JOURNAL OF STRUCTURAL BIOLOGY
ARTICLE NO. SB973873
119, 295–320 (1997)
Actin: From Cell Biology to Atomic Detail
Michel O. Steinmetz, Daniel Stoffler, and Andreas Hoenger1
M. E. Müller Institute for Microscopy, Biozentrum, University of Basel, CH-4056 Basel, Switzerland
Andreas Bremer2
M.E. Müller Institute for Microscopy, Biozentrum, University of Basel, CH-4056 Basel, Switzerland; and Department of Cell Biology,
Duke University Medical Center, Durham, North Carolina 27710
and
Ueli Aebi
M.E. Müller Institute for Microscopy, Biozentrum, University of Basel, CH-4056 Basel, Switzerland
Received September 11, 1995, and in revised form March 14, 1997
and developmentally regulated isoforms (for review,
see Hennessey et al., 1993; Sheterline et al., 1995). In
vivo, roughly equal amounts of actin are unpolymerized (termed G-actin)3 and in filamentous form
(termed F-actin). In vitro, actin is monomeric under
low ionic strength conditions, e.g., 0.2 mM Ca21 or
0.05 mM Mg21. Elevating the ionic strength to
physiological conditions (e.g., 2 mM MgCl2/100 mM
KCl) causes monomeric G-actin to polymerize within
minutes into synthetic F-actin filaments. Figure 1a
illustrates that a macroscopic indication of actin
polymerization is a sharp rise in the viscosity of the
solution: G-actin solutions have a viscosity that is
close to that of water (top cuvette), whereas F-actin
is very viscous, as evidenced by the trapped bubbles
(bottom cuvette). Judged by light and electron microscopy, synthetic F-actin filaments can become many
micrometers long (see Borejdo and Burlacu, 1991;
Burlacu et al., 1992). Micrographs of such synthetic
F-actin filaments are shown after different preparation and electron microscopic imaging: Fig. 1b reveals a negatively stained specimen imaged at high
magnification in a scanning transmission electron
microscope (STEM) by the annular dark-field (ADF)
mode; Fig. 1c displays a completely unstained speci-
Over the past 2 decades our knowledge about
actin filaments has evolved from a rigid ‘‘pearls on a
string’’ model to that of a complex, highly dynamic
protein polymer which can now be analyzed at
atomic detail. To achieve this, exploring actin’s oligomerization, polymerization, polymorphism, and dynamic behavior has been crucial to understanding
in detail how this abundant and ubiquitous protein
can fulfill its various functions within living cells. In
this review, a correlative view of a number of distinct aspects of actin is presented, and the functional implications of recent structural, biochemical, and mechanical data are critically evaluated.
Rational analysis of these various experimental data
is achieved using an integrated structural approach
which combines intermediate-resolution electron
microscopy-based 3-D reconstructions of entire actin filaments with atomic resolution X-ray data of
monomeric and polymeric actin. r 1997 Academic Press
INTRODUCTION
The molecular building blocks of the eukaryotic
cytoskeleton include actin, intermediate filament
proteins, and tubulin together with their respective
associated proteins. Actin is a highly conserved
eukaryotic protein that is expressed in most animals, plants, and fungi in multiple, tissue-specific,
3 Abbreviations used: G-actin, monomeric actin; F-actin, synthetic actin filaments; EM, electron microscope/microscopy; CTEM,
conventional transmission electron microscope/microscopy; STEM,
scanning transmission electron microscope/microscopy; ADF, annular dark-field; BF, bright-field; FESEM, field-emission scanning electron microscope/microscopy; 2-D, 3-D, two-, threedimensional; S-1, myosin subfragment-1; HAS, high-affinity
divalent cation binding site.
1 Present address: Institute for Cell Biology, Swiss Federal
Institute of Technology (ETH), CH-8093 Zürich, Switzerland.
2 Present address: Bellevue Asset Management AG, CH-6301
Zug, Switzerland.
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1047-8477/97 $25.00
Copyright r 1997 by Academic Press
All rights of reproduction in any form reserved.
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STEINMETZ ET AL.
FIG. 1. Actin polymerization, actin filaments, and actin-based motility. (a) (Top) A quartz cuvette containing G-actin at a protein
concentration of 1 mg/ml. (Bottom) The same cuvette 15 min after adding MgCl2 to 2 mM and KCl to 50 mM. Small air bubbles are formed
during polymerization of G- into F-actin. The larger bubbles have been introduced artificially to demonstrate that the viscosity of F-actin is
significantly higher than that of the monomeric G-actin solution. (b–d) Different preparations of F-actin filaments: (b) negatively stained
with 0.75% uranyl formate and imaged in a scanning transmission electron microscope (STEM) by the annular dark-field (ADF) mode; (c)
frozen-hydrated and phase-contrast imaged in a conventional transmission electron microscope (CTEM) (courtesy of Dr. Ron Milligan,
Scripps Research Institute, La Jolla, CA); and (d) freeze-dried/rotary metal-shadowed and imaged by secondary electrons in a
field-emission scanning electron microscope (FESEM) (courtesy of Dr. Roger Wepf, European Molecular Biology Laboratory (EMBL),
Heidelberg, Germany). (e) Actin-based in vitro motility assay. The experimental setup for an in vitro motility assay is relatively simple. A
flow cell, i.e., an open system that allows the exchange of solutions during the experiment, is assembled from a glass slide and a
nitrocellulose-coated coverslip using two parallel lines of silicon grease. A myosin solution is then allowed to flow into the cell. Within
seconds, the myosin molecules bind to the coverslip. After a washing step to remove excess myosin, actin filaments that have been labeled
with fluorescent phalloidin (TRITC-phalloidin; courtesy of Dr. H. Faulstich, Max-Planck Institute for Medical Research, Heidelberg,
Germany) are added to the flow cell. These fluorescent actin filaments bind to the myosin molecules and the motility assay system is now
ready for use. Adding ATP results in a sliding movement of actin filaments. Three video frames (i.e., after t 5 0, 4, and 8 sec of a particular
area of interest) have been recorded after addition of ATP. Three filaments have been marked with arrowheads in each frame to follow their
myosin-driven movement. Scale bar, 25 nm (b–d).
ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
men preserved in a near-native state in a thin film of
amorphous ice and phase-contrast imaged in a conventional transmission electron microscope (CTEM);
and Fig. 1d yields a freeze-dried/rotary metalshadowed sample imaged by secondary electrons
(SE) in a field-emission scanning electron microscope (FESEM).
To reconstitute molecular force generation in a
minimal test tube system, elegant in vitro motility
assays have been developed that require a molecular
motor (e.g., myosin), a molecular ‘‘track’’ (e.g., synthetic F-actin filaments), and fuel (i.e., ATP) (for
details, see Kron et al., 1991). These systems have
significantly advanced our current view of the biochemistry and biophysics of the mechanochemical
ATPase cycle (Finer et al., 1994), the functional
anatomy of the myosin molecule (Lowey et al.,
1993a,b), and the role of the actin filament in generating directed force during muscle contraction or
cellular motility (Prochniewicz and Yanagida, 1990;
Schwyter et al., 1990). Light microscopy allows imaging of dynamic biological processes in their native
aqueous environment with relatively high temporal
resolution (i.e., subsecond regime). As an example, in
Fig. 1e a time-lapse video sequence of unidirectionally moving F-actin filaments driven by myosin
subfragment-1 (S-1) immobilized on a nitrocellulosecoated coverslip is presented. For visualization, the
actin filaments were labeled with fluorescent phalloidin (TRITC-phalloidin) and imaged by a videoenhanced low-light-level camera in an epifluorescence microscope.
Because conditions favoring crystallization of Gactin also induce its polymerization into F-actin
filaments, it has been difficult to obtain 3-D crystals
of pure actin suitable for X-ray diffraction analysis.
To overcome this problem, several investigators have
complexed actin with proteins that inhibit its polymerization and thereby obtained cocrystals. As a
result of these efforts, the atomic structures of actin
in complex with DNase I (Kabsch et al., 1990),
gelsolin segment 1 (McLaughlin et al., 1993), and
profilin (Schutt et al., 1993) have now been determined. Comparison of the different actin structures
has revealed that whereas their overall structures
are remarkably similar, they exhibit a number of
significant local differences which may be due in part
to the different proteins with which they were complexed, but also to the fact that different isoforms
were investigated (i.e., skeletal muscle a-actin in the
case of the actin–DNase I and the actin–gelsolin
segment 1 cocrystals, and bovine b-actin in the case
of the actin–profilin cocrystals). Based on the atomic
structure of skeletal muscle actin determined by
Kabsch et al. (1990), atomic models for synthetic
F-actin filaments (Holmes et al., 1990; Lorenz et al.,
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1993), muscle thin filaments (Lorenz et al., 1995),
and S-1-decorated actin filaments (Rayment et al.,
1993; Schroeder et al., 1993) have been proposed,
and these have been compared with EM-based 3-D
helical reconstructions of corresponding specimens
(e.g., Bremer and Aebi, 1992; Rayment et al., 1993;
Schroeder et al., 1993; Bremer et al., 1994; Egelman
and Orlova, 1995a; Hoenger and Aebi, 1996).
