Some of the data points in
the inset correspond to curves that are not shown on the
main graph for clarity. Conditions were as
described for Fig. 5. a.u., arbitrary units.
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To understand how the interaction of ADF with actin affects the
dynamics of actin filaments, it is important to know the rate of
dissociation of ADF from F-actin. The dissociation of ADF from F-actin
was monitored by the increase in fluorescence linked to the
displacement of ADF bound to NBD-labeled F-actin by an excess of
unlabeled F-actin. The rate constant of the observed first-order process had a value of 0.035 s
l, independent of the
initial saturation level of NBD-labeled F-actin by ADF. Alternatively,
ADF dissociation from NBD-labeled F-actin could be elicited by
phalloidin since ADF and phalloidin bind to F-actin in a mutually
exclusive fashion (7). The observed fluorescence increase was a
monoexponential process at a high concentration of phalloidin. The
first-order rate constant reached a finite limit of 0.1 s
1. This value is higher than the one derived from the
chase experiment described above, indicating that phalloidin first
binds to ADF·F-actin, and then ADF dissociates from the ternary
ADF·F-actin·phalloidin complex.
Human ADF Causes Partial Depolymerization of F-actin in a
pH-dependent Fashion, Qualitatively Similar to Plant
ADF--
Chick or human ADF has been reported to cause complete
depolymerization of F-actin at pH 8.0 (12, 13). This result suggested that ADF was a G-actin-sequestering protein, a conclusion at variance with the ones derived from a more recent study of plant ADF (7) that
shows that ADF interacts with both F- and G-actins. To understand whether vertebrate ADF might have different properties from plant ADF,
analysis of the ADF-induced depolymerization at different pH values was
carried out using a sedimentation assay. The experiments shown in Fig.
7 were conducted either by adding
increasing amounts of human ADF to F-actin at a given concentration (20 µM) or by adding increasing amounts of F-actin to human
ADF maintained at a constant concentration (30 µM). Human
ADF caused a more extensive depolymerization of F-actin than plant ADF
at all pH values; nonetheless, the depolymerization always occurred to
a limited extent. At pH 6.5, a maximum of 2 µM actin was
depolymerized by human ADF, a concentration ~2-fold higher than with
Arabidopsis ADF1 (7). At pH 8.2, only 6-7
µM actin was depolymerized by human ADF
(versus 2.5 µM actin for plant ADF). This
result is at variance with reports by Hawkins et al. (12)
and Hayden et al. (13), but is consistent with the
polymerization properties of ADF·ADP·actin, as shown below.

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Fig. 7.
Human ADF causes only partial
depolymerization of F-actin at high pH. a, samples of
F-actin (13.5 µM), polymerized at pH 8.2 in
ATP-containing F buffer, in the presence of plant or human
(Hu) ADF at the indicated concentrations (in
µM) were centrifuged to sediment F-actin. Supernatants
were submitted to SDS-polyacrylamide gel electrophoresis. The actin
band was visualized by Coomassie Blue stain. The last lane on the
right represents total actin (13.5 µM unsedimented
sample). b, actin at the indicated concentrations was
polymerized at pH 8.2 in the presence of 20 µM plant or
human ADF. The actin band in the electrophoresed supernatants is shown
together with actin standards at the indicated concentrations for easy
visual evaluation of the concentration of unassembled actin in the
samples. Quantitative evaluation using a more expanded calibration
curve (not shown) led to values of 2 and 6.5 µM
unassembled actin at steady state in the presence of
Arabidopsis ADF1 and human ADF,
respectively.
