THE JOURNAL OF BIOLOGICAL CHEMISTRY
© 2001 by The American Society for Biochemistry and Molecular Biology, Inc.
Vol. 276, No. 8, Issue of February 23, pp. 5952–5958, 2001
Printed in U.S.A.
The C-terminal Tail of UNC-60B (Actin Depolymerizing
Factor/Cofilin) Is Critical for Maintaining Its Stable Association
S
with F-actin and Is Implicated in the Second Actin-binding Site*□
Received for publication, August 18, 2000, and in revised form, October 16, 2000
Published, JBC Papers in Press, October 24, 2000, DOI 10.1074/jbc.M007563200
Shoichiro Ono‡§, Amy McGough¶, Brian J. Pope储, Vincent T. Tolbert**, Alice Bui‡, Jan Pohl‡‡,
Guy M. Benian‡, Kim M. Gernert§§, and Alan G. Weeds储
From the ‡Department of Pathology, Emory University, Atlanta, Georgia 30322, the ¶Department of Biological Sciences,
Purdue University, West Lafayette, Indiana 47907, 储Medical Research Council Laboratory of Molecular Biology,
Cambridge CB2 2QH, United Kingdom, **Frederick Douglas High School, Atlanta, Georgia 30030, the ‡‡Microchemical
Facility, Winship Cancer Institute, Emory University, Atlanta, Georgia 30322, and §§BIMCORE, Molecular Graphics,
Emory University, Atlanta, Georgia 30322
Actin depolymerizing factor (ADF)/cofilin changes the
twist of actin filaments by binding two longitudinally
associated actin subunits. In the absence of an atomic
model of the ADF/cofilin-F-actin complex, we have identified residues in ADF/cofilin that are essential for filament binding. Here, we have characterized the C-terminal tail of UNC-60B (a nematode ADF/cofilin isoform) as
a novel determinant for its association with F-actin. Removal of the C-terminal isoleucine (Ile152) by carboxypeptidase A or truncation by mutagenesis eliminated F-actin binding activity but strongly enhanced
actin depolymerizing activity. Replacement of Ile152 by
Ala had a similar but less marked effect; F-actin binding
was weakened and depolymerizing activity slightly enhanced. Truncation of both Arg151 and Ile152 or replacement of Arg151 with Ala also abolished F-actin binding
and enhanced depolymerizing activity. Loss of F-actin
binding in these mutants was accompanied by loss or
greatly decreased severing activity. All of the variants of
UNC-60B interacted with G-actin in an indistinguishable manner from wild type. Cryoelectron microscopy
showed that UNC-60B changed the twist of F-actin to a
similar extent to vertebrate ADF/cofilins. Helical reconstruction and structural modeling of UNC-60B-F-actin
complex reveal how the C terminus of UNC-60B might be
involved in one of the two actin-binding sites.
Actin depolymerizing factor (ADF)1/cofilins are a family of
actin-regulatory proteins ubiquitous among eukaryotes that
are essential for rapid turnover of the actin cytoskeleton in vivo
(1–3). ADF/cofilin enhances the turnover of actin filaments by
* This work was supported by the American Heart Association Southeast Affiliate Grant 9960146V (to S. O.), the National Science Foundation Grant MCB-9728762 (to G. M. B.), and National Institutes of
Health Grant GM59677 (to A. M.). The costs of publication of this
article were defrayed in part by the payment of page charges. This
article must therefore be hereby marked “advertisement” in accordance
with 18 U.S.C. Section 1734 solely to indicate this fact.
□
S The on-line version of this article (available at http://www.jbc.org)
contains coordinates and structures. To view these PDB files, you may
use Rasmol software (http://www.rasmol.org).
§ To whom correspondence should be addressed: Dept. of Pathology,
Emory University, 1639 Pierce Dr., Woodruff Memorial Bldg., Rm.
7109C, Atlanta, GA 30322. Tel.: 404-727-3916; Fax: 404-727-8540; Email: [email protected].
1
The abbreviations used are: ADF, actin depolymerizing factor; CPA,
carboxypeptidase A; Bicine, N,N-bis(2-hydroxyethyl)glycine; MOPS,
4-morpholinepropanesulfonic acid.
both increasing the rate of depolymerization from their pointed
ends (4, 5) and by severing them thereby increasing the number of ends (5–14). These two activities can be uncoupled by
point mutations (15, 16). Binding of ADF/cofilin to F-actin
changes the twist of the filaments (17) and leads to destabilization of the lateral contacts of actin monomers within the
filaments (18). This ADF/cofilin-mediated structural change in
F-actin is important for cooperative binding of ADF/cofilin to
F-actin (17, 19), resulting in the coexistence of bare and ADF/
cofilin-decorated actin filaments, as observed in the lamellipodia of cultured cells (20).
The structural fold of the ADF/cofilin family proteins (21–23)
is similar to that of the individual segments of the gelsolin
family (24). Although the structure of gelsolin segment-1-actin
complex has been solved (25), predicted models of ADF/cofilinactin complex that were based on the topology of this complex
(21, 26) differed significantly and can not be used to predict
F-actin binding. When one of these predicted structures was
extended to develop models of ADF/cofilin-F-actin complex (17),
the C terminus of ADF/cofilin was placed away from the actin
surface. However, alanine-scanning mutagenesis of yeast cofilin (27) revealed that helix ␣4 near the C terminus (Fig. 1b) is
a part of the actin-binding surface that confers filament
binding.
Mutagenesis of yeast cofilin (27) revealed two actin-binding
surfaces as follows: one that is essential for G-actin binding and
a second that is required for F-actin. The essential G-actinbinding surface includes the N terminus, a portion of helix ␣3,
and the turn connecting strand ␤6 and helix ␣4 (27) (Fig. 1b).
