Histochemistry and Cell Biology

, Volume 121, Issue 5, pp 361–369

Actin microdomains on endothelial cells: association with CD44, ERM proteins, and signaling molecules during quiescence and wound healing

Authors

  • P. V. Jensen
    • Division of Cell Biology, Department of Anatomy and PhysiologyThe Royal Veterinary and Agricultural University
    • Division of Cell Biology, Department of Anatomy and PhysiologyThe Royal Veterinary and Agricultural University
Original Paper

DOI: 10.1007/s00418-004-0648-2

Cite this article as:
Jensen, P.V. & Larsson, L. Histochem Cell Biol (2004) 121: 361. doi:10.1007/s00418-004-0648-2

Abstract

During studies of the actin cytoskeleton in cultured endothelial cells we have observed that the luminal side of many cells contains F-actin microdomains that are rich in the hyaluronan receptor CD44 and in ezrin-radixin-moesin (ERM) proteins. A small subpopulation of the domains are also enriched in tyrosine phosphorylated proteins and signaling molecules. Confocal microscopy of rat aortic endothelial cells in situ demonstrated that similar microdomains occur in vivo. During healing of endothelial wounds, characteristic alterations of the actin cytoskeleton occurred. Thus, in many cells close to the wound, focal F-actin branching points appeared. The branching points were similar to the microdomains in that they colocalized with CD44 and ERM proteins, but, in addition, they formed centers for actin filament branching and were associated with phosphorylated protein kinase C α/βII. These colocalization data are consonant with the view that activated PKC is responsible for activating ERM-mediated crosslinking between CD44 and the actin cytoskeleton. Importantly, inhibition of PKC activity decreased staining for phosphorylated ERM proteins, decreased the frequency of F-actin branching points, and inhibited monolayer wound healing. Together, our data show that endothelial cells contain a novel actin cytoskeletal structure, the F-actin microdomain, and suggest that during wound healing such structures become associated with activated signaling molecules and thereby enhance actin cytoskeletal remodeling.

Keywords

Endothelial cellsWound healingActin cytoskeletonCD44ERM proteinsPKC

Introduction

Endothelial cells line the inside of blood vessels and sense changes in blood pressure and flow through hemodynamic forces in the form of shear stress and mechanical strain. Such forces act on the actin cytoskeleton and transduce signals regulating endothelial cell nitric oxide synthase expression as well as endothelial cell proliferation and function (Lehoux and Tedgui 2003; Li et al. 2002; Resnick et al. 2003). In addition, a host of soluble signals, including growth factors, cytokines, hormones, and neurotransmitters, affect endothelial cell function (Bernardini et al. 2003; Li et al. 2003; Rubanyi et al. 2002; Vincent et al. 2003; Walch et al. 2001). One of these functions, the formation of new vessels from preexisting vasculature through angiogenesis, is important to many disease states as well as to normal physiology and wound healing (Battegay 1995; Folkman 1995, 2000; Liotta et al. 1991). Angiogenesis depends upon remodeling of the actin cytoskeleton and endothelial cell motility, phenomena which are regulated by many factors including growth factors and chemokines, and interactions with other cells and with the extracellular matrix (Bernardini et al. 2003; Ingber et al. 1995; Li et al. 2003; Nguyen and D’Amore 2001; Uchida et al. 2000).

During studies of the endothelial actin cytoskeleton we observed that endothelial cells contained luminal F-actin microdomains, which were enriched in the hyaluronan receptor CD44. In addition, the microdomains contained activated (phosphorylated) ezrin-radixin-moesin (phospho-ERM) proteins, which have been shown to crosslink CD44 and other receptors to the actin cytoskeleton (Algrain et al. 1993; Bretscher et al. 2002; Legg and Isacke 1998; Tsukita et al. 1994; Yonemura et al. 1998). A subpopulation of preferentially larger F-actin microdomains were also enriched in signaling molecules and in tyrosine phosphorylated proteins. F-actin microdomains appear not to have been described before and could conceivably act to integrate mechanical and soluble signals during processes like angiogenesis and wound healing. During wound healing of endothelial cell monolayers we demonstrate that many cells close to the wound respond rapidly by forming F-actin branching points enriched in CD44, phospho-ERM proteins and phosphorylated protein kinase C α/βII (phospho-PKC α/βII). The F-actin branching points were similar to F-actin microdomains but, in addition, contained phospho-PKC α/βII and showed a radiating pattern of actin fibers that connected them with the rest of the actin cytoskeleton. These colocalization data suggested that activated PKC was responsible for activating ERM-mediated crosslinking between CD44 and the actin cytoskeleton. In support of this view, inhibition of PKC activity decreased staining for phosphorylated ERM proteins, reduced the frequency of F-actin branching points, and inhibited wound healing. These data are the first to examine the localization of these proteins during wound healing and the inhibitor results together with the colocalization of CD44, phosphorylated ERM proteins, and PKC form strong reasons to propose a functional interaction between these proteins during wound healing.

