Depolarization induces Rho-Rho kinase-mediated myosin light chain phosphorylation in kidney tubular cells

Katalin Szászi,* Gábor Sirokmány,* Caterina Di Ciano-Oliveira, Ori D. Rotstein, and András Kapus

Department of Surgery, The Toronto General Hospital and University Health Network, Toronto, Ontario, Canada

Submitted 28 September 2004 ; accepted in final form 23 April 2005


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
Myosin-based contractility plays important roles in the regulation of epithelial functions, particularly paracellular permeability. However, the triggering factors and the signaling pathways that control epithelial myosin light chain (MLC) phosphorylation have not been elucidated. Herein we show that plasma membrane depolarization provoked by distinct means, including high extracellular K+, the lipophilic cation tetraphenylphosphonium, or the ionophore nystatin, induced strong diphosphorylation of MLC in kidney epithelial cells. In sharp contrast to smooth muscle, depolarization of epithelial cells did not provoke a Ca2+ signal, and removal of external Ca2+ promoted rather than inhibited MLC phosphorylation. Moreover, elevation of intracellular Ca2+ did not induce significant MLC phosphorylation, and the myosin light chain kinase (MLCK) inhibitor ML-7 did not prevent the depolarization-induced MLC response, suggesting that MLCK is not a regulated element in this process. Instead, the Rho-Rho kinase (ROK) pathway is the key mediator because 1) depolarization stimulated Rho and induced its peripheral translocation, 2) inhibition of Rho by Clostridium difficile toxin B or C3 transferase abolished MLC phosphorylation, and 3) the ROK inhibitor Y-27632 suppressed the effect. Importantly, physiological depolarizing stimuli were able to activate the same pathway: L-alanine, the substrate of the electrogenic Na+-alanine cotransporter, stimulated Rho and induced Y-27632-sensitive MLC phosphorylation in a Na+-dependent manner. Together, our results define a novel mode of the regulation of MLC phosphorylation in epithelial cells, which is depolarization triggered and Rho-ROK-mediated but Ca2+ signal independent. This pathway may be a central mechanism whereby electrogenic transmembrane transport processes control myosin phosphorylation and thereby regulate paracellular transport.

membrane potential; Na+-alanine cotransport; epithelium; phosphatidylinositol 3-kinase; LLC-PK1 cells


MYOSIN II-DEPENDENT CELL CONTRACTILITY plays a key role in a variety of cellular functions, including muscle constriction, spreading, motility, resistance to mechanical stress, and the regulation of endothelial and epithelial permeability (19, 27, 30, 47, 51, 54, 60). The regulation of myosin II phosphorylation has been explored extensively in muscle cells, while comparatively much less is known about the underlying mechanisms in "nonexcitable" tissues, particularly epithelia. In smooth muscle cells, electromechanical coupling is the classic mechanism that accounts for the initial activation of myosin-based contractility. During this process, depolarization of the plasma membrane opens voltage-gated Ca2+ channels, leading to elevation of the cytosolic Ca2+ concentration ([Ca2+]i), which in turn activates the Ca2+-calmodulin-dependent myosin light chain kinase (MLCK). MLCK then mono- or diphosphorylates the regulatory myosin light chain (MLC) at Ser19 and Thr18, which results in enhanced actin-myosin interaction and increased myosin ATPase activity (for review, see Ref. 51). While this mechanism is responsible for the phasic (peak) component of smooth muscle contraction, the tonic (sustained) phase of contraction, during which substantial force generation can be maintained at only slightly elevated Ca2+ levels, is mediated by a different process (22) that has been reviewed previously (18). This phenomenon, often referred to as Ca2+ sensitization, involves the activation of the small GTPase Rho and its downstream effector Rho kinase (ROK), which can promote contractility by two mechanisms (41, 45, 46, 50). First, ROK phosphorylates and thereby inactivates MLC phosphatase, an effect that leads to enhanced steady-state MLC phosphorylation by inhibiting dephosphorylation (29, 32). Second, ROK was shown to be able to phosphorylate MLC directly (1, 34, 57) on Ser19 and Thr18, although this effect is thought to have less physiological significance in smooth muscle. Interestingly, in cases of depolarization-induced contraction, both the peak and the sustained phases appear to depend entirely on Ca2+ influx, because both phases are abolished by Ca2+ channel blockers or Ca2+ chelators (41, 46). These findings have led to the notion that membrane depolarization triggers Rho-ROK activation via a Ca2+ signal. Indeed, the observations that in smooth muscle cells a rise in [Ca2+]i can activate Rho (46) and may promote membrane translocation of ROK (58) lend strong experimental support to this concept.

In addition to electromechanical coupling, the other major mode to turn on the myosin-based contractile apparatus is pharmacomechanical coupling (52). During this process, the binding of various neurotransmitters or other mediators to their cell surface receptors provokes voltage-independent MLC phosphorylation. Many of these agonists elicit Ca2+ signals; however, there is strong evidence that they also stimulate the Rho-ROK pathway through partially or entirely Ca2+-independent mechanisms. For example, engagement of certain serpentine receptors activates the {alpha}-subunits of distinct trimeric G proteins (G{alpha}12/13, G{alpha}q), which in turn stimulate Rho-specific guanine nucleotide exchange factors (GEFs) (9, 10, 20). Moreover, a recent study has shown that in fibroblasts lacking MLCK, thrombin or lysophosphatidic acid induces ROK-mediated and Ca2+-independent cell contraction (12).

Cell contractility is thought to play a key role in epithelial physiology as a major regulator of epithelial tissue remodeling (13) and paracellular transport (4, 21, 37, 54, 56). Despite its recognized importance, the regulation of MLC phosphorylation in epithelial cells is poorly characterized. Specifically, the role of membrane depolarization, the Ca2+ dependence of the process, and the relative involvement of MLCK and/or the Rho-ROK pathway have not been elucidated. A possible membrane potential-dependent regulation of MLC in this so-called nonexcitable tissue context may be of particular interest, because epithelia contain a number of electrogenic transporters such as the Na+-amino acid cotransporters (5). Furthermore, it is well known that transmembrane transport processes regulate paracellular transport, but the underlying mechanism remains elusive (55). We considered that depolarization-regulated MLC phosphorylation might serve as a link between electrogenic transmembrane and paracellular transport.

In our previous studies, we found that hyperosmotic stress induces Rho-ROK-mediated MLC phosphorylation in kidney proximal tubular (LLC-PK1) cells (11). Curiously, during the course of these experiments, we noted that certain maneuvers that were expected to depolarize the plasma membrane also led to MLC phosphorylation. This initial finding prompted us to characterize this effect and explore the underlying mechanisms. Our present studies reveal a novel mode of Rho activation and Rho-ROK-dependent MLC phosphorylation in epithelial cells, which is depolarization induced but Ca2+ signal independent. Moreover, we provide evidence that the membrane potential-mediated regulation of cell contractility appears to be relevant physiologically in kidney cells, because the activation of the electrogenic Na+-alanine cotransport leads to Rho activation and MLC phosphorylation.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials and antibodies. ML-7, Y-27632, ionomycin, and thapsigargin, gramicidin D, and LY-294002 were obtained from Calbiochem. Clostridium difficile toxin B was purchased from TechLab, and fura-2 AM and bis-(1,3-dibuthylbarbituric acid)-trimethine oxonol [DIBAC4(3)], 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) were obtained from Molecular Probes. Bovine serum albumin (BSA), urea, glycerol, dithiothreitol (DTT), thioglycolate, trichloroacetic acid (TCA), L-alanine, tetraphenylphosphonium (TPP+), nystatin, and the Fast OPD kit were purchased from Sigma-Aldrich (St. Louis, MO). The following antibodies were used: anti-MLC (SC-9449), anti-phospho-MLC (anti-ppMLC) (Thr18/Ser19) (SC-12896), blocking peptide for anti-phospho-MLC, and anti-RhoA antibody (SC-418) were obtained from Santa Cruz Biotechnology (Santa Cruz, CA), and anti-ppMLC (Thr18/Ser19) was purchased from Cell Signaling Technology (Beverly, MA). Peroxidase- and Cy3-conjugated secondary antibodies were obtained from Jackson ImmunoResearch Laboratories (West Grove, PA).

