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Membrane lipids, EGF receptors, and intracellular signals colocalize and are polarized in epithelial cells moving directionally in a physiological electric field¹

Departments of Biomedical Sciences and
^* Ophthalmology, Institute of Medical Sciences, University of Aberdeen, Aberdeen AB25 2ZD, Scotland, UK

²Correspondence: Department of Biomedical Sciences, Institute of Medical Sciences, University of Aberdeen, Aberdeen AB25 2ZD, Scotland, UK. E-mail: m.zhao@abdn.ac.uk or c.mccaig{at}abdn.ac.uk

SPECIFIC AIMS

Endogenous electric fields (EFs) are common where cell movement occurs, and cells respond to EFs by migrating directionally. We set out to determine whether a physiological EF induces spatially regulated formation, migration, and activation of membrane lipid domains, associated receptors, second messenger signaling molecules, and the actin cytoskeleton in epithelial cells, which could underpin directional cell migration.

PRINICIPAL FINDINGS

1. Asymmetric distribution of membrane lipid, EGF receptors, and signaling molecules in a physiological electric field
Corneal epithelial cells (CECs) from fresh bovine eyes cultured in medium with 10% fetal calf serum (FCS) migrated toward the cathode. In serum-free medium, directional migration was lost. Addition of transforming growth factor (TGF), which activates EGF receptors (EGFRs), causing downstream activation of MAP kinase, restored and significantly enhanced migration directedness and migration rate in a dose-dependent manner. Asymmetric distribution of EGF receptors in live and fixed epithelial cells labeled with specific antibodies was evident (Fig. 1 A–C), and this occurred as early as 10 min after the onset of EF application (Fig. 1B ). Further quantification of the asymmetry showed membrane and juxtamembrane asymmetry of EGFRs in cells cultured in EFs. In most polarized cells, asymmetry of intracellular EGFR staining was also evident.

View larger version (77K): [in this window] [in a new window]   Figure 1. Asymmetry of DiD (A), EGFR (B), dp-ERK (activated MAP kinase), and F-actin (C–P) in corneal epithelial cells exposed to physiological electric fields. A) In an EF of 150 mV/mm, cathodal accumulation (left) of DiD (red) and EGF receptors (green) was colocalized and proportionate (see merged insert and line intensity profile). B) Time lapse images show EGFR accumulated gradually to the cathodal side, within 10 min in an EF (300 mV/mm). C) EGFR accumulated cathodally at the leading edges of a number of cells (150 mV/mm, 1.5 h). G, J, J') ERK (green) was activated preferentially at the cathodal side, together with accumulation of F-actin (red, H, K, K'), and were closely colocalized (I, L–Q). J', K') Surface plots of panels J and K, respectively. Fluorescent intensity is indicated from strongest to weakest by red gradually changing to blue, which illustrates activation of ERK and accumulation of F-actin at the leading, cathodal-facing side of the cell. M–Q) Asymmetry and colocalization were evident throughout the cell from basal to top optical sections. D–F) Asymmetry and close colocalization of dp-ERK and F-actin were not observed in control cells.

To test whether cathodal accumulation of EGFRs was proportional to membrane surface area on the cathodal side of cells, DiD, a dye that stains membrane lipid, was used. Cells were incubated in serum-free medium with DiD (final concentration 3.3 µg/ml, 37°C, 1 h), then exposed to an EF of 150 mV/mm in MEM with 10% FCS for 1 h, fixed, and stained for EGFRs. There was a proportionate cathodal increase of EGFR and of DiD staining. Quantification of an asymmetry index (Ai) for EGFR and DiD indicated that redistribution of EGFR and DiD to the cathodal side was virtually identical (0.20±0.03, n=14 and 0.23±0.03 n=14, respectively; two experiments, P=0.3). (An Ai value > 0 and approaching 1 represents increasing cathodal accumulation of staining; Ai of -1 indicates perfect anodal accumulation and an Ai of 0 indicates no asymmetry.) A Z series of confocal sections taken at 1 µm intervals through the epithelial cells showed equal numbers of sections on cathodal and anodal sides of cells. Each 1 µm section showed greater DiD and EGFR staining cathodally than anodally. This indicates there must be a greater area of membrane on the cathodal side of each 1 µm section. We determined EGFR and DiD fluorescence intensity in control cells, and both showed an even distribution on the left- vs. the right-hand side. One interpretation of these data is that an applied electric field induces a cathodal accumulation of membrane lipids and that the local and directed increase in membrane area this initiates is accompanied by the EGFR, which associates with detergent-insoluble lipid rafts.

