Expression of voltage-gated K+ (Kv) channel genes is regulated by polyamines in intestinal epithelial cells (IEC-6 line), and Kv channel activity is involved in the regulation of cell migration during early restitution by controlling membrane potential (E m) and cytosolic free Ca2+concentration ([Ca2+]cyt). This study tests the hypothesis that RhoA of small GTPases is a downstream target of elevated [Ca2+]cyt following activation of K+ channels by increased polyamines in IEC-6 cells. Depletion of cellular polyamines by α-difluoromethylornithine (DFMO) reduced whole cell K+ currents [I K(v)] through Kv channels and caused membrane depolarization, which was associated with decreases in [Ca2+]cyt, RhoA protein, and cell migration. Exogenous polyamine spermidine reversed the effects of DFMO onI K(v), E m, [Ca2+]cyt, and RhoA protein and restored cell migration to normal. Elevation of [Ca2+]cytinduced by the Ca2+ ionophore ionomycin increased RhoA protein synthesis and stimulated cell migration, while removal of extracellular Ca2+ decreased RhoA protein synthesis, reduced protein stability, and inhibited cell motility. Decreased RhoA activity due to Clostridium botulinum exoenzyme C3 transferase inhibited formation of myosin II stress fibers and prevented restoration of cell migration by exogenous spermidine in polyamine-deficient cells. These findings suggest that polyamine-dependent cell migration is partially initiated by the formation of myosin II stress fibers as a result of Ca2+-activated RhoA activity.
- intracellular calcium
- guanosine 5′-triphosphate-binding protein
- potassium channels
- intestinal epithelial cells
the restoration of normal intestinal mucosal integrity (successful repair of wounds and ulcers) requires epithelial cell decisions that regulate signaling networks controlling gene expression, survival, migration, and proliferation. The process of intestinal epithelial restitution refers to resealing of superficial wounds after injury and occurs as a consequence of epithelial cell migration into the defect, a process independent of cell proliferation (31, 40, 44). This early rapid reepithelialization is a primary repair modality in the gastrointestinal tract and absolutely requires cellular polyamines (31, 44). Polyamines, including spermidine, spermine, and their precursor, putrescine, are organic cations found in all eukaryotic cells and have been implicated in a wide variety of physiological functions (23, 34, 50). The regulation of cellular polyamines is the point of central convergence for the multiple signaling pathways driving epithelial cell motility and proliferation (17, 34). Polyamines accelerate early mucosal restitution of gastric and duodenal mucosal stress erosions in vivo (23, 50, 51) and are essential for the stimulation of cell migration in an in vitro model (25, 26, 52) that mimics the early cell division-independent stage of epithelial restitution.
We (53) recently demonstrated that voltage-gated K+ (Kv) channels are involved in the regulation of polyamine-dependent intestinal epithelial cell migration after wounding by controlling membrane potential (E m) and cytosolic free Ca2+ concentration ([Ca2+]cyt). Polyamines stimulate expression of Kv channels in intestinal epithelial cells, whereas inhibition of ornithine decarboxylase (ODC; the rate-limiting enzyme for polyamine biosynthesis) with α-difluoromethylornithine (DFMO) attenuates Kv channel activity. Because intestinal epithelial cells do not express L-type voltage-dependent Ca2+ channels (VDCC), the polyamine-induced activation of Kv channels causes membrane hyperpolarization, enhances Ca2+ entry by increasing the driving force for Ca2+ influx, raises [Ca2+]cyt, and promotes cell migration during restitution. However, the precise mechanisms by which elevated [Ca2+]cyt mediates polyamine-dependent cell migration after wounding remain to be demonstrated.
The coordinated movement of epithelial cells is a complex process that depends on the cytoskeleton (10, 31). Changes in both the distribution and formation of the cytoskeleton alter the adhesion, spreading, and motility of cells (3, 20, 45). Recently, the regulation of cytoskeletal rearrangements required for directed cell migration has become focused on the Rho family of guanine nucleotide triphosphate (GTP)-binding proteins including RhoA, Rac, and Cdc42 (15, 19, 22, 24, 46). Rho proteins are members of the Ras superfamily of small GTP-binding proteins and function as molecular switches by cycling between an active GTP-bound state and an inactive GDP-bound state (15, 24, 41). Activation of Rho proteins, through GDP-GTP exchange, is stimulated by guanine nucleotide exchange factors, whereas inactivation of the proteins is promoted by GTPase-activating proteins (22, 41, 48). Increasing evidence indicates that activated Rho proteins interact with cellular target proteins or effectors to regulate a signal transduction pathway linking surface receptors to the formation of actomyosin stress fibers and focal adhesions (15, 30, 32, 39, 46). The transformation of RhoA from its inactive GDP-bound form to its active GTP-bound form activates Rho kinase, which results in the formation of actomyosin stress fibers by initiating myosin light chain phosphorylation (1, 19, 22). On the other hand, activation of Rac promotes de novo actin polymerization at the cell periphery to form lamellipodial extensions and membrane ruffles, and activation of Cdc42 results in actin polymerization to form filopodia or microspikes (15, 30).
