measuring patterns, regulation, and biologic consequences...

Gravitational and Space Biology 20(2) June 2007 19

MEASURING PATTERNS, REGULATION, AND BIOLOGIC CONSEQUENCES OF CELLULAR

TRACTION FORCES

Christopher A. Lemmon1,2

and Lewis H. Romer2,3

1Department of Biomedical Engineering,

2Department of Anesthesiology and Critical Care Medicine,

3Departments of Cell Biology and Pediatrics, Johns Hopkins University School of Medicine, Baltimore, MD

21287-4904

ABSTRACT

The exchange of mechanical signals between mammalian cells

and their extracellular matrix microenvironments is a focus of

keen interest for biologists in the diverse fields of development,

vascular disease, tissue engineering, and oncology. The

molecular machinery of cellular mechanical signal response

includes at least three major components: transmembrane

adhesion receptors for extracellular matrix; the microfilament

and microtubule cytoskeletal systems; and the Rho family of

GTP-binding proteins that modulate the density and contraction

state of cytoskeletal elements. Complex signaling pathways link

these three arms of mechanical signal response as they interact

with the extracellular matrix microenvironment.

Arrays of microfabricated silicon posts have provided a system

for reporting the traction force at individual cell-matrix

interaction sites. Volumetric imaging of posts with known

spring constants allows the efficient analysis of maps of cell

traction force vectors and their coincidence with both actin

polymerization and signal molecule localization. The effects of

cytoskeletal contraction on patterns of cell traction forces are

reported here. We have also defined unique patterns of force

generation that are specific for cells of fibroblast, endothelial,

epithelial, and smooth muscle lineages. Cell-generated

mechanical forces directly affect patterning of the matrix milieu

and the mechanical cues that are stored in it. The contributions

of signaling molecules including non-receptor tyrosine kinases

and GTPases to the exchange of mechanical data between the

cell and its environment are discussed. These investigations

have important implications for understanding and optimizing

cellular growth behaviors during tissue development and repair

in normal and microgravity settings.

INTRODUCTION

Restrictions in cell size, shape, and volume are dictated

by the spatial array and stiffness of the immediate

microenvironment. These material properties thereby

become mechanical input signals. The far-reaching

consequences of these signals include the most basic

determinations in the life of the cell and its organism.

Protein synthesis may be completely suppressed in mouse

fibroblasts by preventing cell adhesion (Ben-Ze'ev et al.,

1980), and apoptosis is induced in endothelial cells whose

cell attachment area is restricted to less than that of a 25

µm microbead (Chen et al., 1997). Human mesenchymal

stem cell lineage commitment was limited to adipocyte

differentiation by spatial restriction to surface areas of 1-2

x 103 µm

2 and was consistently altered to an osteogenesis

program by the availability of a more generous

attachment surface area of 1 x 104 µm

2 (McBeath et al.,

2004). More complex developmental schemata apparently

follow similar imperatives. Thus, Xenopus notochord

development is accomplished through patterns of cellular

growth that optimize the balance between flexural

stiffness and isometric force (Keller, 2006), and precise

focusing of compressive and tensile forces may activate

transcription factors, thereby marking localization signals

for cartilage and bone formation during human

craniofacial morphogenesis (Radlanski and Renz, 2006).

A number of recent studies have highlighted the

significance of mechanical signaling in physiologic and

pathologic events. Paszek et al. have demonstrated that

cancerous tumors have a stiffer stroma and elevated Rho-

dependent contractility (Paszek et al., 2005). They

demonstrated that increasing substrate stiffness led to

increased ERK activity, increased Rho activity, and

increase focal adhesion formation, all of which

contributed to a malignant phenotype. In addition,

transformed cells reverted to a non-transformed

phenotype upon Rho-Kinase or ERK inhibition. Baneyx

et al. investigated the effects of traction forces on the

matrix protein fibronectin, and have shown in vitro that

cell traction forces are capable of stretching this protein

(Baneyx et al., 2002). They have constructed a model in

which excessive stretching of fibronectin by cell traction

forces alters integrin-dependent angiogenic signals which

originate from integrin-fibronectin ligation (Vogel and

Baneyx, 2003).

Altered gravity (either microgravity or hypergravity) can

potentially affect cell traction forces and mechanical

signaling in different ways. First, it has been

demonstrated that focal adhesions respond to applied

forces, be it either internally applied via the cytoskeleton

or externally applied by an outside force (Riveline et al.,

2001). The combined mass of the cell and the attached

extracellular matrix will exert a force at these adhesion

sites that is dependent on the degree of gravitational pull

and the orientation of the tissue. Second, it has been

shown that the cytoskeletal architecture, which transmits

cellular force to focal adhesions, is altered in response to

gravity. Studies in primary osteoblasts have shown that

microtubule network height is decreased in a dose-

dependent manner to hyper-gravity (Searby et al., 2005).

