RESEARCH ARTICLE

E-cadherin-mediated force transduction signals regulate globalcell mechanicsIsmaeel Muhamed1, Jun Wu2, Poonam Sehgal2, Xinyu Kong1, Arash Tajik3, Ning Wang3 andDeborah E. Leckband1,2,4,5,*

ABSTRACTThis report elucidates an E-cadherin-based force-transductionpathway that triggers changes in cell mechanics through amechanism requiring epidermal growth factor receptor (EGFR),phosphoinositide 3-kinase (PI3K), and the downstream formation ofnew integrin adhesions. This mechanism operates in addition tolocal cytoskeletal remodeling triggered by conformational changesin the E-cadherin-associated protein α-catenin, at sites ofmechanical perturbation. Studies using magnetic twistingcytometry (MTC), together with traction force microscopy (TFM)and confocal imaging identified force-activated E-cadherin-specificsignals that integrate cadherin force transduction, integrin activationand cell contractility. EGFR is required for the downstream activationof PI3K and myosin-II-dependent cell stiffening. Our findings alsodemonstrated that α-catenin-dependent cytoskeletal remodeling atperturbed E-cadherin adhesions does not require cell stiffening.These results broaden the repertoire of E-cadherin-based forcetransduction mechanisms, and define the force-sensitive signalingnetwork underlying the mechano-chemical integration of spatiallysegregated adhesion receptors.

KEY WORDS: E-cadherin, Integrin, Mechanotransduction, Tractionforce microscopy, Magnetic twisting cytometry, Cell signaling

INTRODUCTIONThe transduction of mechanical stimuli to alter intracellularbiochemical signals enables cells to modulate cell and tissuephysiology in response to force fluctuations propagated through thecell environment or to changes in endogenous contractile,protrusive or compressive forces. Mechanotransduction involvesan increasing repertoire of identified force-sensitive proteins foundin both cells and the extracellular matrix (ECM), and cell surfaceadhesion complexes are common force-transducing elements.Integrins and their ECM ligands are well-established forcetransducers (Geiger and Yamada, 2011). Intercellular adhesivecomplexes not only regulate cell-to-cell cohesion and tissue barrierintegrity, but also transduce force to instruct cell functions, alter cellshape and regulate tissue organization (Desai et al., 2013; Ladouxet al., 2010; le Duc et al., 2010; Leckband and de Rooij, 2014;Lecuit et al., 2011; Liu et al., 2010; Tepass et al., 2002; Tzima et al.,

2005; Yonemura et al., 2010). Fluctuations in tension also triggerjunction remodeling in endothelia and in epithelia, and thesechanges contribute to the mechanical reinforcement of adhesions(Barry et al., 2014, 2015; Liu et al., 2010; Thomas et al., 2013;Yonemura et al., 2010). Studies increasingly reveal howmechanotransduction at cell-to-cell adhesions in vivo impactsupon physiology in different mechanical contexts, such as atinterendothelial junctions near regions of disturbed flow and duringmorphogenesis (Hahn and Schwartz, 2009; Schluck et al., 2013;Weber et al., 2012).

The rudiments of intercellular mechanotransduction mechanismshave been identified in only a few cases (Barry et al., 2014; Collinset al., 2012; Kim et al., 2015; le Duc et al., 2010; Tzima et al., 2005;Yonemura et al., 2010). E-cadherin complexes at epithelialintercellular junctions are force sensitive (Barry et al., 2014; leDuc et al., 2010; Thomas et al., 2013; Yonemura et al., 2010), andα-catenin is an identified force-transducing protein in thesecomplexes (Yonemura et al., 2010). α-Catenin is a crucialmechanical link between homophilic intercellular E-cadherinbonds and the actin cytoskeleton (Barry et al., 2014; Buckleyet al., 2014; Cavey et al., 2008; Desai et al., 2013; Nagafuchi et al.,1991). Experimental evidence supports a mechanism in which theforce-dependent exposure of a cryptic binding site in α-cateninrecruits vinculin, and enables localized actin polymerizationthrough the Mena–VASP complex associated with vinculin(Barry et al., 2014; Buckley et al., 2014; le Duc et al., 2010;Leerberg et al., 2014; Thomas et al., 2013; Yao et al., 2014;Yonemura et al., 2010). This mechanism is consistent withmeasured force-activated changes in the viscoelasticity of E-cadherin adhesions (le Duc et al., 2010), but the stiffeningresponse could also reflect additional force-transductionmechanism(s).

Force-independent cadherin ligation is well known to activate anumber of signaling molecules including Src, phosphoinositide 3-kinase (PI3K), and Rho GTPases (Kovacs et al., 2002; McLachlanet al., 2007; McLachlan and Yap, 2007; Noren et al., 2003;Perez et al., 2008; Ratheesh et al., 2013; Tabdili et al., 2012a;Watanabe et al., 2009). Prior studies have also shown that passiveE-cadherin ligation to E-cadherin-coated beads, without mechanicalperturbation, altered focal adhesions through a mechanism thatinvolved Src and PI3K (Jasaitis et al., 2012). However, a completelyopen question is whether mechanical perturbations activate any ofthese same signals. Moreover, the details of possible force-activatedsignaling pathway(s), the impact of signals on other adhesionproteins in the cell, and their relationship to force- and E-cadherin-dependent changes in measured cell mechanics have yet to bedetermined.

This current study identified an additional E-cadherin-basedmechanotransduction mechanism that activates signal cascades thatincrease cell stiffness through integrin activation. Use of magneticReceived 24 December 2015; Accepted 3 March 2016

1Department of Biochemistry, University of Illinois Urbana Champaign, Urbana, IL61801, USA. 2Department of Chemical and Biomolecular Engineering, University ofIllinois Urbana Champaign, Urbana, IL 61801, USA. 3Department of MechanicalScience and Engineering, University of Illinois Urbana Champaign, Urbana, IL61801, USA. 4Department of Chemistry, University of Illinois Urbana Champaign,Urbana, IL 61801, USA. 5Carl W. Woese Institute of Genomic Biology, University ofIllinois Urbana Champaign, Urbana, IL 61801, USA.

*Author for correspondence (leckband@illinois.edu)

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twisting cytometry (MTC), traction force microscopy (TFM) andfluorescence imaging identified a force-actuated, E-cadherin-ligand-specific signaling cascade that activates distant integrinsand global cell contraction. By identifying early signaling cascadesin E-cadherin mechanotransduction, these findings provide newinsight into correlations between epithelial junction maturation andfocal adhesions (Mertz et al., 2013), and elaborate potential detailsof signaling underlying force independent integrin–cadherincrosstalk (Al-Kilani et al., 2011; Jasaitis et al., 2012).Importantly, this study establishes an additional E-cadherin-basedmechanotransduction mechanism, beyond proximal α-cateninconformation switching and local actin remodeling, thatcoordinates with integrins to regulate cell stiffening.