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ADP·G-actin polymerizes reversibly with a critical concentration of
1.5 µM under physiological ionic conditions (31). The nucleation of ADP·actin is energetically very difficult (32, 33). In
contrast, the nucleation of the ADF·ADP·actin complex to form
ADF-decorated filaments appeared extremely easy (Fig. 8, inset). The assembly
process, monitored turbidimetrically, displayed no detectable lag,
indicating that nuclei formed rapidly. A critical concentration plot
was derived from measurements of the extent of turbidity change at the
end of the reaction (Fig. 8). The critical concentrations for assembly
of ADF·ADP·F-actin from the ADF·ADP·G-actin complex were 3.5 and 8 µM for plant and human ADFs, respectively, at pH 8 under physiological ionic conditions. These data demonstrate that it is
not theoretically possible to depolymerize more than 8 µM
actin with vertebrate ADF, in agreement with the experimental evidence
displayed in Fig. 7.

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Fig. 8.
Critical concentration plots for the
reversible polymerization of ADF·ADP·actin. MgADP·G-actin in
G buffer containing ADP was supplemented with 1 molar eq plus 2 µM excess ADF1 or human ADF and 15 µM excess S6D mutant ADF1 and induced to
polymerize spontaneously by addition of 0.1 M KCl and l
mM MgCl2. The polymerization process was
recorded turbidimetrically at 310 nm. The extent of turbidity change at
the end of the reaction is plotted versus the total
concentration of ADF·ADP·actin. Open circles,
Arabidopsis ADF1; closed
triangles, S6D mutant ADF1; closed
circles, human ADF. Inset, turbidimetric
recording of time courses of spontaneous polymerization of 16 µM ADP·actin in the absence (bottom
curve) and presence (top curve) of 20 µM human ADF.
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Effect of Phosphorylation on ADF Function and Interaction with
F-actin--
The binding of S6D mutant ADF1 to F-actin was
examined by the fluorescence and sedimentation assays. The S6D mutant
ADF1-elicited depolymerization of F-actin was very weak in
the concentration range investigated (0-20 µM).
Fluorescence titration curves of NBD-labeled F-actin by S6D mutant
ADF1 were consistent with an order of magnitude lower
affinity of S6D mutant ADF1 than the wild-type protein for
actin (data not shown). S6D mutant ADF1 bound NBD-labeled
F-actin in a kinetically cooperative fashion like the wild-type
protein. The binding process was 10-fold slower than with wild-type
ADF1. Specifically, when 3 µM NBD-labeled F-actin was reacted with 12 µM ADF1, the lag
time was 18 ms, and the t1/2 of the reaction was 60 ms. When it was reacted with 12 µM S6D mutant
ADF1, the lag time was 200 ms, and the
t1/2 was 0.72 s (data not shown).
Although S6D mutant ADF1 binds G- and F-actins with low
affinity, the question should be raised whether the biological activity is affected by the mutation. Fig. 9 shows
that S6D mutant ADF1 enhances the treadmilling of actin
filaments as all ADFs do (7); however, the effect develops in the
higher range of protein concentration at which the mutant protein binds
to F- and G-actins. In conclusion, the strength of binding of ADF is
weakened by phosphorylation, but the mechanism of binding remains
unchanged. In addition, once bound to actin, the S6D mutant protein
affects the dynamics of filaments in the same manner as the wild-type
protein.

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Fig. 9.
Phosphorylation of ADF lowers its affinity
for actin, but does not affect its function and effect on
treadmilling. The steady-state ATPase activity of F-actin (15 µM) in ATP-containing F buffer at pH 7.6 was measured
following addition of wild-type (WT) ADF1
(squares) or S6D mutant ADF1
(circles) at the indicated concentrations.
