Journ

alof

Cell

Scie

nce

Finding the weakest link – exploring integrin-mediatedmechanical molecular pathways

Pere Roca-Cusachs1,*,`, Thomas Iskratsch2,* and Michael P. Sheetz2,3

1University of Barcelona and Institute for Bioengineering of Catalonia, Barcelona 08028, Spain2Department of Biological Sciences, Columbia University, New York, NY 10027, USA3Mechanobiology Institute of Singapore, National University of Singapore, 117411, Singapore

*These authors contributed equally to this work`Author for correspondence (rocacusachs@ub.edu)

Journal of Cell Science 125, 3025–3038� 2012. Published by The Company of Biologists Ltddoi: 10.1242/jcs.095794

SummaryFrom the extracellular matrix to the cytoskeleton, a network of molecular links connects cells to their environment. Molecules in this

network transmit and detect mechanical forces, which subsequently determine cell behavior and fate. Here, we reconstruct themechanical pathway followed by these forces. From matrix proteins to actin through integrins and adaptor proteins, we review howforces affect the lifetime of bonds and stretch or alter the conformation of proteins, and how these mechanical changes are converted

into biochemical signals in mechanotransduction events. We evaluate which of the proteins in the network can participate inmechanotransduction and which are simply responsible for transmitting forces in a dynamic network. Besides their individual properties,we also analyze how the mechanical responses of a protein are determined by their serial connections from the matrix to actin, their

parallel connections in integrin clusters and by the rate at which force is applied to them. All these define mechanical molecularpathways in cells, which are emerging as key regulators of cell function alongside better studied biochemical pathways.

This article is part of a Minifocus on Mechanotransduction. For further reading, please see related articles: ‘Deconstructing the third dimension – how 3D culturemicroenvironments alter cellular cues’ by Brendon M. Baker and Christopher S. Chen (J. Cell Sci. 125, 3015-3024). ‘Signalling through mechanical inputs – a coordinatedprocess’ by Huimin Zhang and Michel Labouesse (J. Cell Sci. 125, 3039-3049). ‘United we stand – integrating the actin cytoskeleton and cell–matrix adhesions in cellularmechanotransduction’ by Ulrich S. Schwarz and Margaret L. Gardel (J. Cell Sci. 125, 3051-3060). ‘Mechanosensitive mechanisms in transcriptional regulation’ by AkikoMammoto et al. (J. Cell Sci. 125, 3061-3073). ‘Molecular force transduction by ion channels – diversity and unifying principles’ by Sergei Sukharev and Frederick Sachs(J. Cell Sci. 125, 3075-3083).

Key words: Cell adhesion, Cytoskeleton, Mechanotransduction

IntroductionMechanical stimuli that are transmitted between a cell and its

environment determine cell functions, such as proliferation

(Nelson et al., 2005), differentiation (Engler et al., 2006) and

motility (Janmey et al., 2009), as well as key processes in

development (Gorfinkiel et al., 2009; Mammoto and Ingber,

2010), tumorigenesis (Kumar and Weaver, 2009; Paszek et al.,

2005) and wound healing (Fenteany et al., 2000), among others.

Mechanical transmission takes place through different cell

structures, but one of the main components, and definitely the

best studied one, is that of integrin-based cell adhesions. These

adhesions connect extracellular matrix (ECM) proteins to the cell

cytoskeleton through the transmembrane integrins, and a

molecular complex that can contain over a hundred different

types of adaptor molecules (Zaidel-Bar et al., 2007a). Adhesions

withstand and exert forces on the ECM even though they and the

cytoskeleton are dynamic structures. In addition, adhesions are

mechanosensitive elements that convert mechanical stimuli into

biochemical responses (Bershadsky et al., 2003; Riveline et al.,

2001) by sensing and responding to parameters such as matrix

rigidity (Moore et al., 2010). Precisely because of their

complexity, however, the mechanisms behind the mechanical

function of cell–ECM adhesions have been challenging to

unravel. The first step is to dissect where biophysics ends and

biochemistry starts; that is, we must identify the molecules that

exert, withstand, transmit and respond to forces. For those that

respond to force (by changing binding kinetics or by unfolding,

for instance) we must then evaluate whether and how this change

can lead to true force sensing by triggering biochemical signaling

cascades. Only after this is achieved will we be able to

distinguish the primary molecular events that detect forces

from ensuing biochemical signaling networks.

To address this issue, this Commentary will not focus on

signaling networks in adhesions, their assembly and dynamics, or

their detailed composition, topics which have recently been

reviewed elsewhere (Costa and Parsons, 2010; Dubash et al.,

2009; Parsons et al., 2010; Ridley, 2011; Wolfenson et al., 2009).

Rather, we will follow the path that force follows as it is

transmitted from the ECM to the cytoskeleton, analyzing the

molecules likely to be under mechanical stress. We will first

briefly review the molecular machinery that is responsible for

force generation and the biophysical effects that force has on

molecular conformation and binding. We will then analyze the

ECM molecules, integrins and adaptor molecules step-by-step.

For all the components of this protein network, we will review

current knowledge with regard to how they respond to force and

the possible links through which they could participate in force

transmission throughout the cell. We will then discuss the factors

that affect the collective mechanical behavior of all these

molecules that are present in adhesions. Finally, we will

Commentary 3025

Journ

alof

Cell

Scie

nce

discuss the signaling events that could be triggered by putative

mechanosensors. Owing to space limitations, we will not cover

adhesions that are mediated by transmembrane proteins other

than integrins (such as syndecans) or cell–cell adhesions.

First step of force transmission – theextracellular matrixCellular forces are primarily developed by the acto-myosin

cytoskeleton (Box 1) and have a profound effect on molecular

conformation and binding (Box 2). Starting at the cell exterior,

the first elements that forces encounter in their path are ECM

molecules, which form meshworks of polymerized fibers that

constitute the major components of the cell microenvironment.

Among ECM molecules, fibronectin has been the most studied

from a mechanical point of view. Forces of 80–200 pN can

stretch all the fibronectin FNIII domains, which contain the

integrin-binding sites (Table 1). Upon stretching, fibronectin

fibers can extend up to eightfold before breaking (Klotzsch et al.,

2009), which leads to unfolding of the FNIII domains and

exposure of cryptic sites. This unfolding, however, might account

for only a fraction of the fibronectin stretch, which could be

mostly determined by a global conformational change (Lemmon

et al., 2011). In any case, stretching of FNIII domains is a form of

mechanotransduction, because the exposure of the cryptic sites

leads to downstream effects, such as the regulation of the

fibrillogenesis of fibronectin (Sechler et al., 2001), stimulation of

cell growth and contractility (Hocking and Kowalski, 2002), and

stimulation of the digestion of gelatin, type IV collagen, a- and b-

casein and insulin b-chain (Schnepel and Tschesche, 2000).

Force can also change the conformation and distance between the

RGD and PHSRN motifs (two synergistic integrin-binding sites

within FNIII domains), altering integrin recognition and

specificity (Grant et al., 1997; Vogel, 2006). This stretching

and unfolding can be achieved in vivo by cell-generated forces

(Baneyx et al., 2001; Smith et al., 2007) and is controlled by cell

contractility (Baneyx et al., 2002).

Collagen I fibrils can also be stretched with forces of 1–4 nN,

which extrapolates to a force of 4–90 pN per collagen monomer

(Graham et al., 2004). The role of force in collagen function is

unclear, but many of the key binding sites in collagen, including

the GFOGER sequence that can bind to b1 integrin, are cryptic

and are only fully exposed upon proteolysis, partial denaturation

or force application (Orgel et al., 2011; Perumal et al., 2008).

Interestingly, collagens also possess several copies of the RGD

sequence (Davis, 1992), which has different integrin specificity

and is characteristic of other ECM molecules such as fibronectin

(Hynes, 2002). In collagen, the RGD site is only exposed upon

denaturation (which can be caused by force), which could change

integrin specificity and trigger integrin-mediated wound response

(Davis, 1992). The coiled-coil sequences in the fibrinogen a-

helices also unfold under forces of ,100 pN (Table 1), and this

molecular unfolding seems to be the main factor determining the

elasticity of fibrin fibers in blood clots (Brown et al., 2009).

Fibrinogen unfolding can also activate its integrin receptor,

macrophage-1 antigen (MAC1 or integrin aMb2) and promote

inflammation, suggesting that there is a possible mechanosensing

mechanism (Deng et al., 2011). The relevance of mechanical

stimuli for the function of other ECM molecules, including

laminins and vitronectin is essentially unexplored. However, the

list of cryptic binding sites in these molecules that could be

exposed by force is extensive (Davis et al., 2000), opening many

intriguing possibilities.

IntegrinsIntegrins are heterodimeric transmembrane proteins composed of

an a- and b-subunit, and are major molecular links between cells

and the ECM. Different combinations of subunits lead to 24

different types of integrins in humans and determine their

specificity for ECM molecules (Hynes, 2002). Whereas most of

the integrin molecule is located extracellularly, to bind to the

ECM, a short cytoplasmic tail binds to adaptor proteins, as

discussed below. Although force does not unfold integrins, it can

affect them through other mechanisms. First, applied forces can

disrupt integrin–ECM bonds. Such rupture forces have been

studied extensively for the bonds between fibronectin and a5b1

integrin, and fibrinogen and aIIBb3 integrin (see Table 1). Given

the variety of experimental approaches used, the narrow range of

reported unbinding forces (50–100 pN) is remarkable. Integrin–

collagen interactions have been far less studied, but the binding

strength between the GFOGER binding site in collagen and a2b1

integrin has been reported to be 160 pN (Niland et al., 2011),

suggesting that they have a higher resistance to force than

integrin–fibronectin adhesions. When rupture forces are

measured in live cells containing many integrin–ECM bonds,

Box 1. Force generating machinery

Polymerization

Upon binding to actin filaments, actin monomers release energy,

which the filaments can use to generate force and ‘push’ any

structure opposing polymer growth (such as the cell membrane).

Indeed, cells migrate and spread through this mechanism. Actin

polymerization forces at the cell migrating edge (lamellipodium)

are ,2 nN/mm2 (Heinemann et al., 2011; Prass et al., 2006). Of

course, this force generation fluctuates, and the cell also uses a

myriad of molecules, including the Arp2/3 complex, Wiskott–

Aldrich syndrome protein (WASP), cofilin, formins and many

others, to regulate actin dynamics (for a review, see Ridley, 2011).

Polymerizing microtubules can also generate forces of up to 4 pN

(Dogterom and Yurke, 1997), although their contribution to

protrusive forces is much smaller owing to their lower density

and their catastrophic disassembly events. Intermediate filaments

cannot generate directed forces owing to their lack of polarity.

Molecular motors

Molecular motors bind to cytoskeletal filaments; myosins bind to

actin (Sellers, 2000), and kinesins (Hirokawa et al., 2009) and

dyneins (Pfister et al., 2006) bind to microtubules. They undergo

ATP-fueled conformational changes that generate movement and

forces on the filaments. In processive motors such as kinesin 1,

the two tubulin-binding heads coordinate to continuously ‘walk’

over the filament (Alonso et al., 2007), generating forces of up to

6 pN (Kuo and Sheetz, 1993; Meyhofer and Howard, 1995;

Svoboda and Block, 1994), transporting organelles and vesicles

(Hirokawa et al., 2009). Non-processive motors such as myosin II

are not coordinated in the same way and require myosin filaments

to generate forces (Harada et al., 1990). Myosin II filaments

generate large forces in muscle contraction or fibroblast–ECM

adhesion (Cai et al., 2006) of up to 6 pN per myosin head (Finer

et al., 1994; Ishijima et al., 1994; Tyska et al., 1999). Their

concerted effort can generate stresses of tens of nanonewtons per

square meter on the ECM (Balaban et al., 2001; du Roure et al.,

2005; Tan et al., 2003).

Journal of Cell Science 125 (13)3026

Journ

alof

Cell

Scie

nce

the calculated force per bond is much lower (below 10 or even

1 pN) than that obtained from single-molecule measurements

(Friedland et al., 2009; Roca-Cusachs et al., 2009; Walter et al.,

2006). This suggests that in integrin clusters, only a small

fraction of molecules withstand forces (Moore et al., 2010).

The bond between fibronectin and a5b1 integrin shows catch

bond behavior (i.e. a bond that strengthens when force is applied;

see Box 2) in which maximum lifetimes occur at forces of 10–

30 pN (Kong et al., 2009). Such catch bond behavior has not

been observed for aIIb3 (Litvinov et al., 2011) or avb3 integrin.

This could explain the much higher resistance to force of

fibronectin–a5b1-integrin bonds compared with those of

fibronectin–avb3-integrin bonds (Roca-Cusachs et al., 2009)

although the latter are required for force sensing. Thus, although

stable a5b1 catch bonds can regulate adhesion, more fragile avb3

integrin bonds might constitute mechanosensors that undergo

frequent unbinding and rebinding events under force, triggering

downstream events (Arias-Salgado et al., 2005; von Wichert et al.,

2003). In parallel to possible catch bond behaviors, integrins also

undergo conformational changes (activation) that prime them for

binding to the ECM (Luo et al., 2007; Moser et al., 2009).

Activation can also change their affinity for cytoplasmic partners

such as a-actinin (Sampath et al., 1998) and catalyze downstream

events including tyrosine phosphorylation of different substrates

(Legate et al., 2009). Structurally, integrin activation involves

separation of all domains except the ECM-binding head, and can

be triggered by either ECM binding itself (so-called ‘outside-in

activation’) (Kim et al., 2003; Takagi et al., 2002) or binding to

Box 2. Biophysics of unfolding proteins and breaking bonds

Forces promote unfolding [a disruption of secondary or tertiary structure that increases the length of the molecule between the pulling ends

(Bustamante et al., 2004; Kumar and Suan Li, 2010; Sulkowska and Cieplak, 2008)], bond dissociation (slip bonds), or in some cases even bond

strengthening (catch bonds).

Unfolding and slip bonds

Unfolding and unbinding (Figure panel A) can be modeled by two equilibrium states (folded or bound versus unfolded or unbound) that are

separated by a certain distance and an energy barrier (green line, DE) (Bustamante et al., 2004). Applied forces ‘tilt’ the energy landscape (blue

line), reducing the energy barrier between states (orange line, DEF), and thereby decreasing the average time needed for unfolding or unbinding

(Bell, 1978). The rate at which force is applied (the loading rate) increases the average force at which unfolding takes place (Evans and Ritchie,

1999; Rico et al., 2007). This is because the time required to reach a given force will be shorter with fast loading, thereby reducing the likelihood

of an unfolding event during the process. Thus, loading rates must be considered when comparing measurements.