RESULTSForce loading E-cadherin receptors affects cell tractionforcesE-cadherin-mediated mechanotransduction triggers local vinculinrecruitment and actin polymerization at force-loaded E-cadherinreceptors (Barry et al., 2014; Kim et al., 2015; le Duc et al., 2010;Yonemura et al., 2010). This local cytoskeletal remodelingcoincides with increased viscoelasticity of mechanically perturbedE-cadherin adhesions (le Duc et al., 2010). Here, we tested whetherforce-activated E-cadherin signals could also alter cell mechanicsand possibly other adhesion proteins. Combining MTC with TFM(Fig. 1A), we first quantified effects of E-cadherin loading on globalcell contractility and focal adhesion remodeling.TFM quantifies changes in the root mean square traction forces

(ΔRMS traction), from displacement maps of fiduciary beadsembedded in hydrogel substrata relative to positions prior to cellperturbations (Butler et al., 2002). Passive ligation of beadsmodified with recombinant E-cadherin ectodomains (Prakasamet al., 2006) to E-cadherin receptors on MCF7 cells on collagen-coated gels (−Load, Fig. 1B,C) increased cell traction, as reportedpreviously (Jasaitis et al., 2012). However, applying an oscillatingshear stress (force/area) of 8.4 Pa on E-cadherin bead/cell junctionstriggered substantial increases in traction forces, from 144±18 Pa(−Load) to 249±31 Pa (+Load, Fig. 1B,C, Table 1). On fibronectinsubstrates, we similarly observed a 66% increase in ΔRMS traction(P<0.001, Table 1). The force-activated increase exceeded force-independent changes. Representative traction heat maps are inFig. S1A and values are summarized in Table 1. Distributions of themagnitudes of the local substrate strain vectors illustrate the changesin cell traction following E-cadherin receptor loading (Fig. S1B).Force-loading E-cadherin receptors shifted the mode of the tractionstress distribution from 25–50 pN (−Load) to 50–75 pN (+Load).Use of different magnetic bead coatings demonstrated the E-

cadherin-ligand specificity of the force-actuated changes in celltraction (Fig. 1D). Force-loading E-cadherin beads generatedsignificantly greater traction increases than beads coated with (1)poly-L-lysine (PLL), (2) a neutral, non-blocking anti-E-cadherinantibody (Petrova et al., 2012) or (3) the blocking anti-E-cadherinantibody DECMA-1 (Ozawa et al., 1990) (Fig. 1D). The ΔRMStraction changes thus required specific E-cadherin ligation.The traction changes also depended on the substrate coating

(Fig. 1E). With cells cultured on PLL-coated gels, in the presence ofintegrin-blocking antibodies, E-cadherin force loading increasedΔRMS traction by only 134±19 Pa (mean±s.e.m.; Fig. 1E; Table 1).Force-loading PLL beads on cells on PLL-coated gels increased theΔRMS traction by 82±12 Pa. Surprisingly, on E-cadherin-coatedgels, the ΔRMS traction of 172±20 Pa was statistically similar tocells on PLL substrata (Fig. 1E, Table 1).

E-cadherin-specific mechanotransduction triggers focaladhesion remodelingTo determine whether force transduction signals altered focaladhesions, we quantified the number and area of focal adhesions(Gonon et al., 2005; Zamir et al., 1999) based on analyses ofimmunofluorescence images of vinculin at the basal plane. Focaladhesions were distributed mainly at the cell perimeter (Fig. 1F).Upon loading E-cadherin beads, the average focal adhesion area andnumber increased by 35% and 20%, respectively, relative to controls(−Load) (Fig. S1C). Controls with DECMA-1-coated beads, whichdo not trigger adaptive stiffening (Tabdili et al., 2012b) or changesin traction forces (Fig. 1D), did not alter focal adhesionssignificantly (Fig. S1C). Thus, focal adhesion remodeling wasactivated by ligand-dependent E-cadherin mechanotransductionand was not due to general tugging on cell membranes. Thesecombined MTC and TFM measurements thus demonstrated that, inaddition to proximal α-catenin-dependent cytoskeletal remodeling(Barry et al., 2014; le Duc et al., 2010; Yonemura et al., 2010),force-activated E-cadherin-mediated signals alter distal integrin-mediated traction forces.

E-cadherin-mediated adaptive stiffening requires theformation of new integrin adhesionsMTC measurements confirmed that cells exhibit E-cadherin-mediated adaptive stiffening when cultured on ECM proteins oneither glass or on 34 kPa gels. Applying a modulated, orthogonalmagnetic field induces a twisting torque, which displaces the beads.The bead displacements, D, are related to the storage G′ and loss G″moduli of the bead–cell junctions by: T/D=G′+G″. In these studies,G″ did not change; decreasing displacement D reflected an increasein G′, or the stiffness (Wang et al., 1993). Changes in the percentagestiffness are reported relative to the initial modulus of theunperturbed junction. Perturbing E-cadherin beads with a shearstress of 7.2 Pa (force/area) increased MCF7 cell stiffness by ∼35%on glass and gels (Fig. 2A). Control, PLL-coated beads failed toinduce cell stiffening (Fig. 2A).

In endothelial cells, adaptive cell stiffening triggered byPECAM-1 requires the formation of new integrin adhesions(Collins et al., 2012). To test whether E-cadherin-mediatedepithelial cell stiffening similarly requires integrins, MTCmeasurements were conducted with cells on different matrixproteins, in the presence of 10% serum (Fig. 2B). On fibronectinand on collagen I (Col I) substrata, E-cadherin bond loadingincreased cell stiffness by 23±2% and 32±5% (mean±s.e.m.),respectively. On PLL-coated gels in the presence of integrin-blocking antibodies AIIB2 and GOH3, the stiffness change was−3±5%. Somewhat surprisingly, with cells seeded on E-cadherin-coated gels, the stiffness change was −12±8%. Integrin-blockingantibodies were also included in the latter measurements.

To further test whether stiffening required the formation of newintegrin adhesions, cells cultured on matrix-coated glass weretreated with integrin- or fibronectin-blocking antibodies for 25 min,immediately prior to E-cadherin force loading. This brief antibodytreatment (25-min exposure) did not affect the average spread areaof cells or existing focal adhesions (Fig. 2C,D) (Collins et al., 2012;Hall et al., 1990). In cells treated with anti-β1 integrin AIIB2, orwith anti-fibronectin antibody 16G3, the stiffening responses were−4±5% and 3±4%, respectively (Fig. 2E). The non-blocking,control (for 16G3) antibody 13G12 (Nagai et al., 1991) did notsignificantly alter cell stiffening (Fig. 2E).

