Departments of 1 Surgery and 4 Physiology, University of Maryland School of Medicine and 2 Baltimore Veterans Affairs Medical Center, Baltimore, Maryland 21201; and 3 Department of Medicine, School of Medicine, University of California, San Diego, California 92103
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ABSTRACT |
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Polyamines are essential for cell migration
during early mucosal restitution after wounding in the gastrointestinal
tract. Activity of voltage-gated K+ channels (Kv) controls
membrane potential (Em) that regulates cytoplasmic
free Ca2+ concentration
([Ca2+]cyt) by governing the
driving force for Ca2+ influx. This study determined
whether polyamines are required for the stimulation of cell migration
by altering K+ channel gene expression,
Em, and
[Ca2+]cyt in intestinal epithelial
cells (IEC-6). The specific inhibitor of polyamine synthesis,
-difluoromethylornithine (DFMO, 5 mM), depleted cellular
polyamines (putrescine, spermidine, and spermine), selectively
inhibited Kv1.1 channel (a delayed-rectifier Kv channel) expression,
and resulted in membrane depolarization. Because IEC-6 cells did not
express voltage-gated Ca2+ channels, the depolarized
Em in DFMO-treated cells decreased [Ca2+]cyt as a result of reduced
driving force for Ca2+ influx through capacitative
Ca2+ entry. Migration was reduced by 80% in the
polyamine-deficient cells. Exogenous spermidine not only reversed the
effects of DFMO on Kv1.1 channel expression, Em,
and [Ca2+]cyt but also restored
cell migration to normal. Removal of extracellular Ca2+ or
blockade of Kv channels (by 4-aminopyridine, 1-5 mM) significantly inhibited normal cell migration and prevented the restoration of cell
migration by exogenous spermidine in polyamine-deficient cells. These
results suggest that polyamine-dependent intestinal epithelial cell
migration may be due partially to an increase of Kv1.1 channel
expression. The subsequent membrane hyperpolarization raises
[Ca2+]cyt by increasing the driving
force (the electrochemical gradient) for Ca2+ influx and
thus stimulates cell migration.
polyamines; voltage-gated potassium channels; intracellular calcium; membrane potential; intestinal epithelial cells
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INTRODUCTION |
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EPITHELIAL CELLS line the gastrointestinal mucosa and form an important barrier to a wide array of noxious substances in the lumen. The process of mucosal restitution refers to resealing of superficial wounds to this barrier, which occurs as a consequence of epithelial cell migration into the defect, a process that does not require epithelial cell proliferation (40, 41). This early mucosal restitution is a primary repair modality in the gastrointestinal tract and occurs both in vivo (13, 27) and in vitro (33, 40) after wounding. Increasing evidence has demonstrated that the cellular polyamines spermidine and spermine and their precursor putrescine play an important role in the regulation of mucosal restitution and are essential for the maintenance of gastrointestinal mucosal integrity (24, 47). Polyamines accelerate early mucosal restitution of gastric and duodenal mucosal stress erosions in vivo (46, 47) and are required for the stimulation of cell migration in an in vitro (26, 49) model that mimics the early cell division-independent stages of epithelial restitution. The precise mechanism involved in the polyamide-dependent cell migration at cellular and molecular levels is poorly understood.
The ability of intestinal epithelial cells to adopt a variety of shapes and to carry out coordinated and directed movements is a complex process that is regulated through multiple pathways (11, 33). Restitution of wounds in an in vitro model is the result of a series of coordinated cellular events, including lamellipodial extension and retraction, cortical transport of myosin and actin, contraction of transverse fibers, and tail retraction (11, 23, 33). A rise of cytosolic free Ca2+ concentration ([Ca2+]cyt) is a major trigger for cell contraction (5, 10, 44) and is an important stimulus for cell migration (4, 37). Activation of Ca2+/calmodulin-dependent protein kinase II is an important intracellular event that mediates cell migration (4, 37). Indeed, the ability of cells to migrate in response to various factors and chemoatractants can be significantly suppressed by removal of extracellular Ca2+ and/or depletion of intracellularly stored Ca2+ (4, 7, 37).
