Departments of 1 Chemical Engineering, 2 Biomedical Engineering, and 3 Biologic and Materials Sciences, University of Michigan, Ann Arbor, Michigan 48109-2136
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
A number of studies have suggested that externally applied mechanical forces and alterations in the intrinsic cell-extracellular matrix (ECM) force balance equivalently induce changes in cell phenotype. However, this possibility has never been directly tested. To test this hypothesis, we directly investigated the response of the microtubule (MT) cytoskeleton in smooth muscle cells to both mechanical signals and alterations in the ECM. A tensile force that resulted in a positive 10% step change in substrate strain increased MT mass by 34 ± 10% over static controls, independent of the cell adhesion ligand and tyrosine phosphorylation. Conversely, a compressive force that resulted in a negative 10% step change in substrate strain decreased MT mass by 40 ± 6% over static controls. In parallel, increasing the density of the ECM ligand fibronectin from 50 to 1,000 ng/cm2 in the absence of any applied force increased the amount of polymeric tubulin in the cell from 59 ± 11% to 81 ± 13% of the total cellular tubulin. These data are consistent with a model in which MT assembly is, in part, controlled by forces imposed on these structures, and they suggest a novel control point for MT assembly by altering the intrinsic cell-ECM force balance and applying external mechanical forces.
cytoskeleton; mechanotransduction; smooth muscle cells; tubulin; tensegrity
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
MECHANICAL SIGNALS play an essential role in both normal and pathological development of a variety of tissue types, including bone (19), muscle (54), and vascular tissue (46, 60). These mechanical signals occur in vivo in the context of a complex extracellular matrix (ECM), which, by itself, is a critical determinant of the cellular microenvironment. Mechanical signals have been shown to alter cell proliferation (61, 62), production of ECM proteins (4, 8), and gene expression (53), often in an ECM-dependent manner (38, 41, 62). Likewise, signals from the ECM itself have been shown to regulate integrin-mediated signaling events during normal development and neoplastic transformation, generating signals that control growth, differentiation, motility, apoptosis, and matrix synthesis and remodeling (1, 2, 26, 31). Although the chemical composition of the ECM is clearly important, there is increasing evidence that mechanical cues from the matrix may be just as important (10, 32).
Previous studies have shown that altering the mechanics of the matrix without changing the chemical composition is enough to alter cellular behavior (9, 23, 28, 36). The mechanisms utilized by cells to transmit this mechanical information into a change in phenotype remain poorly understood and the subject of ongoing debate. Two dominant, although not necessarily mutually exclusive, paradigms have emerged to explain the responses of cells to changes in the mechanical microenvironment. In the first hypothesis, changes in the mechanics of the ECM trigger a cascade of soluble chemical second messengers, likely derived from the cluster of signaling proteins at the sites of cell adhesion (46, 52). In the second hypothesis, based on models of tensegrity architecture first put forth by Fuller (24, 33, 48), cells exist in an equilibrium balance of forces. According to this hypothesis, altered mechanics of the ECM could directly induce changes in this preexisting cellular force balance, resulting in alterations in the mechanics of the cytoskeleton. These changes may then directly trigger changes in gene expression or influence a variety of signaling pathways (6, 10, 32). An abundant amount of evidence in support of both models has led to increased debate on the subject of mechanotransduction. The cytoskeleton is clearly involved in the response to mechanical forces given that it is altered in response to a variety of externally applied mechanical signals in many different cell types (42, 43, 49, 51, 59). Furthermore, changing the flexibility of an adhesion substrate has been shown (47) to regulate cell migration and focal adhesion structure in a manner that is dependent on the force-generating capabilities of the actin cytoskeleton. In another study (12), an optical trap was used to simulate an increased ECM rigidity, causing strengthening of interaction between integrins and the cytoskeleton proportional to the restraining force of the trap. Likewise, applying forces to integrin ligand-coated beads increased the stiffness of the cytoskeleton in a linear relationship to the amount of stress applied (59).
Microtubules (MTs), one element of the cytoskeleton, may be involved in the mechanotransduction response by altering their assembly in response to compressive forces (5, 34, 49). Two theoretical models predict that altering compressive forces on MTs can alter their state of assembly (5, 30) and hence provide a testable hypothesis that forms the basis of this study. These models suggest that a pseudoequilibrium among tubulin monomers, MTs, and compressive forces inherent on the MTs could be altered by changing the amount of force on the MTs. In this study, the mechanical microenvironment was altered to test these theoretical models directly. We demonstrated control of MT assembly in smooth muscle cells (SMCs) both by applying single external mechanical forces and by changing the density of ECM ligand used for cell adhesion. MT assembly was induced by applying a positive step change in strain to the cell culture surface, imparting a tensile force to the adherent cells. MT disassembly was induced after the application of a negative strain to the substrate, effectively imparting a compressive force to the adherent cells. Changing the density of adsorbed fibronectin in static culture also shifted the pseudoequilibrium set point between tubulin monomeric subunits and MTs, with an increased density of ligand leading to an increased mass of MTs. Combined, these findings support previously published models predicting that MT assembly is regulated by imposed compressive forces.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
SMC culture.
