1 Department of Biomedical Engineering, Boston University,
Boston 02215; 2 Physiology Program, Department of
Environmental Health, Harvard School of Public Health, Boston,
Massachusetts 02115; and 3 Rugjer Bokovi
Institute, 10001 Zagreb, Croatia
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The tensegrity model hypothesizes that cytoskeleton-based microtubules (MTs) carry compression as they balance a portion of cell contractile stress. To test this hypothesis, we used traction force microscopy to measure traction at the interface of adhering human airway smooth muscle cells and a flexible polyacrylamide gel substrate. The prediction is that if MTs balance a portion of contractile stress, then, upon their disruption, the portion of stress balanced by MTs would shift to the substrate, thereby causing an increase in traction. Measurements were done first in maximally activated cells (10 µM histamine) and then again after MTs had been disrupted (1 µM colchicine). We found that after disruption of MTs, traction increased on average by ~13%. Because in activated cells colchicine induced neither an increase in intracellular Ca2+ nor an increase in myosin light chain phosphorylation as shown previously, we concluded that the observed increase in traction was a result of load shift from MTs to the substrate. In addition, energy stored in the flexible substrate was calculated as work done by traction on the deformation of the substrate. This result was then utilized in an energetic analysis. We assumed that cytoskeleton-based MTs are slender elastic rods supported laterally by intermediate filaments and that MTs buckle as the cell contracts. Using the post-buckling equilibrium theory of Euler struts, we found that energy stored during buckling of MTs was quantitatively consistent with the measured increase in substrate energy after disruption of MTs. This is further evidence supporting the idea that MTs are intracellular compression-bearing elements.
cytoskeleton; compression; energy; traction; tensegrity
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
MICROTUBULES (MTs) are structural components of the cytoskeleton that determine cell shape and polarity and that, in cooperation with the actomyosin network, facilitate processes such as cell locomotion and cytokinesis (cf. Refs. 1, 5). Although mechanical measurements in vitro indicate that MTs have high flexural rigidity, which suggests that they may support substantial longitudinal mechanical compression (9, 24, 38), it is not clear whether MTs play a similar role in living cells. The idea that MTs may support a substantial compression as they balance cytoskeleton contraction has become prominent with the emergence of the cellular tensegrity hypothesis (cf. Refs. 15-17). According to this hypothesis, the synergy of contraction and compression forces is essential for normal cellular function. Thus it is of considerable interest to investigate whether MTs do indeed play the role of compression-supporting elements of the cytoskeleton.
A number of previous observations appear to be consistent with the idea that MTs of living cells carry compression. First, for example, in response to cell contraction and mechanical perturbations, MTs buckle (40, 44). Second, in response to disruption of MTs, cells contract (3, 8, 25, 32, 34). Third, there is evidence of compression-induced MT disassembly in cultured smooth muscle cells (33). On the other hand, data from a recent study on cultured fibroblasts (12) suggests that MT-based cytoskeleton exhibits a fluid-like behavior in response to externally applied mechanical disturbance. Furthermore, it has been shown that nocodazole, a chemical that disrupts MTs, causes an increase in myosin phosphorylation in fibroblasts (22) and an increase in the intracellular Ca2+ in vascular smooth muscle cells (30). Consequently, the observed increase in cell contractility in response to nocodazole could be primarily a result of these responses and not a result of the loss of the load-supporting capacity of MTs. Thus the controversy about the role of MTs as a compression-supporting structure of the cytoskeleton needs to be resolved.
In this study, we attempted to elucidate whether MTs indeed carry a substantial compression as they balance cell contraction. We performed quantitative measurements of indices of MT compression in cultured human airway smooth muscle (HASM) cells by utilizing the traction force microscopy technique (42). We analyzed data from these measurements using a novel energetic approach. We found that in cultured HASM cells, compression of MTs balances a significant but relatively small fraction of cell contractile stress during cell maximal activation by histamine.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Traction force microscopy has been used to measure cell traction at the cell-substrate interface (4, 7, 31). From those measurements, one also can calculate the elastic energy stored in a flexible substrate during cell contraction (4). In our companion study (42), we used this technique to measure traction of cultured HASM cells in response to agonist-induced cell contraction. In the present study, we utilized this technique to assess compression in MTs in these cells.
