Physiol Rev
83: 433– 473, 2003; 10.1152/physrev.00026.2002.
Actin Binding Proteins:
Regulation of Cytoskeletal Microfilaments
C. G. DOS REMEDIOS, D. CHHABRA, M. KEKIC, I. V. DEDOVA, M. TSUBAKIHARA,
D. A. BERRY, AND N. J. NOSWORTHY
Institute for Biomedical Research, Muscle Research Unit, Department of Anatomy and Histology,
University of Sydney, Sydney, Australia
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
I. Introduction
A. The cytoskeleton
B. What do we exclude from this review?
II. Structure and Dynamics of Monomeric Actin
A. The first actin crystals
B. Actin microcrystals and tubes
C. Actin monomer structure
D. MreB, an ancestral actin
E. Models of F-actin
F. Assembly of actin filaments
G. Elongation and annealing of F-actin
III. Actin Binding Proteins
A. ADF/cofilin family
B. Profilin family
C. Gelsolin superfamily
D. Thymosins
E. DNase I
F. Capping proteins
G. The Arp2/3 complex
IV. Role of Ternary Complexes in Regulating Actin Cytoskeleton Assembly
A. Cofilin, actin, and DNase I
B. Ternary complexes of actin with two or more ABPs
V. The Cytoskeleton and Pathology
A. Actin
B. Gelsolin
C. Tropomodulin
D. Connections between the extracellular matrix and the nucleus
E. Heart failure
F. Gene arrays
G. Single nucleotide polymorphisms, diseases, and the cytoskeleton
H. Cell signaling and actin microfilaments
VI. Unresolved Issues
A. Atomic structure of F-actin
B. Functional differences between actin isoforms
C. Ternary complexes of monomeric actin with ABPs
D. Undiscovered ABPs
E. Cooperative binding of ABPs along filaments
F. Cytoskeletal proteins in the nucleus
G. Disease and the cytoskeleton
H. Prokaryotic cytoskeletal elements
434
434
434
434
434
434
435
436
437
439
439
440
441
449
450
453
455
456
458
460
461
461
462
462
462
463
463
463
464
464
464
464
464
465
465
465
465
465
465
466
dos Remedios, C. G., D. Chhabra, M. Kekic, I. V. Dedova, M. Tsubakihara, D. A. Berry, and N. J. Nosworthy.
Actin Binding Proteins: Regulation of Cytoskeletal Microfilaments. Physiol Rev 83: 433– 473, 2003; 10.1152/physrev.00026.2002.—The actin cytoskeleton is a complex structure that performs a wide range of cellular functions. In
2001, significant advances were made to our understanding of the structure and function of actin monomers. Many
www.prv.org
0031-9333/03 $15.00 Copyright © 2003 the American Physiological Society
433
434
DOS REMEDIOS ET AL.
of these are likely to help us understand and distinguish between the structural models of actin microfilaments. In
particular, 1) the structure of actin was resolved from crystals in the absence of cocrystallized actin binding proteins
(ABPs), 2) the prokaryotic ancestral gene of actin was crystallized and its function as a bacterial cytoskeleton was
revealed, and 3) the structure of the Arp2/3 complex was described for the first time. In this review we selected
several ABPs (ADF/cofilin, profilin, gelsolin, thymosin ␤4, DNase I, CapZ, tropomodulin, and Arp2/3) that regulate
actin-driven assembly, i.e., movement that is independent of motor proteins. They were chosen because 1) they
represent a family of related proteins, 2) they are widely distributed in nature, 3) an atomic structure (or at least a
plausible model) is available for each of them, and 4) each is expressed in significant quantities in cells. These ABPs
perform the following cellular functions: 1) they maintain the population of unassembled but assembly-ready actin
monomers (profilin), 2) they regulate the state of polymerization of filaments (ADF/cofilin, profilin), 3) they bind to
and block the growing ends of actin filaments (gelsolin), 4) they nucleate actin assembly (gelsolin, Arp2/3, cofilin),
5) they sever actin filaments (gelsolin, ADF/cofilin), 6) they bind to the sides of actin filaments (gelsolin, Arp2/3), and
7) they cross-link actin filaments (Arp2/3). Some of these ABPs are essential, whereas others may form regulatory
ternary complexes. Some play crucial roles in human disorders, and for all of them, there are good reasons why
investigations into their structures and functions should continue.
II. STRUCTURE AND DYNAMICS
OF MONOMERIC ACTIN
A. The Cytoskeleton
Actin is only found in eukaryotes. It comprises a
highly conserved family of proteins that fall into three
broad classes: ␣-, ␤-, and ␥-isoforms. It is mainly located
in the cytoplasm, but it is also present in the nucleus
where it may or may not have motor-associated functions.
The highest concentrations (⬃20% of the total protein) of
actin are in striated muscles; however, significant quantities of actin are present in nonmuscle cells where it plays
a variety of roles including myosin-independent changes
of cells shape, motor-based organelle transport, regulation of ion transport, and receptor-mediated responses of
the cell to external signals.
The internal cytoskeleton of eukaryotic cells is composed of actin microfilaments, microtubules, and intermediate filaments. The cytoskeleton is dynamic and strong,
ever ready to adapt to demands on the cell. An important
property of actin is its ability to produce movement in the
absence of motor proteins. At the cell membrane microfilament assembly protrudes the membrane forward producing the ruffling membranes in actively moving cells.
Microfilaments can also play a passive structural role by
providing the internal stiffening rods in microvilli, maintaining cell shape, and anchoring cytoskeletal proteins.
The major focus of this review is to examine how actin
binding proteins (ABPs) control these processes. Finally,
we raise some interesting and challenging questions for
future research.
B. What Do We Exclude From This Review?
Actin microfilaments provide the “rails” along which
myosin “motors” perform work in a variety of cellular
functions. A major review of myosin motors (263) has
been published, and they are excluded from this review.
Microfilaments cooperate with microtubules via microtubule-associated proteins (MAPs) during the transport of
vesicles and organelles, and this interesting aspect of
microfilament function was recently reviewed (34). Actin
filaments also interact with intermediate filaments, a function that may play an important role in enabling extracellular stimuli to be transmitted to key targets like ribosomes and chromosomes deep within the cell. This field is
an emerging one. It will be covered by Quinlan et al. (235)
and is beyond the scope of this review.
Physiol Rev • VOL
A. The First Actin Crystals
Few people in the field of actin remember that Christine Oriol-Audit was the first to report the crystallization
of actin in the absence of an actin-binding protein (212).1
These crystals were achieved in the presence of polyethylene glycol but were too small to be useful for X-ray
diffraction at that time. If today’s high-powered X-ray
beam facilities were available then, they may well have
yielded the atomic structure of actin in the absence of an
ABP. Perhaps it was a cruel twist of fate that she sadly
died early in 2001 without knowing that the first structure
of uncomplexed actin was soon to be published (216).
The principal obstacle to crystallizing actin was its propensity to spontaneously self-assemble under solvent
conditions conventionally used to grow protein crystals.
B. Actin Microcrystals and Tubes
In their pioneering book on the biophysics of protein
polymerization, Fumio Oosawa and Sho Asakura (210)
1
Christine Oriol-Audit died in January 2001.
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
I. INTRODUCTION
ACTIN AND ACTIN BINDING PROTEINS
435
C. Actin Monomer Structure
Although the cocrystallization of actin with bovine
pancreatic DNase I was first reported in the late 1970s
(176), 13 years passed before the first atomic resolution
(2.4 Å) structure of actin was reported (137). The actin
monomers used in these crystals were lightly digested
with trypsin to remove the COOH-terminal three residues
before crystallization. Because the crystal structure of
DNase I had already been determined at 2.5-Å resolution
(272), it was a relatively simple task to subtract its structure from the actin-DNase I complex to obtain a clear
view of the actin monomer. The resulting structure (illustrated in ribbon form in Fig. 1A) produced a number of
surprises. For example, the large and small domains described at low resolution contained nearly equal numbers
of residues and were nearly equal in size. Both the NH2
and COOH termini were located in the same subdomain.
DNase I makes crystal contacts across the “top” of the
nucleotide cleft, thus explaining how it inhibits nucleotide
exchange. However, since these were cocrystals, did this
structure represent a monomer conformation when it was
not complexed to DNase I?
Monomeric (G-)actin has dimensions of ⬃67 ⫻ 40 ⫻
37 Å and a molecular mass of nearly 43,000 Da. The
overall features of the structure include four quasi-subdomains (here they are numbered 1– 4 but an alternative
nomenclature of IA, IB, IIA, and IIB has an historical
basis, Ref. 266), each having a repeating motif comprising
a multi-stranded ␤-sheet, a ␤-meander, and a right-handed
␤␣␤-unit. About 40% of the structure is ␣-helical. In Figure
1A, the monomer is oriented so that the top has about the
same orientation it would have in the filament with its
“pointed” end directed at the top of the page and the
“barbed” end toward the bottom.
A tightly bound nucleotide lies in a deep cleft in the
center of G-actin. This site is usually occupied by ATP or
ADP-Pi rather than ADP, which binds with ⬃10-fold lower
Physiol Rev • VOL
FIG. 1. A: actin monomer ribbon structure of G-actin from DNase I
cocrystals at 2.8-Å resolution (137). The numbered amino acid residues
were chosen because of their relevance to this review. B: actin monomer
ribbon structure of uncomplexed rabbit skeletal muscle actin at 1.54-Å
resolution (217).
affinity to Mg-G-actin and ⬃200-fold weaker binding with
Ca-G-actin (147). ATP binds as a complex with either
Mg2⫹ [dissociation constant (Kd) 1.2 nM] or Ca2⫹ (Kd 0.12
nM) (79, 270). Mg2⫹, the dominant cation in vivo, determines how tightly or weakly the nucleotide binds. Six or
seven relatively low-affinity divalent cation-binding sites
have been identified that appear to be important for actin
paracrystal (94, 271), and actin microcrystal formation
(73), but their physiological importance is not well understood. A notable feature of the structure is a two-stranded
“hinge” at residues 140 and 338 joining the large and small
domains. Tirion et al. (283) describe a “propeller” motion
between the two domains that produces an opening and
closing of the nucleotide cleft. This putative slewing mo-
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
predicted that actin, like tubulin, would form helically
tubular crystalline assemblies. We subsequently showed
that these structures could be formed in the presence of
the trivalent lanthanide ions, particularly Gd3⫹ (3, 87).
These lanthanide-induced actin tubes (composed of a
single layer of monomers) and microcrystals (bilayered
sheets) provided the first views of the shape of the actin
monomer (3, 73). The computed average image of the
monomer was “pear” shaped, having a “large” and a
“small” domain. Electron diffraction studies revealed
structure out to at least 14.5-Å resolution, but this was
simply not good enough to provide reliable molecular
details (74). In solution, actin filaments can form supramolecular assemblies called paracrystals (99) as well
as forming liquid crystalline arrays (67).
436
DOS REMEDIOS ET AL.
Physiol Rev • VOL
A significant structural difference actin crystallized
alone and complexed with an ABP protein is relatively
large coil-to-␣-helix conversion and a 10° rotation at the
top of subdomain 2. Although it is not clear if these
differences represent native cellular G-actin, Dominguez
et al. are currently completing the analysis of an uncomplexed ATP-G-actin structure that should satisfy their
critics.
The substantial conformational change reported by
Otterbein et al. (217) is well supported by other laboratories including our own. Dedova et al. (81) recently demonstrated, using fluorescence spectroscopy, that significant (1– 4 Å) changes occur in subdomains 1 and 2 when
cofilin and DNase I bind either separately or as a ternary
complex. Several other authors have suggested that actin
can undergo allosteric conformational changes, and this
is discussed in more detail in section IV. The major challenge for structural biologists now lies in relating the
structural changes to functional changes.
D. MreB, an Ancestral Actin
Actin is an essential and ubiquitous cytoskeletal component of all eukaryotic cells and is absent from prokaryotic cells. However, although it has long been suspected
that the evolutionary origins of actin lie in their prokaryotic ancestors, until 2001 no one had reported a credible
candidate. The first suggestion that MreB may be an ancestral actin gene was published 10 years ago (30), and
recently (136), it was localized in Bacillus to form distinct
filaments located close to the cell surface. The atomic
structure of these filaments was more recently reported
by Fusinita van den Ent and colleagues (289) at the MRC,
Cambridge, who provided convincing evidence that MreB
is indeed an ancestral actin gene. This actin look-alike is
present in all nonspherical bacteria.
Amino acid sequence homology to actin is limited to
15% (289), and although its overall size and shape strongly
resemble actin, there are differences. The sequence corresponding to the actin DNase I-binding loop in subdomain 2 is larger, and the COOH terminus, located in
subdomain 1, is 20 residues longer. A 2.1-Å resolution
crystal structure reveals that, like actin, MreB has two
major domains separated by a nucleotide-binding cleft.
Also, like actin, each domain has two subdomains with
essentially the same topology. The structure of MreB is
shown in Figure 2.
MreB can self-assemble into 51-Å-wide filaments that
are about half the diameter of actin microfilaments, but
unlike F-actin, they are linear (nonhelical) polymers resembling the linear polymers seen in actin crystalline
sheets first reported by ourselves (87) and later by others
(3). The axial repeat for MreB microfilaments is 51.1 Å,
somewhat less than the axial repeat of F-actin (55 Å) (Fig.
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
tion is permitted because of this hinge and is viewed as a
rigid body movement between the subdomains.
There are several other atomic structures of actin. In
1993, McLaughlin’s group at the Medical Research Council (MRC) in Cambridge (186) reported the structure of
actin complexed with gelsolin segment 1. This segment
binds to the bottom of the monomer, making contacts
with subdomains 1 and 3 (see discussion below). Unlike
the DNase I-actin structure, the COOH-terminal three residues were not cleaved before crystallization. However,
the DNase I binding loop was not well resolved, presumably because it was not sufficiently stabilized in the absence of DNase I. In the same year Schutt et al. (259)
reported the structure of spleen actin (a ␤-isoform) complexed to profilin at a resolution of 2.55 ÅA. This group
had been refining their structure for many years and
demonstrated that, although there was broad similarity to
the structure of McLaughlin et al. (186), there were significant differences that they attributed to the sequence
differences between the actin isoforms.
The Schutt/Lindberg consortium that captured actin
in a different conformation subsequently reported a
fourth actin-profilin structure. They again used bovine
␤-actin cocrystallized with bovine pancreatic profilin (60).
They named the two structures the open and closed states
and used them to argue that the transition between the
two states represented a structural change that could
convert their “ribbon” polymer into conventional F-actin.
In 2001, the inevitable happened. Dominguez et al. at
the Boston Biomedical Research Institute succeeded in
producing small, high-quality crystals of actin that were
not complexed to ABPs (Fig. 1B). This finding was reported at the Boston meeting of the Biophysical Society
colleagues (216) and was subsequently published in Science (217). They achieved the result that had eluded so
many others by modifying the COOH-terminal cysteinyl
(Cys-374) with a rhodamine label that suppressed the
tendency of actin to polymerize.
Dominguez and colleagues (217) were also the first to
crystallize actin with ADP present in the nucleotide-binding cleft. This was important because we know that the
critical concentration of ADP-G-actin is about an order of
magnitude higher than for ATP or ADP-Pi-G-actin (241).
Kabsch et al. (137) had also produced crystals of ADP-Gactin and showed that the ADP structure was not remarkably different from the ATP form. However, the Kabsch
crystals were formed in the presence of ATP, and the
nucleotide was cleaved to ADP subsequent to crystallization.
The structure of Dominguez et al. significantly differs
from the structure of Kabsch et al., particularly in subdomain 2 where DNase I binds. However, it could be argued
that this Boston/Heidelberg difference is because ADP
was present before the crystals were formed, i.e., before
the crystal contact points were established.
ACTIN AND ACTIN BINDING PROTEINS
437
analogs and control the shape of bacteria. However, we
know nothing of how assembly of this structure is regulated, or whether it binds to motor proteins. Figure 3
illustrates the striking similarities between the crystal
structure of protofilaments of actin (right) and MreB
(left). Nearly identical molecular orientations and contacts are seen between monomers for the two proteins.
Conversion of a pair of actin protofilaments into F-actin is
achieved by simply twisting a pair of actin protofilaments
to conform to the actin filament symmetry (see below).
E. Models of F-actin
3). These polymers form spirals beneath the bacterial cell
wall, and although their precise function remains elusive,
it is likely that MreB filaments behave like microfilament
FIG. 3. A comparison of crystal structure of MreB
(left) with a model of actin protofilaments (right) derived from linear polymers along a single strand of
F-actin. Five monomers are illustrated in both protofilaments. Although the axial repeats between monomers differ, the molecular contacts between monomers in protofilaments of MreB are essentially the
same as the contact between actin monomers along the
two strands of F-actin (van den Ent, personal
communication).
Physiol Rev • VOL
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
FIG. 2. Crystal structure of MreB showing the four-domain motif
seen in the actin monomer structure (136). The NH2 and COOH termini
are located in subdomain 1A (equivalent to subdomain 1 of actin) (F. van
den Ent, personal communication).
The field has made substantial progress in understanding the structure of F-actin. Nearly 40 years ago Jean
Hanson and Jack Lowy were the first to describe the
helical nature of actin filaments (116). In this model,
F-actin can be viewed either as a single-start, left-handed
tightly wound helix of monomers (the so-called genetic
helix) or as a two-start, right-handed long-pitch helix. The
conventional view of F-actin is the two-start helix because
it is probable that the monomer-monomer affinity is stronger along the two long-pitch strands than between those
strands (128).
No atomic structure has yet been determined for
F-actin, although highly plausible models have been proposed based on the model of Holmes et al. (128). Figure 4
illustrates five monomers in a filament. Viewed along the
438
DOS REMEDIOS ET AL.
genetic helix, monomers rotate by ⫺166° and have an
axial translation of 27.5 Å. There are 13 monomers in 6
turns with a pitch of 59 Å yielding a filament diameter of
⬃90 –100 Å (128). Contacts between the monomers along
this helix occur across the diameter of the filament. The
two-start long-pitch helix is right-handed with a somewhat variable half pitch of 360 –390 Å comprising 12–14
monomers per half turn. In this model each monomer is
still surrounded by four others. The center-to-center distance between monomers along this helix is ⬃55 Å. These
filaments have a distinct structural polarity that was first
noticed when the filaments were “decorated” with myosin
Physiol Rev • VOL
fragments, but it can also be seen in good images of
F-actin.
Thus, despite the passage of nearly 40 years and the
best efforts of laboratories in Germany, the United Kingdom, the United States, Japan, and elsewhere, the atomic
structure of F-actin remains elusive. A major part of the
problem is the inherent disorder in linear aggregations of
filaments. Even if they could be aligned with the same
polarity, it is difficult to pack F-actin complexed to phalloidin (a 7-residue peptide that dramatically stabilizes
actin filaments) so there are precise contacts between
filaments. Devices like the innovative trumpet-shaped
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
FIG. 4. The Holmes model of F-actin showing six
monomers, three in each of the two-start long-pitch helix
originally described by Holmes et al. (128).
ACTIN AND ACTIN BINDING PROTEINS
Physiol Rev • VOL
Thus it is now more common to view F-actin as a dynamic, responsive structure than a passive structural element.
F. Assembly of Actin Filaments
The most authoritative dissertation on the biophysics
of F-actin is Oosawa and Asakura’s slim volume, Thermodynamics of the Polymerization of Protein (210). Despite
its age, this book continues to provide a comprehensive
theoretical background for understanding the assembly of
actin into filaments and higher order assemblies. The
conditions under which actin monomers self-assemble in
vitro are well documented (211), although it is fair to say
they are less well understood for in vivo assembly.
Polymerization is essentially a condensation reaction. The main features of this process are 1) a slow initial
association to a dimer that is more likely to rapidly dissociate to monomers than to assemble; 2) the formation
of a stable trimer that represents the nucleus of polymerization, a state where actin assembly is more likely than is
disassembly; and 3) the elongation phase during which
actin monomers are rapidly assembled. Solvent conditions that promote polymerization include high ionic
strength (KCl concentrations ⬎50 mM), neutral or slightly
acidic pH, high Mg2⫹ (rather than high Ca2⫹), and elevated temperature (9, 111, 294), in other words, the conditions found in cells.
In addition to these three processes, actin filaments
are in a continuous state of assembly/disassembly. As a
consequence, in the steady state, a small but finite concentration of actin monomers will be present in any filament population (140, 211). This concentration of free
monomers in equilibrium with a population of actin filaments is referred to as the critical concentration and is
dependent on solvent conditions, particularly on the presence of certain ABPs and/or actin ligands such as phalloidin (80) or latrunculin A (193).
G. Elongation and Annealing of F-actin
Elongation involves association and dissociation of
monomers from the filament. These processes can occur
at either end of the filament, but association predominantly occurs at the barbed end and dissociation at the
pointed end. The nucleotide binding site of monomeric
actin is almost exclusively associated with ATP in vivo.
Conversely, the majority of F-actin subunits contain
bound ADP. While the G-actin bound ATP is readily exchangeable with solvent nucleotides, the ADP of F-actin is
essentially nonexchangeable (121). Hydrolysis of bound
ATP was originally thought to be tightly coupled to the
polymerization process (210, 300); however, subsequent
investigations (44, 46, 77, 221) revealed that a time lag
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
quartz tubes used by David Popp et al. (233) achieved
greatly improved filament alignment that yielded the first
low-angle X-ray diffraction data. More recently, Yuichiro
Maeda (who also worked in Heidelberg and has since
returned to the RIKEN SPring8 Institute synchrotron facility in Japan) and colleagues achieved even better alignment of F-actin without the help of phalloidin using very
high (13.5 Tesla) magnetic fields. They already have structural data out to 5-Å resolution (207), and we can expect
them to report further progress in the near future.
In the meantime, we must content ourselves with the
available models of F-actin based on data from low-resolution electron microscope images. The original model
(128) was created by fitting the atomic structure of the
actin monomer (137) to models based on the limitedresolution X-ray diffraction data from aligned filaments.
This model is illustrated in Figure 4. Subsequently, Michael Lorenz et al. (167) refined the Holmes (128) model
using data from actin mutations. In the same year Tirion
et al. (283) described the propeller motion of the two
major domains about the hinge and then refined the Factin model to take into account domain motion. Thus it
emerged that the actin monomer was not the rigid, static
(double strand of pearls) structure so commonly depicted. Monomers in this model are positioned slightly
tangential to the filament axis, with subdomain 1 (the
major myosin S-1 binding site) being located at the highest radius.
F-actin displays an arrowhead-like appearance when
decorated with myosin subfragment 1 (S-1) (131, 188). For
this reason the opposite ends of the filament have been
named the “barbed” and “pointed” ends. These ends correspond to exposed subdomains 1 and 3 and subdomains
2 and 4, respectively.
Quite a different model was proposed by Schutt,
Lindberg, and colleagues (259) based on crystallographic
principles and known examples from other structures in
biology. In profilin-actin crystals, a “ribbon” structure can
be seen in which monomer subdomains 1 and 2 lie close
to the ribbon axis. Alternating monomers displayed front
and back views. They proposed that monomers were
reoriented to produce F-actin. In essence, this group
turned the model of Holmes et al. (128) inside out by
rotating the actin monomer about 180° in the plane of the
filament axis. The effect of this was to locate subdomains
1 and 2 closer to the filament axis, leaving subdomains 3
and 4 at higher radii. These authors further suggested that
the subdomains were able to undergo significant movements relative to each other and that this could feasibly
be built into a mechanism of contraction (258).
Low-resolution (25–30 Å) electron microscope studies have revealed that the actin filament can exist in
multiple conformations depending on the type of bound
cation and nucleotide, the isoform of actin (213, 214), and
the presence of other proteins bound to actin (182, 218).
439
440
DOS REMEDIOS ET AL.
exists between the incorporation of ATP-G-actin onto the
filament end and hydrolysis of the bound nucleotide. The
kinetics of ATP hydrolysis and release of its product, Pi
(43), suggests that the bound ATP of newly added actin
subunits is hydrolyzed through two distinct sequential
steps as illustrated in the following reaction scheme
ATP-G-actin 7 ATP-F-actin 3
Fast
ADP-Pi-F-actin 7 ADP-F-actin⫹Pi
The rate of ATP hydrolysis was calculated to be 10 times
higher than the rate of Pi release, suggesting that ADP-PiF-actin is the major intermediate in the nucleotide hydrolysis process. Conversion of ATP to ADP-Pi and then to
ADP ⫹ Pi is not random but primarily occurs at the
growing (barbed) ends of filaments (47).
Depolymerization of actin is not simply the opposite
of polymerization, principally because actin cannot regenerate ATP from ADP and Pi (45). Instead, dissociated
ADP-actin subunits rapidly exchange their bound ADP for
ATP in solution (200), a process that is accelerated by
profilin (see below). Polymerization of actin is not dependent on nucleotide hydrolysis. It is possible under special
circumstances to form F-actin from G-actin containing
either no bound nucleotide (141, 227) or a nonhydrolyzable analog of ATP (63). However, nucleotide hydrolysis
is required for the normal function of F-actin.
The structural polarity of the actin filament and the
irreversible nature of ATP hydrolysis during actin assembly have implications for the rate and direction of filament
growth at opposite ends of F-actin. The critical concentration for the pointed end is 12- to 15-fold higher than for
the barbed end under physiological conditions (301). This
difference may result in the unidirectional growth of the
actin filament due to a continual flux of actin subunits
from the pointed to the barbed end of the filament. This
reaction is called “treadmilling” (300).