Vascular SMCs were isolated from thoracic rat aortas with an adaptation
of a previously published technique (49, 50). Routine SMC
culture was performed with medium 199 (M199) supplemented with 10%
fetal bovine serum (FBS), 100 U/ml penicillin, and 0.1 mg/ml
streptomycin (all from GIBCO, Gaithersburg, MD). Immunofluorescence localization of a smooth muscle-specific -actin marker confirmed that these cells were SMCs (data not shown). SMCs between passage 4 and 15 were used in all studies. For serum-free
experiments in this study, cells were cultured in standard
serum-containing medium until they were ready to be used for an
experiment. Cells were then harvested with trypsin followed by either
serum-containing M199 or a soybean trypsin inhibitor solution (0.75 mg/ml) in PBS with 5 mM EDTA. After centrifugation, cells were washed
with PBS for removal of any residual serum, centrifuged again, and
subsequently resuspended in a completely defined serum-free medium
[CSC SF Media Kit supplemented with RocketFuel additive (1:50); Cell
Systems, Kirkland, WA].
Application of external step changes in strain to the cultured SMCs. Cells were plated out at densities between 20,000 and 30,000 cells/cm2 on six-well culture dishes made of silicon rubber (BioFlex plates; Flexcell, Hillsborough, NC). These plates, initially untreated, were coated with a theoretical density of 1 µg/cm2 (accounting for the additional surface area created by the volume of the liquid) of either type I collagen (Vitrogen 100; Cohesion, Palo Alto, CA) or fibronectin (human plasma fibronectin; GIBCO). ECM proteins were adsorbed with the use of a carbonate/bicarbonate buffer (15 mM Na2CO3 and 35 mM NaHCO3, pH 9.4). Cells cultured on these substrates were allowed to attach and spread for ~24 h. Dishes were exposed to step changes in strain with the use of a custom-built device that we have previously described (49). We have previously characterized this device and shown that the strain is linear with respect to vertical displacement and that the strain is a true biaxial strain.
Extraction of MT and total tubulin fractions from cultured cells. MTs and total tubulin were differentially extracted from cultured SMCs as previously described (7, 44, 49). Briefly, to isolate polymeric tubulin (MTs), cells were washed in an MT stabilization buffer (MTSB) and incubated once for 15 min with MTSB plus 0.1% Triton X-100. This treatment permeabilizes the cell membrane, flushing out soluble cytoplasmic proteins, including monomeric tubulin. The remaining cytoskeletal ghosts were subsequently solubilized in a lysis buffer. Total tubulin was extracted by adding lysis buffer directly to intact cells. All extraction steps were performed at 37°C with prewarmed reagents to prevent any cold depolymerization of MTs. We have previously validated these methods by quantifying the distribution of tubulin in SMCs exposed to either nocodazole or paclitaxel, accurately quantifying nocodazole-induced MT depolymerization and paclitaxel-induced MT assembly (49).
Quantification of tubulin mass.
A competitive ELISA technique (58) was performed as
previously described (49). A mouse monoclonal
anti--tubulin antibody (diluted 1:500; Boehringer Mannheim,
Indianapolis, IN), followed by an alkaline phosphatase-conjugated goat
anti-mouse IgG (diluted 1:2,000; Bio-Rad, Hercules, CA), was used to
assay the tubulin content in both Triton X-100-insoluble and total
cellular lysates. A purified bovine brain tubulin standard (Molecular
Probes, Eugene, OR) was assayed in parallel to generate a standard
curve. After the substrate for the enzyme (Bio-Rad) was added to
generate a colored reaction product, plates were read at 405 nm with a
microplate reader (Vmax kinetic microplate reader;
Molecular Devices, Sunnyvale, CA). Absorbance values over a range of
dilutions were used to determine the tubulin concentration in the
experimental samples to ensure that the absorbance values were in the
linear region and that the signal was not saturated. All values for
tubulin mass determined in this study are presented as values relative to control conditions.
Inhibition of tyrosine phosphorylation with genistein. Certain experiments were performed with cells cultured in the presence of a tyrosine phosphorylation inhibitor, genistein (Sigma). SMCs cultured for 1-2 days on static six-well culture dishes in normal serum-containing M199 were subsequently serum starved for 18-36 h to reduce any basal levels of tyrosine phosphorylation caused by the presence of serum. After serum starvation, cells were subjected to genistein (between 10 µM and 1 mM final concentration) for 15 min before serum was added back to their culture environment. The original lyophilized genistein stock was solubilized in DMSO, so an amount of DMSO equivalent to that in the 500 µM genistein-treated sample was added to the untreated cultures to confirm that DMSO alone had no effect on tyrosine phosphorylation. After 15 min of exposure to the serum-containing environment, cells were lysed. Equal protein amounts (1.5 µg of total cellular protein) of these samples were separated by gel electrophoresis. Western blots for phosphotyrosine (using a mouse monoclonal anti-phosphotyrosine antibody, clone PY-20; BD Transduction Laboratories, Lexington, KY) were used to determine the optimal dosage of genistein that reduced total cellular tyrosine phosphorylation without affecting cell adhesion and morphology.
