doi:10.1016/j.jmb.2005.09.021
J. Mol. Biol. (2005) 353, 990–1000
Cofilin Increases the Torsional Flexibility and Dynamics
of Actin Filaments
Ewa Prochniewicz1, Neal Janson2, David D. Thomas1 and
Enrique M. De La Cruz2*
1
Department of Biochemistry
Molecular Biology and
Biophysics, University of
Minnesota, Minneapolis, MN
55455, USA
2
Department of Molecular
Biophysics & Biochemistry
Yale University, New Haven
CT 06520, USA
We have measured the effects of cofilin on the conformation and dynamics
of actin filaments labeled at Cys374 with erythrosin-iodoacetemide (ErIA),
using time-resolved phosphorescence anisotropy (TPA). Cofilin quenches
the phosphorescence intensity of actin-bound ErIA, indicating that binding
changes the local environment of the probe. The cofilin concentrationdependence of the phosphorescence intensity is sigmoidal, consistent with
cooperative actin filament binding. Model-independent analysis of the
anisotropies indicates that cofilin increases the rates of the microsecond
rotational motions of actin. In contrast to the reduction in phosphorescence
intensity, the changes in the rates of rotational motions display non-nearestneighbor cooperative interactions and saturate at substoichiometric cofilin
binding densities. Detailed analysis of the TPA decays indicates that cofilin
decreases the torsional rigidity (C) of actin, increasing the thermally driven
root-mean-square torsional angle between adjacent filament subunits from
w48 (CZ2.30!10K27 Nm2 radianK1) to w178 (CZ0.13!10K27 Nm2
radianK1) at 25 8C. We favor a mechanism in which cofilin binding shifts
the equilibrium between thermal ErIA-actin filament conformers, and
facilitates two distinct structural changes in actin. One is local in nature,
which affects the structure of actin’s C terminus and is likely to mediate
nearest-neighbor cooperative binding and filament severing. The second is
a change in the internal dynamics of actin, which displays non-nearestneighbor cooperativity and increases the torsional flexibility of filaments.
The long-range effects of cofilin on the torsional dynamics of actin may
accelerate Pi release from filaments and modulate interactions with other
regulatory actin filament binding proteins.
q 2005 Elsevier Ltd. All rights reserved.
*Corresponding author
Keywords: cofilin; actin; cooperativity; anisotropy; torsional rigidity
Introduction
Members of the ADF/cofilin family of actinbinding proteins sever actin filaments1–3 and may
accelerate subunit dissociation from the pointed-
ends.4 Severing and enhanced depolymerization
are likely to arise from cofilin-mediated changes in
actin filament structure. ADF/cofilin binding
changes the average twist and subunit tilt of a
filament,5,6 increases the disorder of subdomain 27
Abbreviations used: TPA, transient phosphorescence anisotropy; ErIA, erythrosine iodoacetamide; t, triplet excitedstate lifetime of ErIA; I, phosphorescence emission intensity; Ivv, vertically polarized component of the phosphorescence
emission; Ivh, horizontally polarized component of the phosphorescence emission; A, amplitude; G, instrument
correction factor; R, anisotropy; r0, initial anisotropy; rN, final anisotropy; f, rotational correlation time; qa, angle
between the absorption dipole of bound ErIA and the filament axis; qe, angle between the emission dipole of bound ErIA
and the filament axis; hDxi, rms average fluctuation of the torsion angle between adjacent filament subunits; li, filament
length distribution; k, amplitude reduction factor; a, actin filament radius; h, long-axis height of an individual actin
filament subunit; C, torsional rigidity; a, torsional spring constant; g, rotational frictional coefficient; Dk, rotational
diffusion coefficient; kB, Boltzmann’s constant; h, solvent viscosity.
E-mail address of the corresponding author:
enrique.delacruz@yale.edu
0022-2836/$ - see front matter q 2005 Elsevier Ltd. All rights reserved.
991
Cofilin Affects Actin Filament Torsional Dynamics
and the DNase-binding loop,8 disrupts the interface
between subdomains 1 and 2 along the long-pitch
helix of filaments9 and weakens stabilizing lateral
contacts in the filament.10,11 The change in filament
twist is propagated along the filament to actin
subunits without bound cofilin.6,7
Cofilin binding to actin filaments is cooperative5
and can be described by a nearest-neighbor
cooperative binding model.12 Cooperative interactions may be mediated through conformational
rearrangement of subdomain 2 of actin,12 which
facilitates local changes in subunit tilt observed by
electron microscopy.7
Actin filaments display two types of large-scale
movements: long-axis (flexural) bending and longaxis (torsional) twisting (Figure 1). Bending and
twisting depend on the bound nucleotide and
cation,13,14 interactions with actin-binding proteins,13–16 and actin isoforms.17,18 The flexural
rigidity of actin filaments is larger than the torsional
rigidity so filaments twist more easily than they
bend.
