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Mechanosensing through focal adhesion-anchored intermediate filaments Martin Gregor,*,‡,1 Selma Osmanagic-Myers,*,1 Gerald Burgstaller,*,2 Michael Wolfram,* Irmgard Fischer,* Gernot Walko,* Guenter P. Resch,† Almut Jörgl,* Harald Herrmann,§ and Gerhard Wiche*,3 *Department of Biochemistry and Cell Biology and †Electron Microscopy Campus Science Support Facility, Max F. Perutz Laboratories, University of Vienna, Vienna, Austria; ‡Department of Transgenic Models of Disease, Institute of Molecular Genetics of the ASCR, Prague, Czech Republic; and §Functional Architecture of the Cell, German Cancer Research Center, Heidelberg, Germany Integrin-based mechanotransduction involves a complex focal adhesion (FA)-associated machinery that is able to detect and respond to forces exerted either through components of the extracellular matrix or the intracellular contractile actomyosin network. Here, we show a hitherto unrecognized regulatory role of vimentin intermediate filaments (IFs) in this process. By studying fibroblasts in which vimentin IFs were decoupled from FAs, either because of vimentin deficiency (V0) or loss of vimentin network anchorage due to deficiency in the cytolinker protein plectin (P0), we demonstrate attenuated activation of the major mechanosensor molecule FAK and its downstream targets Src, ERK1/2, and p38, as well as an up-regulation of the compensatory feedback loop acting on RhoA and myosin light chain. In line with these findings, we show strongly reduced FA turnover rates in P0 fibroblasts combined with impaired directional migration, formation of protrusions, and up-regulation of “stretched” high-affinity integrin complexes. By exploiting tension-independent conditions, we were able to mechanistically link these defects to diminished cytoskeletal tension in both P0 and V0 cells. Our data provide important new insights into molecular mechanisms underlying cytoskeleton-regulated mechanosensing, a feature that is fundamental for controlled cell movement and tumor progression.—Gregor, M., Osmanagic-Myers, S., Burgstaller, G., Wolfram, M., Fischer, I., Walko, G., Resch, G. P., Jörgl, A., Herr-

ABSTRACT

Abbreviations: 3D, 3-dimensional; CB, cytoskeleton buffer; DMEM, Dulbecco’s modified Eagle medium; DTSSP, 3,3=dithio-bis-succinimidylproprionate; ECM, extracellular matrix; EGF, epidermal growth factor; EGFP, enhanced green fluorescent protein; FCS, fetal calf serum; FA, focal adhesion; FAK, focal adhesion kinase; FN, fibronectin; GA, glutaraldehyde; GTP, guanosine-5=-triphosphate; HBSS, Hank’s buffered salt solution; IF, intermediate filament; LPA, lysophosphatidic acid; mAb, monoclonal antibody; MAP, mitogen activated protein; MLC, myosin light chain; OA, okadaic acid; P0, plectin-deficient; PAGE, polyacrylamide gel electrophoresis; PBS, phosphate-buffered saline; PDGF, platelet-derived growth factor; PL, polylysine; SDS, sodium dodecyl sulfate; SYN, synergy; V0, vimentin-deficient; WT, wild-type 0892-6638/14/0028-0715 © FASEB

mann, H., Wiche, G. Mechanosensing through focal adhesion-anchored intermediate filaments. FASEB J. 28, 715–729 (2014). www.fasebj.org Key Words: vimentin 䡠 plectin 䡠 integrin activation 䡠 cell motility focal adhesions (fas) reside at the crossroad between the cytoskeleton and the extracellular matrix (ECM). They have the ability to sense forces acting on them, which come either from the contractile actomyosin apparatus in the interior of the cell, or from the ECM. Increased rigidity of the substrate and/or cytoskeletal tension (prestress) result in the reinforcement of adhesions, which become larger in order to resist the applied force (1). The mechanism behind this phenomenon is based primarily on the stretching of proteins, which, as for integrins, results in a transition from a low-affinity to a high-affinity binding state or an unmasking of additional binding sites for vinculin, as in the case of talin (2, 3). A multitude of associated signaling elements, such as focal adhesion kinase (FAK), Src, and RhoA GAPs/GEFs, require tension in order to get fully activated (4, 5). Among many downstream-targets of FAK is RhoA-GAP, which, by down-regulating RhoA, reduces contractility and thus plays a crucial role in the regulation and fine-tuning of FA growth (6, 7). This precisely regulated balance between applied forces and cellular responses appears to be disturbed in the absence of FAK, leading to overactivation of RhoA, more robust FA with 1

These authors contributed equally to this work. Current address: Comprehensive Pneumology Center, Ludwig Maximilians University Munich and Helmholtz Zentrum München, Munich, Germany. 3 Correspondence: Department of Biochemistry and Cell Biology, Max F. Perutz Laboratories, University of Vienna, Dr. Bohrgasse 9, A-1030 Vienna, Austria. E-mail: gerhard. [email protected] doi: 10.1096/fj.13-231829 This article includes supplemental data. Please visit http:// www.fasebj.org to obtain this information. 2

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reduced turnover, and attenuated migration and directionality (8, 9). Besides actin filaments and microtubules (10, 11), vimentin intermediate filaments (IFs), too, have been known for quite a long time to associate with FAs (12). However, the precise role of this interaction is still not understood. Overexpression of vimentin enhances FA dynamics (13), and, in line with this, it was shown that in vimentin-deficient (V0) fibroblasts, FAs were more robust, with actin stress fibers becoming increasingly bundled and lacking geodome structures (14). Consequently, in the absence of vimentin, fibroblasts migrated much more slowly than their wild-type (WT) counterparts and exhibited severely compromised directional migration, leading to a delay in the woundhealing ability of mice (14, 15). When actin stress fibers were disrupted by means of cytochalasin D, WT and V0 cells no longer showed differences in motility, delineating the alterations in actin networks and FAs observed in the absence of vimentin as the cause for migration defects (14). Several recent reports focusing on the cytolinker protein plectin demonstrated a crucial function of the protein in mediating the anchorage of vimentin IFs to FAs (16, 17). Plectin is a major IF-associated protein capable of interlinking IFs of various types and mediating their interaction with actin filaments and microtubules (18). The protein plays a key role in IF-network organization in different cell types, including keratinocytes, myocytes, and fibroblasts (19). Interestingly, plectin-deficient (P0) fibroblasts, similar to V0 cells, exhibited enlarged FAs, prominent actin stress fibers, compromised motility, and delayed wound healing in mice, leading to the suggestion that similar compensatory modes of action operate in both cell systems (20, 21). Previous reports have shown that both P0 and V0 cells exhibit severe reduction in cellular stiffness (prestress) and consequently exert significantly reduced traction forces on ECM (15, 22, 23). These findings are consistent with the so-called tensegrity model, in which cytoskeletal tension requires not only intact actomyosin contractile elements and microtubules acting as “compression struts” but IFs as “tensile stiffeners,” as well (24). Notably, using acrylamide to selectively disrupt the IF network in WT fibroblasts, Wang and Stamenovic (23) provided evidence that the reduction in cytoskeletal tension was indeed due to the absence of IFs and not to alterations of other cytoskeletal networks. Finally, in both cell types (P0 and V0), signal transmission appeared to be compromised, as suggested by longtime force propagation experiments and collagen contraction measurements (14, 22). In the present study, we exploited P0 and V0 cell systems to explore how the anchoring of vimentin to FAs regulates the complex mechanosensing machinery associated with these structures. Our findings implicate a hitherto unrecognized function of vimentin IFs as force transducers, providing efficient integrin-mediated activation of the major mechanosensor FAK and 716

