PKC Regulation of Microfilament Network Organization in Keratinocytes Defined by a Pharmacological Study with PKC Activators and Inhibitors

EXPERIMENTAL CELL RESEARCH ARTICLE NO.

236, 238–247 (1997)

EX973721

PKC Regulation of Microfilament Network Organization in Keratinocytes Defined by a Pharmacological Study with PKC Activators and Inhibitors Benedicte Masson-Gadais, Paul Salers, Pierre Bongrand, and Jean-Claude Lissitzky1 Laboratoire d’Immunologie, Hoˆpital Sainte Marguerite, Unite´ INSERM U387, 270 Boulevard Sainte Marguerite, 13274 Marseille Cedex 9, France

The modulation by PKC activators and inhibitors of adhesion, spreading, migration, actin cytoskeleton organization, and focal complex formation in keratinocytes attaching to type I collagen was studied. Two actin microfilament networks, stress fibers and cortical actin, could be distinguished on the basis of cellular distribution and opposite regulation by growth factors, tyrosine kinase inhibitors, and PKC activators. Stress fiber formation was stimulated by growth factors and by PMA (100 ng/ml) and these stimulations were blocked by tyrosine kinase inhibitors (0.3 mM genistein and 1 m M herbimycin A). By contrast, the cortical network occurred in quiescent cells, was unaffected by tyrosine kinase inhibitors, and was broken down after PKC activation by PMA. Spreading, migration, and actin polymerization were completely blocked while adhesion efficacy was significantly decreased by three specific PKC inhibitors. Half-inhibition of migration was obtained with 0.025, 1, and 3 m M concentrations of calphostin C, chelerytrine chloride, and D-erythrosphingosine, respectively, which are concentrations close to those known to inhibit the PKC kinase function in vitro. Paxillin clustering, which was observed even in the presence of tyrosine kinase inhibitors, disappeared only when actin polymerization was completely impaired, i.e., in cells treated with PKC inhibitors or with both tyrosine kinase inhibitors and PMA, which indicated that focal complex formation was highly dependent on microfilament reorganization. The analysis of these data underscores a major regulation function of PKC in the molecular events involved in growth factor and adhesion-dependent regulation of microfilament dynamics. q 1997 Academic Press INTRODUCTION

Cell adhesion requires a connection between the cytoskeleton and the cell surface receptors interacting 1 To whom correspondence and reprint requests should be addressed. Fax: 0491 75 73 28. E-mail: [email protected].

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0014-4827/97 $25.00 Copyright q 1997 by Academic Press All rights of reproduction in any form reserved.

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with ligands located in the extracellular matrix or on the neighboring cell [1]. The cell adhesion a/b heterodimeric integrin receptors which share a b1 subunit may be linked to the actin microfilament network [2] by talin and a-actinin, which bind to the cytoplasmic domain of the b1 integrin subunit [3, 4] and serve as nucleation sites for a supramolecular assembly of additional cytoskeletal components (tensin, vinculin, cortactin, paxillin) anchored to the actin–microfilament network [5]. In cells adhering to extracellular matrix proteins in vitro, integrins and connecting molecules form clusters called focal contacts, which are associated with actin filaments [5]. Many signaling enzymes, e.g., protein kinases and phosphatases, phospholipid kinases, phospholipases, and GTPases, are also recruited to adhesion sites [6]. They may participate in adhesion regulation and in adhesion-dependent modulation of cell differentiation and proliferation [7]. The molecular process leading to focal contact buildup is not fully understood, although it requires a synergy between actin polymerization and a signal triggered by integrin engagement, possibly the functional activation of tyrosine kinases such as focal adhesion kinase and src, which accumulate in focal contacts [1, 5]. Actin polymerization is controlled by the GTPases cdc42, rac, and rho. These proteins are activated by growth factors [8] and they lead to the production of specific microfilamentous structures since cdc42, rac, and rho induce the formation of filipodia, lamellipodia, and stress fibers, respectively [9]. Thus adhesion involves many molecular interactions and regulation by growth factors via the actin polymerization function of the rho-like GTPases. These interactions may be controlled by protein tyrosine phosphorylations, as suggested by the observations that adhesion and associated cytoskeletal remodelings are altered by tyrosine kinase inhibitors [10]. Tyrosine kinases seem to be involved in several steps of the adhesion process. In fact, integrin engagement is associated with the tyrosine phosphorylation of many proteins which participate in receptor–cytoskeleton linkage [5] and, conversely, ty-

