вход по аккаунту



код для вставкиСкачать
Cell Motility and the Cytoskeleton 42:73–81 (1999)
Effect of Capping Protein, CapZ, on the
Length of Actin Filaments and Mechanical
Properties of Actin Filament Networks
Jingyuan Xu,1 James F. Casella,2 and Thomas D. Pollard3,4*
1Department of Biophysics and Biophysical Chemistry,
Johns Hopkins School of Medicine, Baltimore, Maryland
2Department of Pediatrics, Division of Pediatric Hematology,
Johns Hopkins School of Medicine, Baltimore, Maryland
3Department of Cell Biology and Anatomy,
Johns Hopkins School of Medicine, Baltimore, Maryland
4Salk Institute for Biological Studies, La Jolla, California
We report on how physiological concentrations of capping protein shorten actin
filaments and on the remarkably fluid nature of solutions of such short filaments
even at the high concentrations that exist in cells. We measured the lengths of actin
filaments formed by spontaneous polymerization of highly purified actin monomers by fluorescence microscopy after labeling with rhodamine-phalloidin. The
length distributions are exponential with a mean of about 7 µm (2600 subunits). As
observed previously with less quantitative assays, copolymerization with the actin
capping protein, CapZ, reduces the length of the filaments. At cellular concentrations of capping protein, one filament forms for each molecule of capping protein
and the population of filaments is uniformly short. Using CapZ to vary the length
of actin filaments, we measured how their mechanical properties depend on length.
The stiffness (elastic modulus) of actin filament networks depends steeply on the
length, with long filaments contributing far out of proportion to their numbers to
the stiffness. Even at physiological concentrations (300 µM), networks of filaments
limited to lengths observed in cells with a 1 to 500 molar ratio of CapZ are more
fluid and much less elastic than lower concentrations of longer actin filaments.
Thus the high concentration of short actin filaments in cells must be crosslinked to
produce the observed stiffness of the cortex. Cell Motil. Cytoskeleton 42:73–81,
1999. r 1999 Wiley-Liss, Inc.
Key words: actin; capping protein; CapZ; filaments; mechanical properties; rheology
Actin filaments are major components of the cytoskeleton, which provides the physical basis for the
mechanical properties of cytoplasm. A key consideration
is how actin filaments themselves, exclusive of any
crosslinking proteins, contribute to mechanical properties. To relate physical measurements on purified actin
filaments to conditions inside cells, one must consider the
length of actin filaments.
Polymer length is an important determinant of
mechanical properties [Hvidt and Janmey, 1990; Janmey
r 1999 Wiley-Liss, Inc.
Contract grant sponsor: NIH; Contract grant number: GM-26338;
Contract grant number: AR-40697.
Jingyuan Xu is currently at Department of Chemical Engineering,
Johns Hopkins University, Baltimore, Maryland.
*Correspondence to: Thomas D. Pollard, Salk Institute for Biological
Studies, 10010 N. Torrey Pines Road, La Jolla, CA 92037.
Received 14 September 1998; accepted 10 October 1998
Xu et al.
et al., 1994] and many [Cano et al., 1991; Podolski and
Steck, 1990], but not all [Small et al., 1995; Svitkina et
al., 1997] cellular actin filaments appear to be shorter, ⬍1
µm, than filaments polymerized in vitro from pure actin
monomers. The most thorough study of this point used
gelsolin to control polymer length and concluded ‘‘relatively isotropic F-actin networks are sufficiently strong to
stabilize cells’’ [Janmey et al., 1994]. This hypothesis
suggests that actin filaments at high concentration, estimated to be ⬎200 µM in the cell cortex [Niederman et al.,
1983], might by themselves account for a major part of
the stiffness of the cytoplasm exclusive of crosslinking by
actin-binding proteins.
Given the importance of this point for understanding the physical basis of cytoskeletal function, we reexamined the mechanical properties of actin filaments, using
the muscle capping protein CapZ [Casella et al., 1986,
1987, 1995] to regulate polymer length. CapZ and its
relatives in nonmuscle cells bind with high affinity and
block the fast growing (barbed) end of actin filaments for
both elongation and annealing [Isenberg et al., 1980;
Cooper and Pollard, 1985; Casella et al., 1987; Casella
and Torres, 1994; Caldwell et al., 1989]. Capping proteins
are associated with the barbed ends of actin filaments in
the Z-line of skeletal muscle [Casella et al., 1987] and
block many, if not most, barbed ends of actin filaments in
nonmuscle cells [DiNubile et al., 1995] Capping proteins
also nucleate actin polymerization by stabilizing small
actin oligomers [Cooper and Pollard, 1985; Caldwell et
al., 1989]. The physiological relevance of this nucleating
activity is still in question, because growth is only in the
slow direction. These activities allow capping proteins to
reduce the length of actin filaments. This was shown by
electron microscopy and by qualitative viscosity assays
[Isenberg et al., 1980; Cooper et al., 1984], but the
dependence of the polymer length on capping protein
concentration has not been measured reliably.