Rosner et al. showed that neuroblastoma cells had

increased protrusive lamellar activity when subjected to

____________________

* Correspondence to: Lewis H. Romer

904 Blalock/Pediatric Anesthesiology and Critical Care

Medicine

The Johns Hopkins University School of Medicine

600 North Wolfe Street

Baltimore, MD 21287-4904 USA

Email: [email protected]

Phone: 410-955-7610; Fax: 410-502-5312

C.A. Lemmon and L.H. Romer - Biologic Consequences of Cellular Traction Forces

20 Gravitational and Space Biology 20(2) June 2007

Figure 1 - Reciprocal exchange of mechanical data. A: Mouse embryo fibroblasts were grown on the vulcanized

surfaces of phenylmethyl-siloxane puddles that were prepared in

Rappaport chambers as described in the Methods. Phase

contrast microscopy at three hours revealed wrinkles in this thin

silicon membrane.

B: Mouse embryo fibroblasts were cultivated on fibronectin (25

µg/ml) adsorbed to glass coverslips or poly-dimethylsiloxane

with a base to curing agent ratio of 30 to 1 (“30:1 PDMS”), and

processed for epifluorescence imaging of actin and vinculin.

Surfaces were UV-oxidized prior to plating cells. Merged

images show actin in red and vinculin in green. The stiffer glass

surface enhances actin organization (see detail in the insets in

the top two panels) and matrix adhesion formation. Scale bars

= 20 µm.

hypergravity (2 g) (Rosner et al., 2006). Finally, studies

have shown that gene expression is altered in response to

altered gravitational pull. Shimada et al. showed that

expression of heat shock protein 70 was increased under

both microgravity and hypergravity (Shimada and

Moorman, 2006). Thus it is possible that downstream

signaling pathway activity may be altered by specific

changes in protein expression.

Despite its profound effects, mechanical data exchange

across the cell membrane is readily accessible with

straightforward experiments. Mechanical force is

transmitted and sensed at cell-matrix attachment sites.

Cytoskeletal traction forces that are transmitted to the cell

surroundings can be visualized as wrinkle patterns when

cells are plated onto a deformable silicon sheet (Figure 1

A) (see also (Harris et al., 1980)). The size, number, and

geometric complexity of these wrinkles is directed in part

by the force of cytoskeletal contraction, and this

relationship may change with adhesion maturation and

from cell type to cell type (Beningo et al., 2001;

Pletjushkina et al., 2001). Cells can also sense and

respond to mechanical signals. Organization of the F-actin

cytoskeleton and formation of focal adhesions in

mammalian fibroblasts are significantly altered depending

on the stiffness of the underlying substrate. Cells growing

on a stiff glass surface, which has a Young’s Modulus of

approximately 70 GPa, have well-developed vinculin-

containing focal adhesions after only one hour of

spreading, particularly at the cell periphery, and densely

packed peripheral arrays of F-actin filaments (Figure 1

B). In contrast, cells adherent for one hour to soft

polydimethyl-siloxane with a base to curing agent ratio of

30 to 1 (30:1 PDMS) and a Young’s Modulus of

approximately 1 MPa (Carrillo et al., 2005), lack well-

developed focal adhesions. F-actin fibers in the cells

growing on the softer substrate are thinner and more

uniformly distributed across the cell. The distinct patterns

of F-actin organization that are manifested in these two

very different mechanical environments are shown in

detail in the insets at the upper right corners of the two top

panels in Figure 1 B.

Quantification of forces on deformable substrates has

been achieved by embedding fluorescent beads within the

substrate and tracking bead location before and after

contraction (Dembo and Wang, 1999). The displacement

of each bead within the substrate was used to determine

the traction forces applied to the substrate. This system

was used to demonstrate that the magnitude and direction

of force applied to an individual adhesion correlated with

the direction and size of the adhesion and estimated the

force applied to an individual adhesion to be

approximately 5 nN/ µm2 (Balaban et al., 2001). The

continuous nature of the substrate in this system makes

force quantification computationally complex; the

calculation of forces from the displacement matrix cannot

be solved without making certain assumptions.