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rosine kinases also seem to be involved in the breakdown of adhesion complexes [11]. Obviously the growth factor regulation of GTPase function involves tyrosine kinases since many growth factor receptors are tyrosine kinases. Furthermore, a tyrosine kinase operating downstream of rho appears instrumental in stress fiber formation [12]. Protein kinases C (PKC),2 a family of serine/threonine kinases composed of at least 11 isoforms [13], may also participate significantly in adhesion regulation. Like tyrosine kinases, PKC may be implicated in the signal transduction pathways which are switched on by growth factors and end up in GTPase activation. Schematically, the lipid stimulation of PKC by diacylglycerols or 3*-phosphorylated phosphoinositides which are produced after phospholipases or phosphatidylinositol 3-kinase activation by liganded growth factor receptors [14–16] may stimulate ras [17] and subsequently the rac and rho cascade because rac appears to be a downstream target of ras [18]. PKCs may function more distally in the adhesion process since some PKC isoforms have been shown to translocate to adhesion sites and to bind filamentous actin [19–21]. Furthermore, a yeast equivalent of mammalian PKCs has been shown to be an effector of rho [22], and several molecules which play a role in the membrane targeting of actin fibers such as vinculin [23] and the myristoyl– alanine-rich PKC substrate (MARCKS) [24] are excellent substrates for phosphorylation by PKCs in vitro. Finally, PKC inhibitors have been demonstrated to interfere with adhesion and migration in many cell types: endothelial, melanocytic, and sarcoma [25–30]. Keratinocytes in primary cultures show high phenotypic resemblance to the migrating keratinocytes in wounded skin. By contrast to keratinocytes in steady-state skin, in which only the basal cells in the multistratified epithelium attach to the basement membrane while upper layers differentiate gradually and lose their ability to attach to the extracellular matrix, keratinocytes in wounded skin overexpress integrin receptors and gain the ability to migrate actively on a provisional matrix made of clotted plasma adhesion substrates and interstitial collagens [31]. Similarly cultured keratinocytes show growth factordependent migration on many extracellular matrix proteins and in particular on type I collagen, an event which is mediated by interaction with a2b1 integrins [32, 33]. The aim of the present study was to deter2 Abbreviations used: FITC, fluorescein isothiocyanate; GF/ medium, keratinocyte serum-free medium with insulin and growth factor supplement; GF0 medium, keratinocyte serum-free medium without insulin and without growth factor supplement; HD-BSA, heat-denatured bovine serum albumin; IgG, immunoglobulin; PBS, phosphate-buffered saline; PKC, proteines kinase C; PMA, phorbol 12-myristate 13-acetate; SDS–PAGE, sodium dodecyl sulfate–polyacrylamide gel electrophoresis.

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mine the respective roles of PKC and tyrosine kinases in the molecular processes leading to adhesion, migration, and F-actin reorganization that take place when normal human keratinocytes interact with type I collagen. The study was conducted with pharmacological activators and inhibitors of PKC and tyrosine kinases. We find that the activation of PKC or the inhibition of tyrosine kinases interferes mainly with cell migration while the inhibition of PKC function blocks both migration and adhesion. Interestingly we also find that PKC activation stimulates tyrosine kinase-dependent stress fiber production and a reciprocal breakdown of cortical actin while PKC inhibitors block microfilament assembly. MATERIALS AND METHODS Reagents. Fatty acid-free BSA, phorbol 12-myristate 13-acetate (PMA), chelerythrine chloride, genistein, herbimycin A, calphostin C, type I collagen (rat tail), rhodamine–phalloidin, soybean trypsin inhibitor, nonimmune mouse IgG1 (MOPC), and biochemical reagents (ACS grade) were purchased from Sigma (La Verpille`re France). D-erythro-Sphingosine was obtained from France-Biochem (Meudon, France). Function-blocking antibodies to the integrin a2 subunit (clone Gia) and b1 (clone Lia 1/2), biotinylated or peroxidaseconjugated goat Fab*2 to mouse immunoglobulins, and fluorescein isothiocyanate–streptavidin were from Immunotech (Marseille, France). Keratinocyte serum-free medium (KSFM) and growth factor supplements (epidermal growth factor and bovine pituitary extracts) were obtained from Life Technologies (Cergy Pontoise, France). Monoclonal antibody to paxillin was purchased from Chemicon (Souffelweyersheim, France). Calcein-AM (Molecular Probes), BCA protein assay kit (Pierce), and monoclonal anti-phosphotyrosine antibodies (clone PY20) were obtained from Interchim (Montluc¸on, France). Cell cultures. Human keratinocytes were obtained by dispase and trypsin dissociation of human foreskin, followed by cultivation in KSFM supplemented with growth factors (GF/ medium). Cells were used at 60–80% confluency and between the second and the fourth subcultures. In some instances, cells were deprived of growth factors. For this purpose, cells grown at 60% confluency were washed twice in KSFM without insulin and growth factor supplement (GF0 medium) and incubated in this medium overnight. The same procedure was repeated 6 h before experiments were carried out. Cell treatments with pharmacological reagents. Pharmacological agents were dissolved in dimethyl sulfoxide (DMSO), and stock solutions were aliquoted and frozen at 0207C. Under all experimental conditions, including untreated cells, the final concentration of DMSO in assay medium was kept constant and equal to 0.5% (v/v). Cell treatment with tyrosine kinase inhibitors was performed in a culture dish for 18 h with a combination of genistein and herbimycin A, generally 300 and 1 mM, respectively. Other treatments were performed after cell detachment from culture dishes and resuspension in assay medium either for 30 min at 377C for PKC inhibitors (incubation with calphostin C was performed under fluorescent light for photoactivation) or immediately before plating on collagen for PMA. In some experiments, cells were plated for 2 h in GF/ or GF0 medium with or without tyrosine kinase inhibitors. The medium was then aspirated and replaced by the original medium with or without PMA and incubation was continued for another hour. Viability of the cells after drug treatment was examined by the trypan blue exclusion test and by 51Cr release and it was found in all cases to be better than 90%. Collagen coating of adhesion surfaces. Ninety-six-well plastic dishes used in adhesion assays were coated with 100 ng of type I