We use fluorescence microscopy of actin filaments
labeled with rhodamine-phalloidin [Burlacu et al., 1992]
to establish how copolymerization of highly purified actin
with CapZ affects the length of the filaments. At physiological concentrations each CapZ molecule produces
one filament. By varying the CapZ concentration we
prepared filaments that ranged in mean length from 1.3
µm to 6.7 µm for rheological analysis. The elastic
modulus depends strongly on the mean length. Similar
experiments with gelsolin [Janmey et al., 1994] showed
the same trend, but much higher values at all polymer
lengths. Collaborative experiments with those investigators [Xu et al., 1998] showed that the lower values in the
current paper are more reliable. For the first time, we also
measured the mechanical properties of physiological
concentrations (300 µM) of short (1.3 µm) actin filaments. They have a much lower elastic modulus and are
much more fluid than low concentrations of long filaments formed by spontaneous assembly in vitro. We
conclude, that without crosslinking, the short actin filaments observed in cells cannot account for the stiffness of
the cell cortex.
Buffer G contained 0.2 mM ATP, 0.5 mM dithiothreitol, 0.1 mM CaCl2, 1 mM sodium azide, 2 mM Tris-Cl,
pH 8.0 at 25°C. Concentrated polymerizing buffer
(10 ⫻ KME) contained 500 mM KCl, 10 mM MgCl2, 10
mM EGTA, and 20 mM Tris-Cl, pH 8.0 or 100 mM
imidazole pH 7.0 at 25°C. The Kron et al. [1991]
fluorescence microscopy buffer contained 50 mM KCl, 1
mM MgCl2, 100 mM dithiothreitol, 20 µg/ml catalase, 0.1
mg/ml glucose oxidase, 3 mg/ml glucose, and 2 mM
Tris-Cl, pH 8.0 or 10 mM imidazole pH 7.0 at 25°C.
Protein Purification
Actin was purified from rabbit skeletal muscle
[MacLean-Fletcher and Pollard, 1980]. Each gram of
acetone powder was extracted for 30 min on ice with 20
ml of Buffer G. Insoluble material was pelleted by
centrifugation at 16,000 rpm at 2°C for 30 min and the
pellet resuspended in same volume of Buffer G and
immediately centrifuged. The supernatants were combined and the actin was polymerized by the addition of
MgCl2 to 2 mM and KCl to 50 mM. After 1 h, KCl was
added to 0.8 M and the actin filaments pelleted by
centrifugation at 100,000 g for 2 h at 4°C. The filaments
were depolymerized by dialysis against Buffer G changed
daily for 2 days at 4°C and any remaining filaments were
pelleted by ultracentrifugation. The top two thirds of the
supernatant was gel filtered on a 2.5 ⫻ 110 cm column of
Sephacryl S-300 equilibrated with Buffer G. The fractions at and following the peak were pooled, stored by
dialysis vs. daily changes of fresh buffer G and used
within 5 days. To remove traces of CapZ from this singly
gel filtered actin, the fractions beginning at the midpoint
of the leading edge of the actin peak were pooled,
repolymerized with 50 mM KCl and 2 mM MgCl2,
pelleted, depolymerized and gel filtered a second time
[Casella et al., 1995]. We purified CapZ from chicken
skeletal muscle acetone powder using a KI extraction,
followed by DEAE chromatography, hydroxylapatite
chromatography, gel filtration and sucrose density gradient centrifugation [Casella et al., 1986].
