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Vol. 18, Issue 11, 4519-4527, November 2007
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Department of Physiology, University of Massachusetts Medical School, Worcester, MA 01605
Submitted June 19, 2007;
Revised August 9, 2007;
Accepted August 27, 2007
Monitoring Editor: Carole Parent
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Beloussov et al., 1994) and wound healing (Hinz et al., 2001; Tomasek et al., 2002), and in pathological processes such as fibrosis and carcinogenesis (Paszek et al., 2005). Adherent cells seem capable of both responding to applied mechanical forces (Tzima et al., 2005) and applying contractile forces to probe mechanical properties of the environment (Discher et al., 2005). The downstream responses include changes in migration (Pelham and Wang, 1997; Sheetz et al., 1998; Lo et al., 2000), cell–cell interactions (Guo et al., 2006), proliferation (Wang et al., 2000; Nelson et al., 2005), differentiation (Engler et al., 2004, 2006), and apoptosis (Wang et al., 2000). For cultured adherent cells, focal adhesions are thought to mediate mechanosensing through integrin-mediated anchorage to the extracellular matrix (Larsen et al., 2006). The molecular repertoire of focal adhesions includes >100 adaptor and signaling proteins (Bershadsky et al., 2006), in addition to associated actomyosin bundles that provide mechanical forces for probing the environment (Choquet et al., 1997), inside-out signaling (Chrzanowska-Wodnicka and Burridge, 1996; Schwartz and Ginsberg, 2002), and cell migration (Ridley et al., 2003). However, few details are available concerning how these components work in concert to generate the responses to mechanical signals.
Several aspects have emerged recently as the key features of integrin-mediated mechanosensing. First is the increase in cortical organization and contractility in response to mechanical signals. Application of mechanical forces to integrin-associated beads causes "reinforcement" of cortical resistance (Wang et al., 1993; Choquet et al., 1997), whereas pushing the dorsal cortex or pulling the focal adhesion causes enlargement of focal adhesions (Riveline et al., 2001). In addition, substrate rigidity induces the appearance of large focal adhesions, prominent stress fibers, and strong traction forces (Pelham and Wang, 1997; Discher et al., 2005). Such responses are often accompanied by the induction or enhancement of centripetal cortical movements, which may manifest as flow of actin arcs in lamella or retrograde movements of integrin-associated beads (Felsenfeld et al., 1996). Chemical signals involved in these processes include the activation of small GTPases, including Rho and Rac (Chrzanowska-Wodnicka and Burridge, 1996), and tyrosine kinases, including Src family kinases (Suter and Forscher, 2001) and focal adhesion kinase (FAK; Burridge et al., 1992; Schmidt et al., 1998), as well as the entry of Ca2+ through stretch-activated ion channels (Guharay and Sachs 1984; Lee et al., 1999; Munevar et al., 2004).
A more intriguing response is the redistribution of some focal adhesion proteins upon mechanical stimulations. Several proteins, including zyxin and vasodilator-stimulated phosphoprotein (VASP), are known to localize to a variable extent at focal adhesions (Crawford et al., 1992; Rottner et al., 2001), or to shuttle between focal adhesions and the nucleus (Nix and Beckerle, 1997; Aplin and Juliano, 2001). Recent studies further indicated that mechanical stimulation can induce the change in distribution of zyxin from focal adhesions to either actin stress fibers in cultured fibroblasts (Yoshigi et al., 2005), or the nucleus of vascular smooth muscle cells (Cattaruzza et al., 2004). The latter may be related to the suspected ability of zyxin to regulate gene expression (Degenhardt and Silverstein, 2001). However, the mechanism of redistribution from focal adhesions is unclear, because these studies showed only the steady-state localization after mechanical stimulation.