We tested whether lipid and EGFR asymmetry was a cause or a consequence of asymmetric actin polymerization by inhibiting the latter pharmacologically. Cells were preincubated in serum-free medium with 0.5 µM latrunculin, a potent inhibitor of actin polymerization (37°C, 1 h), then exposed to an EF (150 mV/mm) in MEM with 10% FCS and 0.5 µM latrunculin. Under these conditions, clear asymmetry of membrane lipid and EGFR fluorescence accumulation at the cathodal side were still evident, indicating that both accumulated independent of actin polymerization. The Ai values for EGFR and DiD fluorescence were 0.27 ± 0.07 and 0.17 ± 0.08, respectively (n=7, P<0.05 when compared with no EF control: EGFR, 0.03±0.01, n=10; DiD, 0.04±0.01, n=16). These data indicate that cathodal redistribution of colocalized EGFR and detergent-insoluble lipid was independent of actin polymerization.

2. Polarized activation of MAP kinase ERK1/2, a signaling molecule downstream from the EGFR
We also tested whether asymmetrically distributed EGFRs induce asymmetric intracellular signaling. Using specific monoclonal antibodies against dual phosphorylated ERK1/2 (dp-ERK), we determined that ERK was activated asymmetrically in cells migrating directionally in EFs (Fig. 1D-Q ). Marked cathodal asymmetry of dp-ERK was evident 1.5 h (earliest time tested) after exposure to an EF of 150 mV/mm. The asymmetry was evident in the much brighter staining of the lamellipodium facing the cathode (Fig. 1G-Q ), but non-lamellipodium asymmetry of dp-ERK was evident as well (Fig. 1O-Q ). Cells cultured without an EF did not show this asymmetric staining pattern (Fig. 1D-F ). Figure 1J ', K' dramatically highlights the asymmetry.

Most cells cultured in EFs extended lamellipodia toward the cathode. More than 70% of single cells cultured in an EF showed obvious cathodal dp-ERK asymmetry. This is nearly double the proportion of cells previously reported to show cathodal asymmetry of EGFR (38%). Asymmetric activation of ERK1/2 was quantified. Ai values for F-actin and dp-ERK after field exposure were 0.22 ± 0.03 and 0.21 ± 0.02, respectively (n=17), significantly higher (P<0.001) than for control cells not exposed to EFs (0.04±0.03 and -0.01±0.02, n=20). Activation of ERK was more marked in peripheral regions of EF-exposed cells than in control cells (no EF). dp-ERK staining was present predominantly at the cathode facing lamellipodium in an EF (Fig. 1G-Q ), but was largely perinuclear in control cells (no EF) (Fig. 1D-F ).

Six cells selected randomly from the 17 cells quantified for Ai above were used to compare the asymmetry of F-actin and dp-ERK in lamellipodium, cell body, and whole cell. The Ai’s for F-actin and dp-ERK were higher in lamellipodia than in the cell body. Ai’s for F-actin were 0.24 ± 0.06 at lamellipodia vs. 0.22 ± 0.07 for the whole cell and 0.10 ± 0.06 for the cell body area. Ai for dp-ERK was 0.25 ± 0.04 for lamellipodia vs. 0.10 ± 0.03 for the whole cell (P=0.01) and 0.03 ± 0.02 for cell body (P<0.001). dp-ERK was more closely colocalized with F-actin in EF-exposed cells (Fig. 1I ) than in control cells (Fig. 1F ), with a correlation coefficient between the Ai’s for F-actin and dp-ERK of 0.71 in field treated cells but only 0.25 in control cells (no EF, P<0.01).

In addition to the remarkable lamellipodial asymmetry of dp-ERK in EF exposed cells (Fig. 1G-N ), optical sections showed a higher degree of ERK activation cortically or just beneath the cortical F-actin staining at a cellular level above the lamellipodium (Fig. 1M-Q ). Western blot showed increased ERK1/2 activation when exposed to electric fields. This required the presence of serum.

Cells treated with 0.5 µM latrunculin still showed clear asymmetric activation of ERK in an applied EF, indicating this occurred independent of actin polymerization. The Ai for dp-ERK staining was 0.17 ± 0.03 (n=15), significantly higher than that of no EF controls 0.02 ± 0.06 (n=8) (P=0.02).