The cofilin-induced change in average filament
twist5,6 may modulate the mechanical properties of
actin filaments. We are testing this hypothesis using
transient phosphorescence anisotropy (TPA). TPA
of erythrosin-labeled actin has been successfully
applied to describe the microsecond timescale
dynamics of actin filaments and structural changes
in the region of actin’s C terminus.15,16,18–22 We have
previously shown that (1) TPA of actin cannot be
accounted for by rigid body rotations, but reflects
primarily intra-filament torsional motions that can
be explained in terms of the torsional twist
model, and (2) changes in the phosphorescence
intensity monitor local conformational changes in
actin.15,16,18–20 Here, we demonstrate that cofilin
binding increases the microsecond dynamics and
the torsional flexibility of actin filaments, and that
these effects can propagate to vacant sites on the
filament.
Figure 1. Conformational dynamics of actin filaments.
Schematic representation of actin filament thermal
motions.
Results
Effect of cofilin on time-resolved phosphorescence intensity decays of actin filaments
The phosphorescence intensity decays of
erythrosine iodoacetamide (ErIA)-actin filaments
(Figure 2) were analyzed by fitting data to a sum of
two exponentials (equation (3)) with amplitudes I1
and I2, and the corresponding lifetimes, t1 and t2.
The observed phosphorescence intensity decay is
dominated (I1 w0.8) by a long lifetime (t1Z
w220 ms) component, with a small contribution (I2
w0.2) from a shorter lifetime (t2Zw30 ms, “intermediate”) component. The multiple components
contributing to the decay imply that there exist
multiple (at least two) actin conformations with
microsecond lifetimes. The amplitudes and lifetimes of both the long (t1) and intermediate (t2)
lifetime components are reduced in a cofilin
concentration-dependent manner (i.e. both intensity decays are more rapid with bound cofilin; data
not shown). There is also a rapidly decaying
component (t/4 ms, short lifetime) in the presence
of cofilin, as indicated by the reduction in initial
phosphorescence intensities (Figure 2, inset). We
refer to the amplitude of this rapidly decaying
phase as I3 and the lifetime as t3.
Cofilin binding increases the population of short
(t3) and the intermediate (t2) lifetime components,
and reduces the population of the long lifetime (t1)
species (Figure 3). The mole fractional distribution
of these three components depends sigmoidally on
the cofilin concentration, consistent with cooperative cofilin binding to actin filaments.5,9,12 The
reciprocal partitioning of the short and long lifetime
components (X1 and X2; Figure 3) suggests that
these two states are in equilibrium, and that cofilin
binding shifts the equilibrium distribution of these
two states. Detailed balance requires that if cofilin
shifts the equilibrium between two conformations,
it must bind preferentially to one of the two.
Although equilibrium cofilin binding to actin
filaments is well-described by a nearest-neighbor
Figure 2. Effect of cofilin on the time-resolved
phosphorescence intensity decays of ErIA-actin filaments.
The [ErIA-actin] is 1.8 mM and the [cofilin] are: 0 (black),
0.3 (royal blue), 0.8 (red), 1.0 (green), 1.5 (violet), 2.0
(orange), and 6.0 (light blue) mM. The inset shows the
same data with the time axis plotted on a log scale.
992
Cofilin Affects Actin Filament Torsional Dynamics
anisotropy arise from the modulation of internal
microsecond filament dynamics.
Model-independent analysis: sum of exponentials
Figure 3. Cofilin concentration-dependence of the
observed intensity decay mole fractions (X). (a) Mole
fractions of the slow (X1), intermediate (X2) and fast (X3)
decays contributing to the phosphorescence decays.
(b) Inset of data in (a) to show the fast decaying
component. The continuous lines are drawn for clarity
and have no physical significance.
The effect of cofilin on actin dynamics was first
analyzed in terms of a model-independent fit of the
anisotropy decays to the sum (nZ2) of exponentials
(equation (5)) to obtain the rotational correlation
times (fi), the initial anisotropy (r0) and the final
anisotropy (rN). The correlation times (fi; i.e.
reciprocal rate constants) characterize the rates of
rotational motions, the initial phosphorescence
anisotropy (r0) characterizes the amplitude of local
sub-microsecond motions of the dye and/or
cysteine 374 relative to actin (a lower value means
more mobile), and the final anisotropy (r N)
characterizes the angular amplitudes of microsecond timescale filament motions (i.e. global
motions).
The initial phosphorescence anisotropy of actinbound ErIA (r0Z0.105) is significantly lower than
that of immobilized ErIA (r0Z0.20519) due to fast,
submicrosecond motion of the dye relative to actin.
Cofilin binding reduces the initial anisotropy (r0) in
a non-stoichiometric manner (Figure 5(a)). The
best fits of the data to equation (11) show that a
single bound cofilin affects the initial anisotropy of
90 G24 actin subunits. Thus, the cofilin-mediated
effect can be propagated to vacant subunits in the
cooperativity model,12 the multiple equilibrium
conformations complicate the nearest-neighbor
analysis.
Effect of cofilin on time-resolved phosphorescence anisotropy (TPA) of actin filaments
Cofilin binding has a significant effect on the
phosphorescence anisotropy decays of ErIA-actin
filaments (Figure 4). Light-scattering and sedimentation demonstrate that cofilin does not depolymerize actin under our experimental conditions (data
not shown,12). Therefore, cofilin-induced changes in
Figure 4. Effect of cofilin on the time-resolved
phosphorescence anisotropy decays of ErIA-actin filaments. The [ErIA-actin] is 1.8 mM and the [cofilin] are: 0
(blue), 0.15 (red), 0.8 (green), 1.5 (violet) mM. The inset
shows the same data with the time axis plotted on a log
scale.