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its downstream tension-dependent signaling cascade. Thus, in line with attenuated FAK-activity in P0 fibroblasts, we show strongly reduced FA turnover rates combined with impaired directional migration, formation of protrusion and, notably, up-regulation of highaffinity integrin complexes typical of their “stretched” status. By plating P0 and V0 cells on mutated fibronectin (FN), blocked in its ability to sense tension, these deficits could be traced to diminished cytoskeletal tension, causing perturbations in mechanotransduction. Our data provide novel important insights into the mechanisms of how the cytoskeleton regulates FAK mechanosensing and coordinated cell motility.

MATERIALS AND METHODS Cell culture and transient transfection Immortalized fibroblasts (cell lines) were derived from newborn p53⫺/⫺ plectin WT and P0 mice (25) and used at passages ⱕ 15. Primary cells were derived in a similar manner from mice that were p53⫹/⫹. For the statistical evaluation of data, in general, the cell lines were used. However, all experiments (except for RhoA-pulldown and FA-turnover assays) were confirmed with corresponding primary cell cultures, and data are shown throughout Results and in Supplemental Material. V0 fibroblast cell lines (26) were further subcloned to eliminate desmin- or keratin-expressing cells and were used at maximum passage 30. Cells were grown for 12 h on uncoated plastic dishes unless otherwise stated in the text. Transient transfection was performed with a plasmid encoding enhanced green fluorescent protein (EGFP)tagged paxillin (a kind gift of M. Gimona, University of Salzburg, Salzburg, Austria), using Fugene6 reagent (Roche Applied Science, Mannheim, Germany), according to the manufacturer’s instructions. Okadaic acid (OA) treatment and heat-stress assay For OA treatment, subconfluent cells grown overnight (⬃12 h) were treated with 0.1 ␮g/ml OA (Sigma-Aldrich, St. Louis, MO, USA) for different time periods (see text). For heatstress assays, cells were incubated for 20 min in a water bath at 45°C, after replacement of the cultivation medium with prewarmed Dulbecco’s modified Eagle medium (DMEM) supplemented with 20 mM HEPES (pH 7.4). After heat exposure, the growth medium was removed and replaced by new medium; cells were kept for 2 h at 37°C before being fixed and processed for immunofluorescence microscopy. Immunofluorescence microscopy Cells grown overnight were fixed with methanol and processed for immunofluorescence microscopy, as described previously (27). The following primary antibodies were used: anti-plectin antiserum 46 (1:400; ref. 25), anti-plectin isoform 1f antiserum (1:50; ref. 28), and affinity-purified goat antivimentin antibodies (1:800; ref. 29), kindly provided by P. Traub (University Bonn, Bonn, Germany); mouse monoclonal antibodies (mAbs) to vinculin (1:400; Sigma-Aldrich); mouse mAbs to paxillin (1:1000; BD Biosciences, San Jose, CA, USA); rat mAbs to integrin ␤1 (1:100, clone MB1.2; EMD-Millipore, Billerica, USA); and affinity-purified antibodies to actin (1:100, A 2066; Sigma-Aldrich). As secondary

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antibodies, we used donkey anti-mouse IgM Texas Red, donkey anti-mouse Cy5, donkey anti-mouse Rhodamine Red-X, and donkey anti-goat Cy2 (1:1000; all from Jackson ImmunoResearch Laboratories, West Grove, PA, USA). Specimens were viewed in an LSM 510 laser-scanning microscope (Carl Zeiss MicroImaging, Jena, Germany). For three-dimensional (3D) reconstruction of vimentin filament networks, Z stacks of immunostained fibroblasts were first deconvolved with Huygens Essential 4.0.0 software (Scientific Volume Imaging, Hilversum, The Netherlands) using blind deconvolution. Three-dimensional surface reconstructions of specimens were then rendered using Imaris 7.3.1 software (Bitplane, Zurich, Switzerland). For TIRF imaging, immunolabeled specimens were mounted in phosphate-buffered saline (PBS) and visualized with an HCX PL APO ⫻100, 1.46 NA, TIRF objective using a Leica AM TIRF MC system (Leica Microsystems, Bannockburn, IL, USA). The TIRF angle for all the channels was 69.43°. All images were taken with 90-nm penetration depth for all channels. The laser angle into the objective was 90°. Postacquisition processing was performed with Huygens Essential and Metamorph 7.7 software (Molecular Devices, Sunnyvale, CA, USA). Whole-mount electron microscopy Fibroblasts were grown on formvar-coated coverslips and processed directly for negative-staining whole-mount electron microscopy (30 –32). Briefly, cells were rinsed with prewarmed PBS, extracted, and fixed for 1 min with 0.5% glutaraldehyde (GA), 0.75% Triton X-100 in cytoskeleton buffer (CB; 150 mM NaCl, 5 mM MgCl2, 5 mM EGTA, 5 mM glucose, and 10 mM MES, pH 6.1), and fixed with 1.0% GA supplemented with 10 ␮g/ml phalloidin in CB for ⱖ1 h. Specimens to be labeled for vimentin were sequentially incubated with affinity-purified goat anti-vimentin antibodies (1:800), rabbit anti-goat IgG (1:25), 10-nm gold goat antirabbit IgG (1:25), and postfixed for 30 min in 1% GA in CB. Specimens to be depleted of actin and microtubules were high-salt extracted for enrichment of IFs and further processed according to Small et al. (30). All samples were negatively stained with 4% sodium silicotungstate/0.25% trehalose (pH 7) and visualized using a Jeol 1210 transmission electron microscope (Jeol Ltd., Akishima, Japan) operated at 80 kV acceleration voltage. Digital images were acquired on a SIS Morada CCD camera (Olympus-SIS, Tokyo, Japan). For measurement of filament diameter, random positions on vimentin filaments were selected. Cell lysis and preparation of cell fractions For direct lysis, subconfluent fibroblast cultures, washed twice with PBS, were overlaid with Laemmli sample buffer supplemented with 1 mM Na2VO3, and 1⫻ phosphatase inhibitor cocktail 1 (Sigma-Aldrich). For fractionation, scraped-off cells were collected by centrifugation and gently resuspended in 300 ␮l of ice-cold 0.5% Triton X-100, 10 mM PIPES (pH 7.4), 300 mM sucrose, 100 mM NaCl, 3 mM MgCl2, 3 mM EDTA, 0.2 mM DTT, and protease inhibitor mix (see above), followed by incubation with end-over-end agitation for 20 min at 4°C. After centrifugation (15,800 g, 1 min), the supernatant (cytosolic and membrane fractions) was collected, and pellets (cytoskeletal fraction) were dissolved in sample buffer. Aliquots of total cell lysates or cell fractions containing equal amounts of total protein were subjected to sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE), and, after immunoblotting using peroxidase-coupled secondary antibodies, protein bands were visualized by exposure to X-ray film. Quantitation of bands was performed as described MECHANOSENSING THROUGH VIMENTIN