Figure 2 - Measuring cell traction force. A and B: A mouse embryo fibroblast is shown on an mPAD array. Actin (labeled with phalloidin) appears red, antibody-labeled

fibronectin is blue, and fluorophore-conjugated BSA is green. 3D reconstructions of z-stacks of images are shown following iterative

deconvolution in top (A) and bottom (B) views. mPAD posts are 10 µm high and 3µm in diameter with a post-to-post spacing of 9µm.

C: Surrounding each post with fluorophore signal allows independent imaging of both the anchored (red) and free (blue) ends of each

cantilever post. Superposition of these images provides a measurement of centroid displacement (white arrows).

Calculation of forces can be more directly measured using

the microfabricated post-array detector (mPAD) system:

this assay consists of a substrate of uniformly-spaced,

deformable cantilevers, which deflect when cell traction

forces are applied (Tan et al., 2003; Lemmon et al., 2005;

Li et al., 2007). Cell traction force applied to an

individual mPAD post is linearly proportional to the

deflection and stiffness of that cantilever post. The design

and surface chemistry of the mPAD limits cell attachment

to distinct and discrete locations, which simplifies the

analysis of traction forces. Thus, each cantilever acts as

an individual mechanosensor for cell traction forces.

Fluorescent-labeling of the mPAD post with fluorophore-

conjugated BSA facilitates volumetric imaging of the

entire post surface (Figure 2 A and B, green).

Microcontact-printing of fibronectin onto the mPAD free

ends (Figure 2 A, blue) limits cell attachment to the top

surface. Deflections are calculated by analysis of the BSA

fluorescence images at the anchored and free ends of the

mPAD posts (Figure 2 C). Superposition of the free end

(blue) and anchored end (red) images provides a

measurement of the centroid displacement (white arrows).

A key element in the generation of cellular traction force

is the activation of actomyosin contraction by myosin

light chain phosphorylation (Tan et al., 2003) (also

reviewed in (Romer et al., 2006)). This process is

regulated by the GTP-binding protein Rho, and its target

effector proteins Rho kinase (ROCK) and mDia-1 (Ridley

and Hall, 1992; Nakano et al., 1999; Watanabe et al.,

1999; Wozniak et al., 2003). ROCK increases the

strength of actomyosin contraction by phosphorylating the

myosin light chain, and maintains this increased

contractility by inhibiting myosin light chain phosphatase.

mDia1 then supports the incorporation of contractile

filaments of F-actin and type II myosin into stress fibers –

the organelles that exert traction force on the cytoplasmic

face of cell-matrix adhesions. It is intriguing to note that

complex cellular responses to mechanical stimuli,

including the molecular rearrangements of matrix

adhesions upon exposure to external centripetal pulling

force and the lineage commitment of mesenchymal stem

cells based upon cell adhesion surface area, are dependent

upon mechanical and chemical signaling through mDia-1

and ROCK, respectively (Riveline et al., 2001; McBeath

et al., 2004). Molecules that regulate the activation state

of Rho family GTP-binding proteins provide an additional

level of modulation of cytoskeletal organization,

contraction, and contribution to cell traction forces. These

are the guanine nucleotide exchange factors (GEFs) that

activate Rho by facilitating the exchange of GDP for

GTP, and GTPase activating proteins (GAPs) that

inactivate Rho family members by accelerating their

otherwise very slow GTPase activity. Both GEFs and

GAPs are responsive to mechanical cues in the

extracellular environment (Arthur et al., 2000; Arthur and

Burridge, 2001; Russell et al., 2002; Hornstein et al.,

2004; Tcherkezian and Lamarche-Vane, 2007).

In this communication a technique for quantitative study

of cell traction forces at discrete cell-matrix interaction

sites is described. Data on changing patterns of cell

traction force due to cytoskeletal contraction and

biological diversity are reported. The molecular basis of

mechanical signal response at cell-matrix adhesion sites is

then reviewed, and we propose a biophysical model for

cellular mechanosensing. It is hoped that this

presentation will further new interest and cross-

disciplinary collaborations in the investigation of

mechanical signaling.

METHODS

Cell Culture

Mouse embryo fibroblasts (MEF) and Swiss 3T3 cells

(both obtained from ATCC, Rockville, MD) were

cultivated in DMEM with 10% fetal bovine serum.

Human umbilical vein endothelial cells (HUVEC; VEC

Technologies, Rochester, NY) were cultivated in standard

medium from the same source. MCF10a cells (ATCC)

were cultivated in DMEM with 5% horse serum as

described previously (Lemmon et al., 2005). Bovine



aortic smooth muscle cells (SMC; gift from Donald

Ingber, Harvard) were cultivated in DMEM with 10% calf

serum.