Our working hypothesis is that cell contraction is balanced partly by
traction at the cell-substrate interface and partly by compression of
MTs (Fig. 1). To test this hypothesis, we
used traction force microscopy to measure cell traction in
stimulated cultured HASM cells before and after cell MTs were treated
by colchicine, a drug that disrupts MTs. If this hypothesis holds, then
for a given state of contraction, we predicted that after disruption of
MTs 1) traction would increase due to a transfer of the part
of the contractile stress balanced by MTs to the substrate and
2) energy stored in the substrate would increase due to
transfer of the compression energy of MTs to the substrate. Formal
definitions of the contractile stress, traction, and stored energy are
given later in the text.
|
Cell culture. HASM cells were cultured (105 cells/cm2) with Ham's F-12 medium supplemented with 10% fetal bovine serum, 50 mg/ml gentamicin, and 2.5 µm/ml amphotericin B. Cells at passages 3-6 were used for all experiments. After they reached confluence, cells were serum deprived for 48 h and then plated in serum-free defined medium. Cells were plated sparsely on a 70-µm-thick polyacrylamide elastic gel block coated with collagen type I (0.2 mg/ml). More details about cell culture can be found in our companion paper (42). We chose HASM cells because we showed previously that in this cell type, we can pharmacologically modulate cell contraction in a dose-dependent manner (13).
Traction force microscopy. A detailed description of this technique is given in our companion paper (42). Briefly, the polyacrylamide gel substrate was used as a strain gauge to measure the interfacial cell-substrate traction. Many fluorescent microbeads (0.2-µm diameter) embedded near the gel apical surface served as markers whose displacements were recorded as the adhering cell contracted. The bottom surface of the gel was covalently bonded to a flat, rigid plate, and the lateral surfaces were free. From bead displacements and known elastic properties of the gel (Young's modulus values of 870 and 1,300 Pa and a Poisson's ratio of 0.48), the traction was calculated as described in Refs. 4 and 42.
Protocol. HASM cells plated on a polyacrylamide gel block were first treated with 10 µM histamine, then with 1 µM colchicine plus 10 µM histamine, and, finally, with trypsin. This dose of histamine was shown to produce maximum increases in cell stiffness and myosin light chain phosphorylation. Histamine was added with colchicine to maintain the saturated bath concentration. Trypsin was added until a cell was completely detached from the substrate. Note that before the saturated dose of histamine was added, cells were treated with lower dosages (over 4 min) for the purpose of studies described in our companion paper (42).
Images of fluorescent microbeads were taken at baseline, at 40-s intervals after histamine addition, after colchicine addition, and, finally, after trypsin addition. The image after trypsin was used as the reference (traction-free) image. The displacement field (u) was calculated as follows: u at baseline, after histamine addition, and after colchicine addition was obtained by cross-correlating the corresponding image with the reference image. A detailed description of how u was calculated is given in Ref. 4. The displacement field u was then used to calculate the corresponding traction vector field (t) as described in Refs. 4 and 42; local traction was defined as a local contact force between the cell and the substrate per unit local contact surface area.Calculation of traction, strain energy stored in the substrate,
and the prestress.
The mean traction (
![]() |
(1) |
![]() |
(2) |
Immunosfluorescence staining of MTs. To find out how depolymerization of MTs in response to colchicine progressed with time, we serum deprived HASM cells for 24 h before they were plated overnight in a defined medium on type I collagen-coated (5 µg/ml) Lab-Tek chamber slides. The cells were treated with colchicine (1 µM) for 1, 3, 5, 10, and 15 min and then fixed, permeabilized, and stained using indirect immunofluorescence methods as described in Ref. 29.
Data analysis. Statistical differences were assessed by the paired t-test. Differences with P < 0.05 were considered significant.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Measurements were done on n = 13 cells. Six of
those cells were plated on a soft gel (Young's modulus of 870 Pa) and
seven on a stiff gel (Young's modulus of 1,300 Pa). No significant
difference between data obtained from these two gels was observed, and
therefore all data have been combined. Here we have presented the data
from the point when the traction reached its peak (~3 min after
histamine addition and 3-5 min after colchicine addition). In most
cells, 23 Pa) and ~30% (
Wt
0.13 pJ),
respectively. All of these increases were significant
(P < 0.05). The increases in
|
Results from immunofluorescence staining measurements show that
disruption of MTs was evident 3 min after colchicine addition (Fig.
