1 Department of Biochemistry and Molecular Biophysics, Washington University
School of Medicine, St Louis, MO 63110, USA
2 Department of Pathology and Anesthesiology, St Louis University School of
Medicine, St Louis, MO 63104, USA
* Author for correspondence (e-mail: elson{at}biochem.wustl.edu)
Accepted 2 January 2003
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Summary |
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Key words: Myosin II, Cell spreading, Cell mechanics, Traction force, Contraction, Cytochalasin D
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Introduction |
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Much has recently been learned about the regulation of these forces. For
example, when fibroblasts, initially spherical in suspension, encounter a
substratum, they spread to a more flattened shape. This process is analogous
to the forward extension of lamelllipodia and filopodia by polarized cells as
they migrate over a substratum. This cell spreading is driven by actin
polymerization that is promoted by the Rho-family GTPases Cdc42 and Rac1.
These two molecules initiate polymerization by different pathways
(Worthylake and Burridge,
2001). The former activates the Arp2/3 complex, which nucleates
new actin filaments to form a dendritic actin filament network
(Borisy and Svitkina, 2000
).
The latter uncaps the barbed ends of pre-existing filaments to permit them to
grow further (Hartwig et al.,
1995
). After spreading, the cells form specialized structures such
as stress fibers and focal adhesions that mediate adherence to the substratum.
These processes require myosin-dependent contractile force that is activated
by the RhoA GTPase- and calcium/calmodulin (CaM)-dependent pathways, which
promote the phosphorylation of the regulatory light chain (RLC) of myosin II
in nonmuscle cells (Amano et al.,
1996
; Kimura et al.,
1996
). The processes that regulate extension and adhesion are
coordinated. During the initial cell spreading phase, binding of integrins to
ECM ligands diminishes RhoA activity by activating p190RhoGAP via c-Src and
FAK (Arthur and Burridge, 2001
;
Ren et al., 1999
;
Ren et al., 2000
). This
observation suggests the hypothesis that RhoA is deactivated to diminish the
myosin contractile force that would otherwise impede cell spreading. This
hypothesis predicts that the rate of cell spreading should vary inversely with
myosin activity. In this work, we have tested this hypothesis by correlating
the activity of myosin, measured in terms of the extent of phosphorylation of
its RLC, with the rate of spreading of chicken embryo fibroblasts (CEFs).
As indicated above, current models suppose that the extension of the cell
is propelled by the growth of actin filaments that incorporate actin monomers
at their barbed ends. This hypothesis predicts that the barbed ends of the
participating filaments are free (i.e. are not blocked by barbed-end capping
proteins). If this is true, the filaments should bind cytochalasin D (CD) at
close to nanomolar concentrations in the range of the binding constant
measured for free actin filaments (Cooper,
1987). Therefore, edge extension should be inhibited by CD at
sub-nanomolar concentrations in contrast to the effects of CD on cell shape
and deformability that require much higher (µM) concentrations to compete
with barbed-end capping proteins bound to actin filaments
(Wakatsuki et al., 2001
). We
have tested this hypothesis by determining the concentrations at which CD
inhibits cell spreading.
A further hypothesis is that activation of myosin promotes interaction of myosin with actin filaments anchored to the plasma membrane, thereby increasing cell stiffness and consequently resistance to cell extension driven by actin polymerization. This hypothesis predicts that the stiffness of the cell should be increased by myosin activation. Using a cell indentation assay to measure cell stiffness, we have tested and confirmed this hypothesis.
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Materials and Methods |
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The degree of cell spreading was also measured with different concentrations of CD (Sigma) in the presence of staurosporine (STA) or KT5926 (KT). Each 35 mm dish contained the same concentration of STA (100 nM) or KT (1 µM) and cells (100,000/dish) but a different concentration of CD. The cell spreading was stopped and measured 1 hour after the seeding of cells in each dish as described above. The CD and STA, or CD and KT, were present in the medium throughout the experiment. The half-maximum concentration of CD to inhibit cell spreading was determined by fitting a sigmoidal curve to the data shown in Fig. 3. Stock solution (2 mM) of CD was made in DMSO.
|
Myosin RLC phosphorylation
At each time point during cell spreading, after discarding the culture
medium, the dishes were incubated with a solution containing 1 ml 10% TCA and
10 mM dithiothreitol (DTT). The precipitate was scraped from the dishes and
washed three times each with 1 ml of acetone containing 10 mM DTT. After
centrifugation, the pellets were solubilized in loading buffer containing 9 M
urea, 2 mM DTT, 20 mM Tris, 22 mM glycine and 250 mM sucrose. The
phosphorylated myosin RLCs were separated in 1 mm thick mini-gels containing
40% glycerol, 10% acrylamide, 20 mM Tris and 22 mM glycine. The detailed
experimental procedures are presented elsewhere
(Goeckeler et al., 2000).