Catch bonds

Catch bonds (Figure panel B) strengthen, rather than weaken, with force. This could be because force is either opposed by ‘hook-shaped’

molecules (i), the force aligns binding sites by stretching them (ii), or because it displaces an inhibiting region hiding a cryptic binding domain (iii)

(Thomas et al., 2008). In the hook example (i), without force application, unbinding is much easier if the hook is separated (path 1, DE1)

compared with deforming it (path 2, DE2). Force applied in the direction of hook deformation, however, impairs unbinding in the opposite

direction (DE1F.DE1), but helps to deform the hook (DE2F,DE2). As the bond tends to unbind through the path with the lowest barrier, the

maximum bond strength is when DE1F5DE2F, and will decrease for higher forces. Thus, catch bonds have an optimum force range at which they

are stable, a very interesting feature, with implications discussed in the main text.

ΔEF

ΔE

ΔE2F

ΔE2

ΔE1

Bound Unbound

BoundUnbound

Unbound

No force Force

A

B

Ene

rgy

Distance

Ene

rgy

Distance

No force Force

Energy without force

Energy with force

Effect of force

Energy withoutforce

Energy with force

(i)

(ii)

(iii)

Path 2 – Unbinding in the direction of force

Path 1 – Unbinding againstthe direction of force

ΔE1F

ΔEF

ΔE

ΔE2FEE

ΔE2EE

ΔE1

Bound Unbound

BoundUnbound

Unbound

No force Force

A

B

Ene

rgy

Distance

Ene

rgy

Distance

No force Force

Energy without force

Energy with force

Effect of force

Energy withoutforce

Energy with force

(i)

(ii)

(iii)

Path 2 – Unbinding inthe direction of force

Path 1 – Unbinding againstthe direction of force

ΔE1F

Exploring mechanical molecular pathways 3027

Journ

alof

Cell

Scie

nce

its cytoplasmic partner talin (Bouaouina et al., 2008; Calderwood

et al., 1999), possibly in combination with kindlin (Moser et al.,

2008) (so called ‘inside-out activation’). Integrin activation can

also be triggered by force, as shown for a5b1 catch bonds

(Friedland et al., 2009) and suggested by results of steered

molecular dynamics simulations (Puklin-Faucher et al., 2006).

Because integrin-binding partners, such as talin, filamin and a-

actinin, bind to the cytoplasmic integrin-b tails (but not a-tails)

(Otey et al., 1990; Pfaff et al., 1998), force application from the

cytoskeleton through these partner molecules could induce the

integrin subunit separation necessary for activation (Puklin-

Faucher and Sheetz, 2009). Thus, force-induced integrin

activation can also be considered to be a mechanotransduction

event.

Adaptor proteinsIntegrins, which lack actin-binding domains, are indirectly

connected to the actin cytoskeleton through different protein

complexes (Fig. 1) that form different types of cell–ECM

adhesions, including nascent adhesions, focal complexes, focal

adhesions, podosomes and others (Dubash et al., 2009). These

complexes can contain over a hundred different types of proteins

(Zaidel-Bar et al., 2007a), with many binding in a force-

dependent manner (Pasapera et al., 2010; Schiller et al., 2011).

Here, we will focus on the path that force must follow from

integrins to actin and its effect on the molecules involved.

The connection between integrin and actin can be mediated by

a single molecule, such as talin. Given that the N-terminal head

domain of talin binds to and colocalizes with integrin-b tails, and

the C-terminal domain binds to and colocalizes with actin

(Critchley, 2009; Kanchanawong et al., 2010), talin is ideally

positioned in the force path between integrins and actin. Talin-

depleted fibroblasts initially spread normally, but then retract and

round up instead of developing mature adhesions (Zhang et al.,

2008). This behavior is rescued by full-length talin but not by the

integrin-binding and -activating talin head alone, suggesting that

the force transmitted between integrins and actin by full-length

talin has a key role. Indeed, forces of 12 pN can expose cryptic

Table 1. Mechanical parameters of adhesion proteins

Unfolding Unbinding Stiffnessa

Protein Force (pN) Speed (mm/s) Reference InteractionForce(pN)

Speed(mm/s) Reference pN/nm Reference

ECM

Fn 137 0.2–0.6 (Oberhauseret al., 1998)

Fn–a5b1 69–93b 5 (Li et al.,2003a)

0.5 (Oberhauseret al., 1998)

80–200 0.6 (Oberhauseret al., 2002)

Fn–a5b1 39 0.8 (Sun et al.,2005)

Fb or Fib 150–200 1 (Averett et al.,2009)

Fib–aIIbb3 50–80 0.16–1 (Agnihotriet al., 2009)

1–3 (Brown et al.,2007)

100 (Brown et al.,2007)

Fb–aIIbb3 80–100 80 (Litvinov et al.,2002)

6 (Lim et al.,2008a)

130 0.4–1 (Lim et al.,2008a)

Fib–aIIbb3 80–90 0.8–80 (Litvinov et al.,2005)

RGD–aIIbb3 93 0.2 (Lee andMarchant,2001)

Collagen I 4–90 18000 (Graham et al.,2004)

Col–a2b1 160 0.5 (Niland et al.,2011)

1.5c (Thompsonet al., 2001)

Adaptor proteins

Talin 29–51 0.4 (del Rio et al.,2009)

Actin 2 0.06 (Jiang et al.,2003)

1.5 (del Rio et al.,2009)

a-Actinin Actin 40–78 4–50d (Ferrer et al.,2008)

1700 (Paramoreet al., 2006)

Actin 1.4–44 ? (Miyata et al.,1996)

Filamin 57 5 (Lee et al.,2009)

Actin 40–65 4–50d (Ferrer et al.,2008)

200–800 (Lee et al.,2009)

48 6 se (Chen et al.,2011)

Actin 70 5 (Lee et al.,2009)

50–220 0.37 (Furuike et al.,2001)

Titin kinase 30 1 (Puchner et al.,2008)

aStiffness values were calculated from reported force–extension curves at low extensions, or calculated as first-order approximation from reported worm-likechain parameters at low deformations.

bThe range from inactive to activated is shown.cValue might correspond to more than one molecule in parallel.dLoading rates in pN/second. To convert retraction speeds into loading rates, the stiffness (spring constant) of the system (studied molecules + measuring probe)must be determined. This value will be dominated by the softest component, and is not always reported.

eTime required to unfold protein at constant force.Fn, fibronectin, Fb, fibrinogen; Fib, fibrin; Col, collagen.

Journal of Cell Science 125 (13)3028

Journ

alof

Cell

Scie

nce

binding sites in talin molecules and cause vinculin binding (del

Rio et al., 2009), which might explain why talin is required for

adhesion reinforcement and maturation (Jiang et al., 2003; Roca-

Cusachs et al., 2009). In cells, talin can stretch cyclically from itsfolded length of 50 nm to over 400 nm in a vinculin-dependent

manner, and the stretching cycles correlate with the lifetime of

vinculin in adhesions (Margadant et al., 2011). Thus, vinculin,

which has been reported to withstand forces of up to 2.5 pN(Grashoff et al., 2010), might reinforce the mechanical integrin–

actin connection by simultaneously binding talin and actin

(Burridge and Mangeat, 1984; Menkel et al., 1994). However, the

tension level in vinculin molecules has not been directly relatedto its recruitment to adhesions, and vinculin-null cells have been

reported to have both impaired (Diez et al., 2011) and normal

(Grashoff et al., 2010) focal adhesions and contractile forces.

Therefore, the exact role of vinculin in force transmission and

mechanotransduction remains unclear.

Several other molecules can also simultaneously bind to actin

and integrins, including a-actinin (Pavalko et al., 1991; Pavalkoand LaRoche, 1993), filamin (Hartwig and Stossel, 1975; Loo

et al., 1998; Sharma et al., 1995) and tensin (Calderwood et al.,

2003; Lo et al., 1994). a-Actinin is an antiparallel dimer and a

key actin crosslinker (Sjoblom et al., 2008) that is involved inadhesion maturation (Choi et al., 2008). The crosslinking role of

a-actinin is important in adhesions, because normal adhesion

elongation is recovered in a-actinin-depleted cells when actin

crosslinking is restored through use of a myosin mutant that

constitutively binds actin (Choi et al., 2008). However, thetransmission of forces between integrins and actin by a-actinin

could also have a relevant role. Indeed, inactivating a-actinin in

stress fibers leads to their detachment from focal adhesions(Rajfur et al., 2002), and blocking force transmission with

exogenous a-actinin fragments that contain only the actin- orthe integrin-binding domain, respectively, can lead to thedisassembly of stress fibers and focal adhesions (Pavalko andBurridge, 1991; Triplett and Pavalko, 2006). a-Actinin has

cryptic sites for binding to the smooth muscle protein smitinwithin its spectrin-repeat domains (Chi et al., 2005), althoughthese domains might be too stiff to be stretched physiologically

(Table 1). Filamin is composed of an actin-binding domain and24 immunoglobulin-like domains (Popowicz et al., 2006), andalso forms dimers that can orthogonally crosslink actin, thus

forming three-dimensional meshworks (Flanagan et al., 2001;Hartwig and Shevlin, 1986). This crosslinking activity is crucialfor motility, actin architecture, force generation and rigiditysensing (Byfield et al., 2009; Cunningham et al., 1992; Flanagan

et al., 2001; Lynch et al., 2011). However, the binding of filaminto integrins is also important, as filamin competes with talin forintegrin binding (Kiema et al., 2006), thereby regulating talin-

mediated adhesion (Nieves et al., 2010).

Forces of 40–70 pN can both detach filamin from actin andunfold the immunoglobulin-like domains of filamin (Table 1),

which contain binding sites to multiple partners. This suggeststhat different force-loading rates and directions of forceapplication can lead to either unfolding or unbinding, or tounfolding followed by unbinding as proposed for the actin-

associated cytoskeletal proteins spectrin and ankyrin in red bloodcells (Krieger et al., 2011). Shear strain applied to actin networkscrosslinked with filamin increases integrin binding (Ehrlicher

et al., 2011), probably by unfolding the 21st immunoglobulin-likedomain and exposing a binding site for integrin-b tails (Chenet al., 2009; Pentikainen and Ylanne, 2009). However, strain also

reduces the binding of filamin-A-associated RhoGAP (FILGAP,also known as Rho GTPase-activating protein 24) (Ehrlicheret al., 2011), possibly by separating the two FILGAP-binding

domains present in the filamin dimer. This, therefore, is anelegant mechanism by which a single mechanical stimulus couldregulate the binding of two proteins in opposite ways. Finally,tensin also forms dimers that have multiple actin-binding sites

and a domain that binds to the NPXY motifs on integrin-b tails(Lo, 2004). In fibroblasts, colocalization of tensin with integrinsis much more prominent in mature fibrillar adhesions than in

early focal contacts (Zamir et al., 2000), pointing towards a rolein adhesion maturation. Tensin localization also requires talin andintegrin-linked kinase (ILK) in Drosophila (Torgler et al., 2004),

suggesting that there are interactions between the threemolecules. Expression of a tensin fragment containing the actinhomology 2 region can inhibit adhesion formation and the normal

rearward translocation of a5b1 integrins (Pankov et al., 2000),indicating that tensin normally transmits force from thecytoskeleton to integrins and is necessary for adhesionmaturation. Such a mechanical function is further confirmed by

the fact that the defects observed in tensin-null Drosophila (i.e.wing blisters) only appear after mechanical stimulation (Torgleret al., 2004). However, no information is currently available on

the mechanical properties of tensin or its molecular interactions.

Links formed by more than one molecule can also connectintegrins to actin. For example, ILK binds to integrin-b tails and

connects to actin through an interaction with parvin, forming theILK–pinch–parvin (IPP) complex (Wickstrom et al., 2010). Thiscomplex regulates cell contractility (Wickstrom et al., 2010) and

α-Acti

nin

Filam

in

Tens

in

α βα β α β α β

Vin

α β

ILK

Parvin

Pinch

Arp2/3

FAK

α β

Myo

sin X

Src

p130Cas

Talin

MLCK

Myosin II

(i)

(ii)

(iii)

(iv) (v)(vi)

(vii)

ECM

(viii)

Actin

Fig. 1. The actin–integrin mechanical connection. Known direct and some

indirect connections are shown here. For FAK and p130Cas, possible links

connecting them to integrins and actin are shown. Myosin X (MYO10, not

discussed in the main text) also binds integrin-b tails. Rather than exerting

contractile forces between integrins and actin, however, this myosin binds

integrins to transport them to adhesion sites (Zhang et al., 2004). The steps

indicated in roman numbers show proposed molecular mechanosensing

mechanisms: force could be detected by (i) exposing cryptic domains in ECM

molecules, (ii) increasing the lifetime of fibronectin–a5b1-integrin bonds

(catch bonds) and activating integrins, (iii) increasing the level of

phosphorylation of p130Cas substrate domain, (iv) releasing FAK auto-

inhibition and increasing its tyrosine kinase activity, (v) exposing previously

buried vinculin-binding sites in talin rod domain, (vi) exposing previously

buried binding sites to integrin-b tails or FIP in filamin immunoglobulin-like

domains, (vii) releasing MLCK auto-inhibition and activating myosin II, and

(viii) increasing the lifetime of myosin-IIA–actin bonds (catch bonds).

Exploring mechanical molecular pathways 3029

Journ

alof

Cell

Scie

nce

mechanosensation in touch neurons (Hobert et al., 1999). Kindlinparticipates in integrin activation (Moser et al., 2008) and could

connect to filamin through migfilin (Tu et al., 2003). Given thehigh degree of complexity in the protein interaction networks infocal adhesions (Zaidel-Bar et al., 2007a), several additionalpaths connecting actin to integrins are possible. Force transmitted

through these paths could be sensed by mechanosensory proteins,because several proteins change enzyme activity or becomephosphorylated in response to force (Moore et al., 2010). Several

proteins are also recruited by myosin II contractility, includingFAK, zyxin, vinculin, a-actinin and LIM proteins such as pinch,zyxin, paxillin and migfilin (Pasapera et al., 2010; Schiller et al.,

2011). However, which of the molecules affected by force aretrue mechanosensors (i.e. they respond directly to forces) andwhich simply act downstream, remains unclear. For instance,LIM proteins are likely to bind to exposed sites in

mechanonsensing proteins but are not mechanosensorsthemselves (Schiller et al., 2011). Some insight into thisquestion has been obtained by stretching of membrane-stripped

cytoskeletons, which should eliminate many downstreambiochemical pathways present in live cells. Cytoskeletalstretching leads to increased cytoskeletal binding of paxillin,

focal adhesion kinase (FAK), p130Cas (also known as BCAR1),protein kinase B (Akt), Rap guanine nucleotide exchange factor 1(known as C3G, RAPGEF1 and GRF2) and Crk isoform II

(Sawada and Sheetz, 2002; Tamada et al., 2004). Despite theseobservations, direct evidence for mechanosensing in individualmolecules is scarce. Mechanical stretching of the substratedomain of p130Cas increases its phosphorylation by Src family

kinases in vitro, enabling the binding of several proteins (Sawadaet al., 2006). p130Cas is thus one of the very few demonstratedmechanosensors, although the molecular links through which it

would be stretched in cells are unclear. Force applicationincreases the tyrosine kinase activity of FAK (Tomar andSchlaepfer, 2009), possibly by disrupting an inhibitory

interaction between its FERM and kinase domains (Cooperet al., 2003). FAK connects indirectly to integrins and actin,through proteins such as talin (Chen et al., 1995), paxillin(Hildebrand et al., 1995), or Arp3 (Serrels et al., 2007). In

summary, integrins can be connected to actin through one ormore molecules, but, with the exception of talin, it remainsunclear which of them experience and respond to forces in vivo.