To reach a better understanding of mechanosensing, we have examined the dynamics of actin and several focal adhesion proteins suggested to play a role in mechanosensing, by combining live cell imaging, fluorescence recovery after photobleaching (FRAP), substrate engineering, and mechanical stimulation. We show that actin filaments and some focal adhesion proteins, including zyxin and VASP undergo retrograde fluxes in response to the state of cell migration and mechanical signals. The common aspect of conditions that stimulate the flux lead us to propose a mechanism that controls structural assembly at focal adhesions based on migration- or force-induced structural shear within the focal adhesion. The flux may in turn provide feedback regulation of force production in relation to cell migration, and/or facilitate the delivery of long-range signals released by focal adhesions.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Red Fluorescent protein (RFP)-zyxin was constructed by recloning the enhanced green fluorescent protein (EGFP) construct (Rottner et al., 2001) into the pDsRed1-N1vector (Clontech, Palo Alto, CA). RFP-VASP and EGFP-tagged paxillin (Rottner et al., 2001), and yellow fluorescent protein (YFP)-vinculin were kindly provided by Dr. Juergen Wehland (Deutsche Forschungsgemeinschaft, Braunschweig, Germany), and YFP-dSH2 was kindly provided by Dr. Benny Geiger (Weitzmann Institute; Israel; Kirchner et al., 2003). EGFP-tagged N-fragment of tensin was kindly provided by Dr. Shin Lin (University of California, Irvine). EGFP-actin was purchased from Clontech. NIH3T3 and NRK were nucleofected using Amaxa Nucleofector I and kit R (Amaxa Biosystems, Gaithersburg, MD).
Y-27632 (Calbiochem, San Diego, CA) was prepared as 10 mM stock solution in phosphate-buffered saline (PBS). Stock solutions of 100 mM blebbisatin (Calbiochem), 100 mM genistein (Calbiochem), 10 mM PP2 (Calbiochem), and 10 mM PP3 (Calbiochem) were prepared in DMSO and stored at –20°C. All reagents were diluted from the stock solution into medium immediately before use. The final concentration was 50 µM for Y-27632, 5 µM for blebbisatin, 100 µM for genistein, and 20 µM for PP2 and PP3.
Preparation of Micropatterned Substrates and Local Mechanical Stimulation by Stretching
Patterned substrates with 40- x 40-µm2 islands were generated by spin coating of SU-8 2002 photoresist (MicroChem, Newton, MA) on a dried polyacrylamide film, which was prepared with 50 µl of 10% acrylamide, 0.26% bis-acrylamide spread over a circular area 32 mm in diameter on a glutaraldehyde-activated 40- x 50-mm2 coverslip (Pelham and Wang, 1997). The thickness of the film was 6.2 ± 1.0 µm after air-drying. After further drying at 95°C for 2 min, the surface was spin coated with 200 µl of SU-8. The photoresist was prebaked at 95°C for 2 min and exposed to 365-nm UV from a high-flux UV LED (OTLH-0480-UV) for 1 min through an opaque photomask with an array of 40- x 40-µm2 clear squares (CAD/Art Services, Bandon, OR). After postbaking at 95°C for 2 min, the photoresist was developed in SU-8 developer (MicroChem) for 1 min, rinsed with ethanol, and air-dried. A second layer of 8% polyacrylamide was polymerized on top of the first layer at a volume of 15 µl, to further reduce cell adhesion to areas outside the SU-8 islands (Mader et al., 2007). Omitting bis-acrylamide and N,N,N',N'-tetramethylethylenediamine (TEMED) generated linear polyacrylamide chains that grew off the dried layer of polyacrylamide without covering the SU-8 islands. We subsequently found that the first layer of polyacrylamide may be omitted, by directly spin coating SU-8 on a glutaraldehyde-activated coverslip. After photolithography to generate the island pattern of SU-8 as described above, the remaining area was blocked by grafting 8% linear polyacrylamide (with TEMED) directly onto the exposed glass surface.