Controlling cell-matrix interactions and quantification of spread cell area. For experiments investigating the role of ECM density, SMCs were cultured on different densities of fibronectin on bacteriologic six-well culture dishes in a completely serum-free defined medium (CSC SF Media Kit supplemented with 1:50 diluted RocketFuel supplement, Cell Systems). Briefly, culture dishes were precoated overnight with a theoretical density of 50, 100, 500, or 1,000 ng/cm2 fibronectin. On the day of cell seeding, plates were washed twice with PBS and then blocked in serum-free M199 supplemented with 2% bovine albumin fraction V (GIBCO) for 30 min in a 37°C, 5% CO2 humidified environment. During the blocking step, cells were prepared by routine trypsinization, with the trypsin inhibited by using either serum-containing M199 or a soybean trypsin inhibitor solution as described in SMC culture. After centrifugation, the cells were resuspended in serum-free medium to be plated onto the bacteriologic culture dishes. Cells were seeded to obtain a density of 6,000-20,000 cells/cm2 after attaching and spreading for 24 h. At these densities, cell-to-cell contact was minimized. After 24 h, Triton X-100-insoluble and total protein fractions were extracted as described in Extraction of MT and total tubulin fractions from cultured cells, with parallel wells used for cell counts to normalize the assay results. Polymeric and total tubulin masses were assayed from these samples by using the competitive ELISA. Parallel wells for each condition were stained with the membrane-specific lipophilic dye DiI (Molecular Probes) at a concentration of 10 µg/ml in serum-free M199. Cells were stained for 30 min in a 37°C, 5% CO2 humidified environment before subsequently being fixed. Images of cells viewed on a Nikon Diaphot inverted fluorescent microscope (Nikon, Tokyo, Japan) were captured with the use of a digital camera (Optronix DEI-7000; Optronics Engineering, Goleta, CA) with a digital image grabber board (Scion, Frederick, MD) inside a Macintosh G4 computer. The number of pixels corresponding to the projected cell area in a captured image was quantified by using NIH Image software (National Institutes of Health, Bethesda, MD).
Immunofluorescence localization of 5-integrin in
SMCs.
Immunofluorescence microscopy of the
5-integrin in SMCs
cultured on increasing densities of fibronectin matrix was performed by
using standard methodologies. Cells cultured for 24 h in a serum-free environment on Lab-Tek chamber slides coated with 50, 100, 500, or 1,000 ng/cm2 fibronectin were fixed for 20 min in a
4% formaldehyde solution in a cytoskeletal buffer with sucrose (10 mM
MES, pH 6.1, 138 mM KCl, 3 mM MgCl, and 2 mM EGTA). A rabbit monoclonal
antibody against the
5-integrin receptor (1:100
dilution; Chemicon, Temecula, CA), followed by a secondary
rhodamine-conjugated donkey anti-rabbit antibody (1:100 dilution;
Jackson ImmunoResearch Laboratories, West Grove, PA), was used to
localize the
5-integrin.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Microtubule assembly is modulated by externally applied strain.
In the first part of this study, we directly tested the hypothesis that
tensile forces applied to the adhesion substrate lead to an increase in
net MT assembly. SMCs were cultured in 10% serum-containing medium on
silicon rubber substrates precoated with type I collagen. After 24 h, cells were exposed to a single externally applied 10% step change
in strain and held in the strained position for finite time periods
ranging from 1 min to 1 h. Triton X-100-insoluble (n = 6) and total cellular lysates (n = 6) were obtained from cells held in the stretched position.
Quantification of MT mass with the use of a competitive ELISA revealed
that a positive step change in external strain drives increased MT
assembly (Fig. 1). This response occurred
rapidly (after 1 min of tensile load) and peaked at ~15 min after
application of the strain, with a statistically significant 34 ± 10% increase (P < 0.05) in MT mass over unstretched controls. The response may occur even faster, but the limitations of
the protein extraction methods make resolution of <1 min impossible at
this time. The MT mass remained elevated before returning to near
unstretched levels after 1 h. In all experiments conducted in this
study, quantification of total cellular tubulin levels revealed no
change (data not shown). This finding suggests that the time scale of
these experiments was short enough to prevent de novo protein synthesis
from influencing the level of total cellular tubulin within the cell. A
constant level of total cellular tubulin suggests that MT assembly
occurred from the preexisting cytoplasmic pool of - and
-tubulin
monomers in response to a positive step change in strain applied to the
substrate.
|
|
Strain-driven microtubule assembly occurs to the same extent on
both type I collagen and fibronectin.
We next examined whether strain-driven MT assembly is a widely
applicable mechanotransduction response, observed in a variety of
culture and matrix conditions. To address this hypothesis, we
investigated the dependence of strain-driven MT assembly on the ECM
ligand. Cells cultured on 1 µg/cm2 fibronectin or type I
collagen in a defined, serum-free medium were subjected to a single
positive strain, as in the first set of experiments. Triton
X-100-insoluble (n = 6) and total cellular lysates
(n = 6) were extracted 15 min after a 10% strain was
initiated in the substrate. Quantification of tubulin in these two
types of lysates by competitive ELISA revealed that the MT mass
increased in response to the increased strain for cells cultured on
both ECM molecules studied (Fig.
