A possible role for membrane depolarization in epithelial wound healing

Silvia Chifflet,1 Julio A. Hernández,2 and Silvina Grasso1

1Departamento de Bioquímica, Facultad de Medicina, and 2Sección Biofísica, Facultad de Ciencias, Universidad de la República, Montevideo, Uruguay

Submitted 27 May 2004 ; accepted in final form 17 January 2005


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Linear narrow wounds produced on cultured bovine corneal endothelial monolayers heal by actin cable formation at the wound border and lamellar crawling of cells into the injured area. We report the novel finding that membrane potential depolarization occurs at the leading edge of wounds and gradually extends inward toward the neighboring cells. We have determined that the replacement of extracellular Na+ by choline and the incorporation of phenamil, an inhibitor of the epithelial Na+ channel (ENaC), provoke a decrease in the actin cable and depolarization areas and in the lamellar activity of the wound edges. To the contrary, extracellular Li+ can successfully replace Na+ in the determination of the depolarization and cytoskeletal responses. This finding supports the idea that membrane depolarization, not the increase in intracellular Na+ concentration, is responsible for the formation of the actin cable, a result that is in agreement with previous evidence showing that nonspecific depolarization of the plasma membrane potential (PMP) of epithelial cells may promote characteristic cytoskeletal rearrangements per se (Chifflet S, Hernández JA, Grasso S, and Cirillo A. Exp Cell Res 282: 1–13, 2003). We suggest that spontaneous depolarization of the PMP of the cells at the wound borders determined by a rise in the ENaC activity of these cells constitutes an additional factor in the intermediate cellular processes leading to wound healing in some epithelia.

actin; epithelial sodium channel


THE MAINTENANCE OF THE STRUCTURAL integrity of epithelia is crucial to ensuring the proper functioning of this type of tissue. Epithelial layers respond to accidental or experimental injuries by a complex process of tissue restitution that involves mainly individual or sheet-coordinated cell migration into the denuded area. Two basic mechanisms of wound healing have been described (for reviews, see Refs. 26 and 62). One of them involves the lamellipodial crawling of cells from the wound edge toward the injured area by migration of separated or connected cells (36). In the other mechanism, the cells at the leading edge develop an actin cable at the wound margin that links the cells via insertions at the adherens junctions (26). The coordinated association of the actin cable to other components of the cytoskeleton, such as myosin II and tropomyosin (3), provides this structure with characteristic contractile properties. In the case of circular injuries, the actin belt tightens to close the wound in a manner resembling a purse-string closure. Characteristically, the actin cable mechanism appears to be dominant for the case of wound healing in embryonic epithelia, while injury repair by individual cell migration operates in most adult tissues (17, 26). However, overlapping of the two mechanisms is frequently observed in in situ preparations of some epithelia and also under culture conditions (17, 35). The predominant mechanism of healing might depend on the geometric characteristics of the wound, such as its form, width, and length. Thus, whereas small circular wounds use the purse-string mechanism, large, wide wounds may heal predominantly via lamellipodial crawling of the edge cells (3) and increased proliferation of migrating cells (47, 53).

Numerous factors appear to participate in the initiation and development of the healing process. Although the interruption of intercellular contacts may, in principle, play a role in triggering the repair response, a current view is that growth factor liberation by the injured cells specifically activates signaling pathways of the surviving neighboring cells (27, 51, 62). Also, an early Ca2+ wave of short duration, propagated from the leading edge toward the center of the monolayer, has repeatedly been detected immediately after experimental wounds (27, 51, 55). The activation of several signaling cascades has been reported, possibly triggered by the initiating signals represented by the growth factor liberation or the transient elevation of intracellular Ca2+. Thus, for instance, in the case of the purse-string mechanism, activation of members of the Rho family have been reported to be involved in the formation of the actin cable (9). This protein family also participates in the different stages of individual cell migration in general (for review, see Ref. 48) and particularly in the process of wound healing by lamellipodial crawling (1, 20, 31, 39). In addition, during the past few years, preeminent roles have been recognized for diverse ion transport systems in the establishment of the characteristic cell polarization and cytoskeletal rearrangements that take place during cell migration (18, 54). In the case of corneal epithelial wound healing, the development of induced K+ currents has been suggested to participate in the late stages of tissue restitution (58, 59). Finally, it also has been well established that transtissular electric fields spontaneously arise in wounded epithelia (2, 12), possibly generated by the electrogenic activity of the Na+ pump (40). The small direct currents provoked by these fields participate in the orientation and actin reorganization of migrating cells (34, 37, 41, 50) and may act via a complex interaction with enzymatic cascades (57).