2). The filamentous patterns of MTs
gradually started to disappear and MTs became disorganized as treatment
duration increased from 3 to 15 min (Fig. 2). Because most of our
traction measurements were done 3-5 min after colchicine addition,
it is apparent that structural integrity of a substantial fraction of
MTs was disrupted by this time. However, a fraction of MTs remained
intact even 15 min after colchicine addition (Fig. 2). Note that the
images in Fig. 2 serve only as a qualitative indicator that MTs started to disassemble within the duration of traction measurements.
|
We also calculated t, i.e., the component
of the prestress balanced by the traction, using the algorithm
described in our companion paper (42). We found that
t increased after MT disruption by
colchicine (Table 1). We interpreted this increase in
t as a part of the prestress balanced by
MTs (i.e.,
t =
MT). On average,
t changed by
t
288 Pa, which is ~14% of
t (Table 1); i.e., MTs balanced ~14%
of the prestress. Data from individual cells showed that this
contribution of MTs ranged from ~3 to 30%.
Together, the above results show that disruption of MTs by colchicine
caused significant increases in
We next investigated whether the observed increase in the energy stored
in the gel substrate after the colchicine treatment, Wt
0.13 pJ (Table 1), could be
accounted for by the compression energy (WMT)
stored in MTs before their disruption. To obtain WMT, we used a theoretical approach based on an
energetic analysis.
Energetic considerations.
Our working hypothesis predicts that if MTs indeed carry compression
and if there is no energy loss during MT disruption, then
WMT = Wt.
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Elucidation of biophysical mechanisms by which mechanical stresses are transmitted and balanced within the cell-substrate system is critical for understanding how mechanical signals affect cellular function. Previous studies have shown the role of the substrate in balancing cell contractile stress and how it may affect cell locomotion, spreading, or apoptosis (6, 11, 31). In this study we obtained for the first time prime quantitative data suggesting that MTs also contribute to the balance of the cell contractile stress. According to our measurements, it appears that in maximally activated HASM cells, MTs contribute ~3-30% (~14% on average) to the balance of the contractile prestress. The rest is balanced by the substrate.
In this study we used a novel energetic approach that had several advantages. First, energy is a scalar quantity, independent of the choice of coordinate system, with the property that the total energy is the sum of energies of various components. This simplified mathematical handling of the data. Second, energy analysis requires minimal specifications about cytoskeleton architecture. Results of the energetic analysis showed that 1) MTs buckle as they carry compression, and 2) IFs may play a stabilizing role in this process. These findings appear to be consistent with previous observations. First, it was observed that MTs of living cells buckle during cell contraction (40, 44). Second, the similarity between the observed wavy shape of buckled MTs of living cells and the shape predicted from the theory (2, 37) suggests that in living cells, MTs are indeed stabilized by the surrounding cytoskeleton structures. Finally, experimental data show that there exists mechanical interlinking between MTs and IFs in fibroblasts (36).
Paul et al. (30) found that the increase in contractility in unstimulated arteries is paralleled by a small increase in intracellular Ca2+ after colchicine treatment. They concluded that MTs do not significantly contribute to vascular smooth muscle mechanics but play a role in modulating Ca2+ signal transduction. On the other hand, recent measurements showed that there was no increase in intracellular Ca2+ in response to colchicine treatment of histamine-activated HASM cells, although colchicine alone induced a small increase in intracellular Ca2+ in unstimulated HASM cells (40). Consequently, the observed increase in traction, energy, and prestress after colchicine treatment in HASM cells (Table 1) could not be attributed to an increase in intracellular Ca2+. These results are consistent with published results in isolated arterioles (32).
