2 Laboratory of Cellular Oncology, Signal Transduction Section, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 20892; and 1 Division of Gastroenterology, Research Institute, The Hospital for Sick Children, Departments of Paediatrics and Biochemistry, University of Toronto, Toronto, Ontario, Canada M5G 1X8
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ABSTRACT |
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Treatment of HT-29 cells with phorbol
12-myristate 13-acetate (PMA), an activator of protein kinase C (PKC),
induces MUC2 expression. To investigate the role of PKC in
regulating mucin genes in intestinal cells, we examined the regulation
of MUC1, MUC2, MUC5AC, MUC5B, and MUC6 expression in two human
mucin-producing colonic cell lines, T84 and HT29/A1. T84 and HT29/A1
cells (at 80-90% confluency) were exposed to 100 nM PMA for 0, 3, and 6 h. Twofold or greater increases in mRNA levels for MUC2 and
MUC5AC were observed in both cell lines during this time period,
whereas the levels of MUC1, MUC5B, and MUC6 mRNAs were only marginally affected. These results indicated that PKC differentially
regulates mucin gene expression and that it may be responsible for
altered mucin expression. Our previous results suggested that the
Ca2+-independent PKC- isoform
appeared to mediate PMA-regulated mucin exocytosis in these cell lines.
To determine if PKC-
was also involved in MUC2/MUC5AC gene
induction, HT29/A1 cells were stably transfected with either a
wild-type PKC-
or a dominant-negative ATP-binding mutant of PKC-
(PKC-
K437R). Overexpression of the dominant-negative PKC-
K437R
blocked induction of both mucin genes, whereas PMA-induced mucin gene
expression was not prevented by overexpression of wild-type PKC-
.
PMA-dependent MUC2 mucin secretion was also blocked in cells
overexpressing the dominant-negative PKC-
K437R. On the basis of
these observations, PKC-
appears to mediate the expression of two
major gastrointestinal mucins in response to PMA as well as
PMA-regulated mucin exocytosis.
protein kinase C; signal transduction; phorbol 12-myristate 13-acetate
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INTRODUCTION |
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MUCUS PLAYS AN IMPORTANT role in protecting nonsquamous epithelial surfaces against mechanical damage and in stabilizing the luminal microenvironment of surface cells. Highly glycosylated mucin proteins are the major components of mucus, responsible for its viscoelastic properties. Nine different mucin genes have now been identified in various human tissues. MUC1, MUC2, MUC3, MUC5AC, MUC5B, and MUC6 are expressed in intestinal cells (3, 19, 28, 35, 36). Three mucins, MUC1 (17), the homologue of MUC3 in the rat (14) [although not apparently human MUC3 (10)], and MUC4 (23), have been reported to have classic carboxy-terminal membrane-spanning domains and therefore may be anchored in whole or in part to the surface plasma membrane. The remainder are secreted mucins that are stored in secretory granules within cells and secreted in response to a wide variety of stimuli (8) to form the surface mucus gel. Although individual mucins display tissue specificity, there is growing evidence that expression is subject to regulatory control and that the relative expression of particular mucins may represent a specific response to inflammation or environmental factors. For example, in the adult intestine MUC5AC is normally expressed in mucus-secreting foveolar cells of gastric glands but can be expressed in colonic neoplastic cells, depending on culture conditions (19), and may appear in goblet cells in inflamed duodenum and colon (29).
Regulation of the expression of MUC2 has been explored in HT-29 cells
and shown to be subject to both protein kinase C (PKC)- and protein
kinase A-dependent stimuli (37). The present study was undertaken to
assess the effect of phorbol 12-myristate 13-acetate (PMA) on the
expression of the mucin genes MUC1, MUC2, MUC5AC, MUC5B, and MUC6 and
to determine whether the PKC isoform PKC- was involved, because we
had previously identified PKC-
as the likely isoform mediating
PMA-stimulated mucin exocytosis (12). To examine the role of PKC-
in
PMA-induced mucin gene expression, we have utilized stably transfected
HT29/A1 cells constructed to overexpress either wild-type PKC-
or a
dominant-negative PKC-
mutant.
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MATERIALS AND METHODS |
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Materials. All chemicals were obtained from Sigma (St. Louis, MO) unless otherwise noted. Nytran nylon membranes were obtained from Schleicher & Schuell (Keene, NH).