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collagen contained in 50 ml of 0.05 M Tris, ph 7.4, for 1 h at 377C and then blocked with heat-denaturated 1% BSA in PBS (HD-BSA). Eight-well glass chambers (Lab-Tek, Polylabo, Strasbourg, France) were used for indirect immunofluorescence experiments and cell spreading measurements. They were sequentially washed with 20% H2SO4 in water for 2 h, derivatized with 3-aminopropyltriethoxysilane for 4 min, activated with 0.25% glutaraldehyde in PBS for 30 min, incubated with collagen (20 mg/ml) in PBS overnight at 47C, and blocked with 0.1 M ethanolamine in PBS, pH 7.8, for 30 min and HD-BSA for 1 h, with each of these steps being intervened by rinses with PBS. Adhesion assays. Cells to be assayed were labeled with 10 mCi/ ml [35S]methionine (800 Ci/ml, Amersham) for 18 h. After detachment cells were resuspended in 0.25% KSFM HD-BSA. Cells (30 1 103) treated or untreated with pharmacological agents as described above were seeded in 100 ml of assay medium (0.25% KSFM HDBSA, w/v) in 96-well dishes coated with collagen. After centrifugation (800g for 1 min), cells were incubated at 377C in a cell culture incubator for 5, 15, 30, and 45 min. After two washes, attached cells were lysed with 0.1 ml of 1% SDS (w/v) in water, and radioactivity in the lysates was measured by beta scintillation counting. Specific attachment was estimated as counts in lysates on collagen minus counts in lysates on BSA divided by counts in a seeded cell and multiplied by 100. Quadruplicate determinations were performed for each experimental condition. Cell spreading evaluation. Cells used for cell spreading determination were rendered fluorescent by loading with 5 mg/ml calceinAM for 2 h before detachment, drug treatment, and plating in 8-well glass chambers coated with collagen followed by incubation at 377C in a cell culture incubator. At the given time, chambers were examined with an epifluorescence microscope equipped with a 101 magnifying lens and the resulting images were captured by a SIT camera and digitalized. For each experimental condition, the cell surface was measured in the images of five different fields chosen at random in each of triplicate chambers. The surface occupied by the cells in the image of one field was divided by the number of cells and multiplied by 100. Mean determination corresponded generally to the examination of about 300 cells. Migration assays. Migration quantification was determined by phagokinetic track assay as described in Ref. [34]. The method was adapted for experiments using 24-well culture plates. The gold salts suspension was poured onto 15-mm-diameter glass coverslips deposited at the bottom of the wells. The sedimented gold particles were coated overnight at 47C with 0.3 ml of 10 mg/ml collagen in PBS. After removal of the coating medium, cells were seeded into the wells at a density of 5 1 103 cells per milliliter of GF/ or GF0 0.25% KSFM HD-BSA with or without pharmacological agent. Cells were allowed to migrate for 4 h at 377C in a cell culture incubator and the reaction was stopped with 0.1 ml of formaldehyde. Wells were examined by light microscopy under 201 magnification and the images were captured and digitalized for quantification of the track surfaces left on the gold coat by migrating cells. For each experimental condition, track surface was measured in the images of five independent fields chosen at random in each of the triplicate wells. Image analysis. Images were digitalized with a PC Vision/ card (Imaging Technology, USA) yielding a 512 1 512 frame in 256 gray levels and processed with laboratory-written software which permitted the exploration of the gray-level distribution of the pixels making the image by using a color code as a reporter for progression along the gray scale. When the observed object, i.e., cells or tracks, was colorized, a gray-level threshold value was obtained. The surface of the object could be evaluated by calculating the number of pixels displaying a gray-level value above threshold. Immunofluorescence analysis of microfilaments and paxillin distribution. Cells were plated in eight-well glass slides coated with collagen in the indicated medium. After incubation for 4 h at 377C in a cell culture incubator, medium was aspirated and adherent cells

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were fixed with 3.7% formaldehyde for 15 min, permeabilized with 0.1% Triton X-100 for 2 min, and blocked with 0.1 M ethanolamine, pH 7.4, and 1% BSA. Cells were then either incubated with rhodamine–phalloidin (100 ng/ml) for 30 min or incubated sequentially with 8 mg/ml anti-paxillin monoclonal antibody or nonimmune mouse IgG1 for 2 h followed by 10 mg/ml biotinylated anti-mouse immunoglobulins for 30 min (antibodies were diluted in 5% fat-free dry milk), and then by 1 mg/ml fluorescein-conjugated streptavidin in 1% BSA, and finally with 1 mM biotin. PBS was used as the base of all solutions and intervening rinses, and incubations were performed at room temperature. After mounting in Vectashield (Vector Laboratories, Burlingame, CA) slides were examined under an epifluorescence microscope with appropriate excitation and emission filters under 401 or 201 magnification. Pictures of observed fields were recorded on Kodak TMS 400 ASA film. Immunoblotting of phosphotyrosine proteins. Cells treated as indicated were left to attach at 377C for 30 min on 35-mm plastic culture dishes coated with 10 mg/ml collagen or 50 mg/ml polylysine. After the incubation and after washing twice in PBS, attached cells were lysed in 0.5 ml of a freshly prepared boiling solution composed of 0.06 M Tris, pH 6.8, 1 mM EDTA, 1 mM EGTA, 5 mM benzamidine, 1 mM iodoacetamide, 20 mM sodium fluoride, 10 mM sodium orthovanadate, 1 mM phenylmethanesulfonyl fluoride, 10 mg/ml each of aprotinin, leupeptin, pepsatin, and a2-microglobulin, 20% glycerol, (v/v), and 4% SDS (w/v). After a brief sonication, lysates were heated at 607C for 5 min and centrifuged at 14,500g for 15 min and proteins were measured by microBCA assay. Lysates were analyzed by SDS– PAGE under reducing (20 mM dithiothreitol) conditions. After electrotransfer of gel content to nitrocellulose membranes (Hybond-C/, Amersham, les Ullys, France), according to the supplier’s instructions, membranes were blocked with 5% nonfat dry milk in 0.05 M Tris, 0.15 M NaCl, pH 7.4. Tyrosine-phosphorylated proteins were detected by incubating the blots with 1 mg/ml anti-phosphotyrosine monoclonal antibody (clone PY20) for 2 h at room temperature and then with 1 mg/ml horseradish peroxidase-conjugated anti-mouse immunoglobulins for 15 min followed by visualization with enhanced chemiluminescence (ECL System, Amersham) with Kodak X-OMAT AR film followed by signal quantification on chemiluminograms by image analysis with an Herolab densitometer (OSI, Maurepas, France).