Measurement of Actin Filament Lengths
We stabilized fully polymerized actin filaments
with CapZ (to block the rapidly depolymerizing barbed
end) and with rhodamine-phalloidin (to reduce subunit
Actin Filament Lengths and Mechanical Properties
dissociation at both ends to near zero [Collucio and
Tilney, 1984; Sampath and Pollard, 1991]). Actin was
polymerized by mixing one part of concentrated polymerizing buffer 10 ⫻ KME with nine parts of actin in Buffer
G. After 3 h incubation, we added one CapZ per 500 actin
subunits and one rhodamine-phalloidin (Molecular Probes,
OR) per actin subunit and diluted the sample to 0.3 µM
with fluorescence buffer [Burlacu et al., 1992; Kaufmann
et al., 1992; Käs et al., 1996]. After incubation at room
temperature for 30 min to allow rhodamine-phalloidin
binding [De La Cruz and Pollard, 1994], the labeled actin
was diluted to 2–10 nM with fluorescence buffer, about
10 µL of solution was placed on a microscope slide and
covered with a 20 mm square coverslip coated with
nitrocellulose. To minimize shearing and artifactual fragmentation of filaments during manipulations, we trimmed
the tip of the plastic pipette tip [Burlacu et al., 1992;
Janmey et al., 1994]. We did not fix with aldehydes,
because they damage actin filaments [Lehrer, 1972].
We observed filaments with a Leitz Orthoplan
microscope equipped with a 3-mm BG-38, KP 560 (short
wavelength pass interference filter), 2-mm BG-36 (excitation filter), TK-580 (dichroic mirror), two K-580 (colored
glass barrier filters) and an Olympus 100⫻ (NA 1.25)
objective. We recorded images on Kodak 3200 blackwhite professional film with an exposure of 30 to 60 s.
Images of the filaments were clear enough to measure
filament lengths ⱖ0.3 µm manually on prints at a final
magnification of 3,150⫻. Two independent observers
measured the same length distribution and number average length. In samples with predominantly short filaments ⬍2 µm long, many filaments appeared as fluorescent spots rather than asymmetrical rods, so we measured
their lengths by densitometry. Negative films were digitized with Adobe Photo Shop 3.0. Taking a filament ⬎2
µm long as an internal standard for intensity per unit
length, we used NIH Image 1.6 to measure the intensity
of each fluorescent spot in the whole population on the
same negative. The background was subtracted from the
areas containing each fluorescent filament. The number
average length (Ln ) is defined as Ln ⫽ (1/n)⌺li, where n is
the number of filaments and li is the length of each
filament. The length distributions were approximately
exponential rather than Gaussian, so standard deviation
could not be used to describe the variability. For an
exponential distribution, the fraction of filaments (fi ) with
length l is fi ⫽ ␭ exp (⫺␭li ), the mean length is l/␭ and the
variance (li ) ⫽ (l/␭) 2.
Actin filaments networks are viscoelastic, having
both solid (elastic) and fluid (viscous) properties. They
both store and dissipate mechanical energy. Therefore,
their mechanical properties can be described by rheologi-
cal parameters. For oscillatory deformations, the viscoelasticity is characterized by dynamic elasticity G8 (also
called dynamic storage modulus) and loss shear modulus
G9. G8 is the in-phase ratio of stress/strain and G9 is out of
phase component. These relationships can also described
by the complex modulus 0G*0 ⫽ (G82 ⫹ G92 ) 1/2 and phase
shift ␦ ⫽ tan⫺1 (G9/G8) [Ferry, 1980]. Because G8 ⬎ G9
for actin filaments, 0G*0 mainly reflects the value of G8. A
solid has a phase shift of 0. A viscous liquid has phase
shift of 1.6 rad.
The rheological measurements were made with a
parallel plate Rheometrics RFS II rheometer (Rheometrics, NJ) in the small amplitude (strain ⱕ 2%), forced
oscillation mode [Sato et al., 1985]. Monomeric actin in
Buffer G was mixed with one-tenth volume of 10 ⫻ KME
and immediately placed between the metal plates of the
rheometer to polymerize at 25°C. The plates were sealed
with mineral oil (Sigma, St. Louis, MO) to prevent
sample dehydration. Measurements of G8 and G9 were
made every 30 s using time sweep mode to observe the
gel formation. After G8 and G9 reached a plateau,
frequency sweep mode was used to measure the rheological parameters.
Actin Filament Lengths
We assume that the population of fluorescent filaments attached to a coverslip is equivalent to the population in solution, because within several minutes all
detectable fluorescent filaments in the 25 µm gap between
slide and cover slip attached to the nitrocellulose-coated
glass cover slip, leaving no filaments free in the solution
(Fig. 1). However, the sample on the coverslip underestimates slightly the distribution of lengths in solution,
because about 10% of longer filaments broke into two to
four segments as they bound to the coverslip. The method
may also miss some filaments less than 0.2 µm long due
to their faint fluorescence. The polymers were stable,
since the length distributions and average lengths did not
change between 3 h and 2 days after polymerization.