Materials and methods

Cell culture

Human umbilical vein endothelial cells (HUVECs; ATCC, Manassas, VA) were grown in Kaighn’s F-12 K medium supplemented with 10% fetal calf serum, 100 U/ml penicillin, 100 μg/ml streptomycin, 2 mM l-glutamine (Gibco, Grand Island, NY), and 0.4% endothelial cell growth supplement/heparin (PromoCell, Heidelberg, Germany) at 37°C in a humidified 5% CO2 atmosphere. HUVECs from passages 19–25 were used for all experiments. For studies of intact endothelial cell monolayer cultures, HUVECs were trypsinized, seeded on sterile glass slides (800,000 cells/ml), grown overnight or to confluence, and fixed. Additionally, confluent monolayers were longitudinally wounded using a pipette tip. GF-109203X (bisindolylmaleimide I HCl; Alexis Biochemicals, San Diego, CA) dissolved in dimethylsulfoxide (DMSO; Sigma-Aldrich, St. Louis, MO; final concentration 0.1%) was added to the media to a final concentration of 3 μM or 10 μM. Controls were treated with 0.1% DMSO. Cells were preincubated with inhibitor or vehicle for 1 h prior to wounding and fixed 20 or 60 min after wounding. For studies of effects on wound healing, cultures were fixed 24 or 48 h after wounding. Fixation was in equal parts of methanol/acetone (Baker, Deventer, The Netherlands) or in 3.7% paraformaldehyde (Merck, Darmstadt, Germany). The latter was followed by permeabilization in 1% Triton X-100 (Merck).

Non-cultured endothelial cells

Two adult male Wistar rats received a lethal dose of sodium pentobarbital intraperitoneally and were intracardially perfused with 60 ml of 0.9% sodium chloride followed by 60 ml of 3.7% paraformaldehyde. Specimens from the aorta were postfixed in the same fixative and permeabilized in 1% Triton X-100. Before staining, part of the muscle wall was stripped away with a pair of forceps.

Immunofluorescence and phalloidin staining

To visualize F-actin, cells were stained with Alexa 594-labeled phalloidin (Molecular Probes, Eugene, OR). Primary antibodies included mouse monoclonal antibodies to CD44 (Dako, Glostrup, Denmark), ezrin, phosphatidylinositol 3-kinase (PI3 K), protein tyrosine phosphatase 1B (PTP1B) (Transduction Laboratories, Lexington, KY), phosphotyrosine, insulin receptor substrate 1 (IRS-1), gastrin (Upstate Biotechnology, Lake Placid, NY), VEGF receptor 1 or 2 (Sigma-Aldrich), and affinity-purified rabbit antibodies specific for phosphorylated ezrin (Thr567)/radixin (Thr564)/moesin (Thr558) (phospho-ERM) or phosphorylated protein kinase C α/βII (Thr638/641) (phospho-PKC α/βII) (Cell Signaling Technology, Beverly, MA). The site of antigen–antibody reaction was revealed with goat anti-mouse Ig or goat anti-rabbit Ig antibodies labeled with Alexa 488 or Alexa 594 (Molecular Probes). Controls included use of type-matched monoclonal antibodies, immunoblotting (vide infra), as well as conventional staining controls (Larsson 1988). Double labeling employed primary antibodies raised in different species combined with species-specific second antibodies labeled with contrasting fluorochromes. Analysis of the effects of different treatments on actin and protein immunoreactivities was assessed in blind-coded specimens. Specimens were analyzed either by conventional fluorescence microscopy or in a Molecular Dynamics confocal laser scanning microscope (MultiProbe 2001). All experiments were repeated three to five times. In each experiment, a minimum of two slides were stained with each primary antibody.