Cells. LLC-PK1, a kidney proximal tubule epithelial cell line, and Madin-Darby canine kidney (MDCK) cells, a canine distal tubular epithelial cell line, were obtained from the American Type Culture Collection (Manassas, VA). Both cell lines were maintained in DMEM supplemented with 10% fetal bovine serum and 1% antibiotic suspension (penicillin and streptomycin; Sigma-Aldrich) in an atmosphere containing 5% CO2. Confluent cells were serum depleted for 3 h in HEPES-buffered RPMI medium before the experiments.

Media. HCO3-free RPMI 1640 was buffered with 25 mM HEPES to pH 7.4 (HPMI). The Na+-based medium (referred to as Na+ medium) consisted of (in mM) 130 NaCl, 3 KCl, 1 MgCl2, 1 CaCl2, 5 glucose, and 20 Na+-HEPES (pH 7.4). The K+-based medium (referred to as K+ medium) contained (in mM) 130 KCl, 1 MgCl2, 1 CaCl2, and 20 HEPES, pH 7.4. The Ca2+-free Na+ and K+ media were made by omitting CaCl2 from the corresponding solutions and supplementing them with 1 mM EGTA. For Na+-free choline medium, NaCl was replaced with choline chloride.

Detection of myosin phosphorylation by urea-glycerol polyacrylamide gel electrophoresis. Non-, mono-, and diphosphorylated forms of MLC were separated using nondenaturating urea-glycerol polyacrylamide gel electrophoresis (PAGE) as described in our previous studies (11, 38). Briefly, confluent LLC-PK1 or MDCK cells grown in six-well plates were serum depleted for 3 h and treated as indicated in the figure legends. Subsequently, they were lysed in ice-cold acetone containing 10% TCA and 10 mM DTT, followed by centrifugation for 10 min at 12,500 rpm (4°C). The resulting pellet was washed with pure acetone, allowed to air dry, and dissolved in 60 µl of sample buffer containing 8.02 M urea, 234 mM sucrose, 23 mM glycine, 10.4 mM DTT, 20 mM Tris (pH 8.6), and 0.01% bromphenol blue. Samples were separated on a 12% urea-glycerol gel and blotted onto nitrocellulose membrane. The membrane was blocked with 2% BSA (1 h) and incubated with an anti-MLC primary antibody (1:500 dilution; 1 h). After being washed, the membrane was incubated with a peroxidase-coupled anti-goat antibody (1:3,000 dilution). The blot was washed and developed with the enhanced chemiluminescence kit from Amersham Pharmacia Biotech. Total MLC (i.e., the sum of all 6 bands) was determined by performing densitometry in each lane, and the amount of non-, mono- and diphosphorylated forms was expressed as a percentage of total MLC in the same lane. This normalization allows direct comparison among the samples and prevents any potential errors that might arise from differences in loading. In some experiments, the blots were stripped and reprobed with anti-ppMLC antibody to verify specificity of the antibody.

Detection of myosin phosphorylation using a modified ELISA method. LLC-PK1 or MDCK cells were grown to confluence in 24-well plates. The cells were serum depleted for 3 h and treated as indicated in the figure legends. Subsequently, the cells were fixed with 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) for 20 min, washed three times with PBS, and permeabilized for 20 min with 0.1% Triton X-100 in the presence of 100 mM glycine to quench PFA. After being washed, cells were blocked with 250 µl of PBS containing 2% BSA for 45 min. Next, the cells were incubated with anti-diphospho-MLC (ppMLC) antibody (1:1,000 dilution) in PBS containing 0.2% BSA for 1 h. The cells were then washed five times for 15 min each with PBS and incubated with peroxidase-coupled secondary antibody (1:10,000 dilution) in PBS containing 0.2% BSA. After extensive washing of the cells, binding of the antibodies was detected using the Fast OPD kit. Each well was incubated with 750 µl of OPD reagent for 15 min at room temperature. The reaction was stopped by adding 250 µl of 3 M HCl. The supernatant was collected, and its absorbance was measured at 492 nm using a Beckman Coulter DU 640 spectrophotometer. Each determination was performed in duplicate or in triplicate. The background (absorbance of samples without primary antibody) for each condition was determined on the same plate and subtracted. The signal obtained in control cells was typically about threefold higher than the background. The results are expressed as the relative increase compared with the control. Similar results were obtained with another ppMLC antibody (Cell Signaling Technology; data not shown).

Rho activity assay. The amount of active Rho was determined using an affinity precipitation assay as described previously (11). Briefly, confluent LLC-PK1 cells grown in 10-cm dishes were treated as indicated in the respective figure legends. Cells were lysed in 800 µl of ice-cold lysis buffer containing 100 mM NaCl, 50 mM Tris base (pH 7.6), 20 mM NaF, 10 mM MgCl2, 1% Triton X-100, 0.5% deoxycholic acid, 0.1% SDS, 20 µl/ml protease inhibitor cocktail (BD Pharmingen), 1 mM Na3VO4, and 1 mM PMSF. The lysates were clarified by centrifugation at 12,000 rpm for 1 min at 4°C. After removing 20 µl of sample from each supernatant for determination of total Rho, we incubated the rest of the supernatant at 4°C for 45 min with 10–15 µg of glutathione-Sepharose beads covered with glutathione S-transferase-Rho-binding domain (GST-RBD), followed by extensive washing. Samples for total Rho and the pelleted beads were diluted in Laemmli sample buffer and boiled for 5 min. The proteins were separated using SDS-PAGE (15% gel) and transferred onto nitrocellulose membrane. Blots were blocked with 2% BSA for 1 h, followed by incubation with anti-Rho antibody (1:500 dilution; 1 h). Binding of the antibody was visualized using peroxidase-coupled anti-mouse antibody (1:3,000 dilution) and enhanced chemiluminescence. The amount of captured Rho was quantified by performing densitometry.

Immunofluorescence microscopy. Confluent cells grown on coverslips were serum depleted for 3 h and treated as indicated in the figure legends. Next, the cells were fixed with 4% PFA, washed with PBS, and permeabilized for 20 min with 0.1% Triton X-100 in the presence of 100 mM glycine. This was followed by washing and blocking the cells with 2% BSA for 1 h. The coverslips were then incubated with the primary antibody diluted in PBS containing 0.2% BSA. The primary antibody dilution for all antibodies was 1:100. Bound antibody was detected using the corresponding Cy3-conjugated secondary antibodies (1:1,000 dilution). Staining was visualized using a Nikon Eclipse TE200 microscope (x100 magnification) coupled to a Hamamatsu cooled charge-coupled device camera operated using Simple PCI software.

