Departments of 1 Surgery, 3 Physiology, and 5 Pathology, University of Maryland School of Medicine and 2 Baltimore Veterans Affairs Medical Center, Baltimore, Maryland 21201; and 4 Department of Medicine, School of Medicine, University of California, San Diego, California 92103
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ABSTRACT |
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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
(Em) 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 -difluoromethylornithine
(DFMO) reduced whole cell K+ currents
[IK(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 on
IK(v), Em,
[Ca2+]cyt, and RhoA protein and restored cell
migration to normal. Elevation of [Ca2+]cyt
induced 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.
polyamines; intracellular calcium; guanosine 5'-triphosphate-binding protein; potassium channels; restitution; intestinal epithelial cells
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INTRODUCTION |
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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 (Em) 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 -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 [IK(v)], membrane potential (Em), [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).
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MATERIALS AND METHODS |
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Materials.
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
-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 15-20 were 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/cm2 in 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 IK(v), Em, [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.Electrophysiological measurements.
Whole cell K+ currents (IK) 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 M) 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 optimal
IK(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.
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+]cyt was 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 IK(v) or Em 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 IK(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 on day 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.
Statistical analysis. 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).
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RESULTS |
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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 IK(v) in IEC-6 cells.
Whole cell IK were elicited by depolarizing cells to a series of test potentials ranging from
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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+]cyt
in 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).
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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.
3A, 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. 1Bb). 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).
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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, A
and B, right).
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Effect of ionomycin on cell migration in polyamine-deficient cells.
Figure 7A 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. 7D). 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.
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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. 9A,
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. 9A, b vs. d). Spermidine added concomitantly with DFMO was able to maintain cell
migration at near-normal levels (Fig. 9A, d vs.
e). Treatment with C3 for 6 (Fig. 9A,
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.
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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.
10a), 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.
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DISCUSSION |
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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 cell
IK(v) in IEC-6 cells (Fig. 1). Decreased
IK(v) was associated with a depolarized
Em 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).
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Our findings strengthen the evidence that activation of Kv channels is involved in the regulation of polyamine-dependent intestinal epithelial cell migration by controlling Em 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 Em while the Ca2+ concentration gradient is constant (11, 13, 14, 18). Em is determined by transmembrane K+ permeability (PK) and the K+ gradient across the plasma membrane (18). The K+ gradient is maintained by Na+-K+-ATPase, and PK 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 Em in a variety of cell types (11, 13, 14). When a K+ channel closes or the number of total K+ channels decreases, Em 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, Em 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 IK(v) and depolarized Em in IEC-6 cells. Since IEC-6 cells do not express VDCC, the depolarized Em in polyamine-deficient cells decreases [Ca2+]cyt as 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 IK(v), Em, 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. 8 and 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 inhibits IK(v), depolarizes Em, 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.
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ACKNOWLEDGEMENTS |
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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).
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FOOTNOTES |
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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: jwang{at}smail.umaryland.edu).
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.
Received 11 July 2000; accepted in final form 2 November 2000.
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