When rhodamine-phalloidin labeled actin filaments bound
to coverslips coated with N-ethylmaleimide-inactivated
rabbit skeletal muscle myosin [Warshaw et al., 1990],
their length distribution and number average length were
the same as filaments on nitrocellulose.
Filaments assembled by spontaneous polymerization from doubly gel filtered actin monomers at 24 µM
varied in length from less than 0.3 µm to nearly 100 µm
(Figs. 1A, 2A). The distribution of lengths was approximately exponential with a mean of 6.7 µm. The mean
length was about 20% lower without a second cycle of
purification to remove traces of capping protein [Casella
et al., 1995], even though we used only the fractions from
Xu et al.
Fig. 1. Fluorescence micrographs of filaments of doubly gel filtered actin labeled with rhodaminephalloidin. Conditions: 24 µM purified actin, 50 mM KCl, 1 mM MgCl2, 1 mM EGTA, 180 µM ATP, 0.45
mM DTT, 90 µM CaCl2, 0.9 mM azide, 4 mM Tris-Cl (pH 8.0), 22°C for 3 h. CapZ concentrations: zero
(A); 48 nM (B).
the top of the actin monomer peak from the first gel
filtration column to avoid CapZ, which runs ahead of
actin monomers.
CapZ reduced the length (Fig. 1B) of filaments
assembled from twice gel filtered actin and made their
length distributions more uniform (Fig. 2B–D). Above 20
nM, one filament formed for each CapZ molecule (Fig. 3).
Lower concentrations of CapZ increased the number of
filaments over that produced by spontaneous polymerization, but the number concentration of filaments exceeded
the concentration of capping protein, so some of the
filaments were not capped.
Very high concentrations of short actin filaments are
found in vivo [Small, 1981; Podolski and Steck, 1990;
Cano et al., 1991], so we examined the mechanical
properties of high concentrations of mixtures of actin
with a 1:500 ratio of CapZ (Fig. 6). These short, 1.3 µm
filaments were more fluid-like than actin alone. The phase
shift ␦ was in the range of 0.5 to 0.6 rad at 1 Hz compared
with about 0.3 rad for pure actin. Even at 300 µM actin
the complex modulus 0G*0 was less than 2 Pa. This is
value of the complex modulus 0G*0 observed for 30–40
µM pure actin polymerized in the absence of CapZ.
Mechanical Properties of Actin Filament Networks
Capping Protein Regulation of Polymer Length
Actin filaments alone formed a weak gel with a
complex modulus 0G*0 that depended on the frequency of
oscillation (Fig. 4). Two preparations of doubly gel
filtered actin had the same complex modulus as singly gel
filtered actin filaments, while one preparation had a
slightly higher complex modulus (Fig. 4A). The phase
shift ␦ of all three preparations was around 0.3 rad
(Fig. 4B). Thus, the low concentration of CapZ contaminating actin after one cycle of polymerization and gel
filtration [Casella et al., 1994] had only a small effect on
the mechanical properties of actin filament networks.
Titration of pure actin with CapZ reduced 0G*0
(Fig. 4A) and increased ␦ (Fig. 4B). The complex
modulus depended on the mean length of the filaments
(Fig. 5). A fourfold difference in mean length (1.3 vs. 5.5
µm) resulted in a 30-fold difference in the complex
modulus at a frequency of 0.01 Hz and 10-fold difference
at 0.1–1.0 Hz. The uniform, short filaments behaved more
like a viscoelastic liquid than the exponential distribution
of longer pure actin filaments. The presence of CapZ also
increased the variability in the rheological measurements.
We confirm that copolymerization of actin monomers with capping protein, CapZ in this case, limits the
length of the filaments at steady state. Earlier measurements by electron microscopy showed that actin filaments
are shorter when polymerized in the presence of amoeba
capping protein [Isenberg et al., 1980; Cooper et al.,
1984] and muscle CapZ [Caldwell et al., 1989] but length
distributions were not reported. Since negative staining
underestimates the length of control actin filaments, our
measurements by light microscopy are the only reliable
quantitative assessment of the effect of CapZ on actin
filament length.
