About this Journal Submit a Manuscript Table of Contents
International Journal of Cell Biology
Volume 2012 (2012), Article ID 121424, 13 pages
http://dx.doi.org/10.1155/2012/121424
Review Article

The Plasma Membrane Potential and the Organization of the Actin Cytoskeleton of Epithelial Cells

1Departamento de Bioquímica, Facultad de Medicina, Universidad de la República, Gral. Flores 2125, 11800 Montevideo, Uruguay
2Sección Biofísica, Facultad de Ciencias, Universidad de la República, Iguá 4225 esq. Mataojo, 11400 Montevideo, Uruguay

Received 7 August 2011; Accepted 8 October 2011

Academic Editor: Michael Hortsch

Copyright © 2012 Silvia Chifflet and Julio A. Hernández. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

The establishment and maintenance of the polarized epithelial phenotype require a characteristic organization of the cytoskeletal components. There are many cellular effectors involved in the regulation of the cytoskeleton of epithelial cells. Recently, modifications in the plasma membrane potential (PMP) have been suggested to participate in the modulation of the cytoskeletal organization of epithelia. Here, we review evidence showing that changes in the PMP of diverse epithelial cells promote characteristic modifications in the cytoskeletal organization, with a focus on the actin cytoskeleton. The molecular paths mediating these effects may include voltage-sensitive integral membrane proteins and/or peripheral proteins sensitive to surface potentials. The voltage dependence of the cytoskeletal organization seems to have implications in several physiological processes, including epithelial wound healing and apoptosis.

1. Introduction

The transport of water and solutes across epithelial layers represents a major achievement of biological evolution and constitutes the basis for the existence of higher organisms [1]. To accomplish their transport properties, the epithelial cells acquire characteristic structural and functional features. An epithelial layer must constitute a well-defined macroscopic permeability barrier, which results in the selective transport of solutes and water across the overall tissue. For this, transport epithelia must develop a complex set of cell junctions and a polarized distribution of membrane molecules, which localize at distinct apical and basolateral domains of the plasma membrane [25].

The establishment and maintenance of the polarized epithelial phenotype require a characteristic organization of the cytoskeletal components. There are many cellular effectors involved in the regulation of the cytoskeleton of epithelial cells [611]. Recently, modifications in the plasma membrane potential (PMP) have been shown to participate in the modulation of the cytoskeletal organization. The purpose of this paper is to review evidence relating PMP modifications to changes in the cytoskeletal organization of epithelia, with an emphasis on the actin cytoskeleton, and discuss possible molecular paths mediating these effects. Prior to this, we briefly review the basic characteristics of the cytoskeleton and the generation of plasma membrane potentials in epithelial cells.

2. General Morphological Aspects and Organization of the Cytoskeleton of Epithelial Cells

Mature epithelia are characterized by two fundamental morphological and functional features: a tight cellular packing, supported by the existence of strong adhesive forces between neighboring cells and a polarized cellular phenotype [5]. These two properties are interdependent, since the establishment of intercellular junctions represents the main positional cue triggering the development of cell polarization. All the anchoring junctions, either between cells or between cells and substrate, are associated to cytoskeletal components that are crucial for the junction stability. The characteristic organization of the cytoskeleton of epithelial cells greatly depends upon these interactions with the cell junctions, as depicted in Figure 1. In particular, the actin cytoskeleton associates with diverse cellular structures in mature epithelial cells (Figure 1). In these cells, the most conspicuous actin structure is the circumferential actin belt, a bundle of actomyosin fibers located immediately beneath and associated with the tight and adherens junctions. Here, the microfilaments and the cell junctions interact via a complex set of multifunctional proteins (Figure 2(a)). Actin filaments can also be found along the lateral membrane [1214], where it codistributes with myosin I [1517]. Short actin filaments are part of the spectrin-based membrane skeleton network. Besides its classical role in membrane domain organization [18], this network is involved in the biogenesis of the lateral membrane of epithelial cells and in the maintenance of their columnar shape [19, 20]. Stress fibers at the basal domain are also actomyosin structures associated to focal contacts and other cell-substrate junctions [21]. At the apical domain, parallel bundles of crosslinked actin filaments, extending from the terminal web, constitute the core of the microvilli.

121424.fig.001
Figure 1: Schematic representation of the cytoskeletal organization of a transporting epithelial cell. The scheme shows the cell-cell and cell-substrate junctions connecting the actin, tubulin, and intermediate filament cytoskeletons. See text for details.
fig2
Figure 2: (a) General schematic representation of an anchoring junction. The scheme depicts the basic organization of the proteins comprising an anchoring junction. The three organizational levels (see text) are represented by different colors; modified from Matter and Balda [34]. (b) Scheme representing some of the interactions between ENaC and the cytoskeleton. The channel is kept in its membrane location via the spectrin-based cytoskeleton. In the nonstimulated state, ENaC is bound to F-actin directly and/or by cortactin-Arp2/3 interaction (left panel). Diverse stimuli promote formation of short actin filaments that activate ENaC by direct binding to the channel (right panel). See main text for references. (c) Scheme representing some of the interactions between the Na+-K+ ATPase (NKA) and the cytoskeleton; modified from Bennett and Healy [18].

Polarized epithelial cells exhibit a characteristic pattern of microtubule organization (Figure 1) [6, 22, 23]. Unlike fibroblastic-like cells, where centrosomes are responsible for the nucleation and organization of microtubules, in epithelial cells, most microtubules are acentrosomal. They also present distinctive properties in their behavior, stability, dynamics and regulation [2427]. Typically, in epithelia microtubules organize in long apicobasal-oriented parallel fibers that span the whole length of the cell, with their minus end pointing towards the apical surface and in disordered short filament networks underneath the apical and basal membranes [2831]. This particular arrangement of microtubules is crucial for the targeted traffic of vesicles that sustains the existence of distinctive apical and basolateral domains of epithelial cells [23, 32]. Figure 1 also schematically shows that the other major component of the cytoskeleton, the intermediate filaments, traverses epithelial cells connecting desmosomes and hemidesmosomes [33].

Intercellular anchoring junctions are key structures for epithelial organization and function. Among these, the tight junctions (zonula occludens) selectively seal the intercellular space at the apical side and prevent the exchange of membrane proteins and lipids between the apical and basolateral domains. Underneath the tight junctions, the adherens junctions (zonula adherens) and desmosomes are the main responsible for the mechanical strength of the cell-cell contacts. A schematic description of the structure of anchoring junctions is depicted in Figure 2(a). As emphasized by Matter and Balda [34] for the case of tight junctions, three levels of organization can be recognized. This conception can be generalized to all the anchoring junctions. The first level corresponds to the integral membrane proteins, the second to a set of proteins mediating, among others, the interaction between the cytoskeleton and the adhesion proteins, and the third to the cytoskeletal proteins. The second level is made up of a complex array of soluble proteins, with many of which being capable to transit between other cellular compartments and their juxtamembrane location at the corresponding cell junction [33]. This complex includes transcription factors, regulatory proteins, and scaffolding proteins and is a critical site for the transduction of diverse types of signals [35]. Among these, the acquisition of the epithelial phenotype is triggered by the recruiting of proteins at the second and third junction levels, initially induced by the establishment of cell-cell and cell-substrate contacts [36]. The composition and interactions of the proteins of the second level are highly complex and still remain matters of active investigation. The general organizational pattern shown in Figure 2(a) for the anchoring junctions can also be extended to include the linking between other types of integral membrane proteins, such as ion channels and pumps and the cytoskeleton (see below).

Adherens junctions (AJs) have a central role in the establishment and maintenance of the epithelial phenotype. In this respect, classical cadherins have been recognized to be involved in the establishment of early intercellular contacts and in the organization of microfilaments and microtubules [8, 11, 36]. The association between AJ and these cytoskeletal components is interdependent, since the actin and tubulin cytoskeletons in turn contribute to AJ formation, stability and strength [3743]. Thus, when actin cytoskeleton is disrupted, AJ formation is impaired [38, 39, 44]. Moreover, actin cytoskeletal reorganization, as induced for instance by extracellular calcium deprivation, may determine AJ and tight junction disruption with the consequent loss of the epithelial integrity [45, 46]. Likewise, cell-cell contacts stabilize the plus and minusends of microtubules at the adherens junctions, while the blocking of their dynamic turnover provokes brakeage of cell-cell contacts [26, 43, 47].

Even in the quiescent state, the cytoskeleton of epithelial cells is highly dynamical, undergoing constant assembly and disassembly of its structural units. For this reason, it is subject to complex mechanisms of regulation, many of which involve components of cell-cell junctions and other membrane proteins. Since it is not the purpose of this work to make a thorough revision of this rather involved issue, the reader is referred to several specialized reviews [11, 36, 48, 49]. In Section 5 we will only mention some regulatory pathways that are possible candidates to mediate the responses of the cytoskeleton to modifications in the plasma membrane potential.

3. Interactions between Ionic Transport Systems of the Plasma Membrane and the Cytoskeleton

Besides its associations with the cell junctions, the cytoskeleton also interacts, directly or indirectly, with diverse ion transport systems of the plasma membrane [5053]. Similarly to the case of the cell junctions, many of the associations of the cell membrane channels and transporters with the cytoskeleton are interdependent (for detailed reviews, see [50, 52]). In this way, several transport systems are anchoring sites for the cortical cytoskeleton [5462]. Conversely, the binding to the cytoskeleton modulates diverse ion transport activities [56, 61, 6368]. This interrelation between the cytoskeleton and ionic transport systems plays a relevant role in the physiological properties of transport epithelia [69, 70]. For example, in the renal medulary thick ascending limb the Na+/H+ exchanger regulates bicarbonate absorption by controlling the organization of the actin cytoskeleton [71]. In human syncitiotrophoblast, gelsolin (an actin regulatory protein) stimulates nonselective cation channels of the TRP family in the presence of calcium [72]. In HEK 293 cells, calcium-activated chloride channels require cytoskeletal interactions to achieve full activation [73]. A most noteworthy example of the modulation of ionic transport by the cytoskeleton is provided by volume-sensitive ion transport systems, that modify transport rates in response to changes in cytoskeletal tension [7476]. In several epithelial cells, where water and salt transcellular transport determine modifications in the cellular volume, stretch-activated potassium and chloride channels participate in the regulation of salt transport [7779]. Among other examples of the regulation of ionic conductances by the cytoskeleton, in mammary adenocarcinoma cells a well-organized actin network is necessary for the proper activation of CFTR by cAMP [80].