previously (33). For immunoblotting, the following primary antibodies were used: anti-vimentin (1:10,000), mouse mAbs to ␣-tubulin (B-5-1-2, 1:10,000; Sigma-Aldrich), mouse mAbs to p38 (sc-535, 1:500; Santa Cruz Biotechnology, Santa Cruz, CA, USA), rabbit mAbs to phospho-Thr-180/Tyr-182 p38 (3D7, 1:1000; Cell Signaling Technology, Boston, MA, USA), mouse mAbs to ERK2 (D-2, 1:800; Santa Cruz Biotechnology), mouse mAbs to phospho-Tyr-204 ERK1/2 (E-4, 1:10,000; Santa Cruz Biotechnology), mouse mAbs to phospho-Ser-19 myosin light chain 2 (MLC2; 1:1000; Cell Signaling Technology), anti-c-Src (1:500; Santa Cruz Biotechnology), anti-phospho-Y418 Src (1:1000; Biozol, Eching, Germany), affinitypurified anti-FAK (1:250; Santa Cruz Biotechnology), and anti-phospho-Y397 FAK antisera (1:1000; Biosource, Camarillo, CA, USA). Secondary antibodies used were horseradish peroxidase-conjugated goat anti-rabbit IgG, goat anti-mouse IgG, and donkey anti-goat IgG (1:20,000; all from Jackson ImmunoResearch Laboratories). RhoA- and Rac1-pulldown assays Recombinant GST-TRBD fusion protein encoding the Rhobinding domain of Rhotekin (a kind gift of M. Schwartz, Scripps Research Institute, La Jolla, CA, USA) was used as an activation-specific probe for RhoA–guanosine-5=-triphosphate (GTP) (34). For the detection of GTP-bound RhoA, either unstimulated, or stimulated [10 ␮M lysophosphatidic acid (LPA) for 10 or 30 min], fibroblasts were lysed in 25 mM HEPES/HCl (pH 7.5), 150 mM NaCl, 10 mM MgCl2, 1% (v/v) Nonidet P-40, and 100 ␮M phenylmethylsulfonyl fluoride (pulldown lysis buffer), supplemented with a complete inhibitor pill (Roche). After determining the protein concentration using the bicinchoninic acid assay (Thermo Scientific, Waltham, MA, USA), the supernatant was added to GSTTRBD beads and incubated for 60 min at 4°C. The beads were then washed with ice-cold pull-down lysis buffer and resuspended in sample buffer. Proteins were separated by SDS12.5% PAGE and transferred to nitrocellulose membranes. Equal loading of GST fusion proteins and protein transfer were controlled by Ponceau S staining. Precipitated RhoA proteins were detected using mouse mAbs to RhoA (clone 26C4, 1:500; Santa Cruz Biotechnology) and peroxidaseconjugated anti-mouse IgG (1:20,000) as primary and secondary antibodies, and the enhanced chemiluminescence (ECL) Western blotting detection system (Thermo Scientific). For measuring active Rac1, a recombinant GST-fusion protein encoding the p21-binding domain of PAK1 (GST-PBD, provided by M. Schwartz) and mouse mAbs to Rac1 (1:1,000; BD Transduction Laboratories) were used. Migration, kymography, and FA assembly/disassembly assays Wound healing assays were performed as described previously (35), with minor modifications. In brief, confluent monolayers of WT and P0 dermal fibroblasts were wounded with a yellow Gilson pipette tip, and cells were washed with growth medium. Cell migration was monitored for 12 h after scratching, and images were taken with an EC Plan-Neofluar ⫻10, 0.3-NA objective (Carl Zeiss MicroImaging, New York, NY, USA) in 7-min intervals over the whole migration period. Single-cell motility assays were performed using a motorized Axio Observer Z1 coupled to AxioCam MRm (Carl Zeiss MicroImaging). Cells were plated onto 6-well plates (Costar, Cambridge, MA, USA) at a density of 2.8 ⫻ 105 cells/cm2. Migration of WT and P0 cells was monitored in parallel in a PM S1 incubator (Carl Zeiss MicroImaging) at 37°C and 5% CO2. Recordings of migration started 16 h after plating, and 717

frames were taken with an EC Plan-Neofluar ⫻10, 0.3-NA objective in 10-min intervals over a period of 20 h. Images were processed with Zeiss AxioVision 4.6.3 and analyzed for manual tracking of migrating cells using ImageJ software [W. S. Rasband, U.S. National Institutes of Health (NIH), Bethesda, MD, USA]. To track the whole cell trajectory, cell nuclei were marked for each frame throughout the entire video sequence. Processive indexes were calculated by determining the shortest linear distance between the start and end points compared with the total distance traversed by the cell (36). For statistical evaluation, 50 –70 cells/genotype and experiment were monitored. For kymographic analysis, timelapse movies of migrating single cells were acquired, with frames taken every 15 s for typically 30 min. Kymographs were generated along 1-pixel-wide boxed regions using the Multiple Kymograph plugin (J. Rietdorf and A. Seitz, NIH) for ImageJ software. FA assembly and disassembly were measured as described previously (37). Cells were transfected with EGFP-paxillin and plated onto glass coverslips at a density of 2.8 ⫻ 105 cells/cm2 in phenol red-free DMEM followed by a 16-h spreading period. Time series were acquired on a Zeiss Axiovert S100TV microscope with a Plan-Apochromat ⫻63, 1.4-NA objective (Carl Zeiss MicroImaging) in 3-min intervals over a period of 6 h. Fluorescence intensities of individual adhesions from background-subtracted images were measured over time using MetaMorph 6.3 software (Universal Imaging Corp., Downingtown, PA, USA). The apparent assembly and disassembly constants were determined from semilogarithmic plots of fluorescence intensities as a function of time. Integrin-fibronectin cross-linking assay Expression plasmids encoding fragments (repeats 7–10) of WT (FN-WT) and mutant (FN-R1374/9A) FN type III (a kind gift of H. P. Erickson, Duke University Medical Center, Durham, NC, USA) have been described previously (38). Fragments were expressed in E. coli BL21 (DE3) at 37°C and purified with a Mono-Q column (Amersham Pharmacia, Piscataway, NJ, USA) using standard procedures. For highdensity coating (1 h) of tissue culture-treated polystyrene dishes (39), 30 ␮g/ml of FN fragments, or 10 ␮g/ml of full-length FN (F2006; Sigma-Aldrich) were used. Dishes were then blocked with 1% (w/v) heat-denatured BSA in PBS. For the cross-linking of proteins to the coated dishes, previously described procedures were slightly modified (2, 40). Briefly, fibroblasts to be seeded onto plates were serum starved overnight, trypsinized, and resuspended in Hank’s buffered salt solution (HBSS) supplemented with calcium and magnesium (Life Technologies). Cells were then washed 3 times with HBSS to remove trypsin. Cells (3⫻105) were seeded onto 3.5-cm plates precoated with FN. After 30 min of spreading, cells were incubated with 2 mM 3,3=-dithio-bis-succinimidylproprionate (DTSSP; Thermo Scientific), or just HBSS (control) at 37°C for an additional 30 min, quenched with 50 mM Tris–HCl (pH 7.5) for 10 min and then extracted with 0.1% (w/v) SDS in PBS. For immunofluorescence microscopy, plates were washed with PBS, fixed with methanol/acetone, and FN-cross-linked integrins were analyzed using antibodies to integrin ␤1. For quantitative measurements of the areas covered by cross-linked integrin ␤1, images of integrin ␤1positive structures were obtained from three independent experiments using a ⫻63 objective under constant acquisition settings, avoiding pixel saturation. To measure cellular surface areas vs. integrin ␤1-positive areas, two best-fit lower thresholds were determined from 8-bit images, using the threshold tool of ImageJ and confirmed by visual inspection. Integrin ␤1-positive areas were calculated as percentage of total cell areas. 718