A key assumption of this study was that the mechanical equilibrium and
the energy budget of the cell-substrate system were maintained by means
of a three-way force balance between the contractile elements,
cytoskeleton-based MTs, and the substrate. However, it is very likely
that other cellular structures also may contribute to the balance of
forces and energy budget. First, it was observed recently that stress
fibers of endothelial cells exhibit buckling in response to large
shortening of the substrate (14), suggesting that they may
carry compression. However, stress fibers isolated from fibroblasts and
endothelial cells on average shorten to about 23% of their initial
lengths, suggesting that under normal physiological conditions stress
fibers are indeed in tension (20). Second, we also do not
know the contribution of swelling pressure of the cytoplasm to the
force balance within the cell. However, the contribution of the
cytoplasm to the energy budget at steady state is zero because the
cytoplasm is virtually incompressible. Third, the contributions of the
actin cytoskeleton and myosin cross bridges to the energy budget did
not exceed the order of 102 pJ each (see
APPENDIX). These values are at least an order of magnitude smaller than the measured energy Wt stored in
the substrate and the estimated compression energy
WMT stored in MTs. Fourth, on the basis of data
from in vitro measurements, IF gels have much lower stiffness than
actin gels, except at high strains (18), suggesting that
the energy stored in IFs is substantially smaller than the energy
stored in the actin network, which itself has little contribution to
energy storing. Thus we concluded that in the cell-gel substrate
system, the MTs and the gel substrate had the most prominent
contribution to the energy budget. The other contributions appear to be
less important. Nevertheless, their inclusion would somewhat reduce our
estimate of the contribution of MTs to the cell energy budget.
It was assumed that there was no energy dissipation in the gel
substrate-cell system, i.e., that the system is elastic. Although the
gel behavior has been shown to be almost elastic (31), it is not true for HASM cells, which are known to exhibit a dissipative, viscoelastic behavior (13). Nevertheless, all of our
measurements were done at the steady state when all viscoelastic
stresses are dissipated, and therefore, cellular viscoelasticity should
have little effect on data for the traction, energy, and prestress. On
the other hand, it is likely that during disruption of MTs by
colchicine, a portion of the energy stored in MTs was irreversibly lost. Hence, only a fraction of the energy associated with MTs might be
transferred to the substrate, and thus the observed energy increase
Wt could be an underestimate. This, in turn,
may compensate the overestimates caused by our exclusion of the
contributions of actin, myosin, and IFs to the energy storage of the cell.
The fact that the traction measured on soft gels (Young's modulus of 870 Pa) did not differ significantly from the values obtained on hard gels (Young's modulus of 1,300 Pa) can be explained as follows. First, gel stiffness varied from experiment to experiment: the soft gel range of stiffness was 860-1,000 Pa, whereas the hard gel range of stiffness was 920-1,600 Pa. Thus in some measurements the gel stiffness was not different at all. Second, Yu-Li Wang's group (39) have shown that an increase in gel stiffness by a factor of ~2.4 causes a modest increase in traction of ~50%. Thus it appears that traction measurements are not very sensitive to changes in the substrate stiffness.
We do not know whether colchicine affects actin polymerization and thus the state of stress within the actin network of HASM cells. Earlier studies showed that colchicine does not cause disruption of actin filaments but does cause a small increase in cytoskeleton-associated actin in leukocytes (21) and a significant increase in filamentous actin in fibroblasts (19). The latter finding corresponds to a 1-h period, whereas the colchicine treatment in our measurements did not exceed 10 min. This, in turn, suggests that if there were changes in the amount of actin in our measurements, they might not be large.
A critical review of the assumptions of our theoretical analysis is
given below. The crudest assumptions of the analysis were the length of
MTs of 20 µm and the stiffness of the lateral support of IFs of 8 Pa.
With the assumption that the MTs spread outwardly from the cell
perinuclear region, and taking into account that the length and width
of spread airway smooth muscle cells are roughly 100 and 30 µm,
respectively, our assumption that MTs span an average distance of ~20
µm was not unreasonable. Furthermore, Brodland and Gordon
(2) used the same value in their analysis of MT buckling.