The interrelationship between ion transport systems and the cytoskeleton is also crucial in cell adhesion and migration, processes requiring significant cytoskeletal remodeling and modifications of ion transport. In this respect, it has been shown that inhibition of the CFTR-dependent conductance impairs lamellipodia formation in bronchial epithelial cells [81]. In T-cells, Kv1.3 channels participate in promoting adhesion by establishing complexes with 1 integrin [82]. In neuroblastoma cell lines, TRPM7 channels affect cell adhesion by direct interaction with the actomyosin cytoskeleton [83]. TRPV4 has also been reported to form complexes with the actin cytoskeleton and regulatory kinases, involved in lamellipodial formation [61]. Focal adhesion kinase, an enzyme involved in integrin-mediated focal adhesion, is activated by forming complexes with Kv1.2 channels both in epithelial and nonepithelial cells [84]. The inhibition of either the Na+/H+ exchanger transport activity or its actin cytoskeletal anchoring significantly decreases migration of PS-120 fibroblasts [66]. Also, ENaC has been reported to participate in the processes of cell migration and wound healing in epithelia [85, 86] and other tissues [87]. The molecular nature of this role may imply direct interactions between this channel and cytoskeletal components. In this respect, it has been shown that the alpha subunit of ENaC associates with spectrin [54, 88] and short actin filaments [89]. In addition, there is evidence that the direct binding between ENaC and these filaments modifies the channel conductance [9092]. More recently, Ilatovskaya et al. [68] have shown that, in renal epithelial cells, the actin cytoskeleton regulates ENaC activity via a cortactin-Arp2/3 complex. As an example of the structural relationships between an ion channel and the cytoskeleton, Figure 2(b) resumes diverse evidence in a schematic diagram of the interactions between ENaC and cytoskeletal components during nonactivated and activated conditions [50, 54, 68, 88]. To be noted, the mechanism through which the cytoskeleton promotes modifications in ionic conductance may depend on the particular transport system. For the case of ENaC, Figure 2(b) shows that the channel activation requires direct binding to short actin filaments, produced by direct PKA phosphorylation of actin or via an actin regulatory protein [50]. As another example, in vascular endothelia 4.1 proteins have been proposed to be necessary for the stable expression of TRPC4 in the plasma membrane [93].

The sodium pump has also classically been recognized to establish interactions with the cortical cytoskeleton [94]. More recently, it was shown that this enzyme participates in tight junction assembly in MDCK cells via activation of RhoA and stress fiber formation [9597]. In caveolar structures the sodium pump is the core of a multiprotein complex, the sodium pump signalosome, that contains several proteins involved in cytoskeletal regulation [98]. Moreover, in vitro experiments reinforce the existence of interrelations between the Na+-K+-ATPase and actin [64]. The molecular nature of the interactions between the sodium pump and the cytoskeletal components, and their physiological relevance, are beginning to be known in more detail. In this respect, the critical role of the interaction of ankyrin-G and the spectrin-based actin cytoskeleton in the membrane retention of the sodium pump has been well established [19, 99]. The binding of ankyrin with the α1-subunit of the Na+-K+ ATPase has been shown to be crucial not only for the membrane anchoring of the enzyme, but also for its trafficking in the polarized cell [100]. Based upon the available evidence, the scheme of Figure 2(c) summarizes some of the recognized interactions between the sodium pump and the cytoskeleton in vertebrate cells. In D. melanogaster, Dubreuil and coworkers demonstrated the existence of ankyrin-independent interactions between the sodium pump and spectrin [101, 102].

4. The Plasma Membrane Potential of Epithelial Cells

The classical dogma of the ionic transport properties of epithelia was founded by Ussing and coworkers in the 1950s and has ever since become the basic paradigm of epithelial transport [103105]. According to this model, in its essential terms, the polarized distribution of the sodium pump and sodium channels is the basic process that transforms a homogeneous cell into an epithelial cell, capable of performing net transepithelial transport of salt between two separated compartments (Figure 3). Although the general scheme of Figure 3(b) may apply to different epithelial cell types, it is particularly characteristic of tight epithelia [106]. In this respect, it was recognized that epithelia can be divided into two categories, tight and leaky, according to the electrical resistance of the tight junctions [107]. In addition to this difference, the two types are distinguished by other transport properties. Thus, besides the low electrical resistance, leaky epithelia also exhibit a high water permeability and establish low-transepithelial-potential differences (see below), while the reverse is true for tight epithelia [108].

fig3
Figure 3: Schemes showing some ionic transport systems of the plasma membrane. (a) In symmetric, nonpolarized cells the sodium pump maintains the electrochemical gradients of sodium and potassium across the plasma membrane. The sodium and potassium channels underlie the generation of a diffusion potential across the plasma membrane. Chloride is usually maintained at activities close to equilibrium. (b) In polarized epithelial cells the asymmetric distribution of, mainly, sodium channels and the sodium pump into distinctive apical and basolateral membranes may determine a net transcellular transport of sodium chloride. (ap: apical; bl: basolateral; the orange circles represent the sodium pump, and the other arrows correspond to ionic channels).

In general, since the apical and basolateral domains of epithelial cells have different compositions of ionic transport systems, the mechanisms of generation of the membrane potential differ between them. For the case shown in Figure 3(b), the PMP across the basolateral membrane is approximately given by a diffusion potential dominated by potassium. Among other differential contributions, the sodium pump may be responsible for the generation of the basolateral PMP in an electrogenic fashion, particularly if potassium permeability is low [109]. The composition of the apical membranes greatly varies among the different epithelial cell types. Therefore, the generation of the apical PMP depends on the specific epithelia considered. For example, in intestinal epithelial cells the sodium-glucose electrogenic cotransport may be the major contributor to the apical PMP [110]. The selective modifications in the ionic conductances at the apical or basolateral domains may play physiological roles in diverse epithelia. Thus, in the acinar cells of parotid salivary glands, modifications in the potassium conductance of the apical membrane produce changes in the fluid flow via changes in the apical PMP [111113]. In pancreatic acinar ducts, modifications in the apical depolarization mediated by the sodium-glucose cotransporter modulate the amount of basolateral uptake of chloride and bicarbonate [114]. Another example is given by the basolateral membranes of tracheal and intestinal cells, where potassium channels containing the KCNE family of -subunits control the chloride flux via modifications in the basolateral PMP [115].

In epithelial cells, the differential generation of the PMP at the two membrane domains determines the existence of a net transepithelial potential difference (TPD). As mentioned above, tight epithelia are capable to maintain large TPDs as a consequence, among other factors, of the existence of a large electrical resistance of the paracellular pathway [108]. On the contrary, the TPDs across leaky epithelia are generally small. As an example, Figure 4 shows a scheme of some of the ionic transport systems of corneal endothelium, a typical leaky epithelium that pumps salt and water from the corneal stroma to the aqueous compartment of the eye [116, 117]. As can be seen, the presence at the two domains of sodium-bicarbonate cotransporters with different stoichiometric ratios is the cause for a small TPD (i.e., less than 1 mV). Among other possible roles, as in paracellular ion movement, this TPD may have a relevant function in the mechanism of solute-solvent coupling across corneal endothelium [118].

121424.fig.004
Figure 4: Scheme of a corneal endothelial cell showing electrogenic transport systems of the apical (ap) and basolateral (bl) membranes. Orange circle and arrows as in Figure 3, pale orange circles represent sodium-bicarbonate cotransporters; modified from Montalbetti and Fischbarg [117].

Epithelial cells in culture may acquire some structural and physiological characteristics of the in situ epithelial cells, including the generation of a TPD [119121]. For the case of small TPDs, as is the case of corneal endothelium [116], the employment of voltage-sensitive fluorescent probes provides with global PMP values which are approximately equal to the apical and basolateral PMPs of these cells in culture [117, 122] but does not permit to distinguish between them. Electrophysiological procedures are required to determine the PMP at each one of the plasma membrane domains, both under in situ or culture conditions.

5. Modulation of the Epithelial Actin Cytoskeleton by Modifications in the Plasma Membrane Potential

Several authors have suggested that the plasma membrane potential of nonexcitable cells could play a role in diverse cellular processes [123125]. In MDCK cells, Vaaraniemi et al. [126] found that activation of protein kinase C determined PMP depolarization and reorganization of the spectrin-based and actin cytoskeletons. Consistently with these findings, we showed that the nonspecific modifications of the PMP (i.e., depolarization or hyperpolarization) promote changes in the organization of the actin and tubulin cytoskeletons in bovine corneal endothelial (BCE) cells in culture [127, 128]. In particular, the changes observed for the actin cytoskeleton consisted, for the PMP depolarization, in a gradual loss of the peripheral ring, an increase in F-actin throughout the cytoplasm, appearance of intercellular gaps and, for sufficiently prolonged treatments, eventual cell detachment [127]. Conversely, it was noteworthy to confirm that PMP hyperpolarization determined the opposite response, that is, an increase in the compactness of actin at the peripheral ring and an augmented resistance of intercellular adhesion to diverse destabilizing procedures [128]. These cytoskeletal responses to PMP modifications were characteristic of some cultured epithelia in confluence displaying a typical epithelial phenotype exhibiting, among other characteristics, a well-defined circumferential actin ring, whereas nonconfluent or undifferentiated epithelial cell lines did not manifest a recognizable response [128, 129]. Effects of PMP depolarization on microfilaments were also observed in kidney tubular cells, where the authors demonstrated that Rho activation and the consequent increase in phosphorylation of the light myosin chain are involved [130, 131]. The role of the PMP on the cytoskeletal organization was also supported by the finding that, in vascular endothelia, depolarization decreases cell stiffness by affecting the cortical actin cytoskeleton [132, 133].