MyosinThe chain of forces goes from matrix to integrins to adaptor

proteins bound to actin, and the actin filament network transmitsforces exerted by myosin motors. Myosin I detachment fromactin filaments is decreased by up to 75-fold under forcesgreater than 2 pN, in a mechanism that could contribute to

mechanosensing in hair cells (Laakso et al., 2008). Similarly, themyosin-II–actin interaction behaves like a catch bond, with anoptimal load of 6 pN (Guo and Guilford, 2006). This is also the

maximum force that the motor can exert, indicating that themotor is designed to hold for the longest time at maximal force.External forces might, thus, optimize myosin II binding and

promote further force generation by the cell. Additionally,myosin light chain kinase (MLCK) binds both myosin andactin (Hatch et al., 2001; Sellers and Pato, 1984; Yang et al.,

2006; Ye et al., 1997) and moves rearward with actin during cellspreading (Giannone et al., 2004). MLCK could therefore bestretched between myosin and rearward flowing actin, resulting

in the release of MLCK auto-inhibition and in myosin IIactivation. Mechanoactivation has indeed already been observed

for another member of the MLCK family, the kinase titin, whichis unfolded by 10 nm under forces of 30 pN (Gautel, 2011;Puchner et al., 2008).

Alternative pathwaysIntegrins and the cytoskeleton can also transmit forces without

involving an integrin–actin link. For instance, increases inmembrane tension that result from actin polymerization canactivate contractility and exocytosis (Gauthier et al., 2011), or

regulate the activity of stretch-sensitive ion channels (Sbranaet al., 2008). Mechanosensitive ion channels also associate withb1 integrins (Chao et al., 2011), and might be activated by forces

transmitted through them (Matthews et al., 2010). Forceactivation of these channels leads to adhesion reinforcement,and affects migration, contractility and recruitment of vinculin

(Matthews et al., 2006; Munevar et al., 2004). Besides ionchannels, membrane tension (or ECM deformation) mightregulate integrin spacing, thereby stretching integrincrosslinking proteins such as receptor-type tyrosine-protein

phosphatase alpha (RPTP-a), which can expose a catalytic cleftleading to the activation of Src family kinases (Jiang et al., 2000;von Wichert et al., 2003).

Besides actin, intermediate filaments and microtubules alsohave a role in force transmission, which is much less explored.The intermediate filament protein vimentin localizes to adhesion

sites, and its depletion reduces cell contractility, adhesion sizeand migration (Eckes et al., 1998; Tsuruta and Jones, 2003).Furthermore, vimentin regulates cell extensions through its

interaction with filamin (Kim et al., 2010), depolymerizes uponmyosin II inhibition (Johnson et al., 2007), colocalizes and co-precipitates with a2b1 integrins (Kreis et al., 2005), and is

required for proper b1 integrin localization at adhesions (Ivaskaet al., 2005). Microtubules are also targeted to early adhesionsites (Kaverina et al., 1998), although their depolymerizationleads to formation of stress fibers and focal adhesions (Kaverina

et al., 1998; Liu et al., 1998), and increased contractility(Stamenovic et al., 2002). This result has led to the hypothesisthat microtubules normally carry contractile compressive

stresses, but it could also be explained by there being increasedmyosin contractility upon microtubule depolymerization(Kolodney and Elson, 1995). Thus, although cellular forces are

mostly generated by actomyosin contractility (Cai et al., 2006),microtubules and intermediate filaments could transmitmechanical forces in special cases.

Putting it all together – integrin clusteringIn summary, forces affect a complex series of molecular links that

connect the ECM to the actin cytoskeleton. These links can operatein parallel, given that many different proteins can simultaneouslysupport forces (Fig. 2A). The resulting collective behavior

naturally depends on the rate of force loading and cannotdirectly be predicted from individual bond properties. At lowloading rates, for instance, when a bond breaks, the force

redistributes to the remaining bonds, leading to a fasterdetachment. The average detachment force per bond can then besmaller in a cluster than in an individual bond (Seifert, 2000). At

faster loading rates, rupture is too fast and this effect is notimportant (Sulchek et al., 2005). However, in live cells, maximumforce per bond increases sixfold in pentameric versus monomeric

Journal of Cell Science 125 (13)3030

Journ

alof

Cell

Scie

nce

fibronectin–a5b1 bonds (Roca-Cusachs et al., 2009), indicating

that, beyond physical factors, protein complexes in integrin

clusters stabilize mechanical links. Indeed, integrin cluster

formation involves integrin interactions with kindlin, talin and

phosphatidylinositol (4,5)-bisphosphate [PtdIns(4,5)P2] (Moser

et al., 2009; Saltel et al., 2009), as well as other integrins (Li

et al., 2003b). These interactions require a spacing that is smaller

than 60 nm because larger distances between integrins in

nanopatterned substrates do not sustain cell adhesion (Arnold

et al., 2008; Jiang et al., 2003; Schvartzman et al., 2011).

Once formed, actin filaments grow from clusters, and contractile

forces between them bring them together (Rossier et al., 2010; Yu

et al., 2011; Ghassemi et al., 2012) (Fig. 2B). This requires a link

between adhesion sites and polymerizing actin that could be

provided by formins. Formins are found on dorsal stress fibers that

emanate from focal adhesions and regulate the distribution of focal

adhesions at the lamellum–lamellipodium interface, as well as

focal adhesion turnover (Gupton et al., 2007; Hotulainen and

Lappalainen, 2006). Formins elongate filament barbed ends at

rates of up to 27 nm/second for the formin mDia2 or 120 nm/

second for mDia1, as measured in vitro (Chesarone and Goode,

2009; Kovar et al., 2006). These rates are well within the range of

rearward flows of actin in vivo (see below). Supporting this

hypothesis, formin-mediated polymerization of single actin

filaments has been measured to exert forces greater than 1 pN

(Kovar and Pollard, 2004). Thus, actin polymerization from

clustered integrins could generate forces on adhesions, but details

of the interactions need to be worked out. What is clear, however,

is that the mechanical behavior of adhesion molecules depends not

only on their individual properties but also on their density and

spacing in clusters, their interactions and the force loading rate.

Rearward flow and loading rateAlthough contractile forces pull initial integrin clusters towards

each other, once clusters grow and mature, force is applied

mainly by actin flowing rearward from the leading edge to the

center of cells (Fig. 2C). External stimuli can also apply forces

through the ECM, which can be transmitted to distant points

across a cell (Hu et al., 2003), activating factors such as Src (Na

et al., 2008) or Rac (Poh et al., 2009). However, endogenous

rearward flow appears to regulate adhesion maturation and

traction force generation. This flow is mainly driven by myosin II

contractility because inhibition of myosin decreases the velocity

of actin flow by over two thirds (Cai et al., 2006) and leads to

unbalanced actin polymerization forces that dramatically increase

membrane tension (Gauthier et al., 2011).

In a variety of cell types and depending on the location within

the cell, rearward actin flow is in the range of 2 to 600 nm/second

(Fisher et al., 1988; Forscher and Smith, 1988; Gardel et al.,

2008; Theriot and Mitchison, 1991). Generally, higher flows are

associated with weak or nascent adhesions and low traction

forces on the substrate, and vice versa (Chan and Odde, 2008;

Kress et al., 2007; Zhang et al., 2008). This suggests that high

actin speeds lead to high force loading rates that decrease bond

lifetimes (Box 2) and therefore the ability to transmit force from

actin to adaptor proteins and integrins (Li et al., 2010). This

effect of loading rate would no longer be relevant at speeds below

10 nm/second, at which point the relationship between rearward

flow and traction force becomes direct rather than inverse

(Gardel et al., 2008). Given the transient nature of the mechanical

links, actin flow is coupled best to actin-binding proteins, such as

talin and particularly a-actinin, which also move rearward at a

speed of ,5 nm/second, less efficiently to FAK, paxillin or

Integrins

Adaptors

Myosin

Actin

A

C

Integrins

Adaptors

Actin

ECMKE

KI

KA

FEI

FIA

FAA

v

k1 k3k2s

B

Fig. 2. Summary of mechanical factors in integrin adhesions. (A) Role of stiffness. Whether and where a single connection between integrins and actin unfolds

or breaks under force will depend on the stiffness of the ECM molecule (KE), integrin (KI), and adaptor protein (KA), and the maximum force that can be withstood

by the ECM–integrin bond (FEI), the integrin–adaptor bond (FIA) and the adaptor–actin bond (FAA). These stiffness values will generally increase with protein

stretching, and change after unfolding or unbinding events. To our knowledge, integrin stiffness (KI) or possible unfolding have not been described. The total

stiffness of the chain (k1) will be determined by the softest element in it, and will operate in parallel with other chains in the cluster (k2, k3). The rearward speed v

of the actin layer, multiplied by k1, will provide the rate of force loading in the chain, which will affect both individual rupture forces (FEI, FIA, and FAA) and the

effectiveness with which the different chains in the cluster cooperate to withstand force. The spacing between integrins (s) will also affect cluster formation.

(B) Nascent focal adhesions. In nascent adhesions, adaptor proteins mediate the formation of initial integrin clusters {containing talin, kindlin and PtdIns(4,5)P2},

and might promote actin polymerization from clusters (formins). Myosin II filaments exert local contractile forces between filaments (blue arrows). (C) Focal

adhesion maturation. As adhesions mature and grow, they connect to actin fibers moving rearward towards the cell center due to global myosin contractility (blue

arrows). Rearward moving actin pulls on bound adaptor proteins, which in turn pull on integrins. Because of transient stick-slip bonds, rearward speeds are

reduced with respect to actin to a certain extent in adaptor proteins, and even further in integrins (black arrows).

Exploring mechanical molecular pathways 3031

Journ

alof

Cell

Scie

nce

vinculin, and even less efficiently (,1.3 nm/second) to substrate-bound av integrins (Hu et al., 2007). The efficiency of this

transmission also depends on the ligand density in the ECM(Brown et al., 2006), showing that molecular clustering is adeterminant of this process. Thus, in mature adhesions, transient

stick–slip bonds seem to couple integrins to adaptor proteins andto the actin cytoskeleton (Margadant et al., 2011).

The rearward speeds of proteins, multiplied by their stiffness

(i.e. the ratio between applied force and resulting molecularextension), will determine force-loading rates and thus bondlifetimes (Box 2). Protein stiffness increases as molecules are

extended, often following a worm-like chain behavior (Markoand Siggia, 1995), but constant stiffness values can beapproximated for small extensions (Table 1). In a serialconnection between an ECM molecule, an integrin and an

adaptor protein, the total stiffness would be dominated by thesofter elements (Fig. 2A). For instance, if fibronectin (0.5 pN/nm) is linked to talin (1.5 pN/nm), both elements are soft and

would probably extend and possibly unfold, enablingmechanosensing events. If fibronectin is linked to a-actinin(1700 pN/nm), however, only fibronectin would extend, with a-

actinin simply transmitting the force. This difference couldexplain the reported mechanosensitive unfolding of talin (del Rioet al., 2009) but not of a-actinin. Indeed, talin molecules inadhesions continually extend and relax in concert with the

concomitant binding and release of vinculin (Margadant et al.,2011). However, the timescales of talin stretch–relax cycles (6–16 seconds) do not match those of traction forces (30–

90 seconds) (Ghassemi et al., 2012). This suggests that actinflow might transmit forces through stiff molecules such as a-actinin, while signaling by repeatedly stretching talin, then

unbinding from it, and finally rebinding again. At a loading rateof 2.5 pN/second (0.5 pN/nm and retraction speed of 5 nm/second), for instance, a stretch cycle of 16 seconds would build

a force of 40 pN, sufficient to unfold talin and triggermechanotransduction (Table 1). Given the high retractionspeeds used in talin unfolding experiments (400 nm/second,Table 1), in vivo unfolding forces would however be much

lower. Thus, it remains unclear whether unfolding or unbindingfrom actin (measured at 2 pN at 60 nm/second, Table 1) woulddominate. The interaction between these two events would define

the signaling effectiveness of talin stick–slip cycles. This conceptof stick–slip stretching of molecules as a primary force signalingmechanism has yet to be tested in various systems, but it could

explain how the cell ‘samples’ adhesions over time and generatesintegrated signals to the nucleus or other parts of the cell.

Downstream signalingDownstream signaling from molecular mechanosensing events ismediated by phosphorylation, dephosphorylation, small GTPase

signaling and phospholipid signaling, among others (Fig. 3).Whereas the ensuing signaling networks are vast, we will discusshere examples of known signaling cascades that emanate from

the specific mechanosensing events discussed above.

RPTP-a

The mechanically activated receptor-type tyrosine-proteinphosphatase alpha (RPTP-a, encoded by PTPRA) transmits the

mechanical signal to Src family kinases (SFKs), which areactivated by dephosphorylation (Jiang et al., 2006; Roskoski,2005; von Wichert et al., 2003; Ylinen, 1996; Zheng et al., 2000).

Alternatively, clustering of integrins could also directly activate

SFKs (Arias-Salgado et al., 2003; Yu et al., 2011). SFKs then

phosphorylate tyrosine residues on Rho GTP exchange factors,

such as leukemia-associated Rho GTPase exchange factor

(LARG, also known as Rho GEF factor 12) or Tiam1 (Elias

et al., 2010; Guilluy et al., 2011), resulting in Rho GTPase

activation and cytoskeletal reorganization (Majumdar et al.,

1999; Nobes and Hall, 1995; Rottner et al., 1999). During

adhesion formation in fibroblasts, RPTP-a also interacts with

avb3 integrins to activate the SFK Fyn (Kostic and Sheetz, 2006;

Su et al., 1999; von Wichert et al., 2003). Although the GTP

exchange factors (GEFs) remain unidentified, two Rho GTPases

(Rac1 and Cdc42) are activated downstream of RPTP-a, and they

are known to regulate actin dynamics and focal adhesion turnover

during fibroblast spreading (Arthur and Burridge, 2001; Herrera

Abreu et al., 2008; Jones and Katan, 2007; ten Klooster et al.,

2006). Rho GTPases can also regulate the region of activity of

other Rho GTPases to determine cell morphology (Vega et al.,

2011), activate the ARP2/3 complex (Welch, 1999) or directly

activate formins. In addition, formins can pass on mechanical

signals to the nucleus to induce expression of genes that are under

the control of the serum response element (Young and Copeland,

2010). In some, if not most, of these cases, there appears to be

crosstalk between different small G proteins (Machacek et al.,

2009).