A number of studies have shown that actin filaments are intrinsically dynamic (Aebi et al., 1986;
Carlier, 1991; Bremer et al., 1991, 1994; Steinmetz et
al., 1997) and that this dynamics can be modulated
by effector molecules such as phalloidin or the state
of hydrolysis of the bound nucleotide (Janmey et al.,
1990; Bremer et al., 1991; Bremer and Aebi, 1992;
Orlova and Egelman, 1992; Lorenz et al., 1993;
Lepault et al., 1994; Muhlrad et al., 1994; Steinmetz
et al., 1997). Also, there is ample evidence that
significant structural changes in the actin molecule
may occur upon its interaction with actin-binding
proteins, including myosin and its fragments (Prochniewicz and Yanagida, 1990; Schwyter et al., 1990;
Owen and DeRosier, 1993; McGough et al., 1994;
Schmid et al., 1994; Orlova et al., 1995).
ACTIN IN VIVO
Feuer et al. (1948) first isolated actin as a major
muscle protein. For illustration, in Fig. 2a an ultrathin longitudinal section of a rigor insect flight
muscle reveals a MyAc layer (e.g., Taylor et al., 1984)
in which thin and thick filaments alternate and are
packed with almost crystalline order, thereby maximizing the interaction between the myosin heads
radially projecting from the thick filaments and the
thin filaments for the development of force (for a
detailed review of the more recent history and
theories of muscle contraction, see the review of
Jontes, 1996). In the 1970s, it slowly but definitely
emerged that cytoskeletal actin also plays important
roles unrelated to muscle function (reviewed by dos
Remedios and Barden, 1993; Way and Weeds, 1990;
Pollard, 1986). Accordingly, nonmuscle actin is involved in processes such as cell and intracellular
motility, cell division, and dynamic remodeling of the
cytoskeleton. Moreover, the bacterial pathogen Listeria monocytogene induces actin polymerization into
polar rocket-tail structures within the host cell as a
means to promote its rapid intracellular movement
and its cell-to-cell transfer (for review, see Southwick
and Purich, 1994). As illustrated in Fig. 1e, actin
filaments may also serve as molecular ‘‘tracks’’ for
myosin motors to move along. Furthermore, F-actin
acts as an ATPase, and it interacts with dozens of
actin-binding and regulatory proteins and with small
effector molecules (see below; for review, see Hennessey et al., 1993, and references therein). Last, but not
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FIG. 2. Actin in vivo. (a) Electron micrograph of an ultrathin longitudinal section (for review, see Villiger and Bremer, 1990) through an
insect indirect flight muscle (courtesy of Mary C. Reedy, Duke University Medical Center, Durham, NC). The section is one MyAc layer
thick and yields an almost crystalline array of alternating thick (i.e., myosin-containing) and thin (i.e., actin-containing) muscle filaments.
(b) Goldfish keratocyte labeled with BODIPY-phalloidin. Scales of goldfish were plucked, washed with supplemented medium (amphibian
culture medium from Gibco containing penicillin, streptomycin, neomycin, and amphotericin B), and cultured overnight in the same
medium in tissue culture flasks. Keratocytes that migrate as a single-cell tissue off the scale were separated from contaminating cells and
debris by washing them several times with Ca21-free phosphate-buffered saline (PBS). To dissociate the cells and to detach them from the
plastic substratum of the tissue culture flask, they were incubated with 5 mM EGTA/5 mM EDTA in Ca21-free PBS at pH 7.0 for a few
minutes. The exact time required for quantitative release of the cells was determined by observing the tissue culture flask under the
microscope. The medium containing the cells was then transferred into centrifuge tubes, the cells were spun down, and the pellet was
resuspended in supplemented medium and plated on coverslips, where the cells reattached within minutes. ON, orthogonal network at the
front of the cell; EN, endoplasm, the vesicle-rich part of the cytoplasm that contains the nucleus; FC, focal contacts; RF, retraction fibers. (c)
Electron micrograph of a human KD fibroblast which has been cultured directly on a Formvar-coated EM gold grid prior to detergent
extraction/chemical fixation followed by negative staining with 0.75% uranyl formate. Three distinct cytoskeletal systems can be
distinguished: actin-containing microfilaments (MF), vimentin-type intermediate filaments (IF), and tubulin-containing microtubules
(MT). Along the cell cortex the microfilaments tend to laterally associate into filament bundles, called stress fibers (SF). Scale bars, 100 nm
(a), 10 µm (b), and 250 nm (c).
ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
least, actin filaments may also play a role in compartmentalizing metabolic pathways within cells (for a
review, see e.g., Matsudaira, 1991).
The fluorescence micrograph of a goldfish keratocyte shown in Fig. 2b illustrates that the actin
cytoskeleton defines the overall size and general
shape of this highly motile cell. The actin filaments
in the front of the keratocyte’s lamellipodium are
well organized and form an approximately orthogonal network (ON). Filament length decreases and
filament number increases from the endoplasm that
contains the nucleus (EN) toward the front end of the
lamellipodium. Whereas few long filaments extend
all the way through the lamellipodium, most shorter
filaments are located at its front end (see also
Rinnerthaler et al., 1991; Small et al., 1993). This
observation bears interesting consequences for evaluating possible models of the keratocyte’s motility
that were recently discussed by Small and coworkers (Small et al., 1995). According to these
models, actin filaments bind to focal contacts (FC) at
the cell’s lower surface, thereby mediating adhesion
to the substratum, e.g., the culture dish (Small,
1988). Hence, migrating cells exert force on and pull
themselves along these focal contacts. Incomplete
release of filaments from these contacts frequently
occurs, thus resulting in the formation of retraction
fibers (RF) at the posterior end of migrating cells.
A diagnostic feature of the actin cytoskeleton of
fibroblasts grown in culture is the so-called stress
fiber system that traverses the cell near the plasma
membrane facing the substratum. Stress fibers are
loose bundles of actin-containing microfilaments
probably stabilized and/or held together by bundling
or crosslinking proteins. Stress fibers are typically
50–500 nm wide and 5–10 µm long. As illustrated in
Fig. 2c, in the negatively stained whole mount of a
cultured human KD fibroblast, three distinct cytoskeletal filament systems can be distinguished: actincontaining microfilaments (MF), vimentin-type intermediate filaments (IF), and tubulin-containing
microtubules (MT). Within the cell unaggregated
microfilaments form a loose meshwork, whereas
along the cell cortex they bundle into stress fibers
(SF) which frequently appear to be associated with
microtubules.
THE DOMAIN STRUCTURE OF THE G-ACTIN
MONOMER AND THE MOLECULAR ARCHITECTURE
OF THE F-ACTIN FILAMENT
Actin has long resisted crystallization since conditions typically favoring protein crystallization cause
actin to polymerize into filaments. However, adding
stoichiometric amounts of Gd31 to actin under polymerizing conditions yields 2-D crystals which allowed determination of its 3-D structure to 1.5 nm
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resolution from tilted views of negatively stained
sheet preparations (Aebi et al., 1980, 1981; Smith et
al., 1983; see also Fig. 6 and below). Ten years later,
the 3-D structure of the actin molecule was resolved
to atomic resolution by X-ray diffraction analysis of
3-D crystals of actin in complex with DNase I
(Kabsch et al., 1990), gelsolin segment 1 (MacLaughlin et al., 1993), or profilin (Schutt et al., 1993). The
atomic structure of the actin molecule determined
from the actin–DNase I cocrystals (Kabsch et al.,
1990) is displayed in Fig. 3a in the form of a folded
ribbon representation. As indicated in Fig. 3b, the
entire molecule fits into a box measuring 5.5 3 5.5 3
3.5 nm (cf. Kabsch et al., 1990; Bremer and Aebi,
1992). Each actin subunit is composed of four subdomains, termed 1, 2, 3, and 4. The four subdomains
are primarily held together and stabilized by salt
bridges and hydrogen bonds between the phosphate
groups of the bound nucleotide (i.e., ATP or ADP) and
its associated divalent cation, which is believed to be
Mg21 under physiological conditions. The nucleotide
and its associated divalent cation are localized to the
‘‘heart’’ of the actin molecule (see Figs. 3a and 3b).
The actin subunit is also distinctly polar: subdomain
2 has significantly less and subdomain 1 significantly more mass than the other two subdomains
(see Fig. 3b, where the size of the subdomains
corresponds to their relative masses).
Using the atomic structure of G-actin, atomic
models of the F-actin filament were constructed and
refined so as to minimize the difference of their
computed diffraction patterns with X-ray fiber diffraction patterns to about 8 Å resolution recorded from
oriented gels of F-actin filaments (Holmes et al.,
1990; Lorenz et al., 1993; Holmes, 1995). Without
question these atomic filament models have had a
significant impact on our present understanding of
actin filament structure and dynamics. Nevertheless, 3-D helical reconstructions computed from electron micrographs of both negatively stained (e.g.,
Aebi et al., 1986; Bremer et al., 1991, 1994; see Fig.