Cell culture. T84 human colonic carcinoma cell lines were obtained from the American Type Culture Collection (Rockville, MD). T84 cells were cultured in a 1:1 mixture of DMEM-F12 with 5% FBS, penicillin (100 U/ml), and streptomycin (100 µg/ml). HT29/A1, a mucin-producing subclone of the HT29 human colonic tumor cell line, was obtained from Dr. K.-M. Kreusel (Institut fur Klinische Physiologie, Universitats-klinikum, Frei Universitat Berlin, Berlin, Germany). It has been described previously as HT29/B6 (15). HT29/A1 cells were grown in RPMI 1640 medium containing 10% (vol/vol) FBS, penicillin (100 U/ml), and streptomycin (100 µg/ml). All cell culture regents were from GIBCO BRL (Burlington, ON). Cultures were maintained at 37°C in a humidified atmosphere of 5% CO2 and 95% air.
Extraction of total RNA.
Total RNA was extracted by the method of Chomczynski and Sacchi (5).
Briefly, cultures grown in 10-cm-diameter dishes were washed three
times with ice-cold PBS and lysed directly in the dishes using 2 ml of
solution A (4 M guanidium isothionate, 25 mM sodium citrate,
pH 7.0, 0.5% sarcosyl, and 10 mM -mercaptoethanol). The lysed
monolayer was scraped with a cell scraper to ensure that all material
was released from the dishes. The lysate was transferred to a 15-ml
polypropylene tube, and 0.2 ml of 2 M sodium acetate (pH 4.0), 2 ml
phenol, and 0.4 ml of a chloroform-isoamyl alcohol mixture (49:1) were
added sequentially. The lysate was mixed by inversion after each
addition. After brief vortexing, the mixture was kept on ice for 15 min
and then centrifuged at 12,000 g for
30 min at 4°C. The upper aqueous phase was transferred to a new
tube, and an equal volume of ice-cold isopropanol was added. The
samples were mixed and stored at
20°C overnight. RNA was collected
by centrifugation at 12,000 g for 20 min at 4°C. The supernatant was discarded, and the RNA pellet was
recovered by centrifugation at 12,000 g for 15 min at 4°C, washed twice with 70% ethanol, and dried under vacuum. The dried RNA pellet was
then dissolved in diethylpyrocarbonate-treated water and was quantitated by spectrophotometry at a wavelength of 260 nm.
RNA analysis.
For Northern blot hybridization, RNA samples (15 µg) were denatured
at 65°C for 15 min, applied to a 1.0% agarose gel containing 2.2 M
formaldehyde, and then electrophoresed at 30 V for 12 h. The quality
and the relative abundance of RNA per lane were judged by comparing the
ethidium bromide staining of the ribosomal bands. RNA was transferred
to a Nytran nylon membrane and immobilized by UV cross-linker
(Stratagene). Membranes were incubated at 43°C overnight in
prehybridization buffer [5× sodium chloride-sodium phosphate-EDTA (SSPE), 0.2% SDS, 5× Denhardt's solution, 100 µg/ml sonicated salmon sperm DNA, and 50% formamide]. After
prehybridization, the membranes were hybridized for 36 h at 43°C in
prehybridization buffer containing 10% dextran sulfate and
32P-labeled cDNA (1,000,000 cpm/ml). After hybridization, the membranes were washed three times
with 1× SSPE and 0.1% SDS for 15 min at room temperature and
then washed two times with 0.1× SSPE and 0.1% SDS for 15 min at
55°C. Membranes were exposed to Kodak X-Omat film with intensifying
screens for 6-12 h at 80°C. Slot-blot analysis was
performed by applying 8 µg of total RNA to Nytran nylon membranes
using a slot-blot apparatus (Schleicher & Schuell). Membranes were
incubated at 43°C overnight in prehybridization buffer (5×
SSPE, 0.2% SDS, 5× Denhardt's solution, 100 µg/ml sonicated salmon sperm DNA, and 50% formamide). After prehybridization, the
membranes were hybridized for 36 h at 43°C in prehybridization buffer containing 10% dextran sulfate and
32P-labeled cDNA (1,000,000 cpm/ml). After hybridization, the membranes were washed three times
with 1× SSPE and 0.1% SDS for 15 min at room temperature and
then two times with 0.1× SSPE and 0.1% SDS for 15 min at
55°C. Membranes were exposed to Kodak Biomax film with intensifying
screens for 6-12 h at
80°C. Images were scanned using
an UVP image system, and the densitometric ratio of mucin mRNA to
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA was quantitated
by NIH Image software.