FAK

Initially identified as a Src kinase substrate (Schaller et al., 1992;

Weiner et al., 1993), the tyrosine kinase FAK is a pivotal

signaling molecule for focal adhesion dynamics, and acts as both

a direct mechanosensor, as discussed above, and downstream of

mechanical signaling events (Schober et al., 2007; Wang et al.,

2001). FAK activation is initiated by autophosphorylation at

tyrosine 397, which can be induced by integrin clustering or

binding of tyrosine-phosphorylated peptides (Chen and Chen,

2006; Sieg et al., 1999; Tomar and Schlaepfer, 2009). FAK can

also activate integrins, resulting in adhesion strengthening

(Michael et al., 2009). The activated FAK alone – or in

complex with Src – can then phosphorylate target proteins or

act as scaffold to recruit additional SH2-binding proteins to

adhesions. Central to its function as regulator of adhesion

dynamics, FAK phosphorylates both GEFs and GAPs to

modulate the activity of Rho GTPases (Fig. 3) (see Tomar and

Schlaepfer, 2009). FAK also phosphorylates paxillin, thereby

enhancing focal adhesion turnover (Zaidel-Bar et al., 2007b), and

binding of FAK in a complex with Src to p130Cas results in Cas

activation (Fonseca et al., 2004).

p130Cas

Cas family proteins are hyperphosphorylated downstream of

FAK and SFKs, in a manner that, at least in the case of p130Cas,

is enhanced by mechanical stretching (Cary et al., 1998; Sawada

et al., 2006; Tamada et al., 2004). During cell migration,

phosphorylation of p130Cas at tyrosine 249 acts as a switch for

the formation of a complex with Crk and the Rac1 GEF

DOCK180 (also known as DOCK1). This complex regulates

Rac1 activity and cytoskeletal dynamics (Brugnera et al., 2002;

Gustavsson et al., 2004; Webb et al., 2004). Additionally,

p130Cas associates with the GEF AND34 (also known as

BCAR3) and regulates the activity of the small GTPases Ral,

Journal of Cell Science 125 (13)3032

Journ

alof

Cell

Scie

nce

Rap1 and R-Ras in cell motility (Gotoh et al., 2000; Riggins et al.,

2003).

Other post-translational modifications

Posttranslational modifications other than phosphorylation are

relevant for many cytoskeletal and adhesion proteins, and could

be involved in mechanosensing. Acetylation of a-tubulin

stabilizes microtubules, which can regulate vesicular

trafficking, autophagy or cell motility (Hubbert et al., 2002;

Perdiz et al., 2011; Tran et al., 2007). The addition of arginyl

from tRNA onto the N-terminus of proteins has been detected in

vivo on cytoskeletal proteins including a- and b-actin, myosin

and tubulin (Karakozova et al., 2006; Wong et al., 2007). In actin,

arginylation changes the interaction dynamics with several

binding partners and alters the polymerization properties of the

proteins (Karakozova et al., 2006; Saha et al., 2010).

Modification of proteins with fatty acid or prenyl residues

assists localization to cellular structures and is found on

cytoskeletal and adhesion proteins, such as vinculin or several

members of the formin family (Han et al., 2009; Harris et al.,

2010; Kellie and Wigglesworth, 1987; Resh, 2006).

Nucleocytoplasmic O-GlcNAcylation (i.e. addition of O-

linked N-acetylglucosamine) is especially interesting owing

to its extensive crosstalk with phosphorylation, as both

modifications frequently compete for the same serine or

threonine residues (Wang et al., 2007; Wang et al., 2008;

Wang et al., 2010). The differences in size and charge between

these modifications can alter protein function and stability (Chen

et al., 2006; Liang et al., 2006). Several adhesion and actin-

binding proteins, such as talin, actin-capping protein, actin-

binding LIM proteins (Chalkley et al., 2009; Hagmann et al.,

1992; Wang et al., 2010; Zhu et al., 2001) and striated muscle

isoforms of actin and myosin (Hedou et al., 2009; Ramirez-

Correa et al., 2008) are subjected to O-GlcNAcylation, which

might modulate properties of the cytoskeleton, or alter muscle

contractility (Cieniewski-Bernard et al., 2009). Increased O-

GlcNAcylation is a cellular response to various stress-inducing

agents, whereas heat-shock proteins can associate with the glycan

to protect the modified proteins (Lefebvre et al., 2001; Zachara

et al., 2004). O-GlcNAcylation of adhesion and cytoskeletal

proteins could therefore also constitute a protective mechanism

against tensile stress. Post-translational modifications such as

Rac1GTP

Rac1GDP

β-PIXP

PPPPPPp130Cas

RapGDP

RapGTP

FAK

P

P

P

Crk

DO

CK1

80Src

RhoAGTP

RhoAGDP

P

P

PP

RhoG

EF

p190

membrane protrusion

FAK

P

P

P

Rac1GTP

Formin activation

βα

Rac1GDP

FH2

Stre

tched

talin

Talin

P

PTiam1

PFyn

RPTP-αIntegrins

WAVE

ARP2/3 activation

ARP2/3

Formins

FH

2

Talin

Stretched talin

Vinculin

Myosin II

Adhesion turnover

(i) Attachment/cluster formation

(ii) Rac1 activation(iii) Actin assembly

(iv) Cluster Growth (v) Slip bond release

(vi) β-PIX activation(vii) Cas activation

(viii) Cell spreading/(ix) Adhesion maturation

(x) p190 RhoGEF activation

(xii) Cross regulation (xi) Stress fiber formation

MyosinActin

Fig. 3. Schematic illustration of the signaling processes downstream of force dectection. The upper panel shows RPTP-a-, Fyn- and talin-dependent signaling

events. (i) Cell attachment, integrin clustering and deformation of the extracellular matrix result in RPTP-a activation. This allows the SFK Fyn to access the

catalytic site. Fyn is dephosphorylated at tyrosine 531 and switches to the open conformation. (ii) Fyn subsequently phosphorylates Rac1 GEFs,

such as Tiam1, that in turn activate Rac1. (iii) Rac1 activity induces actin polymerization from integrin adhesion sites by formin family proteins and/or the

ARP2/3 complex. (iv) Rearward actin flow driven by myosin II contractility stretches talin to expose cryptic vinculin-binding sites, which leads to adhesion

strengthening and cluster growth. (v) When high forces are generated by actin rearward flow and contractility, the bond between actin and talin slips and talin

relaxes. The lower panel depicts FAK- and p130Cas-dependent downstream signaling. (vi) Downstream of integrin clustering, FAK is autophosphorylated at

tyrosine 397. Phosphorylated FAK binds to talin and phosphorylates the Rac1 GEF b-PIX (encoded by ARHGEF7) (Karasawa et al., 1982). (vii) FAK also forms a

complex with Src to activate p130Cas. p130Cas is hyperphosphorylated and binds to Crk and DOCK180 to activate small Rho GTPases, such as Rap. (viii) Rac1

induces actin polymerization and cell protrusion, whereas Rap enhances adhesion maturation (ix). (x) Additionally, FAK can activate p190GEF (Lim et al., 2008b)

to enhance RhoA activity, stress fiber formation (xi) and adhesion turnover. (xii) Cross regulation can alter the region of activity of Rho GTPases.

Exploring mechanical molecular pathways 3033

Journ

alof

Cell

Scie

nce

O-GlcNAcylation can thus be expected to receive furtherattention, particularly as its deregulation has been correlatedwith the occurrence of cancer metastasis and invasiveness (Guet al., 2010; Perdiz et al., 2011).

ConclusionsDespite many recent advances, we still lack a clear picture of themolecular mechanical pathways that transmit and detect forcesin adhesions, upstream of biochemical signaling. Simple

biophysical considerations regarding molecular stiffness andbinding forces (Table 1) can help us to identify probablemechanosensors in soft compared with stiff environments, and

separate them from molecules that simply transmit forces.Nevertheless, a systematic analysis of adhesion protein bindingand unfolding under physiological pulling rates is required to clarifywhere and when bonds break and proteins unfold in the network of

links that are connected in series from the ECM to actin, and inparallel in integrin clusters. This seems even more pressing afterconsidering that common pulling rates in single-molecule

experiments (102–104 nm/second, Table 1) are much higher thanphysiological rates (1–16102 nm/second). Such an analysis mightunravel simple but elegant biophysical mechanisms; for instance,

regulation of the rearward speed of cytoskeletal components and theloading rates might trigger different mechanotransduction cascadesby changing the relative resistance to unbinding and unfolding of

different proteins. Differences in loading rates and force levelscould also help to explain the different behaviors characteristic offast events, such as initial adhesion formation, as compared toslower tissue or ECM remodeling. Only when this biophysical

picture is complete, will we be able to reliably and mechanisticallyexplain how the biophysical inputs (i.e. forces) control thebiological outputs of biochemical signaling and effector pathways

that comprise cell behaviors.

AcknowledgementsThe authors thank Simon Moore for helpful discussions.

FundingThe work of our laboratory was supported in part by a grant by theSpanish Ministry of Economy and Competitiveness [grant numberBFU2011-23111].

ReferencesAgnihotri, A., Soman, P. and Siedlecki, C. A. (2009). AFM measurements of

interactions between the platelet integrin receptor GPIIbIIIa and fibrinogen. Colloids

Surf. B Biointerfaces 71, 138-147.

Alonso, M. C., Drummond, D. R., Kain, S., Hoeng, J., Amos, L. and Cross, R. A.(2007). An ATP gate controls tubulin binding by the tethered head of kinesin-1.Science 316, 120-123.

Arias-Salgado, E. G., Lizano, S., Sarkar, S., Brugge, J. S., Ginsberg, M. H. andShattil, S. J. (2003). Src kinase activation by direct interaction with the integrin betacytoplasmic domain. Proc. Natl. Acad. Sci. USA 100, 13298-13302.

Arias-Salgado, E. G., Lizano, S., Shattil, S. J. and Ginsberg, M. H. (2005).Specification of the direction of adhesive signaling by the integrin beta cytoplasmicdomain. J. Biol. Chem. 280, 29699-29707.

Arnold, M., Hirschfeld-Warneken, V. C., Lohmuller, T., Heil, P., Blummel, J.,Cavalcanti-Adam, E. A., Lopez-Garcıa, M., Walther, P., Kessler, H., Geiger, B.

et al. (2008). Induction of cell polarization and migration by a gradient of nanoscalevariations in adhesive ligand spacing. Nano Lett. 8, 2063-2069.

Arthur, W. T. and Burridge, K. (2001). RhoA inactivation by p190RhoGAP regulatescell spreading and migration by promoting membrane protrusion and polarity. Mol.

Biol. Cell 12, 2711-2720.Averett, L. E., Schoenfisch, M. H., Akhremitchev, B. B. and Gorkun, O. V. (2009).

Kinetics of the multistep rupture of fibrin ‘A-a’ polymerization interactions measuredusing atomic force microscopy. Biophys. J. 97, 2820-2828.

Balaban, N. Q., Schwarz, U. S., Riveline, D., Goichberg, P., Tzur, G., Sabanay, I.,

Mahalu, D., Safran, S., Bershadsky, A., Addadi, L. et al. (2001). Force and focaladhesion assembly: a close relationship studied using elastic micropatternedsubstrates. Nat. Cell Biol. 3, 466-472.

Baneyx, G., Baugh, L. and Vogel, V. (2001). Coexisting conformations of fibronectinin cell culture imaged using fluorescence resonance energy transfer. Proc. Natl. Acad.

Sci. USA 98, 14464-14468.

Baneyx, G., Baugh, L. and Vogel, V. (2002). Fibronectin extension and unfoldingwithin cell matrix fibrils controlled by cytoskeletal tension. Proc. Natl. Acad. Sci.

USA 99, 5139-5143.

Bell, G. I. (1978). Models for the specific adhesion of cells to cells. Science 200, 618-627.

Bershadsky, A. D., Balaban, N. Q. and Geiger, B. (2003). Adhesion-dependent cellmechanosensitivity. Annu. Rev. Cell Dev. Biol. 19, 677-695.

Bouaouina, M., Lad, Y. and Calderwood, D. A. (2008). The N-terminal domains oftalin cooperate with the phosphotyrosine binding-like domain to activate beta1 andbeta3 integrins. J. Biol. Chem. 283, 6118-6125.

Brown, A. E., Litvinov, R. I., Discher, D. E. and Weisel, J. W. (2007). Forcedunfolding of coiled-coils in fibrinogen by single-molecule AFM. Biophys. J. 92, L39-L41.

Brown, A. E., Litvinov, R. I., Discher, D. E., Purohit, P. K. and Weisel, J. W. (2009).Multiscale mechanics of fibrin polymer: gel stretching with protein unfolding and lossof water. Science 325, 741-744.

Brown, C. M., Hebert, B., Kolin, D. L., Zareno, J., Whitmore, L., Horwitz, A. R.

and Wiseman, P. W. (2006). Probing the integrin-actin linkage using high-resolutionprotein velocity mapping. J. Cell Sci. 119, 5204-5214.

Brugnera, E., Haney, L., Grimsley, C., Lu, M., Walk, S. F., Tosello-Trampont,

A. C., Macara, I. G., Madhani, H., Fink, G. R. and Ravichandran, K. S. (2002).Unconventional Rac-GEF activity is mediated through the Dock180-ELMO complex.Nat. Cell Biol. 4, 574-582.

Burridge, K. and Mangeat, P. (1984). An interaction between vinculin and talin.Nature 308, 744-746.

Bustamante, C., Chemla, Y. R., Forde, N. R. and Izhaky, D. (2004). Mechanicalprocesses in biochemistry. Annu. Rev. Biochem. 73, 705-748.

Byfield, F. J., Wen, Q., Levental, I., Nordstrom, K., Arratia, P. E., Miller, R. T. andJanmey, P. A. (2009). Absence of filamin A prevents cells from responding tostiffness gradients on gels coated with collagen but not fibronectin. Biophys. J. 96,5095-5102.

Cai, Y. F., Biais, N., Giannone, G., Tanase, M., Jiang, G. Y., Hofman, J. M.,

Wiggins, C. H., Silberzan, P., Buguin, A., Ladoux, B. et al. (2006). Nonmusclemyosin IIA-dependent force inhibits cell spreading and drives F-actin flow. Biophys.