The current study was undertaken to determine the role of RhoA protein in the cellular pathway leading to increased cell migration by elevated [Ca2+]cyt following the induction of K+ channel expression by polyamines during restitution in intestinal epithelial cells (IEC-6 cell line). First, we examined the effects of polyamine depletion on voltage-gated K+ currents [I K(v)], membrane potential (E m), [Ca2+]cyt, and RhoA protein expression in IEC-6 cells. Second, we determined whether manipulating [Ca2+]cyt, either by increase or decrease, altered RhoA protein expression and cell migration in the presence or absence of polyamines. Third, we investigated whether observed Ca2+-induced RhoA protein played a role in the formation of stress fibers and polyamine-dependent cell migration after wounding. Some of these data have been published in abstract form (38).
MATERIALS AND METHODS
Disposable culture ware was purchased from Corning Glass Works (Corning, NY). Tissue culture media and dialyzed fetal bovine serum (dFBS) were obtained from GIBCO-BRL (Gaithersburg, MD), and biochemicals were from Sigma (St. Louis, MO). The primary antibody, an affinity-purified rabbit polyclonal antibody against RhoA, was purchased from Santa Cruz Biotechnology (Santa Cruz, CA). The specific rabbit polyclonal antibody against nonmuscle myosin II was obtained from Biomedical Technologies (Stoughton, MA). Anti-rabbit immunoglobulin G (IgG), fluorescein isothiocyanate isomer (FITC) conjugate, and ionomycin were purchased from Sigma. Clostridium botulinum exoenzyme C3 transferase (C3) was obtained from Upstate Biotechnology (Lake Placid, NY), and α-difluoromethylornithine (DFMO) was a gift from the Merrell Dow Research Institute of Marion Merrell Dow (Cincinnati, OH).
Cell culture and general experimental protocol.
The IEC-6 cell line was purchased from American Type Culture Collection at passage 13. The cell line was derived from normal rat intestine and was developed and characterized by Quaroni et al. (37). IEC-6 cells originated from intestinal crypt cells, as judged by morphological and immunological criteria. They are nontumorigenic and retain the undifferentiated character of intestinal epithelial stem cells.
Stock cells were maintained in T-150 flasks in DMEM supplemented with 5% heat-inactivated FBS, 10 μg/ml insulin, and 50 μg/ml gentamicin sulfate. Flasks were incubated at 37°C in a humidified atmosphere of 90% air-10% CO2. Stock cells were subcultured once a week at 1:20, and medium was changed three times per week. The cells were restarted from original frozen stock every seven passages. Tests for mycoplasma were routinely negative, and passages 15–20were used in the experiments. There were no significant changes of biological function and characterization from passages 15–20.
The general protocol of the experiments and the methods used were similar to those described previously (52). Briefly, IEC-6 cells were plated at 6.25 × 104 cells/cm2in DMEM supplemented with 5% dFBS, 10 μg/ml insulin, and 50 μg/ml gentamicin sulfate. Cells were incubated in a humidified atmosphere at 37°C in 90% air-10% CO2 (vol/vol) for 24 h, and a period of different experimental treatments followed.
In the first series of studies, we examined the effects of polyamine depletion on I K(v), E m, [Ca2+]cyt, and RhoA protein expression in IEC-6 cells. The cells were grown in control cultures and in cultures containing either 5 mM DFMO or DFMO plus 5 μM spermidine for 4 days. The dishes were placed on ice, the monolayers were washed three times with ice-cold Dulbecco's PBS (D-PBS), and then different solutions were added according to the assays to be conducted.
In the second series of studies, we determined the effect of increasing [Ca2+]cyt on RhoA protein expression and cell migration in normal (without DFMO) and polyamine-depleted IEC-6 cells (with DFMO). The Ca2+ ionophore ionomycin was used to increase [Ca2+]cyt, and the measurements for [Ca2+]cyt, RhoA protein synthesis and stability, and cell migration were carried out at various times after treatment with ionomycin with or without wounding.
In the third series of studies, we investigated whether observed Ca2+-induced RhoA protein played a role in the organization of nonmuscle myosin II and polyamine-dependent cell migration after wounding. C3 is an inhibitor for GTP-binding proteins and has been shown to specifically inhibit RhoA activity in epithelial cells (28, 42). C3 was added immediately after wounding to control cultures and to cultures in which ODC was inhibited with DFMO and supplemented with 5 μM exogenous spermidine. Cellular distribution of nonmuscle myosin II and cell migration was assayed 6 and 8 h after treatment.
Whole cell K+ currents (I K) were recorded with an Axopatch-1D amplifier and a DigiData 1200 interface (Axon Instruments, Foster City, CA) by using patch-clamp techniques (54). Patch pipettes (2–4 MΩ) were made on a Sutter electrode puller with the use of borosilicate glass tubes and were fire-polished on a Narishige microforge. Step-pulse protocols and data acquisition were performed with pCLAMP software. Currents were filtered at 1–2 kHz (−3 dB) and digitized at 2–4 kHz with the Axopatch-1D amplifier. To record optimalI K(v), CaCl2 was replaced by equimolar MgCl2 in the bath solution. Series resistance and capacitance were routinely compensated (for 60–80%) by adjusting the internal circuitry of the patch-clamp amplifier. Leakage currents were subtracted with the P/4 protocol in pCLAMP software.
E m in single IEC-6 cells was measured in current-clamp mode (I = 0) by using whole cell patch-clamp techniques. All experiments were performed at room temperature (22–24°C).