3A). The magnitude of the MT
assembly response was similar to that seen for the conditions used in
the first set of experiments (i.e., collagen-coated substrate in a serum-containing medium). The assembly of MTs in response to the tensile strain occurred from the cytoplasmic pool of monomeric tubulin,
as evidenced by the ~80% decrease in the amount of available monomer
between the static and stretched conditions (Fig. 3B). Nearly identical results were obtained in all serum-free studies whether serum-containing M199 or a soybean trypsin inhibitor solution was used to inhibit the action of trypsin after cells were harvested to
be plated out for these experiments (data not shown). These findings
demonstrate that a positive step change in strain drives net MT
assembly for SMCs cultured on either type I collagen or fibronectin,
suggesting that the chemical identity of the matrix may not be critical
in triggering strain-driven MT assembly.
|
Strain-driven microtubule assembly occurs independently of tyrosine
phosphorylation.
A number of studies in the literature have determined that some
cellular responses to applied mechanical forces are dependent on a
number of soluble signaling cascades. Many signals from growth factors
and the ECM converge via phosphorylation of tyrosine residues on
various proteins (31, 52), and many tyrosine
phosphorylation events occur in response to mechanical signals
(46, 51, 60). To eliminate the influence of these events,
we used the inhibitor genistein. Control experiments were first
performed to determine the optimal dosage of genistein for experiments
in which SMCs were subjected to strain. A range of concentrations of
genistein were screened for their ability to reduce serum-induced
tyrosine phosphorylation in SMCs (Fig.
4A), and both 500 µM and 1 mM concentrations of genistein reduced the levels of serum-induced
tyrosine phosphorylation (note the disappearance of bands above 132 kDa
and near 55 kDa and the lower intensity of bands near 80 and 125 kDa).
On the basis of these experiments, a concentration of 500 µM
genistein was used for cells subjected to strain. It should be noted
that serum starvation dramatically reduced the levels of tyrosine
phosphorylation within these cells as well (first 2 lanes in Fig.
4A). SMCs were then plated on collagen-coated BioFlex plates
in the serum-containing M199. After 24 h, cells were then
serum-starved for an additional 24 h before being subjected to a
positive strain in the presence of 500 µM genistein. Cells were
pretreated with genistein for 15 min before being exposed to strain.
Triton X-100-insoluble (n = 6) and total cellular
lysates (n = 6) were extracted 15 min after the strain
was initiated because this was the point at which a maximal response
was seen in the preliminary experiments (see Fig. 1). MTs assembled
(Fig. 4B, left) from the pool of monomeric tubulin (Fig. 4B, right) in the presence of
genistein in the same fashion as control conditions.
|
Control of MT assembly via the ECM density.
Previous studies have indicated that changes in ECM density alter cell
shape via alteration of the cell-ECM force balance (36,
45). According to these studies, an increased ECM ligand density
would support more binding between cells and their substrate, alter
cell shape, and transfer inherent cytoskeletal prestress away from the
cytoskeleton to the ECM. To address this question, we investigated the
effect of ECM density on MT assembly. Cells were cultured on
bacteriologic culture dishes in a completely defined serum-free
environment. Fibronectin was preadsorbed onto the culture dishes at
theoretical densities of 50, 100, 500, and 1,000 ng/cm2
with a carbonate/bicarbonate buffer. After 24 h, Triton
X-100-insoluble (n = 6) and total cellular lysates
(n = 6) were extracted and quantified for tubulin by
using the competitive ELISA. SMCs cultured on 50 ng/cm2
fibronectin had 59 ± 11% of their total cellular tubulin in the form of MTs, while those cultured on 1,000 ng/cm2
fibronectin had 81 ± 13% of their total cellular tubulin in the form of MTs (P < 0.05) (Fig.
5A). Quantification of spread
cell area in these different conditions revealed that the increased MT
mass correlated with a slight but significant increase
(P < 0.05) in cell projected area across these
different ECM conditions (Fig. 5B). To confirm that
presenting increased density of ECM ligand leads to an increase in
cell-ECM bonds, we performed immunofluorescence localization of the
5-integrin receptor. A qualitative increase in the
number and organization of focal adhesions was found as ECM density was
increased (Fig. 5C). SMCs cultured on a range of fibronectin
densities were subsequently subjected to a 10% step increase in
strain. In all matrix conditions studied (from 100 to 5,000 ng/cm2), the mass of MTs increased over static controls,
although the extent of the net assembly of MTs was dependent on the ECM
ligand density (Fig. 6). For cells
cultured on 100 ng/cm2 fibronectin, MT mass increased by
5 ± 10% over static controls. For cells cultured on 500 ng/cm2 fibronectin, the net increase in MT mass was 27 ± 10%, significantly higher than the assembly on the 100 ng/cm2 condition (P < 0.05).