The activities of diverse signaling intermediates are sensible to modifications of the PMP. Thus, in excitable cells, regulators of G protein signaling (RGS), Rho proteins and PKA are activated by the calcium increase provoked by PMP depolarization [134140]. In renal epithelia, however, the activation of Rho determined by depolarization is not mediated by cytosolic calcium increase [130, 141]. Among the integral membrane proteins that could mediate cytoskeletal responses to diverse effectors, the phosphatidylinositol phosphatase Ci-VSP [142] and its homologs [143, 144] contain a voltage sensor in the transmembrane domain [145] and produce PIP2, a well-known regulator of the actin cytoskeleton [146, 147]. Interestingly, this enzyme, present in epithelial and nonepithelial cells [143, 144], is activated by PMP depolarization [142]. More recently, in Xenopus oocytes Zhang et al. [148] described an alternative voltage-sensitive mechanism to increase PIP2 level in response to PMP depolarization, via activation of a PI4 kinase. The G protein-coupled receptors (GPCR) constitute another family of membrane proteins shown to initiate signaling paths leading to actin remodeling [149, 150]. These receptors, activated by a variety of extracellular effectors, are directly regulated by the PMP [151156].

At this point, it should be reminded that the “plasma membrane potential,” as determined from typical electrophysiological procedures, refers to net electrical potential differences between the intra- and extracellular bulk compartments. This difference comprises a series of intermediate electrical potential changes that include surface potentials at membrane proximities and the transmembrane potential [123, 157, 158]. Changes in the surface potentials could affect peripheral proteins, many of them involved in cytoskeletal regulation [123, 158]. For example, diverse putative peripheral proteins can bind to the inner surface of the plasma membrane by electrostatic interactions and modify their degree of attachment in response to modifications in the surface potential, such as MARCKS [159], PTEN [160], K-Ras [161], c-Src: [162], Rac1 [163], and ERM proteins [164]. For the particular case of K-Ras it has been shown that, apart from the inner surface potential, the transmembrane potential also affects its binding to the plasma membrane [165].

The effects of the PMP on the cytoskeletal organization could also be mediated by membrane ionic transport systems directly or indirectly connected to cytoskeletal elements, such as the ones mentioned in Section 3. In principle, there are two main mechanisms through which the PMP could affect the interrelationship between ion transport systems and the cytoskeleton: (a) by modifying ionic currents and thus the ionic environment near the cytoskeletal binding regions, and (b) by determining electroconformational modifications of the ion transporting proteins that can be propagated to the cytoskeletal components, as could be the case for other nontransporting integral membrane proteins. As an example of this latter possibility, several ion channels have been proposed to transmit signals via conformational coupling with integrins, irrespectively of changes in the ionic fluxes [166]. Whatever the mechanism, it must be noted that the particular ion transport system that would mediate the cytoskeletal response to a certain PMP modification (i.e., a hyperpolarization or a depolarization) may depend on the specific procedure employed for the modification. For instance, in a certain cell type PMP hyperpolarization may be achieved by increasing the potassium conductivity, determining augmented potassium efflux, or by increasing the chloride conductivity, producing increased chloride influx. Correspondingly, the intermediate path and specific organizational response of the cytoskeleton to the particular PMP change provoked may also depend on the specific procedure and ionic path (cf. Section 3).

Another possible mechanism mediating the effects of PMP modifications on cytoskeletal rearrangements could be the direct conduction of electrical signals generated at the plasma membrane by the cytoskeletal components themselves. In support of this idea is the finding that actin filaments can propagate electrical signals per se [167].

6. Possible Physiological, Pathological, and Medical Implications of a Regulation of the Actin Cytoskeleton by the Membrane Potential

In principle, it could be expected that the transition of an epithelial cell from a quiescent to a specific secretion or absorption state occurs with characteristic modifications both in cytoskeletal organization and ionic conductances. As suggested from the evidence reviewed in this work, these concurrent modifications could be mediated by the complex regulatory framework provided by the interactions between cytoskeletal components and diverse membrane transport systems. From the results reviewed in the previous section the modifications in the plasma membrane potential could participate in the regulation of the organization of the cytoskeleton of epithelial cells, possibly via effects mediated by ionic transport systems. The examples shown in Section 3 support this notion by describing diverse examples of epithelial cells where ionic transport is associated to cytoskeletal modifications.

As a further physiological counterpart to the results commented in Section 5, we put into evidence that PMP depolarization occurs during wound healing in bovine corneal endothelial cells, as a consequence of the increased expression of the epithelial sodium channel (ENaC), and that it may have a role in the healing process [85]. To be noted, the border cells actively participating in the healing response of epithelia experience characteristic reorganizations of the actin cytoskeleton, which are blocked by ENaC inhibition (ibid). ENaC-dependent PMP depolarization was also observed in the course of wound healing by others in a cell line of human trophoblast [86] and by us in other epithelia in culture (unpublished results). Interestingly, in healing corneal endothelium and epithelium Watsky [168] described a late hyperpolarizing potassium current that, in view of the results described above, could have the role of restituting the membrane potential to its basal value. A role for ENaC in the processes of wound healing was also proposed by Grifoni et al. [87] for smooth vascular muscle cells.

Actin has been found to participate in the development of the apoptotic response [169]. Thus, interference with actin dynamics by inhibition of its depolymerization enhances apoptotic activity in HL-60 cells [170, 171]. However, in T-cells disruption of the actin cytoskeleton promotes caspase-3-mediated apoptosis [172]. A concurrent finding of interest within the conceptual framework of this paper is that cells undergo PMP depolarization in the course of apoptosis, a fact that has been speculated to play a role in the cytoskeletal reorganization that takes place during this process [173].

The finding that hyperpolarization of the PMP determines actin compaction along the adherens junctions and increases junction stability [128] may have application in the design of therapeutic strategies. In this respect, the loss of epithelial intercellular adhesions is at the basis of diverse pathologies [174, 175]. Some of these represent major medical challenges, such as cancer progression [176179] ischemic injuries [180] and bowel inflammatory diseases [181].

7. Concluding Remarks

The modifications in the plasma membrane potential have mostly been classically associated with the physiology of excitable tissues. In both excitable and nonexcitable cells, the PMP is an energetic component of the electrochemical gradients responsible of membrane ionic transport. The findings reviewed in this work contribute to the concept that the PMP may also participate in other cellular processes, including the establishment and maintenance of the morphological and functional features of epithelial cells. In particular, we have emphasized here the possible role of the PMP in the regulation of the actin cytoskeleton. Although some knowledge about signaling pathways involved in the transduction of electrical signals at the plasma membrane to mechanical modifications of the cytoskeleton has been unraveled, the involvement of the cytoskeleton in many relevant physiological cellular phenomena permits to anticipate great progress in this respect in the near future.

Acknowledgments

This work received financial support by grants from CSIC and PEDECIBA (Universidad de la República, Uruguay), ANII (Ministerio de Educación y Cultura, Uruguay), and Comisión Honoraria de Lucha contra el Cáncer (Uruguay). The authors thank two anonymous reviewers for most fruitful suggestions.