At CapZ concentrations above the Kd (0.5–1 nM or
less [Casella et al., 1986, 1987; Caldwell et al., 1989])
and at ratios of actin to CapZ up to 1,000, one filament
forms for every CapZ present during polymerization
(Fig. 3). This is expected from the high affinity of CapZ
for the barbed end of actin filaments and the ability of
capping proteins to promote spontaneous polymerization
of actin monomers by stabilizing transient oligomers
Actin Filament Lengths and Mechanical Properties
Fig. 2. Length distributions of doubly gel filtered actin filaments as a function of CapZ concentration.
Conditions as in Figure 1. CapZ concentrations: zero (A); 4.8 nM (B); 12 nM (C); 48 nM (D). Smooth
curves are the best exponential fits to the data.
[Cooper and Pollard, 1985; Caldwell et al., 1989]. Since
these filaments form rapidly, grow slowly only at their
pointed ends and do not anneal, the steady state population of filaments is short and uniform in size.
When the ratio of actin to CapZ exceeds 1,000,
more filaments form than the concentration of CapZ. This
is expected, because the rate of self-nucleation will
exceed the rate of CapZ-mediated nucleation at low CapZ
concentrations. At a ratio of 5,000 actins per CapZ about
30% of the filaments are capped and 70% are not.
From comparison of length distributions of once
and twice gel filtered actin, we estimate that our singly gel
filtered actin is contaminated with about one part in 104 of
capping protein, similar to that estimated by immunoassay [Casella et al., 1995]. Since the first gel filtration
column removes most of the capping protein activity
[MacLean-Fletcher and Pollard, 1980; Casella et al.,
1987, 1995], actin that has not been gel filtered is more
heavily contaminated.
The dependence of polymer length on the concentration of other capping proteins shows that gelsolin, villin
and scinderin all have a high capacity to nucleate actin
filaments. At molar ratios up to 250 actins per gelsolin,
the length distributions measured by negative staining
Xu et al.
Fig. 3. Dependence of the number average length (Ln ) of actin
filaments on concentration of CapZ. Conditions as in Figure 1.
and electron microscopy show that exactly one filament
forms per gelsolin [Janmey et al., 1986]. At higher ratios
of actin to gelsolin, more than one filament forms per
gelsolin. Spontaneous polymerization is one source of the
excess filaments, but two other factors may contribute.
First, CapZ contaminating once gel-filtered actin will be a
significant fraction of the total nucleating activity at low
concentrations of gelsolin. Second, any breakage during
specimen preparation will over estimate of filament
number. Mixing gelsolin with preformed actin filaments
produced about one filament per gelsolin at ratios of
1,000 according to a light microscopic assay [Burlacu et
al., 1992]. However, a ratio of 500 created many fewer
filaments than gelsolin, so the relationship of gelsolin to
polymer number needs further study. At molar ratios of
actin to villin up to 250, fewer filaments form than the
concentration of villin [Wang and Bonder, 1991], so villin
may be somewhat less efficient at nucleating and capping
than CapZ or gelsolin. At molar ratios up to 100, one
filament formed for each molecule of a smooth muscle
capping protein [Hinssen et al., 1984], but the limited
range of the data precludes comparisons with the nucleating activity of other capping proteins.
Polymer Length and the Mechanical Properties
of Actin Filaments
In agreement with theory [MacKintosh et al., 1995],
pioneering work using semi-quantitative viscometric assays established the general principle that various capping and severing proteins lower the viscosity of actin
filaments by reducing polymer length [Yin and Stossel,
1979; Isenberg et al., 1980; Hasegawa et al., 1980; Craig
and Powell, 1980; Mooseker et al., 1980]. However, the
low and high shear viscosities measured in these experiments are not interpretable physical parameters. The best
quantitative measurements of rheological parameters used
gelsolin to control the polymer length [Janmey et al.,
1994], assuming that one filament forms per gelsolin,
rather than measuring the polymer lengths. This assumption is valid at low, but not high ratios of actin to gelsolin
[Janmey et al., 1986], so we recalculated the polymer
lengths in the 1994 study from the data in the 1986 paper
for comparison with our results where we measured
polymer lengths by light microscopy under the conditions
used in the rheometer.