Strain-induced MT assembly occurred to the same extent in cells
cultured on higher densities of fibronectin (1,000 and 5,000 ng/cm2) as in cells cultured on 500 ng/cm2
(data not shown).
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We have shown that MT assembly can be controlled in SMCs in a
predictable fashion both by applying a step change in external strain
and by altering the density of ECM ligands. Our model for this
additional point of control for MT assembly is summarized in a
schematic representation in Fig. 7. In
this model, tensile forces leading to positive substrate strain promote
net MT assembly in SMCs cultured on flexible substrates, while
compressive forces imparting negative substrate strain drive net MT
disassembly. In parallel, the equilibrium set point between monomeric
and polymeric tubulin can be shifted simply by changing the amount of
available ligand used for cell adhesion.
|
In this study, MTs responded to mechanical strain in the same manner for SMCs cultured on both fibronectin and type I collagen, independent of tyrosine phosphorylation. These findings support the idea that this response is due to the direct mechanical coupling between the ECM and the cytoskeleton. Although we have not exhaustively tested matrix molecules or signaling inhibitors to determine whether the MT response to strain is universal, the finding that the response is the same on fibronectin and collagen is distinct from other mechanical signaling pathways that show a dependence on the identity of the ECM (12, 22, 38, 41, 43). Stretch-induced extracellular signal-regulated kinase activation has been shown to occur only on a fibronectin matrix, suggesting that the particular integrin used to transmit the mechanical signal is critical in determining the fate of the cell (41). Similarly, strengthening of cytoskeleton-integrin linkages in response to an applied force has been shown to be regulated by the tyrosine kinase Src in an ECM-dependent fashion (22). We do not believe that our results are in opposition to these earlier studies. Instead, in response to a mechanical signal, a cell presumably receives many parallel mechanical and chemical signals, all of which must be integrated and understood to generate the appropriate response. Hence, altering the mechanics of the cytoskeleton may not be sufficient alone to explain the ECM-dependent mechanotransduction responses reported in numerous other investigations. However, directly altering the mechanics of the MT network may be permissive for other downstream signaling events, acting in concert with, and not instead of, numerous soluble signaling pathways.
While often overlooked in the study of mechanotransduction, MTs have been studied extensively in the process of neurite extension, clearly contributing to their elongation (17, 18, 37, 55, 63). These findings provided much of the groundwork and initial support for a tensegrity-based model for the cytoskeleton. In a tensegrity model for the cell, the cytoskeleton exists in a dynamic balance of forces, with externally applied mechanical forces superimposed on this preexisting force balance (33-35). Reorganization of the cytoskeleton in response to mechanical forces has been widely reported, particularly for endothelial cells exposed to shear forces (11, 16, 25, 42, 57). Application of a restraining force via optical tweezers (12, 22) and twisting torques with a magnetic twisting device (59) both have been shown to result in a strengthening, or stiffening, of the cytoskeleton. Our findings in this present study support the possibility of MTs acting as compression-resistant struts. Presumably, applying a tensile strain to the substrate reduces the inherent compressive stress on the MTs, favoring a net increase in assembly. By contrast, applying a compressive strain to the substrate increases the compressive stress on the MTs, favoring a net disassembly of MTs.
Numerous investigations have studied MTs in the context of cell contractility (e.g., cell-generated forces) in place of the externally applied forces used in this study. These studies demonstrate that depolymerization of MTs with pharmacological agents (i.e., colchicine or nocodazole) induces formation of actin stress fibers and increased focal adhesions, the hallmarks of contractility (15, 20, 21, 40). The mechanism of increased contractility appears to involve greater myosin light chain phosphorylation, which results from drug-induced MT assembly (39). Other investigators have also demonstrated that changes in the contractility of the cytoskeleton are, in part, due to the small GTP-binding protein RhoA, a central signaling molecule that has been linked to many downstream responses (14, 56). Drug-induced depolymerization of MTs has been shown to trigger an increase in contractility and actin stress fiber formation (13, 20) via activation of Rho (21, 40). It is now believed that cross talk between Rho and Rac may influence the activity of myosin light chain by regulating the activity of myosin light chain phosphatase and kinase (3). All together, this study and past reports indicate MT assembly may be regulated by mechanical signals both directly and via chemical signaling pathways. Changes in MT assembly may influence a number of downstream signaling pathways that subsequently influence cell functions (27).
Our findings in this study disagree with those from a recent study by Heidemann et al. (29) in which local forces were applied to cells expressing green fluorescent protein-tagged MTs with the use of calibrated microneedles. We hypothesize that MTs in cells subjected to whole cell mechanical perturbations via integrin receptors (as in our study) will respond differently than will MTs in cells exposed to local mechanical manipulations. In our system, mechanical signals transduced via the ECM through integrin receptors act on an integrated cytoskeletal network, changing MT assembly. While we cannot rule out alternative explanations to describe how local perturbations in the mechanical microenvironment may affect the cytoskeleton, clearly MT assembly is influenced by mechanical forces in our system.