Micropatterned polyacrylamide gels were prepared by activating the gel surface with 50 mM sulfosuccinimidyl 6 (4'-azido-2'-nitrophenyl-amino) hexanoate (Pierce Chemical, Rockford, IL) in 200 mM HEPES, pH 8.5, as described previously (Wang and Pelham, 1998). Excess liquid on the surface was removed by blowing with nitrogen gas, and the surface was stamped for 1 min with a patterned polydimethylsiloxane stamp inked with collagen. The stamp was made as described by LeDuc et al. (2002), by using a molding made from the SPR 220-3.0 photoresist (Shipley, Marlboro, MA), patterned with the same photomask as for the SU-8 substrates. The stamp was soaked in 100 µg/ml type I collagen (USB, Cleveland, OH) for 45 min and rinsed with PBS briefly. Excess liquid was blown off with nitrogen gas before the stamp was applied to the gel surface. The same procedure was applied to gels with 10% acrylamide/0.26% bis-acrylamide (Young's modulus >50 kPa; referred to as the stiff gel), and gels with 10% acrylamide gel/0.03% bis-acrylamide (Young modulus 
5 kPa; referred to as the soft gel).
Local stretching forces were applied to NIH3T3 cells plated on polyacrylamide gel of 5% acrylamide/0.08% bis-acrylamide as described previously (Lo et al., 2000). Briefly, glass capillary tubing was pulled into needles with a vertical micropipette puller (David Kopf Instruments, Tujunga, CA). The tip of the needle was then melted and shaped with a microforge (Narishige, East Meadow, NY). The blunted needle tip was gently pressed into the gel near the cell and moved away from the cell with a micromanipulator (Leitz, Wetzlar, Germany), to generate pulling forces. The manipulation caused an 10% overall increase in cell length, which was prominent at the end of the cell proximal to the needle but became undetectable at the opposite end (Lo et al., 2000). Based on an estimated Young's modulus of 1–5 kPa for fibroblasts (Engler et al., 2004), the induced cell strain reflected an applied stress of 1–5 x 10–10 N/µm2 cellular cross-sectional area.
Immunofluorescence Staining and Imaging, FRAP, and Image Analysis
Cells were fixed with 4% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA) and permeabilized with 0.2% Triton X-100 in PBS for 10 min. Samples were blocked with 1% bovine serum albumin/PBS for 15 min at room temperature, and then they were incubated for 1 h at 37°C with 1:200 dilution of primary antibodies, against fibronectin (F3648; Sigma-Aldrich), paxillin (H-114; Santa Cruz Biotechnology, Santa Cruz, CA), vinculin (clone VIN11-5; Sigma-Aldrich), FAK (C-20; Santa Cruz Biotechnology), FAK-pY397 (Santa Cruz Biotechnology), phosphotyrosine (clone 4G10; Upstate Biotechnology, Lake Placid, NY), or zyxin (Synaptic Systems, Gottingen, Germany). Alexa dye-conjugated anti-rabbit or anti-mouse secondary antibodies were obtained from Invitrogen (Carlsbad, CA) and applied at a dilution of 1:400. Alexa 488-conjugated phalloidin (Invitrogen) was applied following manufacturer's procedure. Images were collected with a 100x Plan-Nufluor numerical aperture (N.A.) 1.3 oil lens, and a low-light electron multiplication charge-coupled device (CDD) camera iXon DV887DCS-BV (Andor Technology, South Windsor, CT), or a conventional cooled CCD camera (NTE/CCD-512-EBFT; Princeton Instruments, Trenton, NJ), attached to an inverted microscope (Axiovert 200M; Carl Zeiss, Thornwood, NY), equipped with a 100x Plan-Nufluor N.A. 1.3 oil lens and a stage incubator for live cell time-lapse imaging. Fluorescent images were recorded every 2 min.