In previous work (13), we showed that the nonspecific depolarization of the plasma membrane potential (PMP) provokes characteristic modifications in the cytoskeletal organization of bovine corneal endothelial (BCE) cells in culture. To determine possible physiological implications of these findings, we explored whether membrane depolarization participates in the cytoarchitectural modifications that take place in the course of epithelial wound healing. As mentioned above, the dramatic structural changes undergone by cells actively participating in the processes of wound healing require characteristic cytoskeletal reorganization. It has classically been accepted that corneal endothelial monolayers heal, both in vivo and in vitro, predominantly by cell migration (22, 49, 52). This has been described particularly for relatively large wounds (e.g., >3 mm in diameter). However, it has been suggested that narrower endothelial wounds may exhibit zones of actin cable formation at the leading edge (43). In this work, we report that linear wounds narrower than 150 µm produced on cultured BCE cell monolayers heal by a mechanism that simultaneously combines actin cable formation and cell crawling. The actin cable can be detected as early as 1 h after the injury and progressively extends to border almost the totality of the two edges of the wound. We specially report here the novel finding that, in this system, membrane potential depolarization occurs at the leading edge of the wounds and gradually extends inward toward the neighboring cells. We provide evidence that this spontaneous depolarization is involved in the development of the characteristic actin reorganization demonstrated by the healing cells. Thus the replacement of extracellular Na+ by choline (Ch) provokes a marked decrease in both the depolarization areas and actin cable formation. In addition, we obtained evidence suggesting that the membrane depolarization occurs mainly by an increase in epithelial Na+ channel (ENaC)-mediated Na+ permeability. Most remarkably, we found that the incorporation of ENaC inhibitors determines effects analogous to the ones produced by the replacement of the ambient Na+ by Ch and a marked decrease in the lamellar activity of the wound edges. Finally, the finding that Li+ can replace extracellular Na+ in the determination of the depolarization and cytoskeletal responses suggests that membrane depolarization of the leading cells, and not the increase in intracellular Na+, plays a role in the determination of the cytoskeletal reorganization involved in the healing process. The cellular electrical findings reported here might possibly be related to the generation of macroscopic transtissular electrical fields in the course of epithelial wound healing (see above), but the nature of this relationship is not analyzed in this work. The results reported here support the idea suggested previously by several authors (13, 23, 29, 42) that the PMP of nonexcitable cells may play a role as an intermediary of diverse cellular signaling processes.


    MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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Reagents and solutions. All drugs and reagents were purchased from Sigma Chemical (St. Louis, MO) unless otherwise specified. Control solution (CS) contained (in mM) 127 NaCl, 5.4 KCl, 1.02 NaH2PO4, 3.6 CaCl2, 0.8 MgSO4, and 10 HEPES, pH 7.4. Choline chloride (ChCl) solution contained component similar to those of the CS, but with NaCl replaced by equimolar ChCl. The LiCl solution contained ingredients similar to those of the CS, but with NaCl replaced by equimolar LiCl. As needed, phenamil, kept in a 50 mM stock solution in DMSO, was diluted to a final concentration of 50 µM in the culture medium.

Cell cultures and general experimental procedures. BCE cells were obtained and cultured as described previously (13). The biological materials employed to obtain the cellular cultures utilized in this work come from animals butchered at a slaughterhouse, as mentioned in MATERIALS AND METHODS and ACKNOWLEDGMENTS. We did not handle any live animals whatsoever for this study. Briefly, fresh bovine eyes obtained from the slaughterhouse were processed within 4 h of enucleation. The cornea was dissected and treated with trypsin (0.25%) and EDTA (0.02%) in Ca2+- and Mg2+-free PBS for 20–30 min in the tissue culture incubator. The endothelial cells of each cornea were carefully scraped with a blunt spatula and placed in a 35-mm tissue culture plate containing minimum essential medium (MEM) supplemented with 10% serum, 50 µg/ml gentamicin, 0.25 µg/ml amphotericin B, and 50 µg of total protein/ml of retinal extract. For this study, we used cells from passages 15, which were grown on glass coverslips and had achieved visual confluence at least 5 days before the experiments.

Linear incision wounds were created manually on the monolayers using a fresh 21-gauge syringe at a speed of 5 mm/s. After the occurrence of dead cell detachment (see below), the measured width of the wounds was ~150 µm. The injured cells were then kept in medium or in any of the experimental solutions described above for the corresponding time period at 37°C. The fact that the wound-healing processes exhibited similar electrical and cytoskeletal modifications (see below), both in medium and in saline CS lacking HCO3, permitted us to exclude possible effects caused by the lack of this anion involved in several transport processes in BCE cells (7). Figure 1A shows a typical phase-contrast image of a wound 2 h after injury. At this time, it is possible to observe that a wide area of extracellular matrix, which was left denuded as a consequence of the loss of the overlapping dead cells, has not yet been covered by the restituting tissue. Also, some cells at the wound borders exhibit lamellar activity.



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Fig. 1. A: phase-contrast image of a wounded bovine corneal endothelial (BCE) cell monolayer. A linear wound was produced as described in MATERIALS AND METHODS. The image shown was taken 2 h after the wound had been made; by that time, the dead cells had detached from the monolayer. As a result, the underlying extracellular matrix (EM) became exposed. Some of the border cells show lamellar (L) protrusions. B: oxonol-loaded BCE cell monolayer. A BCE cell monolayer was incubated in the presence of Oxonol V as described in MATERIALS AND METHODS. The original image is shown in gray scale. Arrow points to an isolated cell exhibiting a higher fluorescent signal than the rest of the cells. Arrowheads point to a juxtanuclear intracellular region with low signal intensity corresponding to areas with low vesicle densities (see text for details). Bar, 30 µm.

