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INTRODUCTION |
Actin filaments, in conjunction with other cytoskeletal proteins,
are responsible for maintaining the structural integrity of eukaryotic
cells. Phagocytosis, cytokinesis, cell motility, and muscle contraction
all depend on structures assembled from actin. Although muscle
contraction requires stable actin filaments for force generation, much
actin-based motility involves active turnover of filaments. A variety
of actin-binding proteins influence actin polymerization, steady-state
dynamics of monomers and filaments, and the three-dimensional
organization of the filamentous meshwork. At the leading edge of motile
cells, the Arp2/3 complex of actin-related proteins is thought to
initiate polymerization of a network of actin filaments (1, 2) that
turn over on a time scale of minutes (3, 4). This requires the
filaments to depolymerize much more rapidly than pure actin filaments
in vitro. Proteins of the ADF/cofilin family are thought to
promote recycling of actin (for review, see Refs. 5 and 6), because
they enhance the dynamics of actin filaments in Listeria
comet tails (7, 8) and promote depolymerization (7, 9, 10).
A major point of disagreement is whether ADF/cofilin proteins enhance
depolymerization by severing actin filaments, by increasing the rate of
subunit dissociation from one or both ends, or by both severing and
rapid subunit dissociation (reviewed in Ref. 11). Early papers (12-17)
proposed that ADF/cofilin family proteins depolymerize actin filaments
by creating more ends to dissociate subunits. Several lines of evidence
support severing: direct visualization (14), electron microscopy (18,
9), viscometry (18, 14, 9), kinetic analysis of spontaneous
polymerization (10), and fluorescence recovery after photobleaching
(10). On the other hand, Carlier and colleagues (7, 19, 20) argue that the effects of plant or human ADF on muscle actin filaments might be
explained entirely by increased rates of subunit association and
dissociation at the ends of filaments.
To help resolve this controversy, we have done new experiments with
Acanthamoeba actophorin, a member of the ADF/cofilin family. Because the interaction of ADF/cofilin proteins with actin involves a
number of overlapping reactions with actin monomers and actin filaments, our strategy has been to divide the problem into a number of
steps that we can examine in isolation before trying to formulate an
overall mechanism. Ours is also the first systematic comparison of the
interaction of any ADF/cofilin protein with cytoplasmic actin and
muscle actin filaments, an important point, because some of the
reactions differ for the two actins. Our first paper (21) included
equilibrium and rate constants for actophorin binding ATP- and
ADP-actin monomers, for the polymerization of actophorin-actin
complexes, and for the antagonistic effects of actophorin and profilin
(another small protein) on the exchange of nucleotide bound to actin monomers.
Here, we characterize the kinetics and equilibrium binding of
actophorin to actin filaments. Actophorin has a much higher affinity
for muscle actin filaments than amoeba actin filaments. Binding to
muscle actin filaments, but not amoeba actin filaments, is cooperative.
Phosphate or BeF3 bound to ADP-actin filaments inhibits
actophorin binding, so ATP hydrolysis and phosphate dissociation serve
as a timer for binding. Reciprocally, actophorin increases the rate of
phosphate release after ATP hydrolysis. An assay for the number of
filament ends shows that actophorin severs actin filaments in a
concentration- and time-dependent manner. The 10-fold higher affinity of actophorin for ADP-actin monomers than for ADP-actin filaments provides the free energy change to drive depolymerization.
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MATERIALS AND METHODS |
Reagents--
Materials came from the following sources: Sigma,
dithiothreitol (DTT),1 EDTA,
Tris, sodium azide, Me2SO, hexokinase, ATP, ADP,
phalloidin, Sephadex G-25 medium; Molecular Probes (Eugene, OR),
tris-(2-carboxyethyl)phosphine, tetramethylrhodamine maleimide 5'
isomer; Whatman (Maidstone, United Kingdom), DEAE-cellulose DE-52.
Preparation and Labeling of Actophorins--
Wild type and S88C
mutant actophorins (21) in plasmid vector pMW172 were expressed in
Escherichia coli strain BL21 (DE3) pLysS and purified
according to Quirk et al. (22) with 2 mM dithiothreitol in all buffers to avoid cysteine oxidation. Purified actophorins were stored in 10 mM Tris-Cl, pH 7.5, 1 mM EDTA, 2 mM DTT, 1 mM
NaN3. Actophorin S88C was labeled with tetramethylrhodamine maleimide 5' isomer and purified (21).
Other Proteins--
Actin was purified from rabbit skeletal
muscle acetone powder (23) or from Acanthamoeba (24), and
monomeric Ca-ATP-actin was isolated by Sephacryl S-300 chromatography
(25) at 4 °C in G buffer (5 mM Tris-Cl, pH 8.0, 0.2 mM ATP, 0.1 mM CaCl2, 0.5 mM DTT). Actin was labeled on Cys-374 to a stoichiometry of
0.8-1.0 with pyrene iodoacetamide (Ref. 26; as modified by Pollard; see Ref. 24). Mg-ATP G-actin was prepared on ice by addition of 0.2 mM EGTA and an 11-fold molar excess of MgCl2
over actin and used within hours. Actin was polymerized by addition of
1:9 (v/v) 10× KME (500 mM KCl, 10 mM
MgCl2, 10 mM EGTA, 100 mM Tris-Cl, pH 8). ADP-BeF3-actin filaments were prepared by
polymerizing 20 µM Mg-ATP-actin in 0.1 M KCl,
2 mM MgCl2, 5 mM NaF, 150 µM BeCl2 at room temperature for 4 h
(27).