[Ca2+]cyt is controlled by
Ca2+ influx through Ca2+-permeable channels in
the plasma membrane and Ca2+ release from intracellular
Ca2+ stores (mainly sarcoplasmic/endoplasmic reticulum; see
Refs. 4, 7, 34, 44). Ca2+ influx depends on the
Ca2+ driving force or the electrochemical gradient across
the plasma membrane (30, 44). Although the chemical gradient, the ratio of extracellular Ca2+ concentration
([Ca2+]o) to
[Ca2+]cyt, and the Ca2+
equilibrium potential [ECa, which is equal to
12.5 ln([Ca2+]o/[Ca2+]cyt) = 117 131 mV at 25°C] are constant, the
Ca2+ driving force is mainly determined by the electrical
gradient, the difference between membrane potential
(Em) and ECa
(Em - ECa). In other words,
Em is a major determinant of the driving force for
Ca2+ influx.
By controlling the Ca2+ driving force, Em is an important regulator of [Ca2+]cyt in nonexcitable cells, including epithelial cells and lymphocytes (14-16, 29-31). Membrane depolarization decreases the Ca2+ driving force and inhibits Ca2+ influx. In contrast, membrane hyperpolarization increases the Ca2+ driving force and enhances Ca2+ influx. Therefore, in the cells that do not express L-type voltage-dependent Ca2+ channels (VDCC), Ca2+ influx is decreased by membrane depolarization but is increased by membrane hyperpolarization (16, 18, 27, 32, 38). Nonetheless, in excitable cells such as neurons, cardiomyocytes, and muscle cells, VDCC that are opened by membrane depolarization are the major pathway for Ca2+ influx (30, 44). In contrast to the voltage-independent pathway for Ca2+ influx in nonexcitable cells, membrane depolarization opens VDCC and thus increases [Ca2+]cyt in excitable cells (30, 44).
Em is primarily determined by K+ permeability (PK) and the K+ gradient across the plasma membrane (31). Because the K+ gradient is maintained by Na+-K+-ATPase, PK is directly related to the activity and number of membrane K+ channels. Voltage-gated K+ channels (Kv) have been shown to be a major determinant of resting Em in many types of cells (14-16, 31). When K+ channel closes or the number of total K+ channels decreases, Em becomes less negative (i.e., depolarization). When K+ channel opens or the number of total K+ channels rises, Em becomes more negative (i.e., hyperpolarization; see Ref. 31). Therefore, inhibition of K+ channel gene expression would decrease the number of K+ channels and attenuate K+ channel activity. The subsequent membrane depolarization decreases the Ca2+ driving force and thus inhibits Ca2+ influx. Because Ca2+ entry is a major source for [Ca2+]cyt, inhibition of Ca2+ influx would reduce [Ca2+]cyt in cells lacking VDCC (16, 28).
In this study, we proposed to test the hypothesis that K+
channel gene expression, Em, and
[Ca2+]cyt are involved in the
mechanism by which polyamines are required for the stimulation of cell
migration after wounding. First, we determined whether depletion of
cellular polyamines by -difluoromethylornithine (DFMO), a specific
inhibitor of ornithine decarboxylase (ODC, the rate-limiting enzyme for
polyamine biosynthesis), altered K+ channel gene
expression, Em, and
[Ca2+]cyt in intestinal epithelial
cells. Second, we examined the roles of
[Ca2+]cyt and
Em in polyamine-dependent cell migration after wounding.
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MATERIALS AND METHODS |
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Cell culture and general experimental protocol. The intestinal epithelial cell (ICE-6) line was purchased from the American Type Culture Collection (ATCC) at passage 13. The cell line was derived from normal rat intestine and was developed and characterized by Quaroni et al. (39). The nontumorigenic ICE-6 cells originated from intestinal crypt cells, as judged by morphological and immunological criteria, and retained the undifferentiated character of epithelial stem cells.