The results with integrin function-blocking antibodies suggestthat integrin conversion into an active conformation triggers

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Fig. 1. E-cadherin-based mechanotransduction alters cell traction and focal adhesions. (A) Illustration of the experimental setup combining magnetictwisting cytometry (MTC) and traction force microscopy (TFM). An oscillating magnetic field H generates a torque T, which displaces the magnetic beads. Theamplitude of the bead displacement reflects the viscoelastic modulus of the bead–cell junction. Determined changes in cell stiffness or traction changes used cellswith single beads, and excluded the majority of cells with multiple beads or beads at cell–cell contacts. (B) Time sequence of steps in combined MTC andTFMmeasurements. (C) Bar graph indicating changes in traction force (with or without load) exerted by MCF7 cells on collagen-coated polyacrylamide gels withelastic moduli of 8.8 kPa (−Load, n=7 cells; +Load, n=19 cells) and 34 kPa (−Load, n=11 cells, +Load, n=11 cells). (D) Bar graph showing changes in cell tractionafter force-loading beads modified with E-cadherin (E-cad, n=11 cells), poly-L-lysine (PLL, n=18 cells), neutral anti-E-cadherin antibody (Ntrl Ab, n=8 cells), orblocking anti-E-cadherin antibody (DECMA-1, n=8 cells). (E) Bar graph indicating traction changes (ΔRMS traction, Pa) after force-loading E-cadherin beadson cells adhered to collagen (n=11 cells), PLL (n=9 cells), or E-cadherin-coated polyacrylamide gels (34 kPa, n=7 cells). Results obtained with PLL-coated beadson cells adhered to PLL substrata are also shown (n=6 cells). With cells on either PLL- or E-cadherin-coated substrata, the medium contained integrin-blockingantibodies GOH3 and AIIB2. In C–E, the black bar denotes the same data used for statistical comparisons. Data presented are the mean±s.e.m. *P<0.01(Student’s t-test). Two or more independent experiments were performed. (F) Representative confocal immunofluorescence images of vinculin (green) and actin(gold) at the basal plane of cells on collagen-functionalized hydrogels. Cells were probedwith E-cadherin (top) and DECMA-1 (bottom) functionalized beads, with(+Load) and without (−Load) 2 min of force loading. Scale bar: 10 µm.

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increased cell contractility, such that the amplitude of the stiffeningresponse would depend on the population of integrins undergoingthis transition. We tested whether reducing the pool of integrinsactivated by E-cadherin-mediated signals would reduce thestiffening amplitude by pre-activating integrins with MnCl2(0.5 mM) (Cluzel et al., 2005; Mould, 2002). This treatmentreduced the force-dependent stiffening from 32±5% to 12±5%(Fig. 2E). This treatment also generated a higher (than untreatedcells) basal traction force of 315±22 Pa, consistent with integrinactivation.Integrin activation downstream of E-cadherin force transduction

signals was further verified using the GST–fibronectin-III9-11peptide, which specifically binds activated α5β1 integrin (Orret al., 2006; Tzima et al., 2001). Quantitative comparisons ofimmunofluorescence images of whole cells revealed a force-dependent 34±14% increase in peptide labeling, relative tounperturbed (−Load) controls (Fig. 3A,B). Western blotsindicated a 200±10% increase in peptide binding (Fig. 3C,D).The different amplitudes of these changes reflect differences in thesensitivities of the two assays, but both results confirmed integrinactivation, relative to unperturbed controls.

Force-activated E-cadherin-mediated signals required fortraction changesDiverse, independent studies have implicated non-muscle myosin II(MyII, also known as myosin-2), PI3K, Src and/or Rho-associatedprotein kinases (ROCK1 and ROCK2) in force-independent(−Load) crosstalk between cadherins and integrins, as well as inPECAM-1-mediated mechanotransduction (Collins et al., 2012;Jasaitis et al., 2012; Schwartz and DeSimone, 2008; Weber et al.,2011). Cell treatment with blebbistatin or with the ROCK inhibitorY27632 reduced the load-dependent ΔRMS traction by 53% and51%, respectively, relative to untreated cells (Fig. 4A,B). Thesetraction changes were comparable to negative controls withantibody-coated beads (Table 1). Src or PI3K inhibitors reducedthe load-dependent ΔRMS traction by 36% and by 38%,respectively, relative to untreated cells (Fig. 4B). Table 1 alsosummarizes the inhibitor effects on the basal traction (withoutbeads).

The effects of inhibitors on adaptive stiffening were morepronounced (Fig. 4C). Blebbistatin (targeting MyII) and Y27632(targeting ROCK1 and ROCK2) reduced adaptive stiffening from32±5% to 5±6% and 0±3% (mean±s.e.m.), respectively. The Srcinhibitor PP2 reduced the stiffening response to 6±4%. The PI3Kinhibitors LY294002 and Wortmannin reduced the stiffeningresponse to −4±6 and 7±5%, respectively. The Wortmanninconcentration used reportedly does not affect other kinases orphospholipase A2 (Fruman et al., 1998; Orr et al., 2006).

Mechanical perturbation of E-cadherin does not increase SrcactivityE-cadherin ligation (−Load) activates Src kinase (McLachlan et al.,2007), and the Src inhibitor PP2 inhibited E-cadherin actuated cellstiffening (Fig. 4C). However, visualizing changes with a FRET-based, membrane-associated KRas-Src reporter (Wang et al., 2005)indicated that E-cadherin force-loading did not enhance Src activity.The measured CFP-to-YFP emission ratio reports Src activity(Wang et al., 2005). In regions of interest (ROIs) around beads, theload-independent (−Load) Src activity peaked ∼30 min after beadligation, and then dropped to initial levels within ∼45 min (data notshown). To minimize contributions from force-independentsignaling, we measured the CFP-to-YFP ratio during forceloading, at 40 min after bead ligation. However, ∼30 s ofE-cadherin receptor loading did not significantly increase theCFP-to-YFP ratio, relative to the no-load control (Fig. S2). Thus,mechanically perturbing E-cadherin receptors does not increase Srcactivity, but a pool of active Src appears to be essential forE-cadherin-mediated cell stiffening.

Force-loading E-cadherin activates PI3K upstream ofintegrinsTo test whether mechanically stimulated E-cadherin receptorsactivated PI3K, we quantified the accumulation of the PI3K reporterPH-Akt–GFP in ROIs around E-cadherin beads, relative to controlswith PLL beads or cells expressing GFP (Fig. 5A–E). Force-loadingE-cadherin resulted in a 41±0.1% (mean±s.e.m.) increase in thenormalized, mean fluorescence intensity (Fig. 5A–F). Treatmentwith integrin or fibronectin-blocking antibodies abrogated adaptive

Table 1. Effect of bead coating, substrate modulus, and substrate coating on the RMS traction forces

Substrate modulus (kPa)and coating Bead coating

Basaltraction (Pa) s.e.m.

ΔRMS traction−Load (Pa) s.e.m.

ΔRMS traction+Load (Pa) s.e.m.