RESULTS

Collagen-I-Mediated Adhesion, Spreading, and Migration of Keratinocytes: Modulation by Growth Factors, Tyrosine Kinase Inhibitors, and PKC Activators and Inhibitors Type I collagen is known to support attachment and growth factor-dependent migration of keratinocytes [32] that involve interaction with a2b1 integrins [33]. The respective roles of tyrosine kinases and PKC in the signaling pathways involved in these processes were evaluated by examining the effect of pharmacological modifiers of these enzymes on adhesion, spreading, and migration. Adhesion-dependent processes were most dramatically affected by inhibition of PKC function. Three highly specific PKC inhibitors were used: calphostin C and D-erythro-sphingosine, which interact with the protein’s regulatory domain by competing for diacylglycerol binding, and chelerythrine chloride, which interferes with the catalytic function of PKC. For exam-

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FIG. 1. Collagen-I-mediated adhesion, spreading, and migration: modulation by growth factors, tyrosine kinase inhibitors, and PKC inhibitors. Adhesion (A) and spreading (B) were studied under kinetic conditions. Cells labeled with [35S]methionine for adhesion assays or rendered fluorescent with calcein-AM for spreading evaluation, grown in the presence (open symbols) or in the absence of growth factors (closed symbols), untreated (h, j) or treated by 0.3 mM genistein and 1 mM herbimycin A (s, l) or 0.05 mM calphostin C (L, l), were seeded in 96-well culture dishes or in glass chambers coated with 10 mg/ml collagen and incubated at 377C. At the indicated time, adhesion wells were washed and radioactivity in the lysates of adherent cells was counted, while spreading wells were examined under an epifluorescence microscope and the cell surface was measured by computerized analysis of the digitalized images. Mean specific adhesion is expressed as a percentage of counts in input cells [mean of quadruplicate determinations with standard deviations (not shown) less than 10%] and spreading corresponds to the mean cell surface (arbitrary units) 1 100 measured in five fields taken at random in three different chambers where a total of at least 300 cells was counted. (C) Collagen-I-mediated migration of quiescent keratinocytes (0GF, open bar) or cells grown in presence of growth factors (stippled bars) and treated with 1 mM herbimycin A and 0.1 (G.1), 0.2 (G.2), or 0.3 mM (G.3) genistein or with 0.025 mM calphostin C (Calp) relative to the migration of untreated cells (control, Ctl) considered equal to 1. Migration was quantified by image analysis evaluation of the track area left by cells on colloidal gold surfaces. The mean { standard deviation of several independent experiments is shown. In all these experiments, the migration of treated cells was significantly different from that of control cells (Student’s t test, P õ 0.001) except for G.1 (P õ 0.07).

ple, cell treatment with 0.05 mM calphostin C resulted in a 90% blockade of migration, a profound decrease of spreading, and a marked reduction of adhesion (Figs. 1A–1C). Basal and growth factor-stimulated adhesion, spreading, and migration were affected by the compound (Figs. 1A–1C) and quite similar results were obtained with D-erythro-sphingosine and chelerythrine chloride (data not shown). The analysis of migration inhibition by the three drugs in dose–response experiments indicated that these dramatic effects occurred

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in a concentration range consistent with specific inhibition of PKC. Indeed, as may be seen in Fig. 2, halfmaximal inhibition of migration was obtained at a concentrations of 0.02 mM calphostin C, 1 mM chelerythrine, and 3 mM D-erythro-sphingosine, which were very close to the concentrations known to produce half-inhibition of PKC kinase activity in vitro. The activation of PKC by a nondegradable analog of diacylglycerols, the phorbol ester PMA, also resulted in migration impairment. However, by contrast with

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same in cells attached to polylysine and collagen) (Fig. 3A). The second group corresponded to proteins migrating in the 120- and the 80-kDa range, the phosphorylation of which was modulated by adhesion to collagen. The staining intensity of these proteins increased significantly after adhesion to collagen, and phosphorylation stimulation by collagen of these proteins was significant in GF0 cells although it was 40% lower than in GF/ cells (Fig. 3A). As can be seen in Fig. 3B, the phosphophorylation of all these proteins was inhibited 80% by the combination of genistein (0.3 mM) and herbimycin A (1 mM). Together these data suggest that the activation of PKC and inhibition of tyrosine kinases interfere mainly with migration while the adhesion process was also affected by PKC inhibition. We next examined the organization of microfilaments and focal complexes in cells attaching to collagen and their modulation by PKC and tyrosine kinase modifiers. FIG. 2. Dose–response inhibition of collagen-mediated migration by PKC inhibitors. Keratinocytes treated with calphostin C (h), D-erythro-sphingosine (j), and chelerythrine (l) at the indicated concentrations were incubated on colloidal gold coated with collagen for 4 h at 377C. Migration was quantified by computerized image analysis evaluation of the surface of the migration tracks left on gold and is shown expressed as a percentage of the migration of untreated cells. Mean migration { SD of triplicate determinations.