Stock cells were maintained in T-150 flasks in DMEM supplemented with 5% heat-inactivated FBS (Sigma), 10 µg/ml insulin, and 50 µg/ml gentamicin sulfate. Flasks were incubated at 37°C in a humidified atmosphere of 10% CO2 in air. Stock cells were subcultured one time per week at 1:20, and medium was changed three times per week. Tests for mycoplasma were routinely negative. 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 (49). In brief, IEC-6 cells were plated at 6.25 × 104 cells/cm2 in DMEM supplemented with 5% dialyzed FBS, 10 µg/ml insulin, and 50 µg/ml gentamicin sulfate. The cells were incubated at 37°C for 24 h before experimental treatments. In the first series of studies, we examined the effects of polyamine depletion on K+ channel gene expression, Em, and [Ca2+]cyt in IEC-6 cells. The cells were grown in control cultures or cultures containing either DFMO (5 mM) or DFMO plus 5 µM spermidine for 4 days. The dishes were placed on ice, the monolayers were washed three times with ice-cold Dulbeccos's PBS (D-PBS), and then different solutions were added according to the assays to be conducted. In the second series of studies, we investigated the relationship between [Ca2+]cyt and cell migration in normal (without DFMO) and polyamine-depleted IEC-6 cells (with DFMO) treated with or without exogenous polyamine (5 µM spermidine). The Ca2+-free medium or 4-aminopyridine (4-AP, an inhibitor of K+ channels) at different concentrations in the standard DMEM was added immediately after wounding to control cultures and cultures in which ODC was inhibited with DFMO and supplemented with 5 µM exogenous spermidine. Cell migration was assayed 2-6 h after treatment.
RT-PCR. Total RNA was prepared by the acid guanidinium
thiocyanate-phenol-chloroform extraction method (9). The 3 µg of the total RNA were reversely transcribed using the
First-Strand cDNA Synthesis Kit (Pharmacia Biotech) and random hexamers
[pd(N)6 primer]. The reaction mixture was
incubated for 1 h at 37°C and then was heated at 90°C for
5 min to inactivate the RT. Specific primers for Kv channel
(the pore-forming)- and
(the cytoplasmic regulatory)-subunits,
L-type VDCC
1- and
1-subunits, and
transient receptor potential (TRP) channels were designed from the cDNA sequences of the coding regions corresponding to the channel genes (Table 1). PCR was performed by a GeneAmp
PCR System (Perkin-Elmer) using Taq polymerase. The 3 µl of
the first-strand cDNA reaction mixture were used in the PCR reaction.
The cDNA samples were amplified in the thermal cycler under the
following conditions: the mixture was annealed at 55°C (1 min),
extended at 72°C (2 min), and denatured at 94°C (1 min) for 25 cycles. This was followed by a final extension at 72°C (10 min) to
ensure complete product extension. The PCR products were
electrophoresed through a 1% agarose gel, and amplified cDNA bands
were visualized by ethidium bromide staining.
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To quantify the PCR products (the amounts of mRNA) of the Kv and
Ca2+-permeable channels, an invariant mRNA of -actin was
used as an internal control. Immediately after each of the experiments, the optical density (OD) values for each band on the gel were measured
by a Gel Documentation System (UVP, Upland, CA). The OD values in the
K+ or Ca2+ channel signals were normalized to
the OD values in the
-actin signals. The normalized values in the
controls were expressed as 1 arbitrary unit for quantitative comparison
(45).
Western blot analysis of Kv channel protein. Cell samples,
dissolved in SDS sample buffer, were sonicated for 20 s and centrifuged at 2,000 rpm for 15 min. The supernatant was boiled for 10 min and then
was subjected to electrophoresis on 7.5% acrylamide gels according to
Laemmli (43). Each lane was loaded with 20 µg protein equivalents.
After the transfer of protein to nitrocellulose filters, the filters
were incubated for 1 h in 5% nonfat dry milk in 10× 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 1 h in 1% BSA/PBS-T buffer containing 1 µg/ml affinity-purified
polyclonal antibody against Kv1.1 protein (Upstate Biotechnology, Lake
Placid, NY) or -actin (Boehringer Mannheim). The filters were
subsequently washed with 1× PBS-T and were incubated for 1 h with
goat anti-rabbit IgG antibody conjugated to peroxidase. After extensive
washing with 1× PBS-T, the immunocomplexes on the filters were
reacted for 1 min with Chemiluminescene Reagent (NEL-100; NEN).
Finally, the filters were placed in a plastic sheet protector and were
exposed to autoradiography film for 30 or 60 s.
Measurement of Em. Em in single IEC-6
cells grown on 25-mm coverslips was measured using an intracellular
electrode (30-50 M) filled with 3 M KCl. Data were acquired by
an electrometer (Electro 705; WPI) coupled to a computer and a chart
recorder and were analyzed using DATAQ acquisition software.