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Statistical analysis Statistical analysis was carried out on data from independent experiments. P values were calculated using 2-tailed Student’s t test, or 2-tailed Mann-Whitney U test where appropriate. Data are presented as mean ⫾ sem values; error bars in figures represent sem; n values are specified in figure legends.

RESULTS Plectin: a key organizer and reinforcing element of IF networks By anchoring vimentin IFs to FAs, plectin acts as an organizer of cage-like IF networks surrounding the nucleus (17). To test how plectin affects the spatial (3D) organization of vimentin IFs in the proximity of FAs, WT and P0 fibroblasts were subjected to immunofluorescence microscopy (Fig. 1A) with subsequent 3D reconstruction analysis of combined deconvolved images (Fig. 1B, C). Triple staining for vinculin, vimentin, and plectin isoform 1f (P1f) revealed that in WT cells, vimentin IFs generally terminated at centrally, but not peripherally, located FAs, whereas in P0 cells they were often overshooting even the very peripheral FAs (Fig. 1A). Close visual inspection of peripheral FAs (marked by circular dotted lines in Fig. 1A) showed that in ⬃75% of WT cells, IF (vimentin) and FA (vinculin) signals were not overlapping (Fig. 1D, FA⫺), whereas in P0 cells, this fraction was ⬍20%, with ⬎80% of peripheral FAs scoring as vimentin-positive (Fig. 1D, FA⫹); similar results were obtained when primary cells were analyzed (Supplemental Fig. S1B). Three-dimensional rendering of magnified confocal sections in WT cells revealed vimentin IFs that were coated with both vinculin and plectin at their ends, as particularly evident in basal views of the cells (Fig. 1B). In computational 3D reconstructions, both proteins appeared to form pocket-like structures into which filament tips were inserted. This is particularly evident at higher magnification of anchored filament tips (Fig. 1C). In the absence of plectin, more robust vimentin filaments and, consistent with previously published data (21), enlarged and elongated vinculin-containing FAs were observed (Fig. 1A, B, bottom panels). Whereas from the basal view, vimentin IFs were clearly seen overshooting FA, from the apical view, they were seen partly perforating these structures (Fig. 1B, bottom panels). Thus, in P0 cells, vimentin IFs did not terminate at vinculin-containing FAs and seemed not to be affected by them. As shown in Supplemental Fig. S1A, the recruitment of vimentin IFs to plectinpositive FAs could be confirmed using total internal reflection fluorescence microscopy (TIRFM). More robust looking vimentin IFs in P0 cells prompted us to assess their diameter in comparison to that of filaments in WT cells. For this, whole-mount cytoskeletons were prepared by extraction of cells with high salt and detergent, a treatment known to leave only IFs intact, while depleting cells of actin and microtubule

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Figure 1. Vimentin IF network organization and individual filament structure in WT and P0 fibroblasts. Cells were grown for 2 h (A–C) or 12 h (D–F) to subconfluence on plastic uncoated dishes. A–C) Immunolabeling using antibodies for vinculin (pseudoblue), vimentin (green), and plectin isoform 1f (pseudored). A) Note gaps between peripheral vinculin and vimentin filaments in WT, contrary to P0 cells. Dotted circular lines indicate distal ends of peripheral FAs. B, C) Three-dimensional reconstruction analysis of deconvolved magnified images are shown as apical, basal, and orthogonal views. For better visualization of FA-anchored (WT) vs. nonanchored IFs (P0), reconstructed 3D images are shown at angles slightly differing from the corresponding original images. B) Boxes indicate areas where the tips of IFs were surrounded by plectin and vinculin. C) Higher magnification of IF anchorage structure shown in uppermost boxed area in B. Note that on omission of the plectin-specific channel, IF tips are clearly seen inserting into partly transparent vinculin-positive structures with the appearance of pockets (lower panels). Arrowheads, tips of IFs. Arrows, IFs overshooting FAs. Scale bars ⫽ 10 ␮m (A), 5 ␮m (B), 1 ␮m (C). D) Column diagram showing proportions (%) of fibroblasts displaying typical FA⫺ and FA⫹ phenotypes. Cells in which less or more than 25% of all detectable peripheral FAs showed overlapping FA and vimentin signals were scored as FA⫺ and FA⫹ phenotypes, respectively; 300 cells were evaluated from randomly chosen optical fields. Data represent means ⫾ sem; n ⫽ 3. † P ⬍ 0.001. E, F) Statistical evaluation of vimentin filament diameters deduced from actin- and microtubule-depleted cytoskeletons (E), or negatively stained whole-mount preparations after Immunogold labeling (F). Scale bar ⫽ 100 nm. Data represent means ⫾ sem; ⬃100 filaments were evaluated from 5 randomly chosen cells. †P ⬍ 0.001.

structures (30). A statistical analysis of negatively stained specimens revealed an average increase of ⬃5% in IF diameter in P0 compared to WT cells (Fig. 1E, and Supplemental Fig. S1C). When unextracted cytoskeletons (not depleted of actin and tubulin) were inspected, and IFs were identified by anti-vimentin Immunogold labeling, the increase in IF diameter observed in P0 vs. WT specimens (⬃20%) was even more pronounced (Fig. 1F). The cause of this phenomenon remains to be investigated. To test whether the observed alterations of vimentin IF structure and network organization affected the kinetics of IF disassembly, we exposed cells to the serine/threonine phosphatase inhibitor OA, which is known to cause disruption of IFs (41– 43). Monitoring differential extractability (solubility) of vimentin in WT and P0 cells, we found that even without treatment, the solubility of IFs was significantly increased in P0 compared to WT cells (Fig. 2A). On treatment with OA, a sustained increase in IF solubility was observed in P0 (up MECHANOSENSING THROUGH VIMENTIN

to 3 h) but not in WT cells, where IF solubility increased only during the first 1 h, declining thereafter (Fig. 2A). Consistent with these results, in P0 cells, mitogen activated protein (MAP) kinase p38 (which is known to be activated by OA and to phosphorylate vimentin under these conditions; refs. 42, 44) showed a dramatic OA-induced increase in activity despite its comparatively low levels in untreated cells (Fig. 2B). Moreover, assessing stress resilience of IF networks using a heat-shock assay, we observed prominent fragmentation of IFs in P0 cells, contrary to WT cells (Fig. 2C). Both increased disassembly dynamics and lower heat-shock resistance of IFs in P0 cells pointed toward an increased fragility of the IF networks in the absence of plectin. Compromised vimentin network organization hampers cell migration of P0 cells Both vimentin and plectin deficiencies were found to compromise fibroblast cell migration (14, 21). The 719