Helix ␣3 includes two highly conserved basic residues that
have been implicated in monomer binding in several ADF/
cofilin proteins by chemical cross-linking (28), mutagenesis (16,
29, 30), and peptide competition (28, 31). The N terminus is
also involved in actin binding (32), and this region includes the
phosphorylation site Ser-3 (Ser-6 in plant ADF) that is responsible for phosphorylation-dependent inactivation of actin binding by ADF/cofilin (33–36). These regions of the molecule are
close together in the three-dimensional structures and probably form a binding surface that interacts with subdomains 1
and 3 of G-actin (21–23).
There is little detailed information regarding the site on
ADF/cofilin that is required for F-actin binding. Two regions
have been implicated in filament binding as follows: (i) a cluster of charged residues (15, 27) in a loop connecting ␤2 and ␤3
in destrin and at a similar position at the beginning of ␤5 in
yeast cofilin, and (ii) a group of charged residues in helix ␣4 in
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This paper is available on line at http://www.jbc.org
Role of the C Terminus of ADF/Cofilin
yeast cofilin (27). In addition, mutations of conserved tyrosine
residues (Tyr67 and Tyr70 in maize ADF3) in the apolar core of
the molecule results in uncoupling of monomer and filament
binding activities (37).
We previously identified a novel function for the C-terminal
tail of ADF/cofilin (30). The tail is predicted to lie outside helix
␣4 (Fig. 1c) and is not included in the crystal structures of
either yeast cofilin (23) or Acanthamoeba actophorin (38), suggesting structural flexibility in this region. A mutant form of
UNC-60B (a Caenorhabditis elegans ADF/cofilin isoform) that
lacks three C-terminal amino acids (Fig. 1c) shows loss of both
F-actin binding and severing activity but still depolymerizes
F-actin, possibly through its monomer sequestering activity
(30). Importantly, a C. elegans strain that expresses this mutant is defective in proper assembly of actin into myofibrils (30),
indicating that the C-terminal tail of UNC-60B is functionally
significant in vivo. Therefore, in this study, we characterized
the role of the C-terminal tail of UNC-60B in its actin regulating activity.
EXPERIMENTAL PROCEDURES
Homology Modeling of the Structure of UNC-60B—Homology modeling of UNC-60B was based on the known structures of Saccharomyces
cerevisiae cofilin (Protein Data Bank code 1cfy) (23), Acanthamoeba
castellanii actophorin (Protein Data Bank code 1ahq) (22), and Sus
scrofa destrin (Protein Data Bank code 1ak7) (21). Coordinate files were
obtained from Protein Data Bank. The initial alignment, generated by
pattern-induced (local) multiple alignment, showed 10% identical and
57% conserved residues between UNC-60B and cofilin, actophorin and
destrin (Fig. 1a). A model was built based on this initial alignment
using the program Modeller by Sali and Blundel (39).
Evaluation of the initial model, superimposed with the known structures, suggested changes in the alignment in the region of the insertion
at UNC-60B Lys58–Lys65. Multiple cycles of realignment and rebuilding
of the models were completed. Side chain conformations, along the
length of the UNC-60B sequence, were optimized manually to satisfy
individual residue structural functions (i.e. van der Waals packing,
ionic interactions, H bonds between side chain and main chain, etc.) as
seen in the known structures. Additional refinement and minimization
of the model are underway.
Proteins—Recombinant wild-type UNC-60B was purified as described (40). Rabbit muscle actin was purified as described (41), and
G-actin was further purified by gel filtration with Sephacryl S-300 or
purchased from Cytoskeleton Inc. (Denver, CO). We obtained indistinguishable results using either actin preparation. C. elegans actin was
purified from wild-type N2 strain (obtained from the Caenorhabditis
Genetics Center, St Paul, MN) as described (42). Bovine pancreas
carboxypeptidase A (CPA) (EC 3.4.17.1) was purchased from Roche
Molecular Biochemicals.
CPA Treatment of UNC-60B—UNC-60B (1 mg/ml) was incubated
with CPA (20 ␮g/ml) in a buffer containing 0.1 M KCl, 20 mM HEPESNaOH, 1 mM dithiothreitol, pH 7.5, at 25 °C for 3 h. The digestion was
stopped by adding 1,10-phenanthroline (Fisher) at a final concentration
of 0.5 mM.
Electrospray Ionization Mass Spectrometry—The protein solution
was acidified to pH ⬃2 with 10% trifluoroacetic acid, and the protein
was chromatographed on a Jupiter-C4 column (1 ⫻ 150 mm) equilibrated in 0.1% aqueous trifluoroacetic acid and eluted using a linear
gradient of acetonitrile in 0.08% aqueous trifluoroacetic acid. The column eluate was monitored at 210 nm, and the fractions containing the
protein (eluting as a major peak at ⬃30% acetonitrile) were collected for
further analysis. The fractions were mixed 1:1 with isopropyl alcohol/
water/acetic acid (50:50:1) and directly infused at a flow rate of 15
␮l/min into a MicroIon source of model API3000 triple quadrupole mass
spectrometer. Following spectra deconvolution (using the BioSpec
BioReconstruct routine), the mass of the protein was determined.
N-terminal Sequence Analysis—The proteins were subjected to automated Edman degradation in a Procise-cLC sequencer (PE Biosystems)
using a standard manufacturer’s protocol.