Measurement of [Ca2+]cyt. Details of the digital imaging methods employed for measuring [Ca2+]cyt have been published (52). Briefly, IEC-6 cells were plated on 25-mm coverslips and were incubated in culture medium containing 3.3 µM of 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 32-34°C to wash away extracellular dye and 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 using a Nikon Diaphot microscope equipped for epifluorescence. Fluorescent images were obtained using 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 using a Matrix LC imaging board operating in an IBM-compatible computer (66 MHz, 486). 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 every 60 s when [Ca2+]cyt was relatively constant. [Ca2+]cyt was calculated from fura 2 fluorescent emission excited at 380 and 360 nm using the ratio method (34). In most experiments, multiple cells (usually 10-15 cells) were imaged in 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 either recording Em or 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+. The 4-AP (Sigma) was directly dissolved in PSS or culture media on the day of use. The pH in the solution containing 4-AP was adjusted to 7.4 with saturated HCl before each experiment.
The DMEM that was specially customized to exclude NaCl, KCl, and CaCl2 (other ingredients were not changed) was used in the cell migration experiments. Thus we could change the ionic composition in the salt-free DMEM by adding different concentrations of NaCl, KCl, and CaCl2 based on the formula provided by the company (Mediatech, Cellgro, VA). To make Ca2+-free medium supplemented with 5% dialyzed FBS (Sigma), only NaCl and KCl were added to the DMEM, and CaCl2 was replaced by equimolar MgCl2 to keep a constant and consistent osmolarity. EGTA (0.1 mM) was added to chelate residual Ca2+ in the Ca2+-free DMEM.
Measurement of cell migration. The migration assays were carried out as described previously (26, 49), modified for quantitation by computer as described below. IEC-6 cells were plated at 6.25 × 104 cells/cm2 in DMEM/dialyzed FBS with or without DFMO and polyamines on 35-mm dishes thinly coated with Matrigel according to the manufacture's directions and were incubated as described for stock cultures. Four days after plating, approximately one-third of the cell layer was removed with a razor blade, and cell migration was allowed to occur over the denuded area for 2-6 h. Micrographic images of IEC-6 cells along the scratch line were captured with a Macintosh IIci computer equipped with a Data Translation Quick Capture board and COHU 8215 charge-coupled device camera attached to a Nikon TMS inverted microscope with a C mount. A ×10 objective and a ×16 television relay lens were used. Culture dishes were placed on the microscope stage so that the scratch line was horizontal across the lower one-half of the image. Data collection and image analysis were accomplished with National Institutes of Health Image 1.41. Sixteen consecutive images were collected and averaged to reduce background noise. The time period for the 16 consecutive images was ~10-20 s and was automatically regulated by a computer. From the calibrated rule, the number of image pixels per millimeter was determined. Results were reported as the number of cells migrating per millimeter of scratch (cells/mm).
HPLC analysis of cellular polyamines. The cellular polyamine
content was determined as previously described (47). Briefly, after
washing the monolayers three times with ice-cold D-PBS, 0.5 M
perchloric acid was added, and the monolayers were frozen at
80°C until ready for extraction, dansylation, and HPLC
analysis. The standard curve encompassed 0.31-10 µM. Values that
fell >25% below the curve were considered not detectable. Protein
was determined by the Bradford (6) method. The results are expressed as
nanomolar polyamines per milligram of protein (nM/mg).
Statistical analysis. All data are expressed as means ± SE. Autoradiographic and immunofluorescence labeling results were repeated three times. The significance of the difference between means was determined by ANOVA. The level of significance was determined using the Dunnett's multiple range test (17). Differences were considered to be significant at P < 0.05.
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RESULTS |
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Effect of DFMO on cellular polyamines and Kv channel expression. Incubation of IEC-6 cells in media containing 5 mM DFMO, which totally inhibits ODC activity (46, 47), prevented the accumulation of cellular polyamines. The level of putrescine was undetectable on day 2 after treatment. Spermidine was decreased by ~50% on day 2 but was undetectable on day 4. Spermine was less sensitive to the inhibition of ODC and was decreased by ~30% on day 2 and ~65% on day 4 in the DFMO-treated cells.
Depletion of intracellular polyamines by DFMO selectively and
significantly inhibited mRNA expression of the Kv channel -subunit (the pore-forming subunit) Kv1.1 but negligibly affected the
electrically silent Kv channel
-subunit Kv9.3 and the Kv channel
-subunit (Kv
1.1; Fig. 1). The mRNA
level of Kv1.1 channels in cells treated with DFMO for 4 days was only
~5% of the normal value (without DFMO). The decreased mRNA level of
Kv1.1 channels was completely prevented by addition of exogenous
spermidine (5 µM) in the presence of DFMO (Fig.