Figure 5. Cofilin concentration-dependence of the
initial (r0) and final (rN) anisotropies of ErIA actin. The
continuous lines represent the best fits to equation (11)
and indicate that cofilin affects the initial anisotropy of 90
G24 subunits and the final anisotropy of O400 subunits.
Uncertainty bars represent standard deviations from data
sets collected on four separate days. The actin concentration was 1.8 mM. The binding density was estimated
from the data presented in Figure 3, which are
comparable to our previous determinations with pyrene
actin filaments.12
Cofilin Affects Actin Filament Torsional Dynamics
filament (i.e. there are non-nearest-neighbor
cooperative interactions). This range of
cooperativity represents a lower limit, because the
phosphorescence lifetimes and the total intensity
signal (Figure 2) decrease with added cofilin.
Consequently, the effect of cofilin on the observed
anisotropy is less than the effect on the theoretical
anisotropy (equation (7)), so the cooperative unit is
larger than estimated from the concentrationdependence of the observed anisotropy reduction
(i.e. if quenching did not occur, the slope would be
steeper). The low signal-to-noise ratio at high
[cofilin], due to quenching of phosphorescence
intensity (Figure 2), decreases the reliability of the
anisotropy data as cofilin concentration
approaches saturation. The maximum observed
decrease in the initial anisotropy (r0), from 0.105
G0.007 in actin to 0.069 G0.016 in cofilactin
filaments (Figure 5(a)), indicates an increase in the
half-cone angle of the submicrosecond wobble (qc)
from 37(G2) 8 in actin to 47(G4) 8 in actin filaments
saturated with cofilin (Equation (6)).
The final anisotropy (rN) reaches a minimum at a
cofilin binding density /0.1 (Figure 5(b)), indicating that the cofilin-induced increase in the angular
amplitude of the microsecond rotational motions is
also cooperative. The best fit of the data has a large
uncertainty but indicates that the final anisotropy of
several hundred actin subunits (427G355) are
affected by bound cofilin. This maximal effect of
rN at substoichiometric cofilin binding densities is
consistent with long-range, non-nearest-neighbor
cooperative interactions modulating the internal
dynamics of actin filaments. Filament severing,
predicted to occur at low binding densities and
cluster sizes,12 would also lower rN.
Model-dependent analysis: torsional twist
The effect of cofilin on actin dynamics was
analyzed in terms of the torsional twist model
(equation (8)). In this model, the actin filament is
regarded as an array of cylindrical elementary rods
and the observed anisotropy decay results from the
combined global motions, which represent overall
rigid-body tumbling and intrafilament twisting
motions. Therefore, to determine the intrafilament
torsional constant (a) from the TPA data (equation
(7)) accurately, corrections for rigid-body motions
must be made. Thermal bending motions, which
occur on the millisecond timescale, are too slow to
be detected by TPA. Similarly, rigid-body tumbling
motions can contribute to the microsecond TPA
decay only for those filaments that are much shorter
than the typical 4 mm long filaments observed in the
absence of cofilin (Figure 6).15
Since cofilin-induced fragmentation of actin
could produce filaments that tumble on the timescale of TPA measurements, the equilibrium length
distribution (li) of actin filaments at saturating
cofilin was measured by electron microscopy
(Figure 6) and included in the fit function (equation
(8)), allowing us to separate the effect of rigid-body
993
Figure 6. Length distribution of ErI-actin filaments. The
[ErIA-actin] is 1.8 mM and the [cofilin] are: 0 (a) and 4.0 (b)
mM.
rotations of short actin-cofilin filaments on TPA
from the intrafilament motions. The filament radius
is constrained when fitting the data to determine
the torsional constant (a), so our determination of a
was limited to bare and fully decorated filaments,
which have well-defined homogenous radii. The
radius of an actin filament (a) was taken to be 4.5!
10K9 m based on the maximum diameter of 90–95 Å
reported by Holmes.23 The diameter of a cofilindecorated actin filament was determined to be 6.7!
10K9 m by measuring the width of the bare and
cofilin-decorated actin filament.5 The fit value for
the torsional constant (a) increases by a factor of
about 2 when the assumed radius of bare actin is
reduced from 4.5 nm to 3.5 nm.24 With cofilinsaturated actin, when the assumed radius is
reduced from 6.7 nm to 5.7 nm, the torsional
constant increases by w50%. Therefore, although
the best fit values of the torsional constant (a)
depend on assumptions of actin filament radii, the
uncertainty in determining a is no more than a
factor of 2.
Cofilin binding has minimal effects on the
orientations of absorption and emission dipoles of
ErIA (Table 1), but lowers the spring constant (a)
and thus the torsional rigidity C (calculated as ah)
by a factor of about 20 (Table 1). This indicates that
while cofilin-induced changes in the average actin
filament subunit orientation, as detected from the
ErIA probe at the actin C terminus, are subtle (!108
tilt with respect to the long filament axis), filaments
with bound cofilin are torsionally more flexible,
displaying larger and more rapid microsecond
twisting motions than native actin filaments.