Polyclonal rabbit antibody to recombinant myosin II RLC was used to detect the
light chains (Chew et al.,
1998
).
Compounds inhibiting myosin II activities
To inhibit myosin activity, samples were treated with 100 nM STA (Sigma) or
1 µM KT (Calbiochem, La Jolla, CA) dissolved in DMSO (<0.1% of the total
volume of the medium). The cells were not pretreated with any other compounds.
A small amount of medium containing resuspended cells was pipetted into the
medium already mixed with STA or KT at its final concentration. Control
samples contained the same amount of DMSO as the samples treated with STA and
KT.
Confocal immunofluorescence microscopy
F-actin was stained with rhodamine-labeled phalloidin. Myosin II was
immunolocalized using antibodies specific for myosin II
(Goeckeler and Wysolmerski,
1995) and an Alexa-labeled secondary antibody. The fluorescent
images of actin filaments and myosin II were taken using scanning confocal
microscopy (Bio-Rad, Hercules, CA).
Indentation stiffness measurements of single cells
The indenter consists of a vertical glass stylus with a tip about 2 µm
in diameter connected to a linear piezoelectric motor by a glass beam of known
bending constant. The vertical position of the stylus tip is monitored
optically by measuring the light reflected from a flag attached to the tip.
The force exerted on the tip by the resistance of the cell to indentation is
calculated by measuring the bending of the beam
(Zahalak et al., 1990). The
point at which bending of the beam is first detected indicates the initial
contact of the tip with an object. This position indicates the height of the
object relative to the substratum. A reference position was established for
each cell by probing the substratum near the cell.
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Results |
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KT and STA accelerate cell spreading
Can cells spread in the absence of myosin II activity? We used kinase
inhibitors to prevent activation of myosin by phosphorylation of the myosin
RLC, and the effects of the inhibitors on the levels of myosin RLC
phosphorylation were measured. A specific myosin RLC kinase inhibitor KT (1
µM) reduced the amount of the phosphorylated RLC to almost an undetectable
level at 10 minutes after re-plating. Then the level of phosphorylation began
to climb, reaching almost 30% by 2 hours
(Fig. 1). The relatively
nonspecific kinase inhibitor STA (100 nM) suppressed the myosin RLC
phosphorylation to less than 5% for the entire 2-hour measuring period
(Fig. 1).
The inhibitors of myosin RLC phosphorylation, KT and STA, accelerated cell spreading. Although the total projected area at the end of 2 hours was almost indistinguishable for cells in the presence and absence of KT and STA, the difference in spreading area at 30 minutes and 60 minutes between treated and nontreated groups was statistically significant (P<0.05) (Fig. 2). We conclude that myosin II contributes little to the protrusion of the leading edge of a spreading cell. Indeed, cell spreading could occur with an almost undetectable level of myosin II activity. The cell spreading apparently is retarded by the myosin II activity. These observations raise a simple question: what is the major force driving protrusion of cell edges during cell spreading? The most reasonable candidate is actin polymerization. If we could test a specific inhibitor of actin polymerization in combination with the myosin inhibitor, the contribution of actin polymerization to cell spreading would be clarified.
|
Low dose of CD inhibits cell spreading
We used CD to test the importance of actin polymerization in cell spreading
while myosin II activity was suppressed. Although the mechanisms by which CD
affects the organization of the actin cytoskeleton are complex in detail
(Cooper, 1987), at the low
concentrations used in these experiments the main effect of CD arises from its
binding to the barbed end of actin filaments, thereby both inhibiting actin
polymerization and competing with capping proteins for binding to the barbed
ends (Wakatsuki et al., 2001
).