8.8 kPa Collagen E-cad-Fc 74.2 (6) 15 27.9 (7) 4 53.4 (19) 68.8 kPa Collagen PLL 74.2 (6) 15 41.0 (10) 5 45.8 (8) 434 kPa Collagen E-cad-Fc 281.6 (9) 38 144.3 (11) 18 248.6 (11) 3134 kPa Collagen E-cad-Fc

Y27632169.7 (8) 19 146.7 (8) 23 120.7 (12) 17

34 kPa Collagen E-cad-FcBlebbistatin

136.6 (6) 26 175.6 (6) 26 117.1 (9) 16

34 kPa Collagen E-cad-FcLY294002

187.9 (8) 28 107.9 (10) 11 153.8 (7) 20

34 kPa Collagen E-cad-FcPP2

211.9 (8) 25 96.9 (8) 10 158.8 (8) 12

34 kPa Collagen PLL 281.6 (9) 38 170.3 (8) 26 166.3 (18) 1434 kPa Collagen DECMA-1 281.6 (9) 38 169.9 (6) 28 113.6 (8) 2034 kPa Collagen Ntrl Ab 281.6 (9) 38 – – 122.4 (8) 1434 kPa PLL E-cad-Fc 130.6 (7) 16 159.2 (7) 30 134.0 (9) 1934 kPa PLL PLL 130.6 (7) 16 113.3 (7) 13 82.2 (6) 1234 kPa E-cad-Fc E-cad-Fc 134.7 (3) 22 182.4 (7) 19 171.7 (7) 2034 kPa fibronectin E-cad-Fc 347.7 (6) 63 138.4 (19) 9 230.2 (11) 11.2

Data include the basal traction force (BTF) and changes in traction (ΔRMS traction) following bead binding, with or without Load. Values are the mean andstandard error of the mean (s.e.m.). The numbers in parenthesis indicate the number of independent measurements.

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stiffening (Fig. 2E), but not PH-Akt–GFP accumulation at force-loaded E-cadherin beads: mean fluorescence intensities (MFIs)increased by 54±0.1% and by 49±0.1% in cells treated with AIIB2(Fig. 5F) or 16G3, respectively (Fig. 5D–F). GFP did not accumulateat E-cadherin beads, and PH-Akt–GFP did not accumulate at PLLbeads (Fig. 5B,F). In addition, phosphorylation of endogenous Akt(at S473) increased by 21% in western blots of cell monolayersstimulated with E-cadherin beads, compared with a 2.6% changemeasured with PLL bead controls (data not shown). These resultssuggest that mechanically perturbed E-cadherin receptors activatePI3K upstream from integrins.To determine whether focal adhesion remodeling and cell

stiffening corresponded with basal increases in PI3K activity, weimaged PH-Akt–GFP in regions of interest around focal adhesionsat the basal plane. Despite changes in focal adhesion number andsize, the sensor did not reveal any detectable force-dependent

changes in PH-Akt–GFP distributions at the basal plane, abovebackground.

MCF7 cells express a mutant form of PI3K (Vasudevan et al.,2009). Thus, similar studies were performed with MDCK cells – anon-tumorigenic epithelial cell line, which exhibits E-cadherin-mediated adaptive cell stiffening (Barry et al., 2014). With MDCKcells, PH-Akt–GFP similarly accumulated at force-loaded E-cadherin beads, relative to controls (Fig. S3).

EGFR is required for E-cadherin-mediated PI3K activationand adaptive stiffeningReceptor tyrosine kinases, which associate with different cadherins,activate PI3K (Cully et al., 2006), and VEGFR2 and EGFR co-immunoprecipitate with, respectively, VE-cadherin (Coon et al.,2015) and E-cadherin (Pece and Gutkind, 2000; Perrais et al., 2007).To test whether EGFR is required for E-cadherin-mediated PI3K

Fig. 2. E-cadherin-mediated adaptivestiffening requires integrin adhesion.(A) The mean±s.e.m. percentage stiffnesschange versus time during E-cadherinforce loading, with cells on collagen-coated glass (black triangles, n=116beads) or 34 kPa gels (black squares,n=67 beads). Control with poly-L-lysine(PLL) beads (black circles, n=27 beads) isalso shown. Two independentexperiments were performed for allconditions. (B) The mean±s.e.m.percentage stiffness change after 2 min offorce loading with E-cadherin beadsattached to cells adhered to 34-kPa gelscoated with fibronectin (FN, n=9 cells),E-cadherin (E-cad, n=16 cells), poly-L-lysine (PLL, n=22 cells), or collagen (Col,n=13 cells). The negative control was withPLL beads bound to cells on PLL-coatedgels (34 kPa) (n=16 cells). The x-axislabels indicate ‘bead coating/substratecoating’. *P<0.05 (Student’s t-test). Twoindependent experiments were performed.(C) Representative confocalimmunofluorescence images of vinculin(green, top) and actin (gold, bottom) inuntreated MCF7 cells, or cells treated withfunction-blocking antibodies 16G3 orAIIB2 for 25 min. Scale bar: 10 µm. (D) Themean±s.e.m. cell area after brief antibodyexposure is plotted for comparison with C(untreated, n=82 cells; AIIB2, n=79 cells;16G3 treated, n=80 cells). (E) The mean±s.e.m. percentage stiffness change after2 min of E-cadherin force loading onMCF7cells subject to different treatments. Datashown are for untreated MCF7 cells oncollagen-coated glass (black bar, n=116beads), cells treated with antibodies [AIIB2(n=83 beads), 16G3 (n=80 beads), or13G12 (n=56 beads)] for 25 min prior toE-cadherin force loading, and cells treatedfor 25 min with 0.5 mM MnCl2 chloride(n=49 beads). *P<0.05 (Student’s t-test).Two independent experiments wereperformed.

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activation and cell stiffening, cells were treated with the EGFR-specific inhibitor Gefitinib (Paez et al., 2004). At concentrations thatblock EGFR transphosphorylation (Yen et al., 2015), Gefitinibabrogated MCF7 cell stiffening, relative to vehicle controls(DMSO) or untreated cells (Fig. 5G). MDCK cell treatmentreduced adaptive stiffening from 26±5% (mean±s.e.m.; n=47cells, more than three experiments) to 9±3% (P<0.05; n=60 cells,three experiments). Gefitinib also abrogated PH-Akt–GFPrecruitment to force-loaded E-cadherin beads on both MCF7(Fig. 5F) and MDCK cells (data not shown).Gefitinib selectively inhibits EGFR, but not other EGFR-related

receptor tyrosine kinases. However, it has been shown to inhibitother kinases with much higher IC50 values (Brehmer et al., 2005).As an alternative, we therefore treated cells with neutralizing anti-EGF antibody, which specifically inhibits EGF-dependent cellproliferation (Sakaguchi et al., 2008), and found that this treatmentabolished PH-Akt–GFP recruitment to force-loaded E-cadherinreceptors on MCF7 cells (Fig. 5F; n>40, two experiments). Anti-EGF antibody also blocked E-cadherin-mediated adaptive stiffening(Fig. 5G, P<0.05; n=71 cells, three experiments), indicating that cellstiffening is EGF (ligand) dependent.