Immunoblotting

Cells were lysed in 50 mM HEPES buffer, pH 7.6 (Sigma-Aldrich), containing 100 mM NaCl, 10 mM EDTA, 1% Triton X-100 (Merck), 4 mM sodium pyrophosphate (Sigma-Aldrich), 2 mM sodium orthovanadate, 10 mM NaF (Acros, Geel, Belgium), and a Complete tablet of protease inhibitors (Boehringer Mannheim, Hoersholm, Denmark). Aliquots were removed for protein determination and samples were diluted with 4×NuPage sample buffer and NuPage sample reducing agent was added to a final concentration of 10%. Samples were heated to 70°C for 10 min. Lysates were separated on NuPage 10% BisTris gel using NuPage MOPS SDS running buffer in an EI9001-Xcell II Mini cell apparatus. SeeBlue Plus2 prestained protein standard (Novex, San Diego, CA) was used as marker (191–14 kDa). Electroblotting was performed in a semidry blotting apparatus as recommended by the manufacturer (KemEnTec, Copenhagen, Denmark) using 0.2 μm PVDF immunoblot membranes (BioRad, Hercules, CA).

Immunoblots were stained with antibodies detecting CD44, ezrin, phospho-ERM, and phospho-PKC α/βII. Western Breeze chemiluminescence kit (Novex) and Hyperfilm ECL RPN2103 K (Amersham, Buckinghamshire, UK) were used for detection.

Image analysis

Quantitation of monolayer wound healing

Digital images of coded specimens stained with Giemsa (Merck) were acquired using an Olympus BH-2 microscope equipped with a monochrome Sony CCD camera connected to a PC with ImageProPlus software. Images of the wound area were captured through a 10× objective. A rectangle, 820 μm wide and 1,050 μm long, was placed symmetrically over the wound (570.9±37.0 μm in width). All cells within the rectangle, except those touching the lower or left border, were counted. For each slide, three separate areas were counted. Results were presented as means ± standard deviation (SD) and statistics were computed using the Mann-Whitney U-test. Experiments were repeated three times; in the two initial experiments three control and three experimental slides were analyzed, whereas in the final experiment seven slides of each kind were analyzed.

Quantitation of F-actin branching points

Specimens were stained with Alexa 594-labeled phalloidin, coded, and the number of branching points in 100 cells per slide was counted. Only cells bordering the wound were included. Five slides treated with GF-109203X and five slides treated with vehicle only were analyzed. Results were presented as means ± SD and statistics were computed using the Mann-Whitney U-test. In both types of experiments the level of significance was set at 0.05.

Results

Staining of HUVECs for F-actin using fluorescent phalloidin showed that the cells contained actin stress fibers and subcortical actin filaments (Fig. 1A). In addition, many of the cells contained characteristic F-actin microdomains (Fig. 1A). The size of the microdomains varied between 0.7 and 7.5 μm in diameter (mean 3.6±1.9 μm, median 3.5 μm). The vast majority of F-actin microdomains were classified as small (i.e., below 5 μm in diameter). By focusing and by confocal microscopy it was evident that the microdomains occurred on the luminal side of the cells.
Fig. 1A–N

F-actin microdomains of cultured human umbilical vein endothelial cells (HUVECs) contain receptors and signaling molecules. A, B Double staining for F-actin using Alexa 594-labeled phalloidin (A; red) and a type-matched monoclonal control antibody to gastrin (B; green). Note in A the presence of several F-actin microdomains (exemplified by arrows) as well as subcortical bands of actin filaments. Note in B that the control antibody fails to stain the cells. C, D Double staining for F-actin (C; red) and for CD44 (D; green) using indirect immunofluorescence. Note that two F-actin microdomains show staining for CD44 (arrows) and that staining for CD44 also occurs outside the microdomains. E, F Double staining for CD44 (E; red) and phosphorylated ezrin-radixin-moesin (phospho-ERM; F; green). Note that several, but not all, CD44-positive areas also react for phospho-ERM (arrows). G, H Double staining for F-actin (G; red) and phospho-ERM (H; green). Note that the majority of F-actin microdomains stain for phospho-ERM (arrows). Thus, the subpopulation of CD44-positive areas that stained for phospho-ERM in E, F correspond to F-actin microdomains. I, J Double-staining for F-actin (I; red) and phosphorylated protein kinase C α/βII (phospho-PKC α/βII; J; green). Note that a subpopulation of large microdomains stain for PKC α/βII (arrows), which also stains nuclei. K, L Double staining for F-actin (K; red) and tyrosine phosphorylated proteins (L; green). Note staining of two large microdomains (arrows) for tyrosine phosphorylated proteins. In addition, staining for tyrosine phosphorylated proteins is prominent over focal adhesions. M, N Double staining for F-actin (M; red) and IRS-1 (N; green). Note that several large microdomains (exemplified by arrows) stain for insulin receptor substrate 1. Scale bar 25 μm