The dependence of the projected cell area on the concentration of CD was
observed for cells treated with STA and KT. The area measurements were well
fitted to sigmoidal curves plotted on semi-logarithmic scale to yield an
effective half-maximum inhibitory dose (IC50) of CD pretreated with
STA and KT of approximately 3.5 nM and 20 nM, respectively
(Fig. 3). These are much closer
to the estimated dissociation constant (2 nM) for the binding of CD to the
barbed end of actin filaments in vitro
(Cooper, 1987
) than those
estimated or used for F-actin disruption
(Wakatsuki et al., 2001
).
Despite the complexity of the action of CD in vivo
(Cooper, 1987
), the close
match to Kd suggests that the inhibitory effect of CD resulted from its
binding to the barbed ends of actin filaments with little competition from
capping proteins. This further suggests that actin filaments undergoing
polymerization during the protrusion of the leading edge are essentially free
at their barbed ends. Since myosin II-dependent actin reorganization was
suppressed by STA or KT, these results support the hypothesis that the
protrusion of the leading cell edge requires actin polymerization but not
myosin II. Nevertheless, the possible involvement of other motor proteins not
inhibited by STA or KT cannot be ruled out.
Actin and myosin II localization during cell spreading with and
without STA and KT
Active myosin II binds actin filaments and bundles them to form stress
fibers. Myosin II visualized by immunostaining co-localized with actin
filaments labeled with rhodamine-phalloidin. Together, they began to resemble
stress fibers 60 minutes after the initiation of cell spreading without KT and
STA (result not shown). Under the same conditions at 120 minutes, stress
fibers were indicated clearly by a strong co-localization of myosin II with
actin filaments (Fig. 4M).
Suppressing myosin II activity by KT and STA abolished the co-localization of
myosin II with actin filaments. The KT and STA treatments increased the number
of filopodia and induced extension of lamellipodia earlier than in the control
cells (Fig. 4A-C). Thus, cells
treated with KT and STA spread much faster than cells without them
(Fig. 4G-I). The increased rate
of cell spreading in the presence of KT and STA also caused the cells to
flatten earlier. This was visualized in cross-sectional images of cells taken
by confocal microscopy (Fig.
4D-F,J-L). In some cells treated with straurosporine for 60
minutes, the actin filaments began to assume a dispersed distribution and, by
120 minutes, many of the cells began to break apart
(Fig. 4O). A similar breakage
of keratocytes by STA treatment has been reported
(Verkhovsky et al., 1999b).
The cells treated with KT-5926 never show this kind of fragmentation. Instead,
at 120 minutes, a co-localization of myosin II and actin filaments
(Fig. 4N), which correlated
with the increase in myosin II activity, began to appear in KT5926-treated
cells as shown in Fig. 1. The
localization of myosin II and actin with and without STA and KT was consistent
with the activity of myosin II shown in
Fig. 1 and the rate of cell
spreading shown in Fig. 2.
However, at 10 minutes of cell spreading, myosin II was localized diffusely in
the entire cytoplasm with or without STA and KT, despite the difference in
myosin RLC phosphorylation noted above. The association of active myosin II
with F-actin might be difficult to visualize by fluorescence microscopy in the
early stages of cell spreading.
|
Cell deformability
In the early stages of cell spreading, the extent of actin filament
crosslinking by active myosin could be less than at later stages, when stress
fibers are formed (Fig. 4M).
The crosslinking of actin filaments by -actinin increases
viscoelasticity of actin networks (Sato et
al., 1987
). If active myosin II crosslinks actin filaments, the
viscoelastic properties of cells treated with KT should be different from
those of untreated cells. We have suggested that activation of myosin retards
spreading of a cell by increasing its resistance to deformation (i.e. its
stiffness). To test this hypothesis, we measured cell stiffness by an
indentation method in the early stages of spreading (initial 10-30 minutes).
These measurements are similar in concept to atomic force microscopy. The tip
of a probe mounted on a cantilever with known Young's Modulus indents the
cell. The bending of the cantilever is translated into the force required for
indentation (Zahalak et al.,
1990
). This approach yields the heights of cells by comparing the
positions at which the cantilever begins to bend due to initial contacts of
the probe with the cell and with the substratum. In this way, we have
confirmed that KT caused the cells to become flatter than control cells
(Fig. 5A), as was observed in
cross-sectional confocal microscopic images
(Fig. 4D,E,J,K).
|
KT reduced cellular stiffness. The forces required to indent single cells with or without treatment by KT are plotted against % indentation (Fig. 5B). In these measurements, the tip is lowered at a constant rate to indent the cell (`loading') and is then retracted at the same rate (`unloading'). KT reduced both the resistance force and the slope (stiffness) of the loading curve (Fig. 5B).