Actin recruitment to perturbed E-cadherin junctions doesnot require integrin activationWe further tested whether actin accumulation at force-loaded E-cadherin beads – attributed to α-catenin unfurling and vinculinrecruitment (Barry et al., 2014; le Duc et al., 2010; Thomas et al.,

2013; Yonemura et al., 2010) – depends on cell stiffening. Althoughthe anti-β1 antibody AIIB2 abrogated cell stiffening (Fig. 2E), it didnot block actin accumulation at force-loaded E-cadherin beads(Fig. S4). The apparent difference in actin accumulation at E-cadherin beads on AIIB2-treated cells was not due tononspecifically adsorbed matrix protein on the beads, becauseactin accumulation was unaffected by the presence or absence ofserum (Fig. S4C). Conversely, there was no actin accumulationaround PLL beads, in measurements with 10% serum. These resultssuggest that force-activated stiffening of bead–cell junctions mainly

Fig. 3. E-cadherin-force transduction activates α5β1 integrin.(A) Representative fluorescence images of cells treated with the integrinactivationmarker GST–fibronectin (FN)-III9-11, after force loading, washing andthen staining with anti-GST antibody. (B) The mean±s.e.m. fluorescenceintensity (MFI) of whole cells stained with anti-GST-antibody, with or withoutload (−Load, n=112 beads; +Load, n=104 beads). Two independentexperiments were performed. Data were normalized to the −Load condition.(C) Representative western blots for GST–fibronectin-III9-11 and actin.Mechanically stimulated cells were treated with GST-fibronectin-III9-11, lysed,and bound GST-fibronectin-III9-11 was assessed by blotting for GST. Twoindependent experiments were performed. (D) Comparison (mean±s.e.m.) ofpeptide binding in C with or without load. Blots were normalized to actin, andintensities were relative to the −Load condition. *P<0.01 (Student’s t-test).

Fig. 4. Force-activated, E-cadherin-dependent cell stiffening requires Src,PI3K, myosin andROCK. (A) Time sequence for measurements of the impactof inhibitors on ΔRMS Traction and adaptive stiffening. (B) The mean±s.e.m.change in the root mean square traction forces (ΔRMS Traction, Pa), afterforce-loading E-cadherin beads bound to cells treatedwith inhibitors of ROCK1and ROCK2 (Y27632, n=12 cells), myosin II (Blebbistatin, n=9 cells), Srckinase (PP2, n=8 cells), or PI3K (LY294002, n=7 cells). Two independentexperiments were performed. (C) The mean±s.e.m. percentage stiffnesschange, after 2 min of force-loading E-cadherin beads bound to cells treatedwith inhibitors of myosin II (blebbistatin, n=43 beads), ROCK1 (Y27632, n=33beads), Src kinase (PP2, n=39 beads), PI3K (LY294002, n=22 beads;Wortmanin, n=47 beads). For the DMSO controls, n=50 beads, and more than50 beads were analyzed in the PP3 controls. Two independent experimentswere performed. *P<0.05 (Student’s t-test).

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reflects changes in MCF7 cell contractility, rather than localizedcytoskeletal remodeling.

DISCUSSIONFindings reported in this paper identified an additional E-cadherinmechanotransduction pathway, beyond force-dependent α-cateninswitching and local cytoskeletal remodeling (Barry et al., 2014;Leckband and de Rooij, 2014; Leckband et al., 2011; Yonemuraet al., 2010). This mechanism, summarized in Fig. 6, involves anEGFR-dependent signaling pathway that activates PI3K anddownstream integrin ligation, followed by subsequent myosin-II-dependent cell stiffening.The dependence of adaptive stiffening on PI3K, Src and integrins

parallels mechanotransduction mediated by the immunoglobulin

superfamily protein PECAM-1 in endothelia (Collins et al., 2012),but there are differences. One obvious difference is that E-cadherinactivates force transduction in epithelia, which lack PECAM-1 andVE-cadherin. Importantly, these findings also revealed that EGFR isrequired for E-cadherin-mediated force-dependent PI3K activationand cell stiffening. Although we did not explicitly demonstrate thatPI3K activates integrins, prior studies have reported that PI3Kconverts integrins into a high-affinity state independently of theECM (Batra et al., 2012; Katsumi et al., 2005; Orr et al., 2006). Ourresults with the fibronectin-blocking antibody 16G3 showed thatECM ligation and the formation of new adhesions is required for themyosin II-dependent cell contractility.

The abrogation of stiffening in cells on E-cadherin substrata orfollowing treatment with function blocking antibodies confirmed

Fig. 5. E-cadherin-dependent force transduction activates PI3K. (A–E) Representative confocal fluorescence images of PH-Akt–GFP-transfected cells with(+Load) and without (−Load) force-loading beads (white arrows indicate bead locations) coated with (A) E-cadherin (E-cad) or (B) PLL. (C) Controls used E-cadherin beads and cells expressing GFP. (D,E) PH-Akt-GFP recruitment to E-cadherin beads on MCF7 cells treated with (D) integrin-blocking antibody AIIB2,and (E) fibronectin-blocking antibody 16G3. Only cells laden with single beads were used for data analyses. (F) Bar graph of the normalized mean±s.e.m.fluorescence intensity of PH-Akt–GFP in ROIs surrounding E-cadherin beads, without force-loading (No Load, white) and after 4 min of force loading (+Load,black). Values were normalized to the No Load condition. Cells were grown on collagen-coated glass. Results show the normalized fluorescence in the presenceof anti-integrin antibody AIIB2 (−Load, n=158 beads; +Load, n=186 beads, *P<0.001, Student’s t-test), Gefitinib (−Load, n=205 beads; +Load, n=169 beads,*P=0.5, Student’s t-test), or anti-EGF antibody (−Load, n=44 beads; +Load, n=50 beads, P=0.15, Student’s t-test). Controls werewith PLL-coated beads (−Load,n=133 beads; +Load, n=124 beads) or with cells expressingGFP (−Load, n=138 beads; +Load, n=106 beads). Two independent experiments were performed forall conditions. (G) The mean±s.e.m. percentage stiffness change after 2 min of E-cadherin bead loading for untreated cells (black) or cells treated with anti-EGFantibody (n=71) or Gefitinib (n=70). DMSO control is also shown (n=65). Two independent experiments were performed. *P<0.001 (Student’s t-test).