The assumed stiffness of IFs of 8 Pa is consistent with in vitro
measurements on vimentin (18) and keratin
(27) gels. In general, vimentin IF contributes about 20%
of cytoskeleton stiffness in endothelial cells and fibroblasts
(41). However, desmin is the most abundant IF in HASM
cells (10), and few data on mechanical properties of
desmin in these cells are available. Furthermore, the interlinking
between IFs and MTs within the cytoskeleton is facilitated by plectin
(36), whose mechanical properties are not known. It is
likely that other cytoskeleton structures and the viscoelastic
cytoplasm also play a role in stabilizing MTs. Thus the assumed value
of 8 Pa of the stiffness of the lateral support is questionable but
seems to be a good guess for the following reasons. First, a 25%
increase in this value would overly stabilize MTs (i.e., no buckling
would occur), which is contrary to previously observed buckled shapes
of MTs (40, 44), albeit in cells other than smooth muscle
cells. If MTs did not buckle, then the energy associated with their
compression would be much smaller (order of 104 pJ, see
APPENDIX) than the measured value of the energy. Second, a
25% decrease in the assumed value of stiffness of the lateral support
would result in an ~2.5-fold greater value of
WMT than the measured increase in energy,
Wt. We also performed an order-of-magnitude analysis of WMT by using the method described in
the APPENDIX and assuming that l =
O(101) µm, q = O(101) Pa, B = O(101) pN · µm2, and
AMT = O(102)
nm2, where O(10x) denotes
the order of magnitude. We estimated that
WMT = O(10
1-100) pJ. The measured
value of
Wt falls within this range.
In conclusion, our study showed that a greater part of contractile stress of cultured HASM cells was balanced by the substrate, but a significant portion of this stress was balanced by the compression-supporting network of MTs. To our knowledge, these are the first prime quantitative experimental data showing that MTs behave as compression-bearing elements as they balance the contractile stress. On the basis of our energetic analysis, we have concluded that MTs buckle as they carry compression and that, in this process, IFs (and possibly other intracellular structures) stabilize MTs.
Our findings and the findings from our companion study (42) have an important implication on the tensegrity idea. Key features of the cellular tensegrity hypothesis are that the cell stiffness increases in proportion with the cytoskeleton prestress and that the prestress is balanced by intracellular compression-supporting elements (e.g., MTs). Although results from our two studies are consistent with these features, the fact that the MTs balance only a relatively small fraction of the prestress in this spread HASM cell suggests that one may not need to invoke tensegrity to explain the prime feature of contractile cell deformability, i.e., the association of cell stiffness with the cytoskeleton prestress. Consequently, the choice of a model of cell deformability among various prestressed structures (e.g., tensegrity, cable nets, cortical membrane) is likely to depend on the cell type, the extent of cell spreading, or some other factors.
![]() |
APPENDIX |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The energy of buckling of MTs, WMT, is calculated as follows. MTs were depicted as slender elastic cylindrical rods of length l, cross-sectional area AMT, and bending (flexural) rigidity B, supported by a lateral continuous support of IFs of stiffness q. This lateral support effectively reduces the critical buckling length (Lcr) of MTs; Lcr < l. Thus the problem of buckling of a laterally supported strut of length l reduces to the problem of buckling of a simple Euler pin-ended strut of length Lcr with no lateral support, known as elastica (37). This makes the calculation of buckling energy much simpler.
Lcr was obtained from a theoretical relationship
(37) shown in Fig. 3 in a graphical form. The graph in
Fig. 3 was calculated for
q = 8 Pa and B = 21.5 pN · µm2 (9). For l = 20 µm, it was found that Lcr/l = 0.14, and hence Lcr = 2.8 µm. This value was
then used to calculate the critical buckling stress
(Scr) for the Euler elastica as
Scr = (2B)/[(Lcr)2AMT],
where AMT = 190 nm2
(9). It was found that Scr
142 kPa.
|
We next used the theory of postbuckling equilibrium of Euler elastica
(37) to calculate WMT. According to
this theory, the buckling is not a catastrophic event, and compressed
elastica maintains equilibrium after the compression stress exceeds the critical buckling stress, i.e., SMT > Scr (see Fig.
4, inset). The universal
relationship between the compression stress and the chord length
L of the elastica is given in Fig. 4. The area under the
curve corresponds to the energy associated with buckling.
|
Using the value of SMT 152 kPa
determined from the experimental data for prestress (see
Energetic consideration) and Scr
142 kPa, we obtained L/Lcr = 0.87 from Fig. 4. The energy per unit volume
(wMT) of the elastica was obtained as
![]() |
(A1) |
If MTs do not buckle but only shorten under compressive stress
SMT 152 kPa, the energy stored per unit
volume of a single MT is
(SMT)2/2EMT,
assuming that MTs are linearly elastic. Here
EMT
1.2 GPa is the Young's modulus of
a single MT (9). The corresponding energy stored in MTs of
the cytoskeleton is WMT =
MTV(SMT)2/2EMT.