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Figure 2. Increased sensitivity to OA- and heat-shock-induced disruption of vimentin filaments in P0 fibroblasts. A) Dermal fibroblasts were treated with OA for the times indicated. Total cell lysates (L) and detergent-soluble (S) or insoluble (P) cell fractions were subjected to immunoblotting using antibodies to vimentin. Signal intensities of vimentin bands were densitometrically determined, and fraction proportions were calculated as percentage. n ⫽ 5. *P ⬍ 0.05, **P ⬍ 0.01. B) Lysates obtained as in A were analyzed by immunoblotting using antibodies to the unphosporylated/phosphorylated (total), or just the phosphorylated (P-) form of p38. Tubulin, loading control. P-p38 signal intensities, densitometrically determined, were normalized to p38. Values are means ⫾ se; n ⫽ 3. a.u., arbitrary units. *P ⬍ 0.05. C) Fibroblasts were subjected to heat shock at 45°C (20 min) followed by incubation at 37°C for 2 h. Fixed cells were immunolabeled for vimentin. Immunofluorescence contrast-inverted grayscale images of representative cells are shown. Two boxed areas each in a and b are shown as ⬃⫻3.5 images in a=, a⬙, and b=, b⬙, respectively. Arrows in (b=) and (b⬙), fragments of vimentin filaments; arrowheads in (a=) and (a⬙), laterally aligned vimentin filaments; asterisk in (b=), vimentin clump. Scale bars ⫽ 20 ␮m (a, b), 10 ␮m (a=, b=, a⬙, b⬙).

effects of vimentin on cell migration have been demonstrated in two ways. First, the retraction of IFs from the cell periphery was shown to be a necessary prerequisite for lamellipodia formation (45) and second, the forced expression of vimentin led to alterations in FA dynamics (13). Thus, to assess whether plectin-deficiencyinflicted deficits in migration and altered vimentin IF network architecture of P0 cells were mechanistically linked, we analyzed lamellipodia formation in these 720

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cells using immunofluorescence microscopy. Consistent with previous data (17), P0 cells showed less polarization, having fewer protrusions and often just one broad lamellipodium (Fig. 3A). Notably, these lamella regions contained vimentin filaments (Fig. 3A). Using Immunogold electron microscopy, we confirmed the abundance of vimentin IFs in peripheral regions of P0 cells vs. their scarcity in corresponding areas of WT cells (Fig. 3B). Furthermore, we found the bulk of

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Figure 3. Alterations in migration, IF organization, and FA dynamics in plectin-deficient fibroblasts. A) Immunolabeling of vimentin and actin in fibroblasts subjected to a scratch-wound closure assay (see also Fig. 4). Scale bar ⫽ 20 ␮m. B) Whole-mount electron microscopy of negatively stained Immunogold-labeled vimentin IFs in lamella regions of WT and P0 cells. Bottom panels show ⬃⫻17 images of boxed areas in top panels. Scale bar ⫽ 5 ␮m (top panels), 100 nm (bottom panels). C) Individual frames and kymographs from time-lapse movies of WT and P0 cells. Kymographs show lamellipodial activity along the white lines in respective frames. Data are representative of ⱖ10 cells/genotype. Frame scale bar ⫽ 20 ␮m (top panels), kymograph scale bar ⫽ 5 ␮m, time bar ⫽ 3 min (bottom panels). D) Primary fibroblasts isolated in parallel from WT and P0 skin explants were monitored on uncoated dishes for migration over a period of 22 h. Values are means ⫾ sem of migration velocities and processive indices (PI) are shown; PI, defined as linear distance between the start and end points divided by the total distance traversed by a cell, equals 1 for linear migration. Values are based on the analysis of 200 –300 cells/genotype. *P ⬍ 0.05, †P ⬍ 0.001. E) WT and P0 fibroblasts were transfected with EGFP-paxillin and monitored by time-lapse video microscopy for 12 h. Arrows, examples of FAs monitored for fluorescence intensity changes over time. Scale bar ⫽ 10 ␮m. Diagram shows fluorescence intensities, measured as gray values in arbitrary units (a.u.). As tagged paxillin was targeted to peripheral, as well as centrally located, FAs, fluorescence intensities of individual FAs were measured over time without discriminating between their locations. F) Apparent rate constants for FA assembly and disassembly and FA lifetime were determined from semilogarithmic plots of fluorescence intensities as a function of time. Values are means ⫾ sem from 30 individual adhesions monitored per genotype. †P ⬍ 0.001.

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vimentin-positive structures in lamellae not to be anchored at FAs (unpublished observations; see also ref. 17). To investigate whether these phenomena were accompanied by changes in protrusional activity, we performed a kymographic analysis of individual cells. As shown in Fig. 3C, lamellipodial dynamics of WT fibroblasts were characterized by the formation of short

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protrusions and their quick withdrawal, whereas the membrane of P0 fibroblasts extended more slowly over time and rarely withdrew. Since impaired polarization could be expected to affect directionality of cell migration (46), we monitored the migration of single WT and P0 cells by time-lapse video microscopy. In fact, we found that not

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only the migration rates of P0 cells were significantly reduced compared to WT cells (⬃0.75⫻), but also their traversed trajectories were less linear, with processive migration indices of 0.19 for P0 cells and 0.27 for WT cells (Fig. 3D). We next assessed the kinetics of FA dynamics using time-lapse video microscopy of cells transfected with EGFP-paxillin (Fig. 3E). As evident from the intensity graphs shown for two representative FAs monitored (arrows), the paxillin signal in plectin-deficient cells was significantly prolonged over that in WT cells. For quantification, we determined the apparent FA assembly and disassembly rate constants from semilogarithmic plots of fluorescence intensities as a function of time (ref. 37 and Fig. 3E). P0 cells showed significantly reduced assembly (⬃3⫻) and disassembly (⬃5⫻) rates of FAs compared to WT cells, leading to a doubling of FA lifetimes (Fig. 3F). The relatively high reduction of disassembly rate constants calculated for P0 FAs indicated that the larger and more stable FAs observed in mutant cells were primarily due to compromised disassembly dynamics. Taken together, these data implied that the massive accumulation of vimentin filaments in lamella and lack of anchorage to FAs led to impaired polarization and deficits in the FA disassembly process, resulting in impaired motility of P0 cells. Plectin deficiency leads to attenuated FAK signaling To explore signaling pathways that were responsible for the migration and FA turnover deficits of P0 cells, we first analyzed growth factor-activated migration using an ex vivo scratch-wound closure assay. Monitoring migration in the absence of serum [fetal calf serum(FCS)-free], no differences in migration velocities of WT and P0 cells were noticeable (Fig. 4A, B). However, in the presence of FCS or platelet-derived growth factor (PDGF), a potent promigratory growth factor, a ⬃50% increase in migration velocity was observed for WT cells, but only a ⬃25% increase for P0 cells, suggesting that the activation of growth factor-triggered pathways was attenuated in P0 fibroblasts (Fig. 4A, B). FA turnover typically depends on signaling pathways involving FAK and Src kinases, which can be activated