The pseudoequilibrium between tubulin monomers and MTs was altered in this study simply by changing the density of ECM ligand. Changes in the ECM density alone have been shown to trigger changes in cell shape for cultured hepatocytes, modulating the assembly of their MTs (44, 45). However, unlike results in those studies, our results demonstrate that changing fibronectin density causes only slight changes in cell shape for SMCs. We speculate that our finding of increased MT mass with increasing matrix density was not entirely due to the small increase in projected cell area but also to an increased number of cell adhesion sites (as shown in Fig. 5C). It has been hypothesized that increasing the number and organization of focal adhesion sites on higher ECM densities could potentially transfer inherent cytoskeletal prestress away from the MTs and onto the matrix itself, prompting increased MT assembly (34, 44, 45). Such control of MT assembly might be particularly relevant in cell types that do not exist in a dynamic mechanical environment in vivo, allowing changes in ECM composition to trigger changes in cytoskeletal architecture.
In summary, we have shown that MT assembly can be predictably controlled in SMCs by externally applied forces and changes to the ECM, suggesting that control of MT assembly is, in part, mechanical in nature. While the physiological relevance of mechanical control of MT assembly is unclear, changes in MT assembly may be critical to integrate the complex mechanical and chemical signaling events involved in a wide variety of cellular phenomena, including mechanotransduction and cell migration.
![]() |
ACKNOWLEDGEMENTS |
---|
We are especially grateful to Jim Cunningham and Janet Nikolovski for insightful discussions during the preparation of this manuscript.
![]() |
FOOTNOTES |
---|
Financial support from National Institutes of Health (NIH) Grant R01-DE-I3349 is gratefully acknowledged. A. J. Putnam was supported by fellowships from NIH Cellular Biotechnology Training Grant GM-08353 and NIH Organogenesis Training Grant 5T32-HD-07505-02 during the course of this work.
Address for reprint requests and other correspondence: D. J. Mooney, Dept. of Biologic and Materials Sciences, School of Dentistry, Univ. of Michigan, Ann Arbor, MI 48109-1078 (E-mail: mooneyd{at}umich.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.
Received 8 May 2000; accepted in final form 18 September 2000.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Adams, JC,
and
Watt FM.
Regulation of development and differentiation by the extracellular matrix.
Development
117:
1183-1198,
1993
2.
Boudreau, N,
and
Bissell MJ.
Extracellular matrix signaling: integration of form and function in normal and malignant cells.
Curr Opin Cell Biol
10:
640-646,
1998[ISI][Medline].
3.
Burridge, K.
Crosstalk between Rac and Rho.
Science
283:
2028-2029,
1999
4.
Butt, RP,
Laurent GJ,
and
Bishop JE.
Mechanical load and polypeptide growth factors stimulate cardiac fibroblast activity.
Ann NY Acad Sci
752:
387-393,
1995[ISI][Medline].
5.
Buxbaum, RE,
and
Heidemann SR.
A thermodynamic model for force integration and microtubule assembly during axonal elongation.
J Theor Biol
134:
379-390,
1988[ISI][Medline].
6.
Cai, S,
Pestic-Dragovich L,
O'Donnell ME,
Wang N,
Ingber D,
Elson E,
and
De Lanerolle P.
Regulation of cytoskeletal mechanics and cell growth by myosin light chain phosphorylation.
Am J Physiol Cell Physiol
275:
C1349-C1356,
1998
7.
Caron, JM,
Jones AL,
and
Kirschner MW.
Autoregulation of tubulin synthesis in hepatocytes and fibroblasts.
J Cell Biol
101:
1763-1772,
1985[Abstract].
8.
Carver, W,
Nagpal ML,
Nachtigal M,
Borg TK,
and
Terracio L.
Collagen expression in mechanically stimulated cardiac fibroblasts.
Circ Res
69:
116-122,
1991[Abstract].
9.
Chen, CS,
Mrksich M,
Huang S,
Whitesides GM,
and
Ingber DE.
Geometric control of cell life and death.
Science
276:
1425-1428,
1997
10.
Chicurel, ME,
Chen CS,
and
Ingber DE.
Cellular control lies in the balance of forces.
Curr Opin Cell Biol
10:
232-239,
1998[ISI][Medline].
11.
Chien, S,
Li S,
and
Shyy YJ.
Effects of mechanical forces on signal transduction and gene expression in endothelial cells.
Hypertension
31:
162-169,
1998
12.
Choquet, D,
Felsenfeld DP,
and
Sheetz MP.
Extracellular matrix rigidity causes strengthening of integrin-cytoskeleton linkages.
Cell
88:
39-48,
1997[ISI][Medline].
13.
Chrzanowska-Wodnicka, M,
and
Burridge K.
Rho-stimulated contractility drives the formation of stress fibers and focal adhesions.
J Cell Biol
133:
1403-1415,
1996[Abstract].
14.
Clark, EA,
King WG,
Brugge JS,
Symons M,
and
Hynes RO.
Integrin-mediated signals regulated by members of the rho family of GTPases.
J Cell Biol
142:
573-586,
1998
15.
Danowski, BA.
Fibroblast contractility and actin organization are stimulated by microtubule inhibitors.