8a) and frozen hydrated (Milligan et al., 1990) Factin filaments have revealed a remarkable similarity to the atomic models when the latter are represented as electron density maps contoured at
comparable resolution (i.e., 2.5 nm; e.g., Holmes and
Kabsch, 1991; Bremer and Aebi, 1992; Bremer et al.,
1994; Hoenger and Aebi, 1996). Moreover, a recent
reconstruction of Bremer et al. (1994) allowed interpretation of particular structural features at the
level of distinct secondary structural elements, so
that predictions based upon the atomic models could
be verified (see Fig. 8d and below). Hence, EM-based
filament reconstructions have become an invaluable
complement to atomic models, particularly in the
context of investigations trying to assess distinct
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conformational states of the actin polymer in response to various effectors (e.g., Bremer et al., 1991;
Orlova and Egelman, 1992, 1993, 1995; Lepault et
al., 1994; Steinmetz et al., 1997) or to determine the
3-D structure of actin filaments in complex with
actin-binding or regulatory proteins and drugs (e.g.,
Milligan et al., 1990; Bremer et al., 1991; Owen and
DeRosier, 1993; McGough et al., 1994; Schmid et al.,
1994; Orlova et al., 1995; McGough and Way, 1995;
Steinmetz et al., 1997).
Various aspects of a generalized molecular model
of an F-actin filament are presented in Figs. 3d to 3f.
For comparison and critical evaluation of the molecular filament model shown in Fig. 3d, Fig. 3c displays
a helically filtered image (i.e., D(Z, k)-filtered; cf.
Smith and Aebi, 1974; Bremer et al., 1991, 1994) of a
negatively stained synthetic F-actin filament stretch
such as that shown in Fig. 1b. The F-actin filament
can be described as consisting of either two righthanded helices of a relatively long pitch (Fig. 3e) or of
a single left-handed helix of a shallow pitch (Fig. 3f).
Taking into consideration that the intersubunit contacts along the two long-pitch helical strands (right
panel of Fig. 3e) are continuous and approximately
three times stronger than those between them (Erickson, 1989; for reviews, see Bremer and Aebi, 1992;
Holmes and Kabsch, 1991), description of the actin
filament as a two-stranded helical polymer such as
that shown in Fig. 3e makes most physical sense.
Supporting this intersubunit contact pattern, locally
separated, unraveled long-pitch helical strands yielding ‘‘bubbles’’ and ‘‘splayed ends’’ have been reported
(see Fig. 3h; Aebi et al., 1986; Bremer et al., 1991;
Bremer and Aebi, 1992). Both the relative physical
strength of the two types of intersubunit contacts
and the filament unraveling imply that the two
long-pitch helical strands can move relatively independently of each other by ‘‘lateral slipping’’ (Bremer
et al., 1991), a mechanism which involves both a
radial and an angular component (Censullo and
Cheng, 1993; see also below). Nevertheless, the
one-start helix description (see Fig. 3f) of the F-actin
filament yields a simple parametrization of the
filament geometry in terms of a ‘‘helical selection
rule’’ which, in turn, allows elegant 3-D helical
reconstruction to be performed (see also below; cf.
Klug and DeRosier, 1966).
A comparison of the two helix geometries is presented in Figs. 3e and 3f. In the two-start helix
model, there are 13 subunits per right-handed longpitch helical turn of each of the two strands (Fig. 3e,
right panel). Assuming an axial subunit repeat of 5.5
nm, 13 subunits yield a pitch of 13 p 5.5 nm 5 71.5
nm. As a consequence, they cross each other regularly every half pitch, i.e., every 35.75 nm. Figure 3e
also reveals the two long-pitch helical strands to be
axially staggered by half the axial subunit spacing
(i.e., 2.75 nm). The alternative description is as a
5.9-nm pitch one-start ‘‘genetic’’ helix with 13 subunits per 6 left-handed turns (Fig. 3f, right panel).
The schematic view in Fig. 3f reveals that assuming
a subunit spacing of 2.75 nm, 13 subunits in 6 turns
correspond to 13 p 2.75 nm 5 35.75 nm, i.e., the
crossover spacing of the two long-pitch helical strands
(for reviews, see Holmes and Kabsch, 1991; Bremer
and Aebi, 1992).
Asymmetric subunits assemble into intrinsically
polar F-actin filaments by always adding onto the
filament ends in the same orientation with respect to
the filament axis. This filament polarity can be
visualized by stoichiometric binding of myosin S-1 to
F-actin in situ or in vivo, thereby yielding decorated
FIG. 3. The domain structure of the G-actin monomer and the molecular architecture of the F-actin filament. (a) Folding of the actin
molecule represented by ribbon tracing of the a-carbon atoms. An ATP molecule with its associated Ca21 atom are shown in a Van der Waals
sphere representation. (b) Schematic representation of the G-actin molecule in terms of its domain/subdomain organization. At first glance,
the actin molecule as shown in (a) appears to be built of two domains, a ‘‘small’’ domain (i.e., containing boxes 1 and 2) and a ‘‘large’’ domain
(i.e., containing boxes 3 and 4). At a closer look, both the small and the large domains can be further divided into two subdomains each,
called subdomains 1 and 2 (i.e., boxes 1 and 2) and subdomains 3 and 4 (i.e., boxes 3 and 4), respectively. (c) D(Z, k)-filtered image of a
negatively stained synthetic F-actin filament stretch such as shown in Fig. 1b. (d) Generalized structure of the F-actin filament built from
the actin monomer model displayed in (b). The polarity of the actin filament as revealed by the distinct arrowhead pattern upon
stoichiometric decoration with myosin subfragment-1 (S-1) (see g and Fig. 5a) is indicated by short lines (see also c and g). (e) The molecular
architecture of the actin filament may be described by two intertwined right-handed long-pitch helical strands of actin subunits. The pitch
of these long-pitch helical strands is 71.5 nm with 13 subunits per turn (i.e., each having a 5.5-nm axial extent). Seen in projection, the two
strands cross each other every 35.75 nm or every half-pitch (see also c). The axial stagger between the two long-pitch helical strands is 2.75
nm, or one half of the axial extent of a subunit. (f) An alternative description of the molecular architecture of the actin filament is that of a
shallow left-handed one-start helix. This so-called genetic helix has a pitch of 5.9 nm, and it contains 13 subunits in 6 left-handed turns, so
its helical repeat amounts to 35.75 nm or one crossover of the two long-pitch helical strands (see e). (g) (right) Negatively stained
S1-decorated F-actin filament stretch with its ‘‘barbed’’ end to the left and its ‘‘pointed’’ end to the right (see also Fig. 5a). (left)
D(Z, k)-filtered filament projection image such as shown in (c), demonstrating that the ‘‘mini-arrowhead’’ appearance (indicated by short
lines) of the undecorated F-actin filament is opposite to the polarity revealed by the S-1 decoration pattern. (h) Gallery of ADP-F-actin
filament stretches after negative staining with 1% sodium phosphotungstate, pH 7.0. This heavy metal salt, which acts as an inorganic
phosphate analogue (Pi), causes local unraveling of the two long-pitch helical strands so as to produce ‘‘bubbles’’ and ‘‘splayed ends’’ (see
arrowheads). Scale bars, 1 nm (a), 50 nm (g), and 100 nm (h).
ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
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filaments with an ‘‘arrowhead-like’’ appearance
(Moore et al., 1970). The right half of Fig. 3g displays
a short stretch of such a negatively stained S-1decorated F-actin filament with its ‘‘barbed’’ end to
the left and its ‘‘pointed’’ end to the right (see also
Fig. 5a). The polarity revealed by this S-1 decoration
pattern is opposite to that of the ‘‘mini-arrowhead’’
appearance observed in computed EM projection
images of negatively stained F-actin filament
stretches (Bremer et al., 1991; Bremer and Aebi,
1992), such as that illustrated in the left half of Fig.
3g (compare also with Figs. 3c and 3d). This miniarrowhead pattern, in turn, can be used to determine
the polarity of individual F-actin filaments directly
(i.e., without S-1 decoration). For instance, Bremer
et al. (1994) have employed this method for aligning
individual F-actin filament 3-D reconstructions prior
to computing their averaged 3-D reconstructions.
Also, Meyer and Aebi (1990), with a-actinin, and
Faix et al. (1996), with cortexillin I, have used this
direct method to determine whether these F-actin
bundling proteins crosslink actin filaments in a
parallel or an antiparallel fashion.
TESTING PREDICTIONS OF ATOMIC MODELS
A number of mutant actins, obtained by either
selection after random mutagenesis, genetic identification and isolation, or by mutagenesis in vitro have
been produced (for review, see Hennessey et al.,
1993; Sheterline et al., 1995). They allow probing
predictions of atomic models for the structure of the
actin filament (e.g., Lorenz et al., 1993) and dissecting the anatomy of the actin molecule functionally
both in vivo and in vitro.
Many mutant actins display ‘‘antimorphic’’ or
‘‘dominant negative’’ phenotypes in vivo, i.e., the
heterozygous organism exhibits the mutant phenotype. A likely explanation for this observation is that
filament formation involves several binding sites on
the actin molecule (for review, see Sparrow et al.,
1992; Sparrow, 1995), and so, if a point mutation
happens to destroy one of them, the remaining
binding sites on this molecule may still be functional.