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Discussion |
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Both extending and contracting forces are regulated by Rho-family GTPases.
Much recent evidence indicates that actin polymerization drives cell spreading
and forward extension during migration
(Borisy and Svitkina, 2000;
Pollard et al., 2000
). Two
Rho-family GTPases, Rac1 and Cdc42, promote polymerization either by uncapping
the barbed (rapidly polymerizing) ends of pre-existing actin filaments or by
activating the Arp2/3 complex to nucleate new actin filaments at the sides of
pre-existing filaments, thereby forming a dendritic network
(Worthylake and Burridge,
2001
). Another member of this family, RhoA, promotes formation of
stress fibers and focal adhesions by elevating contractile force
(Burridge and Chrzanowska-Wodnicka,
1996
; Elbaum et al.,
1999
). Rho A activates nonmuscle myosin by increasing the level of
phosphorylation of its RLC (Amano et al.,
1996
; Kimura et al.,
1996
).
Depending on their relative magnitudes, the balance of forces arising from
actin polymerization and myosin-dependent contraction will favor either
extension or contraction of the cell margins. Evidently, this balance must
vary during cell locomotion, spreading and adherence on a substrate. Biphasic
regulation of Rho has been demonstrated when Swiss 3T3 cells adhere to
fibronectin-coated substrata (Ren et al.,
1999). A transient inhibition of Rho activity when the cells first
contact the substratum is followed by an increase in Rho activity. The initial
inhibition of Rho requires the tyrosine kinase FAK
(Ren et al., 2000
). In
neutrophils, the mechanism of inhibition appears to operate through the
oncogene c-Src, which by tyrosine phosphorylation activates p190RhoGAP, an
inhibitor of Rho activity (Arthur and
Burridge, 2001
). These observations suggest that early inhibition
of Rho upon contact with integrins might correspondingly inhibit
myosin-dependent contractile forces that would resist cell spreading or the
extension of the leading edge of migrating cells
(Arthur and Burridge, 2001
).
For this hypothesis to be correct, the rate of cell spreading must vary
inversely with myosin activity.
We have directly tested and confirmed this hypothesis by measuring the rate
of spreading under conditions in which myosin activity is either inhibited, at
a baseline level, or activated. In agreement with prediction, the rate of
extension decreases as the level of myosin phosphorylation, and therefore
myosin activity, increases (Figs
1,
2). In neutrophils, contact
with peptides containing an RGD sequence, simulating interaction with an ECM
ligand, had no effect on the activities of Cdc42 and Rac
(Arthur et al., 2000). On the
basis of early observations that edge extension continued even after
contractile forces began to elevate a lamellipodium in the early stages of
`ruffle' formation (Felder and Elson,
1990
), it appears that actin polymerization continues while
myosin-dependent contractile forces increase.
We have observed that the level of myosin phosphorylation was surprisingly
high (50%) in serum-starved cells and that this level at first decreased when
the cells become adherent and then increased back to 50%. In earlier work, it
was observed that while activation of Rho by lysophosphatidic acid was
comparable in suspended and adherent cells, the Rho activity in adherent cells
declined while that of suspended cells remained high
(Ren et al., 1999). This
suggests that there is an adherence-dependent system for regulating myosin
activation through Rho (Arthur and
Burridge, 2001
; Ren et al.,
1999
).
Effects of inhibitors
Our results depend on using the kinase inhibitors KT and STA to reduce
myosin activity. These inhibitors have different target specificities. KT is
specific for the myosin light chain kinase (MLCK)
(Nakanishi et al., 1990) and
hence inhibits calcium-ion-dependent myosin activation. STA has a broad
specificity for many protein kinases including MLCK, Rho kinase, protein
kinase A (PKA), PKC and CaM kinase. However, as we have confirmed
(Fig. 1), STA does inhibit
myosin RLC phosphorylation very effectively
(Sakurada et al., 1998
).