Measurement of [Ca2+]cyt.
Details of the digital imaging methods employed for measuring [Ca2+]cyt have been published previously (55). Briefly, IEC-6 cells were plated on 25-mm coverslips and were incubated in culture medium containing 3.3 μM fura 2-AM for 30–40 min at room temperature (22–24°C) under an atmosphere of 10% CO2 in air. The fura 2-loaded cells were then superfused with standard bath solution for 20–30 min at 32–24°C to wash away extracellular dye and to permit intracellular esterases to cleave cytosolic fura 2-AM into active fura 2. Fura 2 fluorescence (510-nm emission; 380- and 360-nm excitation) from the cells and background fluorescence were imaged with the use of a Nikon Diaphot microscope equipped for epifluorescence. Fluorescent images were obtained with a microchannel plate image intensifier (Amperex XX1381; Opelco, Washington, DC) coupled by fiber optics to a Pulnix charge-coupled device video camera (Stanford Photonics, Stanford, CA).
Image acquisition and analysis were performed with a MetaMorph Imaging System (Universal Imaging). Video frames containing images of fura 2 fluorescence from cells and the corresponding background images (fluorescence from fields devoid of cells) were digitized at a resolution of 512 horizontal × 480 vertical pixels and eight bits with the use of a Matrix LC imaging board operating in an IBM-compatible computer. To improve the signal-to-noise ratio, 8–32 consecutive video frames were usually averaged at a video frame rate of 30 frames/s. Images were acquired at a rate of one averaged image every 3 s when [Ca2+]cytwas changing and one averaged every 60 s when [Ca2+]cyt was relatively constant. [Ca2+]cyt was calculated from fura 2 fluorescent emission excited at 380 and 360 nm by using the ratio method (35). In most experiments, multiple cells (usually 10–15 cells) were imaged in a single field, and one arbitrarily chosen peripheral cytosolic area (4–6 × 4–6 pixels) from each cell was spatially averaged.
Solution and reagents.
A coverslip containing the cells was positioned in the recording chamber (∼0.75 ml) and superfused (2–3 ml/min) with the standard extracellular (bath) physiological salt solution (PSS) for recording either I K(v) or E m or for measuring [Ca2+]cyt. The PSS contained (in mM) 141 NaCl, 4.7 KCl, 1.8 CaCl2, 1.2 MgCl2, 10 HEPES, and 10 glucose buffered to pH 7.4 with 5 M NaOH. In Ca2+-free PSS, CaCl2 was replaced by equimolar MgCl2, and 0.1 mM EGTA was added to chelate residual Ca2+ in the Ca2+-free DMEM. The internal (pipette) solution for recording I K(v) contained (in mM) 125 KCl, 4 MgCl2, 10 HEPES, 10 EGTA, and 5 Na2-ATP (pH 7.2).
Measurement of cell migration.
The migration assays were carried out as described in earlier publications (52, 53). IEC-6 cells were plated at 6.25 × 104/cm2 in DMEM/dFBS with or without 5 mM DFMO and 5 μM spermidine on 60-mm dishes thinly coated with Matrigel according to the manufacturer's instructions and were incubated as described for stock cultures. The cells were fed onday 2 and migration tested on day 4. To initiate migration, we scratched the cell layer with a single-edge razor blade cut to ∼27 mm in length. The scratch began at the diameter of the dish and extended over an area 7–10 mm wide. After the scratch was made, the cell layer was immediately photographed. Care was taken to include some identifying mark on the dish to serve as a future reference point. The dishes were then returned to the incubator, and cell migration was allowed to occur over the denuded area for different time periods. At the end of the desired time, the dishes were removed and rephotographed in the same area as before, and the migrating cells were counted by means of an eyepiece reticle. The migrating cells in six contiguous 0.1-mm squares were counted at ×100 magnification beginning at the scratch line and extending as far out as the cells had migrated. All experiments were carried out in triplicate, and the results were reported as the number of migrating cells per millimeter of scratch. An inverted phase-contrast microscope with an attached Polaroid camera was used for the cell counts and photographs.
Western blot analysis.
Cell samples, dissolved in sodium dodecyl sulfate (SDS) sample buffer (250 mM Tris · HCl, pH 6.8, 2% SDS, 20% glycerol, and 5% mercaptoethanol), were sonicated and centrifuged at 2,000 rpm for 15 min. The protein concentration of the supernatant was measured by the methods described by Bradford (6), and each lane was loaded with 25 μg of protein equivalent. The supernatant was boiled for 5 min and then subjected to electrophoresis on 10% acrylamide gels according to Laemmli (21). Briefly, after the transfer of protein onto nitrocellulose filters, the filters were incubated overnight at 4°C in 5% nonfat dry milk in 1× PBS-Tween 20 [PBS-T; 15 mM NaH2PO4, 80 mM Na2HPO4, 1.5 M NaCl (pH 7.5), and 0.5% (vol/vol) Tween 20]. Immunological evaluation was then performed for 90 min in 1% BSA-PBS-T buffer containing affinity-purified antibody against RhoA protein. The filters were subsequently washed with 1× PBS-T and incubated for 1 h with an IgG second antibody conjugated to peroxidase by protein cross-linking with 0.2% glutaraldehyde. After extensive washing with 1× PBS-T, the immunocomplexes on the filters were reacted for 1 min with chemiluminescence reagent (NEL-100; NEN). Finally, the filters were placed in a plastic sheet protector and exposed to autoradiography film for 30 or 60 s.