Green fluorescent protein (GFP) fluorescence was bleached with an Argon ion laser (Lexel, Palo Alto, CA), by using 10-ms pulses of the 457-nm line at 50 mW. To bleach RFP-zyxin, we used the 514-nm line at a power of 200 mW and 10-ms duration. Fluorescence images before bleaching and during fluorescence recovery were captured every 5 s with spinning-disk confocal optics (QLC100; Solamere Technology, Salt Lake City, UT), using 488 nm for the excitation of GFP and 532 nm for the excitation of RFP and the corresponding filter set provided by the manufacturer. To allow simultaneous bleaching and imaging, the epifluorescence filter cube was mounted with a 525-nm dichroic mirror (525DRLP; Omega Optical, Brattleboro, VT) for RFP signals, or a 475-nm dichroic mirror (z457rdc; Chroma Technology, Brattleboro, VT) for GFP signals. These mirrors allowed the excitation and emission wavelengths for confocal imaging to pass through while directing the bleaching beam at the sample. Conventional fluorescence imaging was carried out with the Omega Optical 100-3 filter set for GFP and Chroma Technology 41002 filter set for RFP. Fluorescence intensity was measured by integrating signals within a user-defined, closed boundary of any shape, using custom software written in C++. Kymographs also were produced and analyzed using custom software, which allowed the user to define the location, orientation, length, and width of a rectangular area in an image series. Images within the defined area were then extracted from the series, reoriented vertically, and combined side by side into a kymograph. All experiments involving single cell time-lapse and immunofluorescence assays have been performed at least three times with consistent results. Statistical significance was determined using Student's t test.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Rottner et al., 2001), may reflect different states of focal adhesions in responses to mechanical signals and/or cell migration. We observed two discrete patterns of the distribution of these proteins near focal adhesions in the lamella region of NIH3T3 cells. In most cells they were confined to the typical plaque-shaped focal adhesions; however, in a small percentage (10% of 100 cells examined) of cells they were present both at focal adhesions and along a "comet tail" that extended away from some focal adhesions toward the cell center. An example of cells in this subpopulation is shown in Figure 1A.
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We noticed that focal adhesions with zyxin tails were located mainly in relatively stationary NIH3T3 cells and in NRK epithelial cells, which were not as motile as NIH3T3 cells. In addition, plots of the number zyxin tails in NIH3T3 cells against the speed of migration showed an inverse relationship (Figure 2A). The dependence on cell migration was further tested by immobilizing NIH3T3 cells on patterned substrates consisting of 40- x 40-µm2 islands of SU-8 photoresist on an otherwise nonadhesive polyacrylamide surface (Figure 2B). Constraining NIH3T3 cells on such substrates induced an increase in the percentage of cells with zyxin tails, from <10% of cells on glass or uniform SU-8 surfaces to 3040% (50 cells examined) on SU-8 islands. Moreover, 86 ± 6% of the tails were located within 23° from the four corners of these square-shaped cells (Figure 2C; based on 250 tails in 10 cells), where ruffling activities and strongest traction forces were concentrated (Parker et al., 2002; Wang et al., 2002).
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Characterization of Retrograde Focal Adhesion Fluxes
We compared the immunofluorescence localization of zyxin with other focal adhesion proteins, including paxillin, vinculin, and FAK. The tail of zyxin and VASP extended far beyond focal adhesions as defined by vinculin or paxillin (Figure 4, A and B), although limited fluxes of these proteins have been detected with total internal reflection fluorescence optics (Brown et al., 2006; Hu et al., 2007). Interestingly, FAK extended further into the tail than vinculin or paxillin (Figure 4C). These observations may reflect differential affinities of focal adhesion proteins with the actin bundle versus the structural scaffold at focal adhesions.
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; Zamir et al., 2000). Immunofluorescence staining indicated that fibronectin colocalized only partially with zyxin tails (Figure 5A). In addition, ventral fibronectin-containing fibrils were localized within a single optical section against the substrate (data no shown), whereas some zyxin tails extended over several optical sections (Figure 5B). Moreover, unlike fibrillar adhesions (Zamir et al., 2000), zyxin tails disappeared rapidly upon the inhibition of contractile force as described below. However, EGFP tagged N-fragment of tensin, which associates with fibrillar adhesions, did localize to zyxin tails (Supplemental Movie 5), suggesting that focal adhesion fluxes may contribute to the formation of fibrillar adhesions.