Vectors and transient transfection. The vector-encoding C3 transferase that we used, a kind gift from Dr. Keith Burridge, was described previously (2). The vector-encoding enhanced green fluorescent protein (EGFP), pEGFP, was purchased from Clontech Laboratories. LLC-PK1 cells were plated on coverslips, and the next day they were cotransfected with cDNA encoding for C3 transferase (1 µg) and GFP (0.2 µg) using FuGENE6 (Roche Molecular Biochemicals) according to the manufacturer’s instructions. Transfection with GFP, which by itself had no effect on ppMLC staining (Fig. 7D; see also our previous study, Ref. 38), was used to visualize transfected cells.



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Fig. 7. Depolarization activates Rho. A: confluent LLC-PK1 cells grown in 10-cm dishes were treated with Na+ or K+ medium. The amount of active Rho was determined using a glutathione-S-transferase-Rho-binding domain (GST-RBD) precipitation assay as described in MATERIALS AND METHODS. The amount of precipitated active Rho was detected using Western blot analysis with an anti-Rho antibody. The blots were quantified by performing densitometry and expressed as the relative increase compared with the values obtained in control cells treated with Na+ medium (n = 11; P < 0.005). B: LLC-PK1 cells grown on coverslips were exposed to Na+ medium or to K+ medium for 15 min. The cells were fixed, permeabilized, and stained using anti-Rho primary and Cy3-labeled secondary antibodies. C: untreated LLC-PK1 cells or cells pretreated with 100 pg/ml Clostridium difficile toxin B for 2 h were preincubated with Na+ medium, followed by 15-min incubation with K+ medium. Myosin phosphorylation was quantified using a ppMLC ELISA as in Fig. 1. Results are expressed as relative increase in ppMLC compared with Na+ that was taken as unity (data not shown) (n = 5; P < 0.05). D: LLC-PK1 cells grown on coverslips were transfected with cDNA for green fluorescent protein (GFP) alone or with C3 transferase (1 µg) and GFP (0.2 µg). After 48 h, the cells were serum depleted and then preincubated for 15 min in Na+ medium followed by K+ medium for 15 min. The cells were fixed and stained with anti-ppMLC as in Fig. 1. ppMLC staining (left) and GFP fluorescence (right) of the same field are shown. The asterisks (top) and dots (bottom) indicate the transfected cells on the corresponding images.

 
Measurement of membrane potential. Changes in membrane potential were detected using the fluorescent potential-sensitive dye DIBAC4(3) (as described in Ref. 63). LLC-PK1 cells grown on coverslips were loaded with 5 µM DIBAC4(3) in Na+ medium for 10 min at 37°C. The coverslips were placed into Attofluor cell chambers (Molecular Probes) and mounted on the stage of a Leica DM/IRB inverted microscope. The fluorescence of a small population of cells was measured using a Delta-RAM dual wavelength illumination and recording system from Photon Technologies International with Felix software. All measurements were obtained at 37°C. The excitation and emission wavelengths were 470 ± 5 and 510 ± 5 nm, respectively. After obtaining a steady-state baseline in Na+ medium, the medium was exchanged for the indicated solution supplemented with 5 µM DIBAC4(3). Each measurement was normalized to the baseline determined in the Na+ medium (resting potential). Calibration of the membrane potential was performed using the monovalent cation ionophore gramicidin (10 µM) and appropriately designed medium as described previously (63). The membrane potential was calculated using the following modified Goldman equation:

Intracellular Na+ and K+ concentrations were assumed to be 140 mM and 10 mM, respectively. Extracellular Na+ and K+ concentrations ([Na+]o + [K+]o, respectively) were set to the values (in mM) 13, 15, 20, 40, 70, or 150 in a solution containing (in mM) 10 HEPES, 1 CaCl2, 1 MgCl2, and varying concentrations of N-methyl-D-glucamine to adjust the osmolarity to 300 mosmol/l.

Measurement of intracellular Ca2+. LLC-PK1 cells grown on coverslips were loaded with fura-2 AM for 30 min in serum-free HPMI at 37°C. The cells were washed and kept in HPMI at 37°C for an additional 15 min to allow complete hydrolysis of the dye. After the coverslips were washed again, ratiometric microfluorometry was performed on a small population of cells using excitation wavelengths of 340 ± 5 nm and 380 ± 5 nm and an emission wavelength of 510 ± 5 nm.

Densitometry. Densitometric analysis of blots was performed using a model GS-690 imaging densitometer and Molecular Analyst software (version 1.5) obtained from Bio-Rad Laboratories (Hercules, CA) as described previously (11).

Statistical analysis. Data are presented as means ± SE of the number of experiments indicated (n) or as representative immunoblots or images of at least three similar experiments. When the data were normalized to the control level, a paired Student’s t-test was performed. In other cases, one-way ANOVA for multiple comparisons or an unpaired t-test was performed, with P < 0.05 considered statistically significant.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Increased extracellular K+ induces MLC phosphorylation in kidney epithelial cells. To assess whether membrane depolarization triggers phosphorylation of MLC in epithelial cells, porcine proximal tubular (LLC-PK1) cells were preincubated in a Na+-based solution (Na+ medium), which was subsequently replaced with a K+-based solution (K+ medium). Initially, we determined MLC phosphorylation by performing urea-glycerol PAGE. As we reported previously (11), there are two MLC isoforms in LLC-PK1 cells, both of which can exist in non-, mono-, and diphosphorylated forms, resulting in three doublets (six bands) when separated using urea-glycerol PAGE. The two uppermost bands correspond to the nonphosphorylated forms, the middle bands correspond to the mono-phosphorylated forms, and the lowermost bands correspond to ppMLC. As shown in Fig. 1A, K+ medium induced a substantial increase in the diphosphorylated form of both isoforms. To quantify the response and allow accurate comparison between the samples independent of any differences in loading, the abundance of the non-, mono-, and diphosphorylated forms was expressed as the percentage of total MLC in each sample (see Fig. 1A). In control cells, incubated in Na+ medium, ~30% of MLC was diphosphorylated and ~40% was nonphosphorylated. In contrast, when cells were challenged with the K+ medium, we observed an approximately twofold rise (to 60%) in ppMLC accompanied by a corresponding approximately twofold decrease (to 20%) in the nonphosphorylated form. Interestingly, the monophosphorylated pool remained essentially unchanged: it accounted for ~30% in both control and K+-stimulated samples. The increase in ppMLC was rapid and sustained. A substantial rise was detected as fast as 1 min, exhibiting a peak at 5 min, followed by a plateau or a slight decay thereafter (Fig. 1B). To routinely measure the extent of myosin phosphorylation, we developed an ELISA-like assay using a ppMLC-specific antibody (see MATERIALS AND METHODS). This approach was faster and easier to quantify than the urea-glycerol PAGE method. Using this ELISA-like assay, we detected a sizable (~2-fold) increase in ppMLC upon stimulation with high K+ (Fig. 1C), which was in excellent agreement with our results obtained with electrophoretic separation (see Fig. 1A). The anti-ppMLC antibody was fully specific for the doubly phosphorylated forms as verified by reprobing the urea-glycerol PAGE-separated proteins with the anti-ppMLC antibody (Fig. 1A, right). Regrettably, the ppMLC antibody could not be used for direct Western blot analysis in SDS gels, because it did not react with the SDS-denatured ppMLC of LLC-PK1 cells.