Measurement of RhoA protein synthesis.
The RhoA protein synthesis was examined by using a [35S]methionine labeling technique (2). Cells were grown in control cultures and in cultures containing 5 mM DFMO for 4 days, washed with the methionine-free medium, and then exposed to 1 μM ionomycin for 2 h. The media were replaced by the cultures containing [35S]methionine (100 μCi/ml) in the presence of ionomycin. The cells were harvested 4 h after incubation with [35S]methionine. In the experiments dealing with removal of extracellular Ca2+ from the culture media, cells were grown in the presence or absence of DFMO for 4 days, exposed to the Ca2+-free medium for 2 h, and then pulse-labeled by incubation with the Ca2+-free medium containing [35S]methionine (100 μCi/ml) for 4 h. The cells were rinsed with cold D-PBS containing 2 mM methionine and were harvested by scraping. Cells were then disrupted by being passed through a 21-gauge syringe needle, and the suspension was centrifuged at 4°C for 10 min. The supernatant (cell lysate) was collected and incubated with a control mouse IgG together with the IgG1 protein G PLUS-agarose for 30 min on a rocker platform with a rotating device at 4°C. Beads were isolated by centrifugation, and the preclear cell lysate was transferred into a new tube. The cell lysate (400 μg) was incubated with anti-RhoA antibody (4 μg) for 1 h at 4°C. The protein G PLUS-agarose was added, and the samples were incubated overnight. Immunoprecipitates were carefully collected after centrifugation at 2,500 rpm for 5 min, and pellets were washed with cold PBS and resuspended in 30 μl of 1× electrophoresis sample buffer. The supernatant fluid was analyzed by SDS-PAGE followed by autoradiography.
Nonmuscle myosin II staining.
Cells were plated at 6.25 × 104/cm2 in chambered slides thinly coated with Matrigel according to the manufacturer's instructions and incubated with medium containing DMEM plus 5% dFBS, 10 μg/ml insulin, and 50 μg/ml gentamicin sulfate. DFMO at a dose of 5 mM with or without 5 μM spermidine was added as treatment. On day 4 after initial plating, approximately one-third of the cell layers were removed diagonally across the chamber slide with a razor blade. The medium was changed to remove floating or damaged cells, and the cells were returned to the incubator for 6 h, during which they began to migrate over the denuded area. The immunofluorescence procedure was carried out according to the method of Vielkind and Swierenga (49) with minor changes. Briefly, the cells were washed with D-PBS and then with D-PBS without Ca2+ and Mg2+(D-PBS-Ca2+-Mg2+) and fixed for 15 min at room temperature in 4% paraformaldehyde diluted with D-PBS. The cells were postfixed for 5 min with ice-cold methanol. The cells were rehydrated in D-PBS-Ca2+-Mg2+ for 30 min at room temperature and then incubated for 1.5 h with the rabbit anti-myosin II IgG used for Western blot analysis and then with anti-rabbit IgG-FITC conjugate for 1 h. The primary antibody recognizes the 200-kDa nonmuscle myosin II in immunoblots of IEC-6 cell extracts and does not cross-react with other cytoskeletal proteins. Nonspecific slides were incubated without antibody to nonmuscle myosin II. After three washes with D-PBS, the slides were mounted with VectaShield mounting medium (Vector Laboratories, Burlingame, CA). Slides were viewed through a Zeiss confocal microscope (model LSM410).
HPLC analysis of cellular polyamines.
The cellular polyamine content was determined as previously described (51). Briefly, after the cells were washed three times with ice-cold D-PBS, 0.5 M perchloric acid was added, and the cells were frozen at −80°C until ready for extraction, dansylation, and HPLC. The standard curve encompassed 0.31–10 μM. Values that fell >25% below the curve were considered undetectable. Protein was determined by the Bradford method (6). The results are expressed as nanomoles of polyamines per milligram of protein.
All data are expressed as means ± SE from six dishes. Autoradiographic and immunofluorescence labeling results were repeated three times. The significance of the difference between means was determined by analysis of variance. The level of significance was determined by using Dunnett's multiple range test (16).
Effect of polyamine depletion on IK(v) and Em in IEC-6 cells.
Exposure of IEC-6 cells to 5 mM DFMO for 4 days, which totally inhibited ODC activity (50, 51), almost completely depleted cellular polyamines. Putrescine and spermidine were undetectable, while spermine was decreased by >65% on day 4 in the DFMO-treated cells (36). Our previous studies (53) showed that depletion of cellular polyamines by DFMO significantly inhibited Kv1.1 channel mRNA and protein expression, which was completely prevented by exogenous spermidine. Using the patch-clamp technique, we further examined the effect of polyamine depletion on the I K(v) in IEC-6 cells.