References

  1. T. Zeuthen, “Molecular mechanisms of water transport,” in Molecular Biology Intelligence Unit, pp. 97–126, R.G. Landes Co., Austin, Tex, USA, 1996.
  2. E. Rodriquez-Boulan and W. J. Nelson, “Morphogenesis of the polarized epithelial cell phenotype,” Science, vol. 245, no. 4919, pp. 718–725, 1989. View at Scopus
  3. C. Yeaman, K. K. Grindstaff, and W. J. Nelson, “New perspectives on mechanisms involved in generating epithelial cell polarity,” Physiological Reviews, vol. 79, no. 1, pp. 73–98, 1999. View at Scopus
  4. W. J. Nelson, “Adaptation of core mechanisms to generate cell polarity,” Nature, vol. 422, no. 6933, pp. 766–774, 2003. View at Publisher · View at Google Scholar · View at Scopus
  5. D. M. Bryant and K. E. Mostov, “From cells to organs: building polarized tissue,” Nature Reviews Molecular Cell Biology, vol. 9, no. 11, pp. 887–901, 2008. View at Publisher · View at Google Scholar · View at Scopus
  6. J. Jaworski, C. C. Hoogenraad, and A. Akhmanova, “Microtubule plus-end tracking proteins in differentiated mammalian cells,” International Journal of Biochemistry and Cell Biology, vol. 40, no. 4, pp. 619–637, 2008. View at Publisher · View at Google Scholar · View at Scopus
  7. J. E. Eriksson, T. Dechat, B. Grin et al., “Introducing intermediate filaments: from discovery to disease,” Journal of Clinical Investigation, vol. 119, no. 7, pp. 1763–1771, 2009. View at Publisher · View at Google Scholar · View at Scopus
  8. K. P. Harris and U. Tepass, “Cdc42 and vesicle trafficking in polarized cells,” Traffic, vol. 11, no. 10, pp. 1272–1279, 2010. View at Publisher · View at Google Scholar · View at Scopus
  9. S. Etienne-Manneville, “From signaling pathways to microtubule dynamics: the key players,” Current Opinion in Cell Biology, vol. 22, no. 1, pp. 104–111, 2010. View at Publisher · View at Google Scholar · View at Scopus
  10. C. M. Niessen, D. Leckband, and A. S. Yap, “Tissue organization by cadherin adhesion molecules: dynamic molecular and cellular mechanisms of morphogenetic regulation,” Physiological Reviews, vol. 91, no. 2, pp. 691–731, 2011. View at Publisher · View at Google Scholar
  11. B. Baum and M. Georgiou, “Dynamics of adherens junctions in epithelial establishment, maintenance, and remodeling,” Journal of Cell Biology, vol. 192, no. 6, pp. 907–917, 2011. View at Publisher · View at Google Scholar
  12. A. L. Hartman, N. M. Sawtell, and J. L. Lessard, “Expression of actin isoforms in developing rat intestinal epithelium,” Journal of Histochemistry and Cytochemistry, vol. 37, no. 8, pp. 1225–1233, 1989. View at Scopus
  13. M. S. Balda, L. Gonzalez-Mariscal, K. Matter, M. Cereijido, and J. M. Anderson, “Assembly of the tight junction: the role of diacylglycerol,” Journal of Cell Biology, vol. 123, no. 2, pp. 293–302, 1993. View at Publisher · View at Google Scholar · View at Scopus
  14. C. J. Maples, W. G. Ruiz, and G. Apodaca, “Both microtubules and actin filaments are required for efficient postendocytotic traffic of the polymeric immunoglobulin receptor in polarized Madin-Darby canine kidney cells,” Journal of Biological Chemistry, vol. 272, no. 10, pp. 6741–6751, 1997. View at Publisher · View at Google Scholar · View at Scopus
  15. J. Breckler and B. Burnside, “Myosin-I in retinal pigment epithelial cells,” Investigative Ophthalmology and Visual Science, vol. 35, no. 5, pp. 2489–2499, 1994. View at Scopus
  16. M. J. Tyska and M. S. Mooseker, “MYO1A (brush border myosin I) dynamics in the brush border of LLC-PK1-CL4 cells,” Biophysical Journal, vol. 82, no. 4, pp. 1869–1883, 2002. View at Scopus
  17. M. C. Wagner, B. L. Blazer-Yost, J. Boyd-White, A. Srirangam, J. Pennington, and S. Bennett, “Expression of the unconventional myosin Myo1c alters sodium transport in M1 collecting duct cells,” American Journal of Physiology, Cell Physiology, vol. 289, no. 1, pp. C120–C129, 2005. View at Publisher · View at Google Scholar · View at Scopus
  18. V. Bennett and J. Healy, “Organizing the fluid membrane bilayer: diseases linked to spectrin and ankyrin,” Trends in Molecular Medicine, vol. 14, no. 1, pp. 28–36, 2008. View at Publisher · View at Google Scholar · View at Scopus
  19. K. Kizhatil and V. Bennett, “Lateral membrane biogenesis in human bronchial epithelial cells requires 190-kDa ankyrin-G,” Journal of Biological Chemistry, vol. 279, no. 16, pp. 16706–16714, 2004. View at Publisher · View at Google Scholar · View at Scopus
  20. K. Kizhatil, J. Q. Davis, L. Davis, J. Hoffman, B. L. M. Hogan, and V. Bennett, “Ankyrin-G is a molecular partner of E-cadherin in epithelial cells and early embryos,” Journal of Biological Chemistry, vol. 282, no. 36, pp. 26552–26561, 2007. View at Publisher · View at Google Scholar · View at Scopus
  21. B. Geiger, A. Bershadsky, R. Pankov, and K. M. Yamada, “Transmembrane extracellular matrix—cytoskeleton crosstalk,” Nature Reviews Molecular Cell Biology, vol. 2, no. 11, pp. 793–805, 2001. View at Publisher · View at Google Scholar · View at Scopus
  22. T. J. Keating and G. G. Borisy, “Centrosomal and non-centrosomal microtubules,” Biology of the Cell, vol. 91, no. 4-5, pp. 321–329, 1999. View at Scopus
  23. A. Musch, “Microtubule organization and function in epithelial cells,” Traffic, vol. 5, no. 1, pp. 1–9, 2004. View at Publisher · View at Google Scholar · View at Scopus
  24. R. Pepperkok, M. H. Bre, J. Davoust, and T. E. Kreis, “Microtubules are stabilized in confluent epithelial cells but not in fibroblasts,” Journal of Cell Biology, vol. 111, no. 6, pp. 3003–3012, 1990. View at Publisher · View at Google Scholar · View at Scopus
  25. E. Shelden and P. Wadsworth, “Observation and quantification of individual microtubule behavior in vivo: microtubule dynamics are cell-type specific,” Journal of Cell Biology, vol. 120, no. 4, pp. 935–945, 1993. View at Publisher · View at Google Scholar · View at Scopus
  26. C. M. Waterman-Storer, W. C. Salmon, and E. D. Salmon, “Feedback interactions between cell-cell adherens junctions and cytoskeletal dynamics in newt lung epithelial cells,” Molecular Biology of the Cell, vol. 11, no. 7, pp. 2471–2483, 2000. View at Scopus
  27. A. Reilein, S. Yamada, and W. J. Nelson, “Self-organization of an acentrosomal microtubule network at the basal cortex of polarized epithelial cells,” Journal of Cell Biology, vol. 171, no. 5, pp. 845–855, 2005. View at Publisher · View at Google Scholar · View at Scopus
  28. R. Bacallao, C. Antony, C. Dotti, E. Karsenti, E. H. K. Stelzer, and K. Simons, “The subcellular organization of Madin-Darby canine kidney cells during the formation of a polarized epithelium,” Journal of Cell Biology, vol. 109, no. 6 I, pp. 2817–2832, 1989. View at Publisher · View at Google Scholar · View at Scopus
  29. M. H. Bre, R. Pepperkok, A. M. Hill et al., “Regulation of microtubule dynamics and nucleation during polarization in MDCK II cells,” Journal of Cell Biology, vol. 111, no. 6, pp. 3013–3021, 1990. View at Publisher · View at Google Scholar · View at Scopus
  30. T. Meads and T. A. Schroer, “Polarity and nucleation of microtubules in polarized epithelial cells,” Cell Motility and the Cytoskeleton, vol. 32, no. 4, pp. 273–288, 1995. View at Publisher · View at Google Scholar · View at Scopus
  31. K. K. Grindstaff, R. L. Bacallao, and W. J. Nelson, “Apiconuclear organization of microtubules does not specify protein delivery from the trans-Golgi Network to different membrane domains in polarized epithelial cells,” Molecular Biology of the Cell, vol. 9, no. 3, pp. 685–699, 1998. View at Scopus
  32. O. A. Weisz and E. Rodriguez-Boulan, “Apical trafficking in epithelial cells: signals, clusters and motors,” Journal of Cell Science, vol. 122, no. 23, pp. 4253–4266, 2009. View at Publisher · View at Google Scholar · View at Scopus
  33. W. W. Franke, “Discovering the molecular components of intercellular junctions—a historical view,” Cold Spring Harbor Perspectives in Biology, vol. 1, no. 3, Article ID a003061, 2009. View at Publisher · View at Google Scholar · View at Scopus
  34. K. Matter and M. S. Balda, “Signalling to and from tight junctions,” Nature Reviews Molecular Cell Biology, vol. 4, no. 3, pp. 225–236, 2003. View at Publisher · View at Google Scholar · View at Scopus
  35. B. N. G. Giepmans and S. C. D. van IJzendoorn, “Epithelial cell-cell junctions and plasma membrane domains,” Biochimica et Biophysica Acta, vol. 1788, no. 4, pp. 820–831, 2009. View at Publisher · View at Google Scholar · View at Scopus
  36. M. Cavey and T. Lecuit, “Molecular bases of cell-cell junctions stability and dynamics,” Cold Spring Harbor Perspectives in Biology, vol. 1, no. 5, Article ID a002998, 2009. View at Publisher · View at Google Scholar · View at Scopus
  37. S. H. Jaffe, D. R. Friedlander, F. Matsuzaki, K. L. Crossin, B. A. Cunningham, and G. M. Edelman, “Differential effects of the cytoplasmic domains of cell adhesion molecules on cell aggregation and sorting-out,” Proceedings of the National Academy of Sciences of the United States of America, vol. 87, no. 9, pp. 3589–3593, 1990. View at Publisher · View at Google Scholar · View at Scopus
  38. C. L. Adams, W. J. Nelson, and S. J. Smith, “Quantitative analysis of cadherin-catenin-actin reorganization during development of cell-cell adhesion,” Journal of Cell Biology, vol. 135, no. 6, pp. 1899–1911, 1996. View at Publisher · View at Google Scholar · View at Scopus