Because of the higher specificity of KT, the relationship between its
inhibition of myosin and the acceleration of spreading (Figs
1,
2) seems relatively
straightforward. Nevertheless, we emphasize that the correlation of myosin
inhibition with acceleration of spreading is seen with both inhibitors. STA
inhibits myosin significantly longer than does KT. Although both STA and KT
initially abolished myosin RLC phosphorylation almost completely,
phosphorylation began to increase after 1 hour of spreading in the cells
treated with KT, whereas STA continued to inhibit the RLC phosphorylation
throughout the entire 2-hour observation period
(Fig. 2). This was also
confirmed visually by localization of myosin II and actin filaments
(Fig. 4). Since STA has a
broader specificity, this difference suggests the involvement of kinases other
than MLCK after 60 minutes of cell spreading. The most attractive candidate is
Rho kinase, which is a target of STA (Feng
et al., 1999). Inhibition of Rho kinase could interfere with RLC
phosphorylation either directly or indirectly via the effect of Rho kinase on
myosin phosphatase (Kureishi et al.,
1997
). The enhancement of endothelial cell spreading by the Rho
kinase-specific inhibitor Y27632 has been observed recently (R.B.W. and G. M.
Grojean, unpublished). Further investigation is required to clarify temporal
differences in the regulation of cell spreading by MLCK and Rho kinase.
It has been reported that 2,3-butanedione monoxime (BDM) inhibits cell
spreading (Sanders et al.,
1999). Since BDM could inhibit ATPase activity of myosin isoforms
other than myosin II, this observation could suggest involvement of different
myosin isoforms in cell spreading. However, in our experiments, BDM (20 mM)
treatment interfered with the adherence of CEFs to the tissue culture dishes
(result not shown). This additional cellular response complicates the
interpretation of the effect of BDM on contractility and myosin II function in
cell spreading. BDM has been reported recently to have nonspecific and weak
inhibitory effects on nonmuscle myosin
(Cheung et al., 2001
).
In addition to confirming the correlation between myosin inhibition and acceleration of spreading, the results obtained using STA are interesting because of the differences observed between its effects and those obtained with KT. These results clearly show that, although we cannot yet identify them, other STA targets are involved in myosin regulation and also in actin polymerization, as indicated below. More work is required to find the kinases responsible for these additional effects.
The actin filaments responsible for spreading are uncapped
To explain the effects of CD on cell shape and mechanical properties, it is
necessary to take into account that most of the actin filaments in a cell are
capped. Therefore, even though the binding of CD to the free barbed ends of
actin filaments occurs with an affinity constant of 2 nM
(Cooper, 1987
), it is
necessary to use CD concentrations in the micromolar range to cause cells to
round up and soften. This discrepancy is readily explained as a consequence of
the competition between CD and endogenous capping proteins that bind the
barbed ends of actin filaments (Wakatsuki
et al., 2001
). By contrast, the concentrations of CD effective for
inhibition of cell spreading pretreated with STA and KT are 3.5 nM and 20 nM
respectively. Therefore, the actin filaments that contribute to spreading are
uncapped, as expected if they are growing by addition of new actin monomers to
their barbed ends. That the concentrations of CD effective to inhibit
spreading are lower in the presence of STA than with KT suggests that
additional STA targets beyond MLCK regulate uncapping activity during initial
stages of cell spreading.
The retardation of cell spreading by active myosin correlates with
increased cell stiffness
Our indentation measurements indicate that the retardation of cell
spreading with increased myosin activity results from a myosin-dependent
increase of cellular stiffness. The increased stiffness resists the protrusive
force supplied by actin polymerization. Increased stiffness due to the action
of nonmuscle myosin has been demonstrated in lymphocytes and
Dityostelium amoebae (Pasternak
and Elson, 1985; Pasternak et
al., 1989
). These studies were carried out by measuring cell
stiffness as the resistance to indentation by a small probe
(Petersen et al., 1982
;
Zahalak, 1986
). We have used
this same approach to measure cell deformation at rates comparable with the
rates at which the cell changes shape. The specific inhibition of MLCK by KT
reduces myosin activity and consequently leads to a reduction of contractile
force and cellular stiffness (Fig.
5B). This supports our hypothesis that KT facilitates cell
spreading by reducing cell stiffness.
A quantitative mechanical model of the cell is not yet available, and so we
cannot provide a definitive structural interpretation of these observations.