It was determined that WMT
0.9 × 10
4 pJ.
The energy stored in the actin network (WMF) was
calculated as follows. It was assumed that the actin filaments were
linearly elastic. In that case, the energy per unit volume of a single actin microfilament is
(SMF)2/2EMF,
where SMF is stress and
EMF 2.6 GPa (9) is the
Young modulus of the filament. The total energy stored in the actin cytoskeleton is therefore equal to WMF =
MFV(SMF)2/2EMF,
where
MF
0.21% is the volume fraction of actin
microfilaments in the cell (cf. Ref. 35) and V = 5,000 µm3 is the cell volume. Stress
SMF was obtained from the data for the net mean
prestress = 2,211 Pa (42), assuming that the
microfilaments form a three-dimensional randomly oriented network where
SMF = 3
MF
(35). It was found that WMF
0.02 pJ.
The energy stored in the myosin cross bridges
(WCB) was obtained as follows. The maximum
energy per cross bridge is eCB = 2.7×108 pJ (26). Thus the total energy
stored in myosin cross bridges of the cell is
WCB =
CBVeCB/vCB, where
CB is the volumetric fraction of the cross bridges in
the cell, vCB is the volume of a cross bridge, and V = 5,000 µm3 is the cell volume. The quantities
CB and vCB were obtained as follows. The
myosin content in the smooth muscle is approximately one-fifth the
myosin content in the skeletal muscle cells, which is 20 µM
(28), and the myosin mass density was assumed to be that
of water. From these data it was obtained that
CB
0.18%. The crude estimate of vCB
10
5 µm3 was obtained from the data for the
cross-bridge geometry (43). Thus it was found that
WCB
0.024 pJ.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. Don Ingber for making his laboratory available for performance of some of the experiments.
![]() |
FOOTNOTES |
---|
This work was supported by National Heart, Lung, and Blood Institute Grants HL-65371 and HL-33009 and by National Aeronautics and Space Administration Grant NAG5-4839.
Address for reprint requests and other correspondence: D. Stamenovi, Dept. of Biomedical Engineering, Boston Univ., 44 Cummington St., Boston, MA 02215 (E-mail:
dimitrij{at}engc.bu.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
10.1152/ajpcell.00271.2001
Received 15 June 2001; accepted in final form 24 October 2001.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Amos, LA,
and
Amos WB.
Molecules of the Cytoskeleton. New York: Guilford, 1991.
2.
Brodland, GW,
and
Gordon R.
Intermediate filaments may prevent buckling of compressively loaded microtubules.
ASME J Biomech Eng
112:
319-321,
1990[ISI][Medline].
3.
Brown, RA,
Talas G,
Porter RA,
McGrouther DA,
and
Estwood M.
Balanced mechanical forces and microtubule contribution to fibroblast contraction.
J Cell Physiol
169:
439-447,
1996[ISI][Medline].
4.
Butler, JP,
Toli-Nørrelykke IM,
and
Fredberg JJ.
Estimating traction fields, moments, and strain energy that cells exert on their surroundings.
Am J Physiol Cell Physiol
282:
C595-C605,
2002
5.
Canman, JC,
and
Bement WM.
Microtubules suppress actomyosin-based cortical flow in Xenopus oocytes.
J Cell Sci
110:
1907-1917,
1997
6.
Chen, CS,
Mrksich M,
Huang S,
Whitesides GM,
and
Ingber DE.
Geometric control of cell life and death.
Science
276:
1425-1428,
1997
7.
Dembo, M,
and
Wang YL.
Stress at the cell-to-substrate interface during locomotion of fibroblasts.
Biophys J
76:
2307-2316,
1999
8.
Gills, JP,
Roberts BC,
and
Epstein DL.
Microtubule disruption leads to cellular contraction in human trabecular meshwork cells.
Invest Opthalmol Vis Sci
39:
653-658,
1998[Abstract].