  39. B. Angres, A. Barth, and W. J. Nelson, “Mechanism for transition from initial to stable cell-cell adhesion: kinetic analysis of E-cadherin-mediated adhesion using a quantitative adhesion assay,” Journal of Cell Biology, vol. 134, no. 2, pp. 549–557, 1996. View at Publisher · View at Google Scholar · View at Scopus
  40. J. Zhang, M. Betson, J. Erasmus et al., “Actin at cell-cell junctions is composed of two dynamic and functional populations,” Journal of Cell Science, vol. 118, no. 23, pp. 5549–5562, 2005. View at Publisher · View at Google Scholar · View at Scopus
  41. F. Pilot, J. M. Philippe, C. Lemmers, and T. Lecuit, “Spatial control of actin organization at adherens junctions by the synaptotagmin-like protein Btsz,” Nature, vol. 442, no. 7102, pp. 580–584, 2006. View at Publisher · View at Google Scholar · View at Scopus
  42. K. Abe and M. Takeichi, “EPLIN mediates linkage of the cadherin-catenin complex to F-actin and stabilizes the circumferential actin belt,” Proceedings of the National Academy of Sciences of the United States of America, vol. 105, no. 1, pp. 13–19, 2008. View at Publisher · View at Google Scholar · View at Scopus
  43. W. Meng, Y. Mushika, T. Ichii, and M. Takeichi, “Anchorage of microtubule minus ends to adherens junctions regulates epithelial cell-cell contacts,” Cell, vol. 135, no. 5, pp. 948–959, 2008. View at Publisher · View at Google Scholar · View at Scopus
  44. V. Vasioukhin, C. Bauer, M. Yin, and E. Fuchs, “Directed actin polymerization is the driving force for epithelial cell- cell adhesion,” Cell, vol. 100, no. 2, pp. 209–219, 2000. View at Scopus
  45. A. I. Ivanov, I. C. McCall, C. A. Parkos, and A. Nusrat, “Role for actin filament turnover and a myosin II motor in cytoskeleton-driven disassembly of the epithelial apical junctional complex,” Molecular Biology of the Cell, vol. 15, no. 6, pp. 2639–2651, 2004. View at Publisher · View at Google Scholar · View at Scopus
  46. A. I. Ivanov, I. C. McCall, B. Babbin, S. N. Samarin, A. Nusrat, and C. A. Parkos, “Microtubules regulate disassembly of epithelial apical junctions,” BMC Cell Biology, vol. 7, article no. 12, 2006. View at Publisher · View at Google Scholar · View at Scopus
  47. S. J. Stehbens, A. D. Paterson, M. S. Crampton et al., “Dynamic microtubules regulate the local concentration of E-cadherin at cell-cell contacts,” Journal of Cell Science, vol. 119, no. 9, pp. 1801–1811, 2006. View at Publisher · View at Google Scholar · View at Scopus
  48. A. I. Ivanov, C. A. Parkos, and A. Nusrat, “Cytoskeletal regulation of epithelial barrier function during inflammation,” American Journal of Pathology, vol. 177, no. 2, pp. 512–524, 2010. View at Publisher · View at Google Scholar · View at Scopus
  49. J. Nance and J. A. Zallen, “Elaborating polarity: PAR proteins and the cytoskeleton,” Development, vol. 138, no. 5, pp. 799–809, 2011. View at Publisher · View at Google Scholar
  50. C. Mazzochi, D. J. Benos, and P. R. Smith, “Interaction of epithelial ion channels with the actin-based cytoskeleton,” American Journal of Physiology, Renal Physiology, vol. 291, no. 6, pp. F1113–F1122, 2006. View at Publisher · View at Google Scholar
  51. P. R. Kiela and F. K. Ghishan, “Ion transport in the intestine,” Current Opinion in Gastroenterology, vol. 25, no. 2, pp. 87–91, 2009. View at Publisher · View at Google Scholar · View at Scopus
  52. A. J. Baines, P. M. Bennett, E. W. Carter, and C. Terracciano, “Protein 4.1 and the control of ion channels,” Blood Cells, Molecules, and Diseases, vol. 42, no. 3, pp. 211–215, 2009. View at Publisher · View at Google Scholar · View at Scopus
  53. H. Yu, Y. Zhang, L. Ye, and W. G. Jiang, “The FERM family proteins in cancer invasion and metastasis,” Frontiers in Bioscience, vol. 16, no. 4, pp. 1536–1550, 2011. View at Publisher · View at Google Scholar
  54. J. B. Zuckerman, X. Chen, J. D. Jacobs, B. Hu, T. R. Kleyman, and P. R. Smith, “Association of the epithelial sodium channel with Apx and α-spectrin in A6 renal epithelial cells,” Journal of Biological Chemistry, vol. 274, no. 33, pp. 23286–23295, 1999. View at Publisher · View at Google Scholar · View at Scopus
  55. V. Bennett and A. J. Baines, “Spectrin and ankyrin-based pathways: metazoan inventions for integrating cells into tissues,” Physiological Reviews, vol. 81, no. 3, pp. 1353–1392, 2001. View at Scopus
  56. S. A. Rajasekaran, L. G. Palmer, K. Quan et al., “Na,K-ATPase β-subunit is required for epithelial polarization, suppression of invasion, and cell motility,” Molecular Biology of the Cell, vol. 12, no. 2, pp. 279–295, 2001.
  57. S. P. Denker and D. L. Barber, “Ion transport proteins anchor and regulate the cytoskeleton,” Current Opinion in Cell Biology, vol. 14, no. 2, pp. 214–220, 2002. View at Publisher · View at Google Scholar · View at Scopus
  58. Y. Noda and S. Sasaki, “Actin-binding channels,” Progress in Brain Research, vol. 170, pp. 551–557, 2008. View at Publisher · View at Google Scholar · View at Scopus
  59. R. Padányi, Y. Xiong, G. Antalffy et al., “Apical scaffolding protein NHERF2 modulates the localization of alternatively spliced plasma membrane Ca2+ pump 2B variants in polarized epithelial cells,” Journal of Biological Chemistry, vol. 285, no. 41, pp. 31704–31712, 2010. View at Publisher · View at Google Scholar
  60. G. H. Lee, T. Ahn, D. S. Kim et al., “Bax inhibitor 1 increases cell adhesion through actin polymerization: involvement of calcium and actin binding,” Molecular and Cellular Biology, vol. 30, no. 7, pp. 1800–1813, 2010. View at Publisher · View at Google Scholar · View at Scopus
  61. C. Goswami, J. Kuhn, P. A. Heppenstall, and T. Hucho, “Importance of non-selective cation channel TRPV4 interaction with cytoskeleton and their reciprocal regulations in cultured cells,” PLoS ONE, vol. 5, no. 7, Article ID e11654, 2010. View at Publisher · View at Google Scholar · View at Scopus
  62. Z. Wang and K. L. Schey, “Aquaporin-0 interacts with the FERM domain of ezrin/radixin/moesin proteins in the ocular lens,” Investigative Ophthalmology & Visual Science, vol. 52, no. 8, pp. 5079–5087, 2011.
  63. E. M. Schwiebert, J. W. Mills, and B. A. Stanton, “Actin-based cytoskeleton regulates a chloride channel and cell volume in a renal cortical collecting duct cell line,” Journal of Biological Chemistry, vol. 269, no. 10, pp. 7081–7089, 1994. View at Scopus
  64. H. F. Cantiello, “Actin filaments stimulate the Na(+)-K(+)-ATPase,” American Journal of Physiology, vol. 269, no. 5, pp. F637–F643, 1995. View at Scopus
  65. B. K. Berdiev, A. G. Prat, H. F. Cantiello et al., “Regulation of epithelial sodium channels by short actin filaments,” Journal of Biological Chemistry, vol. 271, no. 30, pp. 17704–17710, 1996. View at Publisher · View at Google Scholar · View at Scopus
  66. S. P. Denker and D. L. Barber, “Cell migration requires both ion translocation and cytoskeletal anchoring by the Na-H exchanger NHE1,” Journal of Cell Biology, vol. 159, no. 6, pp. 1087–1096, 2002. View at Publisher · View at Google Scholar · View at Scopus
  67. B. Cha and M. Donowitz, “The epithelial brush border Na+/H+ exchanger NHE3 associates with the actin cytoskeleton by binding to ezrin directly and via PDZ domain-containing Na+/H+ exchanger regulatory factor (NHERF) proteins,” Clinical and Experimental Pharmacology and Physiology, vol. 35, no. 8, pp. 863–871, 2008. View at Publisher · View at Google Scholar · View at Scopus
  68. D. V. Ilatovskaya, T. S. Pavlov, V. Levchenko, Y. A. Negulyaev, and A. Staruschenko, “Cortical actin binding protein cortactin mediates ENaC activity via Arp2/3 complex,” FASEB Journal, vol. 25, no. 8, pp. 2688–2699, 2011. View at Publisher · View at Google Scholar
  69. S. Khurana, “Role of actin cytoskeleton in regulation of ion transport: examples from epithelial cells,” Journal of Membrane Biology, vol. 178, no. 2, pp. 73–87, 2000. View at Publisher · View at Google Scholar · View at Scopus
  70. E. A. Papakonstanti and C. Stournaras, “Cell responses regulated by early reorganization of actin cytoskeleton,” FEBS Letters, vol. 582, no. 14, pp. 2120–2127, 2008. View at Publisher · View at Google Scholar · View at Scopus
  71. B. A. Watts, T. George, and D. W. Good, “The basolateral NHE1 Na+/H+ exchanger regulates transepithelial HCO3- absorption through actin cytoskeleton remodeling in renal thick ascending limb,” Journal of Biological Chemistry, vol. 280, no. 12, pp. 11439–11447, 2005. View at Publisher · View at Google Scholar · View at Scopus
  72. N. Montalbetti, Q. Li, G. A. Timpanaro et al., “Cytoskeletal regulation of calcium-permeable cation channels in the human syncytiotrophoblast: role of gelsolin,” Journal of Physiology, vol. 566, no. 2, pp. 309–325, 2005. View at Publisher · View at Google Scholar
  73. Y. Tian, P. Kongsuphol, M. Hug et al., “Calmodulin-dependent activation of the epithelial calcium-dependent chloride channel TMEM16A,” FASEB Journal, vol. 25, no. 3, pp. 1058–1068, 2011. View at Publisher · View at Google Scholar
  74. E. K. Hoffmann and S. F. Pedersen, “Sensors and signal transduction pathways in vertebrate cell volume regulation,” Contributions to Nephrology, vol. 152, pp. 54–104, 2006. View at Publisher · View at Google Scholar · View at Scopus
  75. B. D. Matthews, D. R. Overby, R. Mannix, and D. E. Ingber, “Cellular adaptation to mechanical stress: role of integrins, Rho, cytoskeletal tension and mechanosensitive ion channels,” Journal of Cell Science, vol. 119, no. 3, pp. 508–518, 2006. View at Publisher · View at Google Scholar · View at Scopus