Extracellular Deformation by Cytoskeletal Tension

To evaluate the deformation of extracellular matrix by

cytoskeletal contraction (Figure 1), Rappaport chambers

designed for the stage of an inverted microscope were

assembled with modifications of published specifications

(Burton et al., 1999). A glass ring 1 cm in height and 2.2

cm in outer diameter was attached to a number 0 cover

glass (22 x 22 mm, Thomas) with paraffin, petroleum

jelly, and Lanolin (1:1:1). The glass ring was heated to

60-70o C, and then used to melt the mountant and make a

seal on the cover glass. Phenylmethyl siloxane (#710,

viscosity = 500 centistokes; Dow Corning, Midland, MI)

was spread evenly on the bottom of the chamber and

crosslinked in a standard sputter coater (Polaron E5100)

with a target to specimen distance of 60 mm. The pressure

was reduced to 150 millitorr, the air was replaced with

argon, and the top layer of the siloxane was vulcanized

using gold palladium and a current of 20 mA for 16 sec.

The silicon rubber was exposed to UV light for 4 min to

uncouple the cross-linking of phenyl side chains and

increase the deformability of the silicon rubber surface. 3

hours after plating on this surface, cells were viewed with

a 60x phase contrast objective using the Nikon Eclipse

TE200 microscope and the image capture apparatus

detailed above.

Microfabricated Post Array Detector (mPAD)

Fabrication and Preparation mPADs were fabricated using previously described

techniques (Tan et al., 2003). Briefly, an array of 3-

micron diameter, 10-micron tall posts with a 9-micron by

9-micron grid spacing was fabricated by spin-coating a

positive resist (SU-8) onto a silicon wafer and exposing

the wafer to UV light to polymerize the SU-8. Poly-

dimethylsiloxane (PDMS) (Dow Corning, Midland, MI)

was then poured over the silicon wafer and cured

overnight at 100o C. The silicon wafer was peeled away

from the PDMS, leaving a negative image of the post

array. The negative mold was then treated for 1 minute in

oxygen plasma and coated with [tridecafluoro-1,1,2,2,-

tetrahydrooctyl]-1-trichlorosilane vapor (United Chemical

Technologies, Bristol, PA) under vacuum overnight to aid

in subsequent removal of mPAD substrates. PDMS was

subsequently poured into the negative mold, cured for 20

h at 100o C, and peeled away from the negative mold,

generating a positive feature mPAD substrate.

Prior to cell plating, mPADs were microcontact-printed

and prepared as previously described (Tan et al., 2003;

Lemmon et al., 2005). Briefly, PDMS stamps with a base

to curing agent ratio of 30 to 1 were coated with 50 µg/ml

human plasma fibronectin in PBS (Gibco) and incubated

for 1 h at room temperature. Stamps were then rinsed,

dried, and brought into contact with mPAD substrates that

had been UV-oxidized for 7 min. Stamps were peeled

away from the mPAD, leaving fibronectin on the top

surface of each mPAD post. mPADs were subsequently

incubated in 2 µg/ml cy5-conjugated BSA at room

temperature for 1 h to facilitate mPAD post volumetric

imaging, rinsed in de-ionized water, and incubated in

0.2% Pluronics F-127 to prevent cell attachment to

surfaces other than the top surface of the mPAD posts.

Immunofluorescence labeling, imaging, and analysis

Cells plated onto mPADs were fixed and permeabilized

with 3% paraformaldehyde and 0.5 % Triton X-100 in

PBS. Fixed samples were labeled for

immunofluorescence imaging by incubation with

fluorophore-conjugated phalloidin and polyclonal antisera

against fibronectin (Abcam, Cambridge, MA) for 30 min

at 37 o

C, and then incubated in fluorophore-conjugated,

isotype-specific and affinity cross-adsorbed anti-IgG

antibodies (Chemicon, Temecula, CA) for 30 min at 37o

C. Vinculin was labeled in some cells using a monoclonal

antibody (gift of Alexey Belkin, University of Maryland).

Cells were viewed with a Nikon Eclipse TE200

microscope and an internally cooled 12-bit CCD camera

(CoolSnapHQ, Photometrics, Tucson, AZ). Images were

collected using OpenLab software (Improvision,

Lexington, MA)

Acquired images were exported as 16-bit TIFF images

and read into an original Matlab code designed to analyze

mPAD post deflections, as previously described

(Lemmon et al., 2005). Briefly, acquired images were

imported, and a thresholding algorithm was used to

determine cell area, detect cell edges, and define mPAD

post centroids. Deflections were then calculated and a

vector map of the resulting cell-generated forces was

generated. This original Matlab code is available for

downloading free of charge at:

www.hopkinsmedicine.org/anesthesiology/research/mpad

tools.

