Vinculin binding to actin filaments is thought to be critical for force transduction within a cell, but direct experimental evidence to support this conclusion has been limited. In the present study, we found mutation (R1049E) of the vinculin tail impairs its ability to bind F-actin, stimulate actin polymerization, and bundle F-actin in vitro. Further, mutant (R1049E) vinculin expressing cells are altered in cell migration, which is accompanied by changes in cell adhesion, cell spreading and cell generation of traction forces, providing direct evidence for the critical role of vinculin in mechanotransduction at adhesion sites. Lastly, we discuss the viability of models detailing the F-actin-binding surface on vinculin in the context of our mutational analysis.
Cell migration is a complex process that is critical for embryonic development, wound healing and maintenance of tissue integrity . The current model of cell migration is a multi-step process involving polarization in the direction of a stimulus, extension or protrusion of a membrane, contraction of the cell body and retraction of the cell rear. In previous years, our understanding of cell migration has been significantly advanced by the identification of proteins that govern lamellipodial protrusion [2–8]. This information combined with studies of adhesion plaques has revealed that, once formed, lamellipodial protrusions attach to the substratum by transient adhesion complexes. Some of these transient structures anchor the protrusion and subsequently mature into adhesive structures known as focal adhesions, which support the generation of traction forces necessary to pull the cell body forward and break older adhesions at the cell rear.
The adhesive complexes of migrating cells are rich in the highly conserved focal adhesion protein vinculin. Vinculin consists of a head domain, a short proline rich linker and a tail domain [9,10]. In its ‘closed’ conformation, the vinculin head interacts with its tail. Binding of proteins to the head domain releases this intramolecular interaction allowing vinculin to adopt an ‘open’ conformation. In this open conformation, the vinculin tail (VT) binds actin . Vinculin can bind to and modify existing actin bundles and stimulate the formation of actin bundles and networks , making it an ideal candidate for establishing new actin assemblies and linking them to the existing cytoskeleton.
Multiple models detailing where actin binds the vinculin tail have emerged [13–15]. Work by Janssen et al.  suggests that vinculin binds to F-actin via two distinct binding surfaces within the vinculin tail–an upper and a lower actin monomer binding site. In this model, the R1049 residue is a contact point in the lower site, located just beyond the fifth α-helix, within the vinculin tail. Janssen et al.  also suggest that the region near the C-terminal loop is important for vinculin dimerization, a finding supported by the previous work of Shen et al. . Clustered charge-to-alanine experiments  are fully consistent with the binding sites described in Janssen et al. , and support the structural conclusions that the top site is mostly hydrophobic in nature, whereas the one at the bottom contains a significant amount of electrostatic interactions.
Recent work by others has challenged the vinculin tail:actin interface proposed by Janssen. These studies reveal regions important for actin binding and bundling located outside of the proposed Janssen sites (I997, V1001; [15,17]). In support of this, full-length vinculin harbouring either I997A or V1001A mutations compromise focal adhesion size, number and cell spreading .
With these studies in mind, we tested the Janssen hypothesis that R1049 directly participates in actin binding. We report here that mutating vinculin R1049 directly impacts actin binding, which in turn compromises cell migration, adhesion, spreading and traction force generation.
Purification of VT or RE
pET21a (Novagen) was engineered with a 6His tag and tobacco etch virus (TEV) protease cleavage site fused to the N-terminal region of chicken vinculin amino acids 881–1066 to generate pET21a-VT. pET21a-VT was mutagenized using the Stratagene Quickchange protocol to generate pET21a-RE (R1049E). Both VT and RE proteins were prepared as follows: for expression of the VT, Escherichia coli containing this plasmid were grown to early log phase, and induced with 1 mM IPTG for 3 h at 37°C. Bacteria pellets were resuspended in lysis buffer (50 mM phosphate, 300 mM NaCl, 20 mM imidazole, 5% glycerol, aprotinin (10 μg/ml), pepstatin A (2.5 μg/ml), lysozyme (0.2 μg/ml) and DNa seI (0.02 μg/ml). The bacterial pellet was sonicated twice for 20 s, then insoluble debris was removed by centrifugation at 12000 g for 15 min. The supernatant was filtered through 0.45 μm syringe filters and incubated for 1 h at 4°C with Ni2+-nitrilotriacetate (Ni-NTA)-agarose pre-washed in lysis buffer. The beads were washed twice in lysis buffer, then twice with 50 mM Tris/HCl pH 7.6, 150 mM NaCl. Agarose was resuspended in a suitable volume of 250 mM imidazole in TBS, pH 7.6. The agarose was allowed to settle and the supernatant was passed over a PD-10 column equilibrated with TEV buffer (50 mM Tris/HCl, pH 7.5, 150 mM NaCl, 0.5 mM EDTA, 1 mM DTT). The 6His tags were cleaved from the VT proteins by the addition of 6His-TEV protease (0.16 mg/ml). The 6His-VT proteins were incubated with 6His-TEV overnight at 4°C. Following the incubation, 6His-TEV and remaining 6His-VT were bound to Ni-NTA-agarose beads, and the supernatant was passed over a PD-10 column to exchange the buffer to 50 mM Tris/HCl, pH 7.6, 50 mM NaCl. This buffer was diluted with a final concentration of 20% glycerol and the proteins were stored at −80°C until use. Protein concentration was determined by A280, using the molar absorption coefficient 18470 M−1·cm−1.