Certain transmembrane proteins, including the hyaluronan receptor CD44, are known to be connected to the actin cytoskeleton via bridging ERM proteins (Algrain et al. 1993; Bretscher et al. 2002; Legg and Isacke 1998; Tsukita et al. 1994; Yonemura et al. 1998). We therefore next investigated whether the F-actin microdomains also contained CD44. Double staining using fluorescent phalloidin and a monoclonal antibody to CD44 showed that CD44 immunoreactivity was associated with over 90% of the F-actin microdomains (Fig. 1C, D). In addition, staining for CD44 was frequently observed in areas outside the microdomains (Fig. 1C, D). All staining controls were negative and substitution of the CD44 antibody with a subtype-matched antibody to human gastrin, which is not produced by endothelial cells, revealed no staining (Fig. 1B). Specificity of the antibody was further documented by immunoblotting studies, which showed that the CD44 antibody only reacted with a band of the size expected from CD44 (Fig. 2).
Fig. 2

Western blots of extracts from HUVEC monolayers. Note that staining occurs over bands of the size expected for CD44 (85–90 kDa), ezrin/phospho-ERM (80 kDa), and PKC α (82 kDa)

ERM proteins need to unfold in order to crosslink CD44 to the actin cytoskeleton, and this process is catalyzed by phosphorylation on certain critical threonine residues (Thr567 of ezrin, Thr564 of radixin, Thr558 of moesin; Gautreau et al. 2000; Matsui et al. 1998; Nakamura et al. 1995). We therefore double stained HUVECs with the monoclonal CD44 antibody and an affinity-purified rabbit antibody specific for ERM proteins phosphorylated on such threonine residues (phospho-ERM) (Fig. 1E, F). The majority, but not all, CD44-positive areas stained for phospho-ERM. Since, as noted above, CD44 staining also occurred outside F-actin microdomains, we next double stained monolayers with fluorescent phalloidin and the phospho-ERM antibody in order to determine whether the phospho-ERM-immunoreactive areas corresponded to F-actin microdomains (Fig. 1G, H). By this approach we demonstrated that the vast majority of the F-actin microdomains also were positive for phospho-ERM. Thus, by inference, we may conclude that the vast majority of F-actin microdomains were immunopositive for both CD44 and phospho-ERM. In addition, double staining with fluorescent phalloidin and a monoclonal antibody detecting ezrin showed that ezrin-immunoreactive proteins were associated with most of the F-actin microdomains (data not shown). All controls were negative and immunoblotting studies using the monoclonal ezrin antibody revealed a single immunoreactive band (Fig. 2) corresponding in size to that of ezrin. A similarly sized, but much fainter, band was revealed by the phospho-ERM antibody (Fig. 2).

Together, these data show that HUVECs grown in monolayers contain luminal F-actin microdomains, which are enriched in CD44, possibly by a mechanism involving phospho-ERM-mediated crosslinking between F-actin and CD44.

We next examined the cells for the presence of additional receptors. Immunostaining for VEGF receptor types 1 or 2 revealed that these virtually never occurred clustered in dorsal F-actin microdomains, but occurred diffusely distributed in the cell membrane (data not shown). All staining controls were negative and a subtype-matched irrelevant antibody failed to stain the cells.

PKC isoforms α and θ are known to phosphorylate ERM proteins on the critical threonine residues needed for unfolding (Gautreau et al. 2000; Matsui et al. 1998; Nakamura et al. 1995; Ng et al. 2001; Pietromonaco et al. 1998; Simons et al. 1998) and, in addition, ligands binding to CD44 have been found to activate PKC signaling (Slevin et al. 2002). We therefore examined the actin microdomains for their association with PKC. An antibody to phospho-PKC α/βII preferentially stained some of the larger (>5 μm) F-actin microdomains and, additionally, stained the vast majority of nuclei (Fig. 1I, J). In contrast, little or no staining of the microdomains was obtained with antisera to PI3 K or PTP1B (data not shown). Again, all staining controls were negative and subtype-matched irrelevant antibodies failed to stain the cells. Immunoblotting studies with the phospho-PKC α/βII antibody revealed an immunoreactive band of the size expected for PKC α (Fig. 2).