J Cell Sci
93:
255-266,
1989[Abstract].
16.
Davies, PF,
and
Tripathi SC.
Mechanical stress mechanisms and the cell. An endothelial paradigm.
Circ Res
72:
239-245,
1993[Abstract].
17.
Dennerll, TJ,
Joshi HC,
Steel VL,
Buxbaum RE,
and
Heidemann SR.
Tension and compression in the cytoskeleton of PC-12 neurites. II. Quantitative measurements.
J Cell Biol
107:
665-674,
1988[Abstract].
18.
Drubin, DG,
Feinstein SC,
Shooter EM,
and
Kirschner MW.
Nerve growth factor-induced neurite outgrowth in PC12 cells involves the coordinate induction of microtubule assembly and assembly-promoting factors.
J Cell Biol
101:
1799-1807,
1985[Abstract].
19.
Duncan, RL,
and
Turner CH.
Mechanotransduction and the functional response of bone to mechanical strain.
Calcif Tissue Int
57:
344-358,
1995[ISI][Medline].
20.
Elbaum, M,
Chausovsky A,
Levy ET,
Shtutman M,
and
Bershadsky AD.
Microtubule involvement in regulating cell contractility and adhesion-dependent signalling: a possible mechanism for polarization of cell motility.
Biochem Soc Symp
65:
147-172,
1999[Medline].
21.
Enomoto, T.
Microtubule disruption induces the formation of actin stress fibers and focal adhesions in cultured cells: possible involvement of the rho signal cascade.
Cell Struct Funct
21:
317-326,
1996[ISI][Medline].
22.
Felsenfeld, DP,
Schwartzberg PL,
Venegas A,
Tse R,
and
Sheetz MP.
Selective regulation of integrincytoskeleton interactions by the tyrosine kinase Src.
Nat Cell Biol
1:
200-206,
1999[ISI][Medline].
23.
Folkman, J,
and
Moscona A.
Role of cell shape in growth control.
Nature
273:
345-349,
1978[ISI][Medline].
24.
Fuller, B.
Tensegrity.
Portfolio Artnews Annu
4:
112-127,
1961.
25.
Galbraith, CG,
Skalak R,
and
Chien S.
Shear stress induces spatial reorganization of the endothelial cell cytoskeleton.
Cell Motil Cytoskeleton
40:
317-330,
1998[ISI][Medline].
26.
Giancotti, FG,
and
Ruoslahti E.
Integrin signaling.
Science
285:
1028-1032,
1999
27.
Gundersen, GG,
and
Cook TA.
Microtubules and signal transduction.
Curr Opin Cell Biol
11:
81-94,
1999[ISI][Medline].
28.
Harris, A.
Behavior of cultured cells on substrata of variable adhesiveness.
Exp Cell Res
77:
285-297,
1973[ISI][Medline].
29.
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
30.
Hill, TL.
Microfilament or microtubule assembly or disassembly against a force.
Proc Natl Acad Sci USA
78:
5613-5617,
1981[Abstract].
31.
Howe, A,
Aplin AE,
Alahari SK,
and
Juliano RL.
Integrin signaling and cell growth control.
Curr Opin Cell Biol
10:
220-231,
1998[ISI][Medline].
32.
Huang, S,
and
Ingber DE.
The structural and mechanical complexity of cell-growth control.
Nat Cell Biol
1:
E131-E138,
1999[ISI][Medline].
33.
Ingber, DE.
Cellular tensegrity: defining new rules of biological design that govern the cytoskeleton.
J Cell Sci
104:
613-627,
1993
34.
Ingber, DE.
Tensegrity: the architectural basis of cellular mechanotransduction.
Annu Rev Physiol
59:
575-599,
1997[ISI][Medline].
35.
Ingber, DE,
Dike L,
Hansen L,
Karp S,
Liley H,
Maniotis A,
McNamee H,
Mooney D,
Plopper G,
Sims J,
and
Wang N.
Cellular tensegrity: exploring how mechanical changes in the cytoskeleton regulate cell growth, migration, and tissue pattern during morphogenesis.
Int Rev Cytol
150:
173-224,
1994[ISI][Medline].
36.
Ingber, DE,
and
Folkman J.
Mechanochemical switching between growth and differentiation during fibroblast growth factor-stimulated angiogenesis in vitro: role of extracellular matrix.
J Cell Biol
109:
317-330,
1989[Abstract].
37.
Joshi, HC,
Chu D,
Buxbaum RE,
and
Heidemann SR.
Tension and compression in the cytoskeleton of PC 12 neurites.
J Cell Biol
101:
697-705,
1985[Abstract].
38.
Kim, BS,
Nikolovski J,
Bonadio J,
and
Mooney DJ.
Cyclic mechanical strain regulates the development of engineered smooth muscle tissue.
Nat Biotechnol
17:
979-983,
1999[ISI][Medline].
39.
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].
40.
Liu, BP,
Chrzanowska-Wodnicka M,
and
Burridge K.
Microtubule depolymerization induces stress fibers, focal adhesions, and DNA synthesis via the GTP-binding protein Rho.