Assays for Interaction of Actophorin with Actin
Filaments--
Interaction of actophorin with pyrenyl-labeled actin
filaments was followed by the change in fluorescence with excitation at
366 nm and emission at 387 nm (7). Interaction of
rhodamine-S88C-actophorin with unlabeled actin filaments was followed
by the change in fluorescence anisotropy with excitation at 550 nm and
emission at 574 nm (21). Data were collected with a Alphascan
spectrofluorometer (Photon Technology International, South Brunswick,
NJ). Reactions were initiated by mixing actophorin and actin filaments
manually, or for kinetics experiments with a hand-driven stopped-flow
mixer (Model SFA-12, Hi-tech Scientific Ltd., Salisbury, United Kingdom).
Assay to Measure the Rate of Phosphate Release--
We used the
2amino-6-mercapto-7-methyl purine riboside and purine-nucleoside
phosphorylase assay to measure the release of phosphate during
spontaneous polymerization of actin alone or in presence of actophorin
or phalloidin (28, 29). We first prepared the 1:1 complex Ca-ATP-actin
monomers by treatment of Ca-ATP-actin monomers with AG 1-X4 Resin
(Bio-Rad). Mg-ATP actin monomers were prepared on ice by addition of
0.2 mM EGTA and an 11-fold molar excess of
MgCl2 over the 1:1 complex Ca-ATP-actin monomers and used
within hours. Actin assembly and phosphate release were monitored in
400 µl samples after addition of 1:9 (v/v) 10× KME with a Microplate
Spectrophotometer system, SECTRAmax 250 (Molecular Devices Corporation,
Sunnyvale, CA) using absorbance at 310 nm and 360 nm, respectively. The
increase in absorbance at 360 nm induced by actin polymerization was
used as a blank and subtracted from the absorbance change due to
phosphate release.
Elongation Assay for Actin Filament Number
Concentration--
The assay was carried out in two steps. First,
actophorin was reacted with preformed 2 µM unlabeled
actin filaments for a given time at 25 °C. Then the reaction mixture
was diluted to 0.66 µM polymerized actin in KME buffer,
and 5 µM pyrenyl-actin monomers were added to initiate
growth from the ends of the filaments. Under these conditions, the rate
of spontaneous polymerization is much slower than elongation from the
added filaments. The number concentration of actin filaments
(N) was calculated from the initial rate of polymerization
according to Equation 1,
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(Eq. 1)
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where k+ is the association rate constant
(10 µM
1 s
1),
k
is the dissociation rate constant (1 s
1), and [A] is the concentration of actin monomers.
Under the conditions of this assay, the initial rate was directly
proportional to the concentration of untreated actin filaments.
Data Analysis and Simulation of Kinetics--
Kinetic data were
fit to exponentials with Kaleidagraph software (Synergy Software,
Reading, PA). Binding data were fit with Equation 2,
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(Eq. 2)
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where r is the observed anisotropy,
rcf is the anisotropy of free actophorin,
rcb is the anisotropy of actophorin bound to actin
filaments, [C] is the total concentration of actophorin, [A] is the
total concentration of actin filaments, and Kd is
the dissociation equilibrium constant of the complex.
We used KINSIM (30) to simulate the time course of reactions of
actophorin with actin filaments.
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RESULTS |
Interaction of actin filaments with ADF/cofilin proteins,
including Acanthamoeba actophorin, is complicated by several
overlapping reactions, including filament binding (31, 14, 16, 17, 7,
9, 19), a structural change in the filament (32), severing the filament
(14, 9), and depolymerization of the filament (7, 19). To study these
reactions, we used two fluorescence assays: the fluorescence intensity
of pyrenyl-labeled actin and fluorescence anisotropy of
rhodamine-labeled S88C actophorin. Pyrenyl fluorescence of polymerized
actin is about 20-fold higher than actin monomers (26) and is quenched
by binding of some ADF/cofilin proteins, including actophorin (7). A
complication is that actophorin can change the fluorescence of
pyrenyl-actin filaments by three different mechanisms: (i) binding,
(ii) conformational change, or (iii) depolymerization. With proper
experimental design, these reactions can be separated in time.
Fluorescence anisotropy is sensitive only to the size of the diffusing
species, so binding of rhodamine-actophorin to an actin filament
increases anisotropy more than binding to an actin monomer. The
anisotropy of rhodamine actophorin is 0.14 when free and 0.21 when
bound to an actin monomer (21), whereas the polarization anisotropy of
rhodamine-phalloidin immobilized on an actin filament is about
0.30.2
Reaction of Actophorin with Actin Filaments Followed by
Fluorescence Anisotropy--
Mixing rhodamine-actophorin with an
excess of unlabeled actin filaments results in a biphasic change in
fluorescence anisotropy (Fig.
1A). The first phase is an
exponential increase from 0.14 ± 0.005 to a maximum of 0.235 ± 0.005. The rate and amplitude of this phase depend on the
concentration of actin filaments. The rate constant for the initial
increase in fluorescence anisotropy is in the range of 0.03 µM
1 s
1, the same as measured
more accurately (due to less noisy data) by quenching of the
fluorescence of pyrenyl-actin (Fig. 2).
The second phase is a slow exponential decrease in anisotropy with a
rate constant of 0.012 ± 0.002 s
1, independent of
the concentration of actin filaments. Our interpretation, elaborated
upon under "Discussion," is that the first phase is due to binding
of rhodamine-actophorin to actin filaments, whereas the second phase
results from severing and depolymerization of filaments.