Mutant UNC-60B Proteins—Mutations were introduced in synthetic
oligonucleotides that complement the 3⬘-end of the coding sequence of
the cDNA for UNC-60B. The cDNA fragments that carry mutations
were amplified by polymerase chain reaction using these mutant primers and a common forward primer (5⬘-GATCCCATGGCTTCCGGAGTCAAAGTTG). The amplified DNA fragments were digested by NcoI and
5953
BamHI at the sites introduced by the primers and cloned into pET-3d
(Novagen, Madison, WI). The sequences of the inserts were verified by
DNA sequencing not to contain any polymerase chain reaction-induced
errors. The mutant primers used are 5⬘-CTAGGGATCCTTATCTTTGGTTGGACATCAGGTC for ⌬I, 5⬘-CTAGGGATCCTTATTGGTTGGACATCAGGTCGC for ⌬RI, 5⬘-CTAGGGATCCTTAGATTCTTTAGTTGGACATC for ⌬QRI, 5⬘-CTAGGGATCCTTAGGCTCTTTGGTTGGACATC
for I152A, and 5⬘-CTAGGGATCCTTAGATTGCTTGGTTGGACATC for
R151A. The mutant UNC-60B proteins were expressed and purified as
described previously for wild-type UNC-60B (40).
Actin-binding Assays—Copelleting assays of UNC-60B with rabbit
muscle F-actin were performed as described previously (40) in a buffer
containing 0.1 M KCl, 2 mM MgCl2, 20 mM HEPES-NaOH, 1 mM dithiothreitol, pH 7.5. All protein solutions except F-actin were clarified by
ultracentrifugation prior to the pelleting assays. Ultracentrifugation
was performed with a Beckmen Airfuge at 28 pounds/square inch for 20
min. Assay for nucleation of actin polymerization was performed as
described previously (30) except that rabbit muscle actin was used as
seeds in this study.
Nondenaturing Polyacrylamide Gel Electrophoresis—Nondenaturing
polyacrylamide gel electrophoresis was performed as described by Safer
(43). CaATP-G-actin and UNC-60B were incubated in G-buffer (2 mM
Tris-HCl, 0.2 mM CaCl2, 0.2 mM dithiothreitol, 0.2 mM ATP, pH 7.5) for
30 min at room temperature. The samples were then supplemented
with 0.25 volume of a loading buffer (50% glycerol, 0.05% bromphenol
blue) and electrophoresed using a Bicine/triethanolamine buffer system. The proteins were visualized by staining with Coomassie Brilliant
Blue R-250 (Sigma).
Cryoelectron Microscopy and Modeling of the Structure of UNC-60BF-actin Complex—Decorated filaments were prepared in one of two
ways. In most cases, pre-formed actin filaments (3 ␮M) were incubated
at room temperature for ⬃2 h with 10 ␮M UNC-60B in 20 mM MOPS,
pH 7, 0.1 M KCl, 2 mM MgCl2, 1 mM dithiothreitol, 0.5 mM EGTA.
Although ATP was present in the polymerized F-actin solution, no
additional ATP was added upon dilution into the UNC-60B-containing
buffer, resulting in a final concentration of ⬍0.04 mM ATP. For partial
decoration of filaments, 10 ␮M F-actin was decorated on ice overnight in
the presence of 10 ␮M UNC-60B using the same buffer conditions and
diluted to a final actin concentration of 2.5 ␮M just prior to plunging.
Filament decoration was confirmed by pelleting assays in a Beckman
Airfuge at 20 pounds/square inch for 15 min followed by SDS-polyacrylamide gel electrophoresis. Decorated filaments were rapidly frozen on
holey carbon films in ethane slush cooled with liquid nitrogen and then
imaged using a JEOL 1200EX transmission electron cryomicroscope at
the National Center for Macromolecular Imaging at Baylor College of
Medicine. Micrographs were recorded at an accelerating voltage of 100
kV and a nominal magnification of ⫻ 30,000.
Suitable micrographs were digitized using a Zeiss SCAI film scanner
at a final resolution of 4.7 Å per pixel. Tobacco mosaic virus particles
(0.05 mg/ml; kindly provided by Dr. Ruben Diaz, Florida State University) were included in the samples to provide an internal magnification
standard. The defocus of each micrograph was determined by incoherent averaging of regions of protein embedded in ice (44). The micrographs used in this study were recorded at ⫺2.1- and ⫺2.6-␮m defocus.
Filament images were analyzed using the helical image processing
package PHOELIX (45, 46). Mean crossover lengths were measured
from positions of the first layer line in computed diffraction patterns of
computationally straightened filaments.
For image reconstructions, layer line data were collected to a radial
resolution of 0.032 Å⫺1 (within the first node of the contrast transfer
function) and separated into near and far side data sets. Filaments that
best conformed to the 20:9 selection rule and that showed good symmetry between the near and far sides of the Fourier transform were
selected for further processing. A total of 22 data sets were averaged,
and a three-dimensional structure was calculated by Fourier-Bessel
inversion (47) to a resolution limit of 0.032 Å⫺1 radially using the
following layer lines (l, n): 0, 0; 2, 2; 4, 4; 5, ⫺5; 7, ⫺3; 9, -1; 11, 1; 13,
3; 15, 5. The mean phase residuals during the last round of alignment
of the 22 data sets were 31.8° and 47.6° for the correct and incorrect
polarity, respectively. The resulting reconstruction was aligned to a
previously calculated F-actin reconstruction (17) in Fourier space to aid
in model building.