 
Fluorescence microscopy. To develop the actin, cadherin, and ENaC images, after the corresponding experiments, the wounded monolayers were fixed for 15 min at room temperature with 4% paraformaldehyde in Dulbecco's phosphate-buffered saline (DPBS; pH 7.4) and permeabilized with 0.1% (vol/vol) Triton X-100 in PBS for 15 min. For actin, the cells were incubated with fluorescein isothiocyanate (FITC)-conjugated phalloidin in PBS and 1% bovine serum albumin (BSA) for 15 min at room temperature, washed three times with PBS, and mounted in 80% glycerol (vol/vol) in PBS. For cadherin and ENaC, cells fixed and permeabilized as described above were incubated with an anti-pan-cadherin polyclonal antibody at 1:1,000 dilution and an anti-{alpha}-ENaC polyclonal antibody [Q3560-2 at 1:20 dilution or H-95 (Santa Cruz Biotechnology) at 1:200 dilution], respectively, during 1 h. The anti-{alpha}-ENaC antibody Q3560-2 was generously provided by Dr. Søren Nielsen (University of Aarhus, Aarhus, Denmark). After removal of the primary antibody by washing three times with PBS, we incubated the coverslips with a Cy3-conjugated secondary antibody (Jackson ImmunoResearch, West Grove, PA) for 1 h. The antibodies were then removed, and the coverslips washed in PBS and mounted in glycerol as described above. For all the antibodies, the appropriate dilutions were made in PBS and 1% BSA, and the incubations were performed in a humid chamber. The samples were viewed under a Nikon Optiphot epifluorescence microscope using the appropriate filter set for each probe with a PlanFluor objective and photographed with a Kodak MDS120 digital camera. The camera was controlled using a computer equipped with MDS120 (Kodak Digital Science) and Ulead Photoimpact software (Ulead Systems) as parent application. The images shown are representative of the original data.

The velocity of wound healing was determined on images taken after phalloidin staining. The resulting images were processed with Adobe PhotoShop software as follows: 1) The denuded area was selected with the magic wand tool, and the total number of pixels contained in this area was determined using the histogram command; 2) the quotient between this value and the wound length (expressed in pixels), obtained using the rule tool, was determined and considered to represent the average width of the wound; and 3) the micrometric scale of the corresponding image was used to convert the width value in pixels to micrometers.

Determination of PMP and intracellular Na+ changes. The modifications in the PMP were detected using fluorescence microscopy and the anionic dye Oxonol V [bis-(3-phenyl-5-oxoisoxazol-4-yl) pentamethine oxonol; Molecular Probes, Eugene, OR]. Oxonol V, kept as a 0.7 mM stock solution in ethanol at 4°C, was freshly diluted to a final concentration of 3 µM in the corresponding solution before each experiment. For the experiments, the coverslips containing wounded monolayers were incubated for 30 min at room temperature in the appropriate solution containing 3 µM Oxonol V. In the studies conducted immediately after wounding, the cells were incubated in the oxonol-containing solutions and then injured. In every case, the coverslips were then mounted in a custom-made chamber (13) containing the same solution and placed under a fluorescence microscope. To detect dead cells, all of these solutions also contained 1 µg/ml propidium iodide (PI). The fluorescent images were obtained using a rhodamine filter set and recorded as described in the previous section. Figure 1B shows a typical image of an oxonol-loaded monolayer in which the overall distribution of the dye between the cells is approximately homogeneous, although some isolated cells may exhibit a larger fluorescent signal (Fig. 1B, arrow). As described by Dall'Asta et al. (16), the dye accumulates in the cellular vesicles, thus determining a somewhat heterogeneous intracellular distribution of the fluorescent signal. As also shown in Fig. 1, the cell nuclei appear as negative areas. In our cells, a juxtanuclear area of low vesicle density was frequently observed (Fig. 1B, arrowheads); this was confirmed by staining the intracellular vesicles with acridine orange (results not shown). The higher oxonol intensity displayed by some isolated cells may thus reveal a more depolarized membrane potential, a greater vesicle density, or both. For all of these reasons, the PMP measurements needed to be performed by averaging image regions containing several cells (see below). To enhance the PMP modifications, pseudocolor images were developed from the original ones by transforming the spectrum of intensity of the monochromatic signals into a color spectrum using Scion Image Beta 4.0.2 software.