While agreeing with on basic trends, our measurements of rheological constants as a function of actin
filament length are much lower than in the previous work
[Janmey et al., 1994]. For example, we find 0G*0 is 0.04
Pa for 24 µM actin filaments with a mean length of
2.8 µm. For an assumed mean polymer length of 2.7 µm
(adjusted value 2.2 µm) and a concentration of 48 µM,
Janmey et al. reported a range of values for the elastic
modulus, G8: 0.4 to 1.3 Pa in Figure 1 and 200 Pa in
Figure 2. For filaments 5–6 µm long, we measured 0G*0 ⫽
0.3 Pa for 24 µM actin, similar to Goldmann et al. [1997]
who reported G8 ⫽ 0.2 Pa for 10 µM actin, while Janmey
et al. [1994] measured 500 Pa for filaments 4 µm long at a
concentration of 48 µM.
Two factors are likely to contribute to these large
differences: the actin preparations and the capping protein
used to control the length of the filaments. In a recent
collaboration with the laboratory of P.A. Janmey, we
found that the methods used to prepare and store actin
appear to account for most of the differences reported
previously [Xu et al., 1998]. We now agree that 24 µM
solutions of filaments prepared from fresh, purified actin
have an elastic modulus of about 1 Pa in low amplitude
oscillation experiments at 0.1 to 1.0 Hz. These filaments
have a mean length of about 6 µm, as reported here. The
different capping proteins used to control polymer length
may also contribute to the differences. We used capping
protein copolymerization as a convenient method to
control the length of actin filaments without altering their
conformation [De La Cruz and Pollard, 1996]. Previous
studies [Hvidt et al., 1990; Janmey et al., 1994] used
gelsolin to control polymer length. Gelsolin severs and
caps actin filaments, so it is very effective in regulating
filament length, but in addition, gelsolin causes a conformational change that appears to propagate far along the
filament from the binding site at the barbed end [Prochniewicz et al., 1996]. This change alters the shape of the
subunits [Orlova et al., 1995] and the binding of rhodamine-phalloidin [Allen and Janmey, 1994], so it could
conceivably alter their mechanical properties as well.
Actin Filament Lengths and Mechanical Properties
Fig. 4. Effect of CapZ on the mechanical properties of actin filament networks. Conditions as in Figure 1.
A: The value of the complex modulus 0 G* 0. B: Phase shift ␦. Open square: Singly gel filtered actin. Filled
triangle: Doubly gel filtered actin alone. Filled square: Doubly gel filtered actin polymerized with 4.8 nM
CapZ. Open circle: Doubly gel filtered actin polymerized with 12 nM CapZ. Filled circle: Double gel
filtered actin polymerized with 48 nM CapZ.
oped by MacKintosh et al. [1995] for semi-flexible
polymers like actin. Long filaments, far above the persistence lengths, contribute to the moduli not only from
stiffness and rotational motion but also from bending
movement. Therefore, longer filaments are major contributors of actin networks.
Implications for Actin Filaments in Cells
Fig. 5. Dependence of the mechanical properties of actin filament
networks on polymer length. Conditions as in Figure 1. The length of
filaments of 24 µM doubly gel filtered actin was varied by copolymerization with various concentrations of CapZ as in Figure 3. The
complex modulus 0 G* 0 was measured at a frequency of 0.01 Hz.
The steep dependence of the elastic modulus on
polymer length (Fig. 5) means that long filaments contribute to the mechanical properties far out of proportion to
their length, in general agreement with concepts (but not
with the absolute values of the predicted moduli) devel-
Biochemical and morphological estimates of actin
filament lengths in cells differ [Small, 1981; Podolski and
Steck, 1990; Cano et al., 1991; Small et al., 1995;
Svitkina et al., 1997], but many filaments are shorter than
1 µm. We examined solutions of short actin filaments at
physiological concentrations for the first time, finding
that they have a low elastic modulus and behave more
like a fluid than networks of long actin filaments (Fig. 6).
Below 100 µM actin, the complex modulus depends on
the actin concentration. Above 100 µM, the actin concentration has less effect on the properties of the networks in
three different experiments.
Thus, short actin filaments alone cannot account for
the high elastic modulus of cytoplasm [Evans et al., 1993;
Oliver et al., 1994]. This result emphasizes the importance of actin filament crosslinking proteins in establishing the mechanical properties of the cell and provides an
opportunity for future research. No studies are available
on actin filament crosslinking at physiological concentrations of short filaments.
Xu et al.
Fig. 6. Mechanical properties of high concentrations of short actin filaments. Singly gel filtered actin was
copolymerized with a 1:500 molar ration of CapZ to actin. Conditions as in Figure 1. A: The value of the
complex modulus 0 G* 0. B: Phase shift ␦. Filled circle: 25 µM actin. Open circle: 50 µM actin. Filled
square: 100 µM actin. Open square: 300 µM actin.