2A). The decreased mRNA level of
Kv1.1 channels was paralleled by a decrease in the protein level of
Kv1.1 channels (Fig. 2B). The level of Kv1.1 channel protein
was decreased by ~45% on day 2 and by ~90% on day
4 in cells exposed to DFMO. The Kv1.1 channel protein content
returned to normal levels when DFMO was given together with spermidine.
Putrescine at a dose of 10 µM had an effect equal to spermidine on
the expression of the Kv1.1 channel gene when it was added to cultures
that contained DFMO (data not shown).
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Effect of polyamine depletion on Em. Because
decreased expression of the gene encoding Kv1.1 channels in
polyamine-deficient cells would decrease the number of functional Kv
channels and alter membrane polarization, the next set of
experiments was designed to investigate the effect of polyamine
depletion on resting Em in the presence or absence
of exogenous spermidine. Consistent with the inhibitory effect on the
Kv1.1 channel expression, depletion of cellular polyamines by DFMO
significantly depolarized Em in IEC-6 cells,
whereas addition of spermidine to the cultures in the presence of DFMO
reversed the depolarizing effect (Fig.
3).
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Membrane depolarization would open VDCC and promote Ca2+ influx in excitable cells, including pulmonary artery smooth muscle cells and neurons in which VDCC are major pathways for Ca2+ influx. However, membrane depolarization in nonexcitable cells such as intestinal epithelial cells that do not express VDCC would decrease the Ca2+ driving force and attenuate Ca2+ influx through store-operated Ca2+ channels (SOC) that may be formed by TRP-encoded proteins (34, 53-55).
In IEC-6 cells, the pore-forming (1-) and regulatory
(
1-) subunits of L-type VDCC were not detectable by
RT-PCR analysis, whereas VDCC
1- and
1-subunits were expressed in rat pulmonary artery smooth
muscle cells (Fig. 4A,
top). TRP-1, which encodes a Ca2+-permeable channel
involved in the capacitative Ca2+ entry in mammalian cells
(53-55), was detected in the epithelial cells (Fig. 4A,
top). Although TRP-4 and TRP-6 were detectable in rat pulmonary
artery smooth muscle cells, they were not present in the epithelial
cells (Fig. 4A, bottom). These results indicate that
IEC-cells do not express VDCC but express TRP-1 channels that are
responsible for the capacitative Ca2+ entry (55, 53).
Depletion of intracellular polyamine by DFMO had no effect on mRNA
expression of TRP-1 channels (Fig. 4, B and C).
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Effect of polyamine depletion by DFMO on
[Ca2+]cyt. As described above,
membrane depolarization in the cells that do not express VDCC would
reduce the Ca2+ driving force, inhibit Ca2+
influx, and decrease [Ca2+]cyt.
Indeed, in IEC-6 cells, polyamine depletion by DFMO, which caused
inhibition of Kv1.1 channel expression and membrane depolarization, significantly decreased the resting
[Ca2+]cyt (from 94.3 ± 6.3 to
28.5 ± 4.8 nM, n = 33, P < 0.001) and inhibited the capacitative Ca2+ entry (from 830.3 ± 53.5 to 553.2 ± 37.5 nM, n = 33, P < 0.001; Fig. 5). Cyclopiazonic acid (CPA) induced
an initial transient increase in
[Ca2+]cyt in the absence of
extracellular Ca2+ that was due apparently to
Ca2+ mobilization from intracellular Ca2+
stores (mainly endoplasmic reticulum; see Ref. 34). Addition of
extracellular Ca2+ to the cell superfusate when the
CPA-induced transient rise in [Ca2+]cyt returned to the basal
level (i.e., when the CPA-sensitive intracellular Ca2+
stores were depleted) resulted in a sustained increase in
[Ca2+]cyt because of the
capacitative Ca2+ entry. Membrane depolarization induced by
increasing extracellular K+ concentration to 40 mM
significantly reduced the capacitative Ca2+ entry (Fig.
5A, left) but had no effect on resting
[Ca2+]cyt (because the cells do not
express VDCC). In this experiment, the capacitative Ca2+
entry was estimated by measuring the amplitude of the CPA-induced Ca2+ influx (Fig. 5C, right).