Under in vivo conditions one might expect that all or
most of the actin exists in an assembled (filamentous)
form. Slow addition at the barbed end and even slower
dissociation at the pointed end of the filaments produces
a rate of treadmilling (300) of monomers (⬃2 ␮m/h) that
is ⬃200-fold slower than observed in vivo. Monomers
move progressively along the filament from the barbed
end (the end associated with Z-disks of the sarcomere, or
the cell membrane where ruffling occurs in motile cells)
toward the free pointed end of the filament (located in the
middle of the sarcomere or oriented away from the cell
membrane). ABPs present in vivo regulate different aspects of the assembly/disassembly process. These include
Physiol Rev • VOL
III. ACTIN BINDING PROTEINS
Actin binds a substantial number of proteins collectively called ABPs. Some years ago, Pollard and Cooper
(231) identified a large number of ABPs, and recently we
counted 162 distinct and separate proteins without including their many synonyms or isoforms. No doubt more
will be identified. Many of the known ABPs bind to the
same loci on the surface of actin and therefore can be
expected to compete. A few bind with positive cooperativity and tend to form ternary complexes (see below) but
rather more bind with negative cooperativity. In myofibrils, at least eight sarcomeric proteins bind to the thin
filaments. At least 12 ABPs are membrane-associated proteins, and another nine are membrane receptors or ion
transporters. Thirteen ABPs cross-link actin filaments,
whereas others enable filaments to interact with other
elements of the cytoskeleton. Microfilaments probably do
not interact directly with microtubules and/or intermediate filaments but do so via linker proteins.
Actin also binds ⬃30 other ligands including drugs
and toxins. Thus the sheer number of ligands that have a
significant affinity for actin strongly suggests there is
probably a large number of binding sites that cover much
of the exposed surface of the molecule. A comprehensive
list is available from Actin (266), the Guidebook to the
Cytoskeletal and Motor Proteins (151), and most recently
from the two-volume Molecular Interactions of Actin
(88). Attempts to classify these ABPs leave many “orphans” that do not fit into families, so any attempt to
group them is bound to be somewhat arbitrary. Classifications according to the function of ABPs or their consensus sequences can also be problematic.
Several types of ABPs facilitate disassembly and assembly. Below is a review of the most common ABPs.
Classification of these ABPs can be reduced to seven
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
Slow
filament stabilizers (e.g., tropomyosin), capping proteins
(e.g., CapZ, tropomodulin), ABPs that promote branching
(e.g., Arp2/3), and ABPs that sequester G-actin and thus
maintain a pool of monomers in solution (e.g., thymosin
␤4, profilin).
Finally, filament lengths are affected by fragmentation and annealing. Fragmentation can arise from thermal
motion in vitro, but in vivo much of this will be constrained by the other contents of the cytoplasm. Annealing occurs when an existing filament binds to the appropriate end of a second filament. This has been observed
directly using fluorescent-phalloidin-labeled single filaments (Ishiwata, personal communication) and has been
quantified (2.2 ␮M⫺1䡠s⫺1) under similar conditions (145).
Arp2/3 (discussed below) is believed to create branch
points in actin microfilaments by “capturing” existing filaments.
ACTIN AND ACTIN BINDING PROTEINS
A. ADF/Cofilin Family
Unlike so many of the ABPs, the actin depolymerizing factor (ADF)/cofilin family of proteins is expressed in
Physiol Rev • VOL
virtually all eukaryotic cells. They are relatively small
(15–19 kDa) proteins that exist in multiple isoforms. Their
main functions include the rapid recycling of actin monomers associated with membrane ruffling and with cytokinesis. Different genes encode for ADF and cofilin, but
although it is common to regard them as synonymous,
they are distinctly different.
1. Members of the family
ADF/cofilin was first discovered and purified in 1980
from embryonic chick brain extracts by Bamburg et al.
(20). Since then, the family has grown to include a number
of related proteins, including invertebrate depactin
(named because it depolymerizes actin) (173); porcine
ADF or destrin (destroys F-actin) (191); cofilin (cosediments with filamentous actin) (1), Acanthamoeba actophorin (236); Dictyostelium coactosin (82), Drosophila
twinstar or D-61 (91); unc-60A and unc-60B from Caenorhabditis (185); Xenopus Acs (or XAC1 and XAC2) (2);
and finally Toxoplasma ADF (4). All of these share considerable (30 – 40%) amino acid sequence identity. Furthermore, two other major protein families are related to
ADF through the presence of an ADF homology domain.
One protein has a duplication of this domain and is consequently called twinfilin; the other contains a single ADF
homology domain linked to another motif and encodes
the drebin family of proteins (16).
Despite this somewhat confusing array of homologs,
vertebrates have genes for only two forms, ADF and
cofilin. The names alone suggest that one depolymerizes
F-actin (ADF) while cofilin cosediments with F-actin, but
actually both can elevate the levels of monomeric actin
and both can bind to F-actin. Only one isoform of ADF is
known in mammals (and birds), whereas two are known
for cofilin. ADF and cofilin are clearly different but related
proteins.
Cofilin is diffusely distributed in the cytoplasm of
quiescent cells. However, in active cells, it translocates to
cortical regions where the actin cytoskeleton is highly
dynamic and drives the ruffling of membranes of motile
cells (18, 309), the cleavage furrow of dividing cells (197),
the advancing of neuronal growth cones (18, 198), and
myofibrillogenesis (206). ADF and cofilin are not necessarily expressed at the same level in all tissues. In adults,
ADF is relatively highly expressed in nerves, intestine,
kidney, and testes (18), whereas cofilin levels are higher
in hematopoietic tissues, bone osteoclasts, and fibroblasts (309). In baby hamster kidney cells, ADF constitutes 0.4% of the soluble protein, while cofilin accounts
for 1.3% (148). The total ADF and cofilin amounts to 20
␮M in cells where the total actin concentration is more
than three times higher (67 ␮m) (148) (see Table 1).
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
groups. 1) Monomer-binding proteins sequester G-actin
and prevent its polymerization (e.g., thymosin ␤4, DNase
I). (Note: C. E. Schutt and Uno Lindberg are completing
an extensive review of Actin Monomer Binding Proteins
to be published in the Protein Profile series of Oxford
University Press.) 2) Filament-depolymerizing proteins
induce the conversion of F- to G-actin (e.g., CapZ and
cofilin). 3) Filament end-binding proteins cap the ends of
the actin filament preventing the exchange of monomers
at the pointed end (e.g., tropomodulin) and at the barbed
end (e.g., CapZ). 4) Filament severing proteins shorten
the average length of filaments by binding to the side of
F-actin and cutting it into two pieces (e.g., gelsolin).
(Note: H. Hinssen is completing an extensive review on
the Gelsolin Family to be published in the Protein Profile
series of Oxford University Press.) 5) Cross-linking proteins contain at least two binding sites for F-actin, thus
facilitating the formation of filament bundles, branching
filaments, and three-dimensional networks (e.g., Arp2/3).
6) Stabilizing proteins bind to the sides of actin filaments
and prevent depolymerization (e.g., tropomyosin). 7) Motor proteins that use F-actin as a track upon which to
move (e.g., the myosin family of motors). Here we consider only groups 1– 4.
ABPs are not limited to one class, for example, gelsolin is capable of severing and capping the barbed end of
actin filaments, and the Arp2/3 complex can nucleate
filament formation, elongate filaments, and establish
branch points in actin networks (69). In this review we
have selected eight ABPs that are biologically relevant
and have atomic structures or good working models.
The rapid assembly of actin filaments is the principal
driving force behind many forms of cell locomotion. Cells
can migrate at rates up to ⬃0.5 ␮m/s (264). This means
that filaments must have a net rate of elongation that is
slightly less than 200 monomers/s. Principally this occurs
at their barbed ends where MgATP-actin assembles about
five times faster than MgADP-actin (149). The exchange of
ATP for ADP in filaments is so slow (of the order of days,
Ref. 201) it can be ignored. At steady state, filament length
is constant, but there is a slow turnover (⬃2 ␮m/h) (264)
of monomers due to a treadmilling process that involves
the hydrolysis of ATP to ADP-Pi-actin and then a slower
dissociation of Pi leaving most of the monomers in the
filament with bound MgADP. Thus, if pure actin filaments
treadmill very slowly and if actin assembly is the driver of
cell motility, then the intrinsic rate of treadmilling must
increase in vivo. This is the task of the ABPs.
441
Physiol Rev • VOL
83 • APRIL 2003 •
www.prv.org
(ATP-GA) Kd 0.1
␮M at the
barbed end;
Kd 0.6 ␮M at
the pointed
end (228)
Binding to actin
filaments
Binding to
G-actin
Concentration in
cells
2–5, 40–50, 61–
64, 89–100,
110–112, 130–
131, 166–173,
195–197, 202–
205, 243–247,
266–269, 285–
289, 322–325,
357–360, and
375 (128)
65–70 ␮M (19);
300 ␮M (299)
20% of total
muscle
protein (266)
Weak
375 amino acids;
43 kDa (288)
Amino acids;
molecular
mass
Residues on
actin that bind
ABP (Fig. 6)
Figs. 1, 4, 6
(row 1)
Actin
Relevant figures
Actin Binding
Protein
Mg-ATP Kd 0.5
␮M; Mg-ADP
Kd 5 ␮M
(261); Mg-ATP
Kd 0.1 (6)
Delivers ATP-GA
to the barbed
end of FA
pH ⱕ7, ADP-FA
(0.3 ␮M); ATP-FA
(10 ␮M) (42, 202)
20–100 ␮M (230)
20 ␮M (0.44% of
total protein) (18)
pH 8, ATP-GA Kd 8
␮M; ADP-GA Kd
0.1 ␮M (107, 240)
113, 166, 167,
169, 171–173,
284, 286–288,
290, 354, 355,
361, 364, 369,
371–373 and
375 (259)
125–153 amino
acids; 138
amino acids
14.9 kDa (138)
Figs. 6 (row 2),
9, 10
Profilin
1–12, 25, 147, 167,
288, 292, 328, 334,
341, 344, 345, 348,
357–375 (304)
165–168 amino
acids; 18–19 kDa
(261)
Figs. 5B, 6 (row 1),
7B, 8, 10
ADF/Cofilin
1. Actin binding protein characteristics
Mg-ATP-GA Kd
1.7 ␮M; MgADP-GA ⫽ 80
␮M (42)
Segment 1 to
barbed end Kd
1 ␮M;
segments bind
to 2 GAs 0.07
␮M (257)
5–60 nM at
barbed end
(146); 50 nM
(262)
Weak (41)
Up to 500 ␮M
(249)
1–4, 41, 154, 155,
167 (247, 251)
42 amino acids,
4,963 Da (248)
Figs. 6 (row 2),
14
Thymosin ␤4
⬃5 ␮M
1–18, 112–117,
144, 146, 148,
167, 169, 285–
375, 334, 350,
351, 354, 356–
375, 374 and
375 (186)
Figs. 6 (row 3),
7C, 11, 12A,
12B, 13
720–730 amino
acids in 6
subunits, 80
kDa (86)
Gelsolin
384 (Arp2) ⫹
391 (Arp3) 7
subunit
protein 220
kDa (172)
N/A (actin
subdomains 2
and 4)
␣ ⫽ 286, ␤ ⫽
277 amino
acids; 59 kDa
(38)
2 ␮M Arp2, 5
␮M Arp3 (41,
143)
Binds weakly to
actin dimers
(242)
Kd 10 ␮M (242,
262)
0.6 ␮M (1:500
actin
monomers)
(305)
Only the ␤subunit can
bind GA (53)
1:1, Kd 0.5–1.5
nM (53)
0.05 nM (177)
Kd 0.1 mM (177)
338–348 (156),
360–372 (131)
Fig. 17
Arp2/3 (to
Filaments)
Figs. 6 (row 4),
16
CapZ
(Not known)
39–46, 50, 53,
60–64, 68, 69,
202–204 and
207 (137)
260 amino acids,
29 kDa (222)
Figs. 6 (row 1),
15
DNase I
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
TABLE
442
DOS REMEDIOS ET AL.
83 • APRIL 2003 •
www.prv.org
No
No
Promotes
nucleotide
exchange
(203); PIP2
inhibits
binding (110)
No
Yes
No
PIP2 inhibits
depolymerization
(283); inhibits
nucleotide
exchange (27)
No
Stronger along a
strand
Mg-ATP (Kd 1.2
nM) (79)
PIP2 inhibits
severing (275)
Yes
Yes
No
Yes
Yes
No
Yes
ABP, actin binding protein; Kd, dissociation constant; ADF, actin depolymerizing factor.
No
Yes
Inhibits
nucleation
With cofilin,
disassembly is
fast: 125 s⫺1
(85)
No
No
No
Inserts monomers
0.15 ␮M (262).
No change in off
rate
Cross-links
filaments
Severs FA
Side binding to
F-actin
Important
functional
ligands
Very weak
Inhibits
nucleotide
exchange
(107)
No
No
No
No
No
? Yes
? Yes
PIP/PIP2 inhibit
binding of
CapZ to actin
filaments
(123)
PIP2 inhibits
nucleotide
exchange
(127); inhibits
binding to
actin (130)
Arp2 binds
profilin (143);
activated by
WASP, Scar 1
(103)
No
Yes Kd 0.5 ␮M
No
No
No
No
No
No
No
Kd 10 nM (242,
262)
Yes
Yes (302) near
membrane
(143)
Yes at 70°
Blocks
exchange (Kd
1 nM) (255)
No (Tmod binds
pointed ends)
Yes
Yes
No
No
No
Yes
(disassembles
FA)
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
Physiol Rev • VOL
Caps FA
Nucleates
assembly
Barbed end
effect
Accelerates off rate
at pointed end, 9
s⫺1 (240)
Critical
concentration
is 0.6 ␮M
(228). ATP on
rate 0.5 ␮M/s;
off rate 1 ␮M/
s; ADP on
rate 0.1 ␮M/s;
off rate 0.2
␮M/s
Critical
concentration
is 0.1 ␮M
(228); ATP on
rate 5 ␮M/s;
off rate 1 ␮M/
s; ADP on
rate 1 ␮M/s;
off rate 6 ␮M/
s
N/A
Nucleus is an
actin trimer
Pointed end
effect
ACTIN AND ACTIN BINDING PROTEINS
443
444
DOS REMEDIOS ET AL.
Recently, Vartiainen et al. (291) clarified the ADF/
cofilin nomenclatures for vertebrates. Using mice, they
proposed that cofilin 1 is expressed in most tissues of
embryonic and adult cells, cofilin 2 is expressed in muscle
cells, and ADF is limited to epithelial and endothelial
cells. More importantly, they defined the functional similarities and differences between them.
2. Structure of ADF/cofilin
Porcine destrin was the first structure in this family
of proteins to be solved by NMR spectroscopy (118).
Subsequently, atomic structures of three more members
of the ADF/cofilin family have been determined. A ribbon
representation of yeast cofilin is shown in Figure 5A
Physiol Rev • VOL
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
FIG. 5. A: the atomic structure of yeast cofilin solved
at 1.8 Å showing the NH2 and COOH termini. Putative sites
for binding F-actin and G-actin (purple helix) are located
in the COOH-terminal region of the molecule. The solid
spheres identify residues that are essential for actin binding. [Modified from Fedorov et al. (95).] B: the crystal
structure of yeast cofilin (red) was fitted using molecular
dynamics to the actin monomer structure (grey) using
energy minimization. This approximate orientation is preserved in views of subsequent actin-ABP complexes.
(Model kindly provided by Willy Wriggers.)
ACTIN AND ACTIN BINDING PROTEINS
Physiol Rev • VOL
the presence of muscle and nonmuscle tropomyosins
(17). Tropomyosin is located deeper into the cortex from
the site of active actin assembly and its presence probably
inhibits cofilin binding (28). Cofilin and phalloidin also
compete for binding to F-actin (120), even though their
binding sites are separated by 15–20 Å. McGough et al.
(182) proposed that an allosteric change in the twist of
F-actin induced by cofilin may inhibit phalloidin from
binding between monomers close to the filament axis.
The blue residues in Figure 6 represent the probable
minimal molecular contacts for ADF/cofilin on actin. The
principal motif shown in Figure 5, A and B (a 4-stranded
␤-sheet surrounded by 4 ␣-helices), is also found in an
unrelated sequence of gelsolin segment 1. It is therefore
not surprising that cofilin competes with gelsolin segment
1 and profilin (85) for binding to G-actin. Both profilin and
gelsolin segment 1 bind to the barbed end of the actin
monomer (subdomains 1 and 3), even though only Glu167 and Tyr-169 (see Table 1) are common binding sites.
Because cofilin competes with both proteins, it is possible
that it too interacts with residues in the 162–176 loop of
actin subdomain 3 (see Fig. 1A).
The structural sites to which cofilin binds along the
outside of the filament are not known in atomic detail, so
the blue residues in Figure 6 are based on chemical
cross-linking and other data. At low resolution (25 Å),
ADF/cofilin bridges subdomains 1 and 3 of an upper actin
to subdomains 1 and 2 of a lower actin in a filament. The
major axis of cofilin makes a 30° angle with the plane
normal to the helical axis of the filament. It is centered
(axially) at about the position of subdomain 2 of the lower
actin subunit and radially at the cleft between subdomains 1 and 3 of the upper actin subunit. Electron cryomicroscopy and helical reconstructions of F-actin decorated with cofilin revealed its ability to reduce the angle of
rotation between subunits along the short-pitched lefthanded genetic helix by 3–5° (182). There is no effect on
the axial rise per actin subunit, and consequently, the
long-pitch helical repeat of F-actin is decreased from
⬃360 to 270 Å, and the diameter increases slightly. The
change of helical pitch can even be seen in negatively
stained filaments in the absence of cofilin (Fig. 7a) and in
the presence of stoichiometric cofilin (Fig. 7b). We are
grateful to Roger Craig for providing these micrographs.
3. Cellular functions
A) TREADMILLING. The ADF/cofilin family is almost single-handedly credited for the high rate of treadmilling of
monomers in actin filaments in vivo. This is achieved in
part by increasing (by 30-fold) the off rate (9 s⫺1, Ref. 240)
at the pointed ends of filaments without changing the off
rate at the barbed ends (42). However, in the presence of
profilin (see sect. IIIB), the rate of pointed-end disassembly is even faster (125 s⫺1, Ref. 85). The resulting elevated
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
solved at 1.8 Å (95). Crystal structures have also been
determined for both Acanthamoeba actophorin (161) and
a plant ADF from Arabidopsis thaliana (32). No atomic
structure has been published for any vertebrate cofilin or
for any muscle ADF/cofilin, but the atomic coordinates
for a high-resolution NMR structure for chick cofilin from
embryonic skeletal muscle were recently deposited in the
BioMagRes Bank (http://bmrb.wisc.edu) under BMRB accession number 5177 (11).
Indirect methods have been used to examine the sites
on cofilin that bind to actin. Despite the best efforts of
several groups, these two proteins have not yet been
cocrystallized, and consequently, unlike DNase I, gelsolin
segment 1, and profilin, the precise contact sites between
cofilin and actin are not known. Structural homologies
between cofilin, gelsolin, and profilin strongly suggest
they bind to the same region of G-actin (i.e., to subdomains 1 and 3). However, the functional products of systematic mutagenesis of yeast cofilin indicate that the cofilin-actin interaction is distinctly different.
Competitive binding of a synthetic dodecapeptide
(residues 104 –115 of vertebrate cofilin) has implicated
this region in the interaction of cofilin with monomeric
but not with filamentous actin (308). This peptide is
largely conserved in other members of the ADF/cofilin
family including destrin and depactin, suggesting that it
may be a consensus sequence essential for actin binding
and depolymerizing activities.
Mutagenesis of surface residues for both yeast (158)
and chick (154) cofilin have implicated the NH2-terminal
region in G-actin binding. Zero-length cross-linking of
G-actin to vertebrate cofilin by 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (308) revealed that either Lys-112
and/or Lys-114 of cofilin is in direct contact with actin.
Mutations of these cross-linked residues (192), or the
corresponding Arg-96 and Lys-98 of yeast cofilin, totally
abolished actin binding. Thus a significant fraction of the
surface residues has been implicated in the binding of
cofilin to G- and F-actin (Fig. 5A, circles) and within these
areas, eight or nine residues (solid spheres) are considered essential. Despite the absence of cocrystals, the
cofilin structure has been “docked” onto the Kabsch et al.
(137) actin structure (Fig. 5B) using energy minimization
techniques (304). This figure provides a clear view of the
structural relationship between these two proteins and is
presented so it can be compared with the binding sites for
profilin and DNase I (see below).
Bamburg (16) recently stressed the importance of
switching from an ADF/cofilin system to a tropomyosinbased regulation of F-actin function. Tropomyosin binds
along the grooves of the long pitch helix of F-actin. In
myofibrils, where tropomyosin is constitutively expressed, the binding of tropomyosin and cofilin to actin
filaments is mutually exclusive and competitive (311).
Thus the depolymerizing activity of cofilin is inhibited by
445
446
DOS REMEDIOS ET AL.
Physiol Rev • VOL
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
FIG. 6. The molecular contact sites between actin (shown here as
Van der Waals representations) and ABP sites. “Front” (A) and “back”
(B) views of the monomer are shown where the binding sites are
colored: row 1: yellow, actin monomer-monomer interaction sites; light
green, DNase I; and blue, ADF/cofilin; row 2: dark green, thymosin ␤4;
and red, profilin; row 3: pink, gelsolin segment 1; and black, vitamin
D-binding protein; row 4: red, CapZ sites. There is significant overlap
between the actin binding sites for ADF/cofilin, profilin, gelsolin segment 1, and CapZ and are therefore shown on separate figures. These are
the minimal contacts and are based on chemical cross-linking and
mutagenesis data.
concentration of cofilin-G-actin in the cytoplasm can be
rapidly recycled at the barbed end of filaments, provided
that ATP can replace the actin-bound nucleotide. This
concept is illustrated in Figure 8. In vivo, the concentration of ADF/cofilin is low relative to other ABPs such as
profilin and thymosin ␤4 (see discussion below). ADF/
cofilin “decoration” of actin filaments is probably restricted to their “aged” ends where ATP has been converted to ADP, i.e., some distance from the membrane
surface where the filaments are actively growing (see
below).
B) DEPOLYMERIZATION. At steady state, increasing the
rate of dissociation at the pointed end would not depolymerize actin filaments if the released monomers were able
to reassemble at the barbed end. However, if a barbed-end
capping protein (e.g., CapZ) blocks reassembly, then cofilin will depolymerize actin filaments.
C) NUCLEATION OF POLYMERIZATION. ADF/cofilin can nucleate the assembly of actin, and this function is likely to be
especially important in the presence of barbed-end blockers like CapZ. The ability to nucleate assembly is probably
pH dependent and may vary between different isoforms of
ADF/cofilin.
D) ADP-ACTIN. Binding of ADF/cofilin to actin is regulated, at least in part, by the state of the nucleotide bound
to actin. It binds to ADP-actin with about two orders of
magnitude greater affinity than to ATP-actin (Table 1) or
ADP-Pi-actin, and this is true for both the G- and F-actin
(42) at pH 7.8. Thus cofilin not only promotes the disassembly of ADP-actin monomers from filaments, but it also
binds to released ADP-actin monomers and inhibits the
exchange of their bound nucleotide (202). The off rate at
the pointed ends of filaments is probably the rate-limiting
step in the recycling of disassociated actin subunits (281).
The extent of depolymerization by ADF/cofilin depends
on several factors; for example, different isoforms may
have differing pH sensitivities (240, 183), and other ABPs
can influence the binding of cofilin to actin because they
share binding sites.
E) SEVERING. Marie-France Carlier et al. (42) argue that
the kinetic effects of ADF/cofilin are specific for the ends
of filaments and therefore it cannot sever. However, the
rate of filament fragmentation would not have to be very
great to increase the disassembly rate from ⬃125 s⫺1 in
vitro to the rate required to match the speed of motility
(⬃200 s⫺1). Evidence based on light microscopy (119)
suggests that filaments are rapidly depolymerized by ADF
at slightly alkaline pH. Also, there is a difference in severing activities of recombinant and native cofilin where
the native form is more active than the recombinant cofilin. Recombinant cofilins can vary in their severing activity and can give the impression that they have only
weak severing activity (132). Thus it remains unclear
whether ADF/cofilin can sever actin filaments like true
severing proteins or whether the highly cooperative bind-
ACTIN AND ACTIN BINDING PROTEINS
447
FIG. 7. Electron micrographs of negatively stained
(uranyl acetate) rabbit skeletal muscle actin. a: F-actin
only. b: F-actin ⫹ chick cofilin (molar ratio 1:1). c: F-actin
⫹ human gelsolin (1:0.2). (Micrographs provided by Roger
Craig.)
presence of two cofilins (yeast) per actin subunit. These
were seen by electron microscopy as two distinct sites in
image reconstructions. This raises the question of
whether one or two cofilins bind to actin monomers, a
feature which may depend on the isoform of cofilin. Thus
the question is whether cofilin accelerates treadmilling by
severing or depolymerizing F-actin. Current opinion
seems to favor the severing ability of cofilin.
4. Cellular concentrations
FIG. 8. A cartoon representation of F-actin in the absence (left) and
presence (right) of cofilin. Cofilins (red crescents) preferentially bind to
ADP-containing monomers in the filament and induce a small change in
the angular rotation of monomers in the filament with a consequent
slight increase in filament diameter and decrease in filament length.
Cofilin accelerates the off rate at the pointed ends of the filaments. Actin
monomers are represented as having a small and a large domain and are
color coded according to their bound nucleotide (brown for ATP, orange
for ADP-Pi, and yellow for ADP).
Physiol Rev • VOL
The cellular concentrations of ADF/cofilin are substantially lower than the concentration of actin, so it is
unlikely there is sufficient ADF/cofilin present in cells (20
␮M, Ref. 148) to bind to all polymerized and unpolymerized actins (65 ␮M; see Table 1), even taking into account
its low affinity for ATP- or ADP-Pi-actins. Furthermore,
the binding of ADF/cofilin to filaments is cooperative
(182). If there is not enough ADF/cofilin to act as a monomer-sequestering protein, this role probably belongs to
profilin and thymosin ␤4 (see below).
Relatively few ABPs bind to the pointed compared
with the barbed end of a filament. Cofilin can compete
with 1) spectrin that stabilizes short-actin oligomers (see
below) (254); 2) tropomodulin that caps the pointed end
of tropomyosin-coated actin filaments in muscle and nonmuscle cells (97); 3) DNase I, which binds very tightly to
the pointed end of actin monomers but only weakly to actin
filaments (see sect. IIIE); and 4) new pointed-end ABPs that
are likely to be identified from proteins that currently are
either orphans or have not yet been identified in gene
arrays and two-dimensional gel electrophoresis.