Actin filaments are best described as a continuous
elastic rod.19 However, if the continuous elasticity is
expressed as an elasticity per subunit rise in the
994
Cofilin Affects Actin Filament Torsional Dynamics
Table 1. Effects of cofilin on the structure and dynamics of ErIA-actin filaments
Filament
d
Actin
Cofilactinf
[cofilin]/
[actin]
0
2.2
qa (deg)a
e
44.2 (G1.7)
44.5 (G0.3)
qe (deg)a
C (N m2 radianK1)a
a (N m radianK1)b
hDxic
41.9 (G3.5)
38.1 (G0.5)
2.30 (G1.00)!10
0.13 (G0.06)!10K27
8.36 (G3.6)!10
0.48 (G0.22)!10K19
4.08
16.88
K27
K19
Conditions: 1.8 mM ErIA-actin filaments in 50 mM KCl, 2 mM MgCl2, 2 mM DTT, 0.2 mM ATP, 1 mM NaN3, 20 mM imidazole (pH 6.6),
25 8C.
a
Determined by fitting of the data to equation (7) (subunit height Z5.5 nm).
b
Calculated using equation (9) and a subunit height (h) of 2.75 nm; 1 Nm radianK1Z107 dyn cm radianK1.
c
Calculated using equation (10).
d
Calculated using a filament radius of 4.5 nm.
e
Uncertainties representGone standard deviation from three separate data sets.
f
Calculated using a filament radius of 6.7 nm.
filament (hZ2.75!10K9 m), the root mean squared
average thermal fluctuation of the torsion angle
(hDxi) between adjacent subunits in a filament (i.e.
angular disorder) can be calculated from the
torsional constant (equation (10)25). Cofilin binding
increases the amplitude of the actin subunit torsion
angle fluctuations from 4.08 to 16.88 at 25 8C
(Table 1). This cofilin-induced fourfold increase in
hDxi is independent of the subunit height h, since a is
inversely proportional to h (equation (10)).19 The 48
torsional fluctuation of bare actin filaments
measured here is comparable to the 5–68 angular
disorder observed by electron microscopy image
analysis,26 and more recently using total internal
reflection fluorescence polarization microscopy.27
Effect of subsaturating phalloidin on cofilin binding
and filament dynamics
The ability of phalloidin to cooperatively stabilize
intermonomer bonds in actin28 makes it a useful
tool to examine the role of intersubunit interactions
in the actin–cofilin interaction. Saturating phalloidin has minimal effects on the intensity (Figure 7)
and anisotropy (Figure 7(a) inset) decay of ErIAactin filaments, consistent with previous reports.21,29
Saturating amounts of phalloidin protect actin from
the cofilin-dependent decrease in phosphorescence
intensity and anisotropy (data not shown) because
cofilin does not bind phalloidin-stabilized actin. At
subsaturating phalloidin concentrations (0.1 phalloidin per actin), cofilin can bind actin and
quenches the phosphorescence intensity but minimally affects the changes in anisotropy (Figure 7(b)
inset). This behavior suggests that cofilin binds to
phalloidin-free regions on the filament, locally
quenching the phosphorescence of ErIA, but
phalloidin-induced long-range stabilizing effects
on intermonomer contacts dampen cofilin-induced
changes in torsional flexibility.
The kinetics of cofilin binding to actin filaments
was assayed from the quenching of pyrene actin
fluorescence.12,30 Time-courses of 11.8 mM cofilin
binding to actin filaments displayed a brief lag
phase (Figure 7(c) inset). For simplicity, we treated
the relaxation as a single process even though this is
not an accurate reaction mechanism30 (E.D.L.C. and
W. Cao, unpublished results). The observed time-
course of cofilin (11.8 mM) binding to actin could be
approximated by an exponential with an observed
rate constant of w1.6 sK1. Time-courses of cofilin
binding to phalloidin-stabilized actin filaments
follow double exponentials (Figure 7(c) inset).
Bound phalloidin slowed the observed rate constant of the fast phase and subsaturating phalloidin
concentrations generated maximal inhibition
(Figure 7(c)). The best fit of the data (Figure 7(c))
indicates that a single bound phalloidin inhibits
cofilin binding to 8.6 G0.5 actin subunits in a
filament. This inhibition can be explained by the
stabilizing effect of phalloidin, which extends 10–20
filament subunits.28 The observed rate constants of
the slow phases are 0.002–0.01 sK1, which may be
limited by phalloidin dissociation. Contributions
from photobleaching make it difficult to analyze the
slow rates reliably.
Discussion
Relationship between phosphorescence
intensity, anisotropy and the conformation of
ErIA-actin filaments
The analysis of the effect of cofilin on phosphorescence intensity and anisotropy of ErIA-actin
indicates that cofilin has both local and long-range
effects on actin’s structure and dynamics. Cofilininduced local structural changes in the environment
of the C terminus are indicated by quenching of the
phosphorescence intensity of actin-bound ErIA
(Figures 2 and 3). These changes are probably
facilitated by the proximity (11–30 Å) of cofilin
binding sites to the label on Cys374.5 The subtle
effect of cofilin on the absorption and emission
dipoles of ErIA-actin filaments (Table 1) suggests
that its binding occurs with minimal (!108) tilting
of the ErIA probe with respect to the filament axis,
but the observed decrease in phosphorescence
intensity suggests structural changes in the environment of Cys374 that increase the exposure of the
actin-bound ErIA to quenching by the solvent.