Therefore, the mutant actin molecule may nonproductively interact with wild-type molecules in that it
still incorporates into filaments made of wild-type
molecules but then blocks the addition of further
subunits. Because of this antimorphic behavior, in
vivo effects of mutant actins have often been difficult
to interpret. Fortunately, the Drosophila genome
contains one actin gene which can be mutated without producing a lethal phenotype: Act88F encodes
actin III, a unique actin isoform expressed solely in
the indirect flight muscle (Fyrberg and Donady,
1979; Fyrberg et al., 1983). For in vitro studies,
transgenic flies harboring a mutant actin gene can
be generated by P-element-mediated germline transformation into an Act88F null mutant (see, e.g.,
Drummond et al., 1990).
A mutant actin which has been extensively characterized involves a point mutation of Gly245 to Asp245,
called G245D. Chemical mutagenesis of the diploid
human KD fibroblast strain led to the isolation of the
substrain HuT-14 which is both immortalized and
tumorigenic (Leavitt and Kakunaga, 1980; see also
Leavitt et al., 1982). The HuT-14 substrain expresses
one G245D and one wild-type b-actin allel each
(Vandekerckhove and Weber, 1980). The G245D actin incorporates into stress fibers at lower levels
than wild-type actin, and it alters cell morphology
(Leavitt and Kakunaga, 1980; Leavit et al., 1987).
Figure 4a documents that G245D actin is also polymerization deficient in vitro: The lane marked ‘‘Start’’
reveals the actin mixture isolated from tumors induced in nude mice after inoculation with HUT-14
cells. The lane marked ‘‘Pel’’ contains the pellet of the
polymerized actin mixture, whereas the lane marked
‘‘Sup’’ contains the corresponding supernatant. The
G245D actin clearly enriches over the wild-type
actin in the supernatant (i.e., it is not efficiently
polymerized), whereas the wild-type actin is almost
quantitatively recovered in the pellet. Hence, G245D
actin can selectively be enriched from a mixture of
wild-type and mutant actin by ‘‘cycling’’ (i.e., repeated polymerization and depolymerization of highspeed supernatants). Figure 4b demonstrates that
upon addition of e.g., 2mM MgCl2/50 mM KCl to
G245D mutant actin, instead of polymerizing into
filaments (see inset), it associates into unspesific,
loose aggregates. This clearly demonstrates that
actins mutated at Gly245 (i.e., G245D or G245K)
exhibit a serious polymerization defect (Millonig et
al., 1988; Taniguchi et al., 1988; Aspenstrom et al.,
1992). Consistent with this finding, heterozygous
flies being transgenic for G245D actin produce myofibrils that appear normal on eclosion but eventually
become disrupted over the following 12–14 days, so
that the flies eventually become flightless (Sakai et
al., 1991; Sparrow et al., 1992).
Figure 4c shows residues 243–245 (Pro-Asp-Gly)
at the tip of subdomain 4 of one subunit and residues
322–325 (Pro-Ser-Thr-Met) at the bottom of subdomain 3 of the subunit above it, specifying one of the
major intersubunit contacts along the two long-pitch
helical strands. The polymerization-deficient phenotype of G245D actin is therefore not at all surprising:
as is documented in Fig. 4c, replacing the Gly
residue at position 245 by a negatively charged
residue such as Asp is likely to block polymerization
by a charge mismatch. As is illustrated schematically in Fig. 4d, G245D mutant actin can, however,
still be induced to form ‘‘folded ribbons’’ and sheets
ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
(Millonig et al., 1988). In contrast to F-actin filaments (which are ‘‘upper dimer’’ related), these supramolecular assemblies are ‘‘lower dimer’’ related (see
below and Fig. 6c) and involve intersubunit contacts
distinct from those engaged in filament formation.
A prominent feature in the atomic model of the
F-actin filament (Holmes et al., 1990; refined by
Lorenz et al., 1993) is an extended b-hairpin (i.e.,
residues 262–274). This structure is commonly referred to as the ‘‘hydrophobic plug’’ because the four
central residues (266–269) forming its turn are
hydrophobic. Holmes et al. (1990) suggested a pluglike insertion of this hydrophobic loop across the
filament axis into a complementary ‘‘hydrophobic
pocket’’ provided by the interface between two adjacent subunits of the opposite long-pitch helical strand.
To test this hypothesis, Rubenstein and co-workers
(Chen et al., 1993) mutated a Leu residue, L266, of
the yeast ACT1 gene to Asp (L266D), thereby introducing a hydrophilic residue into the hydrophobic
plug. L266D actin exhibited a cold-sensitive phenotype with a higher critical concentration for polymerization than wild-type actin. This finding indicated
that L266D actin cannot form nuclei at low temperature since the mutated hydrophobic plug can no
longer adopt a proper filament conformation. Independent confirmation for Holmes’ hydrophobic loop
hypothesis came from 3-D reconstructions of negatively stained F-actin filaments by Bremer et al.
(1994) that revealed a distinct high mass density
‘‘bridge’’ across the filament axis that coincided with
the position of the hydrophobic plug (Fig. 8d; see also
below).
OLIGOMERIZATION, POLYMERIZATION, AND
CRYSTALLIZATION OF ACTIN
As documented in Fig. 5c, when monitored by the
increase of the fluorescence signal of actin pyrenated
at Cys374, the time course for polymerizing G-actin
monomers into F-actin filaments is distinctly sigmoidal. It can be divided into at least three distinct
phases: an initial nucleation or lag phase, followed
by an almost linear elongation phase, and finally, a
steady-state phase. Therefore, polymerization is commonly considered a nucleation–elongation process,
involving fast monomer activation, rate-limiting
nucleation, and moderately fast elongation (reviewed by Pollard, 1990; Carlier, 1991; Estes et al.,
1992). The nucleus is defined as the first oligomer
formed during polymerization that is more likely to
grow than to shrink. For actin polymerization, most
available evidence suggests the nucleus to be a
trimer (i.e., involving three subunits along the genetic helix; see Figs. 3f and 5e). During the lag
phase, monomer activation and nucleus formation
predominate, and this process strongly depends on,
303
e.g., the type of divalent cation (Ca21 or Mg21 )
present at the high-affinity divalent cation-binding
site (HAS) of the G-actin molecule (Fig. 11; see also
below). After a significant number of nuclei have
been formed, rapid subunit addition ensues, and the
polymerization reaction progresses into the elongation phase during which the relationship of total
polymer versus time is approximately linear. The
kinetic rate constants for filament elongation strongly
depend on the exact polymerization conditions (e.g.,
pH, temperature, ionic strength, state of hydrolysis
of the bound nucleotide, and type of divalent cation
bound to the HAS of G-actin), but an overall elongation rate of roughly 50 subunits per second and
filament (i.e., corresponding to about 140 nm/sec
filament growth) is probably within the right order of
magnitude (see, e.g., Pollard, 1983). Finally, at steady
state the monomer and polymer concentrations remain invariant, and the monomer concentration
equals the critical concentration for polymerization.
However, subunit exchange between the polymer
and the monomer pools continues by a phenomenon
called ‘‘treadmilling’’ (i.e., subunit flux through the
filament). Possibly, at steady state breaking and
reannealing of filaments may also occur at significant rates (reviewed by Oosawa, 1993).
As was discussed earlier and is depicted in Figs. 3a
and 3b, the 4-domain actin molecule is stabilized by
a tightly bound adenine nucleotide (i.e., ATP or ADP)
and its associated divalent cation (i.e., Ca21 or
Mg21 ). At first glance, treadmilling seems to contradict the laws of thermodynamics, but the polymerization of actin with bound ATP (i.e., ATP–G-actin)
dissipates chemical energy via ATP hydrolysis and is
therefore not strictly reversible (Carlier, 1992). Consistent with this finding, it was first shown by
Wegner (1976) that an energetic difference existed
between the two filament ends. Hence, it is remarkable that ADP–G-actin also polymerizes into filaments, only at an approximately 25-fold higher
critical concentration and at a much slower rate than
ATP–G-actin (Pollard, 1984). ATP hydrolysis increases the rate of subunit exchange: compared to
ATP–G-actin ADP–G-actin dissociates about 10 times
faster from the barbed end of the filament (Korn et
al., 1987; Carlier and Pantaloni, 1988).
Standard isolation protocols (reviewed in Pardee
and Spudich, 1982) yield Ca–ATP–G-actin (i.e., with
Ca21 bound to the HAS of the molecule), whereas in
vivo the majority of G-actin is believed to be Mg–ATP–
G-actin (reviewed in Estes et al., 1992). The affinities
of actin for both ATP and the tightly bound divalent
cation are in the nanomolar range (Estes et al.,
1992). Under most polymerization conditions, ATP
hydrolysis lags behind polymerization (e.g., Pollard
and Weeds, 1984) and because of this uncoupling,
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STEINMETZ ET AL.