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DISCUSSION |
Human ADF and Arabidopsis ADF1, two distant
variants of the ADF/cofilin family, interact with F- and G-actins and
affect actin dynamics in almost identical fashion. The main conclusion
of the direct binding studies using unmodified ADF is that ADF binds essentially ADP·actin with an affinity of 108
M
1 and practically not ATP·actin under
cellular ionic conditions. Therefore, ADF cannot be considered as a
G-actin-sequestering protein as previously thought. The high affinity
of ADF for ATP·G-actin, which was derived from the shift in critical
concentration plots (12, 13), relied on the unverified hypothesis that
ADF was forming a 1:1 complex with ATP·G-actin exclusively and that
this complex was in equilibrium with free ATP·G-actin at the same
critical concentration as in the control curve obtained in the absence of ADF. This interpretation is invalidated by the present and past (7)
results showing that ADF in fact interacts with both ADP·G-actin and
ADP·F-actin, which changes the dynamics of actin and incidentally
affects the steady-state concentration of
ATP·G-actin.2
The fluorescence of NBD-labeled G-actin is not quenched upon binding of
human ADF as it is upon binding of ADF1, indicating that
the environment of Cys-374 on G-actin (but not on F-actin) is not
affected in the same way by the two ADFs. The mutagenesis studies of
Lappalainen et al. (21) show that the G-actin-binding region
in ADF/cofilin contains in particular the loop between strand
6 and
helix
4. The sequence of human ADF diverges significantly from other
ADFs (including ADF1) in this region, which may explain the
difference in the environment of the NBD-derivatized Cys-374 of actin
when different ADFs are bound.
Analysis of the properties of S6D mutant ADF1, which mimics
phosphorylated ADF1, demonstrates that phosphorylation does
not affect the biological activity of ADF per se or its
mechanism of interaction with actin, but reduces its affinity for both
G- and F-actins by 20-fold. The effect of phosphorylation then is equivalent to a 20-fold decrease in the concentration of active protein. However, once bound to actin, S6D mutant ADF1
affects actin dynamics like the unmodified protein. These results
expand the conclusions derived from genetic studies (21) that indicated that phosphorylation did not grossly affect the structure of ADF.
Analysis of the kinetics of interaction of ADF with G- and F-actins is
important in determining which sequence of elementary steps is going to
take place and modify actin dynamics in a living cell, immediately
following reception of a signal leading to ADF activation. This work
shows that binding of ADF to F-actin occurs first, followed (within
2-3 min) by the establishment of a new steady state in which filaments
turn over faster, with an increased accumulation of monomeric actin,
consisting of ADF·ADP·G-actin, ADP·G-actin, and ATP·G-actin.
ADF interacts with ADP·G-actin in rapid equilibrium. The large pool
of dissociated ADF·ADP·G-actin generates an increased amount of
ADP·G-actin and of ATP·G-actin as well, via nucleotide exchange.
Within this view, the increased treadmilling rate is thought to be due
to the increased steady-state concentration of
ATP·G-actin.2 At high ADF concentration, the
ADF·ADP·G-actin complex is stabilized; hence, the decline in
treadmilling rate (Fig. 9) is presumably due to the association flux of
ADF·ADP·G-actin to filament ends, which counteracts and limits the
dissociation flux. The depolymerization induced by ADF, in
ATP-containing F buffer, is limited by the polymerization of
ADF·ADP·actin, which is consistently shown to occur at critical
concentrations of 3.5 and 8 µM for Arabidopsis and human ADFs, respectively (Fig. 8).
ADF binds to and dissociates from F-actin relatively slowly. In
particular, the rate of dissociation of ADF from F-actin is slower than
the rate of depolymerization of ADF·F-actin from the pointed end at
steady state. Therefore, F-actin-bound ADF does not rapidly shuttle
from one filament to the other. Interestingly, ADF binds F-actin in a
kinetically cooperative fashion. The binding of the first molecules of
ADF to a bare filament resembles a nucleation process during which the
structure of the filament is locally disrupted, thus allowing the more
facile subsequent association of other ADFs with the "opened" sites
along the filament. The binding then occurs linearly at a faster rate,
in a zipper-like mode. There is some analogy here to the vectorial
process of ATP hydrolysis on Mg·F-actin (34). The functional
consequence of such a cooperative binding process is that the
enhancement of actin dynamics by ADF (7) occurs essentially on a
fiber-by-fiber basis. In other words, when ADF is added to F-actin in a
1:10 ratio, at the end of the binding process, roughly 10% (in number) of the filaments are entirely decorated by ADF, and 90% are bare, as
was noticed on the electron micrographs (14). Since the ADF-decorated filaments depolymerize rapidly, ADF then visits other filaments in the
population in a treadmilling-assisted process.