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Fig. 1.
Fluorescence anisotropy assay for interaction
of rhodamine S88C actophorin with actin filaments. Conditions were
as follows: 10 mM Tris-HCl, pH 8.0, 50 mM KCl,
2 mM MgCl2, 1 mM EGTA, 0.2 mM ATP, 0.2 mM CaCl2, 0.5 mM DTT, and 3 mM NaN3 at 25 °C.
A, time course of the change in fluorescence anisotropy upon
mixing 0.4 µM rhodamine-S88C-actophorin with 7 µM muscle actin filaments. The noisy curve is
experimental data. Smooth curves are the best single
exponential fits to the rising phase and falling phase of fluorescence
anisotropy. B, dependence of the maximum fluorescence
anisotropy on the concentration of actin filaments. Closed
circles, 2 µM rhodamine-S88C-actophorin plus
Acanthamoeba actin filaments; open circles, 0.5 µM rhodamine-S88C-actophorin plus muscle actin filaments.
Solid lines are the best fits of Equation 2 to the
data.
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Fig. 2.
Fluorescence assay of the time course of the
interaction of unlabeled actophorin with an excess of pyrene-labeled
actin filaments. Conditions were as follows: 10 mM
Tris-HCl, pH 8.0, 50 mM KCl, 2 mM
MgCl2, 1 mM EGTA, 0.2 mM ATP, 0.2 mM CaCl2, 0.5 mM DTT, and 3 mM NaN3 at 25 °C. A, time course
of the change in fluorescence following mixing 0.4 µM
wild type actophorin with 5 µM Acanthamoeba
pyrenyl-actin filaments. The inset shows the initial rapid
decrease in fluorescence on a faster time scale. Noisy
curves are experimental traces. Smooth curve is the
best fit using model 2 and values in Table I. B, dependence
of kobs for the fast phase of the fluorescence
change on the concentration of pyrenyl-actin filaments: closed
circles, Acanthamoeba pyrenyl-actin filaments;
open circles, muscle pyrenyl-actin filaments.
Inset, time course of the dissociation of actophorin from
muscle pyrenyl-actin filaments by competition with unlabeled actin
filaments. A mixture of 2 µM of pyrenyl-actin filaments
and 0.4 µM of actophorin was preincubated for 5 min and
then mixed with 20 µM unlabeled actin filaments to
initiate dissociation of actophorin from the pyrenyl-actin filaments.
C, time course of the change in fluorescence following
mixing 0.6 µM wild type actophorin with 6 µM muscle pyrenyl-actin filaments: dotted
line, in presence of 0.2 mM ADP; noisy
line, in presence of 0.2 mM ATP. Smooth
curves are the best fit using model 2 and values in Table I.
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The amplitude of the maximum increase in fluorescence anisotropy after
mixing rhodamine-actophorin with unlabeled actin filaments depends on
the concentration of filaments (Fig. 1B). A plot of the
maximum amplitude versus the concentration of filamentous actin gives a saturation curve that fits Equation 2. At saturating concentrations of actin filaments the anisotropy is 0.235, higher than
rhodamine-actophorin bound to actin monomers (21) but lower than
expected for an actin filament. The apparent dissociation equilibrium
constant from this analysis depends on the source of the actin. The
affinity of actophorin for rabbit skeletal muscle actin filaments
(Kd = 0.49 µM) is 10-fold higher than for Acanthamoeba actin filaments (Kd = 5.6 µM).
Binding of Actophorin to Actin Filaments Using Fluorescence of
Pyrenyl-actin--
Some ADF/cofilin proteins quench the fluorescence
of pyrenyl-actin filaments. However, this is far from a simple
bimolecular reaction, so the results depend on many variables,
including the source of the actin (Fig. 2B), the presence of
the
-phosphate of ATP bound to actin (see Fig. 4), the ratio of the
reactants (Figs. 2 and 3), and the
presence of ATP or ADP in the buffer (Fig. 2C). These
complications arise from major differences in the actin, including
different affinities and degree of binding cooperativity, but also from
reactions that follow binding, including severing, depolymerization,
and repolymerization.

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Fig. 3.
Fluorescence assay of the time course of
interaction of pyrene-labeled actin filaments with an excess of
unlabeled actophorin. Conditions were as follows: 10 mM Tris-HCl, pH 8.0, 50 mM KCl, 2 mM MgCl2, 1 mM EGTA, 0.2 mM ATP, 0.2 mM CaCl2, 0.5 mM DTT, and 3 mM NaN3 at 25 °C.
A, time course of the change in fluorescence following
mixing of 2 µM pyrenyl-amoeba-actin filaments with
various concentrations of actophorin: 15, 20, 30, and 40 µM from the top to bottom curve,
respectively. B, time course of the change in fluorescence
following mixing of 2 µM pyrenyl-muscle-actin filaments
with various concentrations of actophorin: 4, 6, 8, 10, 12, and 15 µM from the top to bottom curve,
respectively. Inset shows the early time course.
Smooth curves are the best fit using model 1. C, dependence
of kobs of the fluorescence change on the
concentration of actophorin. In the absence of phosphate: open
circles, muscle actin; closed squares, amoeba actin;
and with 68 mM phosphate: closed circles, muscle
actin. The solid lines are manual fits to the data points.
The data point at zero actophorin comes from Fig. 2B.