PKC inhibitors, the inhibition was only partial and a maximal 40% inhibition of migration was obtained with 100 ng/ml PMA (Fig. 1C, P õ 0.001); inhibition was not further increased for concentrations up to 1 mg/ ml (data not shown). Also in contrast to PKC inhibitor effects, spreading was increased by PMA both in GF0 and in GF/ cells while adhesion was little affected (data not shown). The role of tyrosine kinases was investigated by treating keratinocytes with two specific tyrosine kinase inhibitors, 1 mM herbimycin A associated with genistein [6, 35]. A dose-dependent blockade of migration was obtained when the genistein concentration was increased to 0.3 mM, reaching 60% inhibition (Fig. 1C, P õ 0.001). Under these conditions, adhesion and spreading were either slightly decreased in GF/ cells or increased in GF0 cells (Figs. 1A and 1B). Since the effect of tyrosine kinase inhibition appeared relatively modest, the efficacy of the drug combination in inhibiting tyrosine phosphorylation was evaluated by examining the binding of an anti-phosphotyrosine antibody to cell extracts separated by SDS–PAGE and electrotransferred to nitrocellulose. Two major groups of phosphorylated proteins were found. The first group corresponded to proteins migrating in the 180-kDa range, the phosphorylation of which depended on stimulation by growth factors (staining decreased by 80% in GF0 cells) but not on adhesion to collagen (staining was the

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Growth Factors Stimulate Cell Polarization, Stress Fiber Formation, and Ventral Translocation of Adhesion Complexes The organization of the F-actin cytoskeleton was analyzed 3 h after attachment on collagen by epifluorescence microscopy observation of rhodamine–phalloidin binding. In growth factor-starved cells (GF0 conditions), actin microfilaments were underlying the cell periphery in a juxtamembraneous pattern (Fig. 4A). In

FIG. 3. Adhesion and growth factor-dependent protein tyrosine phosphorylation in keratinocytes: modulation by tyrosine kinase inhibitors. Immunoblotting with anti-phosphotyrosine antibody PY20 of total keratinocyte proteins after separation by reducing SDS–6% polyacrylamide gel electrophoresis and electrotransfer to nitrocellulose. (A) Lysates (30 mg protein/lane) of keratinocytes which were grown in the presence of growth factors (lanes 1 and 2) or starved of growth factors (lanes 3 and 4) and incubated for 40 min at 377C in 35-mm culture dishes coated with polylysine (lanes 1 and 3) or with collagen (lanes 2 and 4). (B) Lysates (30 mg protein/lane) of keratinocytes which were grown in the presence of growth factors untreated (lane 1) or treated with 1 mM herbimycin A and 0.3 mM genistein (lane 2), plated in 35-mm culture dishes coated with collagen, and incubated for 40 min at 377C. Numbers on the left-hand side refer to the migration position of molecular weight standards. Band intensities were quantified by densitometry scanning of the chemiluminograms with a Herolab densitometer.

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FIG. 4. Microfilament organization in keratinocytes attaching to type-I collagen: modulation by growth factors, PMA, and tyrosine kinase inhibitors. Keratinocytes grown without (A) or with growth factors (B–F) during the 18 h preceding the experiment were plated in eight-well chambers coated with collagen and incubated for 3 h at 377C in medium without growth factor (A), with growth factors (B, D, E), or with growth factors and tyrosine kinase inhibitors (C, F): 1 mM herbimycin A and 0.3 mM genistein. (D, E, F) Cells were stimulated with PMA (100 ng/ml) either for 3 h (D) or for the last hour of incubation (E, F). Cells were then fixed, permeabilized, and incubated with rhodamine–phalloidin. Typical images taken at 401 magnification are shown.

the presence of growth factors (GF/ conditions), the actin network in most of the cells appeared polarized, displaying a cortical-like distribution on one side and, on the opposite side, a heavy thickening of the actin network made of cytoplasmic fibers, often running parallel to the membrane, resembling stress fibers (Fig. 4B). The distribution of paxillin, a cytoskeletal protein which is known to cluster within adhesion complexes, differed significantly depending on the presence of growth factors. Under GF0 conditions, paxillin appeared as thick patches mainly underlying the periphery of the cell (Fig. 5A). In GF/ conditions, paxillin was condensed as dense spots most frequently polarized to the cell aspect which contained stress fibers and which might correspond to the leading edge of the cell. In this region paxillin clustering was translocated more frequently to the ventral aspect of the cell than to the outer border (Fig. 5B). PKC Activation by Phorbol Esters Stimulates Stress Fiber Formation The examination of the actin cytoskeleton in cells treated with PMA showed that PKC activation resulted in a massive production of stress fibers which occurred both in GF0 (data not shown) and in GF/ cells (Fig. 4D). Under the latter conditions, stress fiber formation was associated with a loss of polarization of the actin network which could be observed in

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untreated cells. The stress fiber network induced by PMA seemed to be composed of a dense perinuclear mesh from which fibers irradiated to the cell periphery. In GF/ cells stimulated with PMA, paxillin was still clustered; however, clusters were found all around the cell margin and their ventral translocation was less frequently observed (Fig. 5D). The Stimulation of Stress Fiber Formation by Growth Factors or PMA Is Inhibited by Tyrosine Kinase Inhibitors Stress fiber formation in fibroblasts is believed to depend on the function of tyrosine kinases implicated upstream and downstream of the GTPase rho [8, 12]. Therefore, the involvement of tyrosine kinases in stress fiber stimulation by growth factors and in PKC activation was investigated by examining the microfilament organization in keratinocytes treated with the tyrosine kinase inhibitors genistein (0.3 mM) and herbimycin A (1 mM). As shown in Fig. 3B, this association allowed a substantial 80% inhibition of protein tyrosine phosphorylations in GF/ cells. Under these conditions, the structure of the actin cytoskeleton was profoundly modified (compare Fig. 4B and Fig. 4C). No stress fibers could be found any longer. Instead the phalloidin staining of the cytoplasm appeared blurry and dotty. By contrast, the cortical actin was maintained and appeared even hypertrophic. Phalloidin clusters were still