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Fig. 1. High extracellular K+ induces myosin light-chain (MLC) phosphorylation in epithelial cells. Kidney proximal tubular (LLC-PK1) cells grown to confluence were serum depleted for 3 h. The cells were preincubated in Na+ medium for 15 min followed by treatment with Na+ or K+ medium for 15 min (A, C–E) or the indicated times (B). A: cells were lysed in an acetone-TCA extraction buffer and subjected to urea-glycerol polyacrylamide gel electrophoresis (PAGE) as described in MATERIALS AND METHODS. MLC was detected using Western blot analysis with anti-MLC (left). n, nonphosphorylated MLC; m, monophosphorylated MLC; d, diphosphorylated MLC. The blot was stripped and reprobed with anti-diphospho-MLC (right). MLC phosphorylation was quantified using densitometry. The total amount of MLC in each lane was determined, and the amount of each phosphorylated form was expressed as the percentage of total MLC. The graph represents means ± SE; n = 11. K+ induced a twofold increase in diphospho-MLC. P < 0.0001. B: cells were treated with K+ medium for the indicated time. MLC phosphorylation was detected and quantified as in A. Data are means ± SE; n ≥ 5 individual experiments. C: MLC phosphorylation was determined using modified ELISA as detailed in MATERIALS AND METHODS. Determinations for both conditions on each plate were performed in triplicate. The background (samples treated identically but stained only with the secondary antibody) was subtracted, and the results for each plate were expressed as relative increase compared with the Na+ medium taken as unity. Data are means ± SE; n = 28. K+ induced a twofold increase in diphospho-MLC. P < 0.0001. D: cells grown on coverslips were treated as indicated and then stained for phospho-MLC (ppMLC) as detailed in MATERIALS AND METHODS. To visualize the accumulation of ppMLC at the periphery (middle; arrows) and in fibers (left), images were obtained at different depths, i.e., at the middle or near the base of the cells. The same camera settings were used for all conditions. E: cells treated with K+ medium were stained with anti-ppMLC in the presence of 1 µg of blocking peptide (left) or with 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) to visualize the nuclei.

 
To determine the spatial distribution of K+-induced MLC phosphorylation, the cells were stained with the ppMLC-specific antibody and visualized using fluorescence microscopy. As shown in Fig. 1D, in the Na+ medium, the cells exhibited only very faint intracellular staining without any discernible structure or sometimes showed some accumulation of ppMLC at the cell periphery (left). In contrast, in K+-treated cells, the overall labeling became much stronger and acquired a specific structure. An enrichment of ppMLC in the submembranous area was often observed (see Fig. 1D, middle, arrows), and increased accumulation was visible in the cytoplasm, with some accumulation in thin, stress fiber-like structures at the base of the cells (Fig. 1D, right). Interestingly, an increase in specific ppMLC staining was also detectable in the nucleus. The staining was specific because an epitope-matching blocking peptide entirely prevented the K+-induced labeling (Fig. 1E).

To test whether the high-K+-provoked MLC response is unique to LLC-PK1 cells or whether it is also present in other kidney epithelial cells, we performed the above experiments on MDCK cells, a canine distal tubular cell line. As shown in Fig. 2, MDCK cells showed both qualitatively and quantitatively similar responses to LLC-PK1 cells. Stimulation with high K+ increased MLC phosphorylation as detected using urea-glycerol PAGE and resulted in enhanced ppMLC staining along intracellular fibers, at the periphery, and in the nucleus.



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Fig. 2. High extracellular K+ induces MLC phosphorylation in Madin-Darby canine kidney (MDCK) cells. MDCK cells were treated with K+ medium, and MLC phosphorylation was detected as in Fig. 1. A: typical urea-glycerol gel and quantitation of n = 3 similar gels. K+ induced a twofold increase in ppMLC. P < 0.005. B: immunofluorescence staining using ppMLC antibody of cells treated with Na+ medium (left) or K+ medium (right).

 
Plasma membrane depolarization induces MLC phosphorylation. To verify that the K+-induced MLC phosphorylation is indeed mediated by depolarization, first we determined the effect of high K+ on the membrane potential in LLC-PK1 cells using DiBAC4(3), a potential-sensitive fluorescent dye. Calibration was performed using the ionophore gramicidin and appropriately composed medium (63) (Fig. 3A). The calculated resting membrane potential of LLC-PK1 cells was approximately –50 mV, which is in excellent agreement with previously published patch-clamp data (14). Replacement of the Na+ medium with K+ medium entirely dissipated the transmembrane potential (Fig. 3B). To further substantiate depolarization, as opposed to high-K+ concentration, as the critical factor per se, we examined the effect of other stimuli that dissipate the transmembrane potential. Toward this end, we permeabilized the membrane for monovalent ions using the ionophore nystatin. This maneuver also induced the collapse of the membrane potential (Fig. 3C) and concomitantly caused a significant increase in the phosphorylation of MLC (Fig. 3D). Because nystatin may affect the internal Cl concentration and the cell volume as well, we applied yet another, more specific strategy. We used the lipid-soluble cation tetraphenylphosphonium (TPP+), which freely permeates cell membranes, thereby collapsing the negative membrane potential as shown on Fig. 3C. Exposure of the cells to TPP+ caused a large shift from nonphosphorylated MLC to ppMLC as shown by urea-glycerol PAGE (Fig. 3E). Quantification with the ppMLC ELISA revealed a nearly twofold increase in ppMLC (Fig. 3F). Morphologically, TPP+ induced an overall increase in cytosolic ppMLC with a robust accumulation at the cell periphery (Fig. 3G). Thus three different manipulations (increased [K+], nystatin treatment, and TPP+), each of which leads to membrane depolarization by distinct mechanisms, induced MLC phosphorylation of similar magnitude. Taken together, these results suggest that plasma membrane depolarization induces strong myosin phosphorylation in kidney epithelial cells.



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Fig. 3. Membrane potential measurements in LLC-PK1 cells. AC: cells were loaded with bis-(1,3-dibuthylbarbituric acid)-trimethine oxonol [DIBAC4(3)] for 10 min. The fluorescence was measured as detailed in MATERIALS AND METHODS. A: typical calibration curves. Inset: correlation between steady-state fluorescence and calculated membrane potential. B and C: arrows indicate where medium was exchanged for Na+ or K+ medium (B) or Na+ medium containing 10 mM tetraphenylphosphonium (TPP+) or 400 U/ml nystatin (C) supplemented with 5 µM DIBAC4(3). DG: plasma membrane depolarization by various means induces MLC phosphorylation in LLC-PK1 cells. D: cells were permeabilized with 400 U/ml nystatin in a Na+-based medium in which [Ca2+]i was buffered to 100 nM as described previously (11). The level of MLC phosphorylation was determined using urea-glycerol PAGE. EG: cells were treated with 1 mM TPP+ in Na+ medium for 15 min. MLC phosphorylation was detected using urea-glycerol PAGE (E), ppMLC ELISA (F), or immunostaining (G) as in Fig. 1. Results in B are expressed as relative increase compared with Na+ medium controls taken as unity (n = 4; P < 0.005).