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Effects of removal of extracellular Ca2+ and membrane
depolarization on polyamine-dependent cell migration. To determine
the role of [Ca2+]cyt in
polyamine-dependent cell migration, two types of experiments were
carried out by using the Ca2+-free medium and the
K+ channel blocker 4-AP (1-5 mM). In the first study,
we examined whether decreasing
[Ca2+]cyt by removal of
extracellular Ca2+ or by membrane depolarization with 4-AP
altered the rate of cell migration after wounding in control cells
(without DFMO). The Ca2+-free medium or 4-AP was added
immediately after wounding and was present during the migration period
(2-6 h). As shown in Fig. 6, exposure
to the Ca2+-free medium or to different doses of 4-AP
significantly decreased the rate of cell migration. Removal of
extracellular Ca2+ and application of 4-AP did not alter
cell viability (data not shown). In the second study, we examined the
effects of removal of extracellular Ca2+ and administration
of 4-AP on the restoration of cell migration by exogenous spermidine in
polyamine-deficient cells. As shown in Fig.
7A, the migration was significantly
decreased in DFMO-treated cells (b vs. c). When
spermidine was added concomitantly with DFMO, it was able to maintain
cell migration at near-normal levels (Fig. 7A, b vs.
d). Treatment with either the Ca2+-free medium
(Fig. 7A, b vs. e) or 4-AP (b vs.
f) during the period of cell migration prevented restoration of
cell migration by spermidine in DFMO-treated cells (Fig. 7, B
and C). There was no apparent loss of cell viability in cells
either treated with DFMO alone, DFMO plus spermidine, or those treated
with DFMO plus the Ca2+-free medium or 4-AP. These results
clearly show that a rise in [Ca2+]cyt due to Ca2+
influx through Ca2+-permeable channels in the plasma
membrane is required for polyamine-dependent migration in intestinal
epithelial cells. Membrane depolarization induced by inhibition of Kv
channels could decrease [Ca2+]cyt
and inhibit polyamine-induced migration in IEC-6 cells.
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DISCUSSION |
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The present study confirms our previous findings (26, 46, 47, 49) that depletion of cellular polyamines by DFMO inhibits cell migration after wounding, which can be completely restored to normal by concomitant treatment with exogenous polyamines. The most significant of the new findings reported in this study is that K+ channel expression is involved in the process by which polyamines stimulate cell migration during intestinal epithelial restitution. Polyamine depletion by DFMO resulted in a significant decrease in Kv1.1 channel gene expression (Figs. 1 and 2) and membrane depolarization (Fig. 3) in IEC-6 cells. Because IEC-6 cells do not express L-type VDCC (Fig. 4), the depolarized Em in polyamine-deficient cells decreased [Ca2+]cyt by reducing the driving force for Ca2+ influx through the capacitative Ca2+ entry pathway (Fig. 5). Furthermore, removal of extracellular Ca2+ or membrane depolarization by blockade of Kv channels with 4-AP inhibited normal cell migration (Fig. 6) and prevented the restoration of cell migration by exogenous spermidine in polyamine-deficient IEC-6 cells (Fig. 7). These results indicate that polyamines are required for the stimulation of cell migration after wounding in association with their ability to modulate K+ channel gene expression that subsequently alters Em and [Ca2+]cyt in intestinal epithelial cells.
Intracellular polyamines and extracellular Ca2+ are required in intestinal epithelial cell migration. Polyamines are ubiquitous organic cations of low molecular weight found in all eukaryotic cells and are intimately involved in a variety of cellular functions, including cell migration, proliferation, and differentiation (24, 42, 47). It has been demonstrated that polyamines are absolutely required for normal repair of gastric and duodenal mucosal stress erosions, and polyamines accelerate repair at least partly through the mechanism involving mucosal epithelial migration (25, 46). However, few specific molecular functions of polyamines in epithelial migration have been described (42, 49). Some of the attributes ascribed to polyamines include the maintenance of the distribution and the formation of cytoskeletal proteins such as actin (26), myosin II (48), and microtubules (3) and regulation of migration-related gene expression (22, 49). Although the observations derived from gastric epithelia in vivo (40, 41) and in vitro (13, 41) have shown the requirement of Ca2+ for mucosal epithelial restitution, there are few studies concerning the relationship between the stimulation of cell migration by polyamines and the K+ channel expression and [Ca2+]cyt regulation.