Whole cell I K were elicited by depolarizing cells to a series of test potentials ranging from −40 to +80 mV in 20-mV increments from a holding potential of −70 mV. Exposure of the cells to 5 mM 4-aminopyridine (4-AP), a K+ channel blocker, significantly and reversibly reduced the whole cell currents, indicating that the currents were 4-AP-sensitiveI K(v). Consistent with the inhibitory effect on the Kv1.1 channel gene expression, polyamine depletion by DFMO (for 4 days) markedly decreased I K(v) (Fig.1, A and Ba), whereas pretreatment of the cells with spermidine completely reversed the inhibitory effect of DFMO on I K(v) (Fig. 1,Ac and Ba). The current-voltage relationships (Fig. 1 Ba) indicate that the DFMO-induced decrease inI K(v) appeared to be voltage dependent; the inhibitory effect was greater at more positive potentials. DFMO treatment decreased I K(v) by 44, 56, 68, and 73% at −40, 0, +40, and +80 mV, respectively. These results suggest that, in addition to regulating the Kv channel expression, polyamines also may modulate the channel function.
Under resting conditions, the membrane input resistance was very high in IEC-6 cells (8.1 ± 3.6 MΩ). Therefore, a small change inI K(v) would cause a large change inE m. Indeed, polyamine depletion by DFMO caused significant membrane depolarization in IEC-6 cells (Fig.1 Bb). Similar to the effect on I K(v), pretreatment of the cells with spermidine abolished the depolarizing effect of DFMO (Fig. 1 Bb). These results suggest that a polyamine depletion-mediated decrease in I K(v)is sufficient to cause a significant membrane depolarization in IEC-6 cells.
Effect of depolarized Em on [Ca2+]cyt and RhoA protein expression.
IEC-6 cells do not express VDCC (53); therefore, membrane depolarization would reduce the Ca2+ driving force, inhibit Ca2+ influx through voltage-independent Ca2+-permeable channels, and decrease [Ca2+]cyt (14, 56, 57). Indeed, membrane depolarization in polyamine-deficient cells significantly decreased the resting [Ca2+]cyt, which was associated with an inhibition of RhoA protein expression (Fig.2). [Ca2+]cytin DFMO-treated cells was ∼50% of normal values (without DFMO), while RhoA protein levels were ∼20% of the control (Fig. 2). Addition of spermidine to the cultures in the presence of DFMO not only reversed the inhibitory effects of polyamine depletion on [Ca2+]cyt but also restored RhoA protein to normal levels. Interestingly, removal of extracellular Ca2+from the culture medium completely prevented the restoration of RhoA protein expression by exogenous spermidine in polyamine-deficient cells (Fig. 2, lane 4). There was no apparent loss of cell viability in cells treated with DFMO alone, DFMO plus spermidine, or spermidine plus the Ca2+-free medium containing DFMO as measured by trypan blue staining method (data not shown).
Effect of increasing [Ca2+]cyt on RhoA protein expression.
To further determine the relationship between [Ca2+]cyt and RhoA activity in intestinal epithelial cells, we examined the effect of increasing [Ca2+]cyt by the Ca2+ionophore ionomycin on RhoA protein expression in normal (without DFMO) and polyamine-deficient cells. Addition of 1 μM ionomycin reversibly increased [Ca2+]cyt by promoting Ca2+ influx in all three groups (Fig.3). In control cells (without DFMO), [Ca2+]cyt was remarkably increased after exposure to ionomycin for 5 min (from 95.7 ± 7.3 to 173.6 ± 15.2 nM, n = 12, P < 0.05). When ionomycin was washed out, [Ca2+]cyt rapidly returned to basal levels. In DFMO-treated cells, the basal level of [Ca2+]cyt was lower than that observed in control cells (from 95.7 ± 7.3 to 59.1 ± 3.9 nM, n = 12, P < 0.05). Exposure to ionomycin also increased [Ca2+]cyt in the presence of DFMO (from 59.1 ± 3.9 to 112.3 ± 9.4 nM,n = 12, P < 0.05), but this response was significantly reduced compared with that of controls (Fig.3 A, left vs. middle). This reduced response of DFMO-treated cells to ionomycin was apparently due to the decreased Ca2+ driving force as a result of membrane depolarization (Fig. 1 Bb). Spermidine given together with DFMO almost completely overcame the change in basal [Ca2+]cyt (from 59.1 ± 3.7 to 84.5 ± 6.9 nM, n = 12, P < 0.05). On the other hand, an ionomycin-mediated increase in [Ca2+]cyt in cells treated with DFMO plus spermidine (133.5 ± 9.2 nM, n = 12) was lower than that observed in control cells exposed to ionomycin (173.6 ± 15.2 nM, n = 12).
Consistent with the augmenting effect on [Ca2+]cyt, administration of ionomycin also increased RhoA protein expression in control and polyamine-deficient cells (Fig. 4). RhoA protein levels in control cells were increased by exposure to ionomycin for 4 and 6 h (from 1 ± 0.05 to 1.75 ± 0.07 and 1.8 ± 0.06, respectively, n = 3, P < 0.05). Although the basal level of RhoA protein in polyamine-deficient cells was lower than that in controls, ionomycin significantly increased RhoA protein in the presence of DFMO (from 0.2 ± 0.01 at 0 h to 0.87 ± 0.06 at 4 h, and 0.85 ± 0.05 at 6 h, n = 3, P< 0.05). The ionomycin-induced RhoA protein levels (values in ionomycin-treated cells − basal value) in DFMO-treated cells were similar to those observed in control cells exposed to ionomycin (e.g., 0.82 ± 0.04 vs. 0.66 ± 0.03 at 6 h). These results indicate that cellular polyamines were involved in the regulation of RhoA activity and that elevation of [Ca2+]cytfollowing activation of Kv channels by cellular polyamine induced RhoA expression in intestinal epithelial cells.