A number of isoforms of cofilin have been described,
and it is possible that not all members of the family
perform the same functions under the same conditions
(e.g., divalent cations and pH) (128). Some cofilins bind
tightly to actin monomers while others do not (Nosworthy, unpublished observations). Some cosediment with
F-actin while others have only a low affinity (120). Some
promote actin assembly while others rapidly depolymerize it (128). Human cofilin seems to be a better nucleator
of assembly whereas ADF is a better polymerizing agent,
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
ing of cofilin at substoichiometric ratios makes the filaments “brittle” at the points where the decorated and
undecorated regions meet (19). Recently, Ed Egelman
and colleagues (104) reported filament severing in the
448
DOS REMEDIOS ET AL.
5. Phosphorylation of ADF/cofilin
The ability of cofilin to bind G-actin is also regulated
by phosphorylation of Ser-3, which is conserved in most
members of the ADF/cofilin family. When ADF/cofilin is
phosphorylated, there is a sharp fall in its affinity for actin
(190). ADP exchange in G-actin is strongly inhibited by
cofilin, but once cofilin is phosphorylated, the released
ADP-actin monomer can exchange with cytoplasmic ATP,
and it is now ready for reincorporation at the barbed end
of a growing filament. Typically, this occurs at the interface of the microfilaments and the membrane at the leading edge of the moving cell. Thus cellular microfilament
turnover can potentially be regulated by cycles of phosphorylation and dephosphorylation. As we will see below,
profilin plays an important role here.
We know that in yeast, phosphorylation is not a
regulatory factor whereas it is in most vertebrates. An
important question is, How much of total cellular ADF/
cofilin is phosphorylated? For most cells, we simply do
not have this information, but in amoeba ⬃30% of actophorin is phosphorylated (29). On balance, it seems likely
that, since not all of ADF/cofilin is constitutively active,
there may be mechanisms other than phosphorylation
that control actin assembly-disassembly in vertebrates.
Phosphorylation of vertebrate cofilin is achieved by
LIM-kinase proteins 1 and 2 (LIMK-1 and LIMK-2). LIMK-1
is predominantly neuronal, whereas LIMK-2 is more widespread (26, 187). Although the signaling pathway for activation of the LIM-kinases is not yet fully understood, it is
known that LIMK-1 is under the control of the small
GTPase Rac (307), whereas LIMK-2 is regulated by the
GTPase cdc42 and rho (21) (see discussion below). LIMKs
are themselves regulated by phosphorylation of a Thr
residue (90). In Acanthamoeba, activation of LIM-kinase
by Pak1 (91) results in phosphorylation of Ser-1 (the
Acanthamoeba version of Ser-3) of actophorin (29). When
Physiol Rev • VOL
expressed in cultured cells, LIMK-1 induces actin reorganization and reverses cofilin-induced depolymerization
(307). Expression of inactive LIMK-1 results in the accumulation of F-actin filaments (7). Phosphorylation of cofilin appears to increase with pH-induced activation of
cofilin, consistent with a compensating homeostatic
mechanism (27).
Although NH2-terminal Ser residues (Ser-1, -3, -4,
and/or -6 in different species) are recognized as the site
for phosphorylation, it is not clear whether the resulting
inhibition of actin binding involves a steric or conformational change. Recently, Blanchoin et al. (29) examined
this question by crystallizing Acanthamoeba actophorin
(ADF/cofilin) in the phosphorylated state. Their structure
was essentially identical to unphosphorylated ADF/cofilin, and they concluded that inhibition of activity by LIMkinase was not due to a conformational change at the
ADF/cofilin-actin interface. Their reported steric blocking
of actin binding was consistent with molecular dynamics
predictions (Fig. 5B) and with site-directed mutant studies. Frustratingly, Ser-1 residue in these crystals could not
be resolved.
Until recently, the mechanism by which cofilin is
dephosphorylated was not well understood. Slingshot, a
protein phosphatase with F-actin binding ability, was
shown by Niwa et al. (204) to dephosphorylate cofilin in
cultured cells and in cell-free assays. In Drosophila, loss
of this enzyme results in a dramatic increase in cellular
levels of both F-actin and phosphorylated cofilin.
6. Inhibition by phosphatidylinositol 4,5-bisphosphate
The activity of ADF/cofilin can be regulated by the
membrane lipids phosphatidylinositol 4-phosphate (PIP)
and phosphatidylinositol 4,5-bisphosphate (PIP2) (309).
These bind to and inhibit the actin-binding domain of
ADF/cofilin at residues 104 –115 and the NH2-terminal
region (154) and suggest that transmembrane signaling by
PIP2 can regulate the function of ADF/cofilin. Little is
known of the molecular nature of this binding site.
7. pH effect
The ability of ADF/cofilin to assemble or disassemble
F-actin is pH dependent in vitro (311). Acidic conditions
(less than pH 6.8) enhance the ability of ADF/cofilin to
stabilize F-actin while at more alkaline pH (⬎7.3), cofilin
can rapidly depolymerize F-actin. Furthermore, the critical concentration of the cofilin-actin complex is known to
be pH dependent, being low (⬃2 ␮M) at pH 6.5 and
significantly higher (7 ␮M) at pH 8.2 (240). Conversely, it
has been suggested that the severing activity of cofilin is
pH independent (89). The physiological significance of
this pH dependency is that actin filaments may be stabilized in the vicinity of membrane Na⫹-H⫹ antiports (16). A
more recent report (27) focused on the role of pH in vivo.
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
and this is reflected in their respective Kd values (42, 240)
(Table 1).
If different isoforms of cofilin have different effects
on microfilament assembly, are they differentially expressed within a cell? Mouse myocyte cells lines express
significant amounts of ADF. Elevated expression of actin
causes a downregulation of ADF and decreases spreading
of these cells without changing the levels of cofilin (256).
Regulating ADF/cofilin expression is likely to be slow (on
a time scale of tens of minutes/hours) and, apart from
being energetically wasteful, it is unlikely to be an effective way of enabling microfilaments to respond quickly to
environmental stimuli. However, because most ABPs bind
with moderate to low affinity (in the micromolar range),
elevated levels of ADF/cofilin could, over time, subtly
shift the balance of ABPs and, by mass action, alter the
state of assembly of cytoskeletal microfilaments.
ACTIN AND ACTIN BINDING PROTEINS
449
Using pH-sensitive fluorescent probes introduced into
mouse fibroblasts (Swiss 3T3 cells), they (27) showed that
by lowering intracellular pH, ADF colocalized with actin
filaments and that when the pH was increased, ADF partitioned more with monomeric actin. In contrast, cofilin is
much less responsive to altered pH, and there are no
reports of structural changes induced in either ADF or
cofilin, despite the fact that its structure has been determined by both NMR spectroscopy (11) and crystallography (158).
8. Nuclear translocation sequence
FIG. 9. A ribbon diagram of the atomic (2.55-Å resolution) structure
of profilin (red) cocrystallized with monomeric actin where the orientation of actin is about the same as seen in Figure 5B (259).
B. Profilin Family
The profilin family of ABPs is found in eukaryotic
cells from Acanthamoeba through to human and in many
but not all species there are two or more profilin genes
(230). Profilins are small proteins with an approximate
molecular mass of 19 kDa (6). They are among the most
highly expressed (20 –100 ␮M, Ref. 37) of the cytoplasmic
proteins and are distributed throughout the cytoplasm.
Like cofilin and DNase I, profilin can be denatured in 8 M
urea and then renatured by dialysis (138), a property used
during its purification.
1. Atomic structure
X-ray diffraction and NMR atomic resolution structures are available for no less than six isoforms of profilin.
All are very similar. Profilin has also been cocrystallized
with actin in two different conformations, the so-called
“closed” and “open” states (60). It was on the basis of
these two structures that Schutt et al. (259) developed
their model of F-actin (discussed above). Within the profilins, the NH2-terminal sequence is conserved whereas
the COOH-terminal region of certain isoforms resembles
gelsolin (see below). The atomic structure of the profilin␥-actin complex is illustrated in Figure 9. Profilin binds to
subdomains 1 and 3 at loci that substantially overlap the
binding sites of gelsolin segment-1 (259). Its main crystal
contacts with actin are as follows: subdomain 1, 113, 354,
Physiol Rev • VOL
355, 361, 364, 369, 371, 373, 375; subdomain 3, 166, 167,
169, 171–173, 284, 286 –288, 290 (see Fig. 6).
2. Cellular functions
Profilin has several cellular functions. It is essentially
a high-affinity (Ka ⫽ 107 M⫺1) monomer-binding protein
(225). It is best known for its ability to promote the
exchange of nucleotide in actin monomers released from
filaments (109) and, because the high (millimolar) concentrations of Mg-ATP in the cytoplasm, it catalyzes the
exchange of ADP for ATP. It achieves this by binding to
subdomains 1 and 3 near the hinge between the two major
domains (see Figs. 6 and 9) and modulates the opening of
the nucleotide cleft. Profilin enhances filament turnover
in the presence of cofilin (85) because the two proteins
act at opposite ends of a filament, with profilin adding
ATP-actin to the barbed end and cofilin dissociating ADPactin from the pointed end (see Fig. 10). Profilin also
inhibits the hydrolysis of ATP bound to actin, thus maintaining actin monomers in a state where they retain a high
affinity for the growing barbed end of filaments (6).
While profilin acts as an effective buffer for high
monomer concentrations in cells, it can promote polymerization by transporting monomers to the fast-growing
barbed ends of filaments. Profilin binds to a site on actin
(see row 2 in Fig. 6) that is inaccessible in the Holmes
model of the filament (128) (see Fig. 4). Thus, after deliv-
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
All vertebrate ADF/cofilin (destrin) sequences contain a putative nuclear localization sequence (NLS) that
enables it to migrate from its normal location in the
cytoplasm to the nucleus under conditions of cellular
stress (203). Heat shock and other forms of stress probably affect a few residues immediately preceding the NLS,
which expose it for subsequent binding to a nuclear transport factor and thus passage through the nuclear pores
(32). Transport is an active process, and actin accompanies the cofilin into the nucleus, although the significance
of this is not clear.
450
DOS REMEDIOS ET AL.
1. Structure of gelsolin and gelsolin/F-actin
FIG. 10. A cartoon representating the cooperative roles of ADF/
cofilin (red crescents) and profilin (blue rhomboids) in regulating the
turnover of actin monomers (yellow) in a microfilament containing ATP
and ADP. Cofilin accelerates the dissociation of monomers from the
pointed ends of filaments. Phosphorylation of cofilin (dark red) dissociates it from ADP-actin and profilin promotes the exchange of ADP for
ATP that facilitates the addition of profilin-ATP-actin at the barbed end.
ering a monomer to the growing end of the filament, it
must either move to a new site that does not block
assembly or completely dissociate. This can occur even
when actin is complexed to thymosin (see below) because of the high exchange rates for both thymosin and
profilin (219). It is not clear why some cells employ thymosin to buffer actin monomers while others use profilin.
Profilin binds only very weakly to the pointed ends of
filaments. Dissociation of profilin from actin is stimulated
by PIP and PIP2 (108); thus profilin may play a role in
transmitting signals between the cell membrane and the
actin cytoskeleton. Profilin binds to Arp2 in the Arp2/3
complex (see sect. IIIG).
These functions are schematically illustrated in Figure 10. Here profilin is shown as blue rhomboids that bind
to the barbed end of F-actin. The blue arrows indicate the
cycle of profilin-actin interactions.
C. Gelsolin Superfamily
Gelsolin belongs to a superfamily of ABPs expressed
in all eukaryotes. The grouping includes (but is not limited to) gelsolin, villin, adseverin (also known as scinderin), CapG, flil, and severin (aka fragmin). All contain at
least one of the 120-amino acid structural repeats and
many have three or six gelsolin repeats (157). For example, villin (92.5 kDa) is a six-domain ABP that regulates
actin assembly in microvilli. It has a Ca2⫹-dependent NH2Physiol Rev • VOL
Gelsolin is an 80-kDa protein consisting of two tandem homologous halves (segments 1–3 and 4 – 6), each
containing threefold repeats (Fig. 11). Its structure in the
absence of Ca2⫹ has been determined at 2.5-Å resolution
(35). The isolated NH2-terminal half can bind to (cap) two
actin monomers and can sever F-actin without the need
for free Ca2⫹. In contrast, the COOH-terminal half binds
a single actin and is Ca2⫹ dependent. The molecule is
compact and globular having dimensions of ⬃85 ⫻ 36
⫻ 55 Å. Segment 1 cocrystallized with monomeric actin is
shown in Figure 12A. It has two bound Ca2⫹ (186). Gelsolin segments 4 – 6 also bind to monomeric actin, and
their relationship to the actin monomer is illustrated in
Figure 12B.
McGough et al. (181) had earlier suggested a mechanism in which segments 2 and 3 bind to the actin filament
while segment 1 wedges itself between two adjacent actin
monomers along the longitudinal axis. This step requires
a rearrangement of the interface between segments 1 and
2 to lengthen the linker between these segments. Segments 4 – 6 reach across the filament to bind a monomer
in the other strand. This step would also require extension
of the convoluted linker between segments 3 and 4.
The following year, on the basis of crystallographic
data, Robinson et al. (243) concluded that Ca2⫹ binding
induces a conformational rearrangement in which segment 6 is flipped over and translated by ⬃40 Å relative to
segments 4 and 5. The structural reorganization tears
apart the continuous ␤-sheet core of segments 4 and 6.
This exposes the actin-binding site on segment 4, enabling
severing and capping of actin filaments to proceed (Fig.
13). Severing of F-actin occurs only when sufficient actinactin bonds are broken (36). The severing of F-actin by
gelsolin is illustrated in Figure 7C where only short segments of actin filaments are observed in the presence of
recombinant gelsolin (electron micrograph courtesy of
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
terminal half (180), whereas CapG (aka gCap39) is a
smaller, three-domain gelsolin analog that cannot sever
F-actin but responds to Ca2⫹ and PIP2 under conditions
where gelsolin is ineffective (312).
Unlike ADF/coflin and profilin, an isoform of intracellular gelsolin can circulate in plasma where it severs
and caps actin filaments released into the circulation (e.g.,
following cell death). The resulting actin monomers and
oligomers are then strongly sequestered by vitamin Dbinding protein (DBP) and ultimately removed from the
circulation in the liver (124). Plasma gelsolin is slightly
larger (83 kDa) than cytosolic gelsolin and is derived by
alternative splicing of a single gene resulting in a short
peptide appended to its NH2 terminus (155). Otherwise,
the sequences of plasma and intracellular gelsolins are
identical.
451
ACTIN AND ACTIN BINDING PROTEINS
Roger Craig, University of Massachusetts Medical Center,
Worcester, MA). Here the molar ratio of gelsolin (segments 1–3) to actin is 0.05:1.
The crystal structure of the F-actin binding domain of
severin, the gelsolin homolog from Dictyostelium discoideum, has recently been described at high (1.75 Å) resolution (243). The structure reveals an ␣-helix-␤-sheet
sandwich that contains two cation-binding sites and Factin binding residues in the equivalent region of gelsolin
segment 2 (234). This segment also mediates the binding
of gelsolin to tropomyosin (172).
2. Functional properties in vitro
A) ACTIN SEVERING.
Gelsolin certainly lives up to its
name. It rapidly dissolves actin gels in vitro by severing,
capping, and nucleating growth. Like cofilin, it can dissociate phalloidin from F-actin (262). These functions are
achieved by attacking filaments from their barbed ends.
Gelsolin is an important ABP because of its high affinity
for actin filaments (Kd 50 nM) (229).
Segments 1–3 and 4 – 6 of intact gelsolin both play an
active role in the severing of F-actin. Binding of gelsolin
FIG. 12. A: crystal ribbon structure of
gelsolin segment 1 (red) and actin. B:
gelsolin segments 4, 5, and 6 and actin.
These figures are shown in the same orientation as Figs. 5B and 9A. Colors are
those used in Fig. 11. (Figures kindly provided by Bob Robinson.)
Physiol Rev • VOL
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
FIG. 11. A ribbon diagram of the
atomic structure of inactive gelsolin
showing segment 1 (red) (containing the
NH2 terminus), segments 2 and 3 (green,
yellow), and segments 4 – 6 (magenta,
green, gold). (Figure kindly provided by
Bob Robinson.)
452
DOS REMEDIOS ET AL.
initiates the severing, but while binding to filaments is
rapid, its severing action is slow. Segments 4 – 6 confer a
Ca2⫹ sensitivity in the micromolar range, and binding
involves a cooperative intramolecular structural rearrangement within gelsolin (229). This can be visualized by
comparing the arrangement of segments 4 – 6 of inactive
gelsolin (Fig. 11) with the active form shown in Figure
12B. A cartoon of active gelsolin segments on F-actin is
shown in Figure 13, which was redrawn from a figure
kindly provided by Bob Robinson (University of Uppsala,
Uppsala, Sweden). Severing is effective even in the presence of tropomyosin and has been used by several investigators to solubilize actin filaments from skeletal muscle
sarcomeres.
B) CAPPING. Not all authors agree that gelsolin is principally a severing protein. Gremm and Wegner (112) argued that at micromolar and submicromolar free Ca2⫹
concentrations, gelsolin acts more as a Ca2⫹-regulated
capping protein. They observed that depolymerization
was mainly caused by dissociation of actin subunits from
the pointed ends of actin filaments and that inhibition of
polymerization was achieved by gelsolin at barbed ends.
The rate of capping at the barbed end was directly proportional to the free Ca2⫹ concentration (112).
After severing, gelsolin remains attached to the
barbed end of the filament as a cap, thereby preventing
the reannealing of actin fragments. Capping prevents
rapid growth at the barbed ends while disassembly proceeds unchecked at the pointed ends, resulting in the
rapid disassembly until an equilibrium is established between capped filaments and free gelsolin (239). Dissociation (uncapping) of gelsolin from these filaments generates numerous polymerization-competent ends that seed
the rapid assembly of new filaments. Thus the cytoskeleton can be rapidly rebuilt in a new direction and cell
movement can be redirected (66).
2⫹
C) CALCIUM DEPENDENCE. Ca
is an important cellular
Physiol Rev • VOL
3. Comparison with ADF/cofilin
ADF/cofilin competes with gelsolin segments 2 and 3
for binding to actin filaments. This suggests they share a
common structural topology for binding to actin (290),
and this was subsequently confirmed by molecular dynamics simulation (304). Both proteins alter the conformation of the actin filament, dissociate phalloidin, sever,
and cap F-actin. They both bind to ADP-G-actin with
higher affinity than to ATP-G-actin, and the binding of
both to actin is inhibited by PIP2 (275). Recent expression
studies suggest both proteins play an important role in
regulating actin dynamics in vivo (268). However, despite
the more powerful severing action of gelsolin, ADF/cofilin
is currently favored as the major regulatory mechanism of
actin cytoskeleton reorganization (16).
4. Knock-out and overexpression
Transgenic gelsolin-null mice have normal embryonic development and longevity but suffer subtle changes
suggesting that it is needed for rapid changes in cell
motility in hemostasis (reduced platelet function), inflammation (delayed neutrophil migration), and wound healing (fibroblasts move more slowly) (303). In these null
mice, gelsolin is an essential effector of rac-mediated
actin dynamics, acting downstream of rac recruitment to
the membrane (10). Gelsolin also appears to play a role in
the initiation of filopodial retraction. An absence of gelsolin delays its retraction and produces “studded” filopodia along growing neurites (169).
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
FIG. 13. A cartoon representation of three monomers in a filament
showing a possible rearrangement of gelsolin segments involved in
cleavage of the actin filament.
trigger for events that involve the cytoskeleton, and gelsolin is also the only known Ca2⫹-dependent severing
protein with a Kd for Ca2⫹ in the micromolar range. The
six motifs contain two sites that bind to G-actin and one
that binds to filaments. The Ca2⫹-dependent monomerbinding site in the COOH-terminal half plays a critical role
both in the cooperative binding to actin by gelsolin and in
its nucleating activity. Actin binding was localized to segment 4 by expressing segments 4, 4 –5, 5, and 5– 6 in
Escherichia coli (232). The nonbinding segments play an
important role in the Ca2⫹ regulation of actin binding and
other activities of gelsolin (232). The segmental arrangements are shown in Figure 10.
An analysis of the gelsolin construct lacking the
COOH-terminal helix indicated the importance of this
part of molecule for Ca2⫹ regulation and supported the
so-called “tail helix latch” mechanism of activation by
Ca2⫹ (170). This latch is primarily responsible for transmitting Ca2⫹ binding from the COOH-terminal half to the
NH2-terminal half of gelsolin. Occupancy of this site
primes gelsolin for severing, but it is unable to sever
because the tail latch is still engaged. Unlatching the tail
helix by micromolar Ca2⫹ releases the final constraint to
initiate the severing cascade (163).
ACTIN AND ACTIN BINDING PROTEINS
453
D. Thymosins
Thymosin ␤4 (T␤4) is a small (molecular mass ⬃5
kDa) protein that belongs to a family of highly conserved
small proteins originally isolated from the thymus gland
but now known to be more widely distributed (93). It
contains 43 residues, ⬃50% of which are charged. The
amino acid composition of T␤4 is remarkably deficient in
hydrophobic residues (168), which suggests that T␤4
probably has very little defined structure in solution.
1. Structure
Until very recently, the structure of T␤4 was based
on a model constructed by Dan Safer et al. (251) who
docked it to the crystal structure of monomeric actin
solved in the presence of DNase I (Fig. 14). These authors
regard this protein as having an essentially unfolded conformation in solution and, although there may be an increase in its ␣-helical content, it remains unfolded once it
binds to actin (251). Based on a range of data (250), it has
been suggested that T␤4 drapes itself around nearly half
the perimeter of the actin. The molecular contact points
between T␤4 and the actin monomer investigated using
zero-length cross-linkers show that Lys-3 of T␤4 is located
close to Glu-167 (in subdomain 3 near the hinge), and that
Lys-18 of T␤4 cross-links to one of the four NH2-terminal
acidic residues of actin (Asp-1 through Glu-4). These two
contacts flank the ␣-helical region of T␤4 at the barbed
end of the monomer. The COOH-terminal region of T␤4
(Lys-38) was cross-linked to Gln-41 at the top of actin
subdomain 2, a known DNase I binding site.
The NMR structure of free T␤4 in solution (75) exhibits a range of conformational preferences. In water it
appears to lack a folded conformation, but a preferential
␣-helical conformation can be observed in solution at
different temperatures and pH. Residues 5–16 putatively
Physiol Rev • VOL
FIG. 14. A modeled complex of thymosin ␤4 and an actin monomer
(78) using an orientation of the actin monomer comparable to Figs. 5A,
9A, and 12. No crystal structure is available for this complex.
bind to the NH2-terminal region of actin and have characteristic temperature-dependent conformations (75).
These data indicate that many parts of T␤4 are not in tight
contact with actin. Similarly, circular dichroism spectra
indicate that free T␤4 is predominantly unstructured but
increases its ␣-helical content when it binds to actin. At
least one research group has made progress with a crystal
structure of T␤4. De La Cruz et al. (78) recently concluded
that T␤4 changes the conformation and structural dynamics (the so-called “breathing”) of actin monomers. The
conformational change may reflect the unique ability of
T␤4 to sequester actin monomers and inhibit nucleotide
exchange (compared with profilin).
2. Cellular distribution and functions
␤-Thymosins are widely distributed in nature. Translational levels can vary from tissue to tissue (262). In
chick brain, concentrations are an order of magnitude
higher than profilin and cofilin. It is abundant in neural
tissue as well as in circulating cells such as platelets,
leukocytes, and macrophages. T␤4 binds stoichiometrically to monomeric actin and inhibits polymerization. In
the developing chick brain, the ratio of ADF to cofilin to
T␤4 increases between embryonic days 13 and 17. Because no other thymosin isoforms were detected in brain
extracts, it is probable that it is the major actin-sequestering protein in this tissue (84). T␤4 mRNA is mainly
expressed in neurons of the hippocampus, neocortex, and
the amygdala, as well as in oligodendrocytes and probably
has a function in the production and remodeling of neu-
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
Overexpression of gelsolin in cultured mouse fibroblasts enhances their migration rate by ⬃125% via gelsolin-promoted actin assembly and disassembly (72). Control of the rate of cell migration is closely linked to the
rates of actin filament severing. In fibroblasts that overexpress gelsolin, there is a direct correlation between
movement velocity and actin turnover rates. This demonstrates that gelsolin is an important control mechanism
for actin cycling in cells (184). Overexpression of gelsolin
in cells also inhibits phospholipase C-␥ activity by competing with it for available PIP2. A rise in free Ca2⫹
increases the affinity of gelsolin for PIP2, and as the level
and availability of PIP2 changes during signaling, crosstalk between PIP2-regulated proteins (e.g., gelsolin and
phospholipase C-␥) provides a mechanism for positive or
negative regulation of the signal transduction cascade
(274).
454
DOS REMEDIOS ET AL.
Physiol Rev • VOL
trostatic contact site with actin. A nonconserved NH2terminal segment preceding the motif exerts an inhibitory
activity on actin polymerization probably by steric hindrance. Substitution of a His or Lys for a Glu at the third
position in the motif converts the peptide from an inhibitor of polymerization into a stimulator. Thus subtle sequence changes in ABPs can have remarkable effects on
their functions.
3. Molecular interaction with actin
Binding of T␤4 to actin is coupled to the dissociation
of bound water molecules, which is greater for CaATPactin than MgATP-actin monomers. The COOH terminus
of T␤4 makes contact with actin at His-40 near the mouth
of the nucleotide cleft and binding slows the 3H exchange
rate, suggesting there are changes in conformation of
actin monomers when it binds. This may reflect the ability
of T␤4 to both sequester actin monomers and inhibit
nucleotide exchange (78).