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Reaction of actophorin with an excess of pyrenyl-actin filaments in ATP
is biphasic: the fluorescence falls rapidly to a value intermediate
between that of polymerized and monomeric actin, followed by a slow
recovery (Fig. 2, A and C), similar to the two
phase change in fluorescence anisotropy of rhodamine-actophorin. Reaction of an excess of actophorin with pyrenyl-actin causes a
monotonic decrease in fluorescence to the level of monomeric actin
(Fig. 3, A and B).
With excess pyrenyl-actin filaments the rapid initial phase is an
exponential decrease in fluorescence (Fig. 2A, inset) at a
rate that depends on the concentration of actin filaments (Fig. 2B). Like Carlier et al. (7), we interpret this
decrease in fluorescence to be due to binding of actophorin to actin
filaments. Plots of the observed rate constant versus
concentration of Acanthamoeba actin filaments are linear
with a slope (association rate constant, k+1) of
0.029 ± 0.003 µM
1 s
1
and a y-intercept (dissociation rate constant,
k
1) of 0.11 ± 0.005 s
1.
The ratio of these rate constants gives a Kd of 3.8 µM for actophorin binding amoeba actin filaments. For
muscle actin, the slope (k+1) is 0.008 ± 0.0005 µM
1 s
1 and the
y-intercept is near zero, too small to estimate accurately by extrapolation. To measure the rate constant for actophorin dissociation from muscle actin filaments, we carried out a chase experiment (Fig. 2B, inset). We bound actophorin to
pyrenyl-actin filaments and then competed it off by adding an excess of
unlabeled actin filaments. The pyrene fluorescence increased during the chase with a rate constant of 0.0050 ± 0.0005 s
1,
independent of concentration of unlabeled actin filaments. Unlabeled actin filaments chase ADF-1 from NBD-labeled actin filaments at a rate
of 0.035 s
1 (19). We interpret this to be the rate of
dissociation of actophorin from the pyrenyl-actin filaments. For muscle
actin, the Kd calculated from the rate constants is
0.6 µM. Both of these equilibrium constants derived from
kinetics agree well with the values from fluorescence anisotropy.
After samples with an excess of pyrenyl-actin filaments reached a
minimum fluorescence, the fluorescence recovered slowly in the presence
of ATP (Fig. 2, A and C). The fluorescence
increased exponentially with a rate constant of 0.015 ± 0.001 s
1 for amoeba actin (Fig. 2A) and 0.0030 ± 0.0005 s
1 for muscle actin (Fig. 2C). The
fluorescence did not recover when the buffer contained ADP rather than
ATP (Fig. 2C).
When an excess of unlabeled actophorin was mixed with amoeba
pyrenyl-actin filaments (Fig. 3A) or muscle pyrenyl-actin
filaments (Fig. 3B), the fluorescence decreased to the level
of monomeric actin and did not recover even in ATP. With amoeba
pyrenyl-actin filaments (Fig. 3A), the time courses fit
single exponentials, and the dependence of the
kobs on the concentration of actophorin was
linear, as expected for a simple bimolecular reaction (Fig. 3C). The slope and intercept give the same rate constants as
the experiment with excess filaments (Fig. 2B).
In contrast, the time course of the reaction of an excess of unlabeled
actophorin with muscle pyrenyl-actin filaments does not fit a single
exponential (Fig. 3B). An initial lag of a few seconds was
followed by a progressively faster decrease in fluorescence to baseline
level. The lag was most pronounced at low concentrations of actophorin.
Both the duration of the lag and the rate of the subsequent decline in
fluorescence depend on the actophorin concentration. In contrast to
experiments with excess pyrenyl-actin filaments (Fig. 2), the
dependence of kobs for the decline in
fluorescent after the lag phase on actophorin was not linear (Fig.
3C). As elaborated upon under "Discussion," these
features suggest that binding of actophorin to muscle actin filaments
is cooperative.
Inhibition of Actophorin Binding to Pyrenyl-actin Filaments by
Phosphate and BeF3--
Maciver et al. (14)
found that inorganic phosphate inhibits the ability of actophorin to
reduce the low shear viscosity of actin filaments but did not establish
whether this was due to inhibition of binding or some subsequent
reaction, such as severing or depolymerization. The half maximal effect
was at 5-10 mM phosphate, similar to the
Kd of 1 mM for phosphate binding to
ADP-actin filaments (33). New kinetic experiments with saturating
phosphate (68 mM) established that actophorin and phosphate
compete for binding muscle ADP-actin filaments (Fig. 4). Mixing an excess of actophorin with
muscle pyrenyl-actin filaments in 68 mM phosphate caused a
slow exponential decrease in fluorescence to the level of monomeric
actin. In contrast to samples without phosphate, the rate is much
slower and there is no lag. As in the absence of phosphate, the
observed rate constants in 68 mM phosphate vary nonlinearly
with actophorin concentration, but the rates are 10 times smaller (Fig.
3C).

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Fig. 4.
Effect of phosphate and BeF3 on
the rate of actophorin binding to pyrene-labeled muscle actin
filaments. Conditions were as follows: 10 mM Tris-HCl,
pH 8.0, 50 mM KCl, 2 mM MgCl2, 1 mM EGTA, 0.2 mM ATP, 0.2 mM
CaCl2, 0.5 mM DTT, and 3 mM
NaN3 at 25 °C with 68 mM potassium phosphate
(stock pH 8.0) or 150 µM BeF3. Dotted
lines, time course of the fluorescence change following mixing of
2 µM pyrenyl-actin filaments in 68 mM of
potassium phosphate with various concentrations of actophorin: 0, 5, 9, 11, and 13 µM from top to bottom.