Pseudo-atomic models of UNC-60B-F-actin were derived using the
actin subunits from the Lorenz filament model (twisted to match the
UNC-60B-induced actin structure) and the homology-based model of
UNC-60B. The UNC-60B model structure was positioned interactively
using the programs O (48) and Iris Explorer (Numerical Algorithms
Group, Downers Grove, IL) running on a Silicon Graphics Indigo R4000
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Role of the C Terminus of ADF/Cofilin
FIG. 1. Homology modeling of the structure of UNC-60B. a,
sequence alignment of UNC-60B, yeast cofilin (Y-cof), actophorin (Actoph), and porcine destrin. Locations of helices ␣1– 4 are indicated by
bold and underlined letters. Colored residues correspond to the ones in
b and c and are explained below. b, crystal structure of yeast cofilin
(Protein Data Bank code 1cfy). Essential residues for actin-binding are
shown in red, and residues that confer F-actin binding are shown in
yellow (27). Note that residues 1–5 and 140 –143 are not visible in the
crystal structure. Therefore, Val6 is colored red instead of the essential
residues 1–5. c, a predicted structure of UNC-60B. Missense mutations
that have been reported by Ono et al. (30) are shown in blue. Residues
in the C-terminal tail (C-tail) that are truncated in the unc-60(r398)
mutation are shown in green. The coordinates of this model
(unc60b_model) are available as Supplemental Material.
work station (Mountain View, CA). The goal was to produce a model
that fit inside the envelope of the electron microscopy reconstruction
well and also taking into account available genetic and biochemical
data concerning the cofilin residues involved in actin contacts (27, 28,
30, 31). Ribbon diagrams were generated using Ribbons 2.65 (49), saved
as Inventor format files, and displayed in IRIS Explorer.
RESULTS
Homology Modeling of the Structure of UNC-60B—The structure of UNC-60B was modeled based on the known structures
of three ADF/cofilin proteins. The model presented in Fig. 1c
incorporated the tertiary structures of the known ADF/cofilin
proteins, a three-layer ␣-␤-␣ sandwich, with an internal fivestranded ␤-sheet. Significant differences in the model resulted
from insertions in the UNC-60B sequence between helix ␣2 and
strand ␤3 (UNC-60B Lys58–Lys65 inserted between Pro58 and
Asn60 of cofilin) and between strands ␤3 and ␤4 (UNC-60B
Arg80–Thr86 inserted between Gly75 and Gly78 of cofilin). There
is also an addition of 4 residues at the C terminus (UNC-60B
Asn149–Ile152), the structure of which is modeled after the
destrin C-terminal structure (residues Leu156–Ala159).
Ile152 of UNC-60B Is Required for Maintaining Stable Association with F-actin—The C-terminal isoleucine at position 152
of UNC-60B was removed by carboxypeptidase, and its effect
on the activity of UNC-60B was examined. We found that CPA
removed only Ile152 because Arg151 is not a substrate of CPA.
Removal of Ile152 was confirmed by electrospray mass spectrometry. Molecular mass of control UNC-60B was determined
as 16,919 atomic mass units that corresponded to residues
2–152 of UNC-60B (the predicted mass, 16,915 atomic mass
units). However, the spectrum of CPA-treated UNC-60B
yielded a major peak of 16,806 atomic mass units and a minor
one of 16,921 atomic mass units. These correspond to residues
2–151 (the predicted mass, 16,802 atomic mass units) and
2–152, respectively. The mass of the major peak was consistent
with that of UNC-60B-⌬I (the experimentally determined
mass, 16,803 atomic mass units), a mutant lacking Ile152. Area
calculations of the peaks from the mass spectra suggested that
⬃17% of the total UNC-60B was not processed with CPA assuming that both forms were ionized to the same extent. Nterminal sequencing of CPA-treated UNC-60B yielded the
identical sequence to control UNC-60B starting from Ala2, indicating that there was no detectable amino- or endopeptidase
activity in the CPA preparation. In addition, there was no
evidence of major alteration in secondary structure as examined by circular dichroism (plots not shown), providing evidence that removal of Ile152 does not significantly disrupt the
structure of the C-terminal helix ␣4.
Interaction of CPA-treated UNC-60B with F-actin was surprisingly different from that of control UNC-60B. Whereas
control UNC-60B cosedimented with rabbit muscle F-actin
without significantly depolymerizing it (Fig. 2a, lane 2, compare with lane 1) (40, 42), CPA-treated UNC-60B increased the
amount of unpolymerized actin in the supernatant and did not
cosediment with F-actin (Fig. 2a, lane 3). Thus, CPA-treated
UNC-60B depolymerizes F-actin and prevents re-polymerization. CPA itself did not cause actin depolymerization (Fig. 2a,
lane 4). Furthermore, preincubation of CPA with its inhibitor,
1,10-phenanthroline, abolished the ability of CPA to change the
activity of UNC-60B (Fig. 2a, lane 5). These results indicate
that removal of the C-terminal Ile from UNC-60B converts it
from a filament-binding protein to a depolymerizing factor.
Nearly complete depolymerization of F-actin was achieved at
molar ratios at greater than 1:1 of CPA-treated UNC-60B to
actin monomer (Fig. 2b). CPA-treated UNC-60B also showed
enhanced actin depolymerizing activity using C. elegans actin
as compared with control UNC-60B (data not shown). Since
rabbit muscle actin consists of a single isoform that has been
very well characterized, we used rabbit muscle actin in the
biochemical experiments to correlate better the biochemical
data with the structural interpretation as described below.
In contrast to the effects on F-actin, CPA treatment of UNC60B did not affect its interaction with G-actin. Both control
UNC-60B and CPA-treated UNC-60B were resolved as a single
band on nondenaturing polyacrylamide gels (Fig. 3, lanes 2–5),
whereas actin gave a doublet and smeared bands (Fig. 3, lane
1). When actin and either control or CPA-treated UNC-60B
were mixed, a third band appeared (Fig. 3, lanes 6 –9, arrows).
The intensity of this band increased as the concentration of
UNC-60B was increased (Fig. 3, lanes 6 –9). This increase was
accompanied by a decrease in the intensities of both actin and
UNC-60B bands. These results indicate that the third band
Role of the C Terminus of ADF/Cofilin
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FIG. 3. G-actin binding of control and CPA-treated UNC-60B.