Computational Analysis of Cell Strain

Cell strain was calculated using finite element analysis. A

grid of rectangular elements was created using the cell

outline, as determined from immunofluorescence of the F-

actin cytoskeleton. The shape and position of each

element in the model was determined from the measured

mPAD deflection maps: the strained position and shape of

each element was obtained from the deflected mPAD post

centroid grid, while the unstrained position and shape of

each element was derived from the undeflected mPAD

post centroid grid. Normal strains and shearing strain

were calculated for each element, and maximum strain

was calculated using Mohr’s Circle equation for 2-

dimensional plane strain:

where γmax is the maximum strain, εx and εy are the normal

strains along each axis, and γxy is the shear strain. Element

size was approximately 0.25 µm2.



Figure 3 - Cytoskeletal contraction changes the magnitude and pattern of cell traction forces. The addition of serum for one hour (data on left) to Swiss 3T3 cells that had been serum-starved for 22 hours induced rapid actin

polymerization and actomyosin contraction as compared to controls without serum (data on right). Epifluorescence imaging of F-actin (A),

mPAD analysis of cell traction force vectors (B), and strain maps (C) were analyzed; representative data are shown for each group. The

blue arrows denote a 10 nN vector for each traction force map shown. Axis labels in C are in pixels, and maximal strain is plotted from -

0.1 (tensile strain, blue) to 0.1 (compressive strain, red).

D: Total exterior force (defined as mPAD posts which are contacted by the cell outline) is summed for both the serum and no serum cases.

The ratio of the average exterior force per mPAD post to the average interior force per mPAD post are shown for each condition (right).



RESULTS

Force Distribution Patterns within Swiss 3T3 Fibroblast

Cells

Serum-starvation of cultured cells leads to decreased actin

stress fiber formation and focal adhesion disassembly, and

re-exposure of these cells to serum reverses these effects

in a Rho-dependent fashion (Giuliano and Taylor, 1990;

Ridley and Hall, 1992; Chrzanowska-Wodnicka and

Burridge, 1994). We wished to study the alterations in

patterns of cell traction forces caused by this serum-

dependent cytoskeletal remodeling. Swiss 3T3 fibroblasts

were plated onto mPADs in the presence of serum for 2h

to allow sufficient spreading and attachment to the

mPADs. Cells were then rinsed twice with serum-free

DMEM, and incubated for an additional 22h in serum-

free conditions. Cells were then either stimulated by

addition of serum for 1h, or left in serum-free conditions

for 1h. As shown in Figure 3A, serum-starved cells

showed a predicted decrease in F-actin fibers while cells

stimulated with serum developed dense F-actin fibers that

crossed the cell. Additionally, serum-activated cells

exhibited increased traction forces at the cell periphery

compared to the control group (Figure 3B). Serum-starved

cells also spread better than serum-activated cells

(Giuliano and Taylor, 1990; Ridley and Hall, 1992;

Chrzanowska-Wodnicka and Burridge, 1994). It is

possible that the decreased traction forces at the periphery

allow for more efficient spreading in these cells.

Finite element modeling was used to predict areas of high

and low strain within the cell. Two grids of rectangular

elements were generated based on mPAD deflection data:

a grid of elements was generated based on the mPAD post

base image, which represented the undeflected position of

mPAD posts, and thus the relaxed element state of the

model. A second grid of elements was generated based on

the mPAD post free-end image, which represented the

deflected position of mPAD posts, and thus the contracted

element state of the model. The x-, y-, and maximum

strains were then computed based on the shape change of

each element from the relaxed to the contracted state.

Maximum strains were plotted for serum-activated cells

as well as control, unstimulated cells (Figure 3C).