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that cell stiffening is integrin dependent. The absence of stiffeningon E-cadherin substrata was unexpected, because E-cadherin-basedadhesion supports traction forces and activates GTPases, Src andPI3K signaling (Kovacs et al., 2002; le Duc et al., 2010; McLachlanand Yap, 2011; Tabdili et al., 2012a). In addition, E-cadherinreceptors are mechanically coupled to actin, and could potentiallytransmit force through the cytoskeleton by stress focusing (Hu et al.,2003). Indeed, RGD-coated beads trigger changes in basal focaladhesions that are attributed to force transmission throughmicrofilaments (Hu et al., 2003). The latter mechanicalmechanism is, however, inconsistent with the inability of apicalE-cadherin perturbations to alter basal E-cadherin adhesions.Similarly, perturbing E-cadherin receptors with DECMA-1-coatedbeads failed to either alter traction forces (Fig. 1D) or triggerstiffening by cells on ECM (le Duc et al., 2010). These resultsindicate that E-cadherin force transduction triggers integrin ligationprimarily through biochemical signals. Results with fibronectin-blocking antibody in turn indicate that ECM ligation is required forsubsequent cell stiffening.E-cadherin force-loading does not appear to locally activate Rho-

dependent contractility through RhoGEFs, as reported at focaladhesions (Guilluy et al., 2011), because the integrin blockadeablated the stiffening response. Similarly, the absence of stiffeningby cells on PLL or on E-cadherin substrata also appears to rule thisout.We did not explicitly identify specific integrins or matrix proteins

required for E-cadherin-mediated cell stiffening. In most studies,cells were initially seeded on collagen-coated substrata but werecultured with 10% serum for at least 6 h prior to measurements.Immunofluorescence images exhibited little fibronectin on thesurrounding matrix, but the cells stained positive for fibronectin,confirming matrix secretion. The anti-β1-integrin antibody AIIB2,which blocks integrin receptors for laminin, collagens andfibronectin (Zaidel-Bar and Geiger, 2010), abrogated cellstiffening. However, the demonstrated activation of α5β1 integrin(fibronectin receptor) and the abrogation of stiffening by treatmentwith anti-fibronectin antibody 16G3 suggest that integrin activationand binding to fibronectin play a major roll in the stiffeningresponse.

It is possible that α-catenin could also participate in this signalingpathway. In an obvious way, α-catenin is essential to sustain tensionacross cadherin adhesions required for force transduction (Barryet al., 2014; Buckley et al., 2014; Thomas et al., 2013). However,these results are intriguing in light of reports that E-cadherin-mediated adaptive stiffening, but not α-catenin conformationswitching, requires the vinculin-binding site of α-catenin (Barryet al., 2014; Kim et al., 2015). Thus, α-catenin might play anadditional role in this additional signaling pathway. Alternatively,its ligands such as vinculin might act as force-sensitive scaffolds forother signaling proteins.

The abrogation of cell stiffening correlated with some apparentreduction in actin accumulation at E-cadherin beads might suggesta positive feedback loop wherein endogenous contractile forcesenhance actin polymerization at perturbed E-cadherin adhesions.Increased contractile forces might simply increase α-cateninunfurling at beads, with a concomitant increase in local actinpolymerization (Leerberg et al., 2014). Alternatively, zyxinfacilitates force-dependent actin polymerization at focaladhesions, and is enriched at stressed adherens junctions (Hirataet al., 2008; Oldenburg et al., 2015). Mechanically stimulating actinfibers also stimulated formin-dependent fiber elongation (Higashidaet al., 2013; Riveline et al., 2001). Assessing whether suchmechanisms underlie our observations is beyond the scope of thisstudy.

The broader physiological implications of this EGFR-dependentmechanism have yet to be established, but our results suggest thatforce perturbs well-documented bidirectional signaling between E-cadherin and EGFR. Blocking EGFR function has been shown topromote desmosome formation and cell adhesion (Lorch et al.,2004; Wheelock and Johnson, 2003). Alternatively, in somecontexts, EGFR activation leads to E-cadherin and β-catenin andp120 catenin phosphorylation and disruption of cytoskeletalinteractions (Hazan and Norton, 1998; Mariner et al., 2004).Conversely, in contact-inhibited proliferation, E-cadherin adhesionsuppresses EGFR signaling (Mariner et al., 2004; Perrais et al.,2007; Qian et al., 2004). E-cadherin appears to block EGFRtrafficking to signaling compartments, by retaining it at themembrane (Curto et al., 2007; McClatchey and Yap, 2012), butcadherin might also suppress EGFR trans phosphorylation (Fedor-Chaikin et al., 2003). Our data suggest that mechanically perturbingE-cadherin might relieve contact-inhibited EGFR signaling.Intriguingly, our results might account for the increase in growth-factor-dependent cell proliferation with increasing substrate rigidityand junctional tension (Kim and Asthagiri, 2011; Kim et al., 2009).

In summary, these results support the model in Fig. 6, whereinE-cadherin-mediated force transduction is mechano-chemicallycoupled to integrin activation in epithelial cells through an EGFR-dependent pathway. This mechanism operates in addition to theproximal, α-catenin dependent cytoskeletal remodeling. Despitefunctional and biochemical differences between E-cadherin andPECAM-1, the similar mechanotransduction signaling suggeststhat these receptors exploit common pathways in different tissuesto transduce force throughout mechano-chemically integratednetworks.

MATERIALS AND METHODSCell lines, protein production and inhibitorsMichigan Cancer Foundation-7 (MCF7) human epithelial breast carcinomaand Madine Darby canine kidney (MDCK) cells were from the ATCC, andwere used as received. The cells were maintained in Dulbecco’s modifiedEagle’s medium (DMEM 4.5 g/l glucose) supplemented with 10% (v/v)

Fig. 6. Proposed model for E-cadherin-mediated globalmechanotransduction. Mechanically stimulated E-cadherin receptorsactivate PI3K through an EGFR-dependent mechanism. PI3K activation isrequired for the downstream formation of new integrin adhesions andconsequent ROCK-dependent activation of myosin II-dependent cellcontractility.

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fetal bovine serum (FBS), 1 mM sodium pyruvate, and 1% (v/v) penicillin–streptomycin (Corning Cell Grow, Manassas, VA) unless otherwise stated.Recombinant E-cadherin extracellular fragment tagged with the C-terminalFc domain of human IgG (Ecad-Fc) was produced as described previously(Prakasam et al., 2006).

Src kinasewas blocked by incubating cells with 10 μMPP2 (Calbiochem,UK) for 30 min, prior to mechanical measurements. PI3K was inhibited bytreating cells with 50 μM LY294002 (Cell Signaling Technologies, Boston,MA) or with 5 nMWortmannin (Santa Cruz Biotechnology, CA) for 1 h and0.5 h, respectively (Orr et al., 2006). At 5 nM, Wortmanin reportedly doesnot inhibit other kinases or phospholipase A2 (Orr et al., 2006; Vlahos et al.,1994). Treatment with 10 μMY27632 (Tocris Biosciences, Bristol, UK) for30 min inhibits ROCK (Shewan et al., 2005). Treatment with 0.05 mMblebbistatin for 1 h inhibits myosin II (Barry et al., 2014). EGFR wasblocked with 15 μM of the non-competitive EGFR-specific inhibitorGefitinib (IRESSA) (Selleckchem, Houston, TX) for 2 h (Yen et al., 2015).Gefitinib is an EGFR-specific, non-competitive inhibitor that blocks EGFRkinase activity (Brehmer et al., 2005; Paez et al., 2004). EGFwas neutralizedwith 5 µg/ml human monoclonal anti-EGF antibody (R&D Systems,Minneapolis, MN: Mouse IgG1 Clone #10825) for 30 min (Sakaguchiet al., 2008). When blocking EGF during MTC measurements, cells werebathed in medium containing anti-EGF antibody.