(Simple linear models such as the linear standard solid cannot explain this
behavior.) Nevertheless, the measurements reported here are sufficient to
confirm the predicted correlations between myosin activation and cell
stiffness and their inverse correlation with rates of cell spreading. The
active retrograde flow of myosin II observed in strongly adherent cells
(Verkhovsky et al., 1999a)
could be another factor that retards cell spreading. Membrane tension is also
one of the physical factors that influence the rate of cell spreading by
opposing actin polymerization (Raucher and
Sheetz, 2000
). Our measurements have demonstrated a shifting
balance between a protrusive force due to actin polymerization and a
myosin-dependent retarding force that also controls the rate of cell
spreading.
Cell spreading and traction force
Cells exert a traction force on the substratum over which they migrate. The
traction force was demonstrated on 2D surfaces by qualitatively observing the
formation of wrinkles on ultra-thin silicon rubber films
(Harris et al., 1981), and
more quantitatively by several groups
(Balaban et al., 2001
;
Dembo et al., 1996
;
Galbraith and Sheetz, 1997
).
The existence of the traction force was first demonstrated in 3D by Stopak and
coworkers observing the contraction of cell-populated collagen matrices (CPM)
(Bell et al., 1979
;
Grinnell, 1994
;
Stopak and Harris, 1982
). We
have directly measured these forces during CPM contraction by connecting the
collagen matrix to an isometric force transducer. Contractile force increased
significantly while cells are spreading into the collagen matrices in the
presence of serum (Wakatsuki and Elson,
2003
). However, we have found that, in a culture medium lacking
serum, cells can spread into a matrix without causing substantial matrix
remodeling and production of contractile force
(Wakatsuki and Elson, 2003
). A
similar result was obtained by tracking the movement of small beads embedded
around the fibroblasts in a collagen matrix
(Roy et al., 1999
). These
observations suggest that a traction force is unnecessary for cell spreading
or that the minimum traction force required for cell spreading could be too
small to detect using the available methods.
Summary
We have analyzed the contribution of myosin II to cell spreading on 2D
substrates. Our results are schematically summarized in
Fig. 6. Reducing myosin II
activity by inhibiting phosphorylation of myosin RLCs increased the rate of
spreading. The protrusion of the lamella observed under myosin II inhibition
was most likely due to actin polymerization (small dark arrow in
Fig. 6), as was demonstrated by
inhibiting cell spreading using low concentrations of CD under conditions of
myosin inhibition. While cells with serum exert large traction forces during
spreading, cells without serum hardly exert any traction forces. The low level
of myosin II activity detected without serum could be required to maintain the
integrity of the actin cytoskeleton (white arrow in
Fig. 6). Consistent with
observations on fish epidermal keratocytes
(Verkhovsky et al., 1999b), we
have shown that STA treatment of CEFs causes them to fragment. The results of
cell indentation measurements suggest that inhibition of myosin activation by
KT decreased cytoplasmic stiffness possibly by decreasing the crosslinking of
actin filaments by myosin. This decrease of stiffness was then responsible for
the observed increased cell deformability and the increased rate of cell
spreading.
|
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Acknowledgments |
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References |
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---|
Amano, M., Ito, M., Kimura, K., Fukata, Y., Chihara, K., Nakano,
T., Matsuura, Y. and Kaibuchi, K. (1996). Phosphorylation and
activation of myosin by Rho-associated kinase (Rho-kinase). J.
Biol. Chem. 271,20246
-20249.
Arthur, W. T. and Burridge, K. (2001). RhoA
inactivation by p190RhoGAP regulates cell spreading and migration by promoting
membrane protrusion and polarity. Mol. Biol. Cell
12,2711
-2720.
Arthur, W. T., Petch, L. A. and Burridge, K. (2000). Integrin engagement suppresses RhoA activity via a c-Src-dependent mechanism. Curr. Biol. 10,719 -722.[CrossRef][Medline]
Balaban, N. Q., Schwarz, U. S., Riveline, D., Goichberg, P., Tzur, G., Sabanay, I., Mahalu, D., Safran, S., Bershadsky, A., Addadi, L. et al. (2001). Force and focal adhesion assembly: a close relationship studied using elastic micropatterned substrates. Nat. Cell Biol. 3,466 -472.[CrossRef][Medline]
Bell, E., Ivarsson, B. and Merrill, C. (1979). Production of a tissue-like structure by contraction of collagen lattices by human fibroblasts of different proliferative potential in vitro. Proc. Natl. Acad. Sci. USA 76,1274 -1278.[Abstract]
Borisy, G. G. and Svitkina, T. M. (2000). Actin machinery: pushing the envelope. Curr. Opin. Cell Biol. 12,104 -112.[CrossRef][Medline]
Burridge, K. and Chrzanowska-Wodnicka, M. (1996). Focal adhesions, contractility, and signaling. Annu. Rev. Cell Dev. Biol. 12,463 -518.[CrossRef][Medline]
Cheung, A. J., Westwood, N. J., Chen, I., Mitchision, T. J. and Straight, A. F. (2001). Blebbistatin: a cell permeable ihibitor of non-muscle myosin II. Mol. Biol. Cell 12, 271a.