Thus, these data show that predominantly some of the larger F-actin microdomains contain phosphorylated PKC α/βII and the immunoblotting results indicate that the isoform detected is PKC α. The involvement of a subpopulation of the F-actin microdomains in active signaling was also underlined by the fact that a subpopulation of preferentially larger domains also stained for phosphotyrosine (Fig. 1K, L) as well as for the signaling intermediate IRS-1 (Fig. 1M, N).

We next examined freshly fixed specimens from rat aorta to determine whether F-actin microdomains were also present on non-cultured endothelial cells. Following dissection of part of the muscle wall, whole-mount specimens were stained and examined by confocal microscopy. A fair number of aortic endothelial cells were found to possess luminal F-actin microdomains that showed staining for CD44 and for phospho-ERM proteins (Fig. 3A–D). Additionally, a subpopulation of predominantly larger domains also stained for phospho-PKC α/βII (data not shown). Staining for other signaling entities was not tested on the aortic specimens. All controls were negative.
Fig. 3A–D

Confocal microscopy of rat aorta double stained for F-actin (A, C; red) and phospho-ERM (B, D; green). The images have been taken at the luminal side (A, B) and 2 μm deeper in the endothelial cell layer (C, D). Note the presence of a luminal, large F-actin microdomain that is immunopositive for phospho-ERM (arrows). Scale bar 15 μm

Staining of wounded HUVEC monolayers with fluorescent phalloidin revealed that areas distant from the wound showed a staining pattern very similar to that of unwounded monolayers. Thus, such cells contained actin stress fibers and subcortical actin filaments. Additionally, many cells contained dorsal F-actin microdomains of various sizes (Fig. 4A). In areas close to the wound (usually within 1–3 cell perimeters), a strikingly different picture emerged. Thus, already 20 min after wounding, the actin cytoskeleton became rearranged and stress fibers disappeared. In many of the cells, focal F-actin branching points, from which actin fibers radiated out, appeared (Fig. 4B). In the following, we will refer to these structures as F-actin branching points in order to distinguish them from the F-actin microdomains detected in confluent, unwounded monolayer cells, which rarely showed connections to the remaining actin cytoskeleton. The F-actin branching points were virtually never detected in unwounded areas or in intact monolayers. In monolayer cultures fixed 60 min after wounding, only a few cells at the wound edge exhibited F-actin branching points.
Fig. 4A–Q

Response of HUVEC monolayers to wounding. A Cell from unwounded monolayer stained with Alexa 594-labeled phalloidin for detecting F-actin. Note staining of subcortical actin filaments and stress fibers as well as of several F-actin microdomains (arrows inserted for orientation). B Phalloidin staining of a cell from the wound edge 20 min after wounding. Note marked reorganization of the actin cytoskeleton and the appearance of focal F-actin branching points (arrows). C–E Wounded monolayer, double stained for CD44 (C; red) and phospho-ERM (D; green, merged image in E). Note colocalization of CD44 and phospho-ERM both at a membrane ruffle (arrowhead) and at spots (arrows). F–H Wounded monolayer double stained for F-actin (F; red) and for phospho-PKC α/βII (G; green, merged image in H). Note staining of F-actin branching points for PKC α/βII (arrows). I–N Wounded monolayers treated either with vehicle only (I–K) or with the PKC inhibitor GF-109203X (L–N) and stained for F-actin (I, L; red) and phospho-ERM (J, M; green, merged images in K, N). Note, in I–K, that phospho-ERM-positive spots colocalize with F-actin branching points (exemplified by arrows) and, in M, that GF-109203X inhibits staining for phospho-ERM. O–Q Wounded monolayer stained for F-actin (O; red) and PTP1B (P; green, merged image in Q). Note that PTP1B is not localized at the F-actin branching points (exemplified by arrows), but occurs in the perinuclear area. Scale bars 10 μm (A, B); 20 μm (C–Q)

The majority of F-actin branching points also demonstrated staining for phospho-ERM proteins (Fig. 4I–K). Additionally, the phospho-ERM-positive branching points also stained for CD44 (Fig. 4C–E) and some of the cells exhibited membrane ruffles that were immunopositive for CD44 and phospho-ERM (Fig. 4C–E). Antibodies recognizing other cellular proteins like PTP1B reacted with other cellular areas, but not with the F-actin branching points, attesting to the specificity of the immune staining (Fig. 4O–Q). Antibodies detecting phospho-PKC α/βII reacted with all F-actin branching points (Fig. 4F–H). Thus, these results demonstrate that phospho-PKC α/βII associates with the focal F-actin branching points of wounded monolayers.