cDNA probes. The 411-bp cDNA from the carboxy-terminal region of MUC2 cloned in our laboratory has been described elsewhere (40). The MUC1 cDNA (656 bp) probe was generated by RT-PCR using the specific primers (sense, 690 5'-AGGCTCAGCTTCTAC-TCTGG-3' 710 and antisense, 1356 5'-GACAGACAGCCAAGGCAATG-3' 1326), described by Chambers and Harris (4). The 271-bp MUC5AC cDNA probe was generated by RT-PCR using the specific primers described by Voynow and Rose (38). A 104-bp cDNA probe was generated by RT-PCR using the sense primer 1651 5'-AGCTCCAAAGCCACTTCCTC-3' 1670 and antisense primer 1754 5'-GGGATGGGTGTAAAGCTGGTAG-3' 1733, based on the updated GenBank sequence of the JER57 clone initially isolated by Dufosse et al. (7) and subsequently identified as part of MUC5B (6). The MUC6 cDNA probe was generated by RT-PCR with primers selected according to the published sequence (31). A 590-bp fragment was produced by specific sense primer 610 5'-GCTTCACCAACAACAAGTTTAAG-3' 630 and antisense primer 1199 5'-CATCATTCAGACAAGCAAAGC-3' 1179. A second round of RT-PCR, using the 590-bp fragment as template, was performed with internal sense primer 797 5'-CCAAGTCTACCAGTAGAGAC-3' 818 and antisense primer 1098 5'-CTTCTGCTTCGATCCACTCA-3' 1079 to generate a 302-bp cDNA probe. The 983-bp GAPDH cDNA probe was generated by RT-PCR using cDNA from human gastric tissue as template. The primer set (sense, 44 5'-TGAAGGTCGGAGTCAACGGATTTGGT-3' 69 and antisense, 1026 5'-CATGTGGGCCATGAGGTCCACCAC-3' 1003) was chosen from the published sequence (33). The specificity of the probes for mRNA of expected molecular mass was confirmed by Northern blot analysis.
RT-PCR. One milligram of total RNA was reverse transcribed with 50 units Moloney murine leukemia virus RT (Perkin Elmer, Foster City, CA), 2.5 µM random hexamer, 1 mM dGTP, 1 mM dATP, 1 mM dTTP, 1 mM dCTP, and 20 units RNase inhibitor in a total volume of 20 µl for 30 min at 42°C. PCR was performed in a 50-µl reaction containing 0.5 units Taq polymerase (Perkin Elmer), 125 µM each dNTP, 2 mM MgCl2, 50 mM KCl, 10 mM Tris · HCl, pH 8.3, and 0.1 mM primers (both sense and antisense). PCR was carried out under the following conditions: 95°C for 5 min, 1 cycle; 94°C for 1 min, 60°C for 1 min, 72°C for 1 min, 30 cycles; and 72°C for 7 min, 1 cycle (Perkin Elmer Cetus DNA thermal cycler 480). The PCR reaction products (10 µl) were separated by electrophoresis on a 2.0% agarose/Tris acetate EDTA gel and stained with ethidium bromide.
Construction of a kinase inactive mutant of PKC-.