Chew, T. L., Masaracchia, R. A., Goeckeler, Z. M. and Wysolmerski, R. B. (1998). Phosphorylation of non-muscle myosin II regulatory light chain by p21-activated kinase (gamma-PAK). J. Muscle Res. Cell Motil. 19,839 -854.[CrossRef][Medline]
Cooper, J. A. (1987). Effects of cytochalasin and phalloidin on actin. J. Cell Biol. 105,1473 -1478.[Medline]
Dembo, M., Oliver, T., Ishihara, A. and Jacobson, K. (1996). Imaging the traction stresses exerted by locomoting cells with the elastic substratum method. Biophys. J. 70,2008 -2022.[Abstract]
Elbaum, M., Chausovsky, A., Levy, E. T., Shtutman, M. and Bershadsky, A. D. (1999). Microtubule involvement in regulating cell contractility and adhesion-dependent signalling: a possible mechanism for polarization of cell motility. Biochem. Soc. Symp. 65,147 -172.[Medline]
Felder, S. and Elson, E. L. (1990). Mechanics of fibroblast locomotion: quantitative analysis of forces and motions at the leading lamellas of fibroblasts. J. Cell Biol. 111,2513 -2526.[Abstract]
Feng, J., Ito, M., Kureishi, Y., Ichikawa, K., Amano, M., Isaka,
N., Okawa, K., Iwamatsu, A., Kaibuchi, K., Hartshorne, D. J. et al.
(1999). Rho-associated kinase of chicken gizzard smooth muscle.
J. Biol. Chem. 274,3744
-3752.
Galbraith, C. G. and Sheetz, M. P. (1997). A
micromachined device provides a new bend on fibroblast traction forces.
Proc. Natl. Acad. Sci. USA
94,9114
-9118.
Goeckeler, Z. M. and Wysolmerski, R. B. (1995). Myosin light chain kinase-regulated endothelial cell contraction: the relationship between isometric tension, actin polymerization, and myosin phosphorylation. J. Cell Biol. 130,613 -627.[Abstract]
Goeckeler, Z. M., Masaracchia, R. A., Zeng, Q., Chew, T. L.,
Gallagher, P. and Wysolmerski, R. B. (2000). Phosphorylation
of myosin light chain kinase by p21-activated kinase PAK2. J. Biol.
Chem. 275,18366
-18374.
Grinnell, F. (1994). Fibroblasts, myofibroblasts, and wound contraction. J. Cell Biol. 124,401 -404.[Medline]
Harris, A. K., Stopak, D. and Wild, P. (1981). Fibroblast traction as a mechanism for collagen morphogenesis. Nature 290,249 -251.[Medline]
Hartwig, J. H., Bokoch, G. M., Carpenter, C. L., Janmey, P. A., Taylor, L. A., Toker, A. and Stossel, T. P. (1995). Thrombin receptor ligation and activated Rac uncap actin filament barbed ends through phosphoinositide synthesis in permeabilized human platelets. Cell 82,643 -653.[Medline]
Kimura, K., Ito, M., Amano, M., Chihara, K., Fukata, Y., Nakafuku, M., Yamamori, B., Feng, J., Nakano, T., Okawa, K. et al. (1996). Regulation of myosin phosphatase by Rho and Rho-associated kinase (Rho-kinase). Science 273,245 -248.[Abstract]
Kureishi, Y., Kobayashi, S., Amano, M., Kimura, K., Kanaide, H.,
Nakano, T., Kaibuchi, K. and Ito, M. (1997). Rho-associated
kinase directly induces smooth muscle contraction through myosin light chain
phosphorylation. J. Biol. Chem.
272,12257
-12260.