 
Depolarization-induced MLC phosphorylation is independent of changes in intracellular [Ca2+]. In many cells, depolarization can open voltage-gated channels, causing Ca2+ influx and a subsequent increase in the cytoplasmic Ca2+. This in turn activates MLCK, thereby increasing MLC phosphorylation. Therefore, we sought to examine whether a similar mechanism was responsible for the depolarization-induced MLC phosphorylation in our epithelial cells. To prevent the contribution of extracellular Ca2+, we induced depolarization in a Ca2+-free medium. As shown in Fig. 4A, under Ca2+-free conditions, high K+ remained able to cause a sizable increase in MLC phosphorylation as detected using urea-glycerol PAGE. Immunofluorescence microscopy showed that depolarization under these conditions caused a marked accumulation of ppMLC at the cell periphery that was similar to or even more pronounced than that observed in the presence of normal extracellular Ca2+ (Fig. 4B; see also Fig. 1D). During the course of these experiments, we noted that compared with resting cells incubated in Ca2+-containing Na+ medium, Ca2+ removal itself resulted in an increase in ppMLC (compare quantitation of urea-glycerol PAGE in Figs. 1A and 4A). Immunofluorescence microscopy also verified that removal of extracellular Ca2+ alone caused increased staining in the cytoplasm with a few visible fibers and submembranous accumulation of ppMLC (Ca2+-free Na+ medium; Fig. 4B, left). These findings were in accord with earlier observations suggesting that Ca2+ removal increases (rather than suppresses) cell contraction (7, 35) and enhances myosin phosphorylation in epithelial cells (26). To quantify these effects, we treated cells on the same multiwell plates with Ca2+-containing or Ca2+-free Na+ and K+ media, and then we measured the amount of ppMLC using the modified ELISA method. The results were normalized to values obtained under Ca2+-containing Na+ (resting) conditions. Indeed, the removal of extracellular Ca2+ by itself increased ppMLC (Fig. 4C). Importantly, these measurements substantiated that subsequent depolarization by high K+ in the absence of extracellular Ca2+ caused an additional ~1.5-fold rise in ppMLC, superimposed on the already elevated basal phosphorylation. Similarly to high K+, TPP+ was also able to induce MLC phosphorylation in the absence of extracellular Ca2+ (Fig. 4, D and E). Collectively, these results suggest that the depolarization-induced MLC phosphorylation does not require extracellular Ca2+, in sharp contrast to the KCl-induced MLC phosphorylation in muscle cells. Curiously, the absence of extracellular Ca2+ triggers MLC phosphorylation, an effect that might be related to the disassembly of Ca2+-dependent intercellular contacts (see DISCUSSION). However, regardless of this modulating effect, membrane depolarization is capable of inducing MLC phosphorylation in the presence or absence of extracellular Ca2+.



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Fig. 4. Depolarization-induced MLC phosphorylation is independent of extracellular Ca2+. A and B: LLC-PK1 cells were preincubated with Na+ medium for 15 min, followed by incubation in Ca2+-free Na+ or K+ medium for an additional 15 min. MLC phosphorylation was determined using urea-glycerol PAGE (A) (n = 7; P < 0.01) or immunofluorescence (B). C: cells grown in 24-well plates were treated with Ca2+-containing Na+ or K+ medium or Ca2+-free Na+ or K+ medium as indicated. The amount of ppMLC was determined using the ELISA method (n = 6). D and E: cells were treated with Ca2+-free Na+ medium with or without 1 mM TPP+. MLC phosphorylation was assessed using urea-glycerol PAGE (D) or ppMLC ELISA (E) as in Fig. 1. Data in E are expressed as relative increase compared with levels obtained in the Ca2+-free Na+ medium (n = 9; P < 0.005).

 
While the fact that depolarization led to MLC phosphorylation in the absence of extracellular Ca2+ excluded the involvement of Ca2+ influx through any voltage-gated channel, it was still conceivable that KCl could increase cytosolic Ca2+ by liberating the ion from intracellular stores. To address this possibility, we measured the changes in [Ca2+]i under our experimental conditions using ratiofluorometry in fura-2-loaded cells. Replacement of Na+ medium by K+ medium did not alter [Ca2+]i (Fig. 5A). To verify that the LLC-PK1 cells were sufficiently loaded with fura-2 and that the system adequately monitored changes in [Ca2+]i, the membranes were permeabilized by the addition of 2 µM ionomycin in Ca2+-free (EGTA containing) medium. This resulted in an immediate, temporary increase in [Ca2+]i due to the release of Ca2+ from intracellular stores, followed by a sharp decline toward a minimal level. Readdition of Ca2+ rapidly elevated the fluorescence in ionomycin-permeabilized cells. These Ca2+ transients verified that, under our loading conditions, changes in [Ca2+]i were readily detectable and the intracellular Ca2+ stores contained releasable Ca2+.



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Fig. 5. Depolarization does not alter intracellular Ca2+ in LLC-PK1 cells. LLC-PK1 cells grown on coverslips were loaded with fura-2 AM for 30 min. The fluorescence was measured in small populations of cells using a PTI spectrophotometer as described in MATERIALS AND METHODS. A: after incubation in Na+ medium, the medium was exchanged for K+ medium (first arrow). At the indicated point, the K+ medium was replaced by Ca2+-free (and 1 mM EGTA-containing) K+ medium supplemented with 2 µM ionomycin. The ionophore initially released Ca2+ from intracellular stores into the cytosol (see transient peak), followed by Ca2+ efflux from the cells (downward deflection). At the end of the measurement, the cells were washed twice with Ca2+-containing K+ medium (arrows) to saturate the dye. B: where indicated by arrows, the Ca2+-containing Na+ medium was exchanged to Ca2+-free Na+ medium, followed by Ca2+-free K+ medium. To check the reactivity of the dye and obtain a maximum, the medium was exchanged to Ca2+-containing K+ supplemented with 2 µM ionomycin at the end of the measurement. CE: increased [Ca2+]i does not induce ppMLC in LLC-PK1 cells. Intracellular Ca2+ was monitored as above. Where indicated, 2 µM ionomycin (C) or 100 nM thapsigargin (TG) (D) was added to Na+ medium. E: cells grown on 24-well plates were treated for 10 min with 2 µM ionomycin or 100 nM TG. Myosin phosphorylation was quantified using ppMLC ELISA as in Fig. 1. Data are expressed as relative increase compared with control cells in Na+ medium (n = 5; none of the columns is significantly different from control).

 
Next, we tested the effect of depolarization on [Ca2+]i in Ca2+-free medium. Removal of extracellular Ca2+ by exchanging the Na+ medium with Ca2+-free Na+ medium caused a small-amplitude, transient blip in [Ca2+]i, which subsided within 1–2 min (Fig. 5B). Subsequent replacement of the Ca2+-free Na+ medium with Ca2+-free K+ medium had no effect on [Ca2+]i. Taken together, these results suggest that the depolarization-induced MLC phosphorylation is not mediated by changes in [Ca2+]i, in either the presence or the absence of extracellular Ca2+.