The PKC-
kinase inactive mutant construct was generated by the PCR
overlap extension method of Ho et al. (11). The following oligonucleotide primers were used in the mutagenesis protocol: 5'
primer, CCGCGTCGACCATGGTAGTGTTCAATGG; mutagen primer A,
CACAGCATA GAC;
mutagen primer B,
GTCTATGCTGT
AAA GAAGGACG; and
3' primer, ATTCGCGCGCTCAGGGCATCAGGTCTTCAC. The altered
sites introduced into primers A and B (underlined nucleotides)
were designed to mutate the original amino acids Lys-437 and Val-438 to
Arg-437 and Ala-438, respectively, and to introduce a new
Apa I restriction site (GGGCCC). The
mutant cDNA fragment generated by the PCR overlap extension procedure
was cloned into the mammalian epitope tagging expression vector
(
MTH), which contains the metallothionein promoter as described by
Olah et al. (25). The PCR reactions were performed with low (
10)
cycle number using the high-fidelity Vent DNA polymerase to minimize
the chance of undesired point mutations. The introduced ATP binding
site mutation was verified in selected clones by restriction digestions
and DNA sequencing. This PKC-
K437R cDNA construct has been verified
by stable transfection into NIH/3T3 fibroblasts in which the expressed
mutant protein was characterized by Western blotting, kinase activity,
and phorbol binding assays as well as by cell fractionation,
immunocytochemistry, and fluorescent microscopy experiments (26).
Transfection of HT29/A1 cells.
HT29/A1 cells were grown in RPMI 1640 containing 10% FBS. The cells
were transfected with either empty expression vector (MTH), with the
MTH expression vector containing the dominant-negative mutant
PKC-
K437R, or with the
MTH PKC-
wild-type vector containing full-length mouse PKC-
cDNA, using lipofectamine (GIBCO BRL), following the procedure recommended by the manufacturer. Stably transfected cell lines were selected with G418 (0.5 mg/ml). Resistant clones (10-20) from each transfection were selected at random and screened for protein expression by Western blot analysis. The
MTH
vector contains a zinc-inducible promoter. Thus transfected cells were
incubated in the presence and absence of 75 µM zinc acetate, as
noted, to induce synthesis of the indicated recombinant proteins.
Preparation of cell extracts for Western blotting.
For protein extraction, cells were washed with ice-cold PBS, scraped
into ice-cold buffer A [0.2% Triton X-100, 20 mM
Tris · HCl, pH 7.5, 0.25 M sucrose, 50 mM
-mercaptoethanol, 1 mM phenylmethylsulfonyl fluoride (PMSF), 200 µg/ml of leupeptin, 5 mM EDTA, and 2 mM EGTA]. The cells were
disrupted using a Dounce homogenizer. Homogenates were kept for 1 h at
4°C and then centrifuged at 100,000 g for 30 min at 4°C. The
supernatant was either used immediately for SDS-PAGE or stored at
80°C. Protein concentration was assayed with Coomassie blue
plus protein reagent (Pierce, Rockford, IL) using BSA as standard.
Western blotting.
Protein samples were analyzed using 10% SDS-PAGE (16). The separated
proteins were electrophoretically transferred to a polyvinylidene
difluoride (PVDF) membrane (Immobilon-P) using a Bio-Rad transfer blot
apparatus by the method of Towbin et al. (32). Nonspecific sites were
blocked by incubation of PVDF membranes with 4% nonfat dry milk
(Bio-Rad) or 3% BSA in 50 mM Tris · HCl, pH 7.5, 0.15 M NaCl, and 0.05% Tween 20 (TBST) for 1 h. The membranes then
were incubated at 4°C overnight with anti-PKC- antibody (GIBCO
BRL) at a dilution of 1:333. After three washes (15 min) with TBST,
secondary antibody [horseradish peroxidase (HRP)-conjugated goat
anti-rabbit IgG, (1:3,000 dilution) or HRP-conjugated rabbit anti-mouse
IgG, (1:5,000 dilution)] was added and incubated for 1 h at room
temperature. Membranes were washed three times, each for 15 min with
TBST, and bound antibody was dectected by the enhanced
chemiluminescence detection system (Amersham, Oakville, ON). When
necessary, blots were scanned by UVP Image System and the captured
image was analyzed for band densities by NIH Image software.
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RESULTS |
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PMA treatment of T84 cells stimulates mucin gene expression.