The thick line is the time course following mixing of 2 µM pyrenyl-ADP-BeF3 actin filaments with 20 µM actophorin. Solid lines are the best single
exponential fits to the data. Fig. 3C (closed
circles) shows the dependence of kobs in 68 mM phosphate on actophorin concentration.
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BeF3 binds ADP-actin filaments with an affinity 3 orders of
magnitude higher than phosphate (27) and completely inhibits the effect
of 20 µM actophorin on the fluorescence of pyrenyl-actin filaments (Fig. 4). The result is similar with filaments stabilized with phalloidin, explaining the ability of phalloidin to protect filaments from actophorin (14). Pelleting experiments confirm that no
actophorin binds actin filaments saturated with BeF3 or phalloidin, ruling out the possibility that actophorin binds without changing the fluorescence of pyrenyl actin.
Actophorin Increases the Rate of Phosphate Release from Actin
Filaments--
The 2-amino-6-mercapto-7-methyl purine
riboside-phosphorylase assay (28, 29) showed that actophorin increases
the rate of phosphate dissociation. Without actophorin, phosphate
release lags behind polymerization, with a rate constant of 0.0022 s
1 (Fig. 5A, in
good agreement with Ref. 29). With a high concentration of actophorin,
phosphate release keeps pace with polymerization, so the actual rate of
phosphate release is faster than the observed rate of 0.032 s
1 (Fig. 5B). According to kinetic
simulations, the minimum rate of phosphate release from filaments
saturated with actophorin is 0.04 s
1 and could be much
faster. Note that actophorin increases the absorbance change used to
follow polymerization. The actophorin concentration dependence of
phosphate dissociation suggests a Kd of about 20 µM for actophorin binding ADP-Pi filaments (Fig. 5C). Addition of actophorin to actin filaments just at
the completion of spontaneous polymerization induces a burst in
phosphate release. In contrast, phalloidin strongly inhibits the rate
of phosphate release in agreement with previous work (34).

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Fig. 5.
Effect of actophorin on the rate of phosphate
release from polymerizing actin. Conditions were as follows: 20 µM Mg-ATP-muscle-actin with varying concentration of
actophorin, 10 mM Tris-HCl, pH 8.0, 50 mM KCl,
2 mM MgCl2, 1 mM EGTA, 0.5 mM DTT, and 3 mM NaN3 at 25 °C.
Noisy thick lines, actin polymerization followed by
absorbance at 310 nm. Noisy thin lines, phosphate release
followed by absorbance at 360 nm. Smooth curves are
exponential fits to the data. Actophorin concentrations were 0 (A) and 40 (B) µM. C,
variation of the rate constant for phosphate release with actophorin
concentration (closed circles). The solid line is
the best manual fit to the data.
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Actophorin Severs Actin Filaments--
A kinetic assay for actin
filament ends confirms that actophorin severs actin filaments (Fig.
6). The experiment consisted of two
steps. First, unlabeled actin filaments were reacted with a range of
actophorin concentrations or with a fixed concentration for a range of
times. These reaction mixtures were used to seed the polymerization of
5 µM pyrene-muscle-actin monomers. This assay measures
the number concentration of filament ends, because the rate of
polymerization is proportional to the concentration of ends, because
the concentration of actophorin in the elongation assay is much lower
than the concentration of pyrenyl-actin monomers, and because
actophorin has little or no effect on the rate of elongation of ATP
actin monomers (21). The fact that the extent of elongation is
unaffected by actophorin (Fig. 6A) is further evidence that
these concentrations of actophorin do not affect the elongation
reactions at either end of filaments.

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Fig. 6.
Actin filament elongation assay for the
effect of actophorin on the number concentration of actin
filaments. Conditions were as follows: both the preincubation and
elongation assays were carried out in 10 mM Tris-HCl, pH
8.0, 50 mM KCl, 2 mM MgCl2, 1 mM EGTA, 0.2 mM ATP, 0.2 mM
CaCl2, 0.5 mM DTT, and 3 mM
NaN3 at 25 °C. The elongation assay contained 5 µM pyrenyl-actin monomers and a 1:3 dilution of the
preincubation mixture. A, time course of pyrenyl-actin
elongation from a reaction mixture of 2 µM unlabeled
muscle actin filaments alone (cross) or 2 µM
actin filaments preincubated with 1 µM actophorin for the
following times: closed circles, 10 s; open
squares, 60 s; closed squares, 120 s;
open circles, 300 s; closed triangles,
600 s; open triangles, 1200 s. Thick
line, time course of polymerization of 2 µM of
muscle pyrenyl-actin. The number concentration of filaments at the end
of the preincubation was calculated from the initial rate of elongation
using Equation 1. B, dependence of the relative rate of
polymerization and the number concentration of filaments in
A on the time of preincubation (closed circles),
and dependence of the number concentration of filaments after 5 min of
preincubation on the concentration of actophorin in the preincubation
mixture with 2 µM unlabeled actin filaments (open
circles).
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Incubation with actophorin increases the rate of elongation from
preformed actin filaments. The increase in elongation rate depends on
the duration of preincubation of the preformed actin filaments with
actophorin (Fig. 6, A and B) and at a fixed time point on the concentration of actophorin (Fig. 6B). We
interpret the increase in the rate of elongation as an increase in the
concentration of filament ends. Assuming no effect on elongation rate
constants, we used Equation 1 to calculate the number concentration of
actin filaments from the initial rate of polymerization. The number of
muscle actin filament ends increased to a maximum after 5 min and then
declined (Fig. 6B). The number of filaments at 5 min increased with the concentration of actophorin up to 10-fold at 4 µM actophorin (Fig. 6B). Maciver et
al. (9) carried out a similar experiment with qualitatively
similar results with actophorin and human ADF. They observed a
transient 5-fold increase in the number of ends after 30 s with
equimolar actophorin and muscle actin filaments.