Cell Adhes Commun
5:
249-255,
1998[ISI][Medline].
41.
MacKenna, DA,
Dolfi F,
Vuori K,
and
Ruoslahti E.
Extracellular signal-regulated kinase and c-Jun NH2-terminal kinase activation by mechanical stretch is integrin-dependent and matrix-specific in rat cardiac fibroblasts.
J Clin Invest
101:
301-310,
1998
42.
Malek, AM,
and
Izumo S.
Mechanism of endothelial cell shape change and cytoskeletal remodeling in response to fluid shear stress.
J Cell Sci
109:
713-726,
1996
43.
Meazzini, MC,
Toma CD,
Schaffer JL,
Gray ML,
and
Gerstenfeld LC.
Osteoblast cytoskeletal modulation in response to mechanical strain in vitro.
J Orthop Res
16:
170-180,
1998[ISI][Medline].
44.
Mooney, DJ,
Hansen LK,
Langer R,
Vacanti JP,
and
Ingber DE.
Extracellular matrix controls tubulin monomer levels in hepatocytes by regulating protein turnover.
Mol Biol Cell
5:
1281-1288,
1994[Abstract].
45.
Mooney, DJ,
Langer R,
and
Ingber DE.
Cytoskeletal filament assembly and the control of cell spreading and function by extracellular matrix.
J Cell Sci
108:
2311-2320,
1995
46.
Osol, G.
Mechanotransduction by vascular smooth muscle.
J Vasc Res
32:
275-292,
1995[ISI][Medline].
47.
Pelham, RJ, Jr,
and
Wang Y.
Cell locomotion and focal adhesions are regulated by substrate flexibility.
Proc Natl Acad Sci USA
94:
13661-13665,
1997
48.
Pugh, A.
Introduction to Tensegrity. Berkeley, CA: Univ. of California Press, 1976.
49.
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
50.
Rothman, A,
Kulik TJ,
Taubman MB,
Berk BC,
Smith CW,
and
Nadal-Ginard B.
Development and characterization of a cloned rat pulmonary arterial smooth muscle cell line that maintains differentiated properties through multiple subcultures.
Circulation
86:
1977-1986,
1992[Abstract].
51.
Schmidt, C,
Pommerenke H,
Durr F,
Nebe B,
and
Rychly J.
Mechanical stressing of integrin receptors induces enhanced tyrosine phosphorylation of cytoskeletally anchored proteins.
J Biol Chem
273:
5081-5085,
1998
52.
Schwartz, MA,
and
Baron V.
Interactions between mitogenic stimuli, or, a thousand and one connections.
Curr Opin Cell Biol
11:
197-202,
1999[ISI][Medline].
53.
Shyy, JY,
and
Chien S.
Role of integrins in cellular responses to mechanical stress and adhesion.
Curr Opin Cell Biol
9:
707-713,
1997[ISI][Medline].
54.
Simpson, DG,
Carver W,
Borg TK,
and
Terracio L.
Role of mechanical stimulation in the establishment and maintenance of muscle cell differentiation.
Int Rev Cytol
150:
69-94,
1994[ISI][Medline].
55.
Tanaka, E,
Ho T,
and
Kirschner MW.
The role of microtubule dynamics in growth cone motility and axonal growth.
J Cell Biol
128:
139-155,
1995[Abstract].
56.
Tapon, N,
and
Hall A.
Rho, Rac and Cdc42 GTPases regulate the organization of the actin cytoskeleton.
Curr Opin Cell Biol
9:
86-92,
1997[ISI][Medline].
57.
Thoumine, O,
Ziegler T,
Girard PR,
and
Nerem RM.
Elongation of confluent endothelial cells in culture: the importance of fields of force in the associated alterations of their cytoskeletal structure.
Exp Cell Res
219:
427-441,
1995[ISI][Medline].
58.
Thrower, D,
Jordan MA,
and
Wilson L.
A quantitative solid-phase binding assay for tubulin.
Methods Cell Biol
37:
129-145,
1993[ISI][Medline].
59.
Wang, N,
Butler JP,
and
Ingber DE.
Mechanotransduction across the cell surface and through the cytoskeleton.
Science
260:
1124-1127,
1993[ISI][Medline].
60.
Williams, B.
Mechanical influences on vascular smooth muscle cell function.
J Hypertens
16:
1921-1929,
1998[ISI][Medline].
61.
Wilson, E,
Mai Q,
Sudhir K,
Weiss RH,
and
Ives HE.
Mechanical strain induces growth of vascular smooth muscle cells via autocrine action of PDGF.
J Cell Biol
123:
741-747,
1993[Abstract].
62.
Wilson, E,
Sudhir K,
and
Ives HE.
Mechanical strain of rat vascular smooth muscle cells is sensed by specific extracellular matrix/integrin interactions.
J Clin Invest
96:
2364-2372,
1995[ISI][Medline].
63.
Zheng, J,
Buxbaum RE,
and
Heidemann SR.
Investigation of microtubule assembly and organization accompanying tension-induced neurite initiation.
J Cell Sci
104:
1239-1250,
1993