T84 human colonic cells were initially used to assess the effect of PMA
on the steady-state mucin mRNA levels. Cells at 80-90% confluency
were exposed to the PKC activator PMA at a concentration of 100 nM for
0-6 h, total RNA was extracted, and RNA slot-blot assays were
performed. Figure 1 shows the slot-blot
hybridization analysis of the effect of PMA treatment on the levels of
the housekeeping gene GAPDH and three mucin genes: MUC1, encoding a
membrane-anchored mucin, and MUC2 and MUC5AC, genes that encode
secretory mucins. The level of MUC1 mRNA showed a minimal change over 6 h. However, the expression of both MUC2 and MUC5AC was considerably
enhanced, particularly at 6 h. PMA treatment did not change the levels
of the GAPDH housekeeping gene. These results were quantitated by comparing density ratios of mucin mRNA to GAPDH mRNA. MUC2 expression increased almost 4-fold, and MUC5AC mRNA levels increased by 25-fold relative to the housekeeping gene, whereas MUC1 mRNA levels increased by <1.5-fold. In T84 cells the mRNA levels of two other secretory mucins, MUC5B and MUC6, were barely detectable and did not change during exposure to PMA (data not shown). Enhanced gene expression was
also not produced by refreshing the culture medium at the time of
adding PMA. These findings support the previous results of Velcich and
Augenlicht (37), which indicated that PMA treatment enhanced the mRNA
levels of MUC2 and importantly establish that a second secretory mucin
gene, MUC5AC, was even more dramatically regulated by a PMA-activated
PKC.
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PMA treatment of HT29/A1 cells also stimulates mucin gene
expression.
Experiments identical to those described above were
performed with HT29/A1 cells, a mucin-producing human colonic cell line known to express mucin at a relatively high level. The induction kinetics of HT29/A1 mucin mRNAs were similar to those found with T84
cells. Figure 2 shows a time course of
mucin gene expression in response to PMA treatment. Of the five mucin
genes examined, only the expression of MUC2 and MUC5AC was increased
more than twofold. In contrast to T84 cells, baseline mRNA levels in
HT29/A1 cells for MUC5B and MUC6 were detectable and were either
minimally increased (MUC6) or reduced (MUC5B) by exposure of these
cells to PMA. Similar to the results obtained with T84 cells, the
levels of MUC1 mRNA were only minimally affected in HT29/A1 cells.
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Overexpression of dominant-negative PKC- K437R
selectively blocks PMA-induced MUC2 and MUC5AC gene expression.
Previously, we suggested that PKC-
is the most likely PKC isoform
involved in regulating PMA-dependent mucin secretion in colonic cancer
cell lines (12) because LS-180 cells, which are selectively low in
PKC-
when compared with T84 and HT29/A1 cells, also responded poorly
to PMA. Although baseline expression of MUC2 and MUC5AC mRNAs were
ample in LS-180 cells, we could find little evidence of induced gene
expression when PMA was added (data not shown). PKC-
therefore
appeared to be a likely candidate for stimulating mucin gene expression
in T84 and HT29 cells.
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Overexpression of wild-type holo PKC-
does not prevent PMA-mediated induction of MUC2 and MUC5AC gene
expression.
To exclude the possibility that overexpression of the
dominant-negative PKC-
protein might have nonspecific effects on
mucin gene expression other than by acting to compete with active
endogenous PKC-
, we also stably transfected HT29/A1 cells with an
MTH expression vector containing wild-type holo PKC-
to generate
cells that would overexpress PKC-
when treated with zinc (Fig.
5A).
Exposure of these PKC-
overexpressor cells to PMA for 3 h resulted
in an increase in MUC2 and MUC5AC gene expression (Fig.
5B), as in wild-type cells. These
results indicate that the endogenous level of PKC-
is sufficient to
mediate PMA induction of mucin gene expression and that significant
overexpression of the PKC-
protein itself does not interfere with
the effects noted with PMA treatment.
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Overexpression of the dominant-negative PKC- K437R
mutant blocks the PMA-induced increase in mucin secretion in HT29/A1
cells.
Because we had previously obtained strong indirect evidence that
PKC-
mediated PMA-dependent mucin secretion in HT29/A1 cells (12),
we compared the secretory response to a standard 100 nM PMA stimulus
between cells infected with PKC-
K437R and two sets of controls,
i.e., cells transfected with the
MTH empty vector and parental
nontransfected cells. Zinc acetate (75 µM) was added to all cultures
6 h before adding PMA to ensure maximal expression of PKC-
K437R. As
in our previous study, the antibody used in the immunoassay was
specific for MUC2 (see MATERIALS AND
METHODS). As shown in Fig.
6, the presence of the dominant-negative
PKC-
K437R mutant virtually eliminated the secretory response. These results strongly support and confirm a