Gelsolin and T␤4 compete for binding to actin, indicating that T␤4 binds to a site close or identical to the
gelsolin segment 1-binding site (13). Figure 6 shows that
the binding loci for gelsolin and T␤4 do not precisely
coincide but that there must be sufficient overlap to explain the competition. If T␤4 is to be a major cellular
actin-sequestering factor, it should bind, among other
things, with a slightly higher affinity (0.7 ␮M) than gelsolin
(⬃1 ␮M) (262), thus subtly shifting the assembly/disassembly equilibrium of actin toward the monomeric state.
The interactions of these two proteins with actin are
complex and require the participation of several complementary peptide sequences. A common motif (I,V)EKFD
in these two proteins exerts a major inhibitory effect on
salt-induced actin polymerization (96).
To make matters more complex, T␤4 and profilin
(and presumably gelsolin) compete with WASP (an activator of Arp2/3) for binding to actin monomers and presumably have overlapping binding sites (see sect. IIIG).
DNase I and T␤4 bind to sites on actin that partially
overlap. The apparent Kd of the actin-T␤4 complex generated from a preformed actin-DNase I complex is 160
␮M. A fivefold excess of DNase I over the preformed
actin-T␤4 complex is necessary to observe a comparable
Kd (129). Thus DNase I can displace T␤4 from the monomer (and probably vice versa).
Chemical cross-linking experiments also indicate
that profilin competes with T␤4 for actin binding. In the
presence of Mg2⫹, profilin promotes assembly of T␤4actin but in the presence of Ca2⫹, profilin was unable to
initiate this polymerization (12). Phalloidin, rabbit skeletal muscle myosin S1, and chicken intestinal myosin I all
polymerize actin from the T␤4-actin complex (15). Thus,
despite the very high affinity of DNase I for actin, its
binding can be inhibited by a lower affinity T␤4. The
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
ronal processes (48). T␤4 is upregulated in the pyramidal
neurons of the hippocampus where its function may be
related to restoration of neurite circuits after focal ischemic damage (292).
T␤4 is present in some cells at concentrations as high
as 0.5 mM (249). High expression levels have been observed in young embryonic muscles but not in adult skeletal muscles. Consequently, it seems that the G-actin pool
in young embryonic skeletal muscle is mainly regulated
by T␤4 and profilin (206). However, T␤4 may become less
important as muscle develops (199). In contrast, there is
evidence that it plays a role in angiogenesis in coronary
smooth muscle, acting as a chemoattractant by stimulating the migration of endothelial cells in vivo (174).
In quiescent NIH 3T3 cells, T␤4 was undetectable,
but after restoring serum to the culture medium where
cells were induced to proliferate, there was a pronounced
increase in T␤4 mRNA (316). Overexpression in these
cells increases F-actin levels while maintaining a constant
level of total actin (273). In contrast, others reported that
a twofold increase in T␤4 in these same cells doubled the
total actin expression but maintained the ratio of G- to
F-actin (110). Unexpectedly, other cytoskeletal proteins
such myosin IIA, ␣-actinin, and tropomyosin also increased nearly twofold (273). The T␤4 cell lines spread
more fully and adhered to the dish more strongly than
vector controls, suggesting a coordinated regulation of
the cytoskeleton by T␤4. In addition, transcriptional levels of several related cytoskeletal proteins also increased.
These observations suggest there is a coordinated cytoskeletal expression of this protein (110).
Like several other ABPs, thymosins bind to and coordinately control the availability of actin monomers for
incorporation into filaments. T␤4 is a contender with
profilin for the title of the universal actin-monomer binding protein. It has a putative actin-binding motif
(LKHAET) (287) found in other ABPs including actobindin and myosin. Nachmias (196), and more recently
Chen et al. (54), reviewed the discovery and primary
structure of T␤4.
Acting alone, T␤4 inhibits polymerization, and like
ADF/cofilin it also inhibits the exchange of the actinbound nucleotide (107). This contrasts with the action of
profilin which, acting alone, facilitates the exchange of
ADP for ATP and promotes actin assembly (109). Thus,
while it acts as a passive buffer of actin monomers, preventing their polymerization, T␤4 can cooperate with profilin to regulate actin assembly (273). However, T␤4 is not
a simple G-actin-sequestering protein because, at least at
high concentrations, it also interacts with F-actin (40).
The actin-binding sites of T␤4 resemble that of another small ABP, actobindin, which also inhibits polymerization. Their binding loci on monomeric actin have been
localized using peptide mimetics (286). Both have an
NH2-terminal hexapeptide motif forming the major elec-
ACTIN AND ACTIN BINDING PROTEINS
diated by G-actin sequestering and involve interaction
with DNase I (114).
NIH 3T3 cells that overexpress (x3) T␤10 have been
shown to have 23–33% more polymerized actin than control cells, and their microfilaments appear thicker. There
is no change in total actin, profilin, or cofilin/ADF content.
These cells are more motile, spread faster, and have
higher chemotactic and wound healing activity (273). Immunofluorescent labeling of peritoneal macrophages
showed that both T␤4 and T␤10 were uniformly distributed within the cytoplasm. Overexpression in fibroblasts
induces extensive loss of stress fibers, indicating that
T␤4 and T␤10 are functionally similar and that both are
effective regulators of actin filament dynamics in living
cells (313).
Both T␤4 and T␤10 are elevated in a human prostate
cancer cell line and correlate positively with the grade of
tumor. Transfection with antisense T␤15 constructs into
rat prostatic carcinoma cells demonstrates that the T␤4
peptide positively regulates cell motility. Increased motility is a critical component of the metastatic pathway (22).
T␤15 is also upregulated in nonmetastatic breast cancer
and even in ductal carcinoma (compared with benign
breast tissue). Consequently, it might represent a potential early marker for breast malignancy (106).
T␤9 and T␤met9 (the minor variants of T␤4) inhibit
polymerization of ATP-actin with identical Kd values (0.7–
0.8 ␮M). Like T␤4, these thymosins bind to ADP-G-actin
with a 100-fold lower affinity than to ATP-G-actin (134).
E. DNase I
4. Other thymosins
Thymosin isoforms are distributed throughout the
vertebrate phyla (247). While T␤4 is the predominant
expressed isoform in mammalian cells, many species produce at least two T␤ isoforms, in some cases in the same
cell (49). Differences in the distributions of the mRNAs
encoding T␤10 and T␤4 suggest there are different roles
for these peptides during mouse early embryogenesis
(49). The levels of T␤10 mRNA strongly increase during
early postimplantation mouse embryogenesis, suggesting
that T␤10 plays an important role in early development.
Changes in T␤10 and T␤4 mRNA expression are also
important in the control of actin dynamics in developed
neurons and glia since T␤10 mRNA is present in different
regions of the rat forebrain, including hippocampus, neocortex, and several brain nuclei (48). T␤10 is also downregulated in failing human hearts with dilated cardiomyopathy (Tsubakihara, unpublished data).
The underlying mechanism for these cellular changes
remains to be elucidated. However, plasmid-driven overexpression of the T␤10 gene is known to increase the
susceptibility of apoptosis in transfected fibroblasts (115).
These proapoptotic properties of thymosins may be mePhysiol Rev • VOL
DNase I is a secretory glycoprotein with an endonuclease activity that cleaves double-stranded DNA to yield
5⬘-phosphorylated polynucleotides. It has a requirement
for both Ca2⫹ and Mg2⫹. Bovine pancreatic DNase I is the
best characterized of a group of DNases. It has a molecular mass of 31 kDa and an optimum pH of 7.8 (152). It
was first purified from pancreas and parotid gland that
contained four DNases (A, B, C, and D), differing according to their carbohydrate side chain and polypeptide components. DNase I is glycosylated at Asn-18 (189). These
factors, together with the presence of sialic acid, result in
a charge heterogeneity that can be detected by isoelectric
focusing (162).
1. Atomic structure
The structure of DNase I was solved crystallographically (272) at 2-Å resolution and was found to have two
Ca2⫹ binding sites. DNase I has a high affinity for actin
(Kd ⫽ 5 ⫻ 108 M⫺1) (177). The crystal structure of the
actin-DNase I complex (with actin in the ATP form) has
been determined at 2.8-Å resolution (137). Most ABPs
interact with actin subdomain I, whereas DNase I binds to
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
explanation for this may be an allosteric conformational
change in the actin monomer as reported by us recently
(81) for DNase I and cofilin.
Competitive binding data suggest that T␤4 also
makes contact with the myosin binding site as well as
with the bottom of subdomain 3 where it overlaps the
binding sites of several ABPs including DBP (Fig. 6, row
3) at the barbed end of the monomer. Therefore, the
distance between the pointed and barbed ends requires
the COOH-terminal half of T␤4 to be in a predominantly
extended conformation (247). Point mutations revealed
that the middle section of T␤4 (residues 17–22) also interacts with actin (267).
Although T␤4 is widely recognized as a monomer
binding protein, at least three reports have demonstrated
its ability to bind to F-actin (14, 40, 273). Addition of 200
␮M T␤4 during actin assembly decreases the critical concentration of free actin. This is consistent with the in vivo
observation that overexpression of T␤4 increases stress
fibers and decreases the amount of monomeric actin
(273). A chemically cross-linked binary complex of T␤4
actin is incorporated into F-actin only when both phalloidin and gelsolin are present (14). T␤4 increases the longpitch helical repeat (from 35 to 40.5 nm determined from
3-dimensional helical reconstructions) of F-actin. Like its
interaction with actin monomers, T␤4 does not “dock”
neatly onto the F-actin model, particularly at the NH2terminal region of actin subdomain 3. This difference may
well be due to conformational changes in both proteins as
suggested recently by at least two authors (14, 78).
455
456
DOS REMEDIOS ET AL.
F. Capping Proteins
15. This ribbon structure of DNase I (red) and actin is derived
from the 1990 2.8-Å crystal structure of the complex (137). The orientation is essentially the same as Figs. 5A, 9A, 12, and 14. Note the
contacts for DNase I are located in subdomain 2.
FIG.
the part corresponding to the pointed end of the actin
(298). The atomic structure of rabbit skeletal muscle actin
complexed with bovine DNase I is shown in Figure 15.
2. Cellular functions
DNase I is an enzyme secreted by exocrine glands
such as the pancreas and the parotid into the alimentary
tract. However, it has a much wider tissue distribution.
This suggests it may play roles other than to hydrolyze
DNA. DNase I, as well as other DNases, has been implicated in apoptosis (223) where it hydrolyzes DNA into
short fragments (DNA laddering).
Lazarides and Lindberg (159) discovered that actin
was a naturally occurring inhibitor of DNase I, and they
were the first to suggest that DNase I may be a cytoskeletal protein. They proposed three possible biological roles
for this interaction: 1) actin controls the nucleolytic activity of this enzyme during the cell cycle, 2) the complex
has a specific function in DNA metabolism, and 3) the
primary function of DNase I is related to the formation
and function of actin filaments rather than to the degradation of DNA. The fact that phosphoinositides can dissociate the binary actin-DNase I complex (309) also suggests that this interaction may have biological relevance.
DNase I has been useful tool in measuring G-actin
Physiol Rev • VOL
Monomers in actin filaments “age” as they progress
from the assembling (barbed) end to the disassembling
(pointed) end of a filament. This aging process is driven
by the chemical energy of Mg-ATP. Free monomers predominantly contain bound ATP that is hydrolyzed to
ADP-Pi only after incorporation at the barbed end, with
the subsequent release of Pi. At the barbed end, the on
rate for ATP monomers (5 ␮M/s) exceeds the off rate (1
␮M/s) while for ADP monomers the corresponding on
rate (1 ␮M/s) is slower than the off rate (6 ␮M/s). Thus,
while monomers at the barbed end nearly all contain ATP,
monomers at the pointed end mostly contain ADP. At the
pointed end, the on rate for ATP monomers (0.5 ␮M/s) is
slower than its off rate (1 ␮M/s) while the off rate for ADP
(0.2 ␮M/s) is faster than its on rate (0.1 ␮M/s) (149).
Because the rates of monomer exchange are 20 times
faster at the barbed than at the pointed end, there is a high
probability that an ATP monomer that adds to this end
will be hydrolyzed before another exchange occurs.
Clearly, capping proteins at these ends will alter monomer kinetics depending on their binding affinity and rate
of exchange.
Many, perhaps most, cells control the length and
number distribution of thin filaments under conditions
where total actin concentration remains approximately
constant. This assembly and disassembly of actin requires
a dynamic equilibrium between actin monomers and the
two, nonequivalent ends of actin filaments. Proteins that
cap or prevent this exchange may variously nucleate,
promote, stabilize, or inhibit assembly. They are therefore
in a strong position to regulate actin polymerization. This
simple concept is illustrated in Figure 16. Here we focus
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
levels in cells (71), although its function in cytoskeletal
dynamics is yet to be elucidated. However, the tight binding and specificity of DNase I for actin and its wide
distribution suggests it has a biological role. It binds very
tightly to G-actin (Kd in the nM range) (266) and essentially removes all free actin monomers from solution. It
binds much more weakly to F-actin (100 ␮M) where it is
a capping protein that increases the rate of dissociation
from the pointed ends of filaments (298).
We recently examined the possible cellular codistribution of DNase I and cofilin in cultured mammalian cells.
Using fluorescent-labeled antibodies to cofilin and DNase
I, we found the cytoplasmic distributions of these two
proteins overlap but are not identical (56). Like T␤4,
DNase I may act as a stabilizer of actin monomers by
effectively removing them from the available pool of
monomers available for assembly. In section IV we raise
the possibility that it may also be a natural modulator of
the action of cofilin on the cytoskeleton.
ACTIN AND ACTIN BINDING PROTEINS
on the most abundant barbed-end capping protein CapZ
and the predominant pointed-end capping protein tropomodulin.
1. CapZ, a barbed-end capping protein
Several different proteins can bind to the barbed end
of filaments. Some of them like gelsolin (and its close
relatives adseverin and fragmin) (254) and profilin have
already been discussed above in a different context. Villin,
CapG, tensin, and fragmin have been reviewed elsewhere (254).
CapZ is present in all eukaryotic cells (65). In skeletal
muscle it is localized at the Z-disk where it is probably
held onto the Z-filaments by a unique protein (see review
in Ref. 165). It is usually prepared from chicken or rabbit
skeletal muscle but, unless G-actin is twice purified by gel
filtration, traces of CapZ remain in the preparation and
affect actin assembly characteristics (52). The difficulty in
preparing actin free of CapZ can be appreciated when one
realizes that one heterodimer of CapZ can cap a filament
containing a thousand or more actin monomers (265).
A) FUNCTIONS OF CAPZ. The reported biological roles of
CapZ are as follows: 1) nucleation of actin assembly, 2)
capture of preexisting filaments, 3) regulation of actin
Physiol Rev • VOL
assembly at the barbed ends of filaments, 4) elimination
of annealing at the barbed ends of filaments (265), and 5)
correct assembly of filaments at the Z-disk (254). CapZ
does not affect the rate of fragmentation of actin filaments
and does not bind to their pointed ends (53). Figure 16
shows CapZ at the barbed end of a filament.
CapZ is a heterodimer comprising an ␣-subunit
(there are at least 2 isoforms in eukaryotic cells) and a
␤-subunit (2 or 3 isoforms), and both subunits are required for effective barbed-end capping of filaments (53).
The ␣1- and ␣2-isoforms result from alternative gene expression (117). Capping proteins containing the ␣1-isoform display fourfold higher affinity for actin than capping
protein containing the ␣2-isoform, although the physiological significance of this remains unclear. The ␤1- and
␤2-isoforms result from alternative splicing of a single
mRNA transcript (255), and although both isoforms demonstrate similar affinity for actin, they are differentially
expressed and distributed. The ␤1-isoform is predominantly expressed in muscle cells and is localized to the
Z-line in striated muscle, whereas the ␤2 is located under
the sarcolemma (51). Consequently, capping protein containing the ␤1-isoform is often referred to as CapZ and is
analogous to ␤-actinin (179). The ␤2-isoform is predominantly expressed in nonmuscle cells where it is localized
to the cell periphery (255).
B) STRUCTURE OF CAPZ. CapZ was recently crystallized
by Yuchiro Maeda and colleagues, and a preliminary report was presented at the SPring8 actin conference held
in Japan in November 2001. Details of this structure are
not yet available but are likely to be published later in
2003. Despite the fact there are no cocrystals of CapZactin, considerable detail is available on the molecular
contacts between these proteins. The contact sites on
actin are illustrated in Figure 6, row 4. Deletion of the
COOH terminus of the ␤-subunit of CapZ eliminates its
capping activity as does the binding of a monoclonal
antibody directed against the COOH-terminal 12 residues
(130). Later, this same group showed that 31– 40%, respectively, of the ␣- and ␤-subunits are not required for strong
binding to actin, suggesting that these regions may be
important for other functions (53).
As its name suggests, capping protein binds tightly to
the barbed end of actin filaments. In vitro, this capping
prevents the exchange of actin subunits from the barbed
end. Furthermore, capping protein facilitates actin polymerization by binding to and stabilizing monomers or
oligomers of actin forming nuclei for elongation of filaments (51).
C) REGULATION OF CAPZ ACTIVITY. In vivo, the capping of
actin filaments is regulated by second messengers, PIP
and PIP2. This is also true for other ABPs including cofilin,
destrin and DNase I (309), profilin (108), and gelsolin
(285) (see Table 1). Upon signal transduction, PIP and
PIP2 promote removal of capping protein from actin fila-
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
FIG. 16. A cartoon representation of the interaction of a barbed-end
(CapZ, purple) and a pointed-end capping protein, tropomodulin
(brown) (Tmod). One CapZ binds to two monomers at the barbed end of
an actin (yellow) filament. The NH2 terminus of Tmod binds tightly to
NH2 terminus of tropomyosin (green) at the pointed end of filaments and
also to one (or two) monomers. The on rate and off rates at the filament
ends are indicated by lengths of the arrows. Two views of the Tmod
molecule [originally published in Krieger et al. (153)], modeled from
X-ray scattering data, are shown at the top of this figure.
457
458
DOS REMEDIOS ET AL.
ments, resulting in localized pockets containing a dramatic increase in the number of free barbed ends. This
allows actin polymerization to proceed in localized regions of the cell. Capping protein is involved in a variety
of biological roles including the activation of platelets and
maintaining the proper assembly of actin filaments at the
Z-line of sarcomere (254).
i.e., at their respective ends. Blocking barbed ends with
cytochalasin D had no effect on filament length, whereas
overexpression of Tmod reduced actin filament length,
possibly because Tmod promotes the conversion of ATP
or ADP-Pi monomers to ADP with a consequent raising of
the critical concentration from 0.1 to 1.0 ␮M (241).
G. The Arp2/3 Complex
2. Tropomodulin, a pointed-end capping protein
Physiol Rev • VOL
Arp2 and Arp3 belong to a recently discovered family
of proteins that are related, at least in a limited way, to the
sequence and structure of actin. Their cellular concentrations are 2 and 5 ␮M, respectively (171). These two actinlike proteins are assembled into the Arp2/3 complex,
which is known to occur in a wide range of organisms
from yeast to mammals. The seven-subunit protein complex comprises Arp2 and Arp3 and five smaller (Arc)
proteins that do not appear to be related to any proteins
currently in the databases. Both Arp2 and Arp3 sequences
have nucleotide-binding and divalent cation-binding regions, but otherwise they are divergent from each other
and from actin (143). Arp2/3 is associated with dynamic
cortical regions of mammalian cells that are rich in cytoskeletal actin (69). It colocalizes with the actin “comet
tails” of Listeria and Shigella where actin polymerization
drives filament assembly and propels these intracellular
bacterial pathogens through the cytoplasm of the host cell
at speeds of 10 –90 ␮m/min (302).
1. Structure
The structure of the Arp2/3 complex at 2.0-Å resolution was only recently described (244). It is remarkable
that a complex this big crystallized at all. Proteins were
extracted from bovine thymus gland, and the isolated
complex was in an inactive state when it was crystallized.
Conversion to an active form is a matter of speculation
but will probably involve a rearrangement of several subunits to allow the Arp2 and Arp3 to be repositioned so
that they can act as nucleation site for the assembly of
new filaments. This rearrangement will also be needed if
the Arp2/3 complex is to “capture” preexisting filaments
and incorporate them with the correct orientation toward
the advancing cell membrane. The structure is now deposited in the Protein Data Bank (PDB accession number
1k8k).
Briefly, the structure has the following features. The
overall shape of the complex is a flat ellipsoid with dimensions of 150 ⫻ 140 ⫻ 70 –100 Å. The structures of Arp2
and Arp3 strongly resemble the actin monomer. Arp2 has
a strong resemblance to subdomains 1 and 2 of actin
except that the interdomain cleft is more open (by ⬃18°)
and contains no bound ATP. It also has a profilin binding
domain (143). Arp3 contains large inserts into subdomains 2, 3, and 4 and has 42 residues more than actin. The
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
Unlike barbed-end capping proteins, pointed-end
capping proteins are something of a rarity. The name
tropomodulin (Tmod) is apt because it binds strongly to
actin in the presence of tropomyosin. With the exception
of Arp2/3 (see sect. IIIG), no other protein binds to the
pointed end of F-actin, and no other protein inhibits filament elongation at this end. Although Tmod binds to
F-actin (it does not bind monomeric actin). Its affinity is
weak (Kd ⬃0.1 ␮M) compared with its affinity for tropomyosin-decorated actin filaments that is about three orders of magnitude stronger (Kd is submicromolar) (297).
For this reason, some authors originally referred to it as a
tropomyosin binding protein (295).
Human Tmod has been cloned and is expressed in a
wide range of tissues with high levels in skeletal and
cardiac muscle (276). It contains 359 amino acids and has
a molecular mass of 40,476 Da determined by mass spectrometry. Overexpression of Tmod in cardiomyocytes
produces short, disorganized, and destabilized thin filaments (277). A Tmod overexpressing transgenic mouse is
now available (278). A related, somewhat large (64 kDa)
protein, leiomodin, is highly expressed in smooth as well
as cardiac and skeletal muscle (62).
Yuichiro Maeda’s group (at the RIKEN SPring8 Institute, Japan) has reported crystals of the actin-binding
COOH-terminal half of Tmod that diffract to 1.45 Å (153).
This structure is now in press in the Biophysical Journal.
The overall shape of a slightly truncated Tmod has been
determined from low-angle X-ray scattering by Fujisawa
et al. (101), also in Maeda’s group. Tmod is 115 Å long and
consists of two domains, one 65 Å long, the other slightly
smaller, where the long axes of the two are tilted by ⬃40°
relative to each other and is illustrated in Figure 16. The
authors proposed a model for Tmod in which the rodshaped NH2-terminal half binds to the NH2 terminus of
tropomyosin, and the COOH-terminal region protrudes
from the pointed end of the actin filament and is slightly
bent toward the actin subunit at the end, capping the
pointed end (101). There appears to be 1–2 Tmod molecules/actin filament (98).
Both CapZ and Tmod can regulate filament length
and assembly in vitro and in vivo. By studying the incorporation of rhodamine-labeled actin monomers into native thin filaments of cardiomyocytes, Littlefield et al.
(164) concluded that both proteins can rapidly exchange,
ACTIN AND ACTIN BINDING PROTEINS
cleft is less open (⬃12°), a feature that lowers its affinity
for ATP. Subdomains 1 and 2 were difficult to resolve in
the crystals, and the authors replaced them with a polyalanine backbone. These and other features give the Arps
distinctive surface features. In mammalian cells, the five
associated proteins are p16, p20, p21, p34, and p40 (proteins are also called ARPC-5, -4, -3, -2, and -1, respectively)
of which the last is a seven-blade propeller structure with
a large insert that may be responsible, along with p20 and
p21, for anchoring the complex to the side of the actin
filament. p20 and p34 are located in the center of the
structure and may move as a block relative to Arp2 (244).
The principal function of Arp2/3 is to create branch
points by nucleating the assembly of filaments near ruffling membranes, a process that needs micromolar concentrations of ATP (195). Arp2/3 is also considered to be
a cross-linking protein, and some authors believe it can
cap the pointed (depolymerizing) ends of actin filaments,
although this is still controversial.
Nucleation of filament assembly by Arp2/3, first observed by Welch et al. (302), is unlike any other ABP.
Extracellular signals such as growth factors are transmitted through G protein-coupled receptors (e.g., Cdc42) to
Wiskott-Aldrich syndrome protein (WASP), the neuronal
homolog of WASP (N-WASP), or suppressor of cAMP
receptor 1 (Scar1). PIP2 may be also involved in the
activation of WASP (242). Activated WASP has sites that
bind both an actin monomer and the Arp2/3 complex.
Arp2/3 then changes its conformation so that it can nucleate the assembly of new actin filaments and assemble
a “daughter” filament at an angle of ⬃70° to the “mother”
filament (Fig. 17). As a consequence, Arp2/3 is localized at
branch points in the cortical filament network, binding
the pointed end of a new (uncapped) daughter filament to
the mother filament thus leaving the barbed (growing)
end available for filament elongation. It is not clear at this
stage whether WASP dissociates from activated Arp2/3 at
the branch point.
Zalevsky et al. (314) recently examined how N-WASP
and Scar1 differentially activate the Arp2/3 complex to
produce actin networks with different three-dimensional
architectures. They found that nucleation was slowest for
Scar1, 16-fold higher for WASP, and 70-fold higher still for
the neuronal analog N-WASP. Their data fit a mathematical model in which one activated Arp2/3 complex, one
actin monomer, and an actin filament combine into a
preactivation complex. This then undergoes a first-order
activation step to become a nucleus. They went on to
FIG. 17. A cartoon of the molecular interactions Arp2/3. Components are as follows: activated Arp2/3 (red), inactive
Arp2/3 (pink), N-WASP (green), Cdc42 (blue), phosphatidylinositol 4,5-bisphosphate (PIP2) (yellow), and actin (orange).