Circular dichroism and dynamic light scattering
For CD analyses, spectra of VT or RE in 50 mM Tris/HCl, pH 7.6, 50 mM NaCl, 20% glycerol were recorded from 190 to 260 nm at 0.1 nm increments on a JASCO J-815 instrument. For dynamic light scattering (DLS) analyses, VT and RE were purified by FPLC. The purified VT or RE samples were then subjected to DLS using a DynaPro NanoStar instrument (Wyatt Technology). Ten measurements were taken and averaged to produce an ‘average measurement’; five average measurements per experiment were then averaged to yield the ‘overall average’ hydrodynamic radius. Three overall averages were then averaged to yield the mean values reported here.
Chemical cross-linking experiments were carried out as in . Briefly, solutions of VT or RE in CL buffer (10 mM sodium phosphate, 100 mM NaCl and 0.02o10 NaN3, pH 7.5) were cross-linked with 100 μM disuccinimidyl suberate (DSS) for 20 min at room temperature. The reactions were quenched by the addition of 50 mM Tris/HCl, pH 7.6.
Purification of yeast actin, pyrene fluorescence and actin-VT co-sedimentation assays
Actin was purified from the lysates of frozen yeast cells by a combination of DNaseI affinity chromatography, DEAE anion exchange chromatography and polymerization/depolymerization cycles as described in . Pyrene fluorescence and co-sedimentation assays were performed as described in . Co-sedimentation assays were spun at either 20000 or 80000 rev./min in a Beckman TL-100 centrifuge using a TLA 100 rotor.
Reactions for visualization using TEM were generally prepared as in . Briefly, for low ionic strength conditions: reactions containing 1 μM G-actin were combined with various concentrations of VT or RE, allowed to reach steady state, then 3 μl of the reaction mixture was pipetted on to a 400 mesh Formvar TEM grid. The grids were stained with 1% uranyl acetate, and visualized using a JEOL TEM 1230. For physiological salt conditions, the reactions were prepared in the same manner as for low salt conditions, except 2 mM MgCl2 and 50 mM KCl was added following the addition of VT or RE.
Generation of stable cell lines
Vinculin-null mouse embryonic fibroblasts (MEFs) were obtained from vinculin-null mice and were the generous gift of Eileen Adamson (Burnham Institute) and maintained in Dulbecco's modified Eagles medium (DMEM) plus 10% FBS, 1% penicillin/streptomycin and 400 μg/ml G418. The vector pLPCX-GFP (Clontech) was engineered to include a GFP tag with a restriction site at its 3′ end, and full-length wild-type vinculin (vinWT) was cloned into this construct to generate pLPCX-GFP-vinWT. Site-directed mutagenesis was carried out on pLPCX-GFP-vinWT using the Stratagene QuickChange protocol to generate pLPCX-GFP-REvin. The entire coding sequence of vinculin was sequenced to confirm the R1049E mutation. The plasmids were transfected into 293-GPG cells using Lipofectamine reagent (Invitrogen) as previously described [19,20]. Virus was collected and added to vinculin-null MEFs. Infected MEFs were selected in 2 μg/ml puromycin, and mass populations of cells were sorted by FACS to achieve the level of expression desired. Expression levels were assayed by immunoblotting using antibodies against vinculin (hVIN1, Sigma) at a concentration of 1:1000 or against the p34-Arc subunit of the Arp2/3 complex as previously described .