This work was supported by NIH grants GM-26338
(to TDP) and AR-40697 (to JFC). JFC was an Established
Investigator of The American Heart Association during
the performance of this work. A Thomas C. Jenkins
Fellowship supported JX. We thank Enrique De La Cruz
for critically reading the manuscript. JX is grateful to
Xing Cao for help with calculations and to Jia Lu for the
independent measurements of the filament lengths.
Allen PG, Janmey PA. 1994. Gelsolin displaces phalloidin from actin
filaments: a new fluorescence method shows that both Ca2⫹ and
Mg2⫹ affect the rate at which gelsolin severs F-actin. J Biol
Chem 269:32916–32923.
Burlacu S, Janmey PA, Borjedo J. 1992. Distribution of actin filament
lengths measured by fluorescence microscopy. Am J Physiol
Caldwell JE, Heiss SG, Mermall V, Cooper JA. 1989. Effects of CapZ,
an actin capping protein of muscle, on the polymerization of
actin. Biochemistry 28:8506–8514.
Cano ML, Lauffenburger DA, Zigmond SH. 1991. Kinetic analysis of
F-actin depolymerization in polymorphonuclear leukocyte lysates indicates that chemoattractant stimulation increases actin
filament number without altering the filament distribution. J
Cell Biol 115:677–687.
Casella JF, Torres MA. 1994. Interaction of CapZ with actin. J Biol
Chem 269:6992–6998.
Casella JF, Maack DJ, Lin S. 1986. Purification and initial characterization of a protein from skeletal muscle that caps the barbed ends
of actin filaments. J Biol Chem 261:10915–10921.
Casella JF, Craig SW, Maack DJ, Brown AE. 1987. 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.
Casella JF, Barron-Casella EA, Torres MA. 1995. Quantitation of CapZ
in conventional actin preparations and methods for further
purification of actin. Cell Motil Cytoskeleton 30:164–170.
Coluccio LM, Tilney LG. 1984. Phalloidin enhances actin assembly by
preventing monomer dissociation. J Cell Biol 99:529–535.
Cooper JA, Pollard TD. 1985. Effect of capping protein on the kinetics
of actin polymerization. Biochemistry 24:793–799.
Cooper JA, Blum JD, Pollard TD. 1984. Acanthamoeba castellanii
capping protein: properties, mechanism of action, immunologic
cross-reactivity, and localization. J Cell Biol 99:217–225.
Craig SW, Powell LD. 1980. Regulation of actin polymerization by
villin, a 95,000 Dalton cytoskeletal component of intestinal
brush border. Cell 22:739–746.
De La Cruz E, Pollard TD. 1994. Transient kinetic analysis of
rhodamine phalloidin binding to actin filaments. Biochemistry
De La Cruz E, Pollard TD. 1996. Kinetics and thermodynamics of
phalloidin binding to actin filaments from three divergent
species. Biochemistry 35:14054–14061.
DiNubile MJ, Cassimeris LU, Joyce M, Zigmond SH. 1995. Actin
filament barbed end capping activity in neutrophil lysates: the
role of capping protein ␤2. Molec Biol Cell 12:1659–1671.
Evans E, Leung A, Zhelev D. 1993. Synchrony of cell spreading and
contraction force as phagocytes engulf large pathogens. J Cell
Biol 122:1295–1300.
Ferry JD. 1980. Viscoelastic properties of polymers. New York: John
Wiley and Sons.
Goldmann WH, Tempel M, Sprenger I, Isenberg G, Ezzell RM. 1997.
Viscoelasticity of actin-gelsolin networks in the presence of
filamin. Eur J Biochem 246:373–379.
Hasegawa T, Takahashi S, Hayashi H, Hatano S. 1980. Fragmin: a
calcium ion sensitive regulatory factor on the formation of actin
filaments. Biochemistry 19:2677–2683.
Actin Filament Lengths and Mechanical Properties
Hinssen H, Small JV, Sobieszek A. 1984. A Ca2⫹-dependent actin
modulator from vertebrate smooth muscle. FEBS Lett 166:
Hvidt S, Janmey PA. 1990. Elasticity and flow properties of actin gels.
Makromol Chem Macromol Symp 39:209–213.
Isenberg G, Aebi U, Pollard TD. 1980. An actin-binding protein from
Acanthamoeba regulates actin filament polymerization and
interactions. Nature (Lond) 288:455–459.
Janmey PA, Peetermans J, Zaner KS, Stossel TP, Tanaka T. 1986.