Inactive Arp2/3 is activated by N-WASP under the control of Cdc42 and PIP2, and filament formation is nucleated. Actin
filaments assemble by addition of monomers at the barbed end. The N-WASP-Arp2/3 complex can bind to the side of the
“mother” filament to create a branch point that also polymerizes. This branching assembly creates an “out-pushing” of
the cell membrane. The branching of “daughter” filaments occurs at an angle of 70° to the mother filament. The solid
black arrows indicate the directions of the polymerization forces.
Physiol Rev • VOL
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
2. Cellular functions
459
460
DOS REMEDIOS ET AL.
Physiol Rev • VOL
“comet tails” (the review by Pantaloni et al. is available at
www.sciencemag.org/cgi/content/full/292/5521/1502/DC1).
The role of WASP/Scar proteins is critical for cytoplasmic organization in a range of organs during development in Drosophila (315) and presumably Arp2/3 and the
WASP family of proteins play essential roles in the development of mammalian cells.
Several reviews have recently been published on the
role of Arp2/3 in actin polymerization and the factors that
control the functions of Arp2/3. These include the role of
WASP family of proteins (50, 125, 242), and a review of
the structure of Arp2/3 itself (31).
Thus Arp2/3 is a complex protein with an even more
complicated method of activation. Single-headed myosin I
motor proteins interact with Arp2/3 via their Scar-homology 3 (SH3) domain (160), but what are motor protein
doing in a process that is usually regarded as an actin-only
process? Certainly there is no evidence to suggest that
Arp2/3 is involved in muscle contraction, although it
clearly plays a role in actin-based motility.
Several other questions remain. What is the nature of
the conformational changes needed to activate Arp2/3?
Can Arp2/3 promote filament growth at the pointed ends
of filaments? What is the role of profilin? What is the role
of ATP hydrolysis, and is this related to changes in the
putative nucleotide cleft in Arp2 or Arp3? Does activation
of filament nucleation produce filament growth at the
barbed or pointed end? Why do branch points occur
where they do? How do Cdc42 and PIP2 cooperate to
release the inhibitory effect of WASP on Arp2/3?
IV. ROLE OF TERNARY COMPLEXES IN
REGULATING ACTIN CYTOSKELETON
ASSEMBLY
The foregoing discussion has probably created the
impression that regulation of assembly and disassembly
of the actin cytoskeleton is complex. In fact, the complexity may be comparable to the system that controls gene
expression in eukaryotic genes where a promoter element
contains the promoter-proximal upstream control elements as well as enhancers and silencer elements. A
further level of regulation is then provided by distal regulatory sequences. DNA-bound proteins that can alter the
state of condensation of the chromatin DNA also influence these DNA sequences. The control of actin cytoskeleton assembly/disassembly may be even more complex.
Just as we do not have a precise understanding of the
control of gene expression, we are also only beginning to
understand the control of location and mechanism of
assembly of the actin cytoskeleton. Not only are there
potentially multiple ABPs, but they are also capable of
simultaneous expression within a single cell. The presence of actin ligands such as ATP or ADP, Ca2⫹ or Mg2⫹
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
show that WASP and Scar1 bind actin and Arp2/3 with
nearly identical affinities but stimulate rates of actin nucleation that vary by almost 100-fold. Differences in nucleation rates were attributed to differences in the number of COOH-terminal acidic amino acids that are responsible for different rates of actin assembly. The authors
therefore concluded that Arp2/3 is not regulated by a
simple on-off switch.
In the vicinity of the ruffling membrane, new monomers are added to the barbed-end of the growing filament
creating a polymerization-based “out-pushing” of the cell
membrane (Fig. 17). The Y-shaped branch points produce
a cross-linked meshwork of filaments that act as a platform against which the actin polymerization can push.
The entire Arp2/3 complex along with WASP have been
observed in electron microscopic images of branching
filaments (293). Mullins et al. (195) refer to this process of
branching filaments as “dendritic nucleation.” Assembly
of monomers at the barbed ends of these filaments generates the force needed to push the ruffling membrane
forward. Branching may occur at the barbed ends or
along the side of the mother filament (or both). Ishiwata
and co-workers (103) recently addressed this question by
the directly visualizing actin assembly using total internal
reflection fluorescence microscopy. In the presence of
Arp2/3 and N-WASP, branching occurred predominantly
along the sides of growing filaments.
Nucleation and branching are carried out close to the
leading edge of motile cells. Arp2/3 is not generally associated with filaments deeper into the cortex of the cell
where the actin filaments are known to bind tropomyosin.
A recent report (28) demonstrated that tropomyosin located deeper into the cytoplasmic cortex inhibits branching and nucleation by WASP-activated Arp2/3, thus restricting its distribution to the cytoplasmic perimeter.
In Listeria, nucleation of new filaments occurs in
response to the expression of ActA that mimics the function of WASP in recruiting Arp2/3. ActA contains 612
amino acids and is anchored into the bacterial membrane.
In a fascinating series of experiments, Cameron et al. (39)
showed that small microspheres coated with purified
ActA could undergo movement generated by actin polymerization. Thus ActA alone can nucleate actin assembly.
The minimum set of proteins and conditions needed for
actin-based motility was recently defined for the propulsion of Listeria by Marie-France Carlier and colleagues
(166). ATP, Arp2/3, actin filaments with bound ADP, ADF/
cofilin, and a capping protein (e.g., CapZ) were all that
was required. ADF/cofilin accelerates the treadmilling of
monomers through the filaments (see above) and drives
the Listeria through the actin-rich cytoplasm of its host
cell. A review from this group (220) highlighted the complexity of the mechanism of actin assembly in cell biology. Clearly other intracellular factors such as profilin and
␣-actinin regulate the rates of actin assembly in these
ACTIN AND ACTIN BINDING PROTEINS
that DNase I may act as a regulator of actin-cofilin interaction and that the ternary complex may therefore be an
important element in the regulation of microfilament assembly (142), effectively removing cofilin from the cellular pool and thereby preventing its normal ability to polymerize. The ternary complex may be destabilized by
other ABPs such as profilin or gelsolin, which alter the
bound nucleotide presumably via an allosteric conformational change. Such a regulatory mechanism would occur
on a slow time scale equivalent to the rate of appearance
and disappearance of DNase I and would depend on the
balance of free ABPs, pH, and divalent cations present in
the cell.
A. Cofilin, Actin, and DNase I
B. Ternary Complexes of Actin With Two
or More ABPs
It is widely accepted that regulation of the rate of
assembly/disassembly (treadmilling) of the actin cytoskeleton is achieved by a single ABP, probably ADF/
cofilin (21). However, the widespread use of DNase I
affinity chromatography to capture complexes of actin
with other ABPs (cofilin, destrin, actobindin, adseverin,
fimbrin, and radixin) suggests that ternary complexes
exist, at least in vitro (133). We recently reported the
formation of ternary complexes of actin, cofilin, and
DNase I (57) using nondenaturing polyacrylamide gel
electrophoresis. The affinity of DNase I for G-actin is high
(nM Kd), significantly higher than for actin-cofilin (␮M)
(266). However, the affinity of cofilin for the binary DNase
I-actin complex is higher than for actin alone, and the
reverse is also true, namely, the affinity of DNase I is
higher for the actin-cofilin complex than for actin alone.
Thus when DNase I is present in a mixture of actin
monomers and the cofilin-actin binary complex, it binds
to the binary complex forming a ternary (cofilin-actinDNase I) complex before it binds to the remaining free
actin (205). This ternary complex was recently suggested
based on colocalization in monkey kidney epithelial
cells (56).
In a report only just completed, we (81) have documented the magnitude of actin conformational changes as
a consequence of separately binding cofilin and DNase I
and in the ternary cofilin-actin-DNase I complex. We used
fluorescence and fluorescence resonance energy transfer
spectroscopy to demonstrate significant conformational
changes in the small domain (subdomains 1 and 2) of
monomeric actin. DNase I binding to actin increases by
⬃1 Å the distance between probes attached to Gln-41 and
Cys-374. When cofilin alone binds to actin, this distance
decreases by ⬃3 Å, but when both bind (ternary complex)
the distance increases by ⬃4 Å. This effect is completely
reversible and is independent of the order of addition.
Thus the effects are not simply additive. We speculated
The actin-DNase I binary complex is unusual but not
unique in its ability to form a ternary complex with cofilin.
Profilin can be chemically cross-linked to actin-DNase I;
however, in the absence of cross-linking, a negative cooperativity was observed (12), i.e., profilin dissociates
DNase I from actin, and vice versa. DNase I and gelsolin
also bind to actin in a negatively cooperative way. Chemical cross-linking of Cys residues between T␤4 and actinDNase also revealed the formation of a ternary complex
(239). These authors found that the Kd of T␤4 for actinDNase I was the same as for actin alone (see Table 1), and
they concluded that the binding loci for these two ABPs
do not overlap. Others (13) reported that T␤4 competes
with gelsolin segment 1, but both are dissociated by the
binding of DNase I to actin. A similar negative cooperativity between spatially separated actin-binding sites was
observed between DNase I and gelsolin segment 1 (12).
Thus, although the binding site for DNase I is physically
separated from the sites for gelsolin segment 1, T␤4, and
profilin, the negatively cooperative nature of their interactions suggests that an allosteric conformational change
may be transmitted along the “length” of the small domain
of actin.
Capping of the pointed ends of actin filaments by
tropomodulin (Tmod, discussed above) appears to involve a ternary complex. Tmod binds only weakly to
F-actin, but the affinity of Tmod for actin increases nearly
three orders of magnitude in the presence of tropomyosin. Rapid exchange of Tmod at this end permits assembly
and disassembly of these filaments. The same is probably
true for CapZ and other proteins that cap the barbed end
of filaments.
In a two-hybrid system, actin interacting protein 1
(Aip1; a 65-kDa protein) was shown to form a ternary
complex with actin and ADF/cofilin (245). In yeast, Aip1
interacts with actin via subdomains 3 and 4 (5), and some
of these residues do not overlap those for ADF/cofilin and
Physiol Rev • VOL
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
can also be important, as well as pH. Under these conditions, it would be surprising if control of the state of
assembly was left to single ABPs as suggested in the
preceding sections. The next level of complexity to consider is whether ternary complexes play a role and if so
how is this achieved. Of course, quaternary and higher
orders of complexity may also exist. There can be no
argument that complexes of more than one ABP control
actin filament functions (see sect. IIIG). Activation by
Ca2⫹ via the tropomyosin-troponin C/I/T complex is now
well established, but there are also proteins that regulate
filament length, anchoring into the Z-disk of sarcomeres,
cap thin filaments, and connect actin to microtubules and
the intermediate filament system.
461
462
DOS REMEDIOS ET AL.
V. THE CYTOSKELETON AND PATHOLOGY
Many of the proteins discussed above are affected by
mutations resulting in the development of significant pathology. Given the central role played by actin and the
ABPs, this outcome is not surprising (see Ref. 21a).
A. Actin
Actin has a remarkably conserved amino acid sequence, probably more than any other protein. This alone
suggests that mutations have been eliminated rather than
tolerated by the genome. Because actins are essential for
motility, force generation, and other important cellular
functions, it is remarkable that so few diseases are associated with primary defects in microfilaments and their
regulators. No doubt, part of the reason is that there are
usually multiple actin genes, i.e., the problem is circumvented by redundancy.
Perhaps the same evolutionary pressures that maintained the highly conserved sequences of actins also protected them from being a major player in the disease
process. Because of its pivotal role, defects or alterations
in actin probably have dramatic effects on survival when
no alternate actin gene is available (e.g., yeast has a single
actin gene which is lethal when disrupted).
A major obstacle to studying the altered properties of
mutant actins is the considerable difficulty cell biologists
have had in expressing actin using standard protein expression systems. Bacteria have no actin of their own and
are therefore a logical choice as an expression system.
Physiol Rev • VOL
Only one report (100) suggested that bacterial expression
could yield native actin, and the findings of this report
have not been repeated. Yeast can be used to express
mutant actins (246), although the yield of the inserted
actin gene product is impractically low.
There are mouse fibroblast cell lines that express
almost pure nonmuscle ␤-actin (252), but these are special cases. Most cells express more than one actin isoform
and, although several methods have been described for
separating them, for example, hydroxyapatite column
chromatography (260), they are technically challenging.
Some actin mutations (for example, removal of the NH2terminal acidic residues) are not essential for the survival
of yeast but are needed to activate myosin ATPase activity (64).
More than 150 papers have reported ⬃300 actin mutations (266). Only a handful deal with human actin mutations, and most of these are deletion mutants near the
COOH terminus. Three at residues 336, 346, and 356 resulted in the production of nonfunctional actin. The remaining deletions at residues 366, 373, and 374 had significantly reduced functionality. In a recent review, Marston and Hodgkinson (178) observed that 13 actin
mutations were known to cause hypertrophic cardiomyopathy (E99K, A331P, P164A, A295S), dilated cardiomyopathy (R312H, E361G), severe nemaline myopathy
(R183C, R256H, N280K, V370F, L94P), and actin myopathy
(G15R, V163L). Clearly, caution is needed in drawing
sweeping conclusions about the functional effects of
these structural changes. However, when these mutant
actins can be expressed in substantial quantities, we can
begin to understand the functional consequences of mutant actins in organs where more than one actin gene is
coexpressed (25).
B. Gelsolin
If the actin cytoskeleton plays a significant role in
disease, and in particular tumors (215), then we would
also expect its associated ABPs to be involved in the
disorder or disease. The functions of gelsolin have been
described above, but it is also a substrate for the apoptotic enzyme caspase-3 (150) that cleaves the actin-binding
domain of gelsolin from its calcium-binding domain. Physical separation of these two domains coincides with
plasma membrane blebbing, one of the hallmarks of apoptosis. Overexpression of the Ca2⫹-independent NH2-terminal half of gelsolin induces apoptosis, whereas gelsolin
null neutrophils have a delayed onset of apoptosis (150).
Cancer cells characteristically lack apoptotic mechanisms and express less gelsolin (157), whereas expression
of mutated (His-321) gelsolin is associated with the loss of
tumorigenicity of transformed cells (102).
Normally, plasma gelsolin functions as a part of the
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
other ABPs. AIP1 alone has little or no filament severing
activity, but it exhibits a cooperative filament severing
activity in the presence of Xenopus ADF/cofilin (245).
This ability of Aip1 to recognize ADF/cofilin bound to
filaments may be important in the modulation of ADF/
cofilin activity.
Drugs like cytochalasin D act directly on the actin
cytoskeleton by altering the rate of assembly/disassembly
at both the pointed and barbed ends of the filaments. A
recent structural analysis of latrunculin-A (193) revealed
that it binds only to actin monomers and prevents polymerization by inducing specific structural changes. The
binding site was not previously known to be an ABP
binding locus. Crystals of actin-gelsolin segment 1 were
soaked in latrunculin-A, and crystallographic refinement
of the resulting ternary complex revealed that it tightly
links subdomains 2 and 4 of actin. Latrunculin-A produces
local rather than global structural changes in actin, allosterically altering the surfaces involved in the assembly of
F-actin. This effect is achieved by restricting the rotations
of subdomains 2 and 4. Thus ternary complexes involving
actin appear to be important for the action of some drugs.
ACTIN AND ACTIN BINDING PROTEINS
actin-scavenging system to assemble and disassemble actin filaments. The D187N and D187Y gelsolin mutations
generate a 71-residue fragment that forms amyloid fibrils
in humans. These mutations were also associated with
familial amyloidosis of the Finnish type (FAF) (139). The
symptoms of FAF patients include the accumulation in
their tissues of amyloid derived from plasma gelsolin
(285). Neither the recombinant wild-type nor the D187N
FAF-associated gelsolin fragments self-associate into
amyloid at pH 7.5, but incubation of either fragment at
low pH values (6.0 – 4.0) produces well-defined fibrils (237).
C. Tropomodulin
D. Connections Between the Extracellular Matrix
and the Nucleus
Maniotis et al. (175) showed that a mechanical tug on
cell surface receptors can immediately change the organization of proteins in the cytoplasm and the nucleus.
They showed that by specifically pulling on cell surface
Physiol Rev • VOL
integrin receptors, the nucleus was elongated in the direction of the pull. Furthermore, they showed that this
effect on the nuclear membrane was mediated by the
actin cytoskeleton. The actin cytoskeleton also provides a
physical connection between ion channels such as ClC3
chloride ion channels in the cell membrane and the interior of the cell (33).
Defects, deficiencies, or deletions of any component
of the linkage between the extracellular matrix (laminin-2, ␣-dystroglycan), trans-sarcolemmal proteins (␣-,
␤-, and ␥-sarcoglycans, ␤-dystroglycan), the cytoskeleton
(dystrophin, dystrobrevin, syntrophin, actin), and the nucleus (emerin, lamin A/C) would dramatically alter this
putative mechanical signaling (175) and thus affect the
mechanical stability of cardiomyocytes.
Altered expression of cytoskeletal, linkage, and extracellular proteins has been reported in hypertrophied
cardiomyocytes (280, 282) and also in failing myocytes
(61, 83, 122). Changes in expression levels of fibronectin,
laminin, fibrillin, fibulin SC1, and decorin, all components
of the extracellular matrix, are upregulated in rat experimental myocardial infarctions (269). A number of monogenetic cardiomyopathies have recently been detailed in a
selected number of families, which include a range of
different genes involved in the surface-nucleus pathway
(55). Thus cytoskeletal proteins may be involved in the
progressive defects leading to heart failure. Hypertrophic
cardiomyopathy appears to be a final common outcome
for gene defects involving an astonishing variety of cytoskeletal and sarcomeric proteins.
E. Heart Failure
Familial hypertrophic cardiomyopathy (FHC) is an
autosomal dominant disorder resulting in increased myocardial mass and myofibrillar disarray. It is the most
common cause of sudden death in the young. Linkage
studies and candidate-gene approaches have revealed disease-causing mutations in more than eight different genes
all encoding sarcomeric proteins. More than 100 mutations have already been identified, and they involve practically every protein component in the sarcomere. Mutations are observed in ␤-myosin heavy chain (105), troponin I (144, 92), troponin T (238), ␣-tropomyosin (282),
myosin-binding protein C (296), regulatory and essential
myosin light chains (226), cardiac ␣-actin (209), titin
(253), and desmin (122, 253). However, these mutations
are present in only half the patients (296), and consequently, there must be other gene mutations that cause
FHC, some of which will probably involve the cytoskeleton. The common phenotype resulting from a wide range
of cytoskeletal/sarcomeric proteins may well prove to be
a reflection of the functional linkage of these proteins to
both the extracellular matrix and the nucleus in cardiomyocytes.
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
Changes in gene expression of this thin filament capping protein produce a surprisingly extensive range of
phenotypes. Increases in Tmod expression levels are associated with the dilation (rather than the hypertrophic)
component in human dilated cardiomyopathy (306). In a
transgenic mouse model, inhibition of Tmod expression
prevents dilation (279) but not hypertrophy of the myocardium. Tmod is a promising development in our understanding of this baffling disorder (278).
Graves’ disease is a complex autoimmune disease
characterized by multiple susceptible loci involving cytotoxic T cells, HLA antigens, and thyroid-stimulating hormone receptors (299). Human Tmod has ⬃40% identity at
the NH2 and COOH termini of the 64-kDa autoantigen
protein identified in Graves’ disease. The insertion of
several homologous repeats in the midsection of the
Graves’ protein together with an extension of the COOH
terminus of Tmod accounts for the differences in length
between the Graves’ protein (572 residues) and tropomodulin (359 residues) (299). It is therefore likely the two
genes evolved from a common ancestral gene (276). The
gene for Tmod was assigned to human chromosome 9 in
a region also known to contain several other genes and
disease loci, for example, an unrelated cytoskeletal gene
for gelsolin.
Finally, two genes encoding for Tmod that map next
to another on human chromosome 15 make them candidates for amyotrophic lateral sclerosis and a dyslexia
gene as well as a candidate gene for limb girdle muscular
dystrophy (70). At this stage it is not clear how these
syndromes are related to Tmod expression disorders, but
the clear link suggests that Tmod is a key gene.
463
464
DOS REMEDIOS ET AL.
(SNPs) from human populations. The idea here is that
SNPs (unlike most other analytical methods) are potentially capable of discovering the cause of disease.
A number of companies have developed technologies
for analyzing SNPs, but the most challenging aspect of the
technology is its ability to accurately detect very small
differences between DNA restriction fragments in which
only one base pair has been replaced (e.g., T instead of A
or G instead of a C base pair, or vice versa). Sequenom
(www.sequenom.com) has developed a method that very
accurately “weighs” the DNA fragments using mass spectrometry which achieves precisions of ⬎99.9%. It is debatable whether this accuracy is really needed, although
their approach was both unusual and effective. They have
identified at least three cytoskeletal proteins that are
statistically associated with specific diseases in specific
human populations. These diseases have a late onset and
are not associated with mortality in infants, juveniles, or
young adults.
F. Gene Arrays
H. Cell Signaling and Actin Microfilaments
Yang et al. (306) used gene array technology to examine altered genes in failing human hearts. They observed an increased expression of gelsolin and a significant upregulation of ␣-B-crystallin, a desmin-associated
protein that protects or chaperones actin and desmin
filaments from stress-induced damage. ␣-B-crystallin is a
member of 27-kDa heat shock protein family. These data
(using Affymetrix gene arrays) were confirmed by us
(using NEN arrays) (135). They support the view that the
actin cytoskeleton is damaged in DCM. Samples of failing
left ventricles have been studied by two-dimensional gel
electrophoresis of human DCM patients (68) and of an
animal model of DCM (122). In another form (familial
hypertrophic) of human heart failure, the expression of
␣-actin, plasma gelsolin, thymosin ␤4, and several other
cytoskeletal proteins (23) changed more than twofold.
Finally, gene arrays have been used to examine an experimental myocardial infarction using failing rat hearts
(269). Cytoskeletal actin and several ABPs (␣-actin, thymosin ␤4 and ␤19, Arp2/3 and a number of other ABPs not
reviewed here) were upregulated. The data clearly implicate the involvement of cytoskeletal and myofibrillar
genes in human heart failure. Thus there is growing evidence for altered expression of cytoskeletal proteins in
heart failure. It is likely that similar results will be reported for the failure of other organs.
G. Single Nucleotide Polymorphisms, Diseases, and
the Cytoskeleton
A relatively new method of determining the connections between human diseases and gene mutations involves analysis of single nucleotide polymorphisms
Physiol Rev • VOL
In mammals, there is a well-defined connection between extracellular signals (like insulin), their membrane
receptors, intracellular signals (Ras and Rac), and the
regulators of actin microfilament assembly (Arp2/3, gelsolin, ADF/cofilin). Because the binding of ligands to
some surface receptors is known to activate genes, the
cytoskeletal links may extend from the membrane all the
way to the nuclei and perhaps even to the chromosomes.
The control of these connections in yeast, Drosophila,
and mammalian cells was recently summarized by Hall
and colleagues (24). The relationships between components of the signaling pathways are complex and are
beyond the scope of this review.
VI. UNRESOLVED ISSUES
A. Atomic Structure of F-actin
It is common to assume that because we know all or
most of the structural atomic details of the actin monomer, the available models of F-actin are equally valid.
How good is this assumption?
It is true that the known atomic structures of monomeric actins are very similar. However, significant differences are beginning to emerge. Nearly all of the available
atomic structures were achieved using actin cocrystallized with an ABP. Recently, a new structure of actin was
reported (217) that is not complexed to an ABP. These
authors observed “a dramatic conformational change in
subdomain 2” compared with previous structures. It will
therefore be of considerable interest to see the effect the
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
Studies of human dilated cardiomyopathy (DCM)
(306) and of mouse models (279) of DCM also point to a
critical role of cytoskeletal elements at the Z-disks in
chamber dilation. Mice lacking the muscle LIM-domain
protein (MLP), a Z-disk protein that interacts directly with
␣-actinin, displayed many of the features of human DCM
(8). Mutations (usually more than one per protein) in
other proteins associated with ␣-actinin, cardiac ␣-actin,
desmin, and titin were subsequently found in familial
forms of DCM (76, 209, 253). Also, mutations of other
Z-disk components in the mouse result in DCM (55, 59).
Mechanical stress on the extracellular matrix leads to
defects in Z-disk formation, sarcomeric stabilization, and
altered muscle-cell signaling, thereby driving the onset of
chamber dilation and heart failure (58). In a dog model of
heart failure, cytoskeletal actin was upregulated while
desmin (a protein in the intercalated disks) was downregulated (122).
ACTIN AND ACTIN BINDING PROTEINS
B. Functional Differences Between Actin Isoforms
The great majority of the several thousand references
on this topic have used rabbit skeletal muscle actin. Only
very small sequence differences exist between different
actin isoforms, and these probably have functional consequences. For example, the myocardium of small mammals like mice and rats expresses almost pure cardiac
␣-actin, whereas cardiomyocytes in larger animals (including humans) express about equal amounts of cardiac
and skeletal ␣-actin (25, 113). Why do larger hearts express skeletal muscle actin? ␥-Actin isoforms are located
just beneath the sarcolemma, whereas only ␣-actins are
incorporated into sarcomeres (113). We do not know if
isoforms can coassemble or whether such filaments have
different effects on activities such as myosin ATPase.
Very little is known about the functional differences between actin isoforms. Even less is known about whether
certain ABPs bind to particular actin isoforms and
whether differences in actin isoform structure determine
which ABPs bind.
The situation is no clearer for ADF/cofilin. We know
there are functional differences (nucleation, elongation,
fragmenting, binding ratios to actin) between ADFs and
cofilins, but comparisons based on existing literature are
difficult because groups investigating these properties often use different (yeast, chick, human) isoforms. There
may also be differences between isoforms within the
cofilins and ADFs, and these will have subtle but important functional effects.
complex involving monomeric actin in vitro, is there a
role for ternary complexes in vivo? The more details we
learn about the ABPs, the more they seem to fit into
families with common cellular functions. Not all DNases
bind actin, so it is still not clear if they can play a universal
role.