J. 91, 3907-3920.

Calderwood, D. A., Zent, R., Grant, R., Rees, D. J., Hynes, R. O. and Ginsberg,M. H. (1999). The Talin head domain binds to integrin beta subunit cytoplasmic tailsand regulates integrin activation. J. Biol. Chem. 274, 28071-28074.

Calderwood, D. A., Fujioka, Y., de Pereda, J. M., Garcıa-Alvarez, B., Nakamoto, T.,Margolis, B., McGlade, C. J., Liddington, R. C. and Ginsberg, M. H. (2003).Integrin beta cytoplasmic domain interactions with phosphotyrosine-binding domains:a structural prototype for diversity in integrin signaling. Proc. Natl. Acad. Sci. USA

100, 2272-2277.

Cary, L. A., Han, D. C., Polte, T. R., Hanks, S. K. and Guan, J. L. (1998).Identification of p130Cas as a mediator of focal adhesion kinase-promoted cellmigration. J. Cell Biol. 140, 211-221.

Chalkley, R. J., Thalhammer, A., Schoepfer, R. and Burlingame, A. L. (2009).Identification of protein O-GlcNAcylation sites using electron transfer dissociationmass spectrometry on native peptides. Proc. Natl. Acad. Sci. USA 106, 8894-8899.

Chan, C. E. and Odde, D. J. (2008). Traction dynamics of filopodia on compliantsubstrates. Science 322, 1687-1691.

Chao, J. T., Gui, P., Zamponi, G. W., Davis, G. E. and Davis, M. J. (2011). Spatialassociation of the Cav1.2 calcium channel with alpha5beta1-integrin. Am. J. Physiol.

Cell Physiol. 300, C477-C489.

Chen, H., Fu, H., Zhu, X., Cong, P., Nakamura, F. and Yan, J. (2011). Improvedhigh-force magnetic tweezers for stretching and refolding of proteins and short DNA.Biophys. J. 100, 517-523.

Chen, H. C., Appeddu, P. A., Parsons, J. T., Hildebrand, J. D., Schaller, M. D. and

Guan, J. L. (1995). Interaction of focal adhesion kinase with cytoskeletal proteintalin. J. Biol. Chem. 270, 16995-16999.

Chen, H. S., Kolahi, K. S. and Mofrad, M. R. (2009). Phosphorylation facilitates theintegrin binding of filamin under force. Biophys. J. 97, 3095-3104.

Chen, S. Y. and Chen, H. C. (2006). Direct interaction of focal adhesion kinase (FAK)with Met is required for FAK to promote hepatocyte growth factor-induced cellinvasion. Mol. Cell. Biol. 26, 5155-5167.

Chen, Y. X., Du, J. T., Zhou, L. X., Liu, X. H., Zhao, Y. F., Nakanishi, H. and Li,

Y. M. (2006). Alternative O-GlcNAcylation/O-phosphorylation of Ser16 inducedifferent conformational disturbances to the N terminus of murine estrogen receptorbeta. Chem. Biol. 13, 937-944.

Chesarone, M. A. and Goode, B. L. (2009). Actin nucleation and elongation factors:mechanisms and interplay. Curr. Opin. Cell Biol. 21, 28-37.

Chi, R. J., Olenych, S. G., Kim, K. and Keller, T. C., 3rd (2005). Smooth musclealpha-actinin interaction with smitin. Int. J. Biochem. Cell Biol. 37, 1470-1482.

Choi, C. K., Vicente-Manzanares, M., Zareno, J., Whitmore, L. A., Mogilner, A.

and Horwitz, A. R. (2008). Actin and alpha-actinin orchestrate the assembly andmaturation of nascent adhesions in a myosin II motor-independent manner. Nat. Cell

Biol. 10, 1039-1050.

Cieniewski-Bernard, C., Montel, V., Stevens, L. and Bastide, B. (2009). O-GlcNAcylation, an original modulator of contractile activity in striated muscle.J. Muscle Res. Cell Motil. 30, 281-287.

Journal of Cell Science 125 (13)3034

Journ

alof

Cell

Scie

nce

Cooper, L. A., Shen, T. L. and Guan, J. L. (2003). Regulation of focal adhesion kinaseby its amino-terminal domain through an autoinhibitory interaction. Mol. Cell. Biol.

23, 8030-8041.

Costa, P. and Parsons, M. (2010). New insights into the dynamics of cell adhesions. Int

Rev Cell Mol Biol 283, 57-91.

Critchley, D. R. (2009). Biochemical and structural properties of the integrin-associatedcytoskeletal protein talin. Annu Rev Biophys 38, 235-254.

Cunningham, C. C., Gorlin, J. B., Kwiatkowski, D. J., Hartwig, J. H., Janmey, P. A.,

Byers, H. R. and Stossel, T. P. (1992). Actin-binding protein requirement for corticalstability and efficient locomotion. Science 255, 325-327.

Davis, G. E. (1992). Affinity of integrins for damaged extracellular matrix: alpha v beta3 binds to denatured collagen type I through RGD sites. Biochem. Biophys. Res.

Commun. 182, 1025-1031.

Davis, G. E., Bayless, K. J., Davis, M. J. and Meininger, G. A. (2000). Regulation oftissue injury responses by the exposure of matricryptic sites within extracellularmatrix molecules. Am. J. Pathol. 156, 1489-1498.

del Rio, A., Perez-Jimenez, R., Liu, R. C., Roca-Cusachs, P., Fernandez, J. M. and

Sheetz, M. P. (2009). Stretching single talin rod molecules activates vinculin binding.Science 323, 638-641.

Deng, Z. J., Liang, M., Monteiro, M., Toth, I. and Minchin, R. F. (2011).Nanoparticle-induced unfolding of fibrinogen promotes Mac-1 receptor activationand inflammation. Nat. Nanotechnol. 6, 39-44.

Diez, G., Auernheimer, V., Fabry, B. and Goldmann, W. H. (2011). Head/tailinteraction of vinculin influences cell mechanical behavior. Biochem. Biophys. Res.

Commun. 406, 85-88.

Dogterom, M. and Yurke, B. (1997). Measurement of the force-velocity relation forgrowing microtubules. Science 278, 856-860.

du Roure, O., Saez, A., Buguin, A., Austin, R. H., Chavrier, P., Silberzan, P. and

Ladoux, B. (2005). Force mapping in epithelial cell migration. Proc. Natl. Acad. Sci.

USA 102, 2390-2395.

Dubash, A. D., Menold, M. M., Samson, T., Boulter, E., Garcia-Mata, R.,

Doughman, R. and Burridge, K. (2009). Chapter 1. Focal adhesions: new angleson an old structure. Int. Rev. Cell Mol. Biol. 277, 1-65.

Eckes, B., Dogic, D., Colucci-Guyon, E., Wang, N., Maniotis, A., Ingber, D.,Merckling, A., Langa, F., Aumailley, M., Delouvee, A. et al. (1998). Impairedmechanical stability, migration and contractile capacity in vimentin-deficientfibroblasts. J. Cell Sci. 111, 1897-1907.

Ehrlicher, A. J., Nakamura, F., Hartwig, J. H., Weitz, D. A. and Stossel, T. P.(2011). Mechanical strain in actin networks regulates FilGAP and integrin binding tofilamin A. Nature 478, 260-263.

Elias, B. C., Bhattacharya, S., Ray, R. M. and Johnson, L. R. (2010). Polyamine-dependent activation of Rac1 is stimulated by focal adhesion-mediated Tiam1activation. Cell Adhes. Migr. 4, 419-430.

Engler, A. J., Sen, S., Sweeney, H. L. and Discher, D. E. (2006). Matrix elasticitydirects stem cell lineage specification. Cell 126, 677-689.

Evans, E. and Ritchie, K. (1999). Strength of a weak bond connecting flexible polymerchains. Biophys. J. 76, 2439-2447.

Fenteany, G., Janmey, P. A. and Stossel, T. P. (2000). Signaling pathways and cellmechanics involved in wound closure by epithelial cell sheets. Curr. Biol. 10, 831-838.

Ferrer, J. M., Lee, H., Chen, J., Pelz, B., Nakamura, F., Kamm, R. D. and Lang,

M. J. (2008). Measuring molecular rupture forces between single actin filaments andactin-binding proteins. Proc. Natl. Acad. Sci. USA 105, 9221-9226.

Finer, J. T., Simmons, R. M. and Spudich, J. A. (1994). Single myosin moleculemechanics: piconewton forces and nanometre steps. Nature 368, 113-119.

Fisher, G. W., Conrad, P. A., DeBiasio, R. L. and Taylor, D. L. (1988). Centripetaltransport of cytoplasm, actin, and the cell surface in lamellipodia of fibroblasts. Cell

Motil. Cytoskeleton 11, 235-247.

Flanagan, L. A., Chou, J., Falet, H., Neujahr, R., Hartwig, J. H. and Stossel, T. P.

(2001). Filamin A, the Arp2/3 complex, and the morphology and function of corticalactin filaments in human melanoma cells. J. Cell Biol. 155, 511-518.

Fonseca, P. M., Shin, N. Y., Brabek, J., Ryzhova, L., Wu, J. and Hanks, S. K. (2004).Regulation and localization of CAS substrate domain tyrosine phosphorylation. Cell.

Signal. 16, 621-629.

Forscher, P. and Smith, S. J. (1988). Actions of cytochalasins on the organization ofactin filaments and microtubules in a neuronal growth cone. J. Cell Biol. 107, 1505-1516.

Friedland, J. C., Lee, M. H. and Boettiger, D. (2009). Mechanically activated integrinswitch controls alpha5beta1 function. Science 323, 642-644.

Furuike, S., Ito, T. and Yamazaki, M. (2001). Mechanical unfolding of single filaminA (ABP-280) molecules detected by atomic force microscopy. FEBS Lett. 498, 72-75.

Gardel, M. L., Sabass, B., Ji, L., Danuser, G., Schwarz, U. S. and Waterman, C. M.

(2008). Traction stress in focal adhesions correlates biphasically with actin retrogradeflow speed. J. Cell Biol. 183, 999-1005.

Gautel, M. (2011). Cytoskeletal protein kinases: titin and its relations in mechanosen-sing. Pflugers Arch. 462, 119-134.

Gauthier, N. C., Fardin, M. A., Roca-Cusachs, P. and Sheetz, M. P. (2011).Temporary increase in plasma membrane tension coordinates the activation ofexocytosis and contraction during cell spreading. Proc. Natl. Acad. Sci. USA 108,14467-14472.

Ghassemi, S., Meacci, G., Liu, S., Gondarenko, A. A., Mathur, A., Roca-Cusachs,

P., Sheetz, M. P. and Hone, J. (2012). Cells test substrate rigidity by localcontractions on sub-micrometer pillars. Proc. Natl. Acad. Sci. USA 109, 5328-5333.

Giannone, G., Dubin-Thaler, B. J., Dobereiner, H. G., Kieffer, N., Bresnick, A. R.

and Sheetz, M. P. (2004). Periodic lamellipodial contractions correlate with rearward

actin waves. Cell 116, 431-443.

Gorfinkiel, N., Blanchard, G. B., Adams, R. J. and Martinez Arias, A. (2009).Mechanical control of global cell behaviour during dorsal closure in Drosophila.

Development 136, 1889-1898.

Gotoh, T., Cai, D., Tian, X., Feig, L. A. and Lerner, A. (2000). p130Cas regulates theactivity of AND-34, a novel Ral, Rap1, and R-Ras guanine nucleotide exchange

factor. J. Biol. Chem. 275, 30118-30123.

Graham, J. S., Vomund, A. N., Phillips, C. L. and Grandbois, M. (2004). Structural

changes in human type I collagen fibrils investigated by force spectroscopy. Exp. Cell

Res. 299, 335-342.

Grant, R. P., Spitzfaden, C., Altroff, H., Campbell, I. D. and Mardon, H. J. (1997).

Structural requirements for biological activity of the ninth and tenth FIII domains ofhuman fibronectin. J. Biol. Chem. 272, 6159-6166.

Grashoff, C., Hoffman, B. D., Brenner, M. D., Zhou, R., Parsons, M., Yang, M. T.,

McLean, M. A., Sligar, S. G., Chen, C. S., Ha, T. et al. (2010). Measuringmechanical tension across vinculin reveals regulation of focal adhesion dynamics.Nature 466, 263-266.

Gu, Y., Mi, W., Ge, Y., Liu, H., Fan, Q., Han, C., Yang, J., Han, F., Lu, X. and Yu,

W. (2010). GlcNAcylation plays an essential role in breast cancer metastasis. Cancer

Res. 70, 6344-6351.

Guilluy, C., Swaminathan, V., Garcia-Mata, R., O’Brien, E. T., Superfine, R. and

Burridge, K. (2011). The Rho GEFs LARG and GEF-H1 regulate the mechanicalresponse to force on integrins. Nat. Cell Biol. 13, 722-727.

Guo, B. and Guilford, W. H. (2006). Mechanics of actomyosin bonds in differentnucleotide states are tuned to muscle contraction. Proc. Natl. Acad. Sci. USA 103,9844-9849.

Gupton, S. L., Eisenmann, K., Alberts, A. S. and Waterman-Storer, C. M. (2007).mDia2 regulates actin and focal adhesion dynamics and organization in the lamellafor efficient epithelial cell migration. J. Cell Sci. 120, 3475-3487.

Gustavsson, A., Yuan, M. and Fallman, M. (2004). Temporal dissection of beta1-integrin signaling indicates a role for p130Cas-Crk in filopodia formation. J. Biol.

Chem. 279, 22893-22901.

Hagmann, J., Grob, M. and Burger, M. M. (1992). The cytoskeletal protein talin is O-glycosylated. J. Biol. Chem. 267, 14424-14428.

Han, Y., Eppinger, E., Schuster, I. G., Weigand, L. U., Liang, X., Kremmer, E.,

Peschel, C. and Krackhardt, A. M. (2009). Formin-like 1 (FMNL1) is regulated byN-terminal myristoylation and induces polarized membrane blebbing. J. Biol. Chem.

284, 33409-33417.

Harada, Y., Sakurada, K., Aoki, T., Thomas, D. D. and Yanagida, T. (1990).Mechanochemical coupling in actomyosin energy transduction studied by in vitromovement assay. J. Mol. Biol. 216, 49-68.

Harris, E. S., Gauvin, T. J., Heimsath, E. G. and Higgs, H. N. (2010). Assembly offilopodia by the formin FRL2 (FMNL3). Cytoskeleton (Hoboken) 67, 755-772.

Hartwig, J. H. and Stossel, T. P. (1975). Isolation and properties of actin, myosin, and anew actinbinding protein in rabbit alveolar macrophages. J. Biol. Chem. 250, 5696-5705.

Hartwig, J. H. and Shevlin, P. (1986). The architecture of actin filaments and theultrastructural location of actin-binding protein in the periphery of lung macrophages.J. Cell Biol. 103, 1007-1020.