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FIG. 5. Paxillin distribution in keratinocytes attaching to type-I collagen: modulation by growth factors, PMA, and tyrosine kinase inhibitors. Keratinocytes grown without (A) or with growth factors (B–F) during the 18 h preceding the experiment were plated in eightwell chambers coated with collagen and incubated for 3 h at 377C in medium without growth factor (A), with growth factors (B, D, E), or with growth factors and tyrosine kinase inhibitors (C, F): 1 mM herbimycin A and 0.3 mM genistein. (D, E, F) Cells were stimulated with PMA (100 ng/ml) either for 3 h (D) or for the last hour of incubation (E, F). Cells were then fixed, permeabilized, and incubated with antipaxillin antibody revealed by double immunofluorescence. Typical images taken at 201 magnification are shown.

observed with tyrosine kinase inhibitor treatment. However, paxillin-containing adhesion complexes were restricted to the cell periphery, suggesting that ventral translocation was impaired (Fig. 5C). To investigate the effect of tyrosine kinase inhibitors on stress fiber stimulation by PMA, keratinocytes were plated on collagen for 2 h with or without tyrosine kinase inhibitors and stimulated with PMA for an additional 1 h. This PMA challenge was sufficient to induce a profuse stress fiber production in control cells (Fig. 4E). However, this stimulation was blocked in cells treated with tyrosine kinase inhibitors (Fig. 4F). Instead the cytoplasm appeared hollow, solely punctured by tiny actin aggregates. Interestingly, the PMA challenge in cells treated with tyrosine kinase inhibitors resulted in a significant thinning of the cortical actin network which vanished almost completely (compare Fig. 4C and Fig. 4F). This was associated with a disappearance of paxillin clusters and by an intensification of cytoplasmic paxillin labelings (compare Fig. 5C and Fig. 5F). PKC Inhibitors Block Actin Microfilament Buildup and Focal Adhesion Formation Thus overactivation of PKCs by phorbol esters resulted in a frank stimulation of stress fiber synthesis. To determine if, under basal conditions, these enzymes

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participated in adhesion-dependent reorganization of microfilaments, cells were treated with specific PKC inhibitors and microfilament organization was examined. As suspected from the profound alteration of adhesion, spreading, and migration caused by PKC inhibitors, the effect of these agents on cell shape and Factin organization was dramatic. As shown in Fig. 6, cells treated with 0.025 mM calphostin C remained rounded up and no specific actin microfilamentous structures or paxillin clusters could be distinguished. Quite similar figures were obtained after cell treatment with chelerythrine and D-erythro-sphingosine at 1 and 3 mM, respectively. DISCUSSION

In this study we show that the remodeling of the F-actin cytoskeleton which occurs when keratinocytes adhere to type I collagen is profoundly modified by pharmacological activators and inhibitors of protein kinase C. Activators stimulate stress fiber production and cortical actin breakdown while inhibitors block microfilament assembly, spreading, and migration and decrease adhesion efficacy. A detailed examination of the data suggests that PKC intervene at several levels in the molecular events which lead to adhesion and remodeling of the actin cytoskeleton.

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FIG. 6. PKC inhibitor calphostin C blocks microfilament assembly. Binding pattern of rhodamine–phalloidin (A) and anti-paxillin antibody (B) in keratinocytes treated with 0.05 mM calphostin C and plated on collagen for 4 h at 377C. Typical images taken at 401 magnification.

Two actin microfilament networks which differed on the basis of cellular distribution, regulation by growth factors, and modulation by tyrosine kinase inhibitors could be distinguished. A first network consisting of transcytoplasmic stress fibers was induced by growth factors and this induction was blocked by tyrosine kinase inhibitors. The second network consisted of fibers closely apposed to the cell outer border, resembling the cortical network described in other cell types. In contrast to stress fibers, this network was also formed in quiescent cells. This finding was surprising because current conceptions of microfilament buildup implicate the stimulation of GTPases by growth factors. Therefore, it is possible that cortical actin synthesis was stimulated by autocrine growth factors or alternatively by signals produced after integrin engagement. In any case, whatever the trigger, the molecular events leading to cortical actin synthesis did not involve tyrosine kinases. Indeed this network was especially obvious and well delineated in cells treated with tyrosine kinase inhibitors at a concentration which resulted in a blockade of stress fiber synthesis and in a considerable reduction of tyrosine phosphorylation of endogeneous proteins. The synthesis regulation of stress fibers and cortical actin by activated PKC occurred in an opposite manner. Indeed, an intense production of stress fibers was induced by PMA and this stimulation was blocked by tyrosine kinase inhibitors. Since stress fiber formation in fibroblasts implicates the stimulation of a tyrosine kinase downstream of rho [12], it is tempting to speculate that the tyrosine kinase-dependent stimulation of stress fibers by PKC operates through activation of rho. Although this hypothesis needs experimental verification, it would be in line with the finding that PMAdependent actin reorganization and adhesion stimulation in leukocytes do not take place when rho is inactivated [36, 37]. By contrast the formation of the cortical network was