Interactions of G-actin with control UNC-60B or CPA-treated UNC-60B
at indicated concentrations were analyzed by nondenaturing polyacrylamide gel electrophoresis. In the presence of both proteins, the complex
of the two proteins emerged as the third bands (arrows). The positions
of actin and UNC-60B are indicated by A and U, respectively.
FIG. 2. Actin-regulating activity of CPA-treated UNC-60B. a,
copelleting assay of F-actin with UNC-60B. F-actin suspension (10 ␮M)
was incubated with buffer (lane 1), control UNC-60B (10 ␮M) (lane 2),
CPA-treated UNC-60B (10 ␮M) (lane 3), CPA (lane 4), or phenanthroline/CPA-treated UNC-60B (10 ␮M) (lane 5). The mixtures were ultracentrifuged, and the supernatants (s) and pellets (p) were analyzed by
SDS-polyacrylamide gel electrophoresis. The positions of actin and
UNC-60B are indicated by A and U, respectively. Molecular mass
markers in kDa are indicated on the left. b, quantitative analysis of the
pelleting assay of F-actin (10 ␮M) with varied concentrations of control
(triangles) or CPA-treated UNC-60B (circles). On the vertical axis are
plotted the sedimented portions (expressed as concentrations assuming
that the pellets were reconstituted in the original volumes) of actin
(open symbols) and control or CPA-treated UNC-60B (filled symbols).
represents the actin-UNC-60B complex. Results were similar
with control or CPA-treated UNC-60B (compare Fig. 3, upper
and lower panels), suggesting that CPA treatment of UNC-60B
did not affect its G-actin binding activity.
The role of Ile152 of UNC-60B was further investigated by
using recombinant mutant UNC-60B proteins. Mutant UNC60B-⌬I, which lacks Ile152, showed substantial actin-depolymerizing activity and cosedimented poorly with F-actin (Fig.
4b). This is in marked contrast to wild-type which binds F-actin
but does not significantly depolymerize it (Fig. 4a). These results are consistent with those using CPA treatment, except
that CPA-treated UNC-60B exhibited stronger actin-depolymerizing activity than UNC-60B-⌬I. UNC-60B-⌬I was indistinguishable from CPA-treated UNC-60B in its interaction
with G-actin, molecular mass, and N-terminal sequence (data
not shown). Treatment of UNC-60B-⌬I with CPA did not
change its behavior (data not shown), suggesting that the
quantitative difference may reflect slightly different folding of
the bacterially expressed ⌬I mutant compared with CPAtreated UNC-60B. Incomplete processing of UNC-60B with
CPA (17% of UNC-60B remained intact) is unlikely to cause
this difference because a mixture of control UNC-60B and
UNC-60B-⌬I at the equivalent ratio showed no difference in
actin depolymerizing activity compared with UNC-60B-⌬I
alone (data not shown).
To investigate further the significance of the isoleucine residue at position 152, we produced the mutant UNC-60B-I152A
in which Ile152 was replaced with Ala. UNC-60B-I152A showed
weaker binding to F-actin than wild-type UNC-60B but had
increased depolymerizing activity compared with the control
(compare Fig. 4, a and c). In the presence of 30 ␮M UNC-60BI152A, 6.6 ⫾ 0.24 ␮M actin was precipitated, and with 30 ␮M
wild-type, 8.0 ⫾ 0.07 ␮M actin was precipitated (Fig. 4, c and a,
respectively). UNC-60B-I152A was indistinguishable from wild
type in its interactions with G-actin (data not shown). These
results suggest that Ala can partially substitute for Ile at
position 152 but is not as efficient as Ile in maintaining stable
association with F-actin. Taken together, these data demonstrate the importance of Ile152 for UNC-60B to make a stable
complex with F-actin and inhibit filament disassembly.
Arg151 of UNC-60B Is Required for F-actin Binding and
Efficient Actin Depolymerization—We investigated the roles of
adjacent C-terminal amino acids of UNC-60B using bacterially
expressed mutant UNC-60B proteins with additional truncations at the C terminus. Mutant UNC-60B-⌬RI cosedimented
poorly with F-actin but showed considerable depolymerizing
activity. This mutant was not as effective as UNC-60B-⌬I (compare Fig. 4, b and d), suggesting that Arg151 is needed for
efficient depolymerizing activity. Additional truncation of Gln150 (UNC-60B-⌬QRI, formerly designated as the r398 mutant
(30)) did not cause any detectable difference in the depolymer-
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Role of the C Terminus of ADF/Cofilin
FIG. 4. Interactions of mutant UNC60B with F-actin. F-actin (10 ␮M) was
incubated with various concentrations of
wild-type (a), ⌬I (b), I152A (c), ⌬RI (d),
⌬QRI (e), or R151A (f) UNC-60B variants.
The mixtures then were examined by
copelleting assays. On the vertical axes
are plotted the sedimented portions (expressed as concentrations assuming that
the pellets were reconstituted in the original volumes) of actin (closed circles) and
wild-type or mutant UNC-60B (open circles). Alterations of UNC-60B at the C
terminus (the C-terminal sequences are
shown at the top of each panel) cause variable F-actin binding and depolymerizing
activities, although all the variants similarly interact with G-actin in nondenaturing polyacrylamide gel electrophoresis
(data not shown). Data shown are
means ⫾ S.D. of three experiments.
izing activity compared with UNC-60B-⌬RI (Fig. 4e). UNC60B-⌬RI and UNC-60B-⌬QRI were indistinguishable from wild
type in their interaction with G-actin (data not shown).