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; Wang et al., 2002). Therefore, mechanical forces, either generated internally by myosin II or applied externally through focal adhesions, may play a role in inducing the fluxes. Y-27632, an inhibitor of the Rho-dependent kinase, and blebbistatin, an inhibitor of myosin II ATPase, both caused strong inhibition of traction forces (Beningo et al., 2006), and focal adhesion fluxes, while reducing zyxin tails to small dot-like structures (Figure 6, A–C, and Supplemental Movie 6). Conversely, application of pulling forces to cells plated on flexible polyacrylamide substrates stimulated the appearance of focal adhesion fluxes (Figure 6, D–G).
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). To determine the effect of substrate rigidity on focal adhesion flux while maintaining a similar cell shape and spread size, we applied soft lithography to polyacrylamide substrates to generate extracellular cell matrix (ECM)-coated islands of different stiffness (Wang et al., 2002). NIH3T3 cells showed prominent zyxin tails on islands of stiff polyacrylamide as they did on photoresist (n = 10 cells; Figure 7A and Supplemental Movie 7), but they formed only small focal adhesions with few zyxin tails on soft substrates (n = 10 cells; Figure 7, B and C).
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; Schmidt et al., 1998), we asked whether tyrosine phosphorylation might be involved in regulating focal adhesion fluxes. Immunofluorescence localization showed that phosphotyrosine was concentrated only at focal adhesions irrespective of the presence of zyxin tails (Figure 8A). Consistent results were obtained with live cells expressing YFP-dSH2, a probe for sites of tyrosine phosphorylation (Kirchner et al., 2003; Figure 8B and Supplemental Movie 8). Interestingly, although FAK was found both at focal adhesions and along the flux, FAK phosphorylated at Tyr397, which is known to activate its kinase activity (Schaller et al., 1994), was localized predominantly at focal adhesions (Figure 8C), consistent with the localization of phosphotyrosine.
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30 min after treatment, without causing disappearance of focal adhesions as shown by RFP-zyxin (Figure 9, A and B, and Supplemental Movie 9), or immunofluorescence (Figure 9G). PP2, which selectively inhibits Src family kinases, also reduced zyxin flux without disassembling focal adhesions (Figure 9, C and D), whereas PP3, an inactive analogue of PP2, had no significant effect (data not shown).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Hotulainen and Lappalainen, 2006).
The maintenance of a steady-state flux requires treadmilling of actin subunits, where subunits are continuously added at focal adhesions and removed at the proximal ends. The process may be regulated by the rates of actin nucleation, elongation, dissociation, and possibly myosin-dependent forces that pull on actin filaments (Medeiros et al., 2006). The dependence of the flux on cell migration and mechanical signals explains why prominent stress fibers are present only in relatively immotile cells (Herman et al., 1981; Tomasek et al., 1982), and in cells on stiff substrates (Pelham and Wang, 1997).
The responses to cell migration and substrate compliance are likely related, because conditions that favor the focal adhesion flux share a common feature, that the actin cytoskeleton undergoes large retrograde displacement relative to the substrate anchorage site, as a result of either contractile forces pulling against a stationary anchorage site (Figure 10D), or anterograde deformation of a flexible substrate (Figure 10C). Conversely, conditions that inhibit the flux, including migration of the cell body (Figure 10A), and large substrate deformability (Figure 10B), are likely to cause a smaller retrograde displacement of the actin cytoskeleton relative to the substrate anchorage site. Therefore, the regulation of the flux may be explained by a structural shear model (Figure 10), where focal adhesion proteins at different distances from the membrane anchorage site are displaced by different distances along the direction of migration or mechanical forces under different conditions, which may in turn regulate tyrosine phosphorylation and actin assembly, e.g., by exposing tyrosine kinase binding domains and/or activating the substrates (Sawada et al., 2006).