Effect of [Ca2+]cyt on RhoA protein synthesis and stability.
To determine the mechanism through which [Ca2+]cyt regulates RhoA expression, we examined changes in the rate of RhoA protein synthesis and the protein stability when [Ca2+]cyt was increased by ionomycin or decreased by removal of extracellular Ca2+from the culture media. In control cells (without DFMO), the rate of newly synthesized RhoA protein was increased by ionomycin but decreased after exposure to the Ca2+-free medium (Fig.5, left). Although the basal level of RhoA protein synthesis in DFMO-treated cells was low, the response of newly synthesized RhoA protein to ionomycin in DFMO-treated cells was similar to that observed in control cells. In addition, removal of extracellular Ca2+ from the medium containing DFMO further decreased the RhoA protein synthesis (Fig. 5, Aand B, right).
Results presented in Fig.6 show that the stability of RhoA protein also was regulated by [Ca2+]cyt in intestinal epithelial cells. The RhoA protein level of control cells declined gradually after protein synthesis was inhibited by administration of cycloheximide. Exposure to ionomycin slightly increased the stability of the RhoA protein (Fig.6 A, a vs. b), but the difference (∼8%) was not statistically significant (Fig. 6 B). In contrast, removal of extracellular Ca2+ from the culture medium significantly decreased the RhoA protein stability compared with that observed in control cells (Fig. 6 A, a vs.c). These data indicate that [Ca2+]cyt not only regulates RhoA protein synthesis but also is involved in the maintenance of RhoA protein stability in IEC-6 cells. These findings suggest that [Ca2+]cyt modulates RhoA expression, at least partially, through posttranscriptional regulation.
Effect of ionomycin on cell migration in polyamine-deficient cells.
Figure 7 A clearly shows that polyamine depletion by treatment with DFMO dramatically decreased cell migration in IEC-6 cells. The number of cells migrating in the DFMO-treated cells was decreased by ∼75% when counted at 4, 6, and 8 h after wounding. In the presence of DFMO, exogenous spermidine restored cell migration nearly to normal levels. Treatment with ionomycin also increased cell migration in all three groups. Ionomycin given immediately after wounding increased the rate of cell migration by ∼20% in control cells (Fig. 7, B and E,a vs. b) and in cells treated with DFMO plus spermidine (Fig. 7 D). Cell migration in DFMO-treated cells also was increased by ionomycin (Fig. 7, C and E,c vs. d). At all the time points studied, the rates of cell migration in DFMO-treated cells exposed to ionomycin were significantly increased compared with those observed in cells treated with DFMO alone. On the other hand, as shown in a previous publication (53), removal of extracellular Ca2+from the culture media significantly decreased the rate of migration in IEC-6 cells (data not shown). These results indicate that a rise in [Ca2+]cyt is required for the stimulation of migration by polyamines in intestinal epithelial cells.
Effect of decreased RhoA activity on cell migration.
To determine the role of Ca2+-induced RhoA in polyamine-dependent cell migration after wounding, we carried out experiments in which C3, a specific inhibitor of the Rho proteins (28, 42), was used. We examined whether inhibition of RhoA activity by treatment with C3 altered the rates of cell migration in control cells and in cells treated with DFMO plus spermidine. Administration of C3 at the concentration of 10 μg/ml for 6 h not only decreased levels of RhoA protein in normal cells (Fig. 8,left) but also prevented the restoration of RhoA protein by exogenous spermidine in DFMO-treated cells (Fig. 8, right). The levels of RhoA protein were decreased by ∼40% after treatment with C3 in normal cells and in cells treated with DFMO plus spermidine. As shown in Fig.9 A,b vs. c, the migration was also decreased when control cells (without DFMO) were exposed to C3. In addition, depletion of cellular polyamines by DFMO decreased cell migration (Fig. 9 A, b vs. d). Spermidine added concomitantly with DFMO was able to maintain cell migration at near-normal levels (Fig. 9 A, d vs.e). Treatment with C3 for 6 (Fig. 9 A,e vs. f, and B) and 8 h (data not shown) during the period of cell migration prevented restoration of cell migration by spermidine in DFMO-treated cells. These results clearly indicate that increased RhoA activity due to elevated [Ca2+]cyt is essential for polyamine-dependent migration in intestinal epithelial cells.
Effect of RhoA activity on distribution of nonmuscle myosin II.
The cellular distribution of nonmuscle myosin II protein was monitored with immunostaining techniques after incubation with 10 μg/ml C3. In control cells (Fig.10 a), long stress fibers traversed the cytoplasm, and a thick network of cortical myosin II fibers was just beneath the plasma membrane. Exposure of control cells to C3 for 6 h significantly decreased the formation of myosin II stress fibers (Fig. 10, a vs. b). The distribution of nonmuscle myosin II stress fibers was sparse and devoid of long stress fiber formation. Polyamine depletion by treatment with DFMO also resulted in reorganization of nonmuscle myosin II in IEC-6 cells (Fig. 10, a vs. c). In DFMO-treated cells, long stress fibers disappeared, and no distinct myosin II stress fibers were observed. Spermidine given together with DFMO restored the distribution of nonmuscle myosin II to near normal (Fig. 10,c vs. d). The distribution of nonmuscle myosin II in cells grown in the presence of DFMO plus spermidine was indistinguishable from that of control cells (Fig. 10, a vs.d). Treatment with C3 for 6 and 8 h completely prevented the restoration of the distribution of nonmuscle myosin II by exogenous spermidine in polyamine-deficient cells (Fig.10, d vs. e and f). The number of long stress fibers was greatly reduced, and in some cells they appeared to be absent.