Maximal strain was plotted from -0.1 (tensile strain, blue)

to 0.1 (compressive strain, red). Strain maps indicated that

serum-activated cells had high levels of peripherally

localized tensile strain, with reduced strain within the

interior of the cell. In contrast, unstimulated cells showed

a much more uniform distribution of moderate

compressive and tensile strain throughout the cell. Bar

graphs in Figure 3D demonstrate the spatial segregation

of cell traction forces seen in the serum-stimulated Swiss

3T3 fibroblasts. Traction forces were separated into two

categories: “exterior” forces, which are applied by the cell

at the cell periphery; and “interior” forces, which are

applied by the cell beneath the perinuclear regions of the

cell. The average exterior force per post in serum-

stimulated cells was 10.44 +/- 0.76 nN (385 posts in 13

cells), while unstimulated cells had a significantly lower

average exterior force per post of 7.99 +/-0.45 nN (296

posts in 10 cells) (p= 0.037). This spatial segregation of

forces can also be seen in the ratio of average exterior

force per mPAD post to average interior force per mPAD

post. Serum-stimulated cells had a ratio of 1.74+/- 0.14

(13 cells), and unstimulated cells had a ratio of 1.10 +/-

0.07 (10 cells) (p = 0.005). Thus, the ratio of exterior to

interior traction force was nearly doubled by serum

exposure.

Figure 4 - Biological diversity in traction forces. A: Representative traction force vector maps are shown for

mouse embryo fibroblasts (MEF), human umbilical vein

endothelial cells (HUVEC), smooth muscle cells (SMC), and

mammary epithelial cells (MCF10a). The blue arrows denote a

10 nN vector for each map shown.

B: The average force per mPAD post and the total cell area

were calculated for each of the 4 cell types. Data indicate that

smooth muscle cells (green) generate a significantly higher

normalized force than other cell types, while HUVEC (blue)

generate a significantly lower normalized force. MCF10a cells

(black) spread significantly less than the other cell types, and

MEF (red) are intermediate for both metrics. Means and

standard deviations for both measurements are shown for each

cell type.

Biological Diversity in Cell Traction Force

Four different cell types were plated onto microfabricated

post array detectors (mPADs) to quantify differences in

traction force and cell spreading between cells of different

lineages as shown in Figure 4. MEF, HUVEC, SMC, and



MCF10a were plated onto mPADs for 24h in serum-

containing media under standard culture conditions.

Results indicated that SMC cells generated the strongest

traction forces, HUVEC generated the weakest traction

forces, and MEF and MCF10a cells generated similar

moderate traction forces. Cell size was also quantified for

each cell type (Figure 4B). Results indicated that HUVEC

spread more efficiently than the other cell types, while

MCF10a spread poorly. Interestingly, an inverse

relationship seems to exist between the normalized

traction force per post and cell size (Figure 4B). Cells

with lower normalized traction forces, such as the

HUVEC, have increased cell size, while cells with higher

normalized forces, such as the SMC, are smaller in size.

The average force per post for the SMC was higher than

for any of the other cell types studied, including the

smaller MCF10a cells.

DISCUSSION

Cells generate, sense, and respond to mechanical signals

(also reviewed in Bershadsky, 2004; Bershadsky et al.,

2006; Giannone and Sheetz, 2006; Romer et al., 2006;

Vogel and Sheetz, 2006). Our data elucidate a new

perspective on cellular mechanical responses – mapping

cell traction forces to discrete points of cell-matrix

contact. Focal adhesions are formed at points of cell

attachment and act as a clutch between the actin

cytoskeleton and the surrounding extracellular matrix;

they are thought to act not only as the point of

transmission of cytoskeletal tension to the surroundings,

but also as the sensor capable of detecting mechanical

stimulus. Mechanical stimuli sensed at focal adhesions

can be internal, as in the application of cytoskeletal

tension, or external, as in the application of environmental

forces. Signaling proteins associated with focal adhesions

transduce mechanical signals into chemical signals; these

chemical signals drive downstream pathways that regulate

cell functions including migration, apoptosis, and cell

cycle progression. Although the transduction of

mechanical signal into chemical signal is thought to be

governed by proteins within transmembrane focal

adhesions as shown in Figure 5A, the actual mechanical

stimulus sensed by the focal adhesion proteins is

modulated by surrounding elements of the system,

including the organization and density of F-actin

filaments, the density and rigidity of extracellular

components, and the size and distribution of the focal

adhesions.

We therefore propose the “3 spring” model depicted in

Figure 5B for the biophysical performance of mechanical

signal transduction complexes at sites of cell adhesion to

extracellular matrix. The cytoskeleton, adhesion complex

and matrix are the 3 component parts of this complex.

Each is conceptualized as a spring that may be

compressed or stretched by contiguous forces, and yet

may reconstitute its initial shape when these extrinsic

forces are removed. The “k” values in the figure denote

independent and individual stiffness coefficients for each

of the components. The model is configured with the 3

springs in series to indicate that differential stiffness

values have a direct impact on neighboring parts of the

compound spring: Increased stiffness in any single

component will increase the deformation of the other

components. Thus, stretch is enhanced in any one of the

triumvirate by either decreasing its own stiffness or by

increasing that of its neighbors. In other words, prestress

or active pulling force applied at either the cytoskeletal or

extracellular environmental ends of the compound

structure will result in a net change in the shape of any

given component that is dependent upon the relative

stiffness of each member of the tripartite spring.