  76. K. Hayakawa, H. Tatsumi, and M. Sokabe, “Actin stress fibers transmit and focus force to activate mechanosensitive channels,” Journal of Cell Science, vol. 121, no. 4, pp. 496–503, 2008. View at Publisher · View at Google Scholar · View at Scopus
  77. B. C. Tilly, M. J. Edixhoven, L. G. J. Tertoolen et al., “Activation of the osmo-sensitive chloride conductance involves P21(rho) and is accompanied by a transient reorganization of the F-actin cytoskeleton,” Molecular Biology of the Cell, vol. 7, no. 9, pp. 1419–1427, 1996. View at Scopus
  78. E. K. Hoffmann, T. Schettino, and W. S. Marshall, “The role of volume-sensitive ion transport systems in regulation of epithelial transport,” Comparative Biochemistry and Physiology, A Molecular and Integrative Physiology, vol. 148, no. 1, pp. 29–43, 2007. View at Publisher · View at Google Scholar · View at Scopus
  79. F. Lang, “Mechanisms and significance of cell volume regulation,” Journal of the American College of Nutrition, vol. 26, no. 5, 2007. View at Scopus
  80. H. Cantiello, “Role of actin filament organization in CFTR activation,” Pflugers Archiv, vol. 443, no. 1, supplement 1, pp. S75–S80, 2001. View at Publisher · View at Google Scholar · View at Scopus
  81. K. R. Schiller, P. J. Maniak, and S. M. O'Grady, “Cystic fibrosis transmembrane conductance regulator is involved in airway epithelial wound repair,” American Journal of Physiology, Cell Physiology, vol. 299, no. 5, pp. C912–C921, 2010. View at Publisher · View at Google Scholar · View at Scopus
  82. M. Levite, L. Cahalon, A. Peretz et al., “Extracellular K+ and opening of voltage-gated potassium channels activate T cell integrin function: physical and functional association between Kv1.3 channels and β1 integrins,” Journal of Experimental Medicine, vol. 191, no. 7, pp. 1167–1176, 2000. View at Publisher · View at Google Scholar · View at Scopus
  83. K. Clark, M. Langeslag, B. Van Leeuwen et al., “TRPM7, a novel regulator of actomyosin contractility and cell adhesion,” EMBO Journal, vol. 25, no. 2, pp. 290–301, 2006. View at Publisher · View at Google Scholar · View at Scopus
  84. J. F. Wei, L. Wei, X. Zhou et al., “Formation of Kv2.1-FAK complex as a mechanism of FAK activation, cell polarization and enhanced motility,” Journal of Cellular Physiology, vol. 217, no. 2, pp. 544–557, 2008. View at Publisher · View at Google Scholar · View at Scopus
  85. S. Chifflet, J. A. Hernández, and S. Grasso, “A possible role for membrane depolarization in epithelial wound healing,” American Journal of Physiology, Cell Physiology, vol. 288, no. 6, pp. C1420–C1430, 2005. View at Publisher · View at Google Scholar · View at Scopus
  86. S. del Mónaco, Y. Assef, and B. A. Kotsias, “Epithelial sodium channel in a human trophoblast cell line (BeWo),” Journal of Membrane Biology, vol. 223, no. 3, pp. 127–139, 2008. View at Publisher · View at Google Scholar · View at Scopus
  87. S. C. Grifoni, K. P. Gannon, D. E. Stec, and H. A. Drummond, “ENaC proteins contribute to VSMC migration,” American Journal of Physiology, Heart and Circulatory Physiology, vol. 291, no. 6, pp. H3076–H3086, 2006. View at Publisher · View at Google Scholar · View at Scopus
  88. D. Rotin, D. Bar-Sagi, H. O'Brodovich et al., “An SH3 binding region in the epithelial Na+ channel (αrENaC) mediates its localization at the apical membrane,” EMBO Journal, vol. 13, no. 19, pp. 4440–4450, 1994.
  89. C. Mazzochi, J. K. Bubien, P. R. Smith, and D. J. Benos, “The carboxyl terminus of the α-subunit of the amiloride-sensitive epithelial sodium channel binds to F-actin,” Journal of Biological Chemistry, vol. 281, no. 10, pp. 6528–6538, 2006. View at Publisher · View at Google Scholar
  90. H. F. Cantiello, J. L. Stow, A. G. Prat, and D. A. Ausiello, “Actin filaments regulate epithelial Na+ channel activity,” American Journal of Physiology, vol. 261, no. 5, pp. C882–C888, 1991. View at Scopus
  91. B. K. Berdiev, R. Latorre, D. J. Benos, and I. I. Ismailov, “Actin modifies Ca2+ block of epithelial Na+ channels in planar lipid bilayers,” Biophysical Journal, vol. 80, no. 5, pp. 2176–2186, 2001. View at Scopus
  92. S. J. Copeland, B. K. Berdiev, H. L. Ji et al., “Regions in the carboxy terminus of α-bENaC involved in gating and functional effects of actin,” American Journal of Physiology, Cell Physiology, vol. 281, no. 1, pp. C231–C240, 2001. View at Scopus
  93. D. L. Cioffi, S. Wu, M. Alexeyev, S. R. Goodman, M. X. Zhu, and T. Stevens, “Activation of the endothelial store-operated ISOC Ca2+ channel requires interaction of protein 4.1 with TRPC4,” Circulation Research, vol. 97, no. 11, pp. 1164–1172, 2005. View at Publisher · View at Google Scholar · View at Scopus
  94. W. J. Nelson and P. J. Veshnock, “Ankyrin binding to (Na++K+)ATPase and implications for the organization of membrane domains in polarized cells,” Nature, vol. 328, no. 6130, pp. 533–536, 1987. View at Scopus
  95. R. G. Contreras, L. Shoshani, C. Flores-Maldonado, A. Lázaro, and M. Cereijido, “Relationship between Na(+),K(+)-ATPase and cell attachment,” Journal of Cell Science, vol. 112, no. 23, pp. 4223–4232, 1999. View at Scopus
  96. S. A. Rajasekaran, L. G. Palmer, S. Y. Moon et al., “Na,K-ATPase activity is required for formation of tight junctions, desmosomes, and induction of polarity in epithelial cells,” Molecular Biology of the Cell, vol. 12, no. 12, pp. 3717–3732, 2001.
  97. I. Larre, A. Lazaro, R. G. Contreras et al., “Ouabain modulates epithelial cell tight junction,” Proceedings of the National Academy of Sciences of the United States of America, vol. 107, no. 25, pp. 11387–11392, 2010. View at Publisher · View at Google Scholar · View at Scopus
  98. W. Schoner and G. Scheiner-Bobis, “Endogenous and exogenous cardiac glycosides: their roles in hypertension, salt metabolism, and cell growth,” American Journal of Physiology, Cell Physiology, vol. 293, no. 2, pp. C509–C536, 2007. View at Publisher · View at Google Scholar · View at Scopus
  99. V. Bennett and J. Healy, “Membrane domains based on ankyrin and spectrin associated with cell-cell interactions,” Cold Spring Harbor Perspectives in Biology, vol. 1, no. 6, Article ID a003012, 2009. View at Publisher · View at Google Scholar · View at Scopus
  100. P. R. Stabach, P. Devarajan, M. C. Stankewich, S. Bannykh, and J. S. Morrow, “Ankyrin facilitates intracellular trafficking of α1- Na+-K+-ATPase in polarized cells,” American Journal of Physiology, Cell Physiology, vol. 295, no. 5, pp. C1202–C1214, 2008. View at Publisher · View at Google Scholar
  101. R. R. Dubreuil, P. Wang, S. Dahl, J. Lee, and L. S. B. Goldstein, “Drosophila β spectrin functions independently of α spectrin to polarize the Na,K ATPase in epithelial cells,” Journal of Cell Biology, vol. 149, no. 3, pp. 647–656, 2000. View at Publisher · View at Google Scholar · View at Scopus
  102. A. Das, C. Base, S. Dhulipala, and R. R. Dubreuil, “Spectrin functions upstream of ankyrin in a spectrin cytoskeleton assembly pathway,” Journal of Cell Biology, vol. 175, no. 2, pp. 325–335, 2006. View at Publisher · View at Google Scholar · View at Scopus
  103. V. Koefoed-Johnsen and H. H. Ussing, “The nature of the frog skin potential,” Acta physiologica Scandinavica, vol. 42, no. 3-4, pp. 298–308, 1958. View at Scopus
  104. S. G. Schultz, “A century of (epithelial) transport physiology: from vitalism to molecular cloning,” American Journal of Physiology, vol. 274, no. 1, pp. C13–C23, 1998. View at Scopus
  105. L. G. Palmer and O. S. Andersen, “The two-membrane model of epithelial transport: Koefoed-Johnsen and ussing (1958),” Journal of General Physiology, vol. 132, no. 6, pp. 607–612, 2008. View at Scopus
  106. L. Reuss, “Ussing's two-membrane hypothesis: the model and half a century of progress,” Journal of Membrane Biology, vol. 184, no. 3, pp. 211–217, 2001. View at Publisher · View at Google Scholar · View at Scopus
  107. E. Frömter and J. Diamond, “Route of passive ion permeation in epithelia,” Nature: New biology, vol. 235, no. 53, pp. 9–13, 1972. View at Scopus
  108. N. K. Wills, L. Reuss, and S. A. Lewis, Epithelial Transport : A Guide to Methods and Experimental Analysis, Chapman & Hall, London, UK, 1996.
  109. J. A. Hernández and S. Chifflet, “Electrogenic properties of the sodium pump in a dynamic model of membrane transport,” Journal of Membrane Biology, vol. 176, no. 1, pp. 41–52, 2000. View at Publisher · View at Google Scholar · View at Scopus
  110. E. M. Wright, D. D.F. LOO, and B. A. Hirayama, “Biology of human sodium glucose transporters,” Physiological Reviews, vol. 91, no. 2, pp. 733–794, 2011. View at Publisher · View at Google Scholar
  111. D. I. Cook and J. A. Young, “Effect of K+ channels in the apical plasma membrane on epithelial secretion based on secondary active Cl- transport,” Journal of Membrane Biology, vol. 110, no. 2, pp. 139–146, 1989. View at Scopus
  112. J. E. Melvin, D. Yule, T. Shuttleworth, and T. Begenisich, “Regulation of fluid and electrolyte secretion in salivary gland acinar cells,” Annual Review of Physiology, vol. 67, pp. 445–469, 2005. View at Publisher · View at Google Scholar · View at Scopus
  113. L. Palk, J. Sneyd, T. J. Shuttleworth, D. I. Yule, and E. J. Crampin, “A dynamic model of saliva secretion,” Journal of Theoretical Biology, vol. 266, no. 4, pp. 625–640, 2010. View at Publisher · View at Google Scholar · View at Scopus