Recently, we demonstrated that expression of Kv channels is regulated by cellular polyamines in intestinal epithelial cells and that inhibition of polyamine synthesis by treatment with DFMO decreases Kv1.1 channel gene expression and inhibits cell migration during restitution (53). In this study, we have advanced our understanding of the role of polyamines in the regulation of Kv channel activity by demonstrating that polyamine depletion decreases whole cellI K(v) in IEC-6 cells (Fig. 1). DecreasedI K(v) was associated with a depolarizedE m and reduced resting [Ca2+]cyt in polyamine-deficient cells. The most significant new finding reported in this article, however, is that RhoA plays an important role in the cellular pathway leading to increased cell migration by elevated [Ca2+]cyt following activation of Kv channel expression by polyamines. Reduction of [Ca2+]cyt by polyamine depletion or removal of extracellular Ca2+ decreased the levels of RhoA protein (Fig. 2), whereas an increase in [Ca2+]cyt by the Ca2+ ionophore ionomycin increased RhoA protein expression (Fig. 4). Furthermore, elevation of [Ca2+]cyt induced by ionomycin stimulated RhoA protein synthesis, while reduction of [Ca2+]cyt following removal of extracellular Ca2+ from the culture media inhibited RhoA protein synthesis and decreased the protein stability (Figs. 5 and 6). Treatment with ionomycin also promoted cell migration in controls and polyamine-deficient cells (Fig. 7). Inhibition of RhoA activity by C3 not only decreased the formation of myosin II stress fibers but also prevented the restoration of cell migration by exogenous spermidine in polyamine-deficient cells (Figs. 9 and 10). These results suggest that elevated [Ca2+]cyt, due to the activation of Kv channels, increases the RhoA activity, which subsequently results in the formation of myosin II stress fibers and stimulates cell migration following an increase in cellular polyamines (Fig.11).
Our findings strengthen the evidence that activation of Kv channels is involved in the regulation of polyamine-dependent intestinal epithelial cell migration by controlling E m and [Ca2+]cyt. Cytoplasmic Ca2+ is an important intracellular second messenger that regulates a large number of physiological functions (4, 47). [Ca2+]cyt undergoes rapid and often substantial fluctuations in response to extracellular first messengers binding to their cognate receptors on target cells. In contrast, extracellular ionized Ca2+ concentration is maintained stably at 1.6∼1.8 mM, ∼10,000–20,000-fold higher than the resting [Ca2+]cyt (50–150 nM) in both excitable and nonexcitable cells. The transmembrane Ca2+electrochemical gradient provides a seemingly inexhaustible supply of Ca2+ for its diverse intracellular function. An increase in [Ca2+]cyt is a trigger for cell migration in a variety of cell types, whereas a decrease in [Ca2+]cyt inhibits cell movement (4, 5,9).
[Ca2+]cyt is controlled by Ca2+influx through Ca2+-permeable channels in the plasma membrane and Ca2+ release from intracellular Ca2+ stores (5, 7, 35). Ca2+influx depends on the Ca2+ driving force (i.e., the electrochemical gradient across the plasma membrane), which is predominantly regulated by E m while the Ca2+ concentration gradient is constant (11, 13, 14,18). E m is determined by transmembrane K+ permeability (P K) and the K+ gradient across the plasma membrane (18). The K+ gradient is maintained by Na+-K+-ATPase, and P K is directly related to the function and number of membrane K+channels. It has been shown that activity of Kv channels is a major determinant of resting E m in a variety of cell types (11, 13, 14). When a K+ channel closes or the number of total K+ channels decreases,E m becomes less negative (i.e., depolarized) and the Ca2+ driving force decreases. When a K+channel opens or the number of total K+ channels increases,E m becomes more negative (i.e., hyperpolarized) and the Ca2+ driving force increases (50). In nonexcitable cells (e.g., intestinal epithelial cells and lymphocytes) that do not express VDCC (14, 29, 53), membrane hyperpolarization raises [Ca2+]cyt by increasing the Ca2+ driving force, whereas membrane depolarization reduces [Ca2+]cyt by decreasing the Ca2+ driving force. Nevertheless, in excitable cells (e.g., neurons and muscle cells) that highly express VDCC (27, 47), membrane depolarization increases [Ca2+]cyt by opening VDCC.
The current studies and our previous findings (53) have demonstrated that polyamines are required for the stimulation of intestinal epithelial cell migration after wounding in association with their ability to regulate [Ca2+]cyt. Inhibition of Kv channel gene expression following a depletion of cellular polyamines decreased I K(v) and depolarized E m in IEC-6 cells. Since IEC-6 cells do not express VDCC, the depolarized E m in polyamine-deficient cells decreases [Ca2+]cytas a result of the reduced driving force for Ca2+ influx and inhibits cell migration (53). Exogenous spermidine not only reverses the effect of DFMO on I K(v),E m, and [Ca2+]cyt but also restores cell migration to normal. Removal of extracellular Ca2+ or blockade of Kv channels inhibits normal cell migration and prevents the restoration of cell migration by exogenous spermidine in the presence of DFMO. These results clearly indicate that elevated [Ca2+]cyt is a major mediator for the stimulation of cell migration following an increase in cellular polyamines.