Increased exposure of the dye to the solvent is also
supported by the increased amplitude of the
nanosecond wobbling motions, which are too fast
for the motions of whole monomers, but are
Cofilin Affects Actin Filament Torsional Dynamics
Figure 7. Effect of substoichiometric phalloidin concentrations on cofilin-dependent changes in phosphorescence intensities and anisotropies. (a) Phosphorescence
intensity decay of actin filaments (blue) and actin
filaments saturated with phalloidin (red). Inset: Anisotropy decay of actin filaments (blue) and actin filaments
saturated with phalloidin (red). Only the first 300 ms are
shown for clarity. (b) Phosphorescence intensity decay of
actin filaments (blue), actin filaments with a molar
equivalent of cofilin (green), and actin filaments equilibrated with a molar equivalent of cofilin and 0.1 molar
equivalent of phalloidin (red). Inset: Anisotropy decay of
actin filaments (blue), actin filaments with a molar
equivalent of cofilin (green), and actin filaments equilibrated with a molar equivalent of cofilin and 0.1 molar
equivalent of phalloidin (red). Only the first 300 s are
shown for clarity. (c) Phalloidin concentration-dependence of the observed rate constant for 11.8 mM cofilin
binding to actin filaments as assayed from the quenching
of pyrene fluorescence. The apparent stoichiometry (n)
obtained from the best fit is 0.116 G0.007 phalloidin
bound per actin. The continuous line is the best fit to
equation (12). Uncertainty bars are within the symbols.
The inset shows time-courses of fluorescence quenching
after mixing 23 mM cofilin with pyrene actin filaments
containing (a) 0, (b) 0.1, (c) 0.4, or (d) 0.8 molar equivalent
of bound phalloidin. The final [ErIA-actin] is 1.8 mM in (a)
and (b) and 0.85 mM in (c).
995
compatible with increased mobility of the C
terminus. These local cofilin-induced structural
changes could contribute to the reported changes
at the interface between subdomain 1 and 29 of
adjacent filament subunits, and because of conformational coupling between subdomain 1 and 231
within an individual subunit, to changes in
subdomain 2 conformation.7
It has been proposed that cofilin binding shifts
the equilibrium distribution of thermal conformers.6,7 The reciprocal, [cofilin]-dependent partitioning of the short and long phosphorescence
lifetime conformational states (Figure 3) is consistent with this hypothesis. However, the cofilininduced increase in torsional amplitude between
adjacent filament subunits (hDxi; Table 1) suggests
that cofilin binding also allows actin filaments to
sample novel conformational states by changing the
filament torsional stiffness. In contrast, cofilin
markedly reduces the variability in subunit
torsion angles observed by electron microscopy.6
This observation suggests that electron
microscopy is sampling the long-range cumulative
component of the angular variability, while the
spectroscopic measurements report the local
rotations of an actin subunit or domain about the
helical axis.
The cofilin-induced cooperative change in phosphorescence anisotropy and the large decrease in
torsional rigidity are likely to reflect the change in
filament twist. Differential scanning calorimetry32
favors a mechanism where cofilin binding
destabilizes (i.e. lowers the thermal transition) the
filament lattice cooperatively, as would be expected
from the increased torsional flexibility at substoichiometric cofilin concentrations. Our spectroscopic observations are also consistent with the
results of electron microscopy, which showed that
cofilin cooperatively changes the twist of the actin
filament.6
The binding of cofilin induces long-range
cooperative changes in the microsecond timescale
actin filament dynamics (Figure 5). The nonnearest-neighbor effects on torsional dynamics
may contribute to the acceleration of Pi33 release
and weak Pi binding of cofilin-actin filaments, and
may influence interaction with other regulatory
proteins, particularly those that are sensitive to the
nucleotide state of actin filaments such as the Arp2/
3 complex.34
Long-range cooperative changes in actin filaments are expected to decrease (i.e. dampen) as the
distance from bound cofilin increases. The number
of subunits that could be affected would be dictated
by the energy change of the two twisted conformations and the energy associated with cofilin
binding. The product of the number of affected
subunits and the free energy change of the
transition could not exceed the free energy associated with cofilin binding. Therefore, the observation
that dozens of subunits are affected by an
individual cofilin molecule favors a mechanism
996
where the twist conformations are comparable in
energy, perhaps thermal conformers.