Cell spreading assays
Glass coverslips were coated with 10 μg/ml fibronectin in PBS overnight at room temperature. The coverslips were rinsed twice with PBS, and blocked with 2% BSA in PBS for 2 h at room temperature, then rinsed twice with PBS. Cells were seeded on the coverslips and allowed to adhere for 25 min or 4 h, then fixed in 3.7% formaldehyde. For 25 min assays, Nomarski images were taken. The outlines of at least 30 cells from three independent experiments were traced in Adobe Photoshop CS4 and converted in to cell area. For 4 h assays, coverslips were permeabilized for 20 min in 0.5% Triton X-100 in Universal Buffer (UB: 150 mM NaCl, 50 mM Tris/HCl, pH 7.6, 0.01% NaN3), blocked for 45 min in 1% BSA in UB, then stained with DAPI (500 ng/ml) and Texas-Red phalloidin (1:750; Invitrogen) for 45 min. Coverslips were rinsed twice in PBS, twice with water, then mounted on to slides using Mowiol (Fisher Scientific).
Cell adhesion assays
Glass coverslips were coated with 10 μg/ml fibronectin as before. Cells were seeded on the coverslips at 50000 cells per coverslip and allowed to adhere for 25 min. Wells of the 24-well plate containing coverslips were vigorously washed three times in PBS, then fixed in the same manner as described. Nine non-overlapping fields were taken of each coverslip at ×10 magnification. Each experiment was performed on duplicate coverslips per condition, and the number of cells relative to the GFP (vinculin-null) control for each experiment was recorded.
Cell migration assays
Glass coverslips were coated with 10 μg/ml fibronectin as before. Cells were seeded on the coverslips and allowed to adhere overnight. Cells were imaged on a Zeiss Axiovert 200 M in an enclosed stage heater and a humidified, 5% CO2 environment. Images were taken every 5 min for a total of 10 h. Only cells visible for a minimum of 4 h were included in the analysis. Tracks were analysed using NIH ImageJ Manual Cell Tracker plugin.
Traction force assays
Microfabricated post array deflection (mPAD) device silicon masters were prepared as described previously . In brief, elastomeric micropost arrays were fabricated using PDMS replica molding. To make microfabricated post array templates, 1:10 PDMS prepolymer was cast on top of silanized mPAD device silicon masters, cured at 110°C for 30 min, peeled off gently, oxidized with oxygen plasma (Plasma-Preen; Terra Universal), and silanized overnight with (tridecafluoro-1,1,2,2,-tetrahydrooctyl)-1-trichlorosilane (Sigma–Aldrich) vapour under vacuum. To make the final PDMS mPAD device, 1:10 PDMS prepolymer was cast on the template, degassed under vacuum for 20 min, and cured at 110°C for 20 h and gently peeled off the template on a #1 25 mm diameter circular coverslip (Electron Microscopy Services). Peeling-induced collapse of the mPADs was rectified by sonication in 100% ethanol, followed by supercritical drying in liquid CO2 using a critical point dryer (Samdri-PVT-3D; Tousimis), as described previously . Flat PDMS stamps were generated by casting 1:30 PDMS prepolymer on flat silanized silicon wafers. Stamps were coated in a saturating concentration of fibronectin (Invitrogen) (50 μg/ml in PBS) for 1 h. These stamps were washed in distilled water and dried under a stream of nitrogen gas. FN-coated stamps were placed in contact with surface-oxidized mPAD substrates (UVO-Model 342; Jelight). mPAD substrates were labelled with 5 μg/ml Δ9-DiI (Invitrogen) in distilled water for 30 min. mPAD substrates were subsequently transferred to a solution of 0.2% Pluronics F127 (Sigma–Aldrich) for 30 min, to prevent nonspecific protein absorption. WT, RE and null eGFP-vinculin MEF cells were seeded in growth medium and then allowed to spread overnight. mPAD substrates were transferred to an aluminium coverslip holder (Attoflour Cell Chamber; Invitrogen) for live cell microscopy and placed in a stage top incubator that regulated temperature, humidity and CO2 (Live Cell; Pathology Devices). Confocal images were taken with a Nikon C2-Confocal Module connected to a Nikon Eclipse Ti inverted microscope using a high magnification objective (CFI Plan Apochromat total internal reflection fluorescence (TIRF) 60 × oil, N.A. 1.45; Nikon). Post images were captured using a 561-nm laser with a 595/50 filter, and vinculin images were captured using a 488 nm laser and 525/50 filter. For force measurements, the top and bottom of the posts were sequentially imaged and the deflection measured. The resulting force, F, was calculated using Euler–Bernoulli beam theory, in which E, D, L and δ are the Young's modulus, post diameter, post height and post deflection respectively:
In the present study, we were motivated to test the consequence of mutating R1049, a vinculin residue implicated in actin binding on cell migration, adhesion, spreading and traction force generation.