Recently, Emmert et al. (12) described Ca2+-independent MLC phosphorylation in MLCK-deficient fibroblasts. To discern the relative contribution that Ca2+ may play in the regulation of MLC phosphorylation in intact kidney epithelial cells, we examined the ability of increased [Ca2+]i to induce MLC phosphorylation. Addition of ionomycin to cells incubated in Ca2+-containing Na+ medium induced an immediate, sharp rise in [Ca2+]i, which remained elevated for the rest of the measurement period (Fig. 5C). Despite the elevated Ca2+ levels, ionomycin only marginally increased MLC phosphorylation as determined by performing ppMLC ELISA (Fig. 5E). Similarly, addition of 100 nM thapsigargin (TG), which caused a smaller but similarly long-lasting increase in [Ca2+]i, also failed to significantly elevate ppMLC (Fig. 5, D and E). Jointly, these data suggest that an increase in [Ca2+]i in LLC-PK1 cells by itself is neither necessary nor sufficient to induce ppMLC.

Depolarization-induced MLC phosphorylation is mediated by the Rho kinase pathway in kidney epithelial cells. We investigated the kinase pathways involved in mediating depolarization-induced MLC phosphorylation. To determine the role of the major myosin-phosphorylating enzyme MLCK, we used ML-7, a potent MLCK inhibitor. Even in the presence of a high (50 µM) concentration of ML-7, K+ was able to induce a sizable increase in ppMLC staining: Immunofluorescence analysis revealed that the peripheral accumulation, the nuclear staining, and the fiberlike structures were all manifest in ML-7-treated cells, similar to controls that were treated with K+ medium only (Fig. 6, A and B). This conclusion was verified by the other two methods as well. When assessed using urea-glycerol PAGE, neither the basal phosphorylation pattern nor its changes upon depolarization were significantly different between control and ML-7-treated cells (Fig. 6D). Indeed, quantitation using ELISA showed no reduction in the depolarization-induced ppMLC content in ML-7-treated cells (Fig. 6E).



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Fig. 6. Depolarization- and Ca2+ removal-induced MLC phosphorylation is mediated by Rho kinase and not by MLC kinase (MLCK). AE: LLC-PK1 cells were pretreated with 50 µM ML-7 for 15 min or 20 µM Y-27632 for 45 min in Na+ medium, followed by either Na+ or K+ medium for 15 min as indicated. The inhibitors were present throughout the treatment. Myosin phosphorylation was detected using immunostaining (AC), urea-glycerol PAGE (D) or ELISA (n = 5; for K+ vs. ML-7 + K+, P > 0.05; for K+ vs. Y-27632 + K+, P < 0.05) (E) as in Fig. 1. F: cells were treated with 20 µM Y-27632 as in C. The medium was changed to a Ca2+-free medium for 15 min. Myosin phosphorylation was detected by immunostaining. G and H: where indicated, cells were pretreated with 20 µM LY-294002 (LY) for 30 min, and the inhibitor remained present during treatment. Cells were treated with Na+ or K+ medium for 15 min. A typical urea gel (G) and quantitation of n = 7 similar gels are shown. ANOVA resulted in the following P values. Na+ vs. K+, P < 0.05; Na+ vs. LY Na+, P > 0.05; Na+ vs. LY K+, P > 0.05; K+ vs. LY Na+, P < 0.001; K+ vs. LY K+, P < 0.01; LY Na+ vs. LY K+, P > 0.05.

 
Next, we tested the potential involvement of another candidate kinase, ROK. In contrast to the lack of an inhibitory effect of ML-7, when cells were pretreated with the ROK inhibitor Y-27632, the depolarization-triggered ppMLC response was largely prevented. Specifically, immunofluorescence microscopy showed a marked inhibition in the K+-induced ppMLC accumulation in all three locations (compare Fig. 6, A and C). The inhibitory effect was also verified using urea-glycerol PAGE (Fig. 6D) and quantified by performing ppMLC ELISA (Fig. 6E), which showed that Y-27632 exerted ~70% inhibition of the depolarization-induced MLC phosphorylation. Interestingly, Y-27632 potently inhibited the Ca2+ removal-induced peripheral MLC phosphorylation as well (Fig. 6F). Taken together, these data support the notion that depolarization-induced MLC phosphorylation is mediated by the Rho kinase pathway and appears to be mostly independent of MLCK.

Depolarization activates Rho, which is necessary for MLC phosphorylation. The ability of Y-27632 to interfere with the depolarization-induced MLC phosphorylation suggests that changes in the membrane potential activate the Rho-Rho kinase pathway. Because ROK can be activated both by Rho-dependent and Rho-independent mechanisms (17), we wished to discern whether depolarization directly affects Rho activation. Toward this end, we performed Rho precipitation assays to assess the amount of active Rho in cell lysates from control and depolarized cells. This approach relies on a fusion protein composed of GST and the RBD of rhotekin, a downstream effector of Rho, which binds the active (GTP-bound) form but fails to react with the inactive (GDP-bound) form of Rho. As shown in Fig. 7A, K+ medium induced substantial Rho activation. Densitometric quantitation of these pull-down assays showed that the K+ medium provoked a 1.82 ± 0.24-fold increase in active Rho compared with the Na+ medium. To visualize the intracellular distribution of Rho, another indicator of its activation, cells were immunostained with an anti-Rho antibody. While in control cells Rho exhibited predominantly cytosolic distribution with occasional membrane localization, after K+ treatment, it showed a marked peripheral staining suggestive of membrane translocation (Fig. 7B). Together, these results imply that in tubular epithelial cells, depolarization activates Rho and induces its redistribution to the cell periphery.

To assess whether depolarization-induced Rho activation is indeed necessary for MLC phosphorylation, we interfered with Rho activation by various means. First, we used C. difficile toxin B, an enzyme that catalyzes the glycosylation of a threonine residue on Rho family GTPases and thereby inactivates them (28). The advantage of this toxin is that it is cell permeable; therefore, it can be introduced into the entire cell population. Cells were incubated with toxin B and exposed to K+ medium. Toxin-treated cells showed typical morphological changes as described in our earlier work (11) but remained firmly attached to the plates. As shown in Fig. 7C, while K+ medium induced a more than twofold increase in ppMLC, as demonstrated earlier, the increase was completely blocked in toxin B-treated cells, suggesting that depolarization requires functional Rho to exert its effect on myosin.

Although toxin B offers the advantage of allowing treatment of the whole cell population, it is not specific for Rho but inhibits all Rho family GTPases. To interfere specifically with endogenous Rho activity, we used Clostridium botulinum C3 transferase, a toxin that selectively ADP ribosylates and thereby inactivates Rho. Because this protein is not cell permeable, we cotransfected the cells with a vector-encoding C3 toxin, together with GFP, to identify the successfully transfected (green) cells. Subsequently, we challenged the monolayer with K+ medium and stained it for ppMLC. Figure 7D shows that in cells expressing GFP only, depolarization induced marked ppMLC accumulation (Fig. 7D, top). In contrast, in cells expressing C3 transferase, the depolarization-induced ppMLC accumulation was completely abolished (Fig. 7D, bottom).

The mechanism whereby the Rho-ROK pathway is activated upon depolarization remains elusive. Because previous studies have shown that phosphatidylinositol 3-kinase (PI3-kinase) can be activated by depolarization (40, 59) and has been implicated as an upstream regulator of the Rho-ROK pathway (39), we wondered whether PI3-kinase might be involved in the depolarization-induced MLC response. To address this question, we used LY-294002, a specific PI3-kinase inhibitor. Figure 6, G and H, shows that LY-294002 suppressed the basal MLC phosphorylation and significantly mitigated the depolarization-provoked rise in ppMLC. This finding is consistent with a contributory role of PI3-kinase as one of the upstream mediators of the potential-sensitive MLC phosphorylation.