PKCs have been strongly implicated in endothelial cell proliferation, migration, and in repair of endothelial cell monolayer wounds (Daviet et al. 1989; Doctrow and Folkman 1987; Kent et al. 1995; Presta et al. 1991; Yamamura et al. 1996). Thus, both PKC α (human endothelial cells; Wang et al. 2002) and θ (rat capillary endothelial cells; Tang et al. 1997) have been shown to be involved in wound healing. Interestingly, precisely these two isoforms of PKC phosphorylate ERM proteins at the critical threonine residues needed for unfolding (Ng et al. 2001; Pietromonaco et al. 1998; Simons et al. 1998). We therefore examined the effects of a specific PKC inhibitor (GF-109203X) on wound healing and phospho-ERM staining of endothelial cell monolayers. At a concentration of 3 μM, GF-109203X showed a small, insignificant (P=0.2) inhibitory effect on wound healing. Moreover, no detectable effect of this dose of inhibitor was observed with respect to phospho-ERM staining. In contrast, a higher concentration of inhibitor (10 μM) markedly inhibited wound healing (Fig. 5) and also eliminated staining for phospho-ERM at the wound edge (Fig. 4L–N). Thus, the dose of GF-109203X required for significant inhibition of wound healing was also needed to inhibit staining of phospho-ERM proteins. Finally, analysis of coded specimens revealed that inhibitor treatment (10 μM) also significantly reduced the number of F-actin branching points detected 20 min after wounding (from 16.20±2.70 to 8.60±4.80; P<0.05).
Fig. 5

Diagram showing the effect of GF-109203X or vehicle (DMSO) on the healing of endothelial cell monolayers. Results from three separate experiments (experiment 1: n=3, 24 h; experiment 2: n=3, 48 h; experiment 3: n=7, 48 h) are presented as means ± SD of the number of cells present in a defined wound area after 24 or 48 h. Note that the PKC inhibitor GF-109203X markedly inhibits wound healing. One star denotes P=0.05; three stars denote P<0.001

Discussion

Our results show that cultured, untransformed HUVECs contain F-actin microdomains that are selectively enriched in certain types of receptors. Since cultured endothelial cells gradually loose their characteristics during passaging, we additionally studied rat aortic endothelial cells in situ and documented that also these cells contained luminally oriented actin microdomains. The presence of actin microdomains seems not to have been noticed previously. The striking colocalization observed between F-actin, CD44, and phospho-ERM proteins in the microdomains may reflect the known interactions between these proteins (Algrain et al. 1993; Bretscher et al. 2002; Legg and Isacke 1998; Tsukita et al. 1994; Yonemura et al. 1998). Thus, ERM proteins can exist both in a folded, inactive form and in an unfolded, active form. In the unfolded form they are capable of crosslinking certain transmembrane proteins, like CD44, to the actin cytoskeleton (Gautreau et al. 2000; Matsui et al. 1998; Nakamura et al. 1995). Importantly, unfolding of ERM proteins requires phosphorylation on critical threonine residues (Thr567 of ezrin, Thr564 of radixin, Thr558 of moesin) and such phosphorylation may be catalyzed by PKC α or θ (Gautreau et al. 2000; Matsui et al. 1998; Nakamura et al. 1995; Ng et al. 2001; Pietromonaco et al. 1998; Simons et al. 1998) or by Rho kinase (Matsui et al. 1998). It is therefore interesting to note that staining for phospho-PKC α/βII was associated with a subpopulation of F-actin microdomains. Moreover, immunoblotting studies with this antiserum revealed a single band migrating at a size corresponding to that of PKC α. Most of the PKC-immunopositive domains were of large size and similar such microdomains also stained for tyrosine phosphorylated proteins and IRS-1, which have been implicated in cell signaling events (reviewed by Hunter 1998). Together these results argue that a small subpopulation of large microdomains were associated with signaling events. This indicates that the latter microdomains are activated, while the majority of small microdomains only contain clustered, unactivated receptors. Importantly, receptor clustering may enhance sensitivity to soluble ligands by virtue of the close proximity of receptors and signaling adaptor molecules (Bray et al. 1998; Kim et al. 2002) and the association with actin microdomains may enhance effects of signaling on the actin cytoskeleton.