D. Undiscovered ABPs
Despite the proliferation of ABPs over recent years
(65 new proteins have been discovered over the past 5
years, Ref. 151), there are still significant areas of the
actin monomer surface that do not appear to be the site
for ABPs (see Fig. 6). Some of this surface may come to
be occupied by known but not yet fully characterized
ABPs. It is unlikely that we have discovered all of the
ABPs, and it seems a safe bet to suggest that as the human
genome and other genomes become refined, at least some
of the “orphan” genes will turn out to be new ABPs.
E. Cooperative Binding of ABPs Along Filaments
There is a spatial separation and organization of filament nucleation, elongation, branching, and other processes involving actin assembly and disassembly, but little or nothing is known about the controls of these processes. For example, we know that Arp2/3 produces a
regular branching of actin microfilaments and that this
regular branching is important for the protrusion of cytoplasmic blebs and fingers (195), but what controls the
distribution of Arp2/3? In muscle thin filaments, binding
of a myosin head at one point alters the probability of a
second head binding up to several monomers along the
filament. What conformational changes are involved here,
and how are they propagated along the filaments? ADF/
cofilin binds cooperatively to F-actin, but does this cooperativity depend on the actin isoform?
F. Cytoskeletal Proteins in the Nucleus
A number of cytoskeletal proteins are found in the
nucleus including cofilin (56, 203) (which has a nuclear
localization peptide) and actin (which lack this sequence). We do not know the functions of these proteins
or how they are regulated.
G. Disease and the Cytoskeleton
C. Ternary Complexes of Monomeric Actin
With ABPs
We have reviewed the evidence for a ternary complex
between actin monomers and two ABPs, cofilin and
DNase I. Although there is good evidence for a ternary
Physiol Rev • VOL
The role of the cytoskeleton and ABPs in disease is
an emerging story. There can be little doubt that several
cytoskeletal proteins are positively associated with significant human disease. Sequenom of San Diego and Gemini
Genomics PLC of Cambridge recently discovered three of
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
new structure from Dominguez et al. (217) has on a model
of F-actin with its bound ADP. Bound nucleotide can have
dramatic effects on the structure of actin and on the
affinities of ABPs for actin. Three-dimensional reconstructions of electron micrographs of actin filaments are
based on low-resolution structural data. Thus the available models make assumptions about the arrangement
and orientation of the monomers in the filament. To date,
most filament models (128, 167, 283) have been constructed using the structures from actin cocrystals with
ABPs, and there is an assumption that this is unaltered in
the filament. Arguably, filaments are the most biologically
relevant form of actin in cells, so there is a real need to
resolve their structure at high resolution. Several groups
are currently working on this most difficult, perhaps intractable, structural problem.
465
466
DOS REMEDIOS ET AL.
these proteins, but their identities have not yet been
disclosed. However, it is possible that identification of
specific disease-associated genes may not produce a complete understanding of the disease process for we know
that a single phenotype like hypertrophic cardiomyopathy
can be the product of mutations in almost any one of the
myofibrillar proteins.
H. Prokaryotic Cytoskeletal Elements
We are grateful to Roger Craig, Roberto Dominguez,
Shin’ichi Ishiwata, Glenn King, Steve Marston, Takashi Obinata,
Tetsura Fujisawa, Alla Kostyukova, Yuichiro Maeda, Bob Robinson, Dan Safer, Fusinita van den Ent, and Willy Wriggers for
many of the figures used here and for valuable contributions on
the manuscript and for discussions.
We are grateful for the support of the National Health and
Medical Research Council of Australia, the Australian Research
Council, the National Heart Foundation of Australia, and the
Medical Foundation of the University of Sydney.
Address for reprint requests and other correspondence: C.
G. dos Remedios, Institute for Biomedical Research, Muscle
Research Unit, Dept. of Anatomy and Histology, F13, Univ. of
Sydney, Sydney 2006, Australia (E-mail: crisdos@anatomy.
usyd.edu.au).
REFERENCES
1. ABE H, ENDO T, YAMAMOTO K, AND OBINATA T. Sequence of cDNAs
encoding actin depolymerizing factor and cofilin of embryonic
chicken skeletal muscle: two functionally distinct actin-regulatory
proteins exhibit high structural homology. Biochemistry 29: 7420 –
7425, 1990.
2. ABE H, OBINATA T, MINAMIDE LS, AND BAMBURG JR. Xenopus laevis
actin-depolymerizing factor/cofilin: a phosphorylation-regulated
protein essential for development. J Cell Biol 132: 871– 885, 1996.
3. AEBI U, FOWLER WE, ISENBERG G, POLLARD TD, AND SMITH TD. Crystalline actin sheets their structure and polymorphism. J Cell Biol
91: 340 –351, 1981.
4. ALLEN ML, DOBROWOWOSKI JM, MULLER H, SIBLEY LD, AND MANSOUR
TE. Cloning and characterization of actin depolymerization factor
from Toxoplasma gondii. Mol Biochem Parasitol 88: 43–52, 1997.
5. AMBERG DC, BASART E, AND BOTSTEIN D. Defining protein interactions with yeast actin in vivo. Struct Biol 2: 38 –35, 1995.
6. AMPE C, MARKEY F, LINDBERG U, AND VANDEKERCKHOVE J. The primary
structure of human platelet profilin: reinvestigation of the calf
spleen profilin sequence. FEBS Lett 228: 17–21, 1988.
7. ARBER S, BARBAYANNIS FA, HANSER H, SCHNEIDER C, STANYON CA,
BERNARD O, AND CARONI P. Regulation of actin dynamics through
phosphorylation of cofilin by LIM-kinase. Nature 393: 805– 809,
1998.
8. ARBER S, HUNTER JJ, ROSS J JR, HONGO M, SANSIG G, BORG J, PERRIARD
JC, CHIEN KR, AND CARONI P. MLP-deficient mice exhibit a disrupPhysiol Rev • VOL
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
It seems likely that the ancestor of eukaryotic actin is
the bacterial protein MreB. This protein can self-assemble
and contributes to structures that resemble straightened
actin rails. This discovery prompts questions like: Are
there MreB-binding proteins, equivalent to ABPs, that
control filament assembly? Do bacteria have motor proteins that employ the MreB “rails”?
tion of cardiac cytoarchitectural organization, dilated cardiomyopathy, and heart failure. Cell 88: 393– 403, 1997.
9. ASAKURA S, KASAI M, AND OOSAWA F. The effects of temperature on
the equilibrium state of actin solutions. J Polymer Sci 44: 35– 49,
1960.
10. AZUMA T, WITKE W, STOSSEL TP, HARTWIG JH, AND KWIATKOWSKI DJ.
Gelsolin is a downstream effector of rac for fibroblast motility.
EMBO J 17: 1362–1370, 1998.
11. BAINS NPS, GORBATYUK VY, NOSWORTHY NJ, ROBSON SA, MACIEJEWSKI
MW, DOS REMEDIOS CG, AND KING GF. Letter to the Editor: backbone
and side chain 1H, 15N, and 13C assignments for chick cofilin.
J Biomol NMR 22: 193–194, 2002.
12. BALLWEBER E, GIEHL K, HANNAPPEL E, HUFF T, JOCKUSCH BM, AND
MANNHERZ HG. Plant profilin induces actin polymerization from
actin: beta-thymosin complexes and competes directly with betathymosins and with negative co-operativity with DNase I for binding to actin. FEBS Lett 425: 251–255, 1998.
13. BALLWEBER E, HANNAPPEL E, HUFF T, AND MANNHERZ HG. Mapping the
binding site of thymosin beta4 on actin by competition with G-actin
binding proteins indicates negative co-operativity between binding
sites located on subdomains 1 and 2 of actin. Biochem J 327:
787–793, 1997.
14. BALLWEBER E, HANNAPPEL E, HUFF T, STEPHAN H, HAENER M,
TASCHNER N, STOFFLER D, AEBI U, AND MANNHERZ HG. Polymerisation
of chemically cross-linked actin:thymosin ␤4 complex to filamentous actin: alteration in helical parameters and visualisation of
thymosin ␤4 binding on F-actin. J Mol Biol 315: 613– 625, 2002.
15. BALLWEBER E, HANNAPPEL E, NIGGEMEYER B, AND MANNHERZ HG.
Induction of the polymerization of actin from the actin:thymosin
beta 4 complex by phalloidin, skeletal myosin subfragment 1,
chicken intestinal myosin I and free ends of filamentous actin. Eur
J Biochem 223: 419 – 426, 1994.
16. BAMBURG JR. Proteins of the ADF/cofilin family: essential regulators
of actin dynamics. Annu Rev Cell Dev Biol 15: 185–230, 1999.
17. BAMBURG JR AND BERNSTEIN BW. Actin and actin-binding proteins in
neurons. In: The Neuronal Cytoskeleton, edited by Burgoyne RD.
New York: Wiley-Liss, 1991, p. 121–160.
18. BAMBURG JR AND BRAY D. Distribution and cellular localization of
actin depolymerizing factor. J Cell Biol 105: 2817–2825, 1987.
19. BAMBURG JR AND DRUBIN DG. Actin depolymerizing factor (ADF)/
cofilin. In: Guidebook to the Cytoskeletal and Motor Proteins, edited by Kreis T and Vale R. Oxford, UK: Oxford Univ. Press, 1999,
p. 19 –24.
20. BAMBURG JR, HARRIS HE, AND WEEDS AG. Partial purification and
characterization of an actin depolymerizing factor from brain.
FEBS Lett 121: 178 –182, 1980.
21. BAMBURG JR, MCGOUGH A, AND ONO S. Putting a new twist on actin:
ADF/cofilins modulate actin dynamics. Trends Cell Biol 9: 364 –370,
1999.
21a.BAMBURG JR AND WIGGON OP. ADF/cofilin and actin dynamics in
disease. Trends Cell Biol 12: 598 – 605, 2002.
22. BAO L, LODA M, JANMEY PA, STEWART R, ANAND-APTE B, AND ZETTER
BR. Thymosin beta 15: a novel regulator of tumor cell motility
upregulated in metastatic prostate cancer. Nat Med 2: 1322–1328,
1996.
23. BARRANS JD, STAMATIOU D, AND LIEW C. Construction of a human
cardiovascular cDNA microarray: portrait of the failing heart. Biochem Biophys Res Commun 280: 964 –969, 2001.
24. BECK T, DELLEY PA, AND HALL MN. Control of the actin cytoskeleton
by extracellular signals. Res Prob Cell Motil 2002 32: 231–262, 2002.
25. BENNETTS BH, BURNETT L, AND DOS REMEDIOS CG. Differential expression of alpha-actin genes within the human heart. J Mol Cell Cardiol 18: 993–996, 1986.
26. BERNARD O, GANIATSAS S, KANNOURAKIS G, AND DRINGEN R. Kiz-1, a
protein with LIM zinc finger and kinase domains, is expressed
mainly in neurones. Cell Growth Differ 5: 1159 –1171, 1994.
27. BERNSTEIN BW, PAINTER WB, CHEN H, MINAMIDE LS, ABE H, AND
BAMBURG JR. Intracellular pH modulation of ADF/cofilin proteins.
Cell Motil Cytoskeleton 47: 319 –336, 2000.
28. BLANCHOIN L, POLLARD TD, AND HITCHOCK-DEGREGORI SE. Inhibition
of the Arp2/3 complex-nucleated actin polymerization and branch
formation by tropomyosin. Curr Biol 11: 1300 –1304, 2001.
29. BLANCHOIN L, ROBINSON RC, CHOE S, AND POLLARD TD. Phosphoryla-
ACTIN AND ACTIN BINDING PROTEINS
30.
31.
32.
33.
34.
35.
37.
38.
39.
40.
41.
42.
43.
44.
45.
46.
47.
48.
49.
50.
51.
52.
53.
Physiol Rev • VOL
54.
55.
56.
57.
58.
59.
60.
61.
62.
63.
64.
65.
66.
67.
68.
69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
NH2-terminal domains of the alpha-1 and beta subunits are not
required for actin capping and alpha 1 beta and alpha 2 beta
heterodimers bind differentally to actin. J Biol Chem 269: 6992–
6998, 1994.
CHEN H, BERNSTEIN BW, AND BAMBURG JR. Regulating actin-filament
dynamics in vivo. Trends Biochem Sci 25: 19 –23, 2000.
CHEN J AND CHIEN KR. Complexity in simplicity: monogenic disorders and complex cardiomyopathies. J Clin Invest 103: 1483–1485,
1999.
CHHABRA D, BAO S, AND DOS REMEDIOS CG. The distribution of cofilin
and DNase I in vivo. Cell Res 12: 207–214, 2002.
CHHABRA D, NOSWORTHY NJ, AND DOS REMEDIOS. The role of ATP, ADP
and divalent cations in the formation of binary and ternary complexes of actin with cofilin and DNase I. Electrophoresis 17: 3863–
3870, 2000.
CHIEN KR. Genomic circuits and the integrative biology of cardiac
diseases. Nature 407: 227–232, 2000.
CHIEN KR. Stress pathways and heart failure. Cell 98: 555–558, 1999.
CHIK JK, LINDBERG U, AND SCHUTT CE. The structure of an open state
of beta-actin at 2.65 Å resolution. J Mol Biol 263: 607– 623, 1996.
COLLINS JF, PAWLOSKI-DAHM C, DAVIS MG, BALL N, DORN GW II, AND
WALSH RA. The role of the cytoskeleton in left ventricular pressure
overload hypertrophy and failure. J Mol Cell Cardiol 28: 1435–1443,
1996.
CONLEY CA, FRITZ-SIX KL, ALMENAR-QUERALT A, AND FOWLER VM.
Leiomodins: larger members of the tropomodulin (Tmod) gene
family. Genomics 73: 127–139, 2001.
COOKE R. The role of the bound nucleotide in the polymerization of
actin. Biochemistry 14: 3250 –3256, 1975.
COOKE RK, BLAKE WT, AND RUBENSTEIN PA. Removal of the amino
terminal acidic residues of yeast actin. J Biol Chem 267: 9430 –9436,
1992.
COOPER JA AND SCHAFER DA. Control of actin assembly and disassembly at filament ends. Curr Opin Cell Biol 12: 97–103, 2000.
COOPER JA AND SCHAFER DA. Control of actin assembly and disassembly at filament ends. Curr Opin Cell Biol 12: 66 –97–103, 1984.
COPPIN CM AND LEAVIS PC. Quantitation of liquid crystalline ordering in F-actin solutions. Biophys J 63: 794 – 807, 1992.
CORBETT JM, WHY HJ, WHEELER CH, RICHARDSON PJ, ARCHARD LC,
YACOUB MH, AND DUNN MJ. Cardiac protein abnormalities in dilated
cardiomyopathy detected by two-dimensional polyacrylamide gel
electrophoresis. Electrophoresis 19: 2031–2042, 1998.
COSSART P. Actin-based motility of pathogens: the Arp2/3 complex
is a central player. Cell Microbiol 2: 195–205, 2000.
COX PR AND ZOGHBI HY. Sequencing, expression analysis, and mapping of three unique human tropomodulin genes and their mouse
orthologs. Genomics 63: 97–107, 2000.
CRAMER LP, BRIGGS LJ, AND DAWE HR. Use of fluorescently labelled
deoxyribonuclease I to spatially measure G-actin levels in migrating and nonmigrating cells. Cell Motil Cytoskeleton 51: 27–38, 2002.
CUNNINGHAM CC, STOSSEL TP, AND KWIATKOWSKI DJ. Enhanced motility in NIH 3T3 fibroblasts that overexpress gelsolin. Science 251:
1233–1236, 1991.
CURMI PMG, BARDEN JA, AND DOS REMEDIOS CG. Actin tube formation: effects of variation in commonly used solvent conditions.
J Muscle Res Cell Motil 5: 423– 430, 1984.
CURMI PMG, VAN DONKELAAR A, TULLOCH PA, AND DOS REMEDIOS CG.
A comparison of the unit cell of dried and hydrated actin microcrystals. In: Actin Structure and Function in Muscle and Nonmuscle Cells, edited by dos Remedios CG and Barden JA. Sydney,
Australia: Academic, 1983, p. 45–52.
CZISCH M, SCHLEICHER M, HORGER S, VOELTER W, AND HOLAK TA.
Conformation of thymosin beta 4 in water determined by NMR
spectroscopy. Eur J Biochem 218: 335– 44, 1993.
DALAKAS MC, PARK KY, SEMINO-MORA C, LEE HS, SIVAKUMAR K, AND
GOLDFARB LG. Desmin myopathy, a skeletal myopathy with cardiomyopathy caused by mutations in the desmin gene. N Engl J Med
342: 770 –780, 2000.
DE LA CRUZ EM, MANDINOVA A, STEINMETZ MO, STOFFLER D, AEBI U,
AND POLLARD TD. Polymerization and structure of nucleotide-free
actin filaments. J Mol Biol 295: 517–526, 2000.
DE LA CRUZ EM, OSTAP EM, BRUNDAGE RA, REDDY KS, SWEENEY HL,
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
36.
tion of Acanthamoeba actophorin (ADF/cofilin) blocks interaction
with actin without a change in atomic structure. J Mol Biol 295:
203–211, 2000.
BORK P, SANDER C, AND VALENCIA A. An ATPase domain common to
prokaryotic cell cycle proteins, sugar kinases, actin, and hsp70 heat
shock proteins. Proc Natl Acad Sci USA 89: 7290 –7294, 1992.
BORTHS EL AND WELCH MD. Turning on the Arp2/3 complex at
atomic resolution. Structure 10: 131–135, 2002.
BOWMAN GD, NODELMAN IM, HONG Y, CHUA NH, LINDBERG U, AND
SCHUTT CE. A comparative structural analysis of the ADF/cofilin
family. Proteins 41: 374 –384, 2000.
BRITTON FC, HATTON WJ, ROSSOW CF, DUAN D, HUME JR, AND HOROWITZ B. Molecular distribution of volume-regulated chloride channels (ClC-2 and ClC-3) in cardiac tissues. Am J Physiol Heart Circ
Physiol 279: H2225–H2233, 2000.
BROWN SS. Cooperation between microtubule- and actin-based motor proteins. Annu Rev Cell Dev Biol 15: 63– 80, 1999.
BURTNICK LD, KOEPF EK, GRIMES J, JONES EY, STUART DI, MCLAUGHLIN
PJ, AND ROBINSON RC. The crystal structure of plasma gelsolin:
implications for actin severing, capping, and nucleation. Cell 90:
661– 670, 1997.
BURTNICK LD, ROBINSON RC, AND CHOE S. Structure and function of
gelsolin. Res Prob Cell Differ 32: 201–211, 2001.
BUSS FC, TEMM-GROVE C, HENNING S, AND JOCHUSH B. Distribution of
profilin in fibroblasts correlates with the presence of highly dynamic actin-filaments. Cell Motil Cytoskeleton 22: 51– 61, 1992.
CALDWELL JE, HEISS SG, MERMALL V, AND COOPER JA. Effects of CapZ,
an actin capping protein of muscle, on the polymerization of actin.
Biochemistry 28: 8506 – 8514, 1989.
CAMERON LA, FOOTER MJ, VAN OUDENAARDEN A, AND THERIOT J.
Motility of ActA protein-coated microspheres driven by actin polymerization. Proc Natl Acad Sci USA 96: 4908 – 4913, 1999.
CARLIER MF, DIDRY D, ERK I, LEPAULT J, VAN TROYS ML, VANDEKERCKHOVE J, PERELROIZEN I, YIN H, DOI Y, AND PANTALONI D. T␤4 is
not a simple G-actin sequestering protein and interacts with F-actin
at high concentration. J Biol Chem 271: 9231–9239, 1996.
CARLIER MF, JEAN C, RIEGER KJ, LENFANT M, AND PANTALONI D.
Modulation of the interaction between G-actin and thymosin beta 4
by the ATP/ADP ratio: possible implication in the regulation of
actin dynamics. Proc Natl Acad Sci USA 90: 5034 –5038, 1993.
CARLIER MF, LAURENT V, SANTOLINI J, MELKI R, DIDRY D, XIA GX, HONG
Y, CHUA NH, AND PANTALONI D. Actin depolymerizing factor (ADF/
cofilin) enhances the rate of filament turnover: implication in actinbased motility. J Cell Biol 136: 1307–1322, 1997.
CARLIER MF AND PANTALONI D. Direct evidence for ADP-Pi-F-actin as
the major intermediate in ATP-actin polymerization. Rate of dissociation of Pi from actin filaments. Biochemistry 25: 211–214, 1986.
CARLIER MF AND PANTALONI D. Actin polymerization and ATP hydrolysis. Adv Biophys 76: 51–73, 1990.
CARLIER MF, PANTALONI D, EVANS JA, LAMBOOY PK, KORN ED, AND
WEBB MR. The hydrolysis of ATP that accompanies polymerization
is essentially irreversible. FEBS Lett 235: 211–214, 1988.
CARLIER MF, PANTALONI D, AND KORN ED. Evidence for an ATP cap
at the ends of actin filaments and its aggregation of the F-actin
steady state. J Biol Chem 259: 9983–9986, 1984.
CARLIER MF, PANTALONI D, AND KORN ED. The mechanism of ATP
hydrolysis accompanying the polymerization of Mg-actin and Caactin. J Biol Chem 262: 3052–3059, 1987.
CARPINTERO P, ANADON R, DIAZ-REGUEIRA S, AND GOMEZ-MARQUEZ J.
Expression of thymosin 4 messenger RNA in normal and kainatetreated rat forebrain. Neuroscience 90: 1433–1444, 1999.
CARPINTERO P, FRANCO DEL AMO F, ANADÓN R, AND GÓMEZ-MÁRQUEZ J.
Thymosin beta10 mRNA expression during early postimplantation
mouse development. FEBS Lett 394: 103–106, 1996.
CARRON E. Regulation of Wiscott-Aldrich syndrome protein and
related molecules. Curr Opin Cell Biol 14: 82– 87, 2000.
CASELLA JF, CRAIG SW, MAACK DJ, AND BROWN AE. CapZ(36/32), a
barbed end actin-capping protein, is a component of the Z-line of
skeletal muscle. J Cell Biol 105: 371–379, 1987.
CASELLA JF, MAACK DJ, AND LIN S. CapZ(36/32), a barbed end actincapping protein, is a component of the Z-line of skeletal muscle.
J Biol Chem261: 10915–10921, 1986.
CASELLA JF AND TORRES MA. Interaction of Cap Z with actin. The
467
468
DOS REMEDIOS ET AL.
79.
80.
81.
82.
83.
84.
85.
86.
87.
88.
89.
90.
91.
92.
93.
94.
95.
96.
97.
98.
99.
100.
101.
102.
Physiol Rev • VOL
103.
104.
105.
106.
107.
108.
109.
110.
111.
112.
113.
114.
115.
116.
117.
118.
119.
120.
121.
122.
123.
124.
(His321)gelsolin isolated from a flat revertant of ras-transformed
cells. Eur J Biochem 229: 615– 620, 1995.
FUJIWARA I, SUETSUGU S, UEMURA S, TAKENAWA T, AND ISHIWATA S.
Visualization and force measurement of branching by Arp2/3 complex and N-WASP in actin filament. Biochem Biophys Res Commun 293: 1550 –1555, 2002.
GALKIN VE, ORLOVA A, LUKOYANOVA N, WRIGGERS W, AND EGELMAN EH.
Actin depolymerizing factor stabilizes an existing state of F-actin
and can change the tilt of F-actin subunits. J Cell Biol 153: 75– 86,
2001.
GEISTERFER-LOWRANCE AA, KASS S, TANIGAWA G, VOSBERG HP, MCKENNA W, SEIDMAN CE, AND SEIDMAN JG. A molecular basis for
familial hypertrophic cardiomyopathy: a beta cardiac myosin heavy
chain gene missense mutation. Cell 62: 999 –1006, 1990.
GOLD JS, BAO L, GHOUSSOUB RA, ZETTER BR, AND RIMM DL. Localization and quantitation of expression of the cell motility-related
protein thymosin beta15 in human breast tissue. Mol Pathol 10:
1106 –1112, 1997.
GOLDSCHMIDT-CLERMONT PJ, FURMAN MI, WACHSSTOCK D, SAFER D,
NACHMIAS VT, AND POLLARD TD. The control of actin nucleotide
exchange by thymosin beta 4 and profilin. A potential regulatory
mechanism for actin polymerization in cells. Mol Biol Cell 3: 1015–
1024, 1992.
GOLDSCHMIDT-CLERMONT PJ, MACHESKY LM, BALDASSARE JJ, AND POLLARD TD. The actin-binding protein profilin binds to PIP2 and inhibits its hydrolysis by phospholipase C. Science 247: 1575–1578, 1990.
GOLDSCHMIDT-CLERMONT PJ, MACHESKY LM, DOBERSTEIN SK, AND POLLARD TD. The mechanism of interaction of human platelet profilin
with actin. J Cell Biol 113: 1081–1089, 1991.
GOLLA R, PHILP N, SAFER D, CHINTAPALLI J, HOFFMAN R, COLLINS L, AND
NACHMIAS VT. Co-ordinate regulation of the cytoskeleton in 3T3
cells overexpressing thymosin-␤4. Cell Motil Cytoskeleton 38: 187–
200, 1997.
GRAZI E AND TROMBETTA G. Effects of temperature on actin polymerized by Ca2⫹. Biochem J 232: 297–300, 1985.
GREMM D AND WEGNER A. Gelsolin as a calcium-regulated actin
filament-capping protein. Eur J Biochem 267: 4339 – 4345, 2000.
GUNNING P, PONTE P, BLAU H, AND KEDES L. Alpha-skeletal and
alpha-cardiac actin genes are co-expressed in adult human skeletal
muscle and heart. Mol Cell Biol 3: 1985–1995, 1983.
HALL AK. Molecular interactions between G-actin, DNase I and the
beta-thymosins in apoptosis: an hypothesis. Med Hypotheses 43:
125–131, 1994.
HALL AK. Thymosin beta-10 accelerates apoptosis. Cell Mol Biol Res
41: 167–180, 1995.
HANSON J AND LOWY J. The structure of F-actin and of actin filaments
isolated from muscle. J Mol Biol 6: 46 – 60, 1963.