It was more difficult to resolve the time course and measure the
concentration of ends in kinetic experiments with
Acanthamoeba actin. Ten seconds of preincubation of
actophorin with amoeba actin filaments increased the rate of elongation
in the second phase of the assay, but the effect was transient. After
600 s, the elongation rate seeded by the mixture of actophorin and
amoeba actin filaments was less than the rate of untreated filaments. This difference, considered in more detail under "Discussion," may
be due to faster binding, severing, and depolymerization of Acanthamoeba actin filaments by actophorin.
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DISCUSSION |
Binding of Actophorin to Actin Filaments--
Mossakowska and Korn
(35) and our previous paper (21) were among the first reports comparing
the interactions of an ADF/cofilin protein with actins from the same
cell and from muscle. Because of its convenience, most laboratories use
skeletal muscle actin (7, 9, 12, 16, 17, 19, 20, 36). Two studies of
yeast cofilin used yeast actin (37, 10) but did not make a systematic
comparison with earlier work with muscle actin (38). Our results show
that a full understanding of the physiological properties of
ADF/cofilin proteins requires experiments with homogeneous systems.
Although actophorin has the same affinity for actin monomers from
muscle and amoeba (21), its interactions with muscle and amoeba actin
filaments differ qualitatively and quantitatively. For example, binding
to muscle actin filaments is cooperative, but binding to amoeba actin
filaments is not. The affinity for muscle ADP-actin filaments is 10 times higher than amoeba actin filaments, due to major differences in
both the association and dissociation rate constants. The association
rate constant is 3 times higher for actophorin binding amoeba ADP-actin
filaments than muscle actin filaments, but actophorin dissociates 20 times faster from amoeba actin. This difference explains why Cooper et al. (18) found that no stable association between
actophorin and amoeba actin filaments. We do not know whether other
ADF/cofilin proteins interact the same or differently with their own
actin and muscle actin.
Kinetics of the Interaction of Actophorin with Actin
Filaments--
Binding of actophorin to actin filaments is complicated
by severing and depolymerization of the filaments and influenced by dissociation of the
-phosphate released by hydrolysis of the bound
ATP. Furthermore, binding to muscle actin filaments is cooperative. Fortunately, using a combination of assays and varying the ratios of
the reactants, it is possible to separate all of these overlapping events in time. The following sections address each phase of the reaction.
Simple Binding of Actophorin to Excess ADP-actin
Filaments--
This is the most straight forward reaction, with
similar results from both fluorescence anisotropy of
rhodamine-S88C-actophorin binding unlabeled actin filaments and
unlabeled actophorin binding pyrenyl-actin filaments. Excess of actin
minimizes the effect of interactions between actophorins bound near to
each other on the same filament. The time course follows a single
exponential, and kobs is directly proportional
to the concentration of actin filaments, as expected for a bimolecular
reaction. The slope gives association rate constants of
k+1 of 0.029 µM
1
s
1 for amoeba actin ADP-actin filaments and 0.008 µM
1 s
1 for muscle ADP-actin
filaments. The dissociation rate constant for amoeba actin, estimated
from the y-intercept, is 0.11 s
1, giving the
same binding constant as that measured in equilibrium experiments by
fluorescence anisotropy (Fig. 1). The ratio of the dissociation rate
constant from muscle actin, estimated from a chase experiment (Fig.
2B, inset), to the association rate constant also agrees
well with the equilibrium experiment. The affinity is 10 times higher
for muscle actin than amoeba actin filaments, due largely to the slower
dissociation of actophorin from muscle actin filaments. These
association rate constants are remarkably low, orders of magnitude
slower than binding of myosin (9 µM
1
s
1) (39),
-actinin (2.5 µM
1 s
1) (40, 41), or gelsolin
(20 µM
1 s
1) (42) to actin
filaments. To account for the slow association rate, we postulate that
bare ADP-actin filaments have few open actophorin binding sites.
Phalloidin also binds slowly because filaments rarely sample the
conformation with an open binding site between the subunits (43,
44).
Excess Actophorin Binding to ADP-actin Filaments--
Excess
actophorin binds amoeba actin filaments in a simple bimolecular
reaction (Fig. 3A), with rate constants the same as in
experiments with excess actin. This shows that binding is not cooperative. On the other hand, when an excess of actophorin binds muscle ADP-actin filaments, the reaction is complicated by effects of
bound actophorin on subsequent binding reactions, as observed for ADF
(19). This positive cooperativity accelerates binding after an initial
lag of <1 s (Fig. 3B). Fortunately, an excess of actophorin
saturates the filaments and reduces the pyrenyl fluorescence to the
level of monomeric pyrenyl-actin, so that subsequent reactions
(including severing and depolymerization) do not produce a signal to
complicate further the analysis of the association reaction.
The simplest interpretation (19) is that initial slow binding of
actophorin accelerates the binding of subsequent actophorins to the
same filament. A two-step mechanism (Scheme
1) accounts for our data.