9.
Gittes, F,
Mickey B,
Nettleton J,
and
Howard J.
Flexural rigidity of microtubules and actin filaments measured from thermal fluctuations in shape.
J Cell Biol
120:
923-934,
1993[Abstract].
10.
Halayko, AJ,
Salari H,
Ma X,
and
Stephens NL.
Markers of airway smooth muscle cell phenotype.
Am J Physiol Lung Cell Mol Physiol
270:
L1040-L1051,
1996
11.
Harris, AK,
Wild P,
and
Stopak D.
Silicon rubber substrata: a new wrinkle in the study of cell locomotion.
Science
208:
177-179,
1980[ISI][Medline].
12.
Heidemann, SR,
Kaech S,
Buxbaum RE,
and
Matus A.
Direct observations of the mechanical behaviors of the cytoskeleton in living fibroblasts.
J Cell Biol
145:
109-122,
1999
13.
Hubmayr, RD,
Shore SA,
Fredberg JJ,
Planus E,
Panettieri RA, Jr,
Moller W,
Heyder J,
and
Wang N.
Pharmacological activation changes stiffness of cultured airway smooth muscle cells.
Am J Physiol Cell Physiol
271:
C1660-C1668,
1996
14.
Hucker, W,
Yin FCP,
and
Costa KD.
The role of cytoskeletal tension in maintaining actin stress fiber integrity (Abstract).
Ann Biomed Eng
28:
S-65,
1999.
15.
Ingber, DE.
Cellular tensegrity: defining new rules of biological design that govern the cytoskeleton.
J Cell Sci
104:
613-627,
1993
16.
Ingber, DE.
The architecture of life.
Sci Am
278:
48-57,
1998[ISI][Medline].
17.
Ingber, DE.
Tensegrity: the architectural basis of cellular mechanotransduction.
Annu Rev Physiol
59:
575-599,
1997[ISI][Medline].
18.
Janmey, PA,
Eutenauer U,
Traub P,
and
Schliwa M.
Viscoelastic properties of vimentin compared with other filamentous biopolymer networks.
J Cell Biol
113:
155-160,
1991[Abstract].
19.
Jung, HI,
Shin I,
Park YM,
Kang KW,
and
Ha KS.
Colchicine activates actin polymerization by microtubule depolymerization.
Mol Cell
7:
431-437,
1997.
20.
Katoh, K,
Kano Y,
Masuda M,
Onishi H,
and
Fujiwara K.
Isolation and contraction of the stress fiber.
Mol Biol Cell
9:
1919-1938,
1998
21.
Keller, HU,
and
Niggli V.
Colchicine-induced stimulation of PMN motility related to cytoskeletal changes in actin, -actinin, and myosin.
Cell Motil Cytoskeleton
25:
10-18,
1993[ISI][Medline].
22.
Kolodney, MS,
and
Elson EL.
Contraction due to microtubule disruption is associated with increased phosphorylation of myosin regulatory light chain.
Proc Natl Acad Sci USA
92:
10252-10256,
1995[Abstract].
23.
Kolodney, MS,
and
Wysolmerski RB.
Isometric contraction by fibroblasts and endothelial cells in tissue culture: a quantitative study.
J Cell Biol
117:
73-82,
1992[Abstract].
24.
Kurachi, M,
Hoshi M,
and
Tashiro H.
Buckling of single microtubule by optical trapping forces: direct measurement of microtubule rigidity.
Cell Motil Cytoskeleton
30:
221-228,
1995[ISI][Medline].
25.
Leite, R,
and
Webb RC.
Microtubule disruption potentiates phenylephrine-induced vasoconstriction in rat mesenteric arterial bed.
Eur J Pharmacol
351:
R1-R3,
1998[ISI][Medline].
26.
Linari, M,
Dobbie I,
Reconditi M,
Koubassova N,
Irving M,
Piazzesi G,
and
Lombardi V.
The stiffness of skeletal muscle in isometric contraction and rigor: the fraction of myosin heads bound to actin.
Biophys J
74:
2459-2473,
1998
27.
Ma, L,
Xu J,
Coulombe PA,
and
Wirtz D.
Keratin filament suspensions show unique micromechanical properties.