HART MC, KORSHUNOVA YO, AND COOPER JA. Vertebrates have conserved capping protein alpha isoforms with specific expression
patterns. Cell Motil Cytoskeleton 38: 120 –132, 1997.
HATANAKA H, OGURA K, MORIYAMA K, ICHIKAWA S, YAHARA I, AND
INAGAKI F. Tertiary structure of destrin and structural similarity
between 2 actin-regulating protein families. Cell 85: 1047–1055,
1996.
HAWKINS M, POPE B, MACIVER SK, AND WEEDS AG. Human actin
depolymerizing factor mediates a pH-sensitive destruction of actin
filaments. Biochemistry 32: 9985–9993, 1993.
HAYDEN SM, MILLER PS, BRAUWEILER A, AND BAMBURG JR. Analysis of
the interactions of actin depolymerizing factor with G- and F-actin.
Biochemistry 32: 9994 –10004, 1993.
HEGYI G, SZILAGYI L, AND BELAGYI J. Influence of the bound nucleotide on the molecular dynamics of actin. Eur J Biochem 175:
271–274, 1988.
HEINKE MY, WHEELER CH, YAN JX, AMIN V, CHANG D, EINSTEIN R,
DUNN MJ, AND DOS REMEDIOS CG. Changes in myocardial protein
expression in pacing-induced canine heart failure. Electrophoresis
20: 2086 –2093, 1999.
HEISS SG AND COOPER JA. Regulation of CapZ, an actin-capping
protein of chicken muscle, by anionic phospholipids. Biochemistry
30: 8753– 8758, 1991.
HELING A, ZIMMERMANN R, KOSTIN S, MAENO Y, HEIN S, DEVAUX B,
BAUER E, KLOVEKORN WP, SCHLEPPER M, SCHAPER W, AND SCHAPER J.
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
SAFER D. Thymosin-beta(4) changes the conformation and
dynamics of actin monomers. Biophys J 78: 2516 –2527, 2000.
DE LA CRUZ EM AND POLLARD TD. Nucleotide-free actin: stabilization
by sucrose and nucleotide binding kinetics. Biochemistry 34: 5452–
5461, 1995.
DANCKER P, LÖW I, HASSELBACK W, AND WIELAND T. Interaction of
actin with phalloidin: polymerization and stabilization of F-actin.
Biochim Biophys Acta 400: 407– 414, 1975.
DEDOVA I, DEDOV V, NOSWORTHY NJ, HAMBLY BD, AND DOS REMEDIOS
CG. Cofilin and DNase I affect the conformation of the small
domain of actin. Biophys J 82: 3134 –3143, 2002.
DEHOSTOS EL, BRADTKE B, LOTTSPEICH F, AND GERISCH G. Coactosin,
a 17 kDa F-actin binding protein from Dictyostelium discoideum.
Cell Motil Cytoskeleton 26: 181–191, 1993.
DE TOMBE PP. Altered contractile function in heart failure. Cardiovasc Res 37: 367–380, 1998.
DEVINENI N, MINAMIDE LS, NIU M, SAFER D, VERMA R, BAMBURG JR,
AND NACHMIAS VT. A quantitative analysis of G-actin binding proteins and the G-actin pool in developing chick brain. Brain Res 823:
129 –140, 1999.
DIDRY D, CARLIER MF, AND PANTALONI D. Synergy between actin
depolymerizing factor cofilin and profilin in increasing actin filament turnover. J Biol Chem 273: 25602–25611, 1998.
DIEFFENBACH CW, SENGUPTA DN, KRAUSE D, SAWZAK D, AND SILVERMAN
RH. Cloning of murine gelsolin and its regulation during differentiation of embryonal carcinoma cells. J Biol Chem 264: 13281–
13288, 1989.
DOS REMEDIOS CG AND DICKENS MJ. Actin microcrystals and tubes
formed in the presence of gadolinium ions. Nature 276: 731–733,
1978.
DOS REMEDIOS CG AND THOMAS DD. (Editors). Molecular Interactions of Actin. Actin Structure and Actin-Binding Proteins. Heidelberg, Germany: Springer-Verlag, 2001, p. 32.
DU J AND FRIEDEN C. Kinetic studies on the effect of yeast cofilin on
yeast actin polymerization. Biochemistry 37: 13276 –13284, 1998.
EDWARDS DC AND GILL GN. Structural features of LIM kinase that
control effects on the actin cytoskeleton. J Biol Chem 274: 11352–
11361, 1999.
EDWARDS KA, MONTAGUE RA, SHEPARD S, EDGAR BA, ERIKSON RL, AND
KIEHART DP. Identification of Drosophila cytoskeletal proteins by
induction of abnormal cell shape in fission yeast. Proc Natl Acad
Sci USA 91: 4589 – 4593, 1994.
ELLIOTT K, WATKINS H, AND REDWOOD CS. Altered regulatory properties of human cardiac troponin I mutants that cause hypertrophic
cardiomyopathy. J Biol Chem 275: 22069 –22074, 2000.
ERICKSON-VITANEN S, RUGGIERI S, NATALINI P, AND HORECKER BL.
Thymosin beta 10, a new analog of thymosin beta 4 in mammalian
tissues. Arch Biochem Biophys 22: 570 –576, 1983.
ESTES JE, SELDEN LA, KINOSIAN HJ, AND GERSHMAN LC. Tightly-bound
divalent cation of actin. J Muscle Res Cell Motil 13: 272–284, 1992.
FEDOROV AA, LAPPALAINEN P, FEDOROV EV, DRUBIN DG, AND ALMO SC.
Structure determination of yeast cofilin. Nat Struct Biol 4: 366 –369,
1997.
FEINBERG J, MERY J, HEITZ F, BENYAMIN Y, AND ROUSTAN C. Correlations between biological activity and structural properties for two
short homologous sequences in thymosin beta4 and gelsolin. Int J
Pept Protein Res 47: 62– 69, 1996.
FOWLER VM AND CONLEY CA. Tropomodulin. In: Guidebook to the
Cytoskeletal and Motor Proteins, edited by Kreis T and Vale R.
Oxford, UK: Oxford Univ. Press, 1999, p. 154 –159.
FOWLER VM, SUSSMAN MA, MILLER PG, FLUCHER BE, AND DANIELS PM.
Tropomodulin is associated with the free (pointed) ends of the thin
filaments in rat skeletal muscle. J Cell Biol 120: 411– 420, 1993.
FOWLER WE AND AEBI U. Polymorphism of actin paracrystals induced by polylysine. J Cell Biol 93: 452– 458, 1982.
FRANKEL S, CONDEELIS J, AND LEINWAND L. Expression of actin in
Escherichia coli. J Biol Chem 265: 17980 –17987, 1990.
FUJISAWA T, KOSTYUKOVA A, AND MAEDA Y. The shapes and sizes of
two domains of tropomodulin, the P-end-capping protein of actintropomyosin. FEBS Lett 498: 67–71, 2001.
FUJITA H, LAHAM LE, JANMEY PA, KWIATKOWSKI DJ, STOSSEL TP, BANNO
Y, NOZAWA Y, MULLAUER L, ISHIZAKI A, AND KUZUMAKI N. Functions of
AND
ACTIN AND ACTIN BINDING PROTEINS
125.
126.
127.
128.
129.
131.
132.
133.
134.
135.
136.
137.
138.
139.
140.
141.
142.
143.
144.
145.
Physiol Rev • VOL
146. KINOSIAN HJ, SELDEN LA, ESTES JE, AND GERSHMAN LC. Kinetics of
gelsolin interaction with phalloidin-stablized F-actin. Rate constants for binding and severing. Biochemistry 35: 16550 –16556,
1996.
147. KINOSIAN HJ, SELDEN LA, ESTES JE, AND GERSHMAN LC. Nucleotide
binding to actin. Cation dependence of nucleotide dissociation and
exchange. J Biol Chem 268: 8683– 8691, 1993.
148. KOFFER A, EDGAR AJ, AND BAMBURG JR. Identification of two species
of actin depolymerizing factor in cultures of BHK cells. J Muscle
Res Cell Motil 9: 320 –328, 1988.
149. KORN ED, CARLIER MF, AND PANTALONI D. Actin polymerization and
ATP hydrolysis. Science 238: 638 – 644, 1987.
150. KOTHAKOTA S, AZUMA T, REINHARD C, KLIPPEL A, TANG J, CHU K,
MCGARRY TJ, KIRSCHNER MW, KOTHS K, KWIATKOWSKI DJ, AND WILLIAMS LT. Caspase-3-generated fragment of gelsolin: effector of
morphological change in apoptosis. Science 278: 294 –298, 1997.
151. KREIS T AND VALE R. (Editors). Guidebook to the Cytoskeletal and
Motor Proteins (2nd ed.). Oxford, UK: Oxford Univ. Press, 1999.
152. KREUDER V, DIECKHOFF J, SITTIG M, AND MANNHERZ HG. Isolation,
characterisation and crystallization of deoxyribonuclease I from
bovine and rat parotid gland and its interaction with rabbit skeletal
muscle actin. Eur J Biochem 139: 389 – 400, 1984.
153. KRIEGER I, KOSTYUKOVA AS, YAMASHITA A, NITANAI Y, AND MAEDA Y.
Crystal structure of the C-terminal half of tropomodulin and structural basis of actin filament pointed-end capping. Biophys J 83:
2716 –2725, 2002.
154. KUSANO K, ABE H, AND OBINATA T. Detection of a sequence involved
in actin-binding in the N-terminal side of cofilin. Mol Cell Biochem
190: 133–141, 1999.
155. KWIATECK O, PAPA II, LEBART MC, AND ROUSTAN C. Interaction of actin
with the capping protein, CapZ from sea bass (Dicentrarchus
labrx) white skeletal muscle. Comp Biochem Physiol B Comp
Biochem 127: 551–562, 2000.
156. KWIATKOWSKI DJ. Functions of gelsolin: motility, signaling, apoptosis, cancer. Curr Opin Cell Biol 11: 103–108, 1999.
157. KWIATKOWSKI DJ, STOSSEL TP, ORKIN SH, MOLE JE, COLTEN H, AND YIN
HL. Plasma and cytoplasmic gelsolins are encoded by a single gene
and contain a duplicate actin-binding domain. Nature 323: 455– 458,
1986.
158. LAPPALAINEN P, FEDOROV EV, FEDOROV AA, ALMO SC, AND DRUBIN DG.
Essential functions and actin-binding surfaces of yeast cofilin revealed by systematic mutagenesis. EMBO J 16: 5520 –5530, 1997.
159. LAZARIDES E AND LINDBERG U. Actin is the naturally occurring inhibitor of deoxyribonuclease I. Proc Natl Acad Sci USA 71: 4724 – 4746,
1974.
160. LECHLER T, JONSDOTTIR GA, KLEE SK, PELLMAN D, AND LI R. A twotiered mechanism by which Cdc42 controls the localization and
activation of an Arp2/3-activating motor complex in yeast. J Cell
Biol 155: 261–270, 2001.
161. LEONARD SA, GITTIS AG, PETRELLA EC, POLLARD TD, AND LATTMAN EE.
Crystal structure of the actin-binding protein actophorin from
Acanthamoeba. Nat Struct Biol 4: 369 –373, 1997.
162. LIAO TH. Bovine pancreatic deoxyribonuclease D. J Biol Chem 249:
2354 –2356, 1974.
163. LIN KM, MEJILLANO M, AND YIN HL. Ca2⫹ regulation of gelsolin by its
C-terminal tail. J Biol Chem 275: 27746 –27752, 2000.
164. LITTLEFIELD R, ALMENAR-QUERALT A, AND FOWLER VM. Actin dynamics
at pointed ends regulates thin filament length in striated muscle.
Nat Cell Biol 3: 544 –551, 2001.
165. LITTLEFIELD R AND FOWLER VM. Defining actin filament length in
striated muscle: rulers and caps or dynamic instability? Annu Rev
Cell Dev Biol 14: 487–525, 1998.
166. LOISEL TP, BOUJEMAA R, PANTALONI D, AND CARLIER MF. Reconstitution of actin-based motility of Listeria and Shigella using pure
proteins. Nature 401: 613– 616, 1999.
167. LORENZ M, POPP D, AND HOLMES KC. Refinement of the F-actin model
against X-ray by the use of a direction mutation algorithm. J Mol
Biol 234: 826 – 836, 1993.
168. LOW TL, HU SK, AND GOLDSTEIN AL. Complete amino acid sequence
of bovine thymosin beta 4: a thymic hormone that induces terminal
deoxynucleotidyl transferase activity in thymocyte populations.
Proc Natl Acad Sci USA 78: 1162–1166, 1981.
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
130.
Increased expression of cytoskeletal, linkage, and extracellular
proteins in failing human myocardium. Circ Res 86: 846 – 853, 2000.
HERRMANNSDOERFER AJ, HEEB PJ, FEUSTEL PJ, ESTES JE, KEENAN CJ,
MINNEAR FL, SELDEN L, GIUNTA C, FLOR JR, AND BLUMENSTOCK FA.
Vascular clearance and organ uptake of G- and F-actin in the rat.
Am J Physiol Gastrointest Liver Physiol 265: G1071–G1081, 1993.
HIGGS HN AND POLLARD TD. Regulation of actin polymerization by
Arp2/3 complex and WASp/Scar proteins. J Biol Chem 274: 325531–
32534, 1999.
HITCHCOCK SE. Actin DNase I interaction. Depolymerization and
nucleotide exchange. J Biol Chem 255: 5668 –5673, 1980.
HOLMES KC, POPP D, GEBHARD W, AND KABSCH W. Atomic model of
the actin filament. Nature 347: 44 – 49, 1990.
HUFF T, ZERZAWY D, AND HANNAPPEL E. Interactions of beta-thymosins, thymosin beta 4-sulfoxide, and N-terminally truncated thymosin beta 4 with actin studied by equilibrium centrifugation,
chemical cross-linking and viscometry. Eur J Biochem 230: 650 –
657, 1995.
HUG C, MILLER TM, TORRES MA, CASELLA JF, AND COOPER JA. Identification and characterization of an actin-binding site of CapZ. J Cell
Biol 116: 923–931, 1992.
HUXLEY HE. Electron microscope studies on the structure of natural
and synthetic protein filaments from striated muscle. J Mol Biol 7:
281–308, 1963.
ICHETOVKIN I, HAN J, PANG KM, KNECHT DA, AND CONDEELIS JS. Actin
filaments are severed by both native and recombinant Dictyostelium cofilin but to different extents. Cell Motil Cytoskeleton 45:
293–306, 2000.
ISENBERG G. Cytoskeleton Proteins: A Purification Manual. Berlin:
Springer-Verlag, 1995.
JEAN C, RIEGER K, BLANCHOIN L, CARLIER MF, LENFANT M, AND PANTALONI D. Interaction of G-actin with thymosin beta 4 and its
variants thymosin beta 9 and thymosin beta met9. J Muscle Res Cell
Motil 15: 278 –286, 1994.
JIANG L, TSUBAKIHARA M, HEINKE MY, YAO M, DUNN NJ, PHILLIPS W,
DOS REMEDIOS CG, AND NOSWORTHY NJ. Heart failure and apoptosis:
electrophoretic methods support data from micro- and macroarrays. A critical review of genomics and proteomics. Proteomics 1:
1481–1488, 2001.
JONES LJF, CARBALLIDO-LÓPEZ R, AND ERRINGTON J. Control of cell
shape in bacteria: helical, actin-like filaments in Bacillus subtilis.
Cell 104: 913–922, 2001.
KABSCH W, MANNHERZ HG, SUCK D, PAI EF, AND HOLMES KC. Atomic
structure of the actin-DNase I complex. Nature 347: 37– 44, 1990.
KAISER DA AND POLLARD TD. Characterization of actin and poly-Lproline binding sites of Acanthamoeba profilin with monoclonal
antibodies and by mutagenesis. J Mol Biol 256: 89 –107, 1996.
KANGAS H, ULMANEN I, PAUNIO T, KWIATKOWSKI DJ, LEHTOVIRTA M,
JALANKO A, AND PELTONEN L. Functional consequences of amyloidosis mutation for gelsolin polypeptide: analysis of gelsolin-actin
interaction and gelsolin processing in gelsolin knock-out fibroblasts. FEBS Lett 454: 233–239, 1999.
KASAI M, ASAKURA S, AND OOSAWA F. The cooperative nature of the
G-F transformation of actin. Biochim Biophys Acta 57: 22–31, 1962.
KASAI M, NAKANO E, AND OOSAWA F. Polymerization of actin free
from nucleotides and divalent cations. Biochim Biophys Acta 94:
494 –503, 1965.
KEKIC M, NOSWORTHY NJ, DEDOVA I, COLLYER CA, AND DOS REMEDIOS
CG. Regulation of the cytoskeleton assembly: a role for a ternary
complex with two actin-binding proteins. Res Prob Cell Differ 32:
165–180, 2001.
KELLEHER JF, ATKINSON SJ, AND POLLARD TD. Sequences, structural
models, and cellular localization of the actin-related proteins Arp2
and Arp3 from Acanthamoeba. J Cell Biol 131: 385–397, 1995.
KIMURA A, HARADA H, PARK JE, NISHI H, SATOH M, TAKAHASHI M, HIROI
S, SASAOKA T, OHBUCHI N, NAKAMURA KOYANAGI T, HWANG THI, CHOO
JA, CHUNG KS, HASEGAWA A, NAGAI R, OKAZAKI O, NAKAMURA H,
MATSUZAKI M, SAKAMOTO T, TOSHIMA H, KOGA Y, IMAIZUMI T, AND
SASAZUKI T. Mutations in the cardiac troponin I gene associated
with hypertrophic cardiomyopathy. Nat Genet 16: 379 –382, 1997.
KINOSIAN HJ, SELDEN LA, ESTES JE, AND GERSHMAN LC. Actin filament
annealing in the presence of ATP and phalloidin. Biochemistry 32:
12353–12357, 1993.
469
470
DOS REMEDIOS ET AL.
Physiol Rev • VOL
194.
195.
196.
197.
198.
199.
200.
201.
202.
203.
204.
205.
206.
207.
208.
209.
210.
211.
212.
213.
214.
215.
216.
the actin-monomer subunit interface to prevent polymerization.
Nat Cell Biol 2: 376 –379, 2000.
MULLENS RD, STAFFORD WB, AND POLLARD TD. Structure subunit
topology and actin-binding activity of the Arp2/3 complex from
Acanthamoeba. J Cell Biol 136: 331–343, 1997.
MULLINS RD, HEUSER JA, AND POLLARD TD. The interaction of Arp2/3
complex with actin: nucleation, high affinity pointed end capping,
and formation of branching networks of filaments. Proc Natl Acad
Sci USA 95: 6181– 6186, 1998.
NACHMIAS VT. Small actin-binding proteins: the beta-thymosin family. Curr Opin Cell Biol 5: 56 – 62, 1993.
NAGAOKA R, ABE H, KUSANO K, AND OBINATA T. Concentration of
cofilin, a small actin-binding protein, at the cleavage furrow during
cytokinesis. Cell Motil Cytoskeleton 30: 1–7, 1995.
NAGAOKA R, ABE H, AND OBINATA T. Site-directed mutagenesis of the
phosphorylation site of cofilin: its role in cofilin-actin interaction
and cytoplasmic localization. Cell Motil Cytoskeleton 35: 200 –209,
1996.
NAGAOKA R, MINAMI N, HAYAKAWA K, ABE H, AND OBINATA T. Quantitative analysis of low molecular weight G-actin-binding proteins,
cofilin, ADF and profilin, expressed in developing and degenerating
chicken skeletal muscles. J Muscle Res Cell Motil 17: 463– 473,
1996.
NEIDL C AND ENGEL J. Exchange of ADP, ATP and 1,N-ethenoadenosine 5⬘-triphosphate on G-actin. Equilibrium and kinetics. Eur
J Biochem 101: 163–169, 1979.
NEWMAN J, ZANER KL, SCHICK KL, GERSHMAN LC, SELDEN LA, KINOSIAN
HG, TRAVIS JL, AND ESTES JE. Nucleotide exchange and rheometric
studies with F-actin prepared from ATP- or ADP-monomeric actin.
Biophys J 64: 1559 –1566, 1993.
NISHIDA E. Opposite effects of cofilin and profilin from porcine
brain on rate of exchange of actin-bound adenosine 5⬘-triphosphate. Biochemistry 24: 1160 –1164, 1985.
NISHIDA E, IIDA K, YONEZAWA N, KOYASU S, YAHARA I, AND SAKAI H.
Cofilin is a component of intranuclear and cytoplasmic actin rods
induced in cultured cells. Proc Natl Acad Sci USA 84: 5262–5266,
1987.
NIWA R, NAGATA-OHASHI K, TAKEICHI M, MIZUNO K, AND UEMURA T.
Control of actin reorganization by Slingshot, a family of phosphatases that dephosphorylate ADF/cofilin. Cell 108: 233–246, 2002.
NOSWORTHY NJ, KEKIC M, AND DOS REMEDIOS CG. The affinity of chick
cofilin for actin increases when actin is complexed with DNase I:
formation of a cofilin-actin-DNase I ternary complex. Proteomics 1:
1513–1519, 2001.
OBINATA T, NAGAOKA-YASUDA R, ONO S, KUSANO K, OHTAKA Y, YAMASHIRO S, OKAADA K, AND ABE H. Low molecular weight G-actin
binding proteins involved in the regulation of actin assembly during
myofibrillogenesis. Cell Struct Funct 22: 181–189, 1997.
ODA T, MAKINO K, YAMASHITA I, NAMBA K, AND MAEDA Y. The helical
parameters of F-actin precisely determined from X-ray fiber diffraction of well-oriented gels. Res Prob Cell Differ 32: 43–58, 2001.
OGAWA Y, TASHIMA M, YUMOTO Y, OKUDA SAWADA T, H, OKUMA M, AND
MARUYAMA Y. Coding sequence of human placenta cofilin cDNA.
Nucleic Acid Res 18: 7169, 1990.
OLSON TM, MICHELS VV, THIBODEAU SN, TAI YS, AND KEATING MT.
Actin mutations in dilated cardiomyopathy, a heritable form of
heart failure. Science 280: 750 –752, 1998.
OOSAWA F AND ASAKURA S. Thermodynamics of the Polymerization
of Protein. New York: Academic, 1975.
OOSAWA F, ASAKURA S, HOTTA K, AND OOI T. G-F transformation of
actin as a fibrous condensation. J Polymer Sci 37: 323–326, 1959.
ORIOL C, DUBORD C, AND LANDON F. Crystallization of native striatedmuscle actin. FEBS Lett 73: 89 –91, 1977.
ORLOVA A, CHEN X, RUBENSTEIN PA, AND EGELMAN EH. Modulation of
yeast F-actin structure by a mutation in the nucleotide-binding
cleft. J Mol Biol 271: 235–243, 1997.
ORLOVA A AND EGELMAN EH. Structural dynamics of F-actin. I.
Changes in the C terminus. J Mol Biol 245: 582–597, 1995.
OSBORNE M AND WEBER K. (Editors). Cytoskeletal Proteins in Tumor Diagnosis. Current Communications. Cold Spring Harbor,
NY: Cold Spring Harbor, 1989.
OTTERBEIN L, GRACEFFA P, AND DOMINGUEZ R. The crystal structure of
G-actin (Abstract). Biophys J 80: 172a, 2001.
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
169. LU M, WITKE W, KWIATKOWSKI DJ, AND KOSIK KS. Delayed retraction
of filopodia in gelsolin null mice. J Cell Biol 138: 1279 –1287, 1997.
170. LUECK A, YIN HL, KWIATKOWSKI DJ, AND ALLEN PG. Calcium regulation
of gelsolin and adseverin: a natural test of the helix latch hypothesis. Biochemistry 39: 5274 –5279, 2000.
171. MACHESKY LM AND MAY RC. Arps: actin-related proteins. Res Prob
Cell Differ 32: 213–229, 2001.
172. MACIVER SK, TERNENT D, AND MCLAUGHLIN PJ. Domain 2 of gelsolin
binds directly to tropomyosin. FEBS Lett 473: 71–75, 2000.
173. MAEKAWA S, NISHIDA E, OHTA Y, AND SAKAI H. Isolation of low
molecular weight actin-binding proteins from porcine brain. J Biochem 95: 377–385, 1984.
174. MALINDA KM, GOLDSTEIN AL, AND KLEINMAN HK. Thymosin 4 stimulates directional migration of human umbilical vein endothelial
cells. FASEB J 11: 474 – 481, 1997.
175. MANIOTIS AJ, CHEN S, AND INGBER DE. Demonstrations of mechanical
connections between integrins, cytoskeletal filaments, and nucleoplasm that stabilize nuclear structure. Proc Natl Acad Sci USA 94:
847– 854, 1997.
176. MANNHERZ HG, KABSCH W, AND LEBERMAN J. Crystals of skeletal
muscle actin: pancreatic DNase I complex. FEBS Lett 73: 141–143,
1977.
177. MANNHERZ HG, GOODY RS, KONRAD M, AND NOWAK E. The interaction
of bovine pancreatic deoxyribonuclease I and skeletal muscle actin. Eur J Biochem 104: 367–379, 1980.
178. MARSTON SB AND HODGKINSON JL. Cardiac and skeletal myopathies:
can genotype explain phenotype? J Muscle Res Cell Motil 22: 1– 4,
2001.
179. MARUYAMA K, KUROKAWA H, OOSAWA M, SHIMOKA S, YAMAMOTO H, ITO
M, AND MARUYAMA K. Beta-actinin is equivalent to CapZ protein.
J Biol Chem 265: 8712– 8715, 1990.
180. MATSUDAIRA P, JAKES R, AND WALKER JE. A gelsolin-like Ca2⫹-dependent actin-binding domain in villin. Nature 315: 248 –250, 1985.
181. MCGOUGH A, CHIU W, AND WAY M. Determination of the gelsolin
binding site on F-actin: implications for severing and capping.
Biophys J 74: 764 –772, 1998.
182. MCGOUGH A, POPE B, CHIU W, AND WEEDS A. Cofilin changes the
twist of F-actin: implications for actin filament dynamics and cellular function. J Cell Biol 138: 771–781, 1997.