Electrogenic L-alanine-Na+ cotransport induces myosin phosphorylation. Proximal tubule epithelial cells contain a large number of apical transporters, many of which are electrogenic (31, 62). Accordingly, Na+-driven cotransport processes, such as Na+-coupled glucose or alanine uptake, have been shown to depolarize the cells (25, 44). Therefore, we next investigated whether physiological membrane potential changes associated with the operation of the Na+-alanine cotransporter were able to induce MLC phosphorylation in LLC-PK1 cells.

Figure 8A shows that the addition of 20 mM L-alanine to the Na+ medium induced a drop in the membrane potential and a concomitant increase in the general ppMLC staining (Fig. 8B). The accumulation of ppMLC was detected, both in the cytoplasm and at the cell periphery. Moreover, we often observed ppMLC-positive thin stress fibers near the basal surface of alanine-treated cells (data not shown). Thus the pattern of ppMLC distribution resembled that observed in the high-K+ medium, albeit that the overall effect appeared weaker. Importantly, similarly to the K+-induced response, the L-alanine-triggered increase in MLC phosphorylation was also entirely prevented by Y-27632, suggesting that ROK is a key mediator of this process as well (Fig. 8B, right). In agreement with the qualitative observations, 10-min exposure of the cells to L-alanine caused a 1.26 ± 0.08-fold increase in ppMLC as measured using the ELISA technique (Fig. 8C, first two columns). Similar MLC phosphorylation was detected in MDCK cells upon the addition of L-alanine (data not shown). We thought that if the effect of L-alanine was indeed due to its electrogenic transport, it should depend on the presence of external Na+. To test this prediction, we examined whether replacement of Na+ with choline would alter the effect of alanine on MLC phosphorylation. Figure 8, A and C, shows that in Na+-free (choline chloride) medium, the addition of alanine did not depolarize the membrane and failed to induce MLC phosphorylation. This observation gives strong support to the notion that the effect of alanine was indeed due to its Na+-dependent uptake into the cells.



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Fig. 8. Electrogenic L-alanine-Na+ cotransport induces MLC phosphorylation. A: membrane potential of LLC-PK1 cells was measured as in Fig. 3. Where indicated by the arrows, the medium was exchanged with Na+ or choline medium containing 20 mM alanine, supplemented with 5 µM DIBAC4(3). B: cells were incubated in Na+ medium with or without 20 mM L-alanine for 10 min. Where indicated, cells were pretreated with 20 µM Y-27632 for 45 min and the inhibitor remained present during treatment with alanine. Cells were fixed and stained with anti-ppMLC. The images were obtained with the same camera settings and are representatives of three different experiments. C: LLC-PK1 cells were treated with 20 mM alanine for 10 min in Na+ medium or Na+-free choline medium. Myosin phosphorylation was quantified using a ppMLC ELISA as in Fig. 1. The results are expressed as relative increase from the respective controls (i.e., cells treated with Na+ or Na+-free choline medium). n = 9; P < 0.03, control vs. alanine in Na+ medium. P > 0.15, for control vs. alanine in choline medium.

 
Having shown that alanine induces Rho kinase-dependent MLC phosphorylation, we asked whether it can also activate Rho. To address this point, we compared the activity and localization of Rho in control and alanine-treated cells using the GTP-Rho precipitation assays and immunostaining, respectively. As shown in Fig. 9A, L-alanine induced a 1.65 ± 0.25-fold increase in the amount of active Rho. Moreover, alanine exposure resulted in a marked subcellular redistribution of endogenous Rho protein; while Rho was mostly cytosolic in control cells, it exhibited strong peripheral accumulation along the membrane in alanine-treated cells (Fig. 9B). In summary, these data suggest that L-alanine is able to induce Rho-Rho kinase-dependent MLC phosphorylation that is likely due to its Na+-dependent uptake and consequent depolarization.



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Fig. 9. L-alanine-Na+ cotransport activates Rho. Confluent LLC-PK1 cells grown in 10-cm dishes (A) or coverslips (B) were treated with or without 20 mM L-alanine added in Na+ medium. A: active Rho was detected as in Fig. 7. The precipitated Rho protein was quantified using densitometry and expressed as the relative increase compared with the optical density values of controls obtained on the same X-ray films (n = 6; P < 0.05). B: coverslips were stained with anti-Rho antibody as in Fig. 7.

 

    DISCUSSION
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According to the widely accepted paradigm, the major physiological role of the membrane potential in nonexcitable cells is to provide the necessary driving force for electrogenic processes and to help maintain the normal intracellular ionic environment. Much less is known about the presumably very important direct roles of the membrane potential (or its changes) in the regulation of biochemical signaling processes and cell morphology (43). In the present work, we provide evidence for the first time that depolarization of the plasma membrane in kidney epithelial cells induces rapid accumulation of the diphosphorylated form of the MLC. Importantly, this reaction is not due to, and does not require, an increase in intracellular Ca2+ and does not appear to be brought about by MLCK. Instead, the activation of the Rho-Rho-kinase pathway is responsible for the elevated MLC phosphorylation.

In smooth muscle cells, the depolarization-induced activation of MLC phosphorylation is entirely dependent on Ca2+ influx (41, 46). On the other hand, Ca2+-independent Rho-ROK-mediated MLC phosphorylation has been observed in fibroblasts (12); however, these effects were provoked by chemical agonists and not by depolarization. Thus the depolarization-induced, Rho-ROK-mediated, but Ca2+ signal-independent MLC phosphorylation represents a new mode of regulation of the contractile apparatus present in kidney epithelial cells.

Several lines of evidence suggest that the triggering factor for MLC phosphorylation is indeed depolarization. Dissipation of the membrane potential by each of three distinct mechanisms (high K+, which eliminates the major driving force for the resting potential; nystatin, which permeabilizes the membrane for monovalent ions; and the lipophilic cation TPP+, which accumulates excessively and at higher concentrations may induce an inside positive diffusion potential) provoked similar changes in ppMLC content and distribution. While these maneuvers might have changed other parameters (e.g., intracellular ion concentrations) as well, and although the participation of these cannot be excluded, the most straightforward interpretation of the data implicates the membrane potential as the critical component. Moreover, we found that a physiological stimulus, L-alanine, that caused depolarization by its electrogenic uptake through Na+-coupled cotransport (25, 31, 44) also induced MLC phosphorylation in the presence but not in the absence of Na+. We observed that the alanine-induced rise in total ppMLC as determined with our newly developed ELISA was less than that observed with the other depolarizing stimuli. This is expected to be so because alanine induced only a partial drop in the membrane potential, in agreement with patch-clamp data (25, 44).

Our findings imply that the membrane potential may directly regulate the cytoskeleton in epithelial cells. Consistent with this notion, in bovine corneal endothelial cells that are functionally equivalent to secretory epithelial cells, depolarization alters the actin cytoskeleton and causes retraction of the cells from each other (8). Although in this study the phosphorylation state of MLC and the underlying signaling mechanisms were not investigated, our present findings are consistent with increased epithelial contractility upon depolarization. Our results suggest a plausible molecular mechanism for these depolarization-provoked structural changes.