Structure and mobility of actin filaments as measured by
quasielastic light scattering, viscosity and electron microscopy.
J Biol Chem 261:8357–8362.
Janmey PA, Hvidt S, Käs J, Lerche D, Maggs A, Sackmann E, Schliwa
M, Stossel TP. 1994. The mechanical properties of actin gels. J
Biol Chem 269:32503–32513.
Käs J, Strey H, Tang JX, Finger D, Ezzell R, Sackmann E, Janmey PA.
1996. F-actin, a model polymer for semiflexible chains in dilute,
semidilute, and liquid crystalline solutions. Biophys J 70:609–
Kaufmann S, Käs J, Goldmann WH, Sackmann E, Isenberg G. 1992.
Talin anchors and nucleates actin filaments at lipid membranes—a direct demonstration. FEBS Lett 314:203–205.
Kron SJ, Toyoshima YY, Uyeda TQP, Spudich JA. 1991. Assays for
actin sliding movement over myosin-coated surfaces. Methods
Enzymol 196:399–416.
Lehrer SS. 1972. Crosslinking of actin and of tropomyosin by
glutaraldehyde. Biochem Biophys Res Commun 48:967–976.
MacKintosh FC, Käs J, Janmey PA. 1995. Elasticity of semiflexible
biopolymer networks. Phys Rev Lett 75:4425–4428.
MacLean-Fletcher S, Pollard TD. 1980. Identification of a factor in
conventional muscle actin preparations which inhibits actin
filament self-association. Biochem Biophys Res Commun 96:
Mooseker MS, Graves TA, Wharton KA, Falco N, Howe CL. 1980.
Regulation of microvillus structure: calcium-dependent solation
and cross-linking of actin filaments in the microvilli of intestinal
epithelial cells. J Cell Biol 87:908–922.
Niederman R, Amrein PC, Hartwig JH. 1983. Three-dimensional
structure of actin filaments and of an actin gel made with
actin-binding protein. J Cell Biol 96:1400–1413.
Oliver T, Lee J, Jacobson K. 1994. Forces exerted by locomoting cells.
Semin Cell Biol 5:139–147.
Orlova A, Prochniewicz E, Egelman EH. 1995. Structure dynamics of
F-actin: II. Cooperativity in structure transitions. J Mol Biol
Podolski JL, Steck TL. 1990. Length distribution of F-actin in
Dictyostelium discoideum. J Biol Chem 265:1312–1318.
Prochniewicz E, Zhang Q, Janmey PA, Thomas DD. 1996. Cooperativity in F-actin: binding of gelsolin at the barbed end affects
structure and dynamics of the whole filament. J Mol Biol
Sampath P, Pollard TD. 1991. Effects of cytochalasin, phalloidin and
pH on the elongation of actin filaments. Biochemistry 30:1973–
Sato M, Leimbach G, Schwarz WH, Pollard TD. 1985. Mechanical
properties of actin. J Biol Chem 260:8585–8592.
Small JV. 1981. Organization of ␣-actinin in the leading edge of
cultured cells: influence of osmium tetroxide and dehydration
on the ultrastructure of actin meshworks. J Cell Biol 91:695–
Small JV, Herzog M, Anderson K. 1995. Actin filament organization in
fish keratocyte lamellipodium. J Cell Biol 129:1275–1286.
Svitkina TM, Verkhovsky AB, McQuade KM, Borisy GG. 1997.
Analysis of the actin-myosin II system in fish epidermal
keratocytes: mechanism of cell body translocation. J Cell Biol
Wang F-S, Bonder EM. 1991. Sea urchin egg villin: identification of
villin in a non-epithelial cell from an invertebrate species. J Cell
Sci 100:61–71.
Warshaw DM, Desrosiers JM, Work SS, Trybus KM. 1990. Smooth
muscle myosin cross-bridge interactions modulate actin filament sliding velocity in vitro. J Cell Biol 111:453–463.
Xu J, Schwarz WH, Käs J, Stossel TP, Janmey PA, Pollard TD. 1998.
Mechanical properties of actin filament networks depend on
preparation, polymerization conditions and storage of actin
monomers. Biophys J 74:2731–2740.
Yin H, Stossel TP. 1979. Control of cytoplasmic action gel-sol
transformation by gelsolin, a calcium-dependent regulatory
protein. Nature 281:583–586.
Без категории
Размер файла
141 Кб
Пожаловаться на содержимое документа