Calibration curves for the membrane potential were performed using the standard procedure of modifying the external K+ concentration in the presence of the selective ionophore valinomycin (24). For this procedure, NaCl in CS was replaced by potassium gluconate and ChCl such that the corresponding external concentrations satisfied the equation [potassium gluconate] + [ChCl] = 137 mM. In every case, valinomycin was added to a final concentration of 2.3 µM. A linear relationship was found between the logarithm of the oxonol fluorescence intensity of the original images and the membrane potential, calculated assuming a Nernst equilibrium for K+ in the interval between 5 and 130 mM of external K+ concentration as predicted by previous studies (16, 24). For the determinations, the intracellular K+ concentration was assumed to be 130 mM. Because of the heterogeneity of the intracellular distribution of the oxonol signal (see above; see also Ref. 16), the quantitative analyses were performed by determining average intensities of the complete original images or of groups of cells within these images. Also for this reason, the pseudocolor images shown have a qualitative character. From the calibration curves, the PMP of the BCE cells in noninjured monolayers kept in CS was determined to be –48.3 mV (SD 8.7; n = 5), a value in fair agreement with previous determinations (63).

The changes in intracellular Na+ concentration ([Na+]i) were determined using the fluorescent intracellular Na+ indicator sodium green tetraacetate (Sodium Green; Molecular Probes). After the experiments, the cells were incubated for 30 min at room temperature in the corresponding solution containing 5 µM Sodium Green. The coverslips were washed three times and mounted in the same solution plus 1 µM PI in a chamber similar to the one used for the determination of PMP changes. The fluorescent signals were analyzed analogously to the oxonol studies (see above), but using a fluorescein filter set.

As suggested by the manufacturers, and as commonly described in the literature (32, 33, 61), the [Na+]i was calculated from the determined value of fluorescence intensity (F) using the following equation

(1)
where, for room temperature, the dissociation constant (Kd) for Sodium Green is 21 mM and Fmax and Fmin are the maximum and minimum fluorescence intensities, respectively. Fmax was determined after incorporation of gramicidin D (1 µg/ml) into CS and Fmin was determined by replacing the total extracellular Na+ with Ch. In every case, the fluorescence intensity values corresponded to the average intensities of the complete original images or of groups of cells within these images. The average [Na+]i under basal conditions was determined to be 14.1 mM (SD 7.3; n = 5), which is in good agreement with previous studies (8, 28).


    RESULTS
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 MATERIALS AND METHODS
 RESULTS
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Actin reorganization and membrane potential changes during wound healing. Confluent BCE cells in culture exhibit a beehive-like pattern of actin and cadherin organization (Fig. 2, A and B) similar to the one observed in vivo (44). After 3 h of healing of a narrow wound, although the general structural aspects were conserved in most of the tissue, in the cells at and near the wound edges, actin displayed conspicuous modifications. The actin bundles at the cell periphery appeared blurred and diffuse, while actin had become more evenly distributed throughout the cytoplasm (Fig. 2C). Previous confocal microscopic studies reported a decrease in the amount of actin associated to adherens junctions in cells near the wound border (45). Most of the leader cells at the wound edges exhibited a thick actin string that connected neighboring cells, except at sites where the cells had developed a lamellar protrusion. Occasionally, some of these cells extended their lamella beneath the actin cable (Fig. 2C; see also Fig. 4, G and I). The cadherin organization of the cells lining the wound appears to have been less affected (Fig. 2D). In this respect, it is interesting to note that the conservation of junctions at the lateral membrane has been noted to be one of the main advantages of wound repair by means of the purse-string mechanism because it permits maintenance of tissue integrity during the healing process (3, 17).



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Fig. 2. Actin and cadherin distribution of wounded BCE monolayers. Double-staining for actin (A and C) and cadherin (B and D) was performed 3 h after wounding. A and B: noninjured monolayers. C and D: injured monolayers. In contrast to control images (A), the cells at the wound borders (C) exhibited less ordered actin distribution. An actin cable runs continuously from cell to cell along the free edge (C, arrowheads), except in areas interrupted by lamellar protrusions (L). While actin exhibited these noticeable changes, comparison of cadherin distribution between control (B) and injured (D) cells revealed less evident modifications. Bar, 30 µm.

 


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Fig. 4. Dependence of actin reorganization and membrane depolarization on extracellular Na+. AD: Wounded monolayers were left at 37°C for the corresponding time periods; incubated in solutions containing Sodium Green tetraacetate, Oxonol V, and PI; and photographed with the corresponding filters. The Sodium Green (A and C) and Oxonol V (B and D) images showing the same field were processed to obtain pseudocolor images. Intensity scale is similar to that in Fig. 3 (see text for details). Times after injury: 1 h (A and B), 2 h (C and D). Note that there is a temporospatial correlation between the depolarization area and the increase in intracellular Na+ concentration ([Na+]i). E and F: wounded monolayers were left at 37°C for 1 h in control solution (CS; E) and in choline (Ch; F) and then incubated in solutions containing Oxonol V and PI. Shown are pseudocolor images of the original monochromatic ones. Intensity scale is the same as described in previous figure legends. Arrow points to PI-stained nucleus. Note the marked decrease in oxonol fluorescence provoked by the replacement of Na+ by Ch. GJ: wounded monolayers were left at 37°C for 1 h in CS (G and I) and in Ch (H and J) and then labeled for actin. Note the marked decrease in the formation of the actin cable provoked by the replacement of Na+ by Ch (H) with respect to the normal Na+ solution (G). At higher magnification, less lamellar activity is observed in the Ch-treated (J) than in the control (I) monolayers. Bars, 60 µm (E and F), 30 µm (AD, G, and H), and 15 µm (I and J).