Comparison of methods used to measure the
torsional rigidity of actin filaments
Several methods have been used to estimate the
angular disorder in the torsion angle between
adjacent filament subunits (hDxi) and the torsional
rigidity (C) of actin filaments, including
electron microscopy,24,35,36 electron paramagnetic
resonance,37,38 transient absorption and phosphorescence anisotropy,15,16,18–21 visualization of
rotational motions of beads attached to actin
filaments,39,40 and total internal reflection
fluorescence polarization microscopy. 27 The
values for the torsional rigidity of actin
filaments obtained with these methods range from
w2!10K27 Nm2 15,16,18–21,24,27,35–38 to w5!10K26
Nm2.39,40 This study estimates the torsional rigidity
of actin filaments in solution as 2.3!10K27 Nm2
(Table 1), comparable to previous spectroscopy,15,16,18–21,37,38 electron microscopy24,35,36
and single-molecule measurements.27 Higher
values were obtained with micromanipulation
methods, using large beads in an optical trap,39,40
which more than likely lead to an overestimate of
torsional rigidity.27
Implications for actin filament severing
The observed local changes at the actin C
terminus may account for the effect of cofilin on
actin filament stability. Normal mode analysis of the
actin filament led to the conclusion that the
torsional flexibility of the whole filament could be
significantly affected by reorientation of only a few
residues, such as in the region of the hydrophobic
plug and subdomain 2;23,41–43 such torsionally
strained filaments fragment more easily than
unloaded filaments.39 Cofilin binding to the C
terminus of actin would interfere with formation
of stabilizing longitudinal contacts established by
subdomains 1 and 2 of adjacent subunits.5–7,9,23,28,44,45
Because subdomain 2 also forms lateral interstrand
contacts through contact with residues 262–274,23,46,47
reorganization of subdomain 2 upon cofilin binding7,8 would disrupt the formation of stabilizing
lateral 10,11 filament interactions as well. By
destabilizing subdomain 2 interactions in the
filament, cofilin makes the overall intersubunit
contacts less stiff. Thus, cofilin serves as a molecular
lubricant that allows actin filaments to adopt
otherwise inaccessible conformations. The combination of compromised lateral and longitudinal
filament contacts would, therefore, destabilize the
filament locally and promote severing.
The increase in the torsional flexibility of cofilinactin filaments (i.e. lower torsional constant, a,
Table 1) and larger intersubunit angular disorder
(hDxi, Table 1) is consistent with proposed cofilininduced disruption of longitudinal and lateral
interactions in actin. It is, therefore, likely that the
Cofilin Affects Actin Filament Torsional Dynamics
increased angular disorder (hDxi) induced by cofilin
binding causes local perturbations in filament
conformation and dynamics that promote severing.
Such a mechanism would account for efficient
filament severing at low cofilin binding densities
and cluster sizes.12
The observed effects of phalloidin provide further
insight into the mechanism of cofilin-induced local
and global changes in actin filament. Phalloidin
binding changes the conformation of subdomain 2
cooperatively,28 dampens the cofilin-dependent
torsional dynamics of actin filaments (Figure 7(b)),
has long-range effects stabilizing intermonomer
bonds, and cooperatively decreases the rate of
cofilin binding (Figure 7(c)), supporting our
hypothesis that the cofilin binding affinity is
dictated by the conformation of subdomain 2 as
well as actin filament torsional dynamics.12 Significant quenching of phosphorescence without binding-related changes in dynamics, as observed at
substoichiometric concentrations of phalloidin
(Figure 7(b)) further suggests that the mechanism
of long-range effects of cofilin on actin’s dynamics
involves destabilization of intermonomer bonds.
Comparison with other actin-binding proteins
The observed effect of cofilin on the microsecond
dynamics of actin represents another example of
cooperative changes in actin filaments induced by
interaction with regulatory proteins. Both gelsolin15
and myosin subfragment 116 cooperatively affect
the conformation and dynamics of actin filaments.
Although each of these proteins affects the environment of the actin C terminus, the changes in TPA
and dynamics are distinct (i.e. myosin increases but
cofilin and gelsolin lower the torsional rigidity),
indicating that the changes are specific to the
structure of the binding interfaces.
Materials and Methods
Proteins
Rabbit skeletal muscle actin was purified as
described.19,48 Recombinant human cofilin was expressed
and purified as described.12,49 All proteins were dialyzed
exhaustively against KMI6.6 buffer (50 mM KCl, 2 mM
MgCl2, 2 mM DTT, 0.2 mM ATP, 1 mM NaN3, 20 mM
imidazole (pH 6.6)) prior to use.
Labeling of actin with optical probes
Actin (48 mM) was polymerized with 50 mM KCl,
20 mM Tris (pH 7.5), and ErIA, freshly dissolved in
dimethylformamide, was added at a concentration of
480 mM. After 2 h incubation at 25 8C, the labeling reaction
was quenched with 5 mM DTT, actin filaments were
centrifuged for 1 h at 200,000g, pellets were suspended in
G buffer and clarified by centrifugation at 350,000g.
Samples were polymerized with 0.1 M KCl and centrifuged for 1 h at 200,000g. Pellets were suspended and
dialyzed against KMI6.6 buffer without magnesium.
997
Cofilin Affects Actin Filament Torsional Dynamics
Pyrene actin was prepared essentially as described.12 The
labeling efficiencies were R90%.
Phosphorescence
Phosphorescence measurements were made at 25 8C in
KMI6.6 buffer supplemented with an oxygen-scavenging
enzyme mixture (36 mg mlK1 catalase, 45 mg mlK1 glucose, 55 mg mlK1 glucose oxidase). Actin filaments and
cofilin-actin filaments were prepared by mixing preformed eryhrosine-labeled (ErIA) actin filaments with a
range of cofilin concentrations and equilibrated at room
temperature for at least 20 min. ErIA was excited at
540 nm with a vertically polarized 10 ns pulse from XeClpumped dye laser (Compex 120, Lambda Physics) using
5 mM coumarin 548 in ethanol, operating at a repetition
rate of 100 Hz. Phosphorescence emission was selected by
a colored glass cut-off 670 nm filter (Corion), detected by a
photomultiplier (R928, Hamamatsu), and digitized by a
transient digitizer (CompuScope 14100, GaGe) using time
resolution of 1 ms/channel, with an analog filter time
constant of 3 ms.