This model assumes that initial slow binding of actophorin to
muscle actin causes a conformational change in the filament that
accelerates further binding of actophorin. C is the concentration of
actophorin, F is the concentration of polymerized actin, *F represents
polymerized actin with a conformational change induced by actophorin,
and *FC represents the complex with actophorin. The rate of the
conformational change induced by actophorin is proportional to
k+1 (rate constant measured for actophorin binding an excess ADP-actin filaments (Fig. 2)) and some power of the
concentration of actophorin. The rate constant for actophorin binding
to *F is an unknown. The first reaction in Scheme 1 is written as a
catalytic, and the concentration of C consumed in forming *F is
ignored, because most C is free in this experiment, so the initial slow
binding inducing the actin filament conformational change does not
affect actophorin concentration.
We varied the two unknowns until computer simulations of the kinetic
curves matched the time course of the change in fluorescence over a
range of actophorin concentrations (Fig. 3B). The initial lag was best fit when the rate of the first step was equal to the
product of the measured association rate constant and the square of the
actophorin concentration. Similar to the results of Ressad et
al. (19) with plant and human ADF, neither the first nor the third
power of the concentration gave good fits. As in their model, the
initial binding of two actophorins in close proximity induces a
conformational change in the filament that accelerates further binding
of actophorin. The subsequent time course was fit best with an
association rate constant, k+2 of 0.075 µM
1 s
1, 10 times larger than
the rate constant for actophorin binding bare muscle ADP-actin
filaments, but only 2.5 times the rate constant for the noncooperative
binding of actophorin to bare amoeba ADP-actin filaments. This
mechanism includes the same concepts as a model proposed for plant and
human ADF binding to muscle ADP-actin filaments (19), although the
rates are substantially lower for actophorin than ADF. The
conformational change induced by actophorin may correspond to the
change in the twist of the filament described by McGough et
al. (32), although additional work is required to prove this point.
This mechanism is attractive for several reasons. First, most subunits
in bare actin filaments are in the standard conformation, whereas
thermal motion allows a few ADP-subunits to sample the twisted
conformation. Second, because binding of ADF/cofilin proteins favors
the twisted conformation, then the twisted conformation is likely to
favor binding of ADF/cofilin proteins. Thus, the standard conformation
may have few sites favoring ADF/cofilin binding, explaining the very
slow association rate. Further, binding of a few ADF/cofilin proteins
will favor (trap) the twisted conformation, making adjacent sites
available for rapid binding, explaining the cooperative binding. If
bound ATP, ADP-Pi, and phalloidin all favor the standard
conformation over the twisted conformation, their inhibition of binding
is explained.
This cooperative mechanism applies to ADF/cofilins binding to muscle
actin filaments, but is it relevant to physiology? The cooperatively
for amoeba actin filaments is far less and binding to bare filaments is
nearly as fast as to activated muscle actin filaments. New experiments
will be required to learn whether this is a general property of
cytoplasmic actin filaments.
Actophorin Severs Actin Filaments--
Elongation experiments
(Fig. 6 and Ref. 9) confirm earlier evidence that actophorin and other
ADF/cofilins sever actin filaments. Severing depends on the
concentration of actophorin, the type of actin, and time. In 5 min, 4 µM actophorin severs each filament into about 10 pieces.
This is similar to the extent of severing observed by Maciver et
al. (9) with actophorin in a similar assay and by Du and Frieden
(10) with yeast cofilin and actin using photobleaching recovery to
estimate length. The accumulation of muscle actin filament ends due to
severing plateaus after 400 s. Several factors may contribute to
the plateau: short filaments may not sever as readily as long filaments
(9), or severing may continue but be balanced by reannealing or the
disappearance of short filaments, because actophorin does not cap and
stabilize severed ends as does gelsolin.
Reactions Subsequent to Actophorin Binding to Actin
Filaments--
Under certain conditions our fluorescence assays
revealed that additional reactions
severing, depolymerization and
repolymerization
follow actophorin binding to actin filaments. We do
not understand most of these reactions nearly as well as binding, but
we know enough to model them approximately. The slow drop in
fluorescence anisotropy (Fig. 1) after binding of rhodamine-actophorin
to unlabeled actin filaments indicates that the actophorin either
dissociates from the filaments or, more likely, that the ADP-actin to
which it is bound is either in a smaller filament (due to severing) or has depolymerized. Neither depolymerization nor severing is
apparent when actophorin is in excess over pyrenyl-ADP-actin filaments, because binding alone reduces the pyrenyl fluorescence to the level of
actin monomers (Fig. 3). Thus, neither severing nor depolymerization causes further change. However, with excess actin, fluorescence drops
only part of the way to the monomer fluorescence (Fig. 2). In ADP, the
fluorescence is steady at this intermediate level, but in ATP the
fluorescence recovers slowly toward the level of polymerized actin.
Others have observed similar recoveries in pyrenyl fluorescence (45,
7). Aizawa et al. (45) did not comment, and Carlier et
al. (7) suggested that the slow increase might be due to a
redistribution of plant ADF from low affinity binding to pyrenyl-actin
subunits to high affinity binding to unlabeled actin subunits in the
filaments. Our binding experiments using fluorescence anisotropy or
pyrene fluorescence show that actophorin has a similar affinity for
unlabeled (Kd = 5.9 µM) and 100%
pyrene-labeled (3.8 µM) actin filaments, ruling out that explanation.