 
The time course of the modifications in PMP is shown in Fig. 3. This figure shows fluorescent images of oxonol-loaded cells obtained during the in vivo experiments. Five minutes after injury, most of the dead cells remained attached to the wound border as revealed by the intense PI staining at this level (Fig. 3A). The few dead cells already detached from the tissue permitted us to observe the underlying cells at the wound borders. In all of these areas, no significant differences in oxonol fluorescence could be observed between these cells and the ones in the rest of the monolayer (Fig. 3A, arrowhead). In turn, this finding suggests that there is no difference in the PMP between these two cell populations at this stage. Approximately 1–2 h after the wound was created, most of the dead cells had detached. The majority of the cells at the wound border started to develop an increase in oxonol fluorescence. These regions usually consisted of a string of cells with a length of three or more contacting cells. A typical image is shown in Fig. 3B. These stringwise depolarization regions became progressively more evident, and their presence could be detected even at very late stages of the healing process. Approximately 3 h after the injury, regions showing more extended depolarization zones began to be detected (Fig. 3C), usually in the form of triangle-shaped areas with the vertices pointing toward the undamaged tissue. In the later stages, the increase in oxonol fluorescence extended inwardly in the monolayer and expanded to several rows of cells (Fig. 3, DF).



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Fig. 3. Time course of membrane depolarization during wound healing of BCE monolayers. Wounded BCE monolayers that had been left at 37°C for the corresponding time periods (except in A) were incubated in solutions containing Oxonol V and propidium iodide (PI; see MATERIALS AND METHODS) and photographed. Times after injury: 1 h (B), 3 h (C), 12 h (D), 24 h (E), and 36 h (F). In A, the intact monolayer was preloaded with Oxonol V and PI, wounded, and immediately photographed. The oxonol images were processed to obtain pseudocolor images (see MATERIALS AND METHODS). The intensity scale is displayed with the top corresponding to white and the bottom to black in the gray-scale image. The strong red signal at the wound border in A corresponds to the PI-stained dead cells; some of them have already detached (arrowhead). The more intense oxonol signal of the cells at the leading edge was already noticeable 1 h after the wound (B) and progressively extended inward toward the undamaged tissue (CF). Note that at the late stages, some of the injured areas already exhibited tissue restitution (F). See text for details. Bars, 60 µm (A, E, and F) and 30 µm (BD).

 
The quantitative analysis of the original images (see MATERIALS AND METHODS) revealed that the more depolarized cells along the wound border had achieved an average membrane potential close to zero. Because the calibration curves for the membrane potential were determined within the range –5 to –90 mV, and because nonlinearity may appear beyond these points (24), it is not possible to state whether some of these cells might even reach positive PMP values.

In the next portions of our present study, we sought to determine the mechanisms involved in the spontaneous depolarization of the PMP demonstrated by the cells at the wound border and to establish whether this depolarization plays a role in the cytoskeletal rearrangements that take place in the course of wound healing.

Dependence of actin reorganization and membrane depolarization on extracellular Na+. To assess whether membrane depolarization at the border cells occurs via an increase in Na+ conductivity, we performed double-staining experiments to study the oxonol signal and [Na+]i (Fig. 4, AD). The pseudocolor images shown in Fig. 4, AD, reveal that, in effect, concomitant with the plasma membrane depolarization, there is an increase in [Na+]i of the cells at the wound edges. Quantitative analysis of the original images (see MATERIALS AND METHODS) yielded an average maximum value for [Na+]i of the cells at the wound border of 81.3 mM (SD 9.1; n = 3). As also shown in Fig. 4, AD, the increase in the Na+ signal propagated toward the rest of the epithelium according to the same pattern observed for membrane depolarization.

Replacement of extracellular Na+ by Ch, a nonpermeant cation, determined a decrease in the depolarization areas (Fig. 4, E and F) and conspicuous modifications in the processes of actin remodeling that takes place during the course of the healing of narrow wounds (Fig. 4, GJ). The oxonol images (Figs. 4, E and F) also reveal that, in general, the fluorescence intensity was lower in the Ch-treated layers (Fig. 4F) than in the ones that healed in the presence of normal extracellular Na+ concentrations (Fig. 4E). Concomitant with the modifications in oxonol fluorescence, the replacement of Na+ by Ch provoked an almost complete interruption of the actin changes observed during the healing process (Fig. 4, GJ). While the control monolayers developed an actin cable at the wound border and exhibited characteristic rearrangements of the actin organization that extended to several rows of cells in the form of relocalization of the peripheral actin toward the cytoplasm (Fig. 4, G and I, see also Fig. 2C), the cells that healed in the presence of Ch did not develop a continuous actin cable and conserved their typical circumferential localization of actin (Fig. 4, H and J). Also, lamellar protrusions were markedly diminished (Fig. 4, I and J). In fact, in the Ch experiments, the border cells exhibited an organizational pattern of actin similar to that observed immediately after wounding (data not shown), although some individual cells may form thin, cablelike structures. The possibility that the observed Ch effects could have been provoked by the stimulation of putative acetylcholine receptors, found to be present in several epithelia (60), was discarded by performing experiments similar to the ones described in this section in the presence of 1 µM atropine or 0.1 mM D-tubocurarine (results not shown).