3. Inhibition of ERK/PI3 kinase activation significantly reduced directed migration
Exposing the epithelial cells to 50 µM LY294002 (PI3 kinase inhibitor) or U0126 (MAP kinase inhibitor) or a mixture of both drugs also significantly decreased the migration rate and the directedness. The average Ai’s for F-actin and dp-ERK of cells cultured in EFs and exposed to the inhibitor U0126 (0.10±0.05, -0.11±0.08, respectively, n=12) were not significantly different from those of the control cells cultured without fields (-0.01±0.04, 0.13±0.10, n=10), indicating that the ERK1/2 inhibitor U0126 prevented asymmetric signaling and that the cathodal redistribution of the F-actin cytoskeleton is downstream of and dependent on asymmetric activation of ERK1/2.

CONCLUSIONS AND SIGNIFICANCE

This study extends an understanding of the mechanisms underpinning EF-directed CEC migration, which requires growth factors and depends on an induced asymmetry of EGF receptors and subsequently of F-actin. It establishes that 1) membrane lipids and EGF receptors accumulate cathodally along with elements of the MAP kinase signaling pathway, which becomes activated asymmetrically in cathodally migrating epithelial cells; 2) cathodal accumulation of lipids, associated receptors, and signaling molecules (ERK1/2) do not depend on, but induce, a cathodal redistribution of F-actin; 3) an EF induces a closer cathodal colocalization of activated ERK1/2 kinase and F-actin; and 4) inhibitors of MAP kinase signaling inhibit directed CEC migration and cathodal asymmetry of activated ERK1/2 and F-actin. Collectively, these findings pinpoint some of the major molecular elements responsible for transducing and effecting EF-induced, directed cell migration, outline the sequence of these events, and allow a comparison with recently established signaling pathways that drive chemotaxis.

To what extent might common principles or mechanisms underpin chemotaxis and electrotaxis? In a gradient of chemoattractant, the social amoeba Dictyostelium discoideum and human neutrophils show strong chemoattraction, but neither concentrates chemoattractant receptors at the leading front of the cell. Neutrophils increase membrane area at the front; this induces a relative increase in receptors compared with other areas, although receptor concentration per unit of cell membrane does not increase. Both cell types also show strong, selective activation and translocation of intracellular signaling molecules (pleckstrin homology domains, for instance) to the side of the cell facing the higher concentration. In both cases, this occurs independent of actin polymerization and is likely to be a cause rather than a consequence. This is illustrated in Fig. 2 A. In Dictyostelium and human neutrophils, chemoattractant receptors are not polarized; rather, the cell responds to a polarized gradient of ligand by translocating membrane and receptors in equal proportion to the leading edge, creating a greater area of highly folded membrane at the front. This induces stronger signals (indicated by thicker arrow) at the leading than at the trailing edge. As a result, signaling molecules are activated in a gradient from the leading to the trailing edge, and this determines directional migration. Similarly, an induced accumulation of membrane lipid and associated receptors cathodally in a physiological EF would induce stronger signaling at the cathodal side (Fig. 2) . This signaling pattern (higher cathodally than anodally) is analogous to the signaling pattern caused by a chemoattractant gradient. In chemotaxis, there is a gradient of ligand extracellularly; this may also arise in a physiological EF. The endogenous EF within the developing prelimb bud region of amphibian embryos induces a directional gradient of microinjected, charged, florescent protein molecules, effectively setting up an EF-induced chemical gradient. In situations where chemotaxis occurs in vivo, endogenous EFs clearly coexist. A further example is at an epithelial wound. An immediate consequence of rupture is local collapse of the transepithelial potential difference, although normal values are maintained 0.5–1 mm away from the wound margin. Inevitably, cells engaged in wound healing do this in the presence of a laterally oriented EF, which may be capable of inducing directed cell movement and a local chemical gradient.

View larger version (40K): [in this window] [in a new window]   Figure 2. Model mechanisms for A) chemotaxis and B) epithelial cell electrotaxis. In chemotaxis, a ligand gradient causes more receptors on one side of the cell to become activated (red) than the other side, resulting in asymmetric signaling, which may amplify the asymmetry (A). In electrotaxis (B), a similar asymmetric signaling may occur due to EF-induced asymmetry of membrane lipids and associated receptors, although ligand may also become redistributed (see text).

In summary, we have shown that asymmetric signaling through the EGFR is critical for directional migration of cultured corneal epithelial cells in a physiological EF, that redundancy and multiple signaling pathways are involved downstream, and that significant similarities exist between the mechanisms underlying chemotaxis and electrotaxis. Because a wound-induced EF is immediate and capable of inducing chemical gradients, it is likely that the two guidance phenomena will be intricately interrelated.

FOOTNOTES

¹ To read the full text of this article, go to http://www.fasebj.org/cgi/doi/10.1096/fj.01-0811fje; to cite this article, use FASEB J. (April 10, 2002) 10.1096/fj.01-0811fje.

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