The R1049E mutant vinculin tail is structurally similar to wild-type vinculin tail
For our in vitro analyses, we used site-specific mutagenesis to generate an R to E substitution at position 1049 in a His-tagged fusion protein containing the wild-type VT residues 881–1066. Except where noted, the His tag was cleaved from the protein. We first examined if mutation of R1049E, herein referred to as RE vinculin, had any effect on vinculin tail structure. For this, both RE and VT purified tail domains were examined by DLS, which estimates the hydrodynamic radius of a protein. The average hydrodynamic radii for VT and RE were 3.0±0.1 nm and 3.0±0.1 nm, which do not differ significantly (Figure 1A). The α-helical content of both VT and RE were also examined by CD. In this method, α-helical content is characterized by valleys in the spectra around 208 and 222 nm. We used a correction factor to set the spectra equal at 208 nm, then corrected every other value by this number and examined the difference in spectra at 222 nm. The spectra track nearly identically (Figure 1B), and the difference in ellipticity was virtually zero (Figure 1B inset). Taken together, these data indicate there are no major structural changes of α-helices in RE compared with VT.
R1049E is structurally similar to wild-type VT
There is speculation that R1049 may contribute to vinculin dimerization. Hence we sought to determine if RE was a dimerization mutant. Work from the Craig Laboratory demonstrated that vinculin tail can dimerize and is cross-linked into higher order oligomers . Hence, we next asked whether RE could be cross-linked to the same extent as VT. Using the same cross-linking approach as Johnson and Craig , we found that the relative amounts of dimer formed when VT or RE are cross-linked was virtually the same (Figure 1C). Thus, RE does not affect dimerization of the vinculin tail. Taken together, these data demonstrate that there are no major structural changes in RE compared with VT.
R1049E is an actin-binding and bundling mutant in physiological ionic strength conditions
We next tested if RE was defective in actin binding under physiological salt concentrations. To assay vinculin tail binding to actin filaments, VT or RE was pre-incubated with 1 μM G-actin, and then F-salt added to initiate actin polymerization. Completed reactions were centrifuged at a speed sufficient to pellet all polymerized actin as well as actin-bound vinculin tail peptide (co-sedimentation). A quantified analysis of pelleted vinculin tail peptide was fit to the quadratic binding equation as previously described  and estimates of dissociation constants (Kd) were generated. Binding data from 80K co-sedimentation assays demonstrated a 6-fold difference in actin binding between VT (Kd=1.31±0.10 μM) and RE (Kd=7.71±0.04 μM) (Figures 2A and 2B). As RE exhibited weaker actin binding, we assessed the ability of VT or RE to stimulate actin filament formation. F-salt was added to reactions containing 1 μM pyrene-labelled G-actin mixed with various concentrations of VT or RE and then actin polymerization measured by the increase in fluorescence over time. At concentrations of either 0.25 μM or 0.5 μM vinculin tail, VT induced actin nucleation more rapidly than RE (Figures 2C and 2D). Together, these data indicate that RE vinculin is compromised in its ability to bind and stimulate actin polymerization under physiological salt concentrations.
RE is an actin binding and polymerization mutant in physiological salt conditions
Next, we examined the extent of vinculin-driven actin bundling. VT or RE was pre-incubated with 1 μM G-actin, and then F-salt concentrations were added to initiate actin polymerization. The completed reactions were centrifuged at a speed sufficient to pellet bundled actin and actin-bound vinculin peptide. Small amounts of thin, two filament thick actin bundles also co-sedimented in this approach . Binding data from 20K co-sedimentation assays yielded vinculin tail:actin dissociation constants that were nearly identical with those generated from 80 K centrifugations (Kd=1.48±0.04 μM for VT and 7.34±0.05 μM for RE) (Figures 3A and 3B). Furthermore, VT exhibited ~35% greater bundling activity than RE (Figure 3C). Since RE was not completely deficient in actin bundling, we examined the samples by TEM. Reactions containing VT had thick, tightly packed bundles, whereas those containing RE had fewer, thinner, more loosely packed bundles (Figures 3D and 3E). These data support the conclusion that RE is defective in both quantity and quality of actin bundling activity under physiological salt conditions.