by growth factors [epidermal growth factor (EGF), PDGF] and/or on integrin-ECM engagement (47). Thus, we assessed FAK activity in WT and P0 cells grown in the presence of serum on uncoated (control) or FN-coated plastic dishes (requiring the engagement of integrins), or on polylysine (PL)-coated plastic dishes (no engagement of integrins). Using antibodies to phospho-Y397 FAK, which detect the activated form of the kinase, FAK activity was found to be reduced by ⬃30% in lysates from P0 compared to WT cells, as long as the cells were grown on uncoated or FN-coated dishes, but not on PL (Fig. 4C). A similar reduction of FAK activity in P0 cells was detected using primary cell cultures grown on plastic dishes (Supplemental Fig. S2A). Using corresponding phospho-specific antibodies, we next measured the activities of Src and ERK1/2, two positively regulated downstream targets of FAK (48) involved in migration. Analyzing cell lines, as well as primary cell cultures, we found both kinases to be significantly down-regulated in P0 cells, consistent with reduced FAK activity (Fig. 4D, E; Supplemental Fig. S2B, C). Another downstream target of FAK, RhoA, plays a key role in the stabilization of actin stress fibers and FAs (49). In contrast to Src and ERK1/2, the activity of RhoA is, however, suppressed by FAK (6). By measuring the amount of active (GTP-bound) RhoA present in quiescent (control) cells and in cells exposed to LPA, we detected significantly increased amounts of active GTP-RhoA in lysates of P0 cells vs. WT cells (Fig. 4F). This was not true for the activity of another GTPase, Rac1, which was unaltered in both cell types (Supplemental Fig. S2D). Surprisingly, on LPA stimulation, larger portions of GTP-RhoA were pulled down from WT compared to P0 cells (Fig. 4F), suggesting that a constitutively higher RhoA activation level in P0 cells leads to reduced responsiveness to LPA treatment. These data are consistent with a previously reported hampered activation of RhoA biosensors in P0 fibroblasts (22). Assessing whether increased RhoA activity in P0 cells correlated with increased phosphorylation of MLC, we measured its phosphorylation on Ser-19,

Figure 4. Analysis of signaling pathways responsible for migration and FA turnover deficits of P0 cells. A) WT and P0 fibroblasts were grown in parallel until reaching confluence (⬃48 h). Subsequently, a scratch wound was inflicted onto monolayers, and cells were exposed (12 h) to either FCS-supplemented or FCS-free medium, as indicated. Lines mark either the original edge of the scratch made (first frames of time-lapse recordings) or the front of migrating cells at the 12 h time point. Scale bar ⫽ 100 ␮m. B) Average migration velocity of the scratch front under the conditions indicated (means⫾sem). n ⫽ 3. *P ⬍ 0.05, **P ⬍ 0.01. C) Fibroblasts were grown on either uncoated plastic (control), PL-, or FN-coated surfaces. For immunoblotting of cell lysates antibodies to phosphorylated (P)-FAK and tubulin (loading control) were used. P-FAK signal intensities, densitometrically determined, were normalized to tubulin (means⫾sem); a.u., arbitrary units. n ⫽ 3. *P ⬍ 0.05, **P ⬍ 0.01. D, E) Proteins in membrane subfractions of fibroblasts (normalized for equal protein contents), were immunoblotted using antibodies to unphosphorylated/phosphorylated (total) Src and ERK2 or phosphorylated versions of Src and ERK1/2; tubulin, loading control. P-Src and P-ERK1/2 signal intensities, densitometrically determined were normalized to total Src and ERK2, respectively (means⫾sem); a.u., arbitrary units. n ⫽ 3. **P ⬍ 0.01. F) Fibroblasts were either untreated (control), or stimulated with LPA for 10 min (10=LPA) or 30 min (30=LPA). Cells were lysed and probed for GTP-bound RhoA by pulldown with GST-TRBD beads. Lysates (total RhoA) and pulldown fractions were immunoblotted using antibodies to RhoA. n ⫽ 3. G) Immunoblotting of cell lysates using antibodies to P-MLC (Ser-19; tubulin, loading control. P-MLC signal intensities (densitometrically determined) were normalized to tubulin (means⫾sem); a.u., arbitrary units. n ⫽ 5. *P ⬍ 0.05. MECHANOSENSING THROUGH VIMENTIN

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which is one of the major phosphoepitopes correlating with increased cell contractility. MLC Ser-19 phosphorylation clearly increased in P0 compared to WT cells (Fig. 4G), implying that attenuated FAK/Src signaling in P0 cells causes a compensatory overactivation of RhoA and MLC, which eventually leads to enlarged, less dynamic FAs slowing down cell migration.

icked by seeding cells without serum for a short time (1 h) on plates coated with a recombinant fragment of FN (FN-R1374/9A) that harbors mutations inactivating the integrin-binding of two major tension-sensing synergy sites, but leaves the tension-independent RGD-binding site intact (for details, see refs. 2, 39, 52). A corresponding FN fragment without these mutations (FN-WT) was used as a control for tension-dependent conditions. As expected, plating on FN-R1374/9A resulted in all cases in an overall decrease in FAK activity, as the so-called tensional activation of the kinase (2) was largely blocked (Fig. 5A). Notably, under these conditions FAK activities were similar in all three cell types, indicating that differences observed on FN-WT were rooted in higher tensional activation of FAK in WT compared to P0 and V0 cells (Fig. 5A). To investigate whether the lack of tensional FAK activation was the cause for the more robust FAs observed in P0 and V0 fibroblasts, cells plated on

Diminished cytoskeletal tension accounts for compromised FAK activation and enlarged FAs in P0 and V0 cells FAK, as one of the primary mechanosensory signaling proteins, is able to sense cytoskeletal tension and adjust its own activity accordingly (2, 50). To test whether the observed diminished activation of FAK was rooted in the reduced cytoskeletal tension (prestress) of P0 and V0 cells (22, 51), we compared the activity of FAK under tension-dependent vs. tension-independent conditions. The tension-independent situation was mim-