Hatch, V., Zhi, G., Smith, L., Stull, J. T., Craig, R. and Lehman, W. (2001). Myosinlight chain kinase binding to a unique site on F-actin revealed by three-dimensionalimage reconstruction. J. Cell Biol. 154, 611-618.

Hedou, J., Bastide, B., Page, A., Michalski, J. C. and Morelle, W. (2009). Mapping ofO-linked beta-N-acetylglucosamine modification sites in key contractile proteins ofrat skeletal muscle. Proteomics 9, 2139-2148.

Heinemann, F., Doschke, H. and Radmacher, M. (2011). Keratocyte lamellipodialprotrusion is characterized by a concave force-velocity relation. Biophys. J. 100,1420-1427.

Herrera Abreu, M. T., Penton, P. C., Kwok, V., Vachon, E., Shalloway, D., Vidali,

L., Lee, W., McCulloch, C. A. and Downey, G. P. (2008). Tyrosine phosphatasePTPalpha regulates focal adhesion remodeling through Rac1 activation. Am. J.

Physiol. Cell Physiol. 294, C931-C944.

Hildebrand, J. D., Schaller, M. D. and Parsons, J. T. (1995). Paxillin, a tyrosinephosphorylated focal adhesion-associated protein binds to the carboxyl terminal

domain of focal adhesion kinase. Mol. Biol. Cell 6, 637-647.

Hirokawa, N., Noda, Y., Tanaka, Y. and Niwa, S. (2009). Kinesin superfamily motorproteins and intracellular transport. Nat. Rev. Mol. Cell Biol. 10, 682-696.

Hobert, O., Moerman, D. G., Clark, K. A., Beckerle, M. C. and Ruvkun, G. (1999).A conserved LIM protein that affects muscular adherens junction integrity andmechanosensory function in Caenorhabditis elegans. J. Cell Biol. 144, 45-57.

Hocking, D. C. and Kowalski, K. (2002). A cryptic fragment from fibronectin’s III1module localizes to lipid rafts and stimulates cell growth and contractility. J. Cell

Biol. 158, 175-184.

Hotulainen, P. and Lappalainen, P. (2006). Stress fibers are generated by two distinctactin assembly mechanisms in motile cells. J. Cell Biol. 173, 383-394.

Hu, K., Ji, L., Applegate, K. T., Danuser, G. and Waterman-Storer, C. M. (2007).Differential transmission of actin motion within focal adhesions. Science 315, 111-115.

Hu, S. H., Chen, J. X., Fabry, B., Numaguchi, Y., Gouldstone, A., Ingber, D. E.,

Fredberg, J. J., Butler, J. P. and Wang, N. (2003). Intracellular stress tomography

Exploring mechanical molecular pathways 3035

Journ

alof

Cell

Scie

nce

reveals stress focusing and structural anisotropy in cytoskeleton of living cells. Am. J.

Physiol. Cell Physiol. 285, C1082-C1090.

Hubbert, C., Guardiola, A., Shao, R., Kawaguchi, Y., Ito, A., Nixon, A., Yoshida,

M., Wang, X. F. and Yao, T. P. (2002). HDAC6 is a microtubule-associateddeacetylase. Nature 417, 455-458.

Hynes, R. O. (2002). Integrins: bidirectional, allosteric signaling machines. Cell 110,673-687.

Ishijima, A., Harada, Y., Kojima, H., Funatsu, T., Higuchi, H. and Yanagida, T.

(1994). Single-molecule analysis of the actomyosin motor using nano-manipulation.Biochem. Biophys. Res. Commun. 199, 1057-1063.

Ivaska, J., Vuoriluoto, K., Huovinen, T., Izawa, I., Inagaki, M. and Parker, P. J.

(2005). PKCepsilon-mediated phosphorylation of vimentin controls integrin recycling

and motility. EMBO J. 24, 3834-3845.

Janmey, P. A., Winer, J. P., Murray, M. E. and Wen, Q. (2009). The hard life of softcells. Cell Motil. Cytoskeleton 66, 597-605.

Jiang, G., den Hertog, J. and Hunter, T. (2000). Receptor-like protein tyrosinephosphatase alpha homodimerizes on the cell surface. Mol. Cell. Biol. 20, 5917-5929.

Jiang, G. Y., Giannone, G., Critchley, D. R., Fukumoto, E. and Sheetz, M. P. (2003).Two-piconewton slip bond between fibronectin and the cytoskeleton depends on talin.Nature 424, 334-337.

Jiang, G. Y., Huang, A. H., Cai, Y. F., Tanase, M. and Sheetz, M. P. (2006). Rigiditysensing at the leading edge through alphavbeta3 integrins and RPTPalpha. Biophys. J.

90, 1804-1809.

Johnson, C. P., Tang, H. Y., Carag, C., Speicher, D. W. and Discher, D. E. (2007).Forced unfolding of proteins within cells. Science 317, 663-666.

Jones, N. P. and Katan, M. (2007). Role of phospholipase Cgamma1 in cell spreadingrequires association with a beta-Pix/GIT1-containing complex, leading to activationof Cdc42 and Rac1. Mol. Cell. Biol. 27, 5790-5805.

Kanchanawong, P., Shtengel, G., Pasapera, A. M., Ramko, E. B., Davidson, M. W.,

Hess, H. F. and Waterman, C. M. (2010). Nanoscale architecture of integrin-basedcell adhesions. Nature 468, 580-584.

Karakozova, M., Kozak, M., Wong, C. C., Bailey, A. O., Yates, J. R., 3rd, Mogilner,

A., Zebroski, H. and Kashina, A. (2006). Arginylation of beta-actin regulates actincytoskeleton and cell motility. Science 313, 192-196.

Karasawa, J., Kuriyama, Y., Kuro, M., Kikuchi, H., Sawada, T. and Mitsugi, T.

(1982). [Monitoring system of cerebral blood flow and cerebral metabolism. Part II.Relationship between internal jugular O2 tention and cerebral blood flow (author’stransl)]. No To Shinkei 34, 239-245.

Kaverina, I., Rottner, K. and Small, J. V. (1998). Targeting, capture, and stabilizationof microtubules at early focal adhesions. J. Cell Biol. 142, 181-190.

Kellie, S. and Wigglesworth, N. M. (1987). The cytoskeletal protein vinculin isacylated by myristic acid. FEBS Lett. 213, 428-432.

Kiema, T., Lad, Y., Jiang, P. J., Oxley, C. L., Baldassarre, M., Wegener, K. L.,

Campbell, I. D., Ylanne, J. and Calderwood, D. A. (2006). The molecular basis offilamin binding to integrins and competition with talin. Mol. Cell 21, 337-347.

Kim, H., Nakamura, F., Lee, W., Hong, C., Perez-Sala, D. and McCulloch, C. A.

(2010). Regulation of cell adhesion to collagen via beta1 integrins is dependent oninteractions of filamin A with vimentin and protein kinase C epsilon. Exp. Cell Res.

316, 1829-1844.

Kim, M., Carman, C. V. and Springer, T. A. (2003). Bidirectional transmembranesignaling by cytoplasmic domain separation in integrins. Science 301, 1720-1725.

Klotzsch, E., Smith, M. L., Kubow, K. E., Muntwyler, S., Little, W. C., Beyeler, F.,

Gourdon, D., Nelson, B. J. and Vogel, V. (2009). Fibronectin forms the mostextensible biological fibers displaying switchable force-exposed cryptic binding sites.

Proc. Natl. Acad. Sci. USA 106, 18267-18272.

Kolodney, M. S. and Elson, E. L. (1995). Contraction due to microtubule disruption isassociated with increased phosphorylation of myosin regulatory light chain. Proc.

Natl. Acad. Sci. USA 92, 10252-10256.

Kong, F., Garcıa, A. J., Mould, A. P., Humphries, M. J. and Zhu, C. (2009).Demonstration of catch bonds between an integrin and its ligand. J. Cell Biol. 185,1275-1284.

Kostic, A. and Sheetz, M. P. (2006). Fibronectin rigidity response through Fyn andp130Cas recruitment to the leading edge. Mol. Biol. Cell 17, 2684-2695.

Kovar, D. R. and Pollard, T. D. (2004). Insertional assembly of actin filament barbedends in association with formins produces piconewton forces. Proc. Natl. Acad. Sci.

USA 101, 14725-14730.

Kovar, D. R., Harris, E. S., Mahaffy, R., Higgs, H. N. and Pollard, T. D. (2006).Control of the assembly of ATP- and ADP-actin by formins and profilin. Cell 124,

423-435.

Kreis, S., Schonfeld, H. J., Melchior, C., Steiner, B. and Kieffer, N. (2005). Theintermediate filament protein vimentin binds specifically to a recombinant integrinalpha2/beta1 cytoplasmic tail complex and co-localizes with native alpha2/beta1 in

endothelial cell focal adhesions. Exp. Cell Res. 305, 110-121.

Kress, H., Stelzer, E. H., Holzer, D., Buss, F., Griffiths, G. and Rohrbach, A. (2007).Filopodia act as phagocytic tentacles and pull with discrete steps and a load-dependent velocity. Proc. Natl. Acad. Sci. USA 104, 11633-11638.

Krieger, C. C., An, X., Tang, H. Y., Mohandas, N., Speicher, D. W. and Discher,

D. E. (2011). Cysteine shotgun-mass spectrometry (CS-MS) reveals dynamicsequence of protein structure changes within mutant and stressed cells. Proc. Natl.

Acad. Sci. USA 108, 8269-8274.

Kumar, S. and Weaver, V. M. (2009). Mechanics, malignancy, and metastasis: theforce journey of a tumor cell. Cancer Metastasis Rev. 28, 113-127.

Kumar, S. and Suan Li, M. (2010). Biomolecules under mechanical force. Phys. Rep.

486, 1-74.

Kuo, S. C. and Sheetz, M. P. (1993). Force of single kinesin molecules measured with

optical tweezers. Science 260, 232-234.

Laakso, J. M., Lewis, J. H., Shuman, H. and Ostap, E. M. (2008). Myosin I can act as

a molecular force sensor. Science 321, 133-136.

Lee, I. and Marchant, R. E. (2001). Force measurements on the molecular interactions

between ligand (RGD) and human platelet aIIbb3 receptor system. Surf. Sci. 491, 433-

443.

Lee, H., Pelz, B., Ferrer, J. M., Kim, T., Lang, M. J. and Kamm, R. D. (2009).

Cytoskeletal deformation at high strains and the role of cross-link unfolding or

unbinding. Cell. Mol. Bioeng. 2, 28-38.

Lefebvre, T., Cieniewski, C., Lemoine, J., Guerardel, Y., Leroy, Y., Zanetta, J. P.

and Michalski, J. C. (2001). Identification of N-acetyl-d-glucosamine-specific

lectins from rat liver cytosolic and nuclear compartments as heat-shock proteins.

Biochem. J. 360, 179-188.

Legate, K. R., Wickstrom, S. A. and Fassler, R. (2009). Genetic and cell biological

analysis of integrin outside-in signaling. Genes Dev. 23, 397-418.

Lemmon, C. A., Ohashi, T. and Erickson, H. P. (2011). Probing the folded state of

fibronectin type III domains in stretched fibrils by measuring buried cysteine

accessibility. J. Biol. Chem. 286, 26375-26382.

Li, F. Y., Redick, S. D., Erickson, H. P. and Moy, V. T. (2003a). Force measurements

of the alpha5beta1 integrin-fibronectin interaction. Biophys. J. 84, 1252-1262.

Li, R. H., Mitra, N., Gratkowski, H., Vilaire, G., Litvinov, R., Nagasami, C., Weisel,

J. W., Lear, J. D., DeGrado, W. F. and Bennett, J. S. (2003b). Activation of

integrin alphaIIbbeta3 by modulation of transmembrane helix associations. Science

300, 795-798.

Li, Y., Bhimalapuram, P. and Dinner, A. R. (2010). Model for how retrograde actin

flow regulates adhesion traction stresses. J. Phys. Condens. Matter 22, 194113.

Liang, F. C., Chen, R. P., Lin, C. C., Huang, K. T. and Chan, S. I. (2006). Tuning the

conformation properties of a peptide by glycosylation and phosphorylation. Biochem.

Biophys. Res. Commun. 342, 482-488.

Lim, B. B., Lee, E. H., Sotomayor, M. and Schulten, K. (2008a). Molecular basis of

fibrin clot elasticity. Structure 16, 449-459.

Lim, Y., Lim, S. T., Tomar, A., Gardel, M., Bernard-Trifilo, J. A., Chen, X. L.,

Uryu, S. A., Canete-Soler, R., Zhai, J., Lin, H. et al. (2008b). PyK2 and FAK

connections to p190Rho guanine nucleotide exchange factor regulate RhoA activity,

focal adhesion formation, and cell motility. J. Cell Biol. 180, 187-203.

Litvinov, R. I., Shuman, H., Bennett, J. S. and Weisel, J. W. (2002). Binding strength

and activation state of single fibrinogen-integrin pairs on living cells. Proc. Natl.

Acad. Sci. USA 99, 7426-7431.

Litvinov, R. I., Bennett, J. S., Weisel, J. W. and Shuman, H. (2005). Multi-step

fibrinogen binding to the integrin (alpha)IIb(beta)3 detected using force spectroscopy.

Biophys. J. 89, 2824-2834.

Litvinov, R. I., Barsegov, V., Schissler, A. J., Fisher, A. R., Bennett, J. S., Weisel,

J. W. and Shuman, H. (2011). Dissociation of bimolecular aIIbb3-fibrinogen

complex under a constant tensile force. Biophys. J. 100, 165-173.

Liu, B. P., Chrzanowska-Wodnicka, M. and Burridge, K. (1998). Microtubule

depolymerization induces stress fibers, focal adhesions, and DNA synthesis via the

GTP-binding protein Rho. Cell Adhes. Commun. 5, 249-255.

Lo, S. H. (2004). Tensin. Int. J. Biochem. Cell Biol. 36, 31-34.

Lo, S. H., Janmey, P. A., Hartwig, J. H. and Chen, L. B. (1994). Interactions of tensin

with actin and identification of its three distinct actin-binding domains. J. Cell Biol.

125, 1067-1075.

Loo, D. T., Kanner, S. B. and Aruffo, A. (1998). Filamin binds to the cytoplasmic

domain of the beta1-integrin. Identification of amino acids responsible for this

interaction. J. Biol. Chem. 273, 23304-23312.

Luo, B. H., Carman, C. V. and Springer, T. A. (2007). Structural basis of integrin

regulation and signaling. Annu. Rev. Immunol. 25, 619-647.

Lynch, C. D., Gauthier, N. C., Biais, N., Lazar, A. M., Roca-Cusachs, P., Yu, C. H.

and Sheetz, M. P. (2011). Filamin depletion blocks endoplasmic spreading and

destabilizes force-bearing adhesions. Mol. Biol. Cell 22, 1263-1273.