RE is an actin bundling mutant in physiological salt conditions
R1049E is an actin-binding and bundling mutant in low ionic strength conditions
We have previously shown that VT is also capable of stimulating actin polymerization under low ionic strength conditions . To assess the ability of RE to trigger filament formation in low salt, we combined various concentrations of VT or RE with 1 μM pyrene-labelled G-actin in the absence of F-salt and then measured polymerization through the increase in fluorescence over time. VT stimulated more actin polymerization than RE at all vinculin tail concentrations examined (i.e. 0.25–10 μM; Figure 4A).
RE is an actin binding, polymerization and bundling mutant in low ionic strength conditions
Next, VT or RE was incubated with 1 μM G-actin in G-buffer, and the resulting product centrifuged at a speed sufficient to pellet actin bundles. In accordance with actin polymerization activity produced in Figure 4A, under the low ionic strength condition the Kd for VT:actin was calculated to be 0.37±0.16 μM and for RE:actin to be 4.48±0.21 μM (Figures 4B and 4C). Furthermore, incubation of actin with VT resulted in 25–40% bundling at concentrations of 0.25 μM, 0.5 μM and 1 μM, which increased to 80% and higher above 2.5 μM VT (Figure 4D). In contrast, the bundling by RE was 20–30% for all concentrations examined. To confirm and visualize this difference in actin bundling, samples were analysed by TEM. The bundles formed when VT was present were several filaments thick and the filaments were tightly packed (Figure 4E). In contrast, those bundles formed when RE was present were thinner, and the filaments were loosely packed (Figure 4F). Along with the studies in Figures 2 and 3, these data demonstrate that RE vinculin tail is compromised in its ability to bind and stimulate the polymerization and bundling of actin filaments in multiple ionic strength conditions.
Vinculin R1049E mutant cells display decreased spreading and adhesion but enhanced migration
We sought to determine if the interaction of vinculin R1049 with actin was a physiologically relevant interaction. Hence, we next sought to examine the consequence of re-expressing GFP-tagged full-length R1049E vinculin (vinRE) in MEFs harvested from the vinculin null mouse . As controls, we also generated lines expressing GFP-tagged full-length vinWT or cytoplasmic GFP (GFP).
After generating and validating the three cell lines by immunoblotting (Figure 5A), we first asked how defective actin binding and bundling affects cell spreading. We performed spreading assays on fibronectin-coated glass at two different time points. Cells were seeded on surfaces coated with fibronectin and allowed to adhere for 25 min or 4 h before fixation and imaging. Consistent with previous studies, we found that cells expressing vinWT spread more robustly and had more prominent actin stress fibres than cells expressing GFP alone at both time points (Figure 5B) . We measured the area of the spread cells and found that at 25 min, vinRE cells exhibit an intermediate ability to spread between that of vinWT and GFP cells (Figure 5C). After 4 h, vinRE cells were still spread to an intermediate degree, ~20% less than vinWT cells, but were more than double the size of the GFP cells (Figures 5B and 5D). The shape of the vinRE cells was comparable with vinWT in that the cells are approximately as long as they are wide, but appeared to have less prominent stress fibres (Figure 5B). This contrasts sharply with the phenotype of the GFP cells, which were typically more than 3-fold longer in one axis than the other (Figure 5B).
Cells expressing R1049E are deficient in spreading, adhesion and migration
Actin stress fibres anchor adhesions to the cytoskeleton, providing stability and support. We hypothesized that the defect in spreading observed in vinRE cells might coincide with a defect in the ability of the cells to adhere. Therefore, we asked what effect the R1049E mutation had on the ability of vinculin re-expressing cells to adhere to a fibronectin-coated substrate. Cells were seeded on fibronectin-coated surfaces, allowed to adhere, and washed. The number of adherent cells was counted and reported relative to the number of adherent GFP cells, which was set equal to one. In this assay, twice as many vinWT cells adhered as vinRE or GFP cells (Figure 5E). This assay did not find a statistical difference between adhesion of vinRE and GFP cells, though vinRE cells tended to adhere better than GFP cells. Thus, the adhesiveness of the vinRE cells is markedly reduced compared with vinWT cells.