Integrin and ECM antibodiesIntegrins were blocked with anti-α6 antibody (GOH3) (20 μg/ml; SantaCruz Biotechnology, SC-19622) and with anti-β1-integrin antibody AIIB2(1:20 dilution) (Barry et al., 2014). The hybridoma producing AIIB2 wasfrom Johan de Rooij (Hubrecht Institute, The Netherlands). The anti-fibronectin antibody 16G3 and the control antibody 13G12, which does notinhibit integrin binding to fibronectin, were from Kenneth Yamada (NIH,Bethesda, MD) (Nagai et al., 1991). As a positive integrin-activationcontrol, surface-expressed unbound integrin dimers were activated with0.5 mM MnCl2 in DMEM (with 10% FBS) for 25 min (Cluzel et al., 2005;Orr et al., 2006).

Magnetic twisting cytometryMTC was used to mechanically perturb E-cadherin receptors, as describedpreviously (Barry et al., 2014; le Duc et al., 2010; Tabdili et al., 2012b).Beads were twisted with an oscillating field of 60 Gauss at 0.33 Hz, whichgenerates a shear stress (force/area) of 7.4 Pa. Prior reports showed that thestiffening increases with the applied twisting torque (Barry et al., 2015; leDuc et al., 2010). In controls, beads were coated with either poly-L-Lysine(PLL), blocking anti-E-cadherin antibody DECMA-1 (Sigma-Aldrich,St Louis, MO, U3254) (Ozawa et al., 1990), or with neutral (non-blocking)anti-E-cadherin antibody (Clone 76D5, from Barry Gumbiner, University ofVirginia, Charlottesville, VA). The neutral antibody binds the ectodomain,but does not alter the E-cadherin-binding affinity (Petrova et al., 2012;Shashikanth et al., 2015). In all cases, the bead coatings were optimized tobind cells and undergo displacements sufficient to measure the viscoelasticmoduli of bead–cell junctions (Barry et al., 2014, 2015; le Duc et al., 2010;Tabdili et al., 2012b; Twiss et al., 2012).

We quantified stiffening or traction changes in cells loaded with singlebeads, although cells with multiple beads were included infrequently. Someimages in this paper show cells with multiple bound beads, but only cellswith single attached beads were primarily included in the analyses (seeFig. S4, for example). In MTC, the specific modulus follows a log-normaldistribution, from which we obtained the mean and s.e.m. To determine thestiffness of individual cells in MTC and TFM measurements, for eachcondition, at least 40 cells were analyzed to obtain sufficient statistics.

Traction force microscopyTraction force microscopy (TFM) measurements used polyacrylamidehydrogels with Young’s moduli of 34 or 8.8 kPa (Tse and Engler, 2010).Proteins were immobilized on Sulfo-SANPAH-activated gels, by overnightincubations at 4°C with (1) 0.2 mg/ml collagen type 1 (Sigma, St Louis,MO), (2) fibronectin (0.2 mg/ml) or (3) PLL (0.2 mg/ml) in immobilizationbuffer (100 mMHEPES, 100 mMNaCl, and 5 mM CaCl2 at pH 8) (Tabdiliet al., 2012b). Fc-tagged E-cadherin extracellular domains were

immobilized as described previously (Tabdili et al., 2012b). After proteinimmobilization, the substrates were rinsed twice with 1× phosphate bufferedsaline (PBS), and sterilized by irradiation (365 nm), for at least 15 min,before seeding cells. E-cad-Fc substrates were blocked with 1% (w/v) BSA(in 1× PBS) for 20 min prior to rinsing and irradiation. MCF7 cells wereharvested using 3.5 mM EDTA in PBS containing 1% (w/v) BSA (Takeichiand Nakagawa, 2001), seeded at 5000–8000 cells/ml onto hydrogels, andallowed to adhere and spread for 6 h at 37°C under 5% CO2, onpolyacrylamide gels with embedded fluorescent microspheres. Theabsolute basal RMS traction force (BTF) was determined from fiduciarybead displacements, relative to the traction-free bead positions after cellremoval (lysis) (Butler et al., 2002). Constrained traction maps and the RMStraction stress (Pa; N/m2) were determined from the bead displacement maps(Butler et al., 2002).

Combined MTC and traction force microscopyMTC measurements were performed as described previously (Barry et al.,2014; le Duc et al., 2010; Tabdili et al., 2012b), except harvested cells(Takeichi and Nakagawa, 2001) were cultured on 34-kPa moduluspolyacrylamide gels with embedded fiduciary fluorescent beads (Tse andEngler, 2010). In combined MTC and TFM measurements, fiduciary beadpositions were imaged after incubating cells with ferromagnetic beads for40 min (∼one bead per cell). The twisting torque was generated with anoscillating (0.3 Hz) field of 70 Gauss, which generates a shear stress (force/area) of 8.4 Pa. The force-induced traction changes (ΔRMS, +Load) weredetermined from bead displacement maps (Butler et al., 2002) measuredimmediately before and after 2 min of force-loading (ΔRMS, +Load).Controls imaged beads after 2 min, but without load (ΔRMS, −Load).Figs 1B and 4A indicate the measurement time sequences. The reportedvalues for the BTF and ΔRMS traction are averages of results obtained withdifferent cells and replicate experiments. The ΔRMS tractions weredetermined from the change in traction forces upon stimulating cells withE-cadherin beads. By contrast, the absolute BTF was measured with cellsfree of ferromagnetic beads, and required cell lysis.

ImagingConfocal immunofluorescenceImmediately after bead loading, cells were fixed with 4% (w/v)paraformaldehyde for 15 min at room temperature. Control cells were notsubjected to bead twisting, but all conditions were otherwise identical. Cellswere permeabilized with 0.1% Triton X-100 for 5 min, blocked in 1% (w/v)BSA for 20 min, and stained with primary antibodies, followed by rinsing, andtreatment with secondary antibodies and phalloidin in 1% (w/v) BSA for 1 h.

Focal adhesions were visualized by immunofluorescence imaging ofvinculin at the basal plane. Vinculin was stained with monoclonalmouse anti-vinculin antibody (1:100, Sigma-Aldrich, V9131), and Alexa-Fluor-488-conjugated goat anti-mouse-IgG (1:200, Sigma-Aldrich,SAB4600388). F-actin was stained with Rhodamine–phalloidin(Invitrogen). Coverslips were mounted using ProLong Gold (Invitrogen),and images were acquired with a laser scanning confocal microscope (LSM700, Zeiss) with a 40× and 1.3 NA oil immersion objective and 488-nmand 555-nm lasers. Confocal images of immunostained vinculin wereanalyzed with a custom MATLAB code (from Brenton Hoffman, DukeUniversity), based on the watershed algorithm (Zamir et al., 1999). Focaladhesion size and number were quantified using the ‘Analyze Particles’function in ImageJ (NIH) with a minimum threshold area of 0.3 μm2

(Gonon et al., 2005).Cell areas were analyzed in ImageJ from acquired DIC images of

respective single MCF7 cells (n=80), with appropriate magnification-specific pixel-to-micron conversion factors. The cells were cultured inmedium containing integrin-blocking antibody (AIIB2) or Mn2+ for 25 minto inhibit or activate integrins, respectively. Cell areas were similarlyquantified after treatment with the fibronectin-blocking antibody 16G3.