ENaC participation in membrane depolarization and actin reorganization during wound healing. Taken together, the results described in the previous sections are highly suggestive that plasma membrane depolarization occurs spontaneously in the cells at the border of narrow wounds produced on cultured BCE monolayers and that this depolarization is involved in the development of the characteristic remodeling of actin. In addition, the evidence also suggests that membrane depolarization occurs mainly as a consequence of an increase in Na+ permeability. To obtain some evidence regarding the possible mechanism underlying the depolarizing response, we explored whether the augmented Na+ permeability could be determined by an increase in ENaC activity of the border cells, because this channel has been detected and demonstrated to play a role in fluid transport in situ and in cultured corneal endothelial cells (28, 38). Figure 5, AD, shows characteristic images of the effects of phenamil, a specific ENaC inhibitor (21), on the PMP and Na+ concentration of the BCE cells 1 h after wounding. The presence of phenamil in the incubation medium strongly inhibits the [Na+]i increase of the leading cells compared with the control cells (Fig. 5, A and C). The depolarization areas are correspondingly decreased (Fig. 5, B and D), suggesting that the ENaC-mediated increase in Na+ conductance represents the main mechanism of plasma membrane depolarization in these cells. After 1 h of healing in the presence of phenamil, the effects of the drug on the cytoskeletal modifications of the border cells (data not shown) are similar to those induced by the replacement of ambient Na+ by Ch (Fig. 4, GJ). In analogous experiments (i.e., maintaining the monolayers in MEM not supplemented with serum) performed for longer periods of time, phenamil started to produce morphological alterations of the cells and eventually cell death. For this reason, to study the effect of phenamil on the velocity of healing, the wounded monolayers were left in MEM supplemented with 5% serum, which permitted us to prolong the experiments for ~6 h without noticeable effects on cellular morphology and viability (Fig. 5, E and F). As shown, under these conditions, the phenamil-treated monolayers exhibited significantly less actin cable and a smaller degree of wound closure. The average velocities determined for the control and phenamil-treated monolayers (see MATERIALS AND METHODS) were 8.7 µm/h (SD 2.3) and 3.2 µm/h (SD 0.9) (data from 3 independent experiments in which a total of 6 fields/experiment were processed), respectively, for each border.



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Fig. 5. Epithelial Na+ channel (ENaC) participation in membrane depolarization during the course of wound healing. AD: wounded monolayers were left in the tissue culture incubator (37°C, 5% CO2) for 1 h in minimum essential medium (MEM; A and B) and in MEM containing 50 µM phenamil (C and D), and afterward they were loaded with Oxonol V, Sodium Green, and PI in CS and photographed for Sodium Green (A and C) and Oxonol V (B and D) fluorescence. Shown are pseudocolor images of the original monochromatic ones. Intensity scales used are the same as those in Figs. 3 and 4. Note that phenamil determined a decrease in both the Sodium Green (C) and Oxonol V (D) fluorescence intensities at the wound borders. E and F: wounded monolayers were left for 6 h in the tissue culture incubator in MEM supplemented with 5% serum without (E) and with (F) 50 µM phenamil, and afterward they were fixed and stained for actin. Note the marked difference in the wound width between the two images. G and H: wounded monolayers were left for 1 h in the tissue culture incubator in CS (G) and in Li+ (H), loaded with Oxonol V and PI, and photographed. Images and intensity scale are the same as in B and D. Note that in both cases, an increase in oxonol fluorescence occurred at the wound borders. I and J: same as G and H, but labeled for actin. Note that in both images (I and J), the actin cable is present. Bar, 30 µm.

 
The findings described thus far permit us to attribute a role for either membrane depolarization or a rise in [Na+]i in the cytoskeletal response to injuries of the cells at the epithelial wound borders. To distinguish these two possibilities, we explored whether the replacement of extracellular Na+ by Li+ is capable of eliciting an analogous cytoskeletal response. In this respect, it has been well established that ENaC possesses a higher conductivity for Li+ than for Na+ ions (25). In principle, if ENaC-induced membrane depolarization is mainly responsible for the actin reorganization, the replacement of the ambient Na+ by Li+ should still determine the cytoskeletal response. In Fig. 5, GJ, we show that membrane depolarization indeed took place under ambient Li+ conditions (Fig. 5, G and H) and, correspondingly, that actin cable formation also occurred (Fig. 5, I and J). The oxonol fluorescent signal was higher and more extended for Li+ than for Na+. Correspondingly, in the presence of Li+, the actin cable appears to be thicker and less interrupted (Fig. 5J) than in the case of the normal Na+ medium (Fig. 5I).