The observations from the current study imply that RhoA of small GTPases is a downstream target of elevated [Ca2+]cyt following activation of Kv channels by polyamines in migrating epithelial cells. Decreased [Ca2+]cyt following inactivation of Kv channels in polyamine-deficient cells was associated with a decrease in RhoA protein (Fig. 2), whereas an increase in [Ca2+]cyt by ionomycin promoted RhoA protein expression (Fig. 4). Figures 5 and 6 further show that posttranscriptional regulation appears to be a key factor increasing the levels of RhoA protein following an elevation in [Ca2+]cyt. Elevation of [Ca2+]cyt induced by ionomycin significantly stimulated the RhoA protein synthesis regardless of the presence or absence of cellular polyamines and slightly slowed down the degradation of RhoA protein. In contrast, reduction of [Ca2+]cyt following removal of extracellular Ca2+ from the culture media not only inhibited the newly synthesized RhoA protein but also decreased the protein stability.
Increased RhoA activity due to elevated [Ca2+]cyt plays a critical role in polyamine-dependent cell migration during early epithelial restitution. Decreased RhoA activity by treatment with C3 inhibits normal cell migration after wounding (in the absence of DFMO) (Figs. 8and 9). These results are consistent with data from other investigators (33, 42), who have found that Rho plays an important role in gastrointestinal mucosal healing and that inactivated RhoA by either treatment with C3 or microinjection of a dominant negative form of RhoA inhibits the rate of normal intestinal epithelial cell migration. An interesting and extended finding obtained in the current study is that decreased RhoA activity also prevents the restoration of cell migration by exogenous spermidine in polyamine-deficient cells (Figs. 8 and 9). Our observations that decreased RhoA activity inhibits migration in cells treated with DFMO and spermidine strongly support the contention that augmented activity of Kv channels following an increase in polyamines results in the stimulation of intestinal epithelial cell migration through the Ca2+-RhoA signaling pathway. In addition, it is not clear at present whether other members of the mammalian Rho subfamily, including RhoB, RhoC, RhoD, RhoE, and RhoG, Rac1 and Rac2, and Cdc42 (12, 15), are regulated by [Ca2+]cyt alterations and are involved in the signal pathway of polyamine-dependent cell migration. For example, it has been shown that the cross talk between Rho and Rac proteins is an important factor controlling the cellular phenotype (8) and that cell motility is dependent on the balance between Rho and Rac activities (8, 15). However, whether the interaction between Rho and Rac proteins plays a role in polyamine-dependent intestinal epithelial cell migration remains to be elucidated.
Our results also show that RhoA regulates polyamine-dependent intestinal epithelial cell migration at least partially by altering the formation of actomyosin stress fibers. After exposure of control cells and cells treated with DFMO plus spermidine to C3, the number of long stress fibers of myosin II decreased significantly, and in some cells they disappeared completely from the cytoplasm, as observed in cells treated with DFMO alone (Fig. 10). Previous reports have shown that RhoA activation has a distinctive effect on the formation of actomyosin stress fibers in many other cell types, including fibroblasts, endothelial cells, astrocytes, and circulating cells such as lymphocytes, mast cells, and platelets (15, 24,46).
In summary, the results obtained from this study demonstrate that RhoA is implicated in the signaling pathway of Ca2+-mediated intestinal epithelial cell migration following activation of Kv channels by increased polyamines during restitution (Fig. 11). Depletion of cellular polyamines by DFMO inhibitsI K(v), depolarizes E m, and reduces [Ca2+]c, which is paralleled by a decrease in RhoA protein in IEC-6 cells. An increase in [Ca2+]cyt promotes RhoA protein expression and stimulates cell migration in the absence of cellular polyamines. Decreased RhoA activity caused by treatment with C3 inhibits the formation of stress fibers and impairs cell migration after wounding. Together, the results reported in the current study and previous studies (53) establish a specific Ca2+-mediated pathway of RhoA activation that is regulated by cellular polyamines. Increased cellular polyamines stimulate Kv channel expression, result in membrane hyperpolarization, and increase the driving force of Ca2+ influx, thus raising [Ca2+]cyt. The Ca2+-induced activation of RhoA increases the formation of actomyosin stress fibers and stimulates intestinal epithelial migration during the early phase of mucosal restitution.
This work was supported by National Heart, Lung, and Blood Institute Grants HL-54043 and HL-64945 (to J. X.-J. Yuan) and National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-45314 and DK-57819 (to J.-Y. Wang) and by a Merit Review Grant from the Department of Veterans Affairs (to J.-Y. Wang). J. X.-J. Yuan is an Established Investigator of the American Heart Association (974009N).
Address for reprint requests and other correspondence: J.-Y. Wang, Dept. of Surgery, Baltimore Veterans Affairs Medical Center, 10 North Greene St., Baltimore, MD 21201 (E-mail:).
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- Copyright © 2001 the American Physiological Society