FIG. 4. Characterization of a mutant human b-actin, called G245D, involving a point mutation of Gly245 to Asp245. (a) SDS–PAGE of a
mixture of wt and G245D actin before and after polymerization with 2 mM MgCl2 and 50 mM KCl: lane 1 (Start), actin mixture before
polymerization; lane 2 (Pel), high-speed (i.e., for 1 hr at 100,000g) pellet of actin mixture after polymerization; and lane 3 (Sup), high-speed
supernatant of actin mixture after polymerization. The enrichment of G245D actin over wt actin in the supernatant is evident, indicating
an altered polymerization behavior of G245D actin compared with wt actin. (b) Electron micrograph of G245D actin after polymerization
with 2 mM MgCl2 and 50 mM KCl, negatively stained with 0.75% uranyl formate. For comparison, wt actin polymerized and prepared for
EM in the same way is shown in the inset. (c) Interface (left, for wt actin; right, for G245D actin) of the actin–actin contact along the
long-pitch helical strands (see also Figs. 3e, 8b, and 8c) at the tip of subdomain 4 (i.e., involving residues P243, D244, and G/D245) of one
subunit and subdomain 3 (i.e., involving residues P322, S323, T324, and M325) of the next subunit above it. The atomically built models for
wt and G245D actin illustrate that replacing the Gly by the larger Asp side chain at amino acid residue 245 is likely to cause a charge
mismatch within the corresponding actin–actin contact along the long-pitch helical strands, thereby interfering with polymerization into
F-actin filaments. (d) Schematic drawing illustrating that while G245D actin fails to polymerize into bona fide F-actin filaments (see b), it
remains capable of forming 2-D crystalline arrays in the presence of stoichiometric amounts of Gd31 (see Figs. 6 and 7; also, Millonig et al.,
1988). Scale bar, 100 nm (b).
ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
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STEINMETZ ET AL.
the state of hydrolysis of the bound nucleotide within
the newly formed actin filaments changes with time.
Since ATP hydrolysis is fast and irreversible and the
release of inorganic phosphate (Pi) is slow and reversible, neither ATP– nor ADP–F-actin but ADP–Pi–Factin is the major intermediate present during polymerization.
Since actin has been the prototype of a protein
readily polymerizing into polar filaments (Fig. 5a), it
came somewhat as a surprise that at least in vitro it
could also form a variety of nonfilamentous polymeric or crystalline supramolecular assemblies (Aebi
et al., 1981), for example, crystalline actin tubes, as
shown in Fig. 6a. Along the same lines, specific and
stoichiometric interaction with actin-binding proteins does interfere with actin’s structural polymorphism: depending on the exact ionic conditions,
stoichiometric mixtures of actin and DNase I will
form either actin–DNase I cocrystals (Mannherz et
al., 1977; reviewed in Mannherz et al., 1992) or
actin–DNase I tubes (Fig. 7e; Fowler et al., 1984).
Moreover, it has become evident that a significant
fraction of G-actin dimerizes at early stages of
polymerization under a wide variety of polymerization conditions (see Newman et al., 1985; Goddette et
al., 1986; Matsudaira et al., 1987; Millonig et al.,
1988; for review, see Grazi, 1989). Compared to the
very careful analyses of monomer activation, nucleation, elongation, and the effects that the nucleotide
and the divalent cation bound to the HAS have on
polymerization rate constants, this dimerization step
went largely unnoticed. Figure 5b depicts a time
course of actin polymerization monitored by covalent
crosslinking with N,N 8-1,4-phenylenebismaleimide
(1,4-PBM) (Knight and Offer, 1978). As polymerization proceeds, an initially formed 86-kDa apparent
molecular weight dimer (i.e., the so called ‘‘lower
dimer,’’ LD) is gradually consumed concomitant with
the formation of a 130-kDa apparent molecular
weight dimer (i.e., the so-called ‘‘upper dimer,’’ UD)
(Millonig et al., 1988; Steinmetz et al., 1997). Biochemical quantities of covalently crosslinked LD and
UD have been purified, and analytical ultracentrifugation has verified that both species are in fact
dimers (Millonig et al., 1988). According to protein
chemical analysis, Lys191 of one subunit is
crosslinked to Cys374 of another subunit in the UD
(Elzinga and Phelan, 1984). The LD crosslink most
likely occurs between Cys374 of both subunits (Millonig et al., 1988). The two covalently crosslinked
actin dimer species are drawn schematically in Fig.
5d, illustrating that the LD can migrate very similarly to a polypeptide chain of twice the molecular
mass of an actin monomer by SDS–PAGE, whereas
the UD cannot adopt a relaxed linear conformation
and hence exhibits an anomalous mobility by SDS–
PAGE.
LD and UD were assayed for their ability to
nucleate actin polymerization, to polymerize into
filaments, and to form paracrystalline and crystalline actin arrays (Millonig et al., 1988). As illustrated
schematically in Fig. 5e, these experiments have
suggested the following assembly pathways: As a
first and fast step, actin monomers form LD at least
as an intermediate under polymerizing conditions.
LDs might assemble further into crystalline arrays
such as sheets (see below and Fig. 7) or they might
dissociate again into monomers once the amount of
free monomer has dropped below its critical concentration for polymerization. In parallel and more
slowly, actin monomers form UD and larger oligomers (trimers, . . .) which efficiently nucleate F-actin
filament polymerization. Interconversion between
LD-based and UD-based supramolecular assemblies
has been observed (e.g., Aebi et al., 1981; Smith et al.,
1983), but it is unclear how this interconversion
occurs. As indicated schematically in Fig. 6c, while
being incompatible with the molecular architecture
of the F-actin filament the LD may be shared among
adjacent filaments (i.e., act as an interfilament dimer)
in actin filament bundles or paracrystalline filament
arrays (Millonig et al., 1988).
Using EM, we have recently documented that
during the process of filament elongation LD might
FIG. 5. Assaying actin polymerization. (a) Structural and functional polarity of the actin filament. Short actin filaments (e.g., after
shearing them by brief sonication or squeezing them through a 25-gauge syringe) have been stoichiometrically decorated with myosin S-1
and used to seed polymerization. The two structurally distinct ends (see Figs. 2 and 8) also differ functionally: the barbed end grows
faster—and hence longer—than the pointed end. (Inset) Upon addition of capZ, an F-actin capping protein (cf. Isenberg et al., 1980),
filament elongation at the barbed end is blocked, indicating that the capping protein preferentially binds to this end and thereby prevents
monomer addition. (b) Assaying actin polymerization by chemical crosslinking (cf. Millonig et al., 1988). Actin polymerization was induced
by adding MgCl2 to 2 mM and KCl to 50 mM. At the time points indicated, aliquots were removed and crosslinked with
N,N 8-1,4-phenylenebismaleimide (1,4-PBM). The samples were then run on 10.5% SDS–PAGE gels. M marks monomer, LD ‘‘lower dimer,’’
and UD ‘‘upper dimer.’’ (c) Assaying actin polymerization by pyrene fluorescence (cf. Cooper et al., 1983). Pyrene actin (i.e., rabbit muscle
actin pyrenated at Cys374) was polymerized with 2 mM MgCl2 and 50 mM KCl, and filament formation was monitored by the fluorescence
increase accompanying polymerization. (d) Schematic representation of the covalent structure of LD and UD produced by chemical
crosslinking of polymerizing actin with 1,4-PBM (see b). (e) Reaction scheme representing the multiple pathways of G-actin (i.e., via
oligomerization into LD and UD) to assemble into 2-D crystalline arrays (i.e., via LD) or F-actin filaments (i.e., via UD) (see also Fig. 6c).
Scale bar, 100 nm (a).
ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
directly be incorporated into growing F-actin filaments via one of its subunits. Gradually, the surplus
monomers dissociate from these ‘‘LD-decorated’’ filaments (propably upon switching of the incorporated
LDs from a G-like to an F-like conformation), thereby
yielding bona fide mature F-actin filaments at steady
state (Steinmetz et al., 1997). These data strongly
suggest that actin polymerization may involve multiple pathways rather than just the commonly assumed nucleation–elongation mechanism (see above).
Based on these recent findings, LD appears to be an
important intermediate in the actin polymerization
pathway and should therefore be considered in any
future models trying to describe this process. Interestingly, recent in vitro studies with gelsolin, actobindin, and swinholide A revealed that these three
actin-binding molecules were able to stabilize an
LD-type actin dimer (Hesterkamp et al., 1993; Bubb
et al., 1994a,b, 1995), thus further supporting the
functional significance of the LD in F-actin filament
polymerization and turnover (Steinmetz et al., 1997;
see also above). In this context, one may speculate
that during dynamic reorganization or turnover of
the actin cytoskeleton, a substantial amount of actin
is present in the form of an LD pool in the cell.
Moreover, incorporation of LD into growing F-actin
filaments might be important in building an actin
meshwork that can rapidly assemble and disassemble in highly motile cells and which might be a
prerequisite for the concerted action of actincrosslinking proteins.
Double-layered actin sheets (Smith et al., 1983)
and actin tubes (Fig. 6; Steinmetz et al., in preparation) are closely related 2-D crystalline actin arrays
(Fig. 7f) which form in the presence of stoichiometric
amounts of the trivalent lanthanide gadolinium. In
both cases, the unit cell of the respective 2-D crystals
contains two actin monomers which are related by a
P2 symmetry axis normal to the crystal plane.
Currently, we are determining the actin–actin interactions within the crystalline tubes at atomic scale
by merging the atomic structure of the G-actin
monomer (Kabsch et al., 1990) with an EM-based
307
3-D reconstruction of the actin tubes (Fig. 6b). In this
context, we are also investigating the possibility of a
structural relationship of the actin–actin contacts
within the sheet or tube dimer with the conformation
of the LD which forms predominantly at the onset of
actin polymerization (see above and Fig. 5c; also
Millonig et al., 1988) and may incorporate with one of
its subunits into growing F-actin filaments (Steinmetz et al., 1997).