The augmented presence of ENaC at the wound borders was demonstrated using indirect immunofluorescence with an anti-{alpha}-ENaC antibody (Fig. 6, AD). Immediately after the wound was created, the ENaC fluorescence intensity was extremely low throughout the cell monolayer (Fig. 6B). At that point, the corresponding actin image (Fig. 6A) exhibited the typical aspect of the initial stages. From then on, the ENaC fluorescent signal increased following a time course and distribution similar to the membrane depolarization and to the increase in [Na+]i. In Fig. 6D, we show a typical image obtained 2 h after injury. As shown, the area exhibiting an increase in the ENaC fluorescence corresponds to the area of actin reorganization and cable formation (Fig. 6C).



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Fig. 6. ENaC immunofluorescence during the course of wound healing. AD: monolayers were fixed immediately after wounding and double-stained for actin (A) and ENaC (B) or left at 37°C for 4 h in CS and then processed for actin (C) and ENaC (D). Bar, 30 µm.

 
Figure 7 displays quantitative findings obtained for the development of the wound border processes under different experimental conditions and after 1 h of healing. In this figure, we have categorized these processes into three distinguishable classes according to the corresponding structure found, i.e., total actin cable, lamellar protrusions alone, and neither of these structures (see Figs. 2C and 4G). The simultaneous presence of actin string and lamellar protrusions (see Fig. 4I) has been included in the total actin cable category. As shown in Fig. 7, both the replacement of extracellular Na+ by Ch and the presence of phenamil in the incubation medium determined similar effects, mainly consisting of a dramatic decrease in the total amount of actin cable formed at the wound borders. The replacement of Na+ by Li+ determined that the total actin cable became practically the only structural modification found after the healing time of these experiments.



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Fig. 7. Effect of different agents on the relative length of actin cable formed at the wound borders in BCE monolayers. Wounded BCE monolayers were left at 37°C for 1 h in CS, in choline chloride (ChCl), in CS containing phenamil (Phe), and in Li+ and then labeled for actin. For each image, the relative length of total actin cable (TAC), border displaying lamellar protrusions (La), and of none of these structures (None) present at the edges of the wounds was quantified by performing direct length measurements on photographs of 10x fields made on lines drawn along the wound border. We considered a lamella to be present when a protrusive structure similar to those shown in Figs. 1 and 2C (L) was observed. Otherwise, if no actin cable was detected, the length of the corresponding zone was included in the "None" category. Each bar represents the average (±SD) of three independent experiments performed in duplicate; a total of 10 fields/experiment were processed.

 
Taken together, the results shown in Figs. 57 suggest that ENaC-dependent PMP depolarization participates in determining the cytoskeletal modifications undergone by the cells actively participating in the process of healing of narrow wounds in BCE monolayers.


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The cellular events leading to the modifications of the actin cytoskeleton in the course of epithelial wound healing are complex and incompletely understood (26, 56, 62). As mentioned above (see Introduction), several factors participating in the initial stages of this response have been described. These include stimuli of a diverse nature triggered by the dramatic alterations provoked by the injuries, such as the disruption of the intercellular junctions and mechanical stimulation (62), the massive liberation of diverse molecules to the medium (e.g., ATP and paracrine factors; see Refs. 26 and 62), the onset of a Ca2+ wave (27, 51, 55), and the generation of local electrical fields (2, 12). After these initial events, a complex signaling cascade is activated (20, 56, 62) that promotes the cellular healing response. In our present work, we have used BCE cells cultured to confluence. Under this condition, the cells exhibited some characteristics similar to those shown in situ. These characteristics include a certain degree of polarization as revealed by the pattern of actin distribution and by some recent evidence suggesting a distinctive localization of certain channels and transporters in the apical and basolateral domains (28). In the present report, we communicate for the first time, to our knowledge, the findings that plasma membrane depolarization occurs spontaneously at the cells on the wound border of one epithelium that heals by the development of an actin cable and that this depolarization is crucial to the cytoskeletal rearrangements that participate in the processes of tissue restoration. We have established that the inhibition of this depolarization determines a significant decrease in the development of the cytoskeletal reorganization and ultimately in the velocity of healing of the tissue. These findings allow us to suggest that the spontaneous depolarization of the PMP of the cells at the wound borders may constitute an additional factor participating in the intermediate cellular processes leading to epithelial wound healing. The possibility that wound-induced membrane depolarization may occur in corneal endothelium has been previously speculated by Watsky (58); this author also proposed a probable role for depolarization in the activation of hyperpolarizing currents at the late stages of wound healing. The fact that PMP depolarization constitutes a prerequisite for the cytoskeletal remodeling is not surprising in view of our previous studies demonstrating that experimental depolarization determines characteristic rearrangements of the actin and tubulin networks in epithelia (13, 14). The interpretation that the cytoskeletal modifications undergone by the BCE cells in the course of wound healing are associated to PMP depolarization is consistent with the totality of the results reported here and, as mentioned above, with previous evidence regarding the responses of cultured BCE cells to experimental depolarization. However, because the probes used in the present work to determine modifications in the PMP and [Na+]i were not ratiometric, the possibility that cell volume changes may have affected the fluorescent measurements to some degree cannot be disregarded completely. These volume changes in the cells at the wound border could be determined, for instance, by modifications in the membrane transport systems that could take place in the course of the healing process.