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Figure 5. Seeding of P0 cells on mutant fibronectin (FN-R1374/9A) abolishes their FA and FAK phenotypes. WT, P0, and V0 fibroblasts were seeded either onto FN-WT or FN-R1374/9A for 1 h in the absence of serum. A) Cell lysates were subjected to immunoblotting using antibodies to P-FAK or unphosphorylated/phosphorylated (total) FAK; tubulin, loading control. Bar graph, densitometrically determined P-FAK:total FAK signal intensities normalized to values obtained for WT cells seeded onto FN-WT (100%). Values are means ⫾ sem n ⫽ 3. *P ⬍ 0.05. B) Cells were immunolabeled for vinculin. Boxes indicate areas of representative FA clusters, shown as enlarged insets (top right corners) after processing for morphometric analysis. Scale bar ⫽ 20 ␮m. C, D) Statistical evaluation of morphometric FA analyses (area and length) of cells plated on FN-WT or FN-R1374/9A; ⬎500 FAs were measured in ⬎10 cells for each cell type and each condition. D) Box and whisker plots indicate the median (middle line in the box), 25th percentile (bottom line of the box), 75th percentile (top line of the box), and 2.5th and 97.5th percentiles (whiskers). *P ⬍ 0.05, **P ⬍ 0.01, †P ⬍ 0.001. 724

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FN-WT and FN-R1374/9A were immunostained for vinculin (Fig. 5B), and the size and length of their FAs were measured morphometrically. As expected, when plated on FN-WT (tension dependent), the subpopulation of cells with enlarged FAs (⬎2 ␮m2 and ⬎2 ␮m in area and length, respectively) was increased in both P0 and V0 compared to WT cells (Fig. 5C). However, on FN-R1374/9A (tension independent), the average area and length of FAs were found to be quite similar among the three cell types, albeit, in general, the values decreased (Fig. 5D). The only exception was the average length of FAs in V0 cells, which remained similarly elevated compared to WT cells. Similar observations were made using antibodies to paxillin (instead of vinculin) to stain for FAs (Supplemental Fig. S3A). Notably, tension-independent conditions appeared not to require plectin and vimentin in FAs, as both proteins retracted and accumulated in perinuclear areas under these conditions (Supplemental Fig. S3C). In summary, tension-independent conditions abolished the differences in FAK activities and FA dimensions between WT, P0, and V0 cells, highlighting the reduced cytoskeletal tension in P0 and V0 cells as the major cause for the observed phenotypic alterations. Compensatory overactivation of integrins in the absence of plectin and vimentin To investigate whether the reduction in integrin-mediated FAK activation in P0 and V0 cells was rooted in the compromised ability of integrins to become activated or to transduce signals, we assessed the activation status of integrins in WT, P0, and V0 cells. For this, we used an assay where cells adhering to FN-WT or FN-R1374/9A are exposed to the chemical cross-linker DTSSP, which cross-links integrins to fibronectin, but solely if they are in their activated state (2). In a slight variation of the original protocol (for details, see Materials and Methods), cells were spread onto dishes that had been densely coated with FN-WT or FN-R1374/9A and, after exposure to the cross-linker and subsequent lysis of the cells, immobilized (fibronectin cross-linked) integrin molecules left behind were visualized by immunofluorescence microscopy using antibodies to integrin ␤1. A number of control experiments confirmed the reliability of this assay. WT and FN-1374/9A were equally efficient in forming high-density coatings, as judged by similarly intense fluorescence signals when using antibodies to fibronectin (Supplemental Fig. S4A). Specific cross-linking of integrins was only observed in the presence of DTSSP cross-linker (unpublished results), and it required high-density coating of culture dishes (in the range of 30 ␮g/ml) similar to conditions described by others (39); at low concentration (5 ␮g/ml), or in control experiments without fibronectin coating, hardly any or no integrin signals were detected (Supplemental Fig. S4B and unpublished results). When the specimens were costained for vimentin, no specific staining was observed, indicating that the extraction of cells with detergent was complete, leaving only exterMECHANOSENSING THROUGH VIMENTIN

nally cross-linked proteins (integrins) behind (unpublished results). Consistent with previous reports showing a crucial role of R1374/9 synergy sites for integrin cross-linking (implying activation; ref. 2), in P0 cells spread onto FN-WT, cross-linked integrin showed massive accumulation at dash-like structures resembling peripheral FAs, whereas on FN-R1374/9A-coated dishes, hardly any such structures were seen, but instead, cross-linked integrins were found as dot-like aggregates diffusely spread over the whole area of the cell (Supplemental Fig. S4A). A statistical evaluation of the cross-linked integrin types (dash- vs. dot-like) revealed that ⬃70% of WT cells spread onto FN-WT exhibited dot-like integrin ␤1-staining patterns, while the remainder showed faint, dash-like structures (Fig. 6A, B). For P0 and V0 cells, it was the other way around, with the majority (⬃80%) of cells spread onto FN-WT showing robust dash-like cross-linked integrins. A significant increase in dashlike cross-linked integrins in P0 vs. WT cells was also observed when, instead of the FN-WT fragment, fulllength fibronectin was used for coating of dishes, indicating that the truncated version of fibronectin acted in a manner similar to that of the full-length protein (Supplemental Fig. S4C). When plated onto FN-R1374/9A-coated dishes, the proportion of dashlike cross-linked integrins in general was reduced and mainly dot-like structures were observed. Notably, however, the marked differences between WT and mutant cells regarding the proportion of cross-linked integrin types (dot-like vs. dash-like) were reduced in the case of V0 cells, or completely abolished in P0 cells (Fig. 6A, B). Similar results were obtained when primary cell cultures (WT and P0) were used instead of immortalized cell lines (Supplemental Fig. S4D). To quantify the amounts of cross-linked (active) integrins in all three cell types, the proportion of integrin ␤1-positive areas vs. mean cellular surface areas was measured using ImageJ software. This method revealed higher values for P0 and V0 compared to WT fibroblasts when spread on FN-WT, whereas no significant differences were observed on FN-R1374/9A (Fig. 6C). This was consistent with the morphological differences observed for cross-linked integrin structures (Fig. 6A, B). Together, these data showed that on FN-WT, but not FN-R1374/9A, more integrins were in their active state in P0 and V0 compared to WT cells. Thus, hyperactivation of integrins in P0 and V0 cells shows that it is not the inability of integrins to become activated, but rather their uncoupled or ineffective signal transmission that leads to the deregulation of major downstream signaling targets and, thereby, to the creation of a compensatory positive feedback loop on themselves.

DISCUSSION Vimentin IFs have been shown to affect the spatial organization, size, function and adhesion strength of 725

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Figure 6. Assessing the activation status of integrins by chemical cross-linking to FN. A) Cells treated as described in Supplemental Fig. S4 were subjected to immunofluorescence microscopy using antibodies to integrin (ITG) ␤1. White and green asterisks indicate dotlike and dash-like integrin clusters, respectively. Scale bar ⫽ 20 ␮m. B) Statistical quantification of cells categorized as shown in A. Values are means ⫾ sem from ⬎50 cells of each genotype from 3 experiments. *P ⬍ 0.05, **P ⬍ 0.01. C) Statistical quantification of integrin ␤1(ITG␤1)positive cross-linked areas using ImageJ software. Values are means ⫾ sem from ⬎50 cells/genotype from 3 experiments shown in A. **P ⬍ 0.01.