J Biol Chem
274:
19145-19151,
1999
28.
MacMahon, TA.
Muscles, Reflexes, and Locomotion. Princeton, NJ: Princeton Univ. Press, 1984.
29.
Mooney, DJ,
Hansen LK,
Langer R,
Vacanti JP,
and
Ingber DE.
Extracellular matrix controls tubulin monomer levels in hepatocytes.
Mol Biol Cell
5:
1281-1288,
1994[Abstract].
30.
Paul, RJ,
Bowman P,
and
Kolodney MS.
Effects of microtubule disruption on force, velocity, stiffness and [Ca2+]i in porcine coronary arteries.
Am J Physiol Heart Circ Physiol
279:
H2493-H2501,
2000
31.
Pelham, RJ, Jr,
and
Wang YL.
Cell locomotion and focal adhesions are regulated by substrate flexibility.
Proc Natl Acad Sci USA
94:
13661-13665,
1997
32.
Platts, SH,
Falcone JC,
Holton WT,
Hill MA,
and
Meininger GA.
Alteration of microtubule polymerization modulates arteriolar vasomotor tone.
Am J Physiol Heart Circ Physiol
277:
H100-H106,
1999
33.
Putnam, AJ,
Cunningham JJ,
Dennis RG,
Linderman JJ,
and
Mooney DJ.
Microtubule assembly is regulated by externally applied strain in cultured smooth muscle cells.
J Cell Sci
111:
3379-3387,
1998
34.
Sheridan, BC,
McIntyre RC, Jr,
Meldrum DR,
Cleveland JC, Jr,
Agrafojo J,
Banerjee A,
Harken AH,
and
Fullerton DA.
Microtubules regulate pulmonary vascular smooth muscle contraction.
J Surg Res
62:
284-287,
1996[ISI][Medline].
35.
Stamenovi, D,
and
Coughlin MF.
The role of prestress and architecture of the cytoskeleton and deformability of cytoskeletal filaments in mechanics of adherent cells: a quantitative analysis.
J Theor Biol
201:
63-74,
1999[ISI][Medline].
36.
Svitkina, TM,
Verkhovsky AB,
and
Borisy GG.
Plectin sidearms interaction of intermediate filaments with microtubules and other components of the cytoskeleton.
J Cell Biol
135:
991-1007,
1996[Abstract].
37.
Timoshenko, SP,
and
Gere JM.
Theory of Elastic Stability (2nd ed.). New York: McGraw-Hill, 1988.
38.
Venier, P,
Maggs AC,
Carlier MF,
and
Pantaloni D.
Analysis of microtubule rigidity using hydrodynamic flow and thermal fluctuations.
J Biol Chem
269:
13353-13360,
1994
39.
Wang, HB,
Dembo M,
and
Wang YL.
Substrate flexibility regulates growth and apoptosis of normal but not transformed cells.
Am J Physiol Cell Physiol
279:
C1345-C1350,
2000
40.
Wang, N,
Naruse K,
Stamenovi D,
Fredberg JJ,
Mijailovich SM,
Toli
-Nørrelykke IM,
Polte T,
Mannix R,
and
Ingber DE.
Mechanical behavior of living cells consistent with the tensegrity model.
Proc Natl Acad Sci USA
98:
7765-7770,
2001
41.
Wang, N,
and
Stamenovi D.
Contribution of intermediate filaments to cell stiffness, stiffening, and growth.
Am J Physiol Cell Physiol
279:
C188-C194,
2000
42.
Wang, N,
Toli-Nørrelykke IM,
Chen J,
Mijailovich SM,
Butler JP,
Fredberg JJ,
and
Stamenovi
D.
Cell prestress. I. Stiffness and prestress are closely associated in contractile adherent cells.
Am J Physiol Cell Physiol
282:
C606-C616,
2002
43.
Warshaw, DM.
The in vitro motility assay: a window into myosin molecular motor.
News Physiol Sci
11:
1-7,
1996
44.
Waterman-Storer, CM,
and
Salmon ED.
Actomyosin-based retrograde flow of microtubules in the lamella of migrating epithelial cells influences microtubule dynamic instability and turnover is associated with microtubule breakage and treadmilling.
J Cell Biol
139:
417-434,
1997