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greatly diminished after PKC activation. This could result from a decrease in cytosolic amounts of actin monomers or some counterregulation related to the profuse stress fiber synthesis occurring under these conditions. However, these explanations are unlikely since the cortical actin was also completely wiped out by PMA when stress fiber synthesis was minimal, i.e., in cells treated with tyrosine kinase inhibitors. This suggests that the formation of the cortical network is negatively regulated by overactivated PKC. Thus phosphorylations by PKC may inhibit the system that drives cortical actin polymerization and/or inactivate membrane proteins which are important for membrane targeting of the cortical network. Prototypical proteins of that kind could be the recently discovered src- and ras-suppressed PKC substrate which appears concentrated in the cortical actin network [38] or the MARCKS which loses microfilament crosslinking activity and translocates from the plasma membrane to the cytosol and lysosomes upon phosphorylation [39]. The cortical network found in quiescent cells was associated with focal adhesion formation as indicated by a significant peripheral clustering of paxillin, an important adaptator protein involved in the targeting to focal complexes of vinculin, focal adhesion kinase, and additional components as well [5, 40]. Focal adhesion formation was maintained as long as microfilament buildup took place. This relationship was best exemplified in cells treated with tyrosine kinase inhibitors where paxillin clustering was still found associated with persisting cortical actin when, on the other hand, it vanished together with the cortical network when the same cells were challenged with PMA. These observations indicate that focal complexes may assemble even when the tyrosine kinase function is inhibited, provided that actin polymerization is unimpaired, thus bringing support to the idea that microfilament dynamics play a main role in focal adhesion formation. Together these data indicate that, in keratinocytes,

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activated PKC gear polymerization of stress fibers and reciprocal breakdown of the cortical actin network. PMA is known to stimulate microfilament remodeling in most cells. In fibroblasts, the drug also stimulates stress fiber synthesis [41]. By contrast, in other cells, the regulation of the actin cytoskeleton by activated PKC may be quite opposite that in keratinocytes. In BSC cells for example, PMA has been reported to stimulate the production of cortical ruffles while stress fibers are dismantled [42]. Interestingly, this dualism is also observed in cells treated with growth factors. Indeed EGF induces stress fiber breakdown and cortical ruffle production in A431 cells while it stimulates stress fiber assembly in fibroblasts [43, 44]. These considerations suggest strongly that the observed actin cytoskeleton remodeling triggered by PMA in keratinocytes is not a pharmacological artifact unrelated to physiology but that it reflects the implication of PKC in the molecular events involved in the regulation of microfilament dynamics by growth factors. Whether a differential expression of diacylglycerol-sensitive PKC participates in the opposite regulations of microfilament remodeling that PMA produces depending on the cell type remains to be determined. Phorbol esters are known to stimulate cell migration possibly by impinging on PKC involved in growth factor-signaling pathways. However, as was already noticed in another study [45], we find that PMA inhibits keratinocyte migration. The reasons for this difference are unclear. However, it could be that the dystrophic F-actin reorganization induced by activated PKC, i.e., an intense synthesis and a random positioning of stress fibers associated with cortical actin breakdown, eventually impairs a spatiotemporal regulation of microfilament dynamics that is essential for optimal migration. Microfilament synthesis and focal adhesion formation in quiescent and growth factor-treated keratinocytes were completely blocked by three different inhibitors, i.e., calphostin C and D-erythro-sphingosine, which prevent diacylglycerol-dependent activation of PKC, and chelerytrine chloride, which interferes with PKC catalytic function. The inhibition of actin polymerization was obviously associated with a dramatic decrease of cell spreading and migration and a significant alteration of adhesion efficacy and all these effects were obtained in a concentration range consistent with a specific inhibition of PKC. These results are consonant with similar observations obtained in several cell types and they raise the possibility that PKC play a decisive role in adhesion-dependent processes in keratinocytes as well. PKC could be implicated at several levels in adhesion regulation. First, they may be involved in a regulation of the function of adhesion receptors by stimulating or maintaining the high-affinity conformation of integrins as it occurs in leukocytes [2]. Second,

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since some PKC isoforms, e.g., a and u, cluster within adhesion sites [19, 46], it is possible that PKC also operate at a postreceptor level where they may participate in linking adhesion receptors to the actin cytoskeleton [47, 48]. Finally, PKC may be directly involved in regulation of actin cytoskeleton remodeling since some diacylglycerol-sensitive PKC isoforms which are expressed in keratinocytes, especially PKC1, have been shown to bind to F-actin or to translocate to microfilaments [20]. This paper presents strong indications that PKC participate in the growth factor regulation of the actin cytoskeleton and in the adhesion process in keratinocytes. The exact isoforms which participate in these regulations, the pathways used for their activation, and their relationship with the molecular events that are known to be of major importance in F-actin remodeling obviously remain to be deciphered. This work was supported by funds from the Association pour la Recherche sur le Cancer (ARC).