The time-resolved phosphorescence intensity I(t) and
anisotropy decays r(t) were calculated according to:
IðtÞ Z Ivv ðtÞ C 2GIvh ðtÞ
(1)
Ivv ðtÞKGIvh ðtÞ
Ivv ðtÞ C 2GIvh ðtÞ
(2)
and
rðtÞ Z
where Ivv(t) and Ivh(t) are vertically and horizontally
polarized components of the emission signal that were
detected at 908 with a single detector equipped with a
Polaroid sheet polarizer alternating between vertical and
horizontal orientations every 500 laser pulses. G is an
instrumentation correction factor, determined by
measuring the anisotropy of ErIA-labeled bovine serum
albumin in 98% glycerol and adjusting G to give a
residual anisotropy value of zero, the theoretical value
for an isotropically tumbling chromophore. The timedependent anisotropy decays were obtained by recording
60 cycles of 1000 pulses (500 in each orientation of the
polarizer) at a laser repetition rate of 100 Hz.
The phosphorescence intensity decays (I(t)) were
analyzed by fitting to a double-exponential:
Kt=t1
IðtÞ Z I1 exp
Kt=t2
C I2 exp
C I3
Ii ti
I1 t1 C I2 t2 C I3 t3
(4)
The lifetime of the rapidly decaying component (t3)
was assumed to be 1 ms, which is the upper limit, since it
is not observed within the 3 ms dead-time for data
acquisition (Figure 2). If the value of t3 is !1 ms,
population of X3 (Figure 3) would be lower.
The anisotropy decays (r(t)) were fitted to doubleexponentials (nZ2) plus a constant (rN):
rðtÞ Z r1 expKt=f1 C r2 expKt=f2 C rN
where, r0Z0.205 for ErIA immobilized in PMMA resin.19
The anisotropy decays (r(t)) were further analyzed in
terms of the theory of Schurr51,52 describing the rotational
diffusion of a flexible filament with mean local cylindrical
symmetry, and applied to TPA of ErIA-actin.19 According
to this model, the filament is regarded as a randomly
labeled array of cylindrical subunits. The anisotropy (r(t))
describes the mean-squared displacements of the subunit
elementary rods (and the rigidly bound probe) due to
combined intrafilament twisting and rigid-body motions
of the whole filament. If the filaments have a broad length
distribution, where each filament of length li is composed
of Ni elementary rods with height h equal to the height of
an individual actin subunit, the anisotropy r(t) reflects the
sum of contributions from the filaments within each
particular group:
rðtÞ Z k
(5)
where ri is the amplitude of the of the ith anisotropy
iZp
nZ2 X
X
A n Cni ðtÞ
(7)
nZ0 iZ1
where k is an “amplitude reduction factor” that accounts
for motions on the timescale more rapid than the time
resolution of detection,19 p is the total number of groups
in the length distribution histograms (Figure 6) and Ān
defines the amplitudes of the motions.
The torsional correlation function (Cni (t)) for filaments
in the length group li is defined as:19
h 2
i
n tkB T
exp K
ðNiC1Þg
Cni ðtÞ Z
Ni C 1
"
#
N
N
i C1
i C1
X
X
2
2 2
Kt=tsi
exp Kn
dsi Qmsi ð1Ke
Þ
!
mZ1
(3)
where Ii is the amplitude and ti is the triplet excited-state
lifetime of the ith relaxation, and t is time. Fits were
performed with unconstrained amplitude and lifetime
parameters, and then with the lifetimes constrained to the
values of F-actin.
Mole fractions (X) of the observed intensity decays
were calculated from:
Xi Z
decay, fi is the rotational correlation time of the ith
relaxation, and rN is the final anisotropy. The initial
anisotropy (r0) is defined as r(tZ0).
The cofilin-induced changes in the initial anisotropy
were analyzed in terms of the wobble-in-a-cone model.
The isotropic wobble of the observed transition dipole of
the probe in a cone was described by the cone half angle
qc:50
r0
cos qc ð1 C cos qc Þ 2
Z
(6)
0:205
2
d2si Z
kB Ttsi
g
sZ2
tsi Z
4a sin
2
g
ðsK1Þp
2ðNiC1Þ
and
sffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
mK 12 ðsK1Þp
2
cos
ð1Kdsi Þ
Qmsi Z
ðNi C 1Þ
ðNi C 1Þ
sffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
1
C dsi
ðNi C 1Þ
(8)
where kB is Boltzmann’s constant (1.381!10K23 Nm KK1),
T is the absolute temperature (298 K), Ni is the number of
subunits in filaments with length li, t is the relaxation
time, d2 is the mean-square amplitude of the sth normal
mode, dsi is the Kronecker delta function, g is the
frictional coefficient for rotation of an elementary rod of
height h about the filament long axis, defined by gZ
4pha2h, h is the solvent viscosity (1 cP), a is the filament
radius (4.5!10K9 m for pure actin and 6.7!10K9 m for
cofilin-decorated actin), a is the intrafilament torsional
998
Cofilin Affects Actin Filament Torsional Dynamics
constant and mZ1, 2,., N iC1. Note that kB T/g
represents the long-axis rotational diffusion coefficient
of an elementary rod, commonly referred to as Dk.