An increase in [Ca2+]cyt is a trigger for cell migration in a variety of cell types, whereas a decrease in [Ca2+]cyt significantly inhibits cell movement (4, 10, 37). The Ca2+- and calmodulin-dependent protein kinase C II plays an important intermediate role in the Ca2+-dependent cell migration (4, 37). Removal of extracellular Ca2+ significantly inhibited cell migration (Figs. 6 and 7), suggesting that Ca2+ influx is required in normal migration in intestinal epithelial cells.
Regulation of [Ca2+]cyt and TRP channels. The transmembrane influx of Ca2+ is mainly determined by the Ca2+ driving force (21). Under physiological conditions, [Ca2+]cyt is 50-150 nM (5), whereas [Ca2+]o is 1.6-1.8 mM. Thus the major determinant of Ca2+ influx is Ca2+ permeability and the Ca2+ electrical potential across the membrane. When cells are hyperpolarized, the driving force for Ca2+ influx rises; when cells are depolarized, the driving force decreases. Because the intestinal epithelial cells do not express L-type VDCC (Fig. 4), Ca2+ influx is decreased by membrane depolarization but increased by membrane hyperpolarization.
In endothelial and epithelial cells, passive Ca2+ leakage, receptor-operated Ca2+ channels (ROC), nonselective cation channels (NSCC), and SOC all contribute to Ca2+ influx (18, 28, 32, 38, 44). Agonist-mediated depletion of intracellular Ca2+ stores induces the influx of extracellular Ca2+, namely capacitative Ca2+ entry (38). The SOC that are involved in the capacitative Ca2+ entry are believed to relate to TRP-encoded channels (53-55). Nonexcitable cells, such as intestinal epithelial cells, lack VDCC (18, 32) but have developed the Ca2+ entry mechanism that is coupled with the depletion of intracellular Ca2+ stores to activate SOC (i.e., TRP channels). Thus the capacitative Ca2+ entry via TRP channels may be a major source of intracellular Ca2+ in endothelial and epithelial cells.
Although IEC-6 cells do not express VDCC, they express the TRP-1 channel (Fig. 4), which is a Ca2+-permeable channel responsible for the capacitative Ca2+ entry (53, 54). The capacitative Ca2+ influx induced by depleting intracellular Ca2+ stores with CPA is huge in IEC-6 cells; the amplitude of the CPA-induced Ca2+ influx (830.3 ± 53.5 nM) was ~3.4 times greater than the amplitude of the CPA-induced Ca2+ release (196.4 ± 10.8 nM; Fig. 5). These data suggest that the capacitative Ca2+ entry through TRP-1 channels may serve as an important source for the agonist-mediated rise in [Ca2+]cyt in intestinal epithelial cells. Polyamine depletion by DFMO did not affect the mRNA expression of TRP-1 channels (Fig. 4, B and C) but significantly attenuated the CPA-induced Ca2+ influx or the capacitative Ca2+ entry (Fig. 5, A and C). These data suggest that polyamines augment the capacitative Ca2+ entry by inducing membrane hyperpolarization, rather than by directly affecting the TRP-1 channel expression.
Determination of Em by Kv channel activity and
expression. Em is a function of the Na+,
K+, and Cl concentration gradients
across the plasma membrane and the relative ion permeability. In
resting cells, Em is controlled primarily by
PK and K+ concentration gradients
because PK > Na+ permeability
(PNa) > PCl
(PK-PNa-PCl = 1.0:0.04:0.45). PK (and thus
Em) is directly related to the whole cell
K+ current, which is dependent on the number of
membrane K+ channels and single-channel (unitary) current
(21, 23). When K+ channel opens or K+ channel
expression rises, Em becomes more negative (i.e.,
hyperpolarization) because of increased PK (14, 31,
50).
Kv channels play an important role in regulating resting
Em in many types of cells (14, 31, 50, 52).