Machacek, M., Hodgson, L., Welch, C., Elliott, H., Pertz, O., Nalbant, P., Abell, A.,

Johnson, G. L., Hahn, K. M. and Danuser, G. (2009). Coordination of Rho GTPase

activities during cell protrusion. Nature 461, 99-103.

Majumdar, M., Seasholtz, T. M., Buckmaster, C., Toksoz, D. and Brown, J. H.

(1999). A rho exchange factor mediates thrombin and Galpha(12)-induced

cytoskeletal responses. J. Biol. Chem. 274, 26815-26821.

Mammoto, T. and Ingber, D. E. (2010). Mechanical control of tissue and organ

development. Development 137, 1407-1420.

Margadant, F., Chew, L. L., Hu, X., Yu, H., Bate, N., Zhang, X. and Sheetz, M. P.

(2011). Mechanotransduction in vivo by repeated talin stretch-relaxation events

depends upon vinculin. PLoS Biol. 9, e1001223.

Marko, J. F. and Siggia, E. D. (1995). Stretching DNA. Macromolecules 28, 8759-

8770.

Matthews, B. D., Overby, D. R., Mannix, R. and Ingber, D. E. (2006). Cellular

adaptation to mechanical stress: role of integrins, Rho, cytoskeletal tension and

mechanosensitive ion channels. J. Cell Sci. 119, 508-518.

Matthews, B. D., Thodeti, C. K., Tytell, J. D., Mammoto, A., Overby, D. R. and

Ingber, D. E. (2010). Ultra-rapid activation of TRPV4 ion channels by mechanical

forces applied to cell surface beta1 integrins. Integr Biol (Camb) 2, 435-442.

Journal of Cell Science 125 (13)3036

Journ

alof

Cell

Scie

nce

Menkel, A. R., Kroemker, M., Bubeck, P., Ronsiek, M., Nikolai, G. and Jockusch,

B. M. (1994). Characterization of an F-actin-binding domain in the cytoskeletalprotein vinculin. J. Cell Biol. 126, 1231-1240.

Meyhofer, E. and Howard, J. (1995). The force generated by a single kinesin moleculeagainst an elastic load. Proc. Natl. Acad. Sci. USA 92, 574-578.

Michael, K. E., Dumbauld, D. W., Burns, K. L., Hanks, S. K. and Garcıa, A. J.(2009). Focal adhesion kinase modulates cell adhesion strengthening via integrinactivation. Mol. Biol. Cell 20, 2508-2519.

Miyata, H., Yasuda, R. and Kinosita, K., Jr (1996). Strength and lifetime of the bondbetween actin and skeletal muscle alpha-actinin studied with an optical trappingtechnique. Biochim. Biophys. Acta 1290, 83-88.

Moore, S. W., Roca-Cusachs, P. and Sheetz, M. P. (2010). Stretchy proteins onstretchy substrates: the important elements of integrin-mediated rigidity sensing. Dev.

Cell 19, 194-206.

Moser, M., Nieswandt, B., Ussar, S., Pozgajova, M. and Fassler, R. (2008). Kindlin-3is essential for integrin activation and platelet aggregation. Nat. Med. 14, 325-330.

Moser, M., Legate, K. R., Zent, R. and Fassler, R. (2009). The tail of integrins, talin,and kindlins. Science 324, 895-899.

Munevar, S., Wang, Y. L. and Dembo, M. (2004). Regulation of mechanicalinteractions between fibroblasts and the substratum by stretch-activated Ca2+ entry.J. Cell Sci. 117, 85-92.

Na, S., Collin, O., Chowdhury, F., Tay, B., Ouyang, M., Wang, Y. and Wang, N.

(2008). Rapid signal transduction in living cells is a unique feature ofmechanotransduction. Proc. Natl. Acad. Sci. USA 105, 6626-6631.

Nelson, C. M., Jean, R. P., Tan, J. L., Liu, W. F., Sniadecki, N. J., Spector, A. A. and

Chen, C. S. (2005). Emergent patterns of growth controlled by multicellular form andmechanics. Proc. Natl. Acad. Sci. USA 102, 11594-11599.

Nieves, B., Jones, C. W., Ward, R., Ohta, Y., Reverte, C. G. and LaFlamme, S. E.

(2010). The NPIY motif in the integrin beta1 tail dictates the requirement for talin-1in outside-in signaling. J. Cell Sci. 123, 1216-1226.

Niland, S., Westerhausen, C., Schneider, S. W., Eckes, B., Schneider, M. F. and

Eble, J. A. (2011). Biofunctionalization of a generic collagenous triple helix with thea2b1 integrin binding site allows molecular force measurements. Int. J. Biochem. Cell

Biol. 43, 721-731.

Nobes, C. D. and Hall, A. (1995). Rho, rac, and cdc42 GTPases regulate the assemblyof multimolecular focal complexes associated with actin stress fibers, lamellipodia,and filopodia. Cell 81, 53-62.

Oberhauser, A. F., Marszalek, P. E., Erickson, H. P. and Fernandez, J. M. (1998).The molecular elasticity of the extracellular matrix protein tenascin. Nature 393, 181-185.

Oberhauser, A. F., Badilla-Fernandez, C., Carrion-Vazquez, M. and Fernandez,

J. M. (2002). The mechanical hierarchies of fibronectin observed with single-molecule AFM. J. Mol. Biol. 319, 433-447.

Orgel, J. P., Antipova, O., Sagi, I., Bitler, A., Qiu, D., Wang, R., Xu, Y. and SanAntonio, J. D. (2011). Collagen fibril surface displays a constellation of sites capableof promoting fibril assembly, stability, and hemostasis. Connect. Tissue Res. 52, 18-24.

Otey, C. A., Pavalko, F. M. and Burridge, K. (1990). An interaction between alpha-actinin and the beta 1 integrin subunit in vitro. J. Cell Biol. 111, 721-729.

Pankov, R., Cukierman, E., Katz, B. Z., Matsumoto, K., Lin, D. C., Lin, S., Hahn, C.and Yamada, K. M. (2000). Integrin dynamics and matrix assembly: tensin-dependent translocation of alpha(5)beta(1) integrins promotes early fibronectinfibrillogenesis. J. Cell Biol. 148, 1075-1090.

Paramore, S., Ayton, G. S., Mirijanian, D. T. and Voth, G. A. (2006). Extending aspectrin repeat unit. I: linear force-extension response. Biophys. J. 90, 92-100.

Parsons, J. T., Horwitz, A. R. and Schwartz, M. A. (2010). Cell adhesion: integratingcytoskeletal dynamics and cellular tension. Nat. Rev. Mol. Cell Biol. 11, 633-643.

Pasapera, A. M., Schneider, I. C., Rericha, E., Schlaepfer, D. D. and Waterman,C. M. (2010). Myosin II activity regulates vinculin recruitment to focal adhesionsthrough FAK-mediated paxillin phosphorylation. J. Cell Biol. 188, 877-890.

Paszek, M. J., Zahir, N., Johnson, K. R., Lakins, J. N., Rozenberg, G. I., Gefen, A.,

Reinhart-King, C. A., Margulies, S. S., Dembo, M., Boettiger, D. et al. (2005).Tensional homeostasis and the malignant phenotype. Cancer Cell 8, 241-254.

Pavalko, F. M. and Burridge, K. (1991). Disruption of the actin cytoskeleton aftermicroinjection of proteolytic fragments of alpha-actinin. J. Cell Biol. 114, 481-491.

Pavalko, F. M. and LaRoche, S. M. (1993). Activation of human neutrophils inducesan interaction between the integrin beta 2-subunit (CD18) and the actin bindingprotein alpha-actinin. J. Immunol. 151, 3795-3807.

Pavalko, F. M., Otey, C. A., Simon, K. O. and Burridge, K. (1991). Alpha-actinin: adirect link between actin and integrins. Biochem. Soc. Trans. 19, 1065-1069.

Pentikainen, U. and Ylanne, J. (2009). The regulation mechanism for the auto-inhibition of binding of human filamin A to integrin. J. Mol. Biol. 393, 644-657.

Perdiz, D., Mackeh, R., Pous, C. and Baillet, A. (2011). The ins and outs of tubulinacetylation: more than just a post-translational modification? Cell. Signal. 23, 763-771.

Perumal, S., Antipova, O. and Orgel, J. P. (2008). Collagen fibril architecture, domainorganization, and triple-helical conformation govern its proteolysis. Proc. Natl. Acad.

Sci. USA 105, 2824-2829.

Pfaff, M., Liu, S., Erle, D. J. and Ginsberg, M. H. (1998). Integrin beta cytoplasmicdomains differentially bind to cytoskeletal proteins. J. Biol. Chem. 273, 6104-6109.

Pfister, K. K., Shah, P. R., Hummerich, H., Russ, A., Cotton, J., Annuar, A. A.,

King, S. M. and Fisher, E. M. (2006). Genetic analysis of the cytoplasmic dyneinsubunit families. PLoS Genet. 2, e1.

Poh, Y. C., Na, S., Chowdhury, F., Ouyang, M., Wang, Y. and Wang, N. (2009).Rapid activation of Rac GTPase in living cells by force is independent of Src. PLoS

ONE 4, e7886.

Popowicz, G. M., Schleicher, M., Noegel, A. A. and Holak, T. A. (2006). Filamins:promiscuous organizers of the cytoskeleton. Trends Biochem. Sci. 31, 411-419.

Prass, M., Jacobson, K., Mogilner, A. and Radmacher, M. (2006). Directmeasurement of the lamellipodial protrusive force in a migrating cell. J. Cell Biol.

174, 767-772.

Puchner, E. M., Alexandrovich, A., Kho, A. L., Hensen, U., Schafer, L. V.,

Brandmeier, B., Grater, F., Grubmuller, H., Gaub, H. E. and Gautel, M. (2008).Mechanoenzymatics of titin kinase. Proc. Natl. Acad. Sci. USA 105, 13385-13390.

Puklin-Faucher, E. and Sheetz, M. P. (2009). The mechanical integrin cycle. J. Cell

Sci. 122, 179-186.

Puklin-Faucher, E., Gao, M., Schulten, K. and Vogel, V. (2006). How the headpiecehinge angle is opened: New insights into the dynamics of integrin activation. J. Cell

Biol. 175, 349-360.

Rajfur, Z., Roy, P., Otey, C., Romer, L. and Jacobson, K. (2002). Dissecting the linkbetween stress fibres and focal adhesions by CALI with EGFP fusion proteins. Nat.

Cell Biol. 4, 286-293.

Ramirez-Correa, G. A., Jin, W., Wang, Z., Zhong, X., Gao, W. D., Dias, W. B.,

Vecoli, C., Hart, G. W. and Murphy, A. M. (2008). O-linked GlcNAc modificationof cardiac myofilament proteins: a novel regulator of myocardial contractile function.Circ. Res. 103, 1354-1358.

Resh, M. D. (2006). Trafficking and signaling by fatty-acylated and prenylated proteins.Nat. Chem. Biol. 2, 584-590.

Rico, F., Roca-Cusachs, P., Sunyer, R., Farre, R. and Navajas, D. (2007). Celldynamic adhesion and elastic properties probed with cylindrical atomic forcemicroscopy cantilever tips. J. Mol. Recognit. 20, 459-466.

Ridley, A. J. (2011). Life at the leading edge. Cell 145, 1012-1022.

Riggins, R. B., Quilliam, L. A. and Bouton, A. H. (2003). Synergistic promotion of c-Src activation and cell migration by Cas and AND-34/BCAR3. J. Biol. Chem. 278,28264-28273.

Riveline, D., Zamir, E., Balaban, N. Q., Schwarz, U. S., Ishizaki, T., Narumiya, S.,

Kam, Z., Geiger, B. and Bershadsky, A. D. (2001). Focal contacts asmechanosensors: externally applied local mechanical force induces growth of focalcontacts by an mDia1-dependent and ROCK-independent mechanism. J. Cell Biol.

153, 1175-1186.

Roca-Cusachs, P., Gauthier, N. C., Del Rio, A. and Sheetz, M. P. (2009). Clusteringof alpha(5)beta(1) integrins determines adhesion strength whereas alpha(v)beta(3) andtalin enable mechanotransduction. Proc. Natl. Acad. Sci. USA 106, 16245-16250.

Roskoski, R., Jr (2005). Src kinase regulation by phosphorylation and depho-sphorylation. Biochem. Biophys. Res. Commun. 331, 1-14.

Rossier, O. M., Gauthier, N., Biais, N., Vonnegut, W., Fardin, M. A., Avigan, P.,

Heller, E. R., Mathur, A., Ghassemi, S., Koeckert, M. S. et al. (2010). Forcegenerated by actomyosin contraction builds bridges between adhesive contacts.EMBO J. 29, 1055-1068.

Rottner, K., Hall, A. and Small, J. V. (1999). Interplay between Rac and Rho in thecontrol of substrate contact dynamics. Curr. Biol. 9, 640.

Saha, S., Mundia, M. M., Zhang, F., Demers, R. W., Korobova, F., Svitkina, T.,

Perieteanu, A. A., Dawson, J. F. and Kashina, A. (2010). Arginylation regulatesintracellular actin polymer level by modulating actin properties and binding ofcapping and severing proteins. Mol. Biol. Cell 21, 1350-1361.

Saltel, F., Mortier, E., Hytonen, V. P., Jacquier, M. C., Zimmermann, P., Vogel, V.,

Liu, W. and Wehrle-Haller, B. (2009). New PI(4,5)P2- and membrane proximalintegrin-binding motifs in the talin head control beta3-integrin clustering. J. Cell Biol.

187, 715-731.

Sampath, R., Gallagher, P. J. and Pavalko, F. M. (1998). Cytoskeletal interactionswith the leukocyte integrin beta2 cytoplasmic tail. Activation-dependent regulation ofassociations with talin and alpha-actinin. J. Biol. Chem. 273, 33588-33594.

Sawada, Y. and Sheetz, M. P. (2002). Force transduction by Triton cytoskeletons.J. Cell Biol. 156, 609-615.

Sawada, Y., Tamada, M., Dubin-Thaler, B. J., Cherniavskaya, O., Sakai, R.,

Tanaka, S. and Sheetz, M. P. (2006). Force sensing by mechanical extension of theSrc family kinase substrate p130Cas. Cell 127, 1015-1026.

Sbrana, F., Sassoli, C., Meacci, E., Nosi, D., Squecco, R., Paternostro, F., Tiribilli,B., Zecchi-Orlandini, S., Francini, F. and Formigli, L. (2008). Role for stress fibercontraction in surface tension development and stretch-activated channel regulation inC2C12 myoblasts. Am. J. Physiol. Cell Physiol. 295, C160-C172.