Fitting r(t) to equation (7) with the values of h, a, h, and
T constrained yields three parameters: qa and qe, the
angles between the absorption and emission dipoles of
the bound dye and the filament axis, respectively, and the
torsional constant a, which characterizes the elastic
properties of actin and reflects the torque force required
to twist a 1 m radius filament by 1 radian (57.38). A larger
torsional constant indicates a greater resistance to
twisting (i.e. more stiff) under applied external rotational
forces.
The torsional rigidity C is defined by the torsional
constant (a) and the long-axis height (h) of an elementary
rod (i.e. filament subunit) as:
C Z ah
(9)
The root-mean-square average fluctuation of the
torsion angle (hDxi in radians, 1 rad Z57.38) between
adjacent filament subunits was calculated from25:
rffiffiffiffiffiffiffiffiffi
kB T
(10)
hDxi Z
a
Equilibrium binding equations
The cofilin concentration-dependence of the initial (r0)
and final (rN) anisotropies were fitted to:
robs Z rCA KðrCA KrA Þð1KvÞn
(11)
where robs is the observed anisotropy (initial or final), rA is
the anisotropy (initial or final) of actin alone and rCA is
that of a cofilin-decorated actin filament, v is the cofilin
binding density (bound cofilin per actin), and n is the
stoichiometry (molar ratio) of cofilin that maximally
affects the robs of actin (i.e. number of actin subunits
affected by bound cofilin).
Pyrene fluorescence
Phalloidin inhibition of cofilin binding was assayed
from the time-courses of pyrene fluorescence quenching12,30 after mixing 11.8 mM cofilin with 0.85 mM pyreneactin filaments (final concentrations after mixing).
Measurements were made at 25.0(G0.1) 8C in KMI6.6
buffer with an Applied Photophysics SX.18MV-R
stopped-flow apparatus. A 400 nm colored glass emission
filter was used to monitor fluorescence (lexZ366 nm).
Time-courses of pyrene fluorescence changes were
fitted to single or double-exponentials. The phalloidinconcentration-dependence of the fast observed rate
constant (kobs) of fluorescence quenching was fitted to:
kobs Z ko C ðkN Kko Þ
0
B
!B
@
½Ph
½A
Kd
C ½A
r
ffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
ffi1
Cn K
½Ph
½A
2n
Kd
C ½A
Cn
2
K4 ½Ph
½A n C
C
A
(12)
where ko is the observed rate constant of cofilin binding to
actin filaments in the absence of phalloidin, kN fluorescence intensity is the observed rate constant of cofilin
binding to actin filaments in the presence of saturating
phalloidin, [Ph] and [A] are the total phalloidin and actin
concentrations respectively, Kd is the apparent dissociation equilibrium constant of phalloidin binding to
rabbit muscle actin filaments under our experimental
conditions (20 nM 53), and n is the stoichiometry (molar
ratio) of bound phalloidin that maximally inhibits cofilin
binding. The stoichiometry (n), initial (ko) and final (kN)
observed rate constants were allowed to float when
fitting.
Electron microscopy
Actin filaments and cofilactin filaments were prepared
by mixing preformed erythrosin-labeled actin filaments
with a range of cofilin concentrations, equilibrated at
room temperature for at least 20 min, adsorbed to glowdischarged carbon-coated copper grids, negatively
stained with 1% (w/v) uranyl acetate and visualized
with a JOEL 100 CX electron microscope at an accelerating
voltage of 80 kV.
Cosedimentation
Samples (200 ml) of ErIA-F-actin (1.8 mM) and cofilin
(0.1, 2 or 8 mM) were prepared with oxygen-removing
enzymes as for TPA experiments. Samples were equilibrated at 25 8C for 10 min then centrifuged for 30 min at
400,000g in a TLA-100 rotor, which is sufficient to pellet
even very short filaments (lavg w0.13 mm; w47 subunits).54
The fraction of ErIA-actin remaining in the supernatant
was determined by measuring absorbance of the probe at
538 nm of samples before centrifugation and supernatants.
Acknowledgements
We thank Dr Wenxiang Cao for engaging
discussions, Adrian O. Olivares and James P.
Robblee for assistance with some of the cofilin
preparations used in this study, and Brian Tucker
for assistance with the kinetics of cofilin binding to
pyrene actin filaments. This work was supported by
a Hellman Family Fellowship (to E.M.D.L.C.),
grants from the American Heart Association
(0235203N to E.M.D.L.C.), the National Science
Foundation (MCB-0216834 to E.M.D.L.C.), and the
NIH (AR32961 to D.D.T.).
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Edited by J. Karn
(Received 24 June 2005; received in revised form 6 September 2005; accepted 9 September 2005)
Available online 26 September 2005