Our data also show that wounding of endothelial cell monolayers brings about a rapid rearrangement of the F-actin cytoskeleton in cells close to the wound edge. This rearrangement is characterized by the gradual rounding up of many of the cells resulting in a more motile phenotype, characterized by numerous membrane ruffles, and in the establishment of characteristic F-actin branching points. Importantly, the focal F-actin branching points stained both for the hyaluronan receptor CD44 and for ERM proteins. The F-actin branching points are similar to microdomains but are associated with branching actin filaments and with phospho-PKC α/βII. These data suggest that F-actin microdomains of cells close to the wound edge become associated with phosphorylated PKC isoforms that activate ERM-mediated crosslinking between CD44 and F-actin. This, in turn, results in increased numbers of connections between the microdomains and the rest of the actin cytoskeleton in the form of F-actin branching points.

Conventional and novel PKC isoforms require phosphorylations on specific threonine or serine residues in order to become activated by diacylglycerol (Liu and Heckman 1998). We therefore used antibodies recognizing PKC α/βII phosphorylated on specific threonine residues. Importantly, this does not necessarily imply that only active PKC α/βII was detected, but shows that molecules that fulfill one requirement for activation were detected. Direct support for a role of PKC in local phosphorylation of ERM proteins and activation of crosslinking between CD44 and F-actin came from use of a PKC inhibitor, which markedly reduced staining for phospho-ERM and reduced the number of F-actin branching points. The inhibitor (GF-109203X) has been reported to be a specific PKC inhibitor and is commonly used in concentrations from 1 to 10 μM, typically with an IC50 in the range of 1–5 μM (Habib et al. 1997; Higaki et al. 1999; Toullec et al. 1991; Villard et al. 1998). This concentration range has been found to be needed in studies of intact cells although studies of purified enzymes show that, in the test tube, the IC50 is considerably lower (Toullec et al. 1991). In agreement with previous studies (Tang et al. 1997; Wang et al. 2002) we found that inhibition of PKC activity markedly inhibited wound healing. However, the dose needed to demonstrate this effect was in the upper range of concentrations commonly used. This could reflect either that the cells studied were relatively poor in taking up the drug or that they metabolized it quickly. Alternatively, it could reflect that the drug also inhibited other enzymatic activities. Whatever the explanation is, it is important to note that the concentrations that were needed for significant inhibition of wound healing were also needed for inhibition of staining for phospho-ERM proteins.

In conclusion, our studies demonstrate a novel actin cytoskeletal structure, the F-actin microdomain. In quiescent cells in vitro and in situ these microdomains associate with CD44 and ERM proteins, but are for the most part not associated with signaling molecules. However, important rearrangements of the localization of the actin cytoskeleton, phospho-ERM proteins, and the hyaluronan receptor CD44 to focal F-actin branching points occur in response to monolayer wounding. To the best of our knowledge this is the first description of such a rearrangement. The colocalization of the F-actin branching points with phospho-PKC α/βII in combination with the inhibitor studies suggest that PKC isoforms are important both for monolayer wound healing, phosphorylation of ERM proteins, and for the very formation of focal F-actin branching points. The morphology of the F-actin branching points was very similar to that observed for F-actin microdomains in HUVEC monolayers. However, the latter did not present extensive connections to the remaining actin cytoskeleton, nor were most of them associated with staining for phospho-PKC α/βII. We therefore suggest that F-actin microdomains on quiescent cells become activated in response to wound healing and that activated PKC α/βII (and possibly additional kinases) serves to induce activation of ERM proteins, so that extensive crosslinks between CD44 and the actin cytoskeleton are formed, resulting in the formation of focal F-actin branching points. This represents an early response to wounding that may be important to the rearrangement of the actin cytoskeleton and, hence, to wound healing per se.

Acknowledgements

Grant support was from the Danish Medical Research Council, the Danish Veterinary and Agricultural Research Council, and the Danish Cancer Society. P.V. Jensen was supported by a PhD scholarship from the Royal Veterinary and Agricultural University. We thank Mrs. B. Traasdahl for performing image analyses.

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© Springer-Verlag 2004