  114. S. Futakuchi, H. Ishiguro, S. Naruse et al., “High glucose inhibits HCO3(-) and fluid secretion in rat pancreatic ducts,” Pflugers Archiv, vol. 459, no. 1, pp. 215–226, 2009. View at Publisher · View at Google Scholar · View at Scopus
  115. P. Preston, L. Wartosch, D. Günzel et al., “Disruption of the K+ channel β-subunit KCNE3 reveals an important role in intestinal and tracheal Cl- transport,” Journal of Biological Chemistry, vol. 285, no. 10, pp. 7165–7175, 2010. View at Publisher · View at Google Scholar
  116. J. Fischbarg and F. P. J. Diecke, “A mathematical model of electrolyte and fluid transport across corneal endothelium,” Journal of Membrane Biology, vol. 203, no. 1, pp. 41–56, 2005. View at Publisher · View at Google Scholar · View at Scopus
  117. N. Montalbetti and J. Fischbarg, “Frequency spectrum of transepithelial potential difference reveals transport-related oscillations,” Biophysical Journal, vol. 97, no. 6, pp. 1530–1537, 2009. View at Publisher · View at Google Scholar · View at Scopus
  118. F. Jorge, “Fluid transport across leaky epithelia: central role of the tight junction and supporting role of aquaporins,” Physiological Reviews, vol. 90, no. 4, pp. 1271–1290, 2010. View at Publisher · View at Google Scholar · View at Scopus
  119. D. Sas, M. Hu, O. W. Moe, and M. Baum, “Effect of claudins 6 and 9 on paracellular permeability in MDCK II cells,” American Journal of Physiology , Regulatory Integrative and Comparative Physiology, vol. 295, no. 5, pp. R1713–R1719, 2008. View at Publisher · View at Google Scholar
  120. M. Avella, O. Ducoudret, D. F. Pisani, and P. Poujeol, “Swelling-activated transport of taurine in cultured gill cells of sea bass: physiological adaptation and pavement cell plasticity,” American Journal of Physiology, Regulatory Integrative and Comparative Physiology, vol. 296, no. 4, pp. R1149–R1160, 2009. View at Publisher · View at Google Scholar · View at Scopus
  121. R. Montesano, H. Ghzili, F. Carrozzino, B. C. Rossier, and E. Feraille, “cAMP-dependent chloride secretion mediates tubule enlargement and cyst formation by cultured mammalian collecting duct cells,” American Journal of Physiology, Renal Physiology, vol. 296, no. 2, pp. F446–F457, 2009. View at Publisher · View at Google Scholar
  122. S. Chifflet and J. A. Hernandez, “Use of standard fluorescence microscopy to assess modifications in the plasma membrane potential and in the intracellular concentration of inorganic ions in cultured cells,” in Modern Research and Educational Topics in Microscopy, A. M. Vilas and J. D. Alvarez, Eds., vol. 1, pp. 229–236, Formatex, 2007.
  123. M. Olivotto, A. Arcangeli, M. Carlà, and E. Wanke, “Electric fields at the plasma membrane level: a neglected element in the mechanisms of cell signalling,” BioEssays, vol. 18, no. 6, pp. 495–504, 1996. View at Scopus
  124. J. A. Hernández and E. Cristina, “Modeling cell volume regulation in nonexcitable cells: the roles of the Na+ pump and of cotransport systems,” American Journal of Physiology - Cell Physiology, vol. 275, no. 4, pp. C1067–C1080, 1998. View at Scopus
  125. F. Lang, G. L. Busch, M. Ritter et al., “Functional significance of cell volume regulatory mechanisms,” Physiological Reviews, vol. 78, no. 1, pp. 247–306, 1998.
  126. J. Vaaraniemi, V. Huotari, V. P. Lehto, and S. Eskelinen, “Effect of PMA on the integrity of the membrane skeleton and morphology of epithelial MDCK cells is dependent on the activity of amiloride-sensitive ion transporters and membrane potential,” European Journal of Cell Biology, vol. 74, no. 3, pp. 262–272, 1997. View at Scopus
  127. S. Chifflet, J. A. Hernández, S. Grasso, and A. Cirillo, “Nonspecific depolarization of the plasma membrane potential induces cytoskeletal modifications of bovine corneal endothelial cells in culture,” Experimental Cell Research, vol. 282, no. 1, pp. 1–13, 2003. View at Publisher · View at Google Scholar · View at Scopus
  128. V. Nin, J. A. Hernández, and S. Chifflet, “Hyperpolarization of the plasma membrane potential provokes reorganization of the actin cytoskeleton and increases the stability of adherens junctions in bovine corneal endothelial cells in culture,” Cell Motility and the Cytoskeleton, vol. 66, no. 12, pp. 1087–1099, 2009. View at Publisher · View at Google Scholar · View at Scopus
  129. S. Chifflet, V. Correa, V. Nin, C. Justet, and J. A. Hernández, “Effect of membrane potential depolarization on the organization of the actin cytoskeleton of eye epithelia. The role of adherens junctions,” Experimental Eye Research, vol. 79, no. 6, pp. 769–777, 2004. View at Publisher · View at Google Scholar · View at Scopus
  130. K. Szászi, G. Sirokmány, C. di Ciano-Oliveira, O. D. Rotstein, and A. Kapus, “Depolarization induces Rho-Rho kinase-mediated myosin light chain phosphorylation in kidney tubular cells,” American Journal of Physiology, Cell Physiology, vol. 289, pp. C673–C685, 2005. View at Publisher · View at Google Scholar
  131. F. Waheed, P. Speight, G. Kawai, Q. Dan, A. Kapus, and K. Szászi, “Extracellular signal-regulated kinase and GEF-H1 mediate depolarization-induced Rho activation and paracellular permeability increase,” American Journal of Physiology, Cell Physiology, vol. 298, no. 6, pp. C1376–C1387, 2010. View at Publisher · View at Google Scholar
  132. H. Oberleithner, C. Callies, K. Kusche-Vihrog et al., “Potassium softens vascular endothelium and increases nitric oxide release,” Proceedings of the National Academy of Sciences of the United States of America, vol. 106, no. 8, pp. 2829–2834, 2009. View at Publisher · View at Google Scholar · View at Scopus
  133. C. Callies, J. Fels, I. Liashkovich et al., “Membrane potential depolarization decreases the stiffness of vascular endothelial cells,” Journal of Cell Science, vol. 124, no. 11, pp. 1936–1942, 2011. View at Publisher · View at Google Scholar
  134. S. S. Grewal, A. M. Horgan, R. D. York, G. S. Withers, G. A. Banker, and P. J. S. Stork, “Neuronal calcium activates a Rap1 and B-Raf signaling pathway via the cyclic adenosine monophosphate-dependent protein kinase,” Journal of Biological Chemistry, vol. 275, no. 5, pp. 3722–3728, 2000. View at Publisher · View at Google Scholar · View at Scopus
  135. M. Ishii, A. Inanobe, S. Fujita, Y. Makino, Y. Hosoya, and Y. Kurachi, “Ca2+ elevation evoked by membrane depolarization regulates G protein cycle via RGS proteins in the heart,” Circulation Research, vol. 89, no. 11, pp. 1045–1050, 2001. View at Scopus
  136. M. Mita, H. Yanagihara, S. Hishinuma, M. Saito, and M. P. Walsh, “Membrane depolarization-induced contraction of rat caudal arterial smooth muscle involves Rho-associated kinase,” Biochemical Journal, vol. 364, no. 2, pp. 431–440, 2002. View at Publisher · View at Google Scholar · View at Scopus
  137. S. Sakurada, N. Takuwa, N. Sugimoto et al., “Ca2+-dependent activation of Rho and Rho kinase in membrane depolarization-induced and receptor stimulation-induced vascular smooth muscle contraction,” Circulation Research, vol. 93, no. 6, pp. 548–556, 2003. View at Publisher · View at Google Scholar · View at Scopus
  138. C. Liu, J. Zuo, E. Pertens, P. B. Helli, and L. J. Janssen, “Regulation of Rho/ROCK signaling in airway smooth muscle by membrane potential and [Ca2+]i,” American Journal of Physiology, Lung Cellular and Molecular Physiology, vol. 289, no. 4, pp. L574–L582, 2005. View at Publisher · View at Google Scholar · View at Scopus
  139. M. H. Roos, W. F. Van Rodijnen, A. A. Van Lambalgen, P. M. Ter Wee, and G. J. Tangelder, “Renal microvascular constriction to membrane depolarization and other stimuli: pivotal role for rho-kinase,” Pflugers Archiv, vol. 452, no. 4, pp. 471–477, 2006. View at Publisher · View at Google Scholar · View at Scopus
  140. M. Fernández-Tenorio, C. Porras-González, A. Castellano, A. Del Valle-Rodríguez, J. López-Barneo, and J. Ureña, “Metabotropic regulation of RhoA/Rho-associated kinase by l-type Ca2+ Channels: new mechanism for depolarization-Evoked mammalian arterial contraction,” Circulation Research, vol. 108, no. 11, pp. 1348–1357, 2011. View at Publisher · View at Google Scholar
  141. C. X. Bai, S. Kim, A. J. Streets, A. C. M. Ong, and L. Tsiokas, “Activation of TRPP2 through mDia1-dependent voltage gating,” EMBO Journal, vol. 27, no. 9, pp. 1345–1356, 2008. View at Publisher · View at Google Scholar · View at Scopus
  142. Y. Murata and Y. Okamura, “Depolarization activates the phosphoinositide phosphatase Ci-VSP, as detected in Xenopus oocytes coexpressing sensors of PIP2,” Journal of Physiology, vol. 583, no. 3, pp. 875–889, 2007. View at Publisher · View at Google Scholar · View at Scopus
  143. H. Neuhaus and T. Hollemann, “Kidney specific expression of cTPTE during development of the chick embryo,” Gene Expression Patterns, vol. 9, no. 8, pp. 568–571, 2009. View at Publisher · View at Google Scholar · View at Scopus
  144. M. Ogasawara, M. Sasaki, N. Nakazawa, A. Nishino, and Y. Okamura, “Gene expression profile of Ci-VSP in juveniles and adult blood cells of ascidian,” Gene Expression Patterns, vol. 11, pp. 233–238, 2011. View at Publisher · View at Google Scholar · View at Scopus
  145. Y. Okamura and J. E. Dixon, “Voltage-sensing phosphatase: its molecular relationship with PTEN,” Physiology, vol. 26, no. 1, pp. 6–13, 2011. View at Publisher · View at Google Scholar