Inhibition of Kv channels by 4-AP causes membrane depolarization,
whereas activation of Kv channels (by nitric oxide) causes membrane
hyperpolarization (1, 12, 51). At molecular levels, Kv channels are
composed of the pore-forming -subunits and the regulatory
-subunits (20). As shown in Fig. 1, at least two Kv channel
-subunits (Kv1.1 and Kv9.3) and a Kv channel
-subunit (Kv
1.1)
are expressed in the IEC-6 cells (other types of K+
channels were not examined). The data from this study clearly demonstrated that depletion of cellular polyamines by DFMO selectively decreased Kv1.1 channel gene expression (Figs. 1 and 2), which was
associated with a significant membrane depolarization (Fig. 3). The
reduced Ca2+ driving force by membrane depolarization
significantly decreased resting
[Ca2+]cyt and inhibited the
CPA-induced [Ca2+]cyt
rise in DFMO-treated cells (Fig. 5), probably through the decrease in
Ca2+ influx via the passive leakage pathway and the
capacitative Ca2+ entry.
It has been demonstrated that Kv channels are also permeable to Ca2+ (8, 19). At resting Em, the Ca2+ influx through K+ channels would increase [Ca2+]cyt by 40 nM/s (8). These results suggest that the enhanced Ca2+ influx through Kv channels may be another mechanism by which polyamines increase [Ca2+]cyt in IEC-6 cells.
Possible mechanisms by which polyamines increase Kv channel
expression. The exact roles of polyamines at the molecular level in
the regulation of Kv channel gene expression remain to be elucidated. Although decreased Kv1.1 channel protein was paralleled by a
significant decrease in its mRNA in polyamine-deficient cells (Fig. 2),
it is not clear at present whether the decreased mRNA level of the Kv1.1 channel gene results from a decrease in the gene transcription or
an alteration of the mRNA stabilization. There is no question that
cellular polyamines play different roles in the expression of various
genes, and their effects are cell type dependent. Polyamines have been
shown to stimulate transcription of the protooncogenes c-myc
and c-jun (36) but to accelerate degradation of transforming growth factor- mRNA in intestinal epithelial cells (35). Further studies are clearly needed to investigate the transcriptional and
posttranscriptional regulation of the Kv channel gene when cellular
polyamines are increased or decreased in IEC-6 cells.
Based on the current findings, we propose a model delineating the
regulation of cell migration on the basis of the polyamine-mediated Kv
channel expression (Fig. 8). In this model,
cellular polyamines are essential for inducing gene expression of the
Kv channels in intestinal epithelial cells. Enhanced Kv channel
expression after increased polyamines would increase the number of
functional Kv channels, increase whole cell Kv currents, and cause
membrane hyperpolarization. Because the intestinal epithelial cells
lack L-type VDCC, membrane hyperpolarization induced by polyamines augments the driving force for Ca2+ influx and raises
[Ca2+]cyt, thereby resulting in
cell migration into the wounding region (Fig. 8). These results are
consistent with the data from other investigators (2) who demonstrate
that administration of putrescine increases
[Ca2+]cyt in rat ascites hepatoma
cells and that the effect of putrescine is not affected by antagonists
of L-type voltage-gated Ca2+ channels.
|
In summary, the data from this study indicate that expression of K+ channels is involved in the stimulation of cell migration by polyamines in IEC-6 cells (Fig. 8). Depletion of cellular polyamines by DFMO resulted in a decrease of Kv1.1 channel expression, membrane depolarization, and a decrease in [Ca2+]cyt, thereby inhibiting IEC-6 cell migration. Exogenous spermidine not only reversed the inhibitory effects of DFMO on the Kv channel gene expression, Em, and [Ca2+]cyt but also restored cell migration to control levels. Decreasing [Ca2+]cyt by removal of extracellular Ca2+ or by membrane depolarization inhibited cell migration. These findings suggest that polyamine-dependent intestinal epithelial cell migration after wounding may be due partially to the enhanced Kv channel (Kv1.1) expression and the resultant membrane hyperpolarization and increased [Ca2+]cyt.
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ACKNOWLEDGEMENTS |
---|
We thank J. E. Seiden and M. J. Viar for technical assistance and L. Tague for assistance with the measurement of polyamines.
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FOOTNOTES |
---|
This work was supported by National Institutes of Health Grants HL-54043 and HL-64945 to J. X.-J. Yuan and DK-45314 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.
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: J.-Y. Wang, Dept. of Surgery, Baltimore VA Medical Center, 10 N. Greene St., Baltimore, MD 21201 (E-mail: jwang{at}smail.umaryland.edu).
Received 30 March 1999; accepted in final form 20 September 1999.
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