Actin-binding proteins have been designed to
modulate actin polymerization, actin filament length,
and the steady-state concentrations of monomer and
polymer in vivo (reviewed in Pollard and Cooper,
1986; Condeelis, 1993; Kreis and Vale, 1993; Stossel,
1993; Hitt and Luna, 1994; Rozycki et al., 1994).
Most nonmuscle cells typically contain on the order
of 0.5 M (approximately 20 mg/ml) actin. Driven by
the ionic strength of the cytoplasm (i.e., 1–2 mM
Mg21 and 75–150 mM K1), this amount of actin
should be quantitatively polymerized into F-actin
filaments. Instead, typically 20–50% of the cytoplasmic actin is present in an unpolymerized form in
nonmuscle cells. Monomer-binding proteins (e.g.,
DNase I) lower the effective G-actin concentration by
sequestering monomers, thereby causing depolymerization of actin filaments. Some actin-binding proteins also catalyze exchange of ADP for ATP (e.g.,
profilin) or have a higher affinity for ATP– than for
ADP–G-actin (e.g., thymosin b4). Usually, proteins
that bind stoichiomerically to actin filaments stabilize them (e.g., tropomyosin, nonmuscle caldesmon).
Most capping proteins (e.g., capZ) bind to the barbed
end of F-actin filaments (see Fig. 5a), thereby slowing down polymer turnover. This, in turn, limits the
average filament length and hence the local viscosity
of the corresponding filament meshwork. Moreover,
it increases the critical concentration for polymerization (which is higher at the pointed end). Severing
proteins (e.g., gelsolin) fragment actin filaments and
bind to the barbed end of one of the fragments,
thereby having the same multiple effects as do
capping proteins (see above). In addition, they accelerate depolymerization by increasing the number of
FIG. 6. Crystalline actin tubes and how the LD and UD might fit into F-actin filaments, actin filament bundles or paracrystals, and 2-D
crystalline actin arrays. (a) High-magnification CTEM view of a stretch of a negatively stained crystalline actin (rabbit muscle) tube which
has been spread-flattened upon adsorption to the EM support film. The inset reveals a low-magnification overview of a crystalline actin
tube preparation. (b) 3-D reconstruction of a crystalline actin tube such as shown in (a) computed from a tilt series recorded from 260° to
160°. The reconstruction yields one unit cell containing two actin molecules related by a twofold axis of symmetry normal to the tube
surface. The surface-rendered actin tube dimer building block has been oriented so as to either exhibit the inside (left) or the outside (right)
of the tube to the viewer. For comparison, above and below the EM-reconstructed actin tube dimer the atomic structure of the actin
molecule (i.e., computed as a surface-rendered electron density map limited to 2 nm resolution) has been oriented so as to best delineate the
different views of the actin monomer within the tube dimer. (c) Attempts to molecularly build the UD and LD, and to fit these two distinct
dimer species into F-actin filaments (UD), actin filament bundles or praracrystals (LD), and crystalline actin sheets (LD). As a model of the
actin subunit, the schematic representation displayed in Fig. 3b was used. Modeling the UD and LD was based mainly on chemical
crosslinking data (Elzinga and Phelan, 1984; Millonig et al., 1988; see also Figs. 5b and 5d) in combination with the atomic structure of the
actin molecule (Kabsch et al., 1990; see also Fig. 3a). Scale bars, 50 nm (a), 2.5 nm (b).
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ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
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STEINMETZ ET AL.
pointed ends. Most severing proteins can sever phalloidin-stabilized actin filaments; however, actophorin
does not (Maciver et al., 1991). F-actin capping and
severing proteins can also stabilize nuclei and thus
increase the number of filaments that are formed.
3-D STRUCTURE, DYNAMICS, AND MECHANICAL
PROPERTIES OF F-ACTIN FILAMENTS:
FACTS AND FICTIONS
The structural disorder of F-actin is manifestated
by the variable crossover spacing and filament diameter along its length and by the local unraveling of
the two long-pitch helical strands (see Fig. 3h and
above). This structural disorder has been the subject
of many investigations and is now generally accepted to represent an intrinsic dynamic property of
the F-actin filament. To formally describe this dynamic behavior of the actin polymer, two models
have been proposed: (1) ‘‘random twist with cumulative angular disorder’’ (Egelman et al., 1982; Stokes
and DeRosier, 1987) and (2) ‘‘compensatory lateral
slipping’’ (Bremer et al., 1991), with the lateral
slipping motion of the two long-pitch helical strands
having both a radial and an angular component
(Censullo and Cheung, 1993). We have found that
the length and corresponding maximum width of
single crossovers determined from STEM ADF images of negatively stained F-actin filaments (Fig. 1b)
appear uncorrelated for many polymerization conditions assayed including stabilization by phalloidin
(Steinmetz et al., 1997). These findings are in accord
with the model of Bremer et al. (1991), and they
further indicate that cumulative lateral slipping
may indeed occur locally over short filament stretches
(i.e., 2–6 subunits). However, over longer filament
stretches local disorder or distortions compensate
each other so that there is no net propagation of
structural or mechanical perturbation along an actin
filament. Hence, we have to conclude that our findings do not support the results of Egelman and
co-workers which proposed allostery and long-range
cooperativity to occur in F-actin filaments (Egelman
and Orlova, 1995a,b), for example, caused by single
gelsolin molecules upon binding to the barbed end of
a filament (Orlova et al., 1995; Prochniewicz et al.,
1996).
To further investigate the structural basis of Factin dynamics, we have critically assessed the
fidelity and significance of a refined and averaged
3-D reconstruction (i.e., a ‘‘consensus’’ reconstruction) produced from a set of F-actin filaments polymerized from Ca–ATP–G-actin with 2 mM MgCl2
and 50 mM KCl. Specifically, we have investigated in
detail how resolution and suboptimal data processing steps may affect distinct structural features of
3-D reconstructions computed from negatively
stained F-actin filaments (Bremer et al., 1994). The
refined and averaged consensus reconstruction from
Bremer et al. (1994) is displayed in Fig. 8a surfacerendered so as to include 100% (left) or 30% (right) of
its nominal molecular mass. In Figs. 8b to 8d, we
have aligned this consensus reconstruction with a
ribbon representation of the atomic model by Holmes
et al. (1990): the fit is remarkable. Among other
structural features, this reconstruction strongly supports the prediction of Holmes et al. (1990) and
Lorenz et al. (1993), who modeled the hydrophobic
plug (see arrows in Fig. 8c) to extend across the
filament axis so as to insert in a plug-like fashion
into a hydrophobic pocket provided by the interface
between two adjacent subunits of the opposite long-
FIG. 8. Toward atomic interpretation of EM-based F-actin filament 3-D reconstructions. (a) An averaged (i.e., over 10 independent,
three-crossover long filament stretches; see Fig. 9a) F-actin filament 3-D reconstruction (see Bremer et al., 1994) surface-rendered to
include 100% (left) and 30% (right) of the nominal molecular mass. (b, c) Comparison of the EM-based consensus reconstruction shown in
(a) with an X-ray diffraction-based atomic model of the F-actin filament (cf. Holmes et al., 1990; Lorenz et al., 1993). The reconstruction was
aligned with the atomic model of the actin filament by Holmes et al. (1990), surface-rendered to include 100% of the nominal molecular
mass, and transparently overlaid onto a ribbon representation of one long-pitch helical strand of the atomic model. The views in (b) and (c)
are rotated by 110° relative to each other. (d) Testing a prediction made by the atomic model of the actin filament. For this purpose, the
reconstruction shown in (a) has been surface-rendered to include 30% of the nominal molecular mass and transparently overlaid onto a
ribbon representation of only the hydrophobic loops of the atomic model. Remarkably, the hydrophobic loop coincides with the ‘‘rung-like’’
intersubunit connectivity across the two long-pitch helical strands of the EM-based filament reconstruction. Note that the filaments in (c)
and (d) are identically oriented. This figure has been adapted from Figs. 7 and 8 of Bremer et al. (1994).
FIG. 12. Locating the phalloidin-binding site within F-actin filaments by stoichiometric binding of an undecagold–phalloidin complex
(i.e., Au11-phalloidin) to the filaments. (a, Top) STEM dark-field micrograph of an unstained, freeze-dried Au11-phalloidin stabilized F-actin
filament. (a, Bottom) As (a, top) but contrast adjusted to display the highest intensities only, i.e., those representing single undecagold
clusters (diameter ,1 nm). This image reveals discrete labeling along the two long-pitch helical strands which are staggered by half a
subunit (i.e., 2.75 nm). The average axial spacing between subsequent gold clusters along one and the same long-pitch helical strand
amounts to 5.5 nm, which corresponds to the axial entent of one actin subunit. (b) An averaged and refined 3-D helical reconstruction
computed from negatively stained Au11-phalloidin-stabilized F-actin filaments (see Fig. 1b) has been surface-rendered to include 100% of
its nominal molecular mass. The location of the undcagold clusters has been determined from a difference map (i.e., Au11-phalloidinstabilized F-actin filament reconstruction minus phalloidin-stabilized F-actin filament reconstruction) and visualized by 1-nm-diameter
gold spheres. Scale bars, 35 nm (a) and 2.75 nm (b).
ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
311
FIG. 7. Actin polymorphism (for details see Millonig et al., 1988). CTEM images of negatively stained samples of various polymeric or
crystalline in vitro assemblies induced from rabbit muscle actin. (a) Bona fide F-actin filaments; (b) actin filaments made of LD instead of
UD; (c) tube-like assemblies induced from a ,35-kDa proteolytic fragment of actin; (d) folded ribbons made of actin LD; (e)
stacked-disk-type actin–DNase I tubes (cf. Fowler et al., 1984); and (f) a mixture of crystalline actin sheets and tubes (cf. Aebi et al., 1981;
see also Fig. 6a). Scale bar, 100 nm (a–f). Reproduced from J. Cell Biol. 106, 785–796 by copyright permission of The Rockefeller University
Press.
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ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
pitch helical strand. When contoured to include 30%
mass, our consensus filament reconstruction reveals
a distinct mass density every 2.75 nm which connects the two long-pitch helical strands laterally like
the ‘‘rungs’’ of a ladder (Figs. 8a and 8d). Remarkably, the hydrophobic loop coincides with this distinct ‘‘bridge’’ of high mass density (Fig. 8d) which
emanates from the region close to the interface
between subdomains 3 and 4 (Fig. 8d, arrows). To
explain our observation of local unraveling (see Fig.
3h) within a double-stranded filament where hydrophobic loops emanate at regular intervals from one
long-pitch helical strand and insert into hydrophobic
pockets provided by the other strand, a molecular
‘‘zipper’’ model is best suited.
It is conceivable that different polymerization conditions may modulate the inherent structural dynamics of the F-actin filament or shift equilibria between
distinct structural/functional states and thus change
the overall filament conformation. Such conformational changes are in principle observable by EM.
However, different polymerization conditions may
produce ‘‘better’’ or ‘‘worse’’ looking filaments in
terms of how suitable they are for 3-D helical reconstruction. Therefore, investigation of subtle conformational changes requires a careful analysis of the
quality of the data and of the reliability of the
processing steps applied, to exclude artifacts which
may simply be caused by different effective resolution of the individual data sets. We addressed this
issue by taking advantage of the increased contrast,
the better signal-to-noise ratio, and the linear contrast transfer characteristics of STEM ADF over
CTEM BF images (Engel and Colliex, 1993) of negatively stained specimens (Fig. 1b) (Bremer et al.,
1994). Obviously, the phenotypes of the individual
refined reconstructions which go into an averaged
reconstruction do affect the phenotype of the average: Figs. 9a and 9b display the 10 three-crossover
long filament stretches that were included in the
refined and averaged consensus reconstruction by
Bremer et al. (1994): they have been surface-
313
rendered to include 100% (a) or 30% (b) of their
nominal molecular mass. In Fig. 9b, these 10 refined
reconstructions have been classified according to the
strength of their intersubunit contacts between the
two long-pitch helical strands: the type A reconstructions reveal little if any connectivity, the type B
reconstructions yield interrupted connectivity, and
the type C reconstructions exhibit uninterrupted
connectivity between the two long-pitch helical
strands. It is worth mentioning that most of the
individual reconstructions look quite similar when
contoured to include 100% mass: in Fig. 9a, compare,
e.g., the type A filament reconstructions with those of
the type C. As pointed out above, producing averages
of the different filament types (Fig. 9c: SA, SB, and
SC) yields subaverages that accentuate the structural features which were initially chosen on the
basis of the overall average (Fig. 9c: S10) as ‘‘earmarks’’ for classification. As with the individual
reconstructions (compare Fig. 9a with Fig. 9b), the
significant differences observed for the averages of
the different filament types at the 30% contouring
level (Fig. 9c), however, become almost invisible at
the 100% contouring level (Fig. 9d).
In addition, we have systematically investigated
the phenotypes of reconstructions that are produced
upon deviation from the optimal helical parameters
or by lowering the nominal resolution of the reconstruction (cf. Bremer et al., 1994). For example, as
revealed in Fig. 10a, small deviations from the
optimal radial position of the helix axis (i.e., 0.1–1.0
nm) yield a substantial redistribution of mass from
several parts of the filament surface into the area
between the two long-pitch helical strands (i.e., see
buildup of a high mass density ‘‘bridge’’ connecting
the two long-pitch helical strands marked by an
arrow). Hence the detailed morphology of the intersubunit contact pattern between the two long-pitch
helical strands is extremely sensitive to small deviations from the optimal radial coordinate of the
filament axis. In fact, it is remarkable that a shift of
as little as 0.06 nm of the radial coordinate of the
FIG. 9. Variations of the intersubunit contact pattern among different F-actin filaments. (a) 3-D reconstructions of 10 three-crossover
long F-actin filament stretches surface-rendered to include 100% of their nominal molecular mass are displayed and enumerated 1 through
10. (b) The same 10 reconstructions are shown surface-rendered to include 30% of their nominal molecular mass, and they have been
classified into three distinctive types of reconstructions according to their intersubunit contact pattern: Type A exhibits strong long-pitch
helical intersubunit contact and little if any contact between the two strands. Type B reveals continuous strong contact along and
interrupted contact between the two long-pitch helical strands. Finally, Type C yields predominant contact between the two strands. (c)
The reconstructions within each of the three types A, B, and C shown in (b) were averaged, and the resulting subaverages SA, SB, and SC
are compared to the overall average S10, all surface-rendered to include 30% of the nominal molecular mass. Arrows identify areas that
clearly differ among the subaverages besides the relative mass density of the intersubunit contacts between the two long-pitch helical
strands (marked by arrowheads) that was used to classify the reconstructions in the first place. (d) The subaverages SA, SB, and SC and
the overall average S10 are displayed surface-rendered to include 100% of the nominal molecular mass. In S10, the arrowhead indicates
the interdomain cleft that separates the larger inner (i.e., subdomains 3 and 4) from the smaller outer (i.e., subdomains 1 and 2) domain
(see Figs. 3a and 3b), whereas the arrow points at the interstrand contact interface. This figure has been adapted from Figs. 4a and 5 of
Bremer et al. (1994).
a
FIG. 10. Phenotypes of F-actin filament reconstructions resulting from deviations from the optimal helical parameters or from different
nominal resolutions (for details, see Bremer et al., 1994). (a) Effects of deviating from the optimal radial coordinate of the filament axis on
the phenotype of the corresponding F-actin filament 3-D reconstructions. For this purpose, the 10 three-crossover long F-actin filament
stretches (see Fig. 9a) have been reconstructed with the filament axis radially shifted as indicated, aligned and averaged, and are compared
to the optimal reconstruction S10 as displayed in Figs. 8a and 9d). The arrow highlights the connectivity between subdomain 4 of one
subunit from one long-pitch helical strand with subdomain 1 of the closest subunit from the other strand. The arrowhead points to a region
of the interstrand interface where structural detail deteriorates when radially displacing the filament axis from its optimal position. (b)
Effect of lowering the nominal isotropic resolution on the phenotype of the F-actin filament 3-D reconstructions. The nominal resolution of
S10 as shown in Figs. 8a and 9d has been reduced from 2.5 to 3.5 or 4.5 nm as indicated. Whereas the arrows highlight the same feature as
in (a), the arrowheads point to the interdomain cleft. All reconstructions in (a) and (b) are shown surface-rendered to include 100% (top in a,
bottom in b) or 30% (bottom in a, top in b) of the nominal molecular mass. This figure has been adapted from Fig. 6 of Bremer et al. (1994).
ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
filament axis (i.e., within a 3-D reconstruction to a
nominal resolution of 2.5 nm) yields distinct changes
of the corresponding reconstruction (see Fig. 10a).
Moreover, lowering the nominal resolution has a
similar effect but, at the same time, it weakens the
long-pitch helix intersubunit contact (see arrow in
Fig. 10b). In addition, lowering the resolution alleviates the cleft between the large and the small
domains of the actin subunit (see arrowhead in Fig.
10b).
We have also assessed the effect of different divalent cations (i.e., Mg21 or Ca21) bound to the highaffinity divalent cation binding site (HAS) of the
G-actin molecule on its polymerization kinetics and
the oligomers that form upon addition of salt, as well
as the mechanical properties and 3-D structure of
the resulting F-actin filaments (Steinmetz et al.,
1997). As illustrated in the left panel (top) of Figs.