We further characterized the role of Arg151 using UNC-60BR151A in which Arg151 was replaced with Ala. UNC-60BR151A cosedimented poorly with F-actin and showed significant actin-depolymerizing activity despite the presence of Ile152
(Fig. 4f). This suggests that Arg151 is required for F-actin
binding by UNC-60B, whereas Ile152 has an accessory function.
Interestingly, there were quantitative differences in the depolymerizing activities of UNC-60B-R151A and the truncated
mutants. At molar ratios ⬍1:1, UNC-60B-R151A depolymerized actin to lesser extents than UNC-60B-⌬I, -⌬RI, or -⌬QRI.
However, at higher molar ratios (⬎2:1), the extent of depolymerization by UNC-60B-R151A was greater than that of
UNC-60B-⌬RI, or -⌬QRI, but slightly weaker than UNC-60B⌬I. Thus, it is possible that these mutants affect actin depolymerization in different ways.
Both Arg151 and Ile152 of UNC-60B Are Required for F-actin
Severing—We explored the role of the C-terminal tail of UNC60B in severing actin filaments, and we found that both Arg151
and Ile152 are required for this activity. Wild-type UNC-60B
increased the ability of F-actin seeds to nucleate actin polymerization by 2.5-fold (Fig. 5), which is strong evidence that
UNC-60B increased the number of ends by severing filaments.
However, this activity was greatly reduced using UNC-60B-⌬I
or UNC-60B-I152A (Fig. 5). The severing activity of these mutants correlates with their weaker F-actin binding activity (Fig.
4c). CPA-treated UNC-60B has no effect on the nucleation rate
(Fig. 5), which again suggests quantitative differences between
CPA-treated UNC-60B and recombinant UNC-60B-⌬I. Thus,
Ile152 of UNC-60B is required for both filament binding and
efficient severing.
Arg151 is also essential for the severing activity of UNC-60B.
Either truncation (UNC-60B-⌬RI and -⌬QRI) or replacement
with Ala (UNC-60B-R151A) abolished this activity (Fig. 5).
These results suggest that both Arg151 and Ile152 are essential
for actin severing activity by UNC-60B.
Structure of UNC-60B-F-actin Complex—We first confirmed
that UNC-60B altered the F-actin twist by computing the Fourier transforms of decorated filaments and measuring the
height of the first layer line that arises from the twist of the
two-start actin helix. UNC-60B produced a mean filament
twist (expressed as rotation per subunit) of 161.6° for C. elegans
Role of the C Terminus of ADF/Cofilin
FIG. 5. Effects of mutant UNC-60B on the nucleating activity of
F-actin seeds. F-actin (10 ␮M) was mixed with the UNC-60B variants
at various molar ratios and used as nuclei at 1.25 ␮M to induce polymerization of pyrene-labeled G-actin. Nucleation rates were determined
as the initial rates of the increase in the pyrene fluorescence, which is
proportional to the number of free barbed ends. The data are expressed
as relative nucleation rates to that of F-actin alone and are means of
three experiments. Error bars are not included in the figure because
they make the graph difficult to read. Standard deviations of these data
are less than 0.27.
F-actin. Filaments prepared using rabbit muscle actin were
twisted to a mean twist of 161.7° per subunit. Thus, as was
found with human cofilin and human ADF (17), UNC-60B
reduces the filament crossover length to about 270 Å irrespective of the source of the actin. Electron cryomicroscopy of filaments decorated with sub-saturating amounts of UNC-60B
confirmed that the C. elegans protein shares the ability of the
human protein to bind filaments cooperatively.
A three-dimensional reconstruction was calculated from electron micrographs of frozen-hydrated UNC-60B-F-actin filaments essentially as described previously (17) (Fig. 6a). This
reconstruction was used to provide a molecular envelope into
which the UNC-60B and actin subunit could be positioned (Fig.
6b). In addition to using the reconstruction as a constraint,
available biochemical and genetic data implicating the long
helix ␣3, the N terminus, and the C-terminal residues (27, 28,
30, 31) were used to help orient the UNC-60B molecule on the
filament.
The resulting orientation is similar to an earlier model of
human ADF in terms of the long helix involvement and its
“polarity” relative to the filament. Residues Val104–Ser113 of
the long helix are still in proximity to actin, but the remaining
7 residues (Val114–Ser120) are rotated away from the filament.
Interestingly, the portion of helix ␣3 that falls near actin includes Ser112 and Ser113; replacement of either of these residues by apolar groups alters F-actin binding (30). Significantly,
both the N and C termini are positioned near the actin interface in the new orientation, consistent with a number of biochemical and genetic studies.
DISCUSSION
In this study, we show that alterations of UNC-60B at the C
terminus inhibit F-actin binding and severing without changing G-actin binding. We found that F-actin binding of UNC-60B
5957
FIG. 6. Structural model for UNC-60B binding to F-actin. a,
reconstruction of UNC-60B-F-actin filament showing four filament
crossovers (40 subunits). b, model of UNC-60B binding to F-actin. Two
actin subunits (red and white ribbons) from the filament are shown. The
asterisks mark the positions of Ser112 (red), Ser113 (blue), and Arg151
(black). The coordinates of this model are available as Supplemental
Material (actin_up, the upper actin; actin_low, the lower actin;
unc60b_comp, UNC-60B.
(Fig. 4) was correlated with the extent of filament severing
(Fig. 5). This suggests that severing occurs as a consequence of
physical distortions in the helix that arise as a result of the
change in twist induced when these proteins bind to the side of
filaments (Fig. 6). This is consistent with previously suggested
models (17, 18).