Nakanishi, S., Yamada, K., Iwahashi, K., Kuroda, K. and Kase, H. (1990). KT5926, a potent and selective inhibitor of myosin light chain kinase. Mol. Pharmacol. 37,482 -488.[Abstract]
Pasternak, C. and Elson, E. L. (1985). Lymphocyte mechanical response triggered by cross-linking surface receptors. J. Cell Biol. 100,860 -872.[Abstract]
Pasternak, C., Spudich, J. A. and Elson, E. L. (1989). Capping of surface receptors and concomitant cortical tension are generated by conventional myosin. Nature 341,549 -551.[CrossRef][Medline]
Petersen, N. O., McConnaughey, W. B. and Elson, E. L. (1982). Dependence of locally measured cellular deformability on position on the cell, temperature, and cytochalasin B. Proc. Nat. Acad. Sci. USA 79,5327 -5331.[Abstract]
Pollard, T. D., Blanchoin, L. and Mullins, R. D. (2000). Molecular mechanisms controlling actin filament dynamics in nonmuscle cells. Annu. Rev. Biophys. Biomol. Struct. 29,545 -576.[CrossRef][Medline]
Raucher, D. and Sheetz, M. P. (2000). Cell
spreading and lamellipodial extension rate is regulated by membrane tension.
J. Cell Biol. 148,127
-136.
Ren, X. D., Kiosses, W. B. and Schwartz, M. A.
(1999). Regulation of the small GTP-binding protein Rho by cell
adhesion and the cytoskeleton. EMBO J.
18,578
-585.
Ren, X. D., Kiosses, W. B., Sieg, D. J., Otey, C. A.,
Schlaepfer, D. D. and Schwartz, M. A. (2000). Focal adhesion
kinase suppresses Rho activity to promote focal adhesion turnover.
J. Cell Sci. 113,3673
-3678.
Roy, P., Petroll, W. M., Cavanagh, H. D. and Jester, J. V. (1999). Exertion of tractional force requires the coordinated up-regulation of cell contractility and adhesion. Cell Motil. Cytoskeleton 43,23 -34.[CrossRef][Medline]
Sakurada, K., Seto, M. and Sasaki, Y. (1998). Dynamics of myosin light chain phosphorylation at Ser19 and Thr18/Ser19 in smooth muscle cells in culture. Am. J. Physiol. 274,C1563 -1572.[Medline]
Sanders, L. C., Matsumura, F., Bokoch, G. M. and de Lanerolle,
P. (1999). Inhibition of myosin light chain kinase by
p21-activated kinase. Science
283,2083
-2085.
Sato, M., Schwarz, W. H. and Pollard, T. D.
(1987). Dependence of the mechanical properties of
actin/-actinin gels on deformation rate. Nature
325,828
-830.[CrossRef][Medline]
Stopak, D. and Harris, A. K. (1982). Connective tissue morphogenesis by fibroblast traction. I. Tissue culture observations. Dev. Biol. 90,383 -398.[Medline]
Verkhovsky, A. B., Svitkina, T. M. and Borisy, G. G. (1999a). Network contraction model for cell translocation and retrograde flow. Biochem. Soc. Symp. 65,207 -222.[Medline]
Verkhovsky, A. B., Svitkina, T. M. and Borisy, G. G. (1999b). Self-polarization and directional motility of cytoplasm. Curr. Biol. 9,11 -20.[CrossRef][Medline]
Wakatsuki, T. and Elson, E. L. (2003). Reciprocal interactions between cells and extracellular matrix during remodeling of tissue constructs. Biophys. Chem.
Wakatsuki, T., Schwab, B., Thompson, N. C. and Elson, E. L.
(2001). Effects of cytochalasin D and latrunculin B on mechanical
properties of cells. J. Cell Sci.
114,1025
-1036.
Worthylake, R. A. and Burridge, K. (2001). Leukocyte transendothelial migration: orchestrating the underlying molecular machinery. Curr. Opin. Cell Biol. 13,569 -577.[CrossRef][Medline]
Zahalak, G. I. (1986). A comparison of the mechanical behavior of the cat soleus muscle with a distribution-moment model. J. Biomech. Eng. 108,131 -140.[Medline]
Zahalak, G. I., McConnaughey, W. B. and Elson, E. L. (1990). Determination of cellular mechanical properties by cell poking, with an application to leukocytes. J. Biomech. Eng. 112,283 -294.[Medline]