Integrin activation assaysCell incubation with 20 μg/ml of GST–fibronectin-III9-11 peptide (fromMartin Schwartz, Yale School of Medicine, New Haven, CT) for 30 minwas used to assess α5β1 integrin activation (Orr et al., 2006; Tzima et al.,

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2001). For immunofluorescence imaging, cells were fixed and stained withmouse anti-GST antibody (1:100, Thermo Scientific, MA4-004), and withAlexa-Fluor-488-conjugated goat anti-mouse-IgG antibody (1:200, Sigma-Aldrich, SAB4600388). Cover slips were mounted with Prolong Gold(Invitrogen), and imaged by laser scanning confocal microscopy (ZeissLSM 700, 40×1.3 NA oil immersion objective). Total cell fluorescence wasquantified with ImageJ. Western blots similarly assessed peptide binding(Tzima et al., 2001). Controls used unperturbed cells (−Load). To activateintegrins, cells were bathed in DMEM (with 10% FBS) containing 0.5 mMMnCl2 for 25 min before E-cadherin-force loading (Cluzel et al., 2005; Orret al., 2006). The BTF and stiffening response of MnCl2-activated cells wasdetermined as described above.

PI3K activation and actin accumulation at force-loaded E-cadherin beadsPI3K activation was visualized with the GFP-tagged pleckstrin homologydomain of Akt (PH-Akt–GFP) (Collins et al., 2012). MCF7 or MDCKcells on collagen-coated glass were transiently transfected with PH-Akt–GFP (plasmid from Yingxiao Wang, UCSD, San Diego, CA). Theaccumulation of PH-Akt–GFP and of Phalloidin-stained actin with orwithout load (4 min) was quantified from background-subtractedfluorescence images (Barry et al., 2014, 2015). Background-subtractedmean fluorescence intensities (MFI), were normalized by measurementsunder no-load conditions (−Load). PH-Akt–GFP controls also usedcells transiently transfected with GFP (EGFP-N1, Clontech, MountainView, CA).

To confirm that PH-Akt–GFP accumulation reflected an increase inendogenous Akt phosphorylation during E-cadherin force-loading, we lysedthe serum-starved MCF7 monolayer after 4 min of E-cadherin beadtwisting. Lysates were blotted for Akt phosphorylation at S473 with apan-specific antibody (1:1000, R&D Systems, Minneapolis, MN: catalog#AF887) overnight at 4°C. Densitometry scans, using ImageJ quantifiedchanges in phospho-Akt levels, relative to PLL bead controls (Tzima et al.,2001). Actin was used as a loading control.

Dynamic fluorescence imaging of Src activityDynamic changes in Src kinase activity at cadherin adhesions werevisualized with a membrane-targeted FRET-based KRas-Src reporter(Yingxiao Wang, UCSD, San Diego CA) (Seong et al., 2013; Wanget al., 2005). Transiently transfected MCF7 cells were cultured on collagen-I-coated glass-bottomed dishes (Glass #1.5, Cell E&G, Houston TX) inPhenol-Red-free DMEM supplemented with 0.5% FBS, 1 mM sodiumpyruvate, and 1% (v/v) penicillin-streptomycin (Corning Cell Grow,Manassas, VA), for 6 h prior to the measurements, in order to reduceserum-induced Src kinase activity (McLachlan et al., 2007; Wang et al.,2005). Cells were then incubated with E-cadherin functionalizedferromagnetic beads for 40 min. An inverted Leica microscope (40×magnification, 0.55 NA air objective) coupled with a Dual View system(Optical Insights) acquired CFP and YFP images simultaneously with aCCD camera (Hamamatsu). The filter sets included CFP (excitation, S430/25, emission S470/30) and YFP (excitation, S500/20, emission S535/30).The emission filter set uses a 515-nm dichroic mirror to split the twoemission images, and the exposure time was 300 ms.

The time-dependent CFP-to-YFP ratio, which is proportional to activeSrc, was monitored for 30 s. The 30 s time frame includes ∼10 s prior toforce-loading, a time delay of ∼10 s between switching on the oscillatingmagnetic field and acquiring FRET images, and an additional 10 s duringbead twisting. The acquired stacked images were then uniformly contrast-enhanced, using ImageJ. Next, the ratios of the CFP to YFP emission in aregion of interest (ROI) around the beads were calculated, using a customMATLAB code (available on request). Each image was backgroundcorrected, using the custom MATLAB program, which negates manualbias when choosing an arbitrary region for background correction. Theprogram uniformly subtracted the minimal fluorescent pixel intensity fromeach of the acquired CFP and YFP images. Control data were acquiredunder identical conditions, but without bead loading. The CFP-to-YFPratios in the ROIs were normalized to the average CFP-to-YFP ratio priorto bond shear. In additional controls, cells were treated with the Srcinhibitor PP2.

StatisticsThe mean±s.e.m. are reported in the text and figure legends. P-values werecalculated from two-tailed Student’s t-tests, using Microsoft Excel, with*P<0.05 defining statistically significant differences. The standard errors fora set of replicate experiments were calculated using the pooled standarddeviations of each experiment, as indicated in the text and figure legends.The sample size was not computed when designing the study. The numberof beads analyzed to assess protein recruitment (confocalimmunofluorescence) or adaptive stiffening and the number of replicateexperiments required for statistical significance were based on extensiveprior studies. Similarly, the number of traction force measurements requiredfor statistical significance (n≥6) was based on prior experience (Barry et al.,2014) and published literature (Butler et al., 2002).

AcknowledgementsWe thank Saiko Rosenberg for technical assistance, Ryan Huang for codedevelopment, Prof. Hoffman (Duke University) for focal adhesion analysis software,and Kenneth Yamada for 16G3 and 13G12 antibodies.

Competing interestsThe authors declare no competing or financial interests.

Author contributionsI.M. designed and conducted experiments, analyzed data, edited and wrote themanuscript. J.W. designed, conducted, analyzed experiments and edited themanuscript. P.S. designed, conducted and analyzed results. X.K. assisted with dataanalysis, A.T. assisted with experiments, N.W. provided technical advice, D.E.L.designed experiments, analyzed data, and edited, wrote and finalized themanuscript.

FundingThis work was supported by the National Science Foundation [grant numbers CMMI1029871 and 1462739 to D.E.L.]; and National Institutes of Health [grant numberR01 GM097443 to D.E.L. and N.W.]. J.W. was supported by the Shen PostdoctoralFellowship. Deposited in PMC for release after 12 months.

Supplementary informationSupplementary information available online athttp://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.185447/-/DC1

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