Figure 5 – Cellular Mechanical Detection. The constituents and biophysical inter-relationships of the

proposed mechanical detection system are shown.

A: Intermittent and distinct transmembrane cell-matrix adhesion

complexes link the cytoskeleton and the matrix. They provide

discrete and spatially precise inputs that are integrated at both

the intracellular and the extracellular levels.

B: The cytoskeleton, transmembrane adhesion complex, and

matrix act as three springs of independent stiffness (k values) in

the proposed model. The deformability of each of these three

components is affected by the relative stiffness of the other two,

and also by prestress and subsequent force generation at both

the cytoskeletal and extracellular ends of the compound spring.



An exciting challenge to the student of mechanical signal

transduction is the paucity of detail currently known about

the molecular mechanisms that drive it, despite rapid

expansion in the field. Here follows a discussion of

candidate mechanosensors in each of the 3 components of

the “3 spring” complex (please also see Table I). To

begin, GEFs regulate Rho family activation states in the

cytoskeletal portion of this complex. One GEF that is an

attractive candidate mechanosensor molecule is the ~720

kDa, multidomain protein obscurin. Ankyrin-like repeats

that could tie activation of its Dbl homology domain to

membrane deformation and subcellular localization to

sarcomeric z-lines in developing cardiac myocytes

suggest that it may regulate tension, growth and

development in the heart in response to mechanical

stimuli (Russell et al., 2002; Borisov et al., 2003).

Another protein that may sculpt the cytoskeleton in

response to extracellular mechanics including plasma

membrane stretch is the Rho effector mDia-1 (Riveline et

al., 2001; Kozlov and Bershadsky, 2004). This formin is

a processive capper of growing actin filaments that may

act as a molecular elastic bridge that translates the

potential energy created by membrane stretch into an

impetus for focused G-actin subunit addition to

contiguous ends of F-actin.

TABLE I

Cell-Matrix Adhesion Components of the Cellular

Mechanical Response

Mechanical

Subunits

Candidate

Molecular

Signals

Effector Systems

Cytoskeleton RhoGEFs,

Rho,

Rac,

ROCK,

mDia1

actin polymerization

& crosslinking,

actomyosin

contraction,

microtubule targeting

Adhesion

Complex

integrins,

Src,

FAK,

p130Cas

integrin engagement

& clustering,

lateral mobility of

membrane proteins,

adhesion dynamics

Extracellular

Matrix

fibronectin,

laminin,

collagen

fibrillogenesis,

matrix density,

matrix mechanics

Many studies have led to a focus upon multi-

phosphocomponent signaling complexes at focal

adhesions as potential substrates for mechanical signal

response (Giannone and Sheetz, 2006). Assembly of

these complexes is enhanced by tyrosine phosphorylation

on scaffolding units that associate with SH2 domains of

signaling proteins including additional kinases (see

Romer et al., 2006 for review). The cytoplasmic non-

receptor tyrosine kinase FAK is also a substrate for Src,

and is one of these phosphocomponent signaling proteins

and has been implicated as a director of cell motility in

response to environmental mechanics (Li et al., 2002).

p130Cas is another Src substrate and has been the target

of elegant analysis of its mechanical signaling potential

(Sawada et al., 2006). The central substrate domain,

positioned between the amino-terminal SH3 domain and

the carboxy-terminal Src-binding domain, was shown to

be specifically tyrosine-phosphorylated by Src family

kinases and Abl in response to stretching to ~110% of

baseline cell-matrix interface. This stretch-activated

phosphorylation was not due to changes in the activities

of the kinases studied, but reflected enhanced substrate

susceptibility. This mechanically triggered molecular

event was then followed by the activation of the Rap1

GTPase, indicating the next step in signal propagation.

A third focal adhesion component with remarkable

responses to mechanical stimuli is zyxin. This LIM

domain protein interacts with p130 Cas, and has been

shown to mobilize from focal adhesions and enhance

fibroblast stress fiber alignment after 15 minutes of cyclic

stress (Yoshigi et al., 2005). Further, reduction in

cytoskeletal tension also triggers alteration of baseline

zyxin association with matrix adhesion sites in fibroblasts

(Lele et al., 2006). In vascular smooth muscle cells, 30

minutes of cyclic stretch induced nuclear translocation of

zyxin, where it was linked to the upregulation of

endothelin B expression (Cattaruzza et al., 2004).