183. MCGOUGH A, POPE B, AND WEEDS A. The ADF/cofilin family: acceleration of actin reorganization. Res Prob Cell Differ 32: 135–154,
2001.
184. MCGRATH JL, OSBORN EA, TARDY YS, DEWEY CF JR, AND HARTWIG JH.
Regulation of the actin cycle in vivo by actin filament severing. Proc
Natl Acad Sci USA 97: 6532– 6537, 2000.
185. MCKIM KS, MATHESON C, MARRA MA, WAKARCHUK MF, AND BAILLIE DL.
The Caenorhabditis elegans Unc-60 gene encodes proteins homologous to a family of actin-binding proteins. Mol Gen Genet 242:
346 –357, 1994.
186. MCLAUGHLIN PJ, GOOCH JT, MANNHERZ HG, AND WEEDS AG. Structure
of gelsolin segment-1-actin complex and the mechanism of filament
severing. Nature 364: 685– 692, 1993.
187. MIZUNO K, OKANO I, OHASHI K, NUNOUE K, KUMA K, MIYATA T, AND
NAKAMURA T. Identification of a human cDNA clone encoding a
novel protein kinase with two repeats of the LIM/double zinc finger
motif. Oncogene 9: 1605–1612, 1994.
188. MOORE PB, HUXLEY HE, AND DEROSIER DJ. Three-dimensional reconstruction of F-actin, thin filaments and decorated thin filaments. J
Mol Biol 50: 279 –295, 1970.
189. MOORE S. Pancreatic DNase. Enzymes 14: 281–296, 1981.
190. MORGAN TE, LOCKERBIE RO, MINAMIDE LS, BROWNING MD, AND BAMBURG JR. Isolation and characterization of a regulated form of actin
depolymerizing factor. J Cell Biol 122: 623– 633, 1993.
191. MORIYAMA K, NISHIDA E, YONEZAWA N, SAKAI H, MATSUMOTO S, IIDA K,
AND YAHARA I. Destrin, a mammalian actin-depolymerizing protein,
is closely related to cofilin. Cloning and expression of porcine brain
destrin cDNA. J Biol Chem 265: 5768 –5773, 1990.
192. MORIYAMA K, YONEZAWA N, SAKAI H, YAHARA I, AND NISHIDA E. Mutational analysis of an actin-binding site of cofilin and characterization of chimeric proteins between cofilin and destrin. J Biol Chem
267: 7240 –7244, 1992.
193. MORTON WM, AYSCOUGH KR, AND MCLAUGHLIN PJ. Latrunculin alters
ACTIN AND ACTIN BINDING PROTEINS
Physiol Rev • VOL
240.
241.
242.
243.
244.
245.
246.
247.
248.
249.
250.
251.
252.
253.
254.
255.
256.
257.
258.
259.
260.
261.
262.
ternary complex of DNase I, actin and thymosin beta4. FEBS Lett
387: 132–136, 1996.
RESSAD F, DIDRY D, XIA GX, HONG Y, CHUA NH, PANTALONI D, AND
CARLIER MF. Kinetic analysis of the interaction of actin-depolymerizing factor (ADF)/cofilin with G- and F-actins. J Biol Chem 273:
20894 –20902, 1998.
RICKARD JE AND SHETERLINE P. Cytoplasmic concentrations of inorganic phosphate affect the critical concentrations for actin assembly in the presence of cytochalasin D or ADP. J Mol Biol 191:
273–280, 1986.
RIDLEY A. What initiates actin polymerization? Genome Biol 1: 1– 6,
2000.
ROBINSON RC, MEJILLANO M, LE VP, BURTNICK LD, YIN HL, AND CHOE
S. Domain movement in gelsolin: a calcium-activated switch. Science 286: 1939 –1942, 1999.
ROBINSON RC, TURBEDSKY K, KAISER DA, MARCHAND JB, HIGGS HN,
CHOE S, AND POLLARD TD. Crystal structure of Arp2/3 complex.
Science 294: 1679 –1684, 2001.
RODAL AA, TRETREAULT JW, LAPPALAINEN P, DRUBIN DG, AND AMBERG
DC. Aip 1p interacts with cofilin to disassemble actin filaments.
J Cell Biol 145: 1251–1264, 1999.
RUBENSTEIN PA. The functional importance of multiple actin isoforms. Bioessays 12: 309 –315, 1990.
SAFER D AND CHOWRASHI PK. Beta-thymosins from marine invertebrates: primary structure and interaction with actin. Cell Motil
Cytoskeleton 38: 163–171, 1997.
SAFER D, ELZINGA M, AND NACHMIAS VT. Thymosin beta 4 and Fx, an
actin-sequestering peptide, are indistinguishable. J Biol Chem 266:
4029 – 4032, 1991.
SAFER D AND NACHMIAS VT. ␤ Thymosins as actin binding peptides.
Bioessays16: 473– 479, 1994.
SAFER D AND NACHMIAS VT. Thymosin ␤4. In: Guidebook to the
Cytoskeletal and Motor Proteins, edited by Kreis T and Vale R.
Oxford, UK: Oxford Univ. Press, 1999, p. 149 –151.
SAFER D, SOSNICK TR, AND ELZINGA M. Thymosin beta 4 binds actin
in an extended conformation and contacts both the barbed and
pointed ends. Biochemistry 36: 5806 –5816, 1997.
SAKIYAMA S, FUJIMURA S, AND SAKIYAMA H. Absence of gamma-isoactin expression in the mouse fibroblast cell line. J Biol Chem 256:
31–33, 1981.
SATOH M, TAKAHASHI M, SAKAMOTO T, HIROE M, MARUMO F, AND
KIMURA A. Structural analysis of the titin gene in hypertrophic
cardiomyopathy: identification of a novel disease gene. Biochem
Biophys Res Commun 262: 411– 417, 1999.
SCHAFER DA AND COOPER JA. Control of actin assembly at filament
ends. Annu Rev Cell Dev Biol 11: 497–518, 1995.
SCHAFER DA, KORSHUNOVA YO, SCHROER TA, AND COOPER JA. Differential localization and sequence analysis of capping protein betasubunit isoforms of vertebrates. J Cell Biol 127: 453– 465, 1994.
SCHEVZOV G, LLOYD C, HAILSTONES D, AND GUNNING P. High level
expression of transfected beta- and gamma-actin genes differentially impacts on myoblasts cytoarchitecture. J Cell Biol 117: 775–
785, 1993.
SCHOEPPER B AND WEGNER A. Rate constants and equilibrium constants for binding of actin to the 1:1 gelsolin-actin complex. Eur
J Biochem 202: 1127–1131, 1991.
SCHUTT CE, LINDBERG U, MYSLIK J, AND STRAUSS N. Molecular packing
in profilin:actin crystals and its implications. J Mol Biol 209: 735–
746, 1989.
SCHUTT CE, MYSLIK JC, ROZYCKI MD, GOONESKERE N, AND LINDBERG U.
The structure of crystalline profilin-beta-actin. Nature 365: 810 –
816, 1993.
SEGURA M AND LINDBERG U. Separation of non-muscle isoactins in
the free form or as profilin complexes. J Biol Chem 259: 3949 –3954,
1984.
SELDEN LA, ESTES JA, AND GERSHMAN LC. High affinity divalent
cation binding to actin. Effect of low affinity salt binding. J Biol
Chem 264: 9271–9277, 1989.
SELDEN LA, KINOSIAN HJ, NEWMAN J, LINCOLN B, HURWITZ C, GERSHMAN
LC, AND ESTES JE. Severing of F-actin by the amino-terminal half of
gelsolin suggests internal cooperativity in gelsolin. Biophys J 75:
3092–3090, 1998.
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
217. OTTERBEIN L, GRACEFFA P, AND DOMINGUEZ R. The crystal structure of
uncomplexed actin in the ADP state. Science 293: 708 –711, 2001.
218. OWEN C AND DEROSIER D. A 13-angstrom map of the actin-scruin
filament from the Limulus acrosomal process. J Cell Biol 123:
337–344, 1993.
219. PANTALONI D AND CARLIER MF. How profilin promotes actin filament
assembly in the presence of thymosin beta 4. Cell 75: 1007–1014,
1993.
220. PANTALONI D, LE CLAINCHE C, AND CARLIER MF. Mechanism of actinbased motility. Science 292: 1502–1506, 2001.
221. PARDEE JD, SIMPSON PA, STRYER L, AND SPUDICH JA. Actin filaments
undergo limited exchange at physiological salt conditions. J Cell
Biol 94: 316 –324, 1982.
222. PAUDEL HK AND LIAO TH. Comparison of the three primary structures of deoxyribonuclease isolated from bovine, ovine, and porcine pancreas. Derivation of the amino acid sequence of ovine
DNase and revision of the previously published amino acid sequence of bovine DNase. J Biol Chem 261: 16012–16017, 1986.
223. PEITSCH MC, POLZAR B, STEPHAN H, CROMPTON T, MACDONALD HR,
MANNHERZ HG, AND TSCHOPP J. Characterization of the endogenous
deoxyribonuclease involved in nuclear DNA degradation during
apoptosis (programmed cell death). EMBO J 12: 371–377, 1993.
224. PERELROIZEN I, CARLIER MF, AND PANTALONI D. Binding of divalent
cation and nucleotide to G-actin in the presence of profilin. J Biol
Chem 270: 1501–1508, 1995.
225. PERELROIZEN I, DIDRY D, CHRISTENSEN H, CHUA NH, AND CARLIER MF.
Role of nucleotide exchange and hydrolysis in the function of
profilin in actin assembly. J Biol Chem 271: 12302–12309, 1996.
226. POETTER K, JIANG H, HASSANZADEH S, MASTER SR, CHANG A, DALAKAS
MC, RAYMENT I, SELLERS JR, FANANAPAZIR L, AND EPSTEIN ND. Mutations in either the essential or regulatory light chains of myosin are
associated with a rare myopathy in human heart and skeletal
muscle. Nat Genet 13: 63– 69, 1996.
227. POLLARD TD. Polymerization of ADP-actin. J Cell Biol 99: 769 –777,
1984.
228. POLLARD TD. Rate constants for the reactions of ATP- and ADPactin with the ends of actin filaments. J Cell Biol 103: 2747–2754,
1986.
229. POLLARD TD. Introduction. In: Guidebook to the Cytoskeletal and
Motor Proteins, edited by Kreis T and Vale R. Oxford, UK: Oxford
Univ. Press, 1999, p. 3–11.
230. POLLARD TD. Profilins. In: Guidebook to the Cytoskeletal and Motor
Proteins, edited by Kreis T and Vale R. Oxford, UK: Oxford Univ.
Press, 1999, p. 117–120.
231. POLLARD TD AND COOPER JA. Actin and actin-binding proteins: a
critical evaluation of mechanisms and function. Annu Rev Biochem
55: 987–1035, 1986.
232. POPE B, MACIVER S, AND WEEDS A. Localization of the calciumsensitive actin monomer binding site in gelsolin to segment 4 and
identification of calcium binding sites. Biochemistry 34: 1583–1588,
1995.
233. POPP D, LEDNEV VV, AND JAHN W. Methods of preparing well oriented
sols of F-actin containing filaments suitable for X-ray diffraction. J
Mol Biol 197: 679 – 684, 1987.
234. PUIUS YA, FEDOROV EV, EICHINGER L, SCHLEICHER M, AND ALMO SC.
Mapping the functional surface of domain 2 in the gelsolin superfamily. Biochemistry 39: 5322–5331, 2000.
235. QUINLAN R, HUTCHISON C, AND LANE B (Editors). Intermediate Filament Proteins (3rd ed.). Oxford, UK: Oxford Univ. Press. In press.
236. QUIRK S, MACIVER SK, AMPE C, DOBERSTEIN SK, KAISER DA, VANDAMME
J, VANDEKERCKHOVE JS, AND POLLARD TD. Primary structure of and
studies on Acanthamoeba actophorin. Biochemistry 32: 8525–
8533, 1993.
237. RATNASWAMY G, KOEPF E, BEKELE H, YIN HL, AND KELLY JW. The
amyloidogenicity of gelsolin is controlled by proteolysis and pH.
Chem Biol 6: 293–304, 1999.
238. REDWOOD C, LOHMANN K, BING W, ESPOSITO GM, ELLIOTT K, ABDULRAZZAK H, KNOTT A, PURCELL I, MARSTON S, AND WATKINS H. Investigation of a truncated cardiac troponin T that causes familial hypertrophic cardiomyopathy: Ca2⫹ regulatory properties of reconstituted thin filaments depend on the ratio of mutant to wild-type
protein. Circ Res 86: 1146 –1152, 2000.
239. REICHERT A, HEINTZ D, ECHNER H, VOELTER W, AND FAULSTICH H. The
471
472
DOS REMEDIOS ET AL.
Physiol Rev • VOL
286.
287.
288.
289.
290.
291.
292.
293.
294.
295.
296.
297.
298.
299.
300.
301.
302.
303.
304.
305.
306.
307.
ulatory proteins to mixed lipid bilayers. Eur J Biochem 263: 85–92,
1999.
VANCOMPERNOLLE K, GOETHALS M, HUET C, LOUVARD D, AND VANDEKERCKHOVE J. G- to F-actin modulation by a single amino acid
substitution in the actin binding site of actobindin and thymosin
beta 4. EMBO J 11: 4739 – 4746, 1992.
VANCOMPERNOLLE K, VANDEKERCKHOVE J, BUBB MR, AND POLLARD TD.
The interfaces of actin and Acanthamoeba actobindin. J Biol Chem
266: 15427–15431, 1991.
VANDEKECKHOVE J AND WEBER K. At least six different actins are
expressed in a higher mammal: an analysis based on the amino acid
sequence of the amino-terminal tryptic peptide. J Mol Biol 126:
783– 802, 1978.
VAN DEN ENT F, AMOS LA, AND LÖWE J. Prokaryotic origin of the actin
cytoskeleton. Nature 413: 39 – 44, 2001.
VAN TROYS M, DEWITTE D, VERSCHELDE JL, GOETHALS M, VANDEKERCKHOVE J, AND AMPE C. Analogous F-actin binding by cofilin and
gelsolin segment 2 substantiates their structural relationship. J Biol
Chem 272: 32750 –32758, 1997.
VARTIAINEN MK, MUSTONEN T, MATTILA P, OJALA PJ, THESLEFF MK,
PARTANEN J, AND LAPPALAINEN P. The three mouse actin-depolymerizing factor/cofilins evolved to fulfil cell-type-specific requirements
for actin dynamics. Mol Biol Cell 13: 183–194, 2002.
VARTIAINEN N, PYYKONEN I, HOKFELT T, AND KOISTINAHO J. Induction of
thymosin ␤(4) mRNA following focal brain ischemia. Neuroreport
7: 1613–1616, 1996.
VOLKMANN N, AMANN KJ, STOILOVA-MCPHIE S, EGILE C, WINTER DC,
HAZELWOOD L, HEUSER JE, LI R, POLLARD TD, AND HANEIN D. Structure
of Arp2/3 complex in its activated state and in actin filament branch
junctions. Science 293: 2456 –2459, 2001.
WANG F, SAMPOGNA RV, AND WARE B. pH dependance of actin
self-assembly. Biophys J 55: 293–298, 1989.
WATAKABE A, KOBAYASHI R, AND HELFMAN DM. N-tropomodulin: a
novel isoform of tropomodulin identified as the major binding
protein to brain tropomyosin. J Cell Sci 109: 2299 –2309, 1996.
WATKINS H, CONNER D, THIERFELDER L, JARCHO JA, MACRAE C, MCKENNA WJ, MARON BJ, SEIDMAN JG, AND SEIDMAN CE. Mutations in the
cardiac myosin binding protein-C gene on chromosome 11 cause
familial hypertrophic cardiomyopathy. Nat Genet 11: 434 – 437,
1995.
WEBER A, PENNISE CR, BABCOCK GG, AND FOWLER VM. Tropomodulin
caps the pointed ends of actin filaments. J Cell Biol 127: 1627–1635,
1994.
WEBER A, PENNISE CR, AND PRING M. DNase I increases the rate
constant of depolymerization at the pointed (⫺) end of actin filaments. Biochemistry 33: 4780 – 4786, 1994.
WEETMAN AP. Graves’ disease. N Engl J Med 343: 1236 –1248, 2000.
WEGNER A. Head to tail polymerization of actin. J Mol Biol 108:
139 –150, 1976.
WEGNER A AND ISENBERG G. Twelvefold difference between the
critical concentrations of the two ends of actin filaments in physiological salt conditions. Proc Natl Acad Sci USA 80: 4922– 4925,
1983.
WELCH MD, IWAMATSU A, AND MICHISON TJ. Actin polymerization is
induced by Arp2/3 protein complex at the surface of Listeria
monocytogenes. Nature 385: 265–269, 1997.
WITKE W, SHARPE AH, HARTWIG JH, AZUMA T, STOSSEL TP, AND KWIATKOWSKI DJ. Haemostatic, inflammatory, and fibroblast responses
are blunted in mice lacking gelsolin. Cell 81: 41–51, 1995.
WRIGGERS W, TANG JX, AZUMA T, MARKS PW, AND JANMEY PA. Cofilin
and gelsolin segment-1: molecular dynamics simulation and biochemical analysis predict a similar actin binding mode. J Mol Biol
282: 921–932, 1998.
XU J, CASELLA JF, AND POLLARD TD. Effect of capping protein, CapZ,
on the length of actin filaments and mechanical preperties of actin
filaments networks. Cell Motil Cytoskeleton 42: 73– 81, 1999.
YANG J, MORAVEC CS, SUSSMAN MA, DIPAOLA NR, FU D, HAWTHORN L,
MITCHELL CA, YOUNG JB, FRANCIS GS, MCCARTHY PM, AND BOND M.
Decreased SLIM1 expression and increased gelsolin expression in
failing human hearts measured by high-density oligonucleotide
arrays. Circulation 102: 3046 –3052, 2000.
YANG N, HIGUCHI O, OHASHI K, NAGATA K, WADA A, KANGAWA K,
NISHIDA E, AND MIZUNO K. Cofilin phosphorylation by LIM-kinase 1
83 • APRIL 2003 •
www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
263. SELLERS J. Myosin. In: Protein Profile (2nd ed.), edited by Sheterline
P. Oxford, UK: Oxford Univ. Press, 1999.
264. SELVE N AND WEGNER A. Rate of treadmilling of actin filaments in
vitro. J Mol Biol 187: 627–231, 1986.
265. SEPT D, XU J, POLLARD TD, AND MCCAMMON JA. Annealing accounts
for the length of actin filaments formed by spontaneous polymerization. Biophys J 77: 2911–2919, 1999.
266. SHETERLINE P, CLAYTON J, AND SPARROW JC (Editors). Actin. In:
Protein Profile (4th ed.). Oxford, UK: Academic, 1998.
267. SIMENEL C, VAN TROYS M, VANDEKERCKHOVE J, AMPE C, AND DELEPIERRE M. Structural requirements for thymosin beta4 in its contact
with actin. An NMR-analysis of thymosin beta4 mutants in solution
and correlation with their biological activity. Eur Biophys J 267:
3530 –3538, 2000.
268. SOUTHWICK FS. Gelsolin and ADF/cofilin enhance the actin dynamics of motile cells. Proc Natl Acad Sci USA 97: 6936 – 6938, 2000.
269. STANTON LW, GARRARD LJ, DAMM D, GARRICK BL, LAM A, KAPOUN AM,
ZHENG Q, PROTTER AA, SCHREINER GF, AND WHITE RT. Altered patterns of gene expression in response to myocardial infarction. Circ
Res 86: 939 –945, 2000.
270. STRZELECKA-GOLASZEWSKA H. Divalent cations, nucleotides and actin
structure. Res Prob Cell Differ 32: 23– 42, 2001.
271. STRZELECKA-GOLASZEWSKA H, PROCHNIEWICZ E, AND DRABIKOWSKI W.
Interaction of actin with divalent cations. I. The effect of various
cations on the physical state of actin. Eur J Biochem 88: 219 –227,
1978.
272. SUCK W, OEFNER C, AND KABSCH W. Three-dimensional structure of
bovine pancreatic DNase I at 2.5 Å resolution. EMBO J 3: 2423–
2430, 1984.
273. SUN HQ, KWIATKOWSKA K, AND YIN HL. ␤-Thymosins are not simple
actin monomer buffering proteins. Insights from overexpression
studies. J Biol Chem 271: 9223–9230, 1996.
274. SUN HQ, LIN KM, AND YIN HL. Gelsolin modulates phospholipase C
activity in vivo through phospholipid binding. J Cell Biol 13: 811–
820, 1997.
275. SUN HQ, WOOTEN DC, JANMEY PA, AND YIN HL. The actin side-binding
domain of gelsolin also caps actin filaments. Implications for actin
filament severing. J Biol Chem 35: 5229 –5237, 1994.
276. SUNG LA, FAN Y, AND LIN CC. Gene assignment, expression, and
homology of human tropomodulin. Genomics 34: 92–96, 1996.
277. SUSSMAN MA, BAQUE S, UHM CS, DANIELS MP, PRICE RL, SIMPSON D,
TERRACIO L, AND KEDES L. Altered expression of tropomodulin in
cardiomyocytes disrupts the sarcomeric structure of myofibrils.
Circ Res 82: 94 –105, 1998.
278. SUSSMAN MA, WELCH S, GUDE N, KHOURY PR, DANIELS SR, KIRKPATRICK
D, WALSH RA, PRICE RL, LIM HW, AND MOLKENTIN JD. Pathogenesis of
dilated cardiomyopathy: molecular, structural, and population
analyses in tropomodulin-overexpressing transgenic mice. Am J
Pathol 155: 2101–2113, 1999.
279. SUSSMAN MA, WELCH S, WALKER A, KLEVITSKY R, HEWETT TE, WITT SA,
KIMBALL TR, PRICE R, LIM HW, AND MOLKENTIN JD. Hypertrophic
defect unmasked by calcineurin expression in asymptomatic tropomodulin overexpressing transgenic mice. Cardiovasc Res 46:
17–19, 2000.
280. TAGAWA H, WANG N, NARISHIGE T, INGBER DE, ZILE MR, AND COOPER
G. Cytoskeletal mechanics in pressure-overload cardiac hypertrophy. Circ Res 80: 281–289, 1997.
281. TEUBNER A AND WEGNER A. Kinetic evidence for a readily exchangeable nucleotide at the terminal subunit of the barbed ends of actin
filaments. Biochemistry 37: 7532–7538, 1998.
282. THIERFELDER L, WATKINS H, MACRAE C, LAMAS R, MCKENNA W, VOSBERG HP, SEIDMAN JG, AND SEIDMAN CE. Alpha-tropomyosin and
cardiac troponin T mutations cause familial hypertrophic cardiomyopathy: a disease of the sarcomere. Cell 77: 701–712, 1994.
283. TIRION MM, BENAVRAHAM D, LORENZ M, AND HOLMES KC. Normal
modes as refinement parameters for the F-actin model. Biophys J
68: 5–12, 1995.
284. TSUTSUI H, ISHIHARA K, AND COOPER G. Cytoskeletal role in the
contractile dysfunction of hypertrophied myocardium. Science 260:
682– 687, 1993.
285. TUOMINEN EKJ, HOLOPAINEN JM, CHEN J, PRESTWICH MACHILLER GD,
KINNUNEN PR, PKJ, AND JANMEY PA. Fluorescent phosphoinositide
derivatives reveal specific binding of gelsolin and other actin reg-
ACTIN AND ACTIN BINDING PROTEINS
308.
309.
311.
312.
and its role in rac-mediated actin reorganization. Nature 393: 809 –
812, 1998.
YONEZAWA N, NISHIDA E, IIDA K, KUMAGAI H, YAHARA I, AND SAKAI H.
Inhibition of actin polymerization by a synthetic dodecapeptide
patterned on the sequence around the actin-binding site of cofilin.
J Biol Chem 266: 10485–10499, 1991.
YONEZAWA N, NISHIDA E, IIDA K, YAHARA I, AND SAKAI H. Inhibition of
the interactions of cofilin, destrin, and deoxyribonuclease I with
actin by phosphoinositides. J Biol Chem 265: 8382– 8386, 1990.
YONEZAWA N, NISHIDA E, KOYASU S, MAEKAWA S, OHTA Y, YAHARA I, AND
SAKAI H. Distribution among tissues and intracellular localization of
cofilin, a 21 kDa actin-binding protein. Cell Struct Funct 12: 443–
452, 1987.
YONEZAWA N, NISHIDA E, AND SAKAI H. pH control of actin polymerization by cofilin. J Biol Chem 260: 14410 –14412, 1985.
473
313. YU FX, JOHNSTON PA, SUDHOF TC, AND YIN HL. gCap39, a calcium ionand polyphosphoinositide-regulated actin capping protein. Science
250: 1413–1415, 1990.
314. YU FX, LIN SC, MORRISON-BOGORAD M, AND YIN HL. Effects of thymosin ␤ 4 and thymosin beta 10 on actin structures in living cells.
Cell Motil Cytoskeleton 27: 13–25, 1994.
315. ZALEVSKY J, LEMPERT L, KRANITZ H, AND MULLINS RD. Different WASP
family proteins stimulate different Arp2/3 complex-dependent actin-nucleating activities. Curr Biol 11: 1903–1913, 2001.
316. ZALLEN JA, COHEN Y, HUDSON AM, COOLEY L, WIESCHAUS E, AND
SCHEJTER ED. SCAR is a primary regulator of Arp2/3-dependent
morphological events in Drosophila. J Cell Biol 156: 689 –701,
2002.
317. ZALVIDE JB, ALVAREZ CV, VIDAL A, DIEGUEZ C, VEGA FV, AND
DOMINGUEZ F. Regulation of thymosin beta 4 mRNA levels during
cell proliferation. Cell Prolif 28: 85–91, 1995.
Downloaded from http://physrev.physiology.org/ by 10.220.33.5 on July 4, 2017
Physiol Rev • VOL
83 • APRIL 2003 •
www.prv.org