The concurrent findings of this work that phenamil is a potent inhibitor of the depolarization response and of the actin rearrangement, that Li+ can replace Na+ in the generation of the response, and that the ENaC immunofluorescent signal increases in the course of the healing process support the idea that the augmented activity of this channel may be responsible for the increase in [Na+]i and the concomitant plasma membrane depolarization exhibited by the border cells. In this respect, it has been demonstrated that modifications in the number and open probability of ENaC mediates PMP depolarization responses in cultured alveolar epithelium (30). On the basis of our findings, a plausible hypothesis of the sequence of events leading to membrane depolarization could consider the transcriptional activation of genes involved in ENaC expression and its membrane insertion. This activation could be promoted, for instance, by the liberation of growth factors and/or by the intracellular Ca2+ increase that take place immediately after the injury (see above). The wound-induced increase in intracellular Ca2+ has indeed been reported to activate genes involved in the regulation of cellular motility (55). On the basis of our present results, the appearance of the ENaC immunofluorescent signal seems to follow a time course similar to that of the depolarization area. This supports the idea that, for both the border and neighboring cells, membrane depolarization is achieved via an increase in ENaC activity and not by the intercellular propagation of electrical or chemical signals. This notion is further supported by our finding that heptanol and octanol, commonly used blockers of gap junctions (11), did not affect the propagation of the depolarization areas (results not shown). Also in this respect, it is interesting to note that the early Ca2+ wave, which could promote the initiation of the healing response, does not seem to be propagated via gap junctions (27).

The participation of diverse ion channels and transporters in cell motility and migration is well documented (for review, see Ref. 54). Although several possible mechanisms have been suggested to account for this participation, there is a considerable body of evidence relating ion transport systems and the cytoskeleton (18, 19, 46, 54). Thus, several reports point to the crucial role of the ion channels and transporters in the anchoring of the cortical cytoskeleton (4, 19). In turn, the cytoskeleton exerts control over ion transport activity by modulating ionic currents (5, 10, 18). This interrelationship may underlie the participation of some ion transport systems in cell migration. For instance, in the case of the Na+/H+ exchanger, both the cytoskeletal anchoring and the ion translocation are necessary for cell migration and wound healing of PS-120 fibroblasts (18). Similarly, ENaC may participate in actin cable-mediated wound healing via an analogous interaction with some cytoskeletal components. In this respect, it has been reported that the {alpha}-subunit of ENaC associates with spectrin, a member of the cortical cytoskeletal network (64). Also, direct interaction of ENaC with actin results in modifications of the channel conductance (6, 15).

Finally, we remark that, under the conditions used in this study, linear narrow wounds produced in corneal endothelial monolayers predominantly heal by a purse-string mechanism. In this mechanism, the adherens junctions play a key role by providing the sites of anchorage of the actin cable, which allows the constitution of a continuous functional string all along the wound border (17). As emphasized above, one of the central findings of this study is that the development of the actin string is dependent on plasma membrane depolarization of the border cells. This finding is consistent with recent results showing that the actin reorganization induced by the nonspecific depolarization of the PMP of undamaged cultured epithelium occurs during the initial stages concomitant with conservation of the cadherin disposition (14).

In summary, the results described herein permit us to conclude that the actin cable formation and reorganization observed at the edges of narrow wounds produced on BCE monolayers are determined by the spontaneous depolarization of the PMP of the cells present at or near the wound borders. The depolarization seems to be generated by a rise in the ENaC activity of these cells as suggested by the fact that inhibitors of this channel greatly suppress the electrical and cytoskeletal responses, as well as by the increase in the immunofluorescent signal of the channel observed after the production of the wound. Also, the fact that Li+ ions can replace Na+ ions in the determination of the response supports the concept that the membrane depolarization, not the increase in the [Na+]i, is mainly responsible for it. This is consistent with our previous observations that the nonspecific membrane depolarization of confluent BCE cells and other epithelia in culture elicits a characteristic reorganization of the actin cytoskeleton (13). Hence, the findings of the present study further support the idea, suggested by several authors (13, 23, 29, 42), that the PMP of nonexcitable cells may play a role in the regulation of diverse cellular processes.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by Programa de Desarrollo de las Ciencias Básicas and Comisión Sectorial de Investigación Científica (Universidad de la República, Montevideo, Uruguay).


    ACKNOWLEDGMENTS
 
We thank Dr. Søren Nielsen (University of Aarhus, Aarhus, Denmark) for kindly providing anti-{alpha}-ENaC polyclonal antibody (Q3560-2), Susana Tolosa for expert technical assistance, and Frigoríficos Las Piedras and Lorsinal (Uruguay) for supplying fresh bovine eyes.


    FOOTNOTES
 

Address for reprint requests and other correspondence: S. Chifflet, Depto. de Bioquímica, Facultad de Medicina, Universidad de la República, Gral Flores 2125, 11800 Montevideo, Uruguay (E-mail: schiffle{at}mednet.org.uy)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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