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; Hu et al., 2007), and they may serve as a feedback mechanism that adjusts structural assembly, possibly coupled with the generation of contractile forces, in response to the needs of cell migration. The mechanism will provide the strongest propulsive forces for slow-moving cells on stiff substrates, which are known to induce strong resistive adhesions as well as traction forces (Guo et al., 2006). A second potential function of a regulated flux at focal adhesions is to serve as a slippage clutch for cell migration (Smilenov et al., 1999; Hu et al., 2007). Retrograde flux may reflect disengagement of the clutch, when contractile forces are consumed in the transport of actin filaments rather than transmitted to the substrate for cell migration.
Focal adhesion fluxes may also be involved in the formation of fibrillar adhesions and the assembly of a fibronectin network to guide cell migration in tissues (Boucaut et al., 1990; Darribere and Schwarzbauer, 2000). Although the flux does not correlate exactly with fibrillar adhesions in their localization and sensitivity to cell contractile forces (Zamir et al., 2000), the colocalization of zyxin with tensin and the partial colocalization with fibronectin in NIH3T3 suggest that the flux may serve as a precursor for the formation of fibrillar adhesions in fibroblasts (Zamir et al., 2000; Figure 4D), through recruitment and retrograde transport of 
5
1 integrins along the flux followed by fibronectin binding (Pankov et al., 2000). Because focal adhesion fluxes also exist in epithelial cells, which produce different extracellular matrix proteins than fibroblasts, the fluxes may play a general role in organizing the extracellular matrix. Further evaluation of the role of fluxes in three-dimensional environment may elucidate this possible function of fluxes.
An alternative function of the focal adhesion flux is to facilitate the transport of signals from focal adhesions in response to mechanical cues, as suggested by impaired responses of zyxin–/– cells to ECM anchorage and mechanical stimulations (Yoshigi et al., 2005; Hoffman et al., 2006). Although focal adhesions have been recognized as an important source of signals with multiple downstream effects such as activation of the mitogen-activated protein kinase pathway and eventually gene expression (Giancotti and Ruoslahti, 1999), how these signals propagate is unclear. A diffusion-like mechanism seems inefficient within a crowded cytoplasmic environment, and transport along the flux of actin filaments may serve as a much more effective mechanism. Particularly relevant is the relocation of zyxin from focal adhesions into the nucleus of vascular smooth muscle cells in response to mechanical signals, where zyxin itself may serve as a transcription activator (Degenhardt and Silverstein, 2001). In addition, retrograde flux of FAK may have a profound effect on the state of phosphorylation at focal adhesions and actin filaments, which in turn affects the maturation and turnover of focal adhesions (Webb et al., 2004). Our results further suggest that the flux of focal adhesion proteins requires tyrosine phosphorylation, which, in conjunction with the small GTPases Rac and Rho, represents the primary molecular switches in response to integrin–ECM anchorage and mechanical signals (Burridge et al., 1992; Schmidt et al., 1998). Tyrosine phosphorylation regulates the dynamic turnover of focal adhesion proteins through a number of substrates including paxillin, p130Cas, and FAK (Carragher and Frame, 2004). In addition, Rho activities show a transient decrease followed by an increase upon integrin signaling, which may be responsible for the activation of formin that in turn nucleates the assembly of actin filaments at focal adhesions (Wallar and Alberts, 2003). Understanding the regulation of focal adhesion flux is likely to shed light on a wide range of activities that involve cell migration and mechanosensing.
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ACKNOWLEDGMENTS |
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Footnotes |
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The online version of this article contains supplemental material at MBC Online (http://www.molbiolcell.org). 
Address correspondence to: Yu-li Wang (yuli.wang{at}umassmed.edu).
Abbreviations used: FAK, focal adhesion kinase; FRAP, fluorescence recovery after photobleaching; VASP, vasodilator-stimulated phosphoprotein.
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