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FAs (for review, see ref. 53). However, the underlying mechanisms of these actions have remained elusive. Because of their remarkable ability to greatly stretch beyond their original length, vimentin IFs have been suggested to sustain tensile forces of actin stress fibers and act as stretch sensors, thereby regulating forcedependent recruitment of different molecules to their surfaces (53). These properties have prompted several authors to suggest a role of vimentin filaments in mechanotransduction (14, 23, 54 –56), and in an elegant study, it has been shown that vimentin IF can mediate a direct mechanical force transfer from cellsurface integrins to the cell interior (57). Analyzing P0 and V0 fibroblasts, we show here for the first time that the anchorage of IFs at FAs is a strict prerequisite for efficient integrin-mediated mechanotransduction and subsequent activation of the major mechanosensory molecule FAK. Abrogation of this anchorage leads to the slowdown of FA dynamics and impairment of migration, similar to the situation of IF deficiency. On the basis of experiments in which we abolished the ability of cells to sense tension exerted from the exterior of the cell, we suggest that the absence of vimentin at FAs reduces the tension in actin filaments in fibroblasts and leads to inefficient mechanotransduction in the cell. FAs have long been known to be the main anchor points for the actin cytoskeleton, but it has only re726

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cently begun to be appreciated that they may act in a similar way as docking sites of vimentin IFs. According to previous reports, the multifunctional cytolinker protein plectin is the crucial molecule that mediates the attachment of vimentin IFs to FAs (16, 17, 58). Our study provides evidence that the lack of vimentin IFs at FAs not only has consequences for the structural organization of the vimentin network (which loses its cagelike appearance), but also affects individual filaments as they display increased average diameters. Larger diameters would support the idea that the filaments in P0 cells are under reduced tension and, thus, less stretched, consistent with in vitro data (59). In addition, our study shows that vimentin IFs not being anchored at FAs not only show altered morphology and networking capacity, but also are less resistant toward collapse in response to various types of stress. Our study unveils an essential role for vimentin and plectin in FAK activation on integrin engagement. Residing at the crossroads between integrin signaling and various other signaling events, FAK activates several other kinases, including Src and MAP kinases, and it exerts a negative feedback loop onto RhoA activity by up-regulating its negative effector RhoA GAP (50, 60, 61). Our analysis of plectin-deficient fibroblasts uncovered changes in the entire signaling cascade downstream of FAK, which lead to inefficient activation of Src, ERK1/2, and p38, and notably, to a

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dampening of the negative-feedback loop acting on RhoA. The resulting enhancement of RhoA activity coupled with the activation of (FA-promoting) MLC provides a plausible explanation for the increase in stress fibers and FAs known for some time to be caused by plectin deficiency (21). Actomyosin-generated tension has been shown to be a major regulator of FAK and many other adhesionassociated signaling molecules (5). To understand how vimentin filaments and their targeting to FAs influence adhesion-dependent signaling, we have adopted the actomyosin contraction model postulated by Deguchi et al. (62). According to this model, strain will be generated within the actomyosin gel only if it is held in place by physical constraints (Fig. 7). These constraints are provided by attachments to the ECM on the one hand and to the cytoskeleton or other intracellular structures on the other. Extracellular attachment is mediated via integrins, which can occupy two binding domains on FN, one containing the RGD sequence, the other containing synergy sites (SYN). Occupation of the RGD site results in basal FAK and Src activation that does not depend on intrinsic cellular tension (prestress). Integ-

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rin’s SYN site occupation occurs on cytoskeletal tension and leads to a significant amplification of the FAK and Src activation signals. Reduced cytoskeletal tension (prestress) has been reported for both P0 and V0 cells, correlating well with our finding of reduced FAK activation in these cells. Our study revealed that under tension-independent conditions there were no longer differences observed between P0, V0, and WT cells regarding the activation of FAK and the size of FAs. This pointed toward diminished tension in actomyosin fibers of P0/V0 cells being the cause for altered signaling. We suggest that vimentin IFs act as a provider of physical constraints for the actomyosin system along with other cytoskeletal components. However, to perform this function, vimentin IFs need to be anchored at FAs and, additionally, form a robust rather rigid plectin-interlinked network structure (cage) in the central part of the cell (schematically depicted by the upright bars on the righ-hand side of the panels in Fig. 7). The absence of these constraints, due to the loss of either the IF network altogether (as in the case of V0), or of its anchoring and networking proteins (P0), results in reduced cellular prestress and diminished ability of cells to sense tension. Consequently, FAK, the major sensor of cytoskeletal tension, is being less activated, and thus less effective, in exerting a negative feedback loop on RhoA and MLC. In turn, this leads to RhoA overactivation and increased actin stress fiber formation. In fact, the increased proportion of cross-linked integrins (increased SYN site occupation) monitored in P0 cells points toward an attempt of the cells to compensate for diminished tension by elevating actin stress fiber formation and, thereby, increasing the proportion of active (tensed) integrins. In all, this model implies that vimentin IFs bear a major part of the physical constraints required for actomyosin gels to be effective and, thereby, serve for mechanosensing, i.e., fine-tuning of the cell’s ability to detect and respond to forces. Our findings reveal that FA turnover, polarization, and directional cell migration, the major FAK-mediated migratory determinants (6, 9, 61), are perturbed in plectin-deficient fibroblasts, in line with a reduction of their FAK activity. Activated FAK, on the other hand, is considered as a marker for oncogenic transformation and metastasis (63) and an increasing number of studies shows that plectin and vimentin are upregulated in various cell carcinomas, leading (in conjunction with FAK-regulated ERK1/2) to increased invasiveness and metastasis potential of cells, and poor survival rate of patients (64, 65), Hence, by identifying a novel mechanistic link between vimentin IF targeting to FAs and integrin-mediated FAK signaling in regulating cell migration, our study sheds new light on plectin/vimentinmediated tumor progression. The authors thank H. P. Erickson (Duke University Medical Center, Durham, NC, USA), M. Gimona (University of Salzburg, Salzburg, Austria), M. Schwartz (Scripps Research 727

Institute, La Jolla, CA, USA), and P. Traub (University Bonn, Bonn, Germany) for generously providing materials, and A. J. Ridley, J. V. Small, and R. G. Valencia for sharing expertise. This work was supported by Austrian Science Research Fund grants P20744-B11 and I413-B09 (Multilocation DFG-Research Unit 1228 subproject). M.G. received support from MEYS (7AMB13AT012 and OP RDI CZ.1.05/1.1.00/02.0109 BIOCEV).

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Mechanosensing through focal adhesion-anchored intermediate filaments.

Integrin-based mechanotransduction involves a complex focal adhesion (FA)-associated machinery that is able to detect and respond to forces exerted ei...
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