As cell spreading and adhesion are both crucial for proper cell migration, we analysed the migration of the three cell lines on fibronectin-coated surfaces. Cells were seeded and allowed to adhere overnight then imaged every 5 min for a total of 10 h. The movies were analysed using NIH ImageJ Manual Cell Tracker plugin. It is well known that vinculin-null cells display enhanced migration compared with those expressing vinculin, and our results here agree with the published data [21,24]. As expected, the GFP cells displayed enhanced migration on fibronectin compared with vinWT cells, with an average velocity of 1.3±0.09 μm/min. The vinWT cells migrated significantly slower with an average velocity of 0.81±0.05 μm/min (Figure 5F). The vinRE cells migrated at an intermediate velocity, 1.0±0.07 μm/min. We also noted that many of the vinRE cells appeared to have ‘trailing tails’ and some were ‘snapped back’ by these tails after attempts to crawl forward.
Vinculin R1049E mutant cells exert less traction force than WT cells
Cell migration depends upon a balance of forces at the leading and trailing edges, requiring that old adhesions must be disassembled in order for cells to move forward. The intermediate phenotype observed in vinRE cells led us to next examine if the traction force exerted by vinRE cells differed from vinWT cells. We used mPADs to measure the traction forces generated by the three cell lines . Cells were seeded overnight on to fibronectin-coated mPADs and developed focal adhesions (Figure 6A). The vinWT cells spread more than both vinRE and GFP cells (Figure 6B). Knowing the micropost stiffnesses, we measured post deflections for vinWT, vinRE and GFP cell lines to quantify cell traction forces. Figure 6(A) shows images of the fibronectin-coated posts (red), with the cell outline visible, and the recruitment of GFP-vinculin (green) to focal adhesions. Force vectors (cyan) were calculated from the deflection of the microposts. As observed previously , the magnitude of the forces varied across the cell, with the highest forces located at the cell periphery. Figure 6(C) presents box-whisker plots for the total traction force per cell, which represents the sum of the magnitudes of the force vectors for each cell. As observed previously the vinculin-null (GFP) cells generated traction force (~50 nN), confirming that vinculin is not required for force transmission at focal adhesions. Expression of WT vinculin increased the traction forces 5-fold compared with the GFP controls, demonstrating that vinculin enhances transmission of traction forces. The expression of the R1049E mutant (vinRE cells) increased the traction force 3-fold compared with the GFP cells, indicating that the R1049E mutant vinculin is capable of enhancing traction forces, though not to the degree of vinWT. Taken together, these data demonstrate that actin-binding deficient vinculin R1049E protein inadequately restores cellular traction forces.
Cells expressing R1049E are defective in generating traction forces
We also generated traction force-cell area plots (Figures 6D, 6E and 6F), as Dumbauld et al.  had previously demonstrated cell area and cytoskeletal tension coupling. We evaluated traction force-cell area coupling as another metric for assessing how vinculin expression modulates traction force generation. There is strong correlation between cell area and traction force generation for null, vinRE and vinWT cells. As observed previously, vinculin null cells exhibited a linear relationship between cell area and traction force, indicating that vinculin is not required for cell area–traction force coupling. This result also suggests other focal adhesion components can play a role in traction force generation, such as talin-actin force transfer . Both vinRE and vinWT expression, however, significantly enhanced coupling between cell area and traction force generation, as demonstrated by the nearly 3-fold increase in the regression slope for both vinRE and vinWT compared with the null cells. Based on the similar traction force–cell area slopes between vinRE and vinWT, this also suggests that the vinRE spreading defect contributes to the weaker total traction force generation compared with the vinWT cells.
In 2006 Janssen et al.  proposed a model for how vinculin binds actin. A more recent model for actin binding to vinculin  called into question the validity of the Janssen Model. In the current study, we determined the consequence of mutating a vinculin residue identified by Jansen et al. as critical for actin binding. Our in vitro data show that this mutant form of vinculin disrupts actin binding and bundling. Furthermore, expression of our mutant vinculin in cells lacking vinculin leads to a reduced ability of cells to spread, adhere and generate traction forces; these cells migrate at enhanced rates. Based on this new information, we prioritize which aspects of the current vinculin tail:actin models are most likely.