We found that depolarization resulted in the accumulation of ppMLC at the cell periphery, in fine fibers at the base of the cells, and in the nucleus. While increased contractility in the central areas and at the membrane may affect a variety of functions, including paracellular permeability (see below), the importance of nuclear ppMLC remains elusive. Nonetheless, the nuclear signal appears to be specific because it was eliminated by a specific blocking peptide and was effectively suppressed by the Rho kinase inhibitor Y-27632. In accordance with our findings, Ueda et al. (57) described specific nuclear staining by a ppMLC antibody in HELA cells that was eliminated by preadsorbing the antibody with its specific antigen.

Our results suggest that the depolarization-provoked MLC phosphorylation is mediated by the Rho-ROK pathway. This conclusion is supported by the findings that 1) depolarization activated Rho and induced its translocation to the periphery, 2) inhibition of Rho by either C. difficile toxin B or the more specific C3 transferase eliminated the depolarization-triggered MLC phosphorylation, and 3) the ROK inhibitor Y-27632 largely prevented the MLC response. While the mechanism of depolarization-induced Rho activation and peripheral translocation remains unknown, it appears to be tissue specific because it was found to be Ca2+ dependent in smooth muscle cells (46) but not in epithelia. Interestingly, Rac, another member of the Rho family, was also shown to be translocated to the membrane upon depolarization in endothelial cells (49). Because depolarization activates PI3-kinase in certain cells (40, 59) and has been suggested to act upstream of the Rho-ROK pathway (39), we investigated whether PI3-kinase might participate in the MLC response. Our findings that LY-294002 reduced the basal and depolarization-stimulated MLC phosphorylation are consistent with such a role. Conceivably, PI3-kinase may facilitate small GTPases by stimulating their GEFs. Indeed, several of the identified GEFs harbor lipid-binding PH domains (13) that could interact with and are activated by the products of PI3-kinase. Alternatively, changes in the membrane potential might modulate the exposed charges or the composition of the lipid bilayer, directly influencing the binding and/or activity of GEFs. Further work is needed to validate these possibilities. It is noteworthy that we and others have shown that Rac, Cdc42, and Rho are also responsive to changes in cell volume and intracellular ionic strength (6, 11, 36). Taken together, these data imply that the Rho family small GTPases are not only regulated by biochemical changes but also sensitive to physical parameters.

While Rho is the major activator of ROK, the kinase can be stimulated by Rho-independent pathways as well, such as by caspase-mediated cleavage (48). However, we found no evidence for the involvement of this mechanism, because ROK was not cleaved in depolarized cells, and caspase inhibitors did not interfere with MLC phosphorylation (data not shown).

The relative contribution of various kinases in MLC phosphorylation appears to differ significantly between tissues. In smooth muscle cells, MLC is phosphorylated predominantly by the Ca2+-dependent MLCK, while the major target of ROK is thought to be myosin phosphatase (51). Accordingly, MLC phosphorylation is Ca2+ dependent and is inhibited by MLCK blockers (41, 64). In contrast, in epithelial cells, we found no change upon depolarization in intracellular Ca2+, and ML-7 had only a marginal effect on the MLC response. The applied dose of ML-7 likely inhibited MLCK because it abolished spreading, an MLCK-dependent process (19, 47) (C Di Ciano-Oliveira and A Kapus, unpublished observations). Furthermore, [Ca2+]i elevation failed to increase MLC phosphorylation, arguing against the role of MLCK as the centrally regulated element of the pathway. Similar findings were reported in chicken embryonic cells, in which [Ca2+]i rise induced only a small, transient MLC phosphorylation (33). These observations, together with the strong inhibitory effect of Y-27632, suggest that ROK acts not only by the inhibition of MLC phosphatase but also by directly phosphorylating MLC. This conclusion is consistent with previous reports showing that Rho kinase can directly diphosphorylate MLC in nonmuscle cells and can mediate effective MLC phosphorylation even in the absence of MLCK (12, 57). A novel and intriguing regulator of myosin activity is the protein kinase C-mediated inhibitor CPI-17. Both ROK and PKC have been shown to phosphorylate CPI-17, which in turn enhances myosin phosphorylation by inhibiting myosin phosphatase (23). Future studies are warranted to assess the role of CPI-17 in the depolarization-induced increase in contractility.

Interestingly, removal of extracellular Ca2+ itself promoted myosin phosphorylation. This finding is consistent with earlier reports showing that removal of extracellular Ca2+ causes an immediate contraction of MDCK cells (15, 35). Because we found that Ca2+ removal-triggered MLC phosphorylation was also inhibited by Y-27632, we suggest that Rho-ROK activation is the underlying mechanism. Rho activation under these conditions might be related to the disassembly of intercellular contacts. Accordingly, the inverse process, i.e., the formation of cadherin-based cell-cell adhesions, was reported to strongly downregulate Rho activity (42). Importantly, the Rho-ROK-MLC system remains sensitive to the membrane potential and was further stimulated by depolarization even after Ca2+ removal.

Finally, we consider the physiological importance of the depolarization-induced MLC phosphorylation in epithelia. Cell contractility, governed by the phosphorylation state of MLC, is a key modulator of many epithelial functions, including the regulation of apical ion transporters (53) and the permeability of tight junctions (TJs) (21, 56) that are crucial determinants of the paracellular transport pathway. While basal Rho activity and contractility are required to maintain the integrity of intercellular contacts (16, 24), increased MLC phosphorylation is usually associated with loosening of the TJ and decreased transepithelial resistance (21, 54, 56). It has long been known that certain Na+-coupled membrane transport processes (e.g., Na+-glucose or Na+-amino acid cotransport) induce substantial increases in paracellular permeability in intestinal and renal epithelia (4, 55, 61). Remarkably, the same transport processes have been shown to induce increased myosin phosphorylation and the formation of a perijunctional contractile actomyosin ring at the apical pole (4, 37, 55). These structural changes were suggested to underlie the increase in TJ permeability (54). However, the mechanism whereby Na+-coupled membrane transport leads to enhanced MLC phosphorylation has not been elucidated. On the basis of our results that alanine activates Rho and provokes Y-27632-sensitive MLC phosphorylation, we propose the following mechanism. The initiation of the Na+-coupled electrogenic transport depolarizes the apical membrane, which leads to Rho activation and subsequent ROK-mediated myosin phosphorylation, results in increased TJ permeability. This intriguing mechanism would explain how transmembrane and paracellular transport processes are coupled and coordinated. Consistent with this proposition, recently a novel TJ-associated Rho-GEF was described, the overexpression of which resulted in augmented paracellular permeability (3). Future studies are warranted to explore the mechanism of depolarization-induced Rho activation and to validate the proposed membrane potential-dependent regulation of paracellular permeability.


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 ABSTRACT
 MATERIALS AND METHODS
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This work was supported by an operating grant from the Canadian Institutes of Health Research (CIHR) (to A. Kapus). K. Szászi is a recipient of a CIHR Senior Research fellowship, and C. Di Ciano-Oliveira is supported by a Canada Graduate Scholarship.


    FOOTNOTES
 

Address for reprint requests and other correspondence: A. Kapus, St. Michael’s Hospital, 30 Bond St., Queen Wing, Rm. 7009, Toronto, ON, Canada M5B 1W8 (e-mail: kapusa{at}smh.toronto.on.ca)

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

* K. Szászi and G. Sirokmány contributed equally to this work. Back


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