Transmembrane integrins respond to extracellular matrix

ligand binding with long range allosteric rearrangements

that increase ligand affinity. In addition, integrin

clustering at sites of adhesion augments the adhesion

strength or avidity (Hynes, 2002). Both of these

mechanisms are apparently involved in the spatial

rearrangement of new ligand formation and the

downstream signaling through Shc that result from shear

stress exposure in human endothelial cells (Jalali et al.,

2001). These effects were abrogated by function-

blocking antibodies, and this additional finding indicates

an active role for matrix ligand presentation in the

regulation of integrin function.

Studies on the third component of the “3 spring” model –

the matrix – underline the active role that this partner has

in mechanical signaling to the cytoskeleton and beyond.

Fibronectin matrix density has been shown to modulate

cytoskeletal tension. Thus, a five-fold increase in the

density of fibronectin matrix presentation to bovine

vascular smooth muscle cells (from .02 to .1 µg/cm2)

induced an ~40% increment in myosin light chain

phosphorylation and increased cytoskeletal prestress to

250% of levels measured at the lower matrix density

(Polte et al., 2004).

The impact of Rho-modulated cytoskeletal contraction

and subsequent cell traction force transmission on

fibronectin fibrillogenesis provides a lucid application of

the “3 spring” model. Transmembrane adhesion

complexes transmit cell traction force to the fibronectin

matrix causing unfolding of its complex quaternary

structure and the exposure of fibronectin type III domain



repeats that form a “cryptic site” that favors fibril

assembly (Zhong et al., 1998). Fluorophore-labeling of

the seventh and fifteenth fibronectin type III domain

repeats allowed direct measurement of this molecular

unfolding by fluorescence resonance energy transfer.

Loss of signal exchange between the donor and receptor

fluorophores in this system indicated that cell traction

force transmission accomplished a > 10 nm distraction

between these two folded arms of the fibronectin

molecule (Baneyx et al., 2002). Other studies indicate

that laminin assembly may also be modulated by

mechanical traction from portions of the molecule that are

anchored to transmembrane integrins, and that these cell-

bound laminin molecules may also facilitate the

organization of collagen fibrils in the matrix environment

(Aumailley et al., 2003). Collagen fibrils also experience

direct actomyosin-driven rearrangements as they are

dragged across the fibroblast surface to be incorporated

into complex arrays that are essential mechanical

templates for efficient wound repair or normal embryonic

development (Kadler, 2004; Meshel et al., 2005).

Our data and those of others (Balaban et al., 2001;

Lemmon et al., 2005) indicate that cells generate forces in

the range of 1-5 nN per focal adhesion, and that focal

adhesions can sense and respond to forces of this

magnitude. Based on this information and an estimate of

cell mass, we can predict whether or not an individual cell

would sense a gravitational force resulting from its own

mass. Assuming an average cell volume of 5000 µm3 and

a cell density approximately equal to the density of water

(1 g/ml), the mass of a cell would be approximately 5 ng.

These assumptions are comparable to measured values of

osteoblast cell size and density (Searby et al., 2005). This

would result in a gravitational force of 50 pN. Assuming

that this force is equally distributed across all focal

adhesions, and that there is an average of 100 adhesions

per cell, the force at each focal adhesion would be 0.5 pN.

This would be 2000 times smaller than what could be

sensed at an individual focal adhesion. However, based on

this calculation, an inertial mass of approximately 5 µg

applied over the surface of a cell could be sensed at

individual focal adhesions – a value that is within range

for mammalian cells subjected to normal locomotion

(Lindemann, 2003).

In summary, mechanochemical coupling in cell-matrix

adhesion involves 3 partners: the cytoskeleton, the

transmembrane adhesion complex, and the extracellular

matrix. The cytoskeletal contraction and details of

cytoskeletal function that are cell-type specific regulate

cell traction force. Finally, a dynamic reciprocal

regulatory relationship exists between cell traction forces

and the extracellular matrix microenvironment, and this

may be modulated by inertial loads that are available in

most multicellular organisms.

ACKNOWLEDGEMENTS

The authors thank Dr. Jack van Loon for the opportunity

to learn from the ASGSB, the editors for their guidance,

Dr. Elena Doutova for her work on the Rappaport

chamber studies, and Dr. Fumin Chang for helpful

discussions and careful reading of the manuscript. This

work was supported by grants DE13079, HL058064, and

AI061042 from the NIH, and the Funds for Medical

Discovery from JHU to LR.

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