The present study is the first to show that the actin binding and bundling by vinculin is required for the generation of traction forces. A previous study showed that a mutant form of vinculin deficient in actin binding was compromised in its ability to stiffen when forces were applied to integrins . Collectively, these data suggest that the actin binding and bundling properties of vinculin are required for force transmission. In our study (Figures 5 and 6), cells with defects in force transmission migrated faster than the vinculin-null MEFs re-expressing vinWT. We found that the vinRE cells migrated faster than wild-type re-expressers, a phenotype reminiscent of parental vinculin null MEFs. Previously it has been shown that migration speed exhibits a biphasic dependency on adhesive force . Hence it is possible that the increased migration rate in the RE cells results from changes in their adhesivity.
In the traction force studies, we found some striking differences between the spreading of cells on fibronectin-coated mPAD devices that measure traction forces when compared with the same cells on fibronectin-coated glass surfaces. On the stiff substrate glass, we consistently observed that vinRE cells show a spreading phenotype that was intermediate between vinWT and GFP cells (Figures 5C and 5D). In contrast, on the mPADs the difference between vinWT and vinRE is more pronounced with the vinRE more closely mirroring the GFP cells (Figure 6). This observation is consistent with previous work showing that fibroblasts spread more on stiffer substrates whereas cells on softer substrates tend to adopt a more spherical phenotype . Consequently, the differences we observe in cell spreading on the two substrates can likely be attributed to their pliability.
It is striking that in all of the biological events we measured, the RE mutant form of vinculin is not completely devoid of activity. Rather, it produces phenotypes that are intermediate between vinWT and vinculin-null (GFP) cells. These intermediate phenotypes are likely due to two possibilities. First, we speculate that the intermediate phenotype could be due to retention of some actin binding and bundling capability in R1049E. The examination of other mutations in vinculin, I997A and V1001A , supports the idea that the greater the actin binding capacity, the better the cell spreading. A second possible explanation for the intermediate phenotypes lies in the ability of vinculin to bind proteins that modulate actin dynamics. Vinculin recruits the Arp2/3 complex (an actin nucleating protein) and VASP (an anti-capper of actin filaments) to focal adhesions. Cells re-expressing P878A vinculin that fails to bind Arp2/3 displayed a number of defects, including decreased cell spreading and lamellipodial extension, an increased length to width ratio that is more similar to vinculin-null cells, and enhanced migration . As is the case with our R1049E mutant, the phenotypes of the P878A mutant are intermediate between WT and vinculin-null cells, indicating that it, too, preserves some of the other functions of vinculin.
We considered the possibility that our R1049E mutation might affect vinculin binding to other tail ligands. However, the binding sites on VT for Ptd Ins (4, 5) P2 Raver1 and paxillin are all distant from R1049. In the case of the focal adhesion protein paxillin, the binding site is within VT residues 881–1000 . The VT-binding portion of Raver1, a heterogeneous nuclear ribonucleoprotein, has been co-crystallized with VT, and the four residues that form the interface are within helices two and three of VT–distant from R1049 . Finally, VT associates with PtdIns (4, 5) P2 , but it is the N-terminal strap and nearby region of VT that is thought to be important for this interaction. As before, R1049E is on the back side of the VT protein, distant from the N-terminal strap. Based on this information, in conjunction with our studies demonstrating that the secondary structure and dimerization capability of R1049E are not perturbed, it is unlikely that our mutation disrupts binding to these VT-associated proteins.
The surface on vinculin involved in actin binding is the subject of much discussion in the field, and several models have appeared in the literature over the past 10 years. The first, the Janssen Model , describes the interaction between chicken vinculin and rabbit skeletal muscle actin, and is based on the reconstruction of negative stain electron microscopy and diffraction data. The Janssen Model proposes two distinct surfaces on vinculin tail as contact points for an upper and lower actin monomer of a filament. In this model, R1049 lies within the lower actin monomer binding site.
Second and third models were proposed by Thompson et al. . To generate these models, the crystal structure of vinculin tail (PDB code 1QKR) and the atomic model of F-actin (PDB code 3MFP) were manually docked into negatively stained 3D EM reconstruction of the rabbit skeletal muscle actin filaments decorated with chicken vinculin tail (Thompson Manual Fit Model) or manually docked and refined using discrete molecular dynamics (DMD; Thompson DMD Model). These newer models propose an alternative interface for vinculin binding to actin. Mutational studies of this alternative vinculin interface (i.e. I997 or V1001) identified actin binding defects . Thompson et al. also mutated a vinculin residue (I948) and found no actin binding deficiency in this mutant. These observations led them to challenge the Janssen Model despite the fact, that although I948 is adjacent to the Janssen predicted interface, this residue has less than 15% probability of interacting with F-actin in the Janssen Model  and was not considered to be part of the interface. Charge-to-alanine mutation clusters scanning studies  do not show any significant effects on actin binding in the regions indicated by Thompson et al. as an alternative interface, but clusters that do show significant effects on actin binding match well with regions of high interaction probability predicted by Janssen et al. .
In the present study, we present evidence that mutation of a residue predicted to be in the interface with over 50% probability by the Janssen Model (i.e. R1049) diminishes actin binding and bundling and has profound effects on the events required for cell migration. Based on this new information, we have re-evaluated which of the existing models most probably reflect how vinculin binds F-actin. In Figures 7(A)–7(C), we show the three models with the actin filaments aligned in the same orientation and with the I997, V1001 and R1049 residues highlighted. In all three models one molecule of the vinculin tail (grey) is interacting with two actin monomers (light green and dark green). In the Janssen Model, the side chain of R1049 is 5.1 Å (1 Å=0.1 nm) away from E100 on the actin monomer suggesting that it is poised to form an electrostatic interaction (Figure 7D). In further support of the existence of such an interaction vinculin binds actin approximately four times tighter under low ionic strength conditions (Figure 4B, Kd=0.37±0.16 μM) than physiological salt conditions (Figures 2A and 3A, Kd=1.31±0.10 μM, Kd=1.48±0.04 μM). Finally, in support of the Janssen Model, R1049 vinculin is proposed to be in direct contact with actin monomer in a MDS of vinculin binding to actin . The Thompson Manual Fit and Thompson DMD Model both have R1049 more than 9.0 or 20.1 Å away from the nearest residue side chain on the actin monomer (Figures 7E and 7F). Moreover, the nearest residue in the Manual Fit Model is a proline residue, which is not expected to form an electrostatic interaction. Hence, our data agree with the Janssen Model more than either of the models proposed by Thompson et al. .
R1049E is a residue predicted to contact actin by the Janssen Model
Collectively, these data [13,15–17,33] raise the question as to which model more accurately represents vinculin binding to F-actin. Data from four groups, including ours [13,16,33] support residues or regions identified as important for the vinculin:actin interaction by Janssen et al. . Two other groups have published data that implicate residues outside of the Janssen Model as important for the vinculin:actin interaction [15,17]. Clearly, the picture is more complex than originally thought, and further mutational studies are needed to resolve how vinculin binds actin with an emphasis on determining if there are multiple binding conformations.
Karry Jannie designed and performed experiments, analysed data and drafted the manuscript. Shawn Ellerbroek and Dennis Zhou designed and performed experiments, analysed data and edited the manuscript. Sophia Chen and David Crompton performed and analysed experiments and edited the manuscript. Andrés García and Kris DeMali conceived experiments and edited the manuscript. All authors approved the manuscript in its final form.
We would like to thank Eileen Adamson for the generous gift of reagents. We are especially grateful to Niels Volkmann, Dorit Hanein and Peter Rubenstein for their thoughtful discussions and critical comments on the work. We thank Kuo Kuang Wen for experimental advice during the initial phases of this project. We thank Lokesh Gakar for assistance with DLS analyses. We thank Xu Liu for assistance with CD analyses and Ernie Fuentes for help with Pymol and discussion of structural models.
This work is supported by the National Science Foundation [grant number 1120478 (to K.A.D.)]; the National Institutes of Health R01 [grant number GM065918 (to A.J.G.); and the NSF Graduate Research Fellowship award (to D.W.Z.). Research reported in this publication was also supported by the National Cancer Institute of the National Institutes of Health under Award Number P30CA086862 to the Holden Comprehensive Cancer Center at the University of Iowa. The data presented herein were obtained in the Central Microscopy and Flow Cytometry Facilities, which are Carver College of Medicine/Holden Comprehensive Cancer Center core research facilities at the University of Iowa. These Facilities are funded through user fees and the generous financial support of the Carver College of Medicine, Holden Comprehensive Cancer Center and Iowa City Veteran's Administration Medical Center.