REFERENCES 1. Luna, E. J., and Hitt, A. L. (1992) Science 258, 955. 2. Hynes, R. O. (1992) Cell 69, 11. 3. Horwitz, A., Duggan, K., Buck, C., Beckerie, M., and Burridge, K. (1986) Nature 320, 531. 4. Otey, C. A., Pavalko, F. M., and Burridge, K. (1990) J. Cell Biol. 111, 721. 5. Clark, E. A., and Brugge, J. S. (1994) Science 268, 233. 6. Miyamoto, S., Teramoto, H., Coso, O. A., Gutkind, S. J., Burdelo, P. D., Akimaya, S. K., and Yamada, K. M. (1995) J. Cell Biol. 131, 791. 7. Roskelley, C. D., Srebrow, A., and Bissel, M. J. (1995) Curr. Opin. Cell Biol. 7, 736. 8. Nobes, C. D., Hawkins, P., Stephens, L., and Hall, A. (1995) J. Cell. Sci. 108, 225. 9. Nobes, C. D., and Hall, A. (1995) Cell 81, 53. 10. Burridge, K., Turner, C. E., and Romer, L. H. (1992) J. Cell Biol. 119, 893. 11. Crowley, E., and Horwitz, A. F. (1995) J. Cell Biol. 131, 525. 12. Ridley, R. J., and Hall, A. (1994) EMBO J. 13, 2600. 13. Dekker, L. V., and Parker, P. J. (1994) Trends Biochem. Sci. 19, 73. 14. Nishizuka, Y. (1992) Science 258, 607. 15. Liu, B., Wasiuddin, A. K., Hannun, Y. A., Timar, J., Taylor, J. D., Lundy, S., Butovich, I., and Honn, K. V. (1995) Proc. Natl. Acad. Sci. USA 92, 9323. 16. Divecha, N., and Irvine, R. F. (1995) Cell 80, 269. 17. Thomas, M. S., DeMarco, M., D’Archangelo, G., Halegoua, S., and Brugge, J. S. (1992) Cell 68, 1031. 18. Joneson, T., White, M. A., Wigler, M. H., and Bar-Sagi, D. (1996) Science 271, 810. 19. Jaken, S., Leach, K., and Klauck, T. (1989) J. Cell Biol. 109, 697. 20. Prekeris, R., Mayhew, M. W., Cooper, J. B., and Terrian, D. M. (1996) J. Cell. Biol. 132, 77.

eca

PKC REGULATION OF MICROFILAMENT NETWORK IN KERATINOCYTES 21. Blobe, G. C., Stribling, S. D., Fabbro, D., Stabel, S., and Hannun, Y. A. (1996) J. Biol. Chem. 271, 15823. 22. Kamada, Y., Qadota, H., Python, C. P., Anraku, Y., Ohya, Y., and Levin, D. E. (1996) J. Biol. Chem. 271, 9193. 23. Werth, D. K., Niedel, J. E., and Pastan, I. (1983) J. Biol. Chem. 258, 11423. 24. Aderem, A. (1992) Cell 71, 713. 25. Mascardo, R. N., Thopson, W., Gallo, M. A., Chambers, T. C., and Eilon, G. (1990) J. Cell. Physiol. 142, 401. 26. Friedlander, M., Brooks, P. C., Shaffer, R. W., Kincaid, C. M., Varner, J. A., and Cheresh, D. A. (1995) Science 270, 1500. 27. Klemke, R. L., Yebra, M., Bayna, E. M., and Cheresh, D. A. (1994) J. Cell Biol. 127, 859. 28. Chun, J.-S., Ha, M., and Jacobson, B. S. (1996) J. Biol. Chem. 271, 13008. 29. Vuori, K., and Ruoshlahti, E. (1993) J. Biol. Chem. 268, 21459. 30. Liu, B., Timar, J., Howlett, J., Diglio, C. A., and Honn, K. V. (1991) Cell Regul. 2, 1045. 31. Gailit, J., and Clark, R. A. F. (1994) Curr. Opin. Cell Biol. 6, 717. 32. Ando, Y., and Jensen, P. J. (1993) J. Invest. Dermatol. 100, 633. 33. Scharffetter-Kochanek, K., Klein, C., Heinen, G., Mauch, K., Schaefeer, T., Adelman-Grill, B., Goerz, G., Fusenig, N. E., Krieg, T. M., and Plewig, G. (1992) J. Invest. Dermatol. 98, 3. 34. Albrecht-Buehler, G. (1977) Cell 11, 395. 35. Miyamoto, S., Akiyama, S. K., and Yamada, K. M. (1995) Science 267, 883.

36. Laudanna, C., Campdell, J. J., and Butcher, E. C. (1996) Science 271, 981. 37. Tominaga, T., Sugie, K., Hirata, M., Morii, N., Fukata, J., Uchida, A., Imura, H., and Narumiya, S. (1993) J. Cell Biol. 120, 1529. 38. Lin, X., Tombler, E., Nelson, P. J., Ross, M., and Gelman, I. H. (1996) J. Biol. Chem. 271, 28430. 39. Allen, L.-A. H., and Aderem, A. (1995) EMBO J. 14, 1109. 40. Brown, M. C., Perrotta, J. A., and Turner, C. E. (1996) J. Cell Biol. 135, 1109. 41. Woods, A., and Coochman, J. R. (1992) J. Cell. Sci. 101, 277. 42. Schliwa, M., Nakamura, T., Porter, K. R., and Euteneuer, U. (1984) J. Cell Biol. 99, 1045. 43. Peppelenbosch, M. P., Tertoolen, L. G. J., Hage, W. J., and de Laat, S. W. (1993) Cell 74, 565. 44. Peppelenbosch, M. P., Qiu, R.-G., de Vries-Smits, A. M. M., Tertoolen, L. G. J., de Laast, S. W., McCormick, F., Hall, A., Symons, M. H., and Bos, J. L. (1995) Cell 81, 849. 45. Ando, Y., Lazarus, G. S., and Jensen, P. J. (1993) J. Cell. Physiol. 156, 487. 46. Monks, C. R. F., Kupfer, H., Tamir, I., Barlow, A., and Kupfer, A. (1997) Nature 385, 83. 47. Shaw, L. M., Messier, J. M., and Mercurio, A. M. (1990) J. Cell Biol. 110, 2167. 48. Lewis, J. M., Cheresh, D. A., and Schwartz, M. A. (1996) J. Cell Biol. 134, 1323.

Received April 1, 1997 Revised version received July 7, 1997

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