The assembly of ADF·F-actin from the high affinity
ADF·ADP·G-actin complex is a simple case of reversible
polymerization. It is remarkable that nucleation of ADP·actin is
greatly facilitated by ADF, whereas the critical concentration for
assembly is increased. This apparent paradox indicates that, although
the actin-actin bonds involved in filament elongation are globally
weakened by ADF, the energetically unfavorable actin-actin bonds
involved in nucleation are strengthened by ADF. Structural and
biochemical studies as well as the atomic model of the actin filament
suggest that the longitudinal actin-actin bonds play a major role in
filament stability (35-37) and that the lateral bonds are involved in
filament nucleation (38-41). Within this view, in binding to F-actin,
ADF would bridge two neighboring subunits along the small pitch helix, thus stabilizing lateral actin-actin bonds, whereas longitudinal bonds
would be destabilized in ADF-decorated filaments, thus accounting for
an overall increase in critical concentration. The destabilization of
longitudinal bonds would account for the contorted appearance (7, 42)
and increased twist (14) of ADF-decorated filaments. The low viscosity
of solutions of ADF·F-actin is also consistent with a decreased
rigidity due to the destabilization of longitudinal bonds.
Similar conclusions can be derived from the analysis of the kinetics of
ADF binding to F-actin maintained in different structural states by
phalloidin, by proteolytic modification, or by capping by gelsolin. The
highly ordered structure of the filament stabilized by phalloidin or
Pi/BeF3
is not favorable
for ADF binding. In contrast, ADF binds faster and in a non-cooperative
fashion to gelsolin-capped filaments or to filaments assembled from
subtilisin-cleaved actin, which both display a bridge of density
connecting the two strands of the long pitch helix (27), which suggests
that the structure of the filaments in which lateral bonds are
strengthened is favorable for ADF binding. If ADF were binding
preferentially to bent regions of filaments, as proposed to account for
its putative severing function (11), opposite results would have been
obtained, i.e. ADF would have bound less easily to short
rod-like gelsolin-capped filaments and to subtilisin-cleaved F-actin,
which has a large persistence length (30). The cooperative change in
structure of the filament linked to ADF binding, accompanied by a
torsional movement (14) and a possible decrease in length, is expected to cause a large mechanical constraint if the filament is immobilized between two anchorage points. This constraint may result in breakage of
the filament, which has in fact been observed in video microscopy experiments when filaments were immobilized on myosin heads in rigor
prior to addition of ADF (11). In conclusion, our work brings more
support to the view (28, 29, 37, 43) that the variability of the
F-actin structure can be modulated by actin-binding proteins.
The fact that ADF binds faster to gelsolin-capped filaments may be
physiologically relevant in actin-based motility, as follows. The
forward movement of the leading edge is thought to be due to the
steady-state barbed end growth of non-capped filaments at the membrane,
whereas filaments that dissociate from the membrane become capped and
undergo depolymerization from their pointed ends at the rear of the
lamellipodium (15). The present results suggest that ADF may enhance
the treadmilling process supporting protrusion of the lamellipodium by
binding selectively to the capped filaments and activating their
depolymerization, whereas the uncapped filaments at the front would be
less accessible to ADF and therefore would keep a stiffer structure
appropriate for pseudopod extension. The presence of rigid stretches of
F-actin·ADP·Pi subunits (30) at the growing barbed
ends, which remain ADF-free, would further enhance the functional
sorting of filaments. Further experiments will have to be designed to
test the possibility that, in modulating the structure of the
filaments, ADF may as well generate spatial order in the cell,
e.g. in selecting different types of myosin motors to
translocate particles to and from different subcellular locations.