Schaller, M. D., Borgman, C. A., Cobb, B. S., Vines, R. R., Reynolds, A. B. and

Parsons, J. T. (1992). pp125FAK a structurally distinctive protein-tyrosine kinaseassociated with focal adhesions. Proc. Natl. Acad. Sci. USA 89, 5192-5196.

Schiller, H. B., Friedel, C. C., Boulegue, C. and Fassler, R. (2011). Quantitativeproteomics of the integrin adhesome show a myosin II-dependent recruitment of LIMdomain proteins. EMBO Rep. 12, 259-266.

Schnepel, J. and Tschesche, H. (2000). The proteolytic activity of the recombinantcryptic human fibronectin type IV collagenase from E. coli expression. J. Protein

Chem. 19, 685-692.

Schober, M., Raghavan, S., Nikolova, M., Polak, L., Pasolli, H. A., Beggs, H. E.,Reichardt, L. F. and Fuchs, E. (2007). Focal adhesion kinase modulates tensionsignaling to control actin and focal adhesion dynamics. J. Cell Biol. 176, 667-680.

Schvartzman, M., Palma, M., Sable, J., Abramson, J., Hu, X. A., Sheetz, M. P. andWind, S. J. (2011). Nanolithographic control of the spatial organization of cellularadhesion receptors at the single-molecule level. Nano Lett. 11, 1306-1312.

Exploring mechanical molecular pathways 3037

Journ

alof

Cell

Scie

nce

Sechler, J. L., Rao, H., Cumiskey, A. M., Vega-Colon, I., Smith, M. S., Murata, T.and Schwarzbauer, J. E. (2001). A novel fibronectin binding site required forfibronectin fibril growth during matrix assembly. J. Cell Biol. 154, 1081-1088.

Seifert, U. (2000). Rupture of multiple parallel molecular bonds under dynamic loading.Phys. Rev. Lett. 84, 2750-2753.

Sellers, J. R. (2000). Myosins: a diverse superfamily. Biochim. Biophys. Acta 1496, 3-22.

Sellers, J. R. and Pato, M. D. (1984). The binding of smooth muscle myosin light chainkinase and phosphatases to actin and myosin. J. Biol. Chem. 259, 7740-7746.

Serrels, B., Serrels, A., Brunton, V. G., Holt, M., McLean, G. W., Gray, C. H.,

Jones, G. E. and Frame, M. C. (2007). Focal adhesion kinase controls actinassembly via a FERM-mediated interaction with the Arp2/3 complex. Nat. Cell Biol.

9, 1046-1056.Sharma, C. P., Ezzell, R. M. and Arnaout, M. A. (1995). Direct interaction of filamin

(ABP-280) with the beta 2-integrin subunit CD18. J. Immunol. 154, 3461-3470.Sieg, D. J., Hauck, C. R. and Schlaepfer, D. D. (1999). Required role of focal adhesion

kinase (FAK) for integrin-stimulated cell migration. J. Cell Sci. 112, 2677-2691.Sjoblom, B., Salmazo, A. and Djinovic-Carugo, K. (2008). Alpha-actinin structure and

regulation. Cell. Mol. Life Sci. 65, 2688-2701.Smith, M. L., Gourdon, D., Little, W. C., Kubow, K. E., Eguiluz, R. A., Luna-

Morris, S. and Vogel, V. (2007). Force-induced unfolding of fibronectin in theextracellular matrix of living cells. PLoS Biol. 5, e268.

Stamenovic, D., Mijailovich, S. M., Tolic-Nørrelykke, I. M., Chen, J. X. and Wang,N. (2002). Cell prestress. II. Contribution of microtubules. Am. J. Physiol. Cell

Physiol. 282, C617-C624.Su, J., Muranjan, M. and Sap, J. (1999). Receptor protein tyrosine phosphatase alpha

activates Src-family kinases and controls integrin-mediated responses in fibroblasts.Curr. Biol. 9, 505-511.

Sulchek, T. A., Friddle, R. W., Langry, K., Lau, E. Y., Albrecht, H., Ratto, T. V.,

DeNardo, S. J., Colvin, M. E. and Noy, A. (2005). Dynamic force spectroscopy ofparallel individual Mucin1-antibody bonds. Proc. Natl. Acad. Sci. USA 102, 16638-16643.

Sułkowska, J. I. and Cieplak, M. (2008). Stretching to understand proteins - a survey ofthe protein data bank. Biophys. J. 94, 6-13.

Sun, Z., Martinez-Lemus, L. A., Trache, A., Trzeciakowski, J. P., Davis, G. E., Pohl,U. and Meininger, G. A. (2005). Mechanical properties of the interaction betweenfibronectin and alpha5beta1-integrin on vascular smooth muscle cells studied usingatomic force microscopy. Am. J. Physiol. Heart Circ. Physiol. 289, H2526-H2535.

Svoboda, K. and Block, S. M. (1994). Force and velocity measured for single kinesinmolecules. Cell 77, 773-784.

Takagi, J., Petre, B. M., Walz, T. and Springer, T. A. (2002). Global conformationalrearrangements in integrin extracellular domains in outside-in and inside-outsignaling. Cell 110, 599-11.

Tamada, M., Sheetz, M. P. and Sawada, Y. (2004). Activation of a signaling cascadeby cytoskeleton stretch. Dev. Cell 7, 709-718.

Tan, J. L., Tien, J., Pirone, D. M., Gray, D. S., Bhadriraju, K. and Chen, C. S.

(2003). Cells lying on a bed of microneedles: an approach to isolate mechanical force.Proc. Natl. Acad. Sci. USA 100, 1484-1489.

ten Klooster, J. P., Evers, E. E., Janssen, L., Machesky, L. M., Michiels, F., Hordijk,P. and Collard, J. G. (2006). Interaction between Tiam1 and the Arp2/3 complexlinks activation of Rac to actin polymerization. Biochem. J. 397, 39-45.

Theriot, J. A. and Mitchison, T. J. (1991). Actin microfilament dynamics inlocomoting cells. Nature 352, 126-131.

Thomas, W. E., Vogel, V. and Sokurenko, E. (2008). Biophysics of catch bonds. Annu.

Rev. Biophys. 37, 399-416.Thompson, J. B., Kindt, J. H., Drake, B., Hansma, H. G., Morse, D. E. and Hansma,

P. K. (2001). Bone indentation recovery time correlates with bond reforming time.Nature 414, 773-776.

Tomar, A. and Schlaepfer, D. D. (2009). Focal adhesion kinase: switching betweenGAPs and GEFs in the regulation of cell motility. Curr. Opin. Cell Biol. 21, 676-683.

Torgler, C. N., Narasimha, M., Knox, A. L., Zervas, C. G., Vernon, M. C. andBrown, N. H. (2004). Tensin stabilizes integrin adhesive contacts in Drosophila. Dev.

Cell 6, 357-369.Tran, A. D., Marmo, T. P., Salam, A. A., Che, S., Finkelstein, E., Kabarriti, R.,

Xenias, H. S., Mazitschek, R., Hubbert, C., Kawaguchi, Y. et al. (2007). HDAC6deacetylation of tubulin modulates dynamics of cellular adhesions. J. Cell Sci. 120,1469-1479.

Triplett, J. W. and Pavalko, F. M. (2006). Disruption of alpha-actinin-integrininteractions at focal adhesions renders osteoblasts susceptible to apoptosis. Am. J.

Physiol. Cell Physiol. 291, C909-C921.Tsuruta, D. and Jones, J. C. (2003). The vimentin cytoskeleton regulates focal contact

size and adhesion of endothelial cells subjected to shear stress. J. Cell Sci. 116, 4977-4984.

Tu, Y., Wu, S., Shi, X., Chen, K. and Wu, C. (2003). Migfilin and Mig-2 link focaladhesions to filamin and the actin cytoskeleton and function in cell shape modulation.Cell 113, 37-47.

Tyska, M. J., Dupuis, D. E., Guilford, W. H., Patlak, J. B., Waller, G. S., Trybus,

K. M., Warshaw, D. M. and Lowey, S. (1999). Two heads of myosin are better thanone for generating force and motion. Proc. Natl. Acad. Sci. USA 96, 4402-4407.

Vega, F. M., Fruhwirth, G., Ng, T. and Ridley, A. J. (2011). RhoA and RhoC have

distinct roles in migration and invasion by acting through different targets. J. Cell

Biol. 193, 655-665.

Vogel, V. (2006). Mechanotransduction involving multimodular proteins: converting

force into biochemical signals. Annu. Rev. Biophys. Biomol. Struct. 35, 459-488.

von Wichert, G., Jiang, G., Kostic, A., De Vos, K., Sap, J. and Sheetz, M. P. (2003).

RPTP-alpha acts as a transducer of mechanical force on alphav/beta3-integrin-

cytoskeleton linkages. J. Cell Biol. 161, 143-153.

Walter, N., Selhuber, C., Kessler, H. and Spatz, J. P. (2006). Cellular unbinding

forces of initial adhesion processes on nanopatterned surfaces probed with magnetic

tweezers. Nano Lett. 6, 398-402.

Wang, H. B., Dembo, M., Hanks, S. K. and Wang, Y. L. (2001). Focal adhesion

kinase is involved in mechanosensing during fibroblast migration. Proc. Natl. Acad.

Sci. USA 98, 11295-11300.

Wang, Z., Pandey, A. and Hart, G. W. (2007). Dynamic interplay between O-linked

N-acetylglucosaminylation and glycogen synthase kinase-3-dependent phosphoryla-

tion. Mol. Cell. Proteomics 6, 1365-1379.

Wang, Z., Gucek, M. and Hart, G. W. (2008). Cross-talk between GlcNAcylation and

phosphorylation: site-specific phosphorylation dynamics in response to globally

elevated O-GlcNAc. Proc. Natl. Acad. Sci. USA 105, 13793-13798.

Wang, Z., Udeshi, N. D., Slawson, C., Compton, P. D., Sakabe, K., Cheung, W. D.,

Shabanowitz, J., Hunt, D. F. and Hart, G. W. (2010). Extensive crosstalk between

O-GlcNAcylation and phosphorylation regulates cytokinesis. Sci. Signal. 3, ra2.

Webb, D. J., Donais, K., Whitmore, L. A., Thomas, S. M., Turner, C. E., Parsons,

J. T. and Horwitz, A. F. (2004). FAK-Src signalling through paxillin, ERK and

MLCK regulates adhesion disassembly. Nat. Cell Biol. 6, 154-161.

Weiner, T. M., Liu, E. T., Craven, R. J. and Cance, W. G. (1993). Expression of focal

adhesion kinase gene and invasive cancer. Lancet 342, 1024-1025.

Welch, M. D. (1999). The world according to Arp: regulation of actin nucleation by the

Arp2/3 complex. Trends Cell Biol. 9, 423-427.

Wickstrom, S. A., Lange, A., Montanez, E. and Fassler, R. (2010). The ILK/PINCH/

parvin complex: the kinase is dead, long live the pseudokinase! EMBO J. 29, 281-291.

Wolfenson, H., Henis, Y. I., Geiger, B. and Bershadsky, A. D. (2009). The heel and

toe of the cell’s foot: a multifaceted approach for understanding the structure and

dynamics of focal adhesions. Cell Motil. Cytoskeleton 66, 1017-1029.

Wong, C. C., Xu, T., Rai, R., Bailey, A. O., Yates, J. R., 3rd, Wolf, Y. I., Zebroski,

H. and Kashina, A. (2007). Global analysis of posttranslational protein arginylation.

PLoS Biol. 5, e258.

Yang, C. X., Chen, H. Q., Chen, C., Yu, W. P., Zhang, W. C., Peng, Y. J., He, W. Q.,

Wei, D. M., Gao, X. and Zhu, M. S. (2006). Microfilament-binding properties of N-

terminal extension of the isoform of smooth muscle long myosin light chain kinase.

Cell Res. 16, 367-376.

Ye, L. H., Hayakawa, K., Kishi, H., Imamura, M., Nakamura, A., Okagaki, T.,

Takagi, T., Iwata, A., Tanaka, T. and Kohama, K. (1997). The structure and

function of the actin-binding domain of myosin light chain kinase of smooth muscle.

J. Biol. Chem. 272, 32182-32189.

Ylinen, P. (1996). Acetabular cup loosening–peeling off of the plasma-sprayed porous

coating in 2 cases. Acta Orthop. Scand. 67, 508-510.

Young, K. G. and Copeland, J. W. (2010). Formins in cell signaling. Biochim. Biophys.

Acta 1803, 183-190.

Yu, C. H., Law, J. B., Suryana, M., Low, H. Y. and Sheetz, M. P. (2011). Early

integrin binding to Arg-Gly-Asp peptide activates actin polymerization and

contractile movement that stimulates outward translocation. Proc. Natl. Acad. Sci.

USA 108, 20585-20590.

Zachara, N. E., O’Donnell, N., Cheung, W. D., Mercer, J. J., Marth, J. D. and Hart,

G. W. (2004). Dynamic O-GlcNAc modification of nucleocytoplasmic proteins in

response to stress. A survival response of mammalian cells. J. Biol. Chem. 279,

30133-30142.

Zaidel-Bar, R., Itzkovitz, S., Ma’ayan, A., Iyengar, R. and Geiger, B. (2007a).

Functional atlas of the integrin adhesome. Nat. Cell Biol. 9, 858-867.

Zaidel-Bar, R., Milo, R., Kam, Z. and Geiger, B. (2007b). A paxillin tyrosine

phosphorylation switch regulates the assembly and form of cell-matrix adhesions.

J. Cell Sci. 120, 137-148.

Zamir, E., Katz, M., Posen, Y., Erez, N., Yamada, K. M., Katz, B. Z., Lin, S., Lin,

D. C., Bershadsky, A., Kam, Z. et al. (2000). Dynamics and segregation of cell-

matrix adhesions in cultured fibroblasts. Nat. Cell Biol. 2, 191-196.

Zhang, H., Berg, J. S., Li, Z., Wang, Y., Lang, P., Sousa, A. D., Bhaskar, A., Cheney,

R. E. and Stromblad, S. (2004). Myosin-X provides a motor-based link between

integrins and the cytoskeleton. Nat. Cell Biol. 6, 523-531.

Zhang, X., Jiang, G., Cai, Y., Monkley, S. J., Critchley, D. R. and Sheetz, M. P.

(2008). Talin depletion reveals independence of initial cell spreading from integrin

activation and traction. Nat. Cell Biol. 10, 1062-1068.

Zheng, X. M., Resnick, R. J. and Shalloway, D. (2000). A phosphotyrosine

displacement mechanism for activation of Src by PTPalpha. EMBO J. 19, 964-978.

Zhu, W., Leber, B. and Andrews, D. W. (2001). Cytoplasmic O-glycosylation prevents

cell surface transport of E-cadherin during apoptosis. EMBO J. 20, 5999-6007.

Journal of Cell Science 125 (13)3038