Cryoelectron microscopy and structural modeling of UNC60B-F-actin complex suggest that the C terminus is involved in
the second actin-binding site. The alterations at the C terminus
are not likely to cause major disruption of the structural fold,
because we were unable to detect any changes in far UV circular dichroism or in their interactions with G-actin. However,
we cannot exclude the possibility of minor conformational
changes caused by these alterations. The structures of the
C-terminal regions of yeast cofilin (23) and actophorin (22) are
disordered in the crystals, which suggests that the C terminus
is not well ordered although it may be stabilized upon binding
to F-actin. The fact that Ile152 is a substrate of CPA indicates
that this residue is exposed on the surface of the molecule and
readily accessible to its active site.
Our reconstructions and structural modeling suggest that
both the N and C termini are positioned near the actin interface, but the predicted interactions they make with the filament are very different. The N terminus is positioned near the
subdomain 1 of the upper actin subunit, whereas the C terminus falls near subdomain 2 of the lower actin. Since cofilin and
profilin compete for binding to monomeric actin (8, 50), and
profilin binds on the “bottom” of actin between subdomains 1
and 3, this implies that the N terminus, but not the C terminus,
is involved in G-actin interactions. The C terminus, on the
other hand, primarily influences F-actin interactions. Mutational analysis of human cofilin suggested that F-actin binding
occurs initially through this site with the lower actin, followed
by interaction with the upper actin subunit via the G-actin site
(15). Interestingly, subdomain 2 is generally accepted as a
5958
Role of the C Terminus of ADF/Cofilin
molecular “sensor” for the nucleotide state (51, 52). The finding
that F-actin binding by actophorin increases the rate of phosphate release (12) raises the possibility that the C terminus is
directly involved in the nucleotide sensitivity of cofilin for actin.
It will be interesting to find out whether the C terminus is
involved in the activation of phosphate release. If the C-terminal tail of UNC-60B is flexible, preliminary molecular graphic
simulations suggested that it could extend into the ATP-binding pocket of actin and interact with the nucleotide-binding
site.
By using these mutants, we have also shown enhanced depolymerizing activity. The mechanism by which this occurs is
uncertain, but our preliminary experiments using gelsolincapped filaments show that these mutants activate subunit
dissociation from the pointed ends of filaments. The extent of
depolymerization differed between different mutants even in
the absence of any severing. This supports recent reports that
depolymerization and severing activities of ADF/cofilin can be
uncoupled (15, 16). Detailed kinetic analysis of depolymerization by our mutant versions of UNC-60B should provide further
insight into this mechanism.
Our data suggest that the C terminus of ADF/cofilin is the
structural determinant of the quantitative differences in Factin binding and actin-depolymerizing activities that have
been observed for different ADF/cofilin family members (5, 7,
19, 40, 53). The C-terminal region is highly divergent in both
sequence and length (54). For example, human ADF is a much
more potent depolymerizing agent than human cofilin (55), and
the largest region of sequence divergence is located in the
C-terminal 20% of the protein. Moreover, two C. elegans isoforms, UNC-60A and UNC-60B, show far greater sequence
divergence and functional difference than the two human isoforms, with the biggest differences in sequence seen near the C
terminus (40, 56). It will be interesting to examine the structural basis of these differences and the extent to which various
C-terminal sequences are involved in functional diversity.
Acknowledgments—We thank Sharon Langley for technical assistance in DNA sequencing. Instrumentation Grants RR12878 and
RR13948 were from the National Center for Research Resources of the
National Institutes of Health to Emory Microchemical Facility. The
electron microscopy facilities at Baylor College of Medicine are supported by a grant (to Wah Chiu) from the National Center for Research
Resources of the National Institutes of Health.