After actophorin binds ADP filaments, the pyrenyl fluorescence recovers
partially in presence of ATP. Our interpretation is that the
intermediate fluorescence is due to a steady state that includes
accelerated depolymerization of ADP-actin filaments, exchange of ADP on
dissociated monomers for ATP and repolymerization of ATP-actin to form
filaments that transiently do not bind actophorin owing to their
content of ATP and ADP-Pi subunits. To test this hypothesis, we simulated a single cycle reaction of actophorin with
pyrenyl-ADP-actin filaments followed by recovery using the mechanism in
Scheme 2,
where C is actophorin, A is actin monomer, T is ATP, D is ADP, P
is Pi, E is filament ends, and F is polymerized actin.
Although this scheme appears complicated, most of the rate and
equilibrium constants are known (Table I)
except for k
6 (the rate of dissociation of
actophorin-ADP-actin subunits from filaments), k
12 (the rate of dissociation of ADP-actin
from filaments partially saturated with actophorin), and
k+9 (the rate of severing).
Simulations of the time course of pyrenyl-actin fluorescence fit the
observations remarkably well (Fig. 2, A and C),
using the measured rate constants and reasonable values for the number of ends present at the beginning of the experiments (0.6 nM
for muscle actin and 2.9 nM for amoeba actin) and estimates
of unknown rate constants: k+9 = 0.05 s
1 and k
12 = 40 s
1
for amoeba actin; k+9 = 0.02 s
1
and k
12 = 20 s
1 for muscle
actin. The values of k+9 agree well with the rate of severing measured directly k+9 = 0.012 s
1(Fig. 6B). In experiments with an excess of
actin filaments over actophorin, most depolymerizing subunits are
ADP-actin rather than actophorin-ADP-actin subunits. Consequently, the
data do not constrain the value of k
6. In
agreement with these observations, the simulations show no recovery
with ADP. In ATP, dissociating ADP-actin exchanges ADP for ATP, and
owing to the low critical concentration for ATP actin, it repolymerizes
at the ends of the numerous severed filaments with a concomitant increase of pyrene fluorescence. ATP hydrolysis and phosphate release
are slow enough for the polymer concentration to recover partially
without rebinding actophorin. The shape of the curves in Fig. 2,
A and C, constrains k
12
to between 30 and 50 s
1 for amoeba actin and between 15 and 25 s
1 for muscle actin. Varying the value of each of
the unknown parameters more than 30% yielded theoretical curves that
failed to fit the full set of experimental curves, even if the values
of other unknowns were varied in a compensatory fashion.
The estimated rate constants are reasonably robust and agree well with
previous work (20, 10). The value proposed for k
12 is the same order of magnitude as the rate
of depolymerization of ADP-actin subunits from the barbed ends, but
more than 1 order of magnitude larger than dissociation from the
pointed ends without ADF/cofilin binding. Available evidence suggests
that ADF/cofilins promote both severing and depolymerization from
pointed ends (7, 19, 20) and perhaps barbed ends as well. Severing and
depolymerization reactions still need more work, but the large
difference in affinity of actophorin for amoeba ADP-actin monomers
relative to ADP-actin filaments provides a thermodynamic basis for
severing amoeba actin more effectively than muscle actin filaments,
which bind actophorin almost as tightly as muscle actin monomers.
Phosphate Dissociation as a Timer for Polymer
Destruction--
Occupation of the
-phosphate site in actin
filaments by phosphate or BeF3 inhibits actophorin binding.
This explains how millimolar concentrations of phosphate protect actin
filaments from the effects of actophorin on the low shear viscosity
(14, 9) and of ADF1 on binding and depolymerization (7). A lack of
actophorin binding to ADP-Pi filaments and phalloidin-ADP
filaments in pelleting assays establishes that the absence of a
fluorescence signal is due to a lack of binding rather than failure of
bound actophorin to induce a fluorescence change in pyrenyl
ADP-Pi-actin filaments.
Phosphate and actophorin compete for binding ADP-actin filaments. As
expected from detailed balance, BeF3 is a much stronger inhibitor of actophorin binding, because it binds much more tightly to
the
-phosphate position than phosphate. The thermodynamic relationships remain to be determined by further experiments, but the
kinetic consequences are already clear: tightly bound BeF3
reduces the rate of actophorin binding more than 2 orders of magnitude,
weakly bound phosphate inhibits the rate of actophorin binding more
than 10-fold, and actophorin stimulates the rate of phosphate release
more than 10-fold. Actophorin binding may change the subunit
conformation to favor the proposed "back-door" pathway of phosphate
dissociation (46).
Competition between actophorin and phosphate makes binding of excess
actophorin to ADP-Pi muscle actin filaments (Fig. 4) qualitatively different from binding to muscle ADP-actin filaments (Fig. 3B). Binding is not only much slower but is also less
cooperative. The time course follows a single exponential and the
dependence of kobs on actophorin concentration
gives an association rate constant of 0.003 µM
1 s
1, three times lower
even than the initial slow binding of actophorin to excess ADP-actin
filaments (Fig. 2). At no concentration did we observe a lag followed
by rapid cooperative binding, so overall, the rate of binding is 15 times slower to ADP-Pi actin filaments than ADP-actin
filaments at 15 µM actophorin (Fig. 4).
Obviously, the situation during active polymerization in the presence
of actophorin is complicated, because not only do ATP hydrolysis and
phosphate release regulate actophorin binding, but actophorin affects
phosphate release. Further work is required to understand the detailed
relationships and their consequences for actin filament severing and
depolymerization. At the very least, by enhancing phosphate release
actophorin accelerates the cycle of polymerization depolymerization.
Phosphorylation of ADF/cofilins by LIM kinase inhibits binding to actin
filaments (47, 48)3 and is
another potential avenue of regulation.