11a and 11b, monitored by the increase of the
fluorescence signal of pyrenated actin (i.e., at
Cys374), the lag phase of Ca–ATP–G-actin appears
much more pronounced than that of Mg–ATP–Gactin, while the elongation rate remains similar for
both conditions. Both low-magnification views (i.e.,
middle panels (top left) of Figs. 11a and 11b) as well
as high-magnification views (i.e., middle panels (top
right) of Figs. 11a and 11b) document that the gross
morphologies of the two types of F-actin filaments
are very similar. The flexibility of the actin filaments
was determined from the average crossover spacing
according to Bremer et al. (1991) (i.e., middle panels
(bottom left) of Figs. 11a and 11b), and the apparent
persistence length was measured according to Orlova and Egelman (1993) (i.e., based on Landau and
Lifshitz (1958); see middle panels (bottom right) of
Figs. 11a and 11b). These data clearly document that
the type of divalent cation bound to the HAS of
G-actin does not significantly affect the mechanical
properties of the resulting F-actin filaments, a finding being consistent with results derived from thermal fluctuations of F-actin filaments in solution
(Isambert et al., 1995). These results are strongly
supported by our refined and averaged 3-D reconstructions of corresponding F-actin filaments. As
illustrated in the right panels of Figs. 11a and 11b,
no significant structural differences were revealed
between Mg– and Ca–F-actin (for details, see Steinmetz et al., 1997). In contrast, Egelman and coworkers (Orlova and Egelman, 1993; Egelman and
Orlova, 1995b) aimed to document that F-actin
flexibility can be increased up to four times, depending on the ionic conditions applied. Also, their corresponding 3-D helical reconstructions revealed substantial structural differences, a finding which our
data failed to confirm (right pannels of Figs. 11a and
11b).
315
We have also investigated the effect of phalloidin
on actin polymerization. For this purpose, Ca–ATP–
G-actin was polymerized with 100 mM KCl in the
presence of a 2:1 molar excess of phalloidin over
actin (Fig. 11c). As documented by the time course of
the pyrene fluorescence increase in the left panel
(top) of Fig. 11c, phalloidin drastically shortens the
lag phase of Ca–ATP–G-actin, and it accelerates the
elongation rate by a factor of about 2 (compare with
Fig. 11b). Filaments polymerized from Ca–ATP–Gactin in the presence of phalloidin (Fig. 11c, middle
and right panels) differ from their native counterparts (compare with Fig. 11b) mainly in three aspects (for details and discussion, see Steinmetz et al.,
1997): (1) As revealed by the middle panel of Fig. 11c,
their mean crossover spacing increases by 2 nm
(bottom left), and their apparent persistence length
more than doubles (bottom right). (2) As marked by
the arrowhead in the right panel of Fig. 11c (filament
reconstruction contoured to include 100% mass), the
distinct ‘‘groove’’ residing between the two long-pitch
helical strands becomes ‘‘padded’’ with additional
mass. And (3) as indicated by the two arrowheads in
the right panel of Fig. 11c (filament reconstruction
contoured to include 30% mass), the intersubunit
contacts both along and between the two long-pitch
helical strands become more massive. Our EM-based
3-D reconstructions of phalloidin-stabilized Ca–Factin filaments (see also Steinmetz et al., 1997) are
in good qualitative agreement with a corresponding
atomic model computed by Lorenz et al. (1993).
Lorenz et al. (1993) also determined the possible
binding site of phalloidin within their atomic filament model. Their placement of the toxin molecule is
in agreement with its EM-based localization (Bremer
et al., 1991; refined by Steinmetz et al., 1997; see also
below), its chemical mapping (Vandekerckhove et al.,
1985), and its localization predicted by mutational
analysis (Drubin et al., 1993).
Recently, we have engineered an undecagoldtagged phalloidin derivative (Au11-phalloidin; in collaboration with Dr. H. Faulstich, Max-Planck Institute for Medical Research, Heidelberg, Germany) in
order to directly map the binding site of this bicyclic
heptapeptide within the F-actin filament (Steinmetz
et al., manuscript in preparation). As documented in
Fig. 12a, the ,1-nm-diameter gold clusters can be
visualized directly by STEM ADF of unstained freezedried Au11-phalloidin-stabilized F-actin filaments.
Remarkably, the gold particles are lining up the two
long-pitch helical strands at distinct, 5.5-nm intervals and are axially spaced by 2.75 nm (i.e., due to
the 2.75-nm axial stagger of the two long-pitch
helical strands). At first glance, a refined and averaged 3-D reconstruction computed from STEM ADF
images of negatively stained Au11-phalloidin-stabi-
316
STEINMETZ ET AL.
ACTIN FILAMENT STRUCTURE, POLYMERIZATION, AND POLYMORPHISM
lized F-actin filament stretches (Fig. 12b) looked
indistinguishable from a corresponding 3-D reconstruction computed from just phalloidin-stabilized
F-actin filaments (see Fig. 11c, right panel). However, a difference map computed from these two
filament reconstructions after proper scaling yielded
a single positive mass peak per actin subunit on the
filament surface. In Fig. 12b, the difference peaks
are shown mapped onto the refined and averaged
Au11-phalloidin-stabilized F-actin filament 3-D reconstruction as 1-nm-diameter gold spheres. As the
Au11-cluster is attached to the bicyclic heptapeptide
via a ,1.7-nm-long linker, the gold particles are
expected to appear displaced from the actual phalloidin-binding sites along the actin filament. Taking
the location of the phalloidin molecule within the
F-actin filament as proposed by Lorenz et al. (1993),
we have fitted an atomic model of the undecagold–
phalloidin complex into our EM-based Au11-phalloidin-stabilized F-actin filament 3-D reconstruction.
Whereas Lorenz’s orientation of the phalloidin molecule caused the attached gold cluster to sterically
collide with an adjacent actin molecule, rotation of
the phalloidin moiety by approximately 180° about
an axis roughly parallel to the filament axis yielded
and almost perfect coincidence of the gold cluster
with the unique difference peak such as shown in
Fig. 12b.
In summary, we have optimized our F-actin filament preparation and EM data aquisition, and we
have improved the fidelity and reproducibility of our
semiautomated helical data processing protocol.
Taken together, these steps have become a powerful
and valuable tool, particularly to more systematically assess distinct conformational states of the
F-actin polymer in response to various effectors (e.g.,
the type of the tightly bound divalent cation, the
state of hydrolysis of the bound nucleotide, and the
binding of small drugs such as phalloidin, etc.) or to
more faithfully determine the 3-D structure of actin
filaments in complex with actin-binding or regulatory proteins and molecules.
CONCLUDING REMARKS
Our recent advances in specimen preparation,
imaging techniques, and data analysis, processing,
and merging now open exciting new avenues toward
317
a more profound understanding of the structural
basis and the complex mechanisms which underlay
the process of actin polymerization, dynamics, and
network formation. Our integrated approach presented here allows for a direct correlation of structural data with biochemical findings and mechanical
properties to systematically investigate the distinct
steps involved in F-actin assembly, dynamics, and
turnover. Our novel polymerization data indicate
that F-actin filament formation involves additional
pathways besides the commonly proposed nucleation–condensation mechanism, and they suggest a direct
involvement of the so-called ‘‘lower dimer’’ (LD) in
the polymerization process. Currently, we are investigating the structure and the exact role of the LD
during actin filament assembly, dynamics, and turnover, as well as its functional significance in vivo. We
have now refined our data analysis and processing
methods so as to produce averaged EM-based 3-D
reconstructions which, in combination with X-ray
diffraction data, allow us to analyze the F-actin
filament structure to atomic detail. This integrated
approach represents a powerful tool to assess distinct conformational states of the actin polymer and
to determine the 3-D structure of the F-actin filament in functionally important complexes with actinbinding proteins and even small effector molecules
such as phalloidin. By combining the power of X-ray
diffraction and/or nuclear magnetic resonance spectroscopy data of individual components at atomic
resolution with EM-based 3-D reconstructions of
entire supramolecular assemblies at intermediate
resolution, this approach is generally applicable and
plays an invaluable role in building and refining
atomic models. In the case of actin, we will continue
to systematically apply these methods to eventually
solve the atomic structure of the F-actin filament
and to gain more detailed insight into its dynamic
properties and functional involvement in various
cellular processes.
We thank Drs. Ron Milligan, Roger Wepf, and Mary Reedy for
allowing us to use some of their EM data, and we are grateful to
Dr. Cora-Ann Schoenenberger for critical reading of the manuscript. This work was supported by the Canton Basel-Stadt and
the M.E. Müller Foundation of Switzerland, by a research grant
from the Swiss National Science Foundation (31-39691.93), and
FIG. 11. Correlation between G-actin conformation, oligomerization, and polymerization and F-actin mechanical properties and 3-D
structure: Mg– and Ca–ATP–G-actin were polymerized with 100 mM KCl (a, b), and Ca–ATP–G-actin was polymerized with 100 mM KCl
in the presence of a 2:1 molar ratio of phalloidin over actin (c). For each of these three polymerization conditions, the following data were
collected: a polymerization time course monitored by pyrene fluorescence (left panel, top), and by intermolecular crosslinking with 1,4-PBM
(left panel, bottom); a low-magnification CTEM (middle panel, top left), and a high-magnification STEM ADF (middle panel, top right) view
of negatively stained F-actin filaments; a crossover spacing frequency distribution (middle panel, bottom left) and an apparent persistence
length plot (middle panel, bottom right); and an averaged 3-D reconstruction surface-rendered to include either 100% (right panel, left) or
30% (right panel, right) of the nominal molecular mass. This figure has been composed from data reported by Steinmetz et al. (1997).
318
STEINMETZ ET AL.
by a postdoctoral fellowship awarded to A. Bremer by the Human
Frontier Science Program (HFSP).
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