REFERENCES
1. Bamburg, J. R., McGough, A., and Ono, S. (1999) Trends Cell Biol. 9, 364 –370
2. Bamburg, J. R. (1999) Annu. Rev. Cell Dev. Biol. 15, 185–230
3. Carlier, M. F., Ressad, F., and Pantaloni, D. (1999) J. Biol. Chem. 274,
33827–33830
4. Carlier, M. F., Laurent, V., Santolini, J., Melki, R., Didry, D., Xia, G. X., Hong,
Y., Chua, N. H., and Pantaloni, D. (1997) J. Cell Biol. 136, 1307–1322
5. Maciver, S. K., Pope, B. J., Whytock, S., and Weeds, A. G. (1998) Eur. J. Biochem. 256, 388 –397
6. Mabuchi, I. (1983) J. Cell Biol. 97, 1612–1621
7. Nishida, E., Maekawa, S., and Sakai, H. (1984) Biochemistry 23, 5307–5313
8. Maciver, S. K., Zot, H. G., and Pollard, T. D. (1991) J. Cell Biol. 115, 1611–1620
9. Hawkins, M., Pope, B., Maciver, S. K., and Weeds, A. G. (1993) Biochemistry
32, 9985–9993
10. Hayden, S. M., Miller, P. S., Brauweiler, A., and Bamburg, J. R. (1993)
Biochemistry 32, 9994 –10004
11. Du, J., and Frieden, C. (1998) Biochemistry 37, 13276 –13284
12. Blanchoin, L., and Pollard, T. D. (1999) J. Biol. Chem. 274, 15538 –15546
13. Ressad, F., Didry, D., Egile, C., Pantaloni, D., and Carlier, M. F. (1999) J. Biol.
Chem. 274, 20970 –20976
14. Ichetovkin, I., Han, J., Pang, K. M., Knecht, D. A., and Condeelis, J. S. (2000)
Cell Motil. Cytoskeleton 45, 293–306
15. Pope, B. J., Gonsior, S. M., Yeoh, S., McGough, A., and Weeds, A. G. (2000) J.
Mol. Biol. 298, 649 – 661
16. Moriyama, K., and Yahara, I. (1999) EMBO J. 18, 6752– 6761
17. McGough, A., Pope, B., Chiu, W., and Weeds, A. (1997) J. Cell Biol. 138,
771–781
18. McGough, A., and Chiu, W. (1999) J. Mol. Biol. 291, 513–519
19. Ressad, F., Didry, D., Xia, G. X., Hong, Y., Chua, N. H., Pantaloni, D., and
Carlier, M. F. (1998) J. Biol. Chem. 273, 20894 –20902
20. Svitkina, T. M., and Borisy, G. G. (1999) J. Cell Biol. 145, 1009 –1026
21. Hatanaka, H., Ogura, K., Moriyama, K., Ichikawa, S., Yahara, I., and Inagaki,
F. (1996) Cell 85, 1047–1055
22. Leonard, S. A., Gittis, A. G., Petrella, E. C., Pollard, T. D., and Lattman, E. E.
(1997) Nat. Struct. Biol. 4, 369 –373
23. Fedorov, A. A., Lappalainen, P., Fedorov, E. V., Drubin, D. G., and Almo, S. C.
(1997) Nat. Struct. Biol. 4, 366 –369
24. Sun, H. Q., Yamamoto, M., Mejillano, M., and Yin, H. L. (1999) J. Biol. Chem.
274, 33179 –33182
25. McLaughlin, P. J., Gooch, J. T., Mannherz, H. G., and Weeds, A. G. (1993)
Nature 364, 685– 692
26. Wriggers, W., Tang, J. X., Azuma, T., Marks, P. W., and Janmey, P. A. (1998)
J. Mol. Biol. 282, 921–932
27. Lappalainen, P., Fedorov, E. V., Fedorov, A. A., Almo, S. C., and Drubin, D. G.
(1997) EMBO J. 16, 5520 –5530
28. Yonezawa, N., Nishida, E., Iida, K., Kumagai, H., Yahara, I., and Sakai, H.
(1991) J. Biol. Chem. 266, 10485–10489
29. Moriyama, K., Yonezawa, N., Sakai, H., Yahara, I., and Nishida, E. (1992)
J. Biol. Chem. 267, 7240 –7244
30. Ono, S., Baillie, D. L., and Benian, G. M. (1999) J. Cell Biol. 145, 491–502
31. Van Troys, M., Dewitte, D., Verschelde, J. L., Goethals, M., Vandekerckhove,
J., and Ampe, C. (1997) J. Biol. Chem. 272, 32750 –32758
32. Sutoh, K., and Mabuchi, I. (1989) Biochemistry 28, 102–106
33. Agnew, B. J., Minamide, L. S., and Bamburg, J. R. (1995) J. Biol. Chem. 270,
17582–17587
34. Moriyama, K., Iida, K., and Yahara, I. (1996) Genes Cells 1, 73– 86
35. Abe, H., Obinata, T., Minamide, L. S., and Bamburg, J. R. (1996) J. Cell Biol.
132, 871– 885
36. Smertenko, A. P., Jiang, C. J., Simmons, N. J., Weeds, A. G., Davies, D. R., and
Hussey, P. J. (1998) Plant J. 14, 187–193
37. Jiang, C. J., Weeds, A. G., Khan, S., and Hussey, P. J. (1997) Proc. Natl. Acad.
Sci. U. S. A. 94, 9973–9978
38. Lappalainen, P., and Drubin, D. G. (1997) Nature 388, 78 – 82
39. Sali, A., and Blundell, T. L. (1993) J. Mol. Biol. 234, 779 – 815
40. Ono, S., and Benian, G. M. (1998) J. Biol. Chem. 273, 3778 –3783
41. Pardee, J. D., and Spudich, J. A. (1982) Methods Enzymol. 85, 164 –181
42. Ono, S. (1999) Cell Motil. Cytoskeleton 43, 128 –136
43. Safer, D. (1989) Anal. Biochem. 178, 32–37
44. Zhou, Z. H., Hardt, S., Wang, B., Sherman, M. B., Jakana, J., and Chiu, W.
(1996) J. Struct. Biol. 116, 216 –222
45. Whittaker, M., Carragher, B. O., and Milligan, R. A. (1995) Ultramicroscopy
58, 245–259
46. Schroeter, J. P., and Bretaudiere, J. P. (1996) J. Struct. Biol. 116, 131–137
47. DeRosier, D. J., and Moore, P. B. (1970) J. Mol. Biol. 52, 355–369
48. Jones, T. A., Zou, J. Y., Cowan, S. W., and Kjeldgaard, M. (1991) Acta Crystallogr. Sect. A 47, 110 –119
49. Carson, M., and Bugg, C. E. (1986) J. Mol. Graphics 4, 121–122
50. Blanchoin, L., and Pollard, T. D. (1998) J. Biol. Chem. 273, 25106 –25111
51. Kim, E., Motoki, M., Seguro, K., Muhlrad, A., and Reisler, E. (1995) Biophys.
J. 69, 2024 –2032
52. Moraczewska, J., Strzelecka-Golaszewska, H., Moens, P. D., and dos
Remedios, C. G. (1996) Biochem. J. 317, 605– 611
53. Nishida, E., Muneyuki, E., Maekawa, S., Ohta, Y., and Sakai, H. (1985)
Biochemistry 24, 6624 – 6630
54. Lappalainen, P., Kessels, M. M., Cope, M. J., and Drubin, D. G. (1998) Mol.
Biol. Cell 9, 1951–1959
55. Yeoh, S., Pope, B., Mannherz, H. G., and Weeds, A. G. (1999) Mol. Biol. Cell 10,
(suppl.) 156
56. McKim, K. S., Matheson, C., Marra, M. A., Wakarchuk, M. F., and Baillie,
D. L. (1994) Mol. Gen. Genet. 242, 346 –357