  146. R. P. Bhattacharyya, A. Reményi, B. J. Yeh, and W. A. Lim, “Domains, motifs, and scaffolds: the role of modular interactions in the evolution and wiring of cell signaling circuits,” Annual Review of Biochemistry, vol. 75, pp. 655–680, 2006. View at Publisher · View at Google Scholar · View at Scopus
  147. T. Takenawa, “Phosphoinositide-binding interface proteins involved in shaping cell membranes,” Proceedings of the Japan Academy Series B, vol. 86, no. 5, pp. 509–523, 2010. View at Publisher · View at Google Scholar · View at Scopus
  148. X. Zhang, X. Chen, C. Jia, X. Geng, X. Du, and H. Zhang, “Depolarization increases phosphatidylinositol (PI) 4,5-bisphosphate level and KCNQ currents through PI 4-kinase mechanisms,” Journal of Biological Chemistry, vol. 285, no. 13, pp. 9402–9409, 2010. View at Publisher · View at Google Scholar · View at Scopus
  149. M. Cotton and A. Claing, “G protein-coupled receptors stimulation and the control of cell migration,” Cellular Signalling, vol. 21, no. 7, pp. 1045–1053, 2009. View at Publisher · View at Google Scholar · View at Scopus
  150. S. H. Lee and R. Dominguez, “Regulation of actin cytoskeleton dynamics in cells.,” Molecules and cells, vol. 29, no. 4, pp. 311–325, 2010. View at Publisher · View at Google Scholar · View at Scopus
  151. M. Cohen-Armon and M. Sokolovsky, “Evidence for involvement of the voltage-dependent Na+ channel gating in depolarization-induced activation of G-proteins,” Journal of Biological Chemistry, vol. 268, no. 13, pp. 9824–9838, 1993. View at Scopus
  152. Y. Ben-Chaim, O. Tour, N. Dascal, I. Parnas, and H. Parnas, “The M2 muscarinic G-protein-coupled receptor is voltage-sensitive,” Journal of Biological Chemistry, vol. 278, no. 25, pp. 22482–22491, 2003. View at Publisher · View at Google Scholar · View at Scopus
  153. J. Martinez-Pinna, I. S. Gurung, C. Vial et al., “Direct voltage control of signaling via P2Y1 and other Gαq-coupled receptors,” Journal of Biological Chemistry, vol. 280, no. 2, pp. 1490–1498, 2005. View at Publisher · View at Google Scholar · View at Scopus
  154. Y. Ben-Chaim, B. Chanda, N. Dascal, F. Bezanilla, I. Parnas, and H. Parnas, “Movement of 'gating charge' is coupled to ligand binding in a G-protein-coupled receptor,” Nature, vol. 444, no. 7115, pp. 106–109, 2006. View at Publisher · View at Google Scholar · View at Scopus
  155. L. Ohana, O. Barchad, I. Parnas, and H. Parnas, “The metabotropic glutamate G-protein-coupled receptors mGluR3 and mGluR1a are voltage-sensitive,” Journal of Biological Chemistry, vol. 281, no. 34, pp. 24204–24215, 2006. View at Publisher · View at Google Scholar · View at Scopus
  156. K. Sahlholm, D. Marcellino, J. Nilsson, K. Fuxe, and P. Arhem, “Voltage-sensitivity at the human dopamine D2S receptor is agonist-specific,” Biochemical and Biophysical Research Communications, vol. 377, no. 4, pp. 1216–1221, 2008. View at Publisher · View at Google Scholar
  157. S. Genet, R. Costalat, and J. Burger, “A few comments on electrostatic interactions in cell physiology,” Acta Biotheoretica, vol. 48, no. 3-4, pp. 273–287, 2000. View at Publisher · View at Google Scholar · View at Scopus
  158. T. Yeung and S. Grinstein, “Lipid signaling and the modulation of surface charge during phagocytosis,” Immunological Reviews, vol. 219, no. 1, pp. 17–36, 2007. View at Publisher · View at Google Scholar · View at Scopus
  159. U. Dietrich, P. Kruger, and J. A. Kas, “Structural investigation on the adsorption of the MARCKS peptide on anionic lipid monolayers—effects beyond electrostatic,” Chemistry and Physics of Lipids, vol. 164, no. 4, pp. 266–275, 2011. View at Publisher · View at Google Scholar
  160. R. E. Redfern, D. Redfern, M. L. M. Furgason, M. Munson, A. H. Ross, and A. Gericke, “PTEN phosphatase selectively binds phosphoinositides and undergoes structural changes,” Biochemistry, vol. 47, no. 7, pp. 2162–2171, 2008. View at Publisher · View at Google Scholar · View at Scopus
  161. J. F. Hancock, H. Paterson, and C. J. Marshall, “A polybasic domain or palmitoylation is required in addition to the CAAX motif to localize p21(ras) to the plasma membrane,” Cell, vol. 63, no. 1, pp. 133–139, 1990. View at Publisher · View at Google Scholar · View at Scopus
  162. C. T. Sigal, W. Zhou, C. A. Buser, S. McLaughlin, and M. D. Resh, “Amino-terminal basic residues of Src mediate membrane binding through electrostatic interaction with acidic phospholipids,” Proceedings of the National Academy of Sciences of the United States of America, vol. 91, no. 25, pp. 12253–12257, 1994. View at Publisher · View at Google Scholar · View at Scopus
  163. M. A. O. Magalhaes and M. Glogauer, “Pivotal advance: phospholipids determine net membrane surface charge resulting in differential localization of active Rac1 and Rac2,” Journal of Leukocyte Biology, vol. 87, no. 4, pp. 545–555, 2010. View at Publisher · View at Google Scholar · View at Scopus
  164. K. Hamada, T. Shimizu, T. Matsui, S. Tsukita, S. Tsukita, and T. Hakoshima, “Structural basis of the membrane-targeting and unmasking mechanisms of the radixin FERM domain,” EMBO Journal, vol. 19, no. 17, pp. 4449–4462, 2000. View at Scopus
  165. G. A. Gomez and J. L. Daniotti, “Electrical properties of plasma membrane modulate subcellular distribution of K-Ras,” FEBS Journal, vol. 274, no. 9, pp. 2210–2228, 2007. View at Publisher · View at Google Scholar · View at Scopus
  166. A. Arcangeli and A. Becchetti, “Complex functional interaction between integrin receptors and ion channels,” Trends in Cell Biology, vol. 16, no. 12, pp. 631–639, 2006. View at Publisher · View at Google Scholar · View at Scopus
  167. J. A. Tuszyński, S. Portet, J. M. Dixon, C. Luxford, and H. F. Cantiello, “Ionic wave propagation along actin filaments,” Biophysical Journal, vol. 86, no. 4, pp. 1890–1903, 2004.
  168. M. A. Watsky, “Nonselective cation channel activation during wound healing in the corneal endothelium,” American Journal of Physiology, Cell Physiology, vol. 268, no. 5, pp. C1179–C1185, 1995. View at Scopus
  169. V. E. Franklin-Tong and C. W. Gourlay, “A role for actin in regulating apoptosis/programmed cell death: evidence spanning yeast, plants and animals,” Biochemical Journal, vol. 413, no. 3, pp. 389–404, 2008. View at Publisher · View at Google Scholar · View at Scopus
  170. J. Y. Rao, Y. S. Jin, Q. Zheng, J. Cheng, J. Tai, and G. P. Hemstreet, “Alterations of the actin polymerization status as an apoptotic morphological effector in HL-60 cells,” Journal of Cellular Biochemistry, vol. 75, no. 4, pp. 686–697, 1999. View at Publisher · View at Google Scholar · View at Scopus
  171. D. P. Cioca and K. Kitano, “Induction of apoptosis and CD10/neutral endopeptidase expression by jaspamide in HL-60 line cells,” Cellular and Molecular Life Sciences, vol. 59, no. 8, pp. 1377–1387, 2002. View at Publisher · View at Google Scholar · View at Scopus
  172. H. Suria, L. A. Chau, E. Negrou, D. J. Kelvin, and J. Madrenas, “Cytoskeletal disruption induces T cell apoptosis by a caspase-3 mediated mechanism,” Life Sciences, vol. 65, no. 25, pp. 2697–2707, 1999. View at Scopus
  173. R. Franco, C. D. Bortner, and J. A. Cidlowski, “Potential roles of electrogenic ion transport and plasma membrane depolarization in apoptosis,” Journal of Membrane Biology, vol. 209, no. 1, pp. 43–58, 2006. View at Publisher · View at Google Scholar · View at Scopus
  174. E. M. Fish and B. A. Molitoris, “Alterations in epithelial polarity and the pathogenesis of disease states,” New England Journal of Medicine, vol. 330, no. 22, pp. 1580–1588, 1994. View at Publisher · View at Google Scholar · View at Scopus
  175. M. Cereijido, R. G. Contreras, D. Flores-Benítez et al., “New diseases derived or associated with the tight junction,” Archives of Medical Research, vol. 38, no. 5, pp. 465–478, 2007. View at Publisher · View at Google Scholar · View at Scopus
  176. P. Cowin, T. M. Rowlands, and S. J. Hatsell, “Cadherins and catenins in breast cancer,” Current Opinion in Cell Biology, vol. 17, no. 5, pp. 499–508, 2005. View at Publisher · View at Google Scholar · View at Scopus
  177. C. Salon, S. Lantuejoul, B. Eymin, S. Gazzeri, C. Brambilla, and E. Brambilla, “The E-cadherin-beta-catenin complex and its implication in lung cancer progression and prognosis,” Future Oncology, vol. 1, no. 5, pp. 649–660, 2005. View at Publisher · View at Google Scholar · View at Scopus
  178. N. Bonitsis, A. Batistatou, S. Karantima, and K. Charalabopoulos, “The role of cadherin/catenin complex in malignant melanoma,” Experimental Oncology, vol. 28, no. 3, pp. 187–193, 2006. View at Scopus
  179. U. Cavallaro, S. Liebner, and E. Dejana, “Endothelial cadherins and tumor angiogenesis,” Experimental Cell Research, vol. 312, no. 5, pp. 659–667, 2006. View at Publisher · View at Google Scholar · View at Scopus
  180. T. A. Sutton, H. E. Mang, S. B. Campos, R. M. Sandoval, M. C. Yoder, and B. A. Molitoris, “Injury of the renal microvascular endothelium alters barrier function after ischemia,” American Journal of Physiology, Renal Physiology, vol. 285, no. 2, pp. F191–F198, 2003. View at Scopus
  181. M. G. Laukoetter, P. Nava, and A. Nusrat, “Role of the intestinal barrier in inflammatory bowel disease,” World Journal of Gastroenterology, vol. 14, no. 3, pp. 401–407, 2008. View at Publisher · View at Google Scholar · View at Scopus