INTRODUCTION

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A large body of recent work has focused on how cells convert applied mechanical tension into cytoplasmic signals that regulate cell metabolism and transcription (mechanotransduction) (1-3). Important elements of mechanotransduction include cellular adaptation and survival in the face of increased environmental force (4). Indeed, cells undergo dramatic internal structural changes in response to increased environmental forces (5). These mechanoprotective, structural adaptations enable cells to maintain membrane integrity, cell shape, and adhesion to extracellular matrix molecules (4, 6). For example, periodontal ligament fibroblasts function in a much more mechanically stressed environment than gingival fibroblasts and have a nearly 2-fold higher proportion of actin in filamentous form (7). This example indicates that cells are able to sense and adapt to environmental tension in part through cytoskeletal adaptations. NIH/3T3 fibroblasts up-regulate their attachment strength to extracellular matrix ligands if increased tension is applied through integrins (8), indicating that cells not only sense changes in applied extracellular loads but can rapidly reinforce cytoskeletal linkages locally at force application sites. Consistent with these data we have demonstrated that fibroblasts undergo localized actin assembly during isolated force application through focal adhesion complexes (6).

Localized cortical actin mechanoprotective responses presumably involve actin-binding proteins. The cross-linking and bundling activities of actin-binding proteins can increase the structural strength and integrity of the cortical actin network (9). Among the most efficient actin cross-linking proteins is actin-binding protein 280 (ABP-280),¹ a 540-kDa dimeric protein first identified in macrophages (10) and present in other tissues including most non-muscle cells (11). A homologous protein, filamin, first purified from skeletal muscle (12, 13) shares extensive sequence homology with ABP-280 but is encoded by a separate gene and displays different abilities to cross-link or bundle F-actin. ABP-280, filamin, dystrophin, spectrin, -actinin, ABP-120, and fimbrin are part of an actin cross-linking superfamily (14) that share a common actin-binding site (11). Some members of this superfamily (spectrin and dystrophin) may act to mechanically reinforce the cell membrane (15) and thereby enhance membrane stability during increased membrane tension. Notably, ABP-280 cross-links actin and links actin to integral membrane proteins (16, 17), thereby providing increased cellular cortical rigidity (18). Thus ABP-280 and other actin cross-linking and bundling proteins may be key structural elements that stabilize the cell membrane by facilitating interactions between the cortical actin cytoskeleton and the plasma membrane.

The importance of structural interactions between the cortical cytoskeleton and the plasma membrane in ion channel regulation has been recognized previously (3, 19, 20). Specifically, ABP-280 has been implicated in regulating the conductance of ion channels activated by cell swelling (21). Since chronically high Ca²⁺ entry is known to be toxic to cells (22), it is likely that cells in mechanically active environments must have evolved adaptive mechanisms to regulate the sensitivity of SACs to chronic or prolonged high membrane tension. We have suggested previously that the open probability of SACs is dependent on the rigidity of the membrane cortex which is determined by specific cytoskeletal proteins and their organization (6). In this study we have characterized a localized, force-induced actin recruitment to focal adhesions and have examined the dependence of this recruitment on ABP-280. In this article, we test the hypothesis that ABP-280 mediates a mechanoprotective mechanism that desensitizes SACs and is important for cell survival in the face of applied physical stress.

	EXPERIMENTAL PROCEDURES

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Reagents-- Anti-filamin mouse monoclonal antibody (clone FIL2), cross-reactive with ABP-280, anti-filamin rabbit polyclonal antibody, anti-vinculin mouse monoclonal antibody (clone VIN-1), -actinin mouse monoclonal antibody (clone BM 75.2), -actin monoclonal antibody (clone AC-15), fluorescein isothiocyanate-conjugated goat anti-mouse antibodies, and TRITC-phalloidin were purchased from Sigma. Mouse monoclonal antibody to ₂-integrin was purchased from Calbiochem (clone P1E6). Anti-phosphoserine antibody was purchased from Zymed (San Francisco, CA). Anti-villin mouse monoclonal antibody (clone ID2C3) was purchased from Biodesign International (Kennebunk, MA). Anti-MARCKS mouse monoclonal antibody was purchased from Upstate Biotechnology (Lake Placid, NY). Fura-2/AM was purchased from Molecular Probes (Eugene, OR). All antibodies were against human antigens. Cytochalasin D was purchased from Sigma. Bisindolylmaleimide and calphostin C were purchased from Calbiochem.

Cell Culture-- Human gingival fibroblasts were derived from primary explant cultures as described (7). Cells from passages 6-19 were grown as monolayers in T-75 flasks (3). Twenty-four hours before each experiment cells were harvested with 0.01% trypsin, and 2 × 10⁵ cells were plated into 60-mm diameter culture dishes (Falcon, Becton Dickinson, Mississauga, ON). The cells were sub-confluent prior to all experiments.

Force Generation-- A force generation model was used as described previously (3, 6, 23). Briefly, a ceramic permanent magnet (grade 8, 2.2 × 9.6 × 11 cm; Jobmaster, Mississauga, ON) was used to generate physical forces that lasted longer than 5 s as described previously (6). For all experiments the pole face was parallel with and 2 cm from the cell/culture dish surface unless specified. At this distance the force generated was 0.48 pN/µm² (6). A constant force of varying durations was used for all experiments, which induced an upward force on the cell. Thus, applied forces were perpendicular to the dorsal surface of the cell. The force generated by application of the magnetic field to a 4.5-µm bead was determined as described previously (3, 23).

Electron Microscopy-- Fibroblasts were permeabilized with 10% PHEM (0.6 M Pipes, 0.25 M Hepes, 0.1 M EGTA, 20 mM MgCl₂), 0.75% Triton X-100, and 1% glutaraldehyde. After 30 min, samples were embedded in Lowicryl-K4M, and thin sections were placed on nickel grids. Gold-conjugated (15 nm diameter) goat anti-mouse IgG was obtained from Zymed (San Francisco, CA). Sections were blocked with a 0.2% gelatin, 0.1% BSA in TBS solution for 1 h. The grids were washed in PBS, 0.1% BSA buffer and placed on a 25-ml drop of anti-filamin/ABP-280 monoclonal IgG antibody (10 mg/ml, diluted in PBS, 0.1% BSA buffer) and incubated for 1 h at room temperature. Samples were washed as described above. The grids were placed on a 25-µl drop of the secondary gold-conjugated goat anti-mouse IgG (1:20, diluted in PBS, 0.1% BSA). The grids were stained with uranyl acetate and lead citrate and observed under an electron microscope (Hitachi-60). The number of gold particles per 0.25-µm² area was counted to determine the labeling index of the cell sections for each group. Background counts (sections only incubated with the secondary antibody) demonstrated less than 0.5 beads per µm².

Immunofluorescence-- Cells grown on coverslips were fixed with 3.7% formaldehyde in PBS for 10 min, stained with TRITC-phalloidin, and examined under both phase contrast and fluorescent 20 × objectives on a Diaphot microscope (Nikon). Images were observed on a microscope connected to a CCD camera (Pentamax, Princeton Instruments, NJ) and stored on a computer. Images were processed using the software package Winview 1.6.1 (Princeton Instruments). For antibody staining cells were permeabilized in 0.3% Triton X-100 in PBS for 15 min at room temperature. Cells were incubated with primary antibody (anti-filamin; 1:80 dilution) for 1 h at 37 °C, washed 3 times with PBS containing 0.03% Triton and 0.2% BSA, and incubated with fluorescein-conjugated goat anti-mouse (1:100 dilution). Nonspecific control staining was performed on the same slide using secondary antibody only. Coverslips were washed with PBS and mounted with an anti-fade mounting medium (ICN).

Fluorescence Quantitation of Actin-- Images of TRITC-phalloidin-stained fibroblasts acquired in Winview were assessed for F-actin accumulation/enrichment at bead binding sites using the pixel fluorescence function in the Winview 1.61 software. Paraformaldehyde-fixed cells from no force and force-treated samples were stained with rhodamine-phalloidin; the cells were imaged, and the rhodamine fluorescence (F-actin) level (average pixel intensity) at bead sites on a given cell were divided by the average pixel intensity for the entire cell. This provided a measure of the percent F-actin enrichment at bead binding sites.

Intracellular [Ca²⁺]-- Measurement of intracellular calcium ion concentration ([Ca²⁺]_i) was conducted as described previously (3). Briefly, cells on coverslips were incubated at 37 °C with 3 mM fura-2/AM (Molecular Probes, Eugene, OR) for 20 min and then at room temperature for 10 min. Whole cell [Ca²⁺]_i measurements were obtained with a dual excitation, microscope-based spectrofluorimeter (Photon Technology Int., London, ON). A variable aperture, intra-beam mask was used to restrict measurements to single cells. Estimates of [Ca²⁺]_i independent of the precise intracellular concentration of fura-2 were calculated from dual excitation emitted fluorescence as described (24). As demonstrated previously by image analysis (3) and by microscopic evaluation between exposures, repeated applications of force did not remove attached beads. Force application during calcium measurements was applied as described previously (3). Briefly, an electromagnet with a pole extension was used to focus and direct the magnetic field to the cell of interest.

Isolation of Bead Complexes-- Proteins enriched in bead-associated focal adhesion complexes were assessed as described previously (6, 25). Briefly, cells and attached beads were collected by scraping cells into ice-cold cytoskeleton extraction buffer (CSKB, Triton X-100, 50 mM NaCl, 300 mM sucrose, 3 mM MgCl₂, 20 µg/ml aprotinin, 1 µg/ml leupeptin, 1 µg/ml pepstatin, 1 mM phenylmethylsulfonyl fluoride, 10 mM Pipes (pH 6.8)). The isolation procedure was carried out at 4 °C using a side-pull magnetic isolation apparatus (Dynal, Lake Placid, New York). The cell-bead suspension was sonicated for 10 s (output setting, 3, power 15%; Sonifier 185, Branson) and homogenized in a 2-ml Dounce homogenizer (20 strokes). The magnetic beads were pelleted and washed 3 times with CSKB prior to protein analysis. Equal numbers of beads from both "force" and "no force" samples were suspended in equal volumes of sample buffer and boiled for 5 min to remove protein from the beads. Protein levels were estimated using densitometry of Western blots. Standard curves were performed for each protein from isolates obtained from beads, and a large concentration range (1-20 µg) of protein was loaded on each lane. The standard curves demonstrated a linear relationship between protein loading levels, and densitometry measurements for all the proteins that were examined between 0 and 20 µg (Pearson correlation R² values were between 0.89 and 0.97 for all proteins tested). The optimal protein concentration loaded per gel was 10 µg maximum, which was well within the linear range as determined from each of the standard curves.

Inhibitors-- We assessed the dependence of actin rearrangement on calcium ions by incubating cells with BAPTA/AM at 3 µM for 45 min at 37 °C prior to force application. Previous pilot experiments have shown that this reduces [Ca²⁺]_i to <50 nM and blocks ligand-induced calcium fluxes. We assessed the dependence of actin rearrangement on actin assembly by incubating cells with cytochalasin D at 1 µM for 30 min at 37 °C prior to force application.

Immunoblotting and Immunoprecipitation-- ABP-280, villin, MARCKS, -actinin, ₂-integrin, and serine-phosphorylated proteins were identified by immunoblotting. Isolated proteins were separated by SDS-polyacrylamide gel electrophoresis (7% acrylamide) and transferred to nitrocellulose. Blots were blocked for 1 h with 5% skim milk in PBS and incubated in the indicated antibody for 1 h at room temperature. Blots were washed three times with 0.5% Tween/PBS for 10 min, incubated with goat anti-mouse horseradish peroxidase or anti-rabbit horseradish peroxidase (Amersham Corp.) for 1 h, washed 5 × in PBS/Tween, and developed by chemiluminescence (Amersham Corp.). Serine-phosphorylated ABP-280 was identified by immunoprecipitation and immunoblotting. Isolated protein from plated samples were incubated with protein G-Sepharose beads that had been incubated with anti-filamin (polyclonal;10 mg/ml beads) overnight at 4 °C. Beads were collected by centrifugation in a microcentrifuge. The precipitate was washed six times. Protein was separated from beads by heating at 65 °C for 10 min in 2 × Laemmli buffer.

Phagocytosis-- Phagocytosis was analyzed as described previously (29). Briefly polystyrene microbeads (2 µm diameter; Molecular Probes, Eugene, OR) were coated with collagen as described above. Cells were incubated with beads at a ratio of 4 beads per cell for 3 h. Cell were trypsinized to create a cell suspension and remove externally attached beads. Ten minutes prior to analysis propidium iodide (50 mg/ml) was added to cell suspensions to estimate the proportion of non-viable cells. Internalization of beads and cell viability was assessed by dual color flow cytometry as described previously (30). Briefly, cell samples were analyzed with a FACStar Plus flow cytometer (Becton Dickinson FACS Systems, Mountain View, CA) at a sheath pressure of 11 p.s.i. and with excitation from an Innova 70 argon laser (Coherent Laser, Palo Alto, CA) at light regulation mode setting of 250 milliwatts and a wavelength of 488 nm. Emitted fluorescence was split between two detectors through a short pass 560 beam splitter (all filters and beam splitters from Omega Optical Inc., Brattleboro, VT) and a 530DF30 filter for green fluorescence (phagocytosed beads) and a 625DF38 filter for red fluorescence. Photomultiplier tube voltage settings were determined for each experiment on the basis of thresholds established from appropriate negative and positive control samples.

Motility Assay-- Confluent monolayer cultures of gingival fibroblasts on 60-mm dishes were "wounded" by creating a uniform cell-free wound using a scalpel (31). The wound length was approximately 1 mm in length. The maximum migration distance for untreated control cells was 500 µm over the assay period. Hepes-buffered -minimum essential medium containing 15% fetal bovine serum was added to the culture, and migration of the cells into the wound was visualized by time lapse cinemicrography (Panasonic AG6050 recorder; Nikon microscope) of the wound area for 15 h with a 20 × objective (Nikon). Tracings of the wound area were made before and after force application, and the cell-free area was calculated using NIH IMAGE (version 1.6) on a MacIntosh computer. The migratory index was expressed as the percent area of the original wound that was repopulated with cells.

Statistical Analysis-- For all assays, three or more separate experiments were performed; means ± S.E. were calculated for continuous variables, and comparisons were made by unpaired t tests or analysis of variance as indicated.

	RESULTS

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F-actin Accumulation-- We studied force-induced actin assembly locally at the force transfer sites by isolating the proteins associated with collagen-coated magnetic beads. Identification by Western blot of cell membrane proteins bound to the magnetic beads showed that the bead attachment sites were enriched with vinculin, actin, and ₂-integrin but that the levels of vinculin and ₂-integrin did not change over time of force exposure (20 min; Fig. 1A). In contrast, there was a marked increase (1.5-2-fold) in the amount of actin associating with the beads (Fig. 1A). The actin accumulation following force application was confirmed by electron micrographs of bead-containing sections from fibroblasts exposed to force. Prominent filament bundles were observed in close proximity to collagen beads from force-treated samples but were absent in samples incubated only with the collagen beads (Fig. 1B). Lower magnification micrographs demonstrated increased density of actin fiber bundles in cells exposed to force with some of the bundles oriented toward the substrate surface (i.e. parallel to the applied force). To verify the Western blot data a fluorescence-based image analysis technique was developed to visualize and quantify the enrichment of F-actin at bead attachment sites. The average pixel intensity around beads of rhodamine-phalloidin-stained samples was enriched 1.5-fold in force samples compared with no force samples (No force: 1.15 ± 0.074; Force: 1.79 ± 0.089; p < 0.001; mean ± S.E.).

View larger version (65K): [in this window] [in a new window]   Fig. 1.   A, Western blots of proteins isolated from collagen-coated magnetic beads before and after application of force to gingival fibroblasts. Blots demonstrate that beads bind through 2-integrins and associate with focal adhesion proteins (vinculin). Force induces increased actin to associate with integrin bound beads without increasing levels of the adhesion complex proteins. Protein from equal numbers of beads was loaded in each lane. B, schematic diagram demonstrating the approximate dorsal-apical level of micrographs. 1, electron micrographs of the most dorsal surface of gingival fibroblasts bound with magnetic beads. Micrographs of force samples clearly demonstrate increased filament assembly associating with the bead complex compared with no force samples. Note that beads were removed during sectioning (clear zone). A bead fragment is still present in the right corner of the force micrograph (bar = 0.5 µm). 2, lower magnification micrographs demonstrate increased fiber bundles in force-exposed cells (bar = 40 µm).

View larger version (31K): [in this window] [in a new window]   Fig. 2.   Western blots demonstrating that the force-induced increase of actin at bead binding sites is dependent on calcium ions and actin assembly. Proteins from gingival fibroblasts were obtained by the bead isolation technique (see "Experimental Procedures"). Elimination of intracellular and extracellular calcium ions (3 µM BAPTA/AM and calcium-free buffer) prevented force-induced actin accumulation. Prevention of actin assembly with cytochalasin D (1 µM) also prevented the force-induced actin accumulation.

Actin-binding Proteins-- Western blots of the proteins isolated from the beads were screened for a number of actin-binding proteins that have been implicated in F-actin cross-linking and F-actin membrane association. Proteins that were probed included ABP-280 (18), -actinin (32), villin (33), ₂-integrin, and MARCKS (34). Densitometry of Western blots indicated that only ABP-280 was significantly enriched at the force transfer sites after force application (F-actin: No force (NF) versus force (F), p < 0.01; ABP-280: NF versus F, p < 0.01; all other proteins: NF versus F, p > 0.20; Fig. 3A).

View larger version (43K): [in this window] [in a new window]   Fig. 3.   A, densitometry of Western blots of lysates probed for -actin, ABP-280 (non-muscle filamin), -actinin, villin, MARCKS, and 2-integrin. Proteins from gingival fibroblasts were obtained by the bead isolation procedure. Western blots were scanned, and the density of the protein in the force samples was divided by the density in the control (No Force) sample. Densitometry of Western blots demonstrates that only ABP-280 increases at the bead-membrane complex during constant force exposure, whereas the levels of the other actin-binding proteins and the 2-integrin of the collagen receptor remain constant (results are representative of at least three separate experiments; error bars are S.E.; asterisks indicate p < 0.01 comparing Force to No Force accumulation). B, bead/protein co-localization: high magnification paired images of individual beads: fluorescence images/F-actin (right) and corresponding phase contrast image (left) showing bead size and location relative to actin/ABP-280 accumulation. Force induces ABP-280 accumulation at membrane/bead sites. Note minimal ABP-280 accumulation at bead/membrane sites in no force samples. Immunofluorescence microscopy of TRITC-phalloidin-stained cells demonstrates force-dependent actin accumulation at membrane/bead sites. Minimal detectable F-actin accumulation directly at bead sites is observed at bead/membrane sites in no force samples. Although some actin stress fibers can be seen in the vicinity of the bead, they do not appear to coincide with the bead. C, force-induced ABP-280 increase at bead binding sites is dependent on calcium ions and actin assembly. Elimination of intracellular and extracellular calcium ions (3 µM BAPTA/AM and calcium-free buffer) prevented the force-induced ABP-280 accumulation. Prevention of actin assembly with cytochalasin D (1 µM) also prevented the force-induced ABP-280 accumulation.

Regulation of ABP-280-- ABP-280 has been described as a phosphoprotein (35), and the regulatory mechanism for the localization and actin association of ABP-280 involves serine phosphorylation (36, 37). To determine if ABP-280 was serine-phosphorylated in gingival fibroblasts, we immunoprecipitated ABP-280 and Western blotted for serine-phosphorylated proteins in whole cell lysates. ABP-280 was serine-phosphorylated during force application (Fig. 4A). Control experiments showed that force did not increase total cell ABP-280 levels over the experimental time frame (data not shown). We also examined ABP-280 levels and the serine phosphorylation of ABP-280 in proteins prepared from beads (Fig. 4B). Force increased ABP-280 levels 4-fold in bead preparations. Serine-phosphorylated ABP-280 was increased 8-fold by force indicating that the increased phosphorylated ABP-280 seen at bead sites was not simply due to the presence of more ABP-280 but also because of an increased number of phosphorylated serine residues. As we hypothesized that serine phosphorylation of ABP-280 may be a critical regulatory step in force-induced actin cross-linking, the protein kinase C (PKC) inhibitor bisindolylmaleimide (BIM; 5 µM) (26) was used to determine if force-induced serine phosphorylation was mediated through PKC (28). BIM decreased the force-induced serine phosphorylation of ABP-280 (Fig. 4A) and reduced the level of force-induced ABP-280 accumulation at bead sites (Fig. 4B). BIM also reduced the amount of actin accumulating at the force transfer sites. Notably, the only other serine-phosphorylated protein present in the bead isolates was approximately 70 kDa, but the level of serine phosphorylation and accumulation of this 70-kDa protein at bead binding sites appeared to be independent of PKC since BIM did not affect its accumulation or phosphorylation state. To verify the role of PKC in the force-induced actin reorganization, a second specific PKC inhibitor calphostin C was used (27). Calphostin inhibited the force-induced serine phosphorylation, ABP-280, and actin accumulation to the same extent as the BIM (data not shown).

View larger version (44K): [in this window] [in a new window]   Fig. 4.   A, gingival fibroblasts were exposed to the indicated treatment combinations of force (20 min) and the PKC inhibitor bisindolylmaleimide (BIM; 5 µM), and ABP-280 was immunoprecipitated from fresh cell lysates. The blot was probed with an anti-phosphoserine antibody. Note the marked increase in ABP-280 serine phosphorylation following force application which is inhibited by treatment with the PKC inhibitor. Equal ABP-280 was loaded in each lane as determined by parallel Western blot. B, proteins isolated from the beads were probed with the indicated antibody. Note the increase of serine-phosphorylated ABP-280, actin, and a serine-phosphorylated 70-kDa protein following force application. Inhibition of ABP-280 serine phosphorylation by BIM decreased the association of ABP-280 and actin association with the beads but not the 70-kDa protein which may be phosphorylated by a different force-activated kinase.

ABP-280 Knockouts-- To verify that the actin accumulation was dependent on ABP-280, a melanoma cell line that does not express ABP-280 (M2) was used to examine the force-induced actin redistribution. By using the single cell fluorescence method described above, we found a >95% increase in force-induced actin accumulation at bead sites from ABP-280+ cells (A7; p < 0.01) compared with only a 12% increase in the ABP-280-deficient cells (p > 0.1; Fig. 5).

View larger version (25K): [in this window] [in a new window]   Fig. 5.   Filamentous actin accumulation at bead sites. TRITC-phalloidin-stained actin was quantified using Winview software, and the average pixel fluorescence was quantified at bead sites. Bead-derived fluorescence was divided by the average pixel fluorescence for the entire cell (RIF). Note that force increased the actin enrichment at bead sites to a much greater extent in cells expressing ABP-280 (A7) compared with ABP-280-deficient cells (M2). Data are representative of at least three separate experiments (mean ± S.E.). Asterisk indicates p < 0.01 with respect to No force sample.

SACs-- As suggested previously (3, 6), the cortical actin and the force-induced actin accumulation in particular may play a role in regulating stretch-activated ion channel (SAC) activity. To assess this potential downstream mechanoprotective effect, we measured calcium ion influxes through SACs following a brief (1-s) force pulse. Net calcium influx levels in fibroblasts following the actin/ABP-280 reorganization demonstrated a 68% decrease in the stretch-induced calcium ion influx (Fig. 6A; No preforce versus Preforce: p < 0.01). The force-induced cytoskeletal dependent decrease was abolished when the cells were treated with cytochalasin D which as we have demonstrated previously prevents force-induced actin accumulation (7) (Fig. 6A). We determined if there was a force time/dose-dependent reduction in the decrease of the SAC response to force. Increasing the length of the force exposure induced a time-dependent decrease in SAC responses (Fig. 6B).

View larger version (20K): [in this window] [in a new window]   Fig. 6.   A, calcium influx through stretch-activated channels in gingival fibroblasts. The peak calcium influx after a 1-s force application is demonstrated here. Pre-force samples were exposed to 20 min of constant force (0.48 pN/µm2) prior to calcium monitoring during a 1-s force application (0.1 pN/µm2). The contribution of the actin cytoskeleton to regulation of stretch-sensitive cation-permeable channels was assessed by incubating cells with cytochalasin D (1 µM, 20 min; Sigma) prior to force application or during the 20-min pre-force application, a protocol that causes a 3-fold reduction of cortical actin in human gingival fibroblasts (7). Each value represents the mean ± S.E. of at least four determinations. Asterisks indicate p < 0.01 with respect to control, determined by two-way analysis of variance. B, time course of the mean increase in [Ca2+]i after a 1-s stretch demonstrating that in gingival fibroblasts pretreated with constant force for the indicated times there was a time-dependent decrease in the stretch-activated calcium-permeable channel activity. C, force-induced calcium influx in melanoma cell lines. ABP-280-deficient cells (M2) demonstrate a 70% greater force-induced calcium influx compared with the control ABP-280 expressing cells (A7). Following 20 min of force application the ABP-280+ cells demonstrate a greater than 85% decrease in the calcium influx compared with ABP-280 cells which only demonstrate a 50% decrease in calcium influx.

View larger version (12K): [in this window] [in a new window]   Fig. 7.   A, viability of melanoma cells as assessed by flow cytometry and propidium iodide (PI) staining. Viable cells excluded propidium iodide and were therefore not detected in the red channel. Application of force induced a 93% increase of propidium iodide-stained cells in ABP-280 negative (M2) cell line compared with a 46% increase in ABP-280 expressing (A7) cells (p < 0.01). B, actin-dependent processes. Phagocytosis is reduced during force application in gingival fibroblasts. Phagocytosis was measured using a collagen fluorescent bead assay and flow cytometry to assess the proportion of collagen phagocytosing cells. The reduction of phagocytosis was dependent on the magnitude of the force. C, force-dependent reduction of motility in gingival fibroblasts. The effect was dependent on the magnitude of the applied force. Motility was assessed using an in vitro wound system in which the ability of cells to migrate into a cell-free zone was measured over 15 h with or without force. The migratory index is expressed as the percent area of the cell-denuded wound which is repopulated during the experiment.

Actin-dependent Processes: Phagocytosis and Motility-- To determine the potential downstream effects of the force-induced cytoskeletal rearrangement, cellular functions dependent on the assembly of cortical actin filaments and extracellular matrix adhesion were investigated during force application in gingival fibroblasts. Assays for collagen phagocytosis and cell motility were performed during force application. Using a well characterized fluorescent bead phagocytosis assay that requires actin assembly (38), we found that although fibroblasts were under tension there was a significant inhibition of collagen bead phagocytosis. This inhibition was dependent on the magnitude of the applied force (Fig. 7B). At a stress level of 0.4 pN/µm² there was an 80% reduction in phagocytosis. Propidium iodide exclusion was used to verify that the decrease in phagocytosis was not due to the inclusion of dying cells in the assay.

	DISCUSSION

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The important question of how cells protect themselves and adapt to increased environmental tension remains largely unanswered. By using a novel force application model (6), we studied actin recruitment to focal-like contacts through which the tension was applied. Our principal finding is that force induces a local actin accumulation at force transfer sites that is dependent on the co-localization and modification of ABP-280. Previous studies have focused on whole cell cytoskeletal changes during force application (5). These types of studies do not provide information on local changes in the cell cortex that may be important for signal transduction and cellular mechanoprotective processes. We have used a magnetic bead model that allowed the application of defined, localized forces directly through integrin receptors to the actin cytoskeletal complex. This method also permits isolation of proteins that localize to these force transfer sites and thereby facilitates analyses of local changes during increased membrane and cytoskeletal tension. By using atomic force microscopy, we have previously demonstrated that this model induces localized changes in actin assembly that result in increased local rigidity. The localized changes in actin assembly occurred without causing detectable changes in global actin architecture or the global cellular balance of actin monomer/filament (6).

Role of ABP-280-- The force-induced actin accumulation was not a result of increased integrin clustering or increased number of focal contacts since vinculin and ₂-integrin levels in Western blot analyses of bead-derived proteins were unchanged after force application. Consequently we endeavored to determine if any actin-binding proteins were components of the local force-induced assembly and accumulation. Examination of a group of actin-binding proteins that have been previously associated with regulating actin form and architecture showed that of the proteins examined, only ABP-280 was enriched at the force application sites. The importance of this ABP-280 enrichment in mediating actin accumulation was demonstrated in a human melanoma cell line that does not express ABP-280. These cells did not accumulate actin after application of force; however, when ABP-280 was expressed in these same cells, the localized mechanoprotective actin response was restored. The ABP-280 accumulation explains in part the localized increase in membrane rigidity following force application. ABP-280 has been shown to increase the rigidity of actin solutions in vitro (39). Furthermore, cells expressing ABP-280 have more than a 2-fold greater elastic modulus (cortical rigidity) compared with corresponding ABP-280-deficient cells (18).

ABP-280 Regulation-- We demonstrated that the force-induced cytoskeletal response is dependent on calcium ions and actin polymerization since both chelation of free cytoplasmic calcium ions and cytochalasin D treatment inhibited the accumulations of both ABP-280 and actin. However, since ABP-280 was required for actin accumulation, we sought to determine what specific regulatory pathway was involved in the force response. As ABP-280 is a phosphoprotein with more than 380 serine/threonine residues (11), we hypothesized that phosphorylation may be an important regulatory process for localized accumulation in the bead complex (37). The data showed that ABP-280 was phosphorylated on serine residues following force application. From the amino acid sequence of ABP-280, 33 potential PKC sites have been deduced (11), and this suggested that ABP-280 may be phosphorylated by PKC. Indeed 10 of the 33 PKC phosphorylation sites are clustered near the N terminus which contains the actin-binding domain (36). This observation supports the idea that PKC may be involved in regulating the ability of ABP-280 to bind actin. We found that BIM and calphostin C, potent inhibitors of PKC, reduced the serine phosphorylation induced by force and also reduced the amount of ABP-280 localizing to the bead/force application site. This finding suggests that serine phosphorylation plays an important role in regulating ABP-280 force-induced actin binding and that PKC is one of the kinases involved in this event. Support for this regulatory mechanism comes from Wu and co-workers (36) who demonstrated the existence of four phosphorylated forms of ABP-280 in platelets and showed that the more phosphorylated form is able to cross-link twice as much actin as the lesser phosphorylated forms.

SAC-- Mechanoprotective responses likely involve regulation of stretch-activated ion channels (SAC) since chronic force application without SAC desensitization could lead to pathologically high calcium levels (22). As we have previously demonstrated (3, 6), the actin cytoskeleton does play a regulatory role in SAC activation. In the present report, cells with localized cytoskeletal accumulation exhibited decreased SAC activity, an effect that was reversed by cytochalasin D. To determine if ABP-280 plays a role in regulating SAC sensitivity, we studied the stretch-induced calcium influx in ABP-280-deficient cells. There was a markedly increased calcium influx in the ABP-280-deficient cells compared with the ABP-280+ cells suggesting that ABP-280 reduces the open probability of SACs possibly through a tension absorption mechanism. Previous work with the same ABP-280-deficient cell line demonstrated increased basal permeability to K⁺ ions and the lack of a regulatory volume decrease in response to osmotically induced stretch. These alterations were thought to be caused by deficient linkages between the actin cortex and the membrane (43). Consistent with this hypothesis we suggest that ABP-280 and actin interactions are part of a sensing mechanism required to regulate ion transport at the plasma membrane.

Membrane Stabilization-- Applied force apparently shifts the actin monomer/filament equilibrium in cortical regions toward gelation which in turn promotes the formation of a protective shell. The protective nature of this response is suggested by the observation that compared with ABP-280+ cells, ABP-280-deficient cells demonstrated significant elevations of propidium iodide staining after increasing membrane tension indicative of plasma membrane disruption. The increased gelation in the cortical region also affects downstream actin-dependent events such as motility and phagocytosis. In an in vitro motility model, increased gelation due to higher levels of ABP-280 is associated with inhibition of filament velocity and reductions in the number of moving filaments (44). ABP-280-induced gelation of the actin cytoskeleton also dramatically inhibits the rate of gel contraction (45). Based on this previously mentioned work and our data, we suggest that the force-induced cross-linking of cortical actin filaments decreases actin filament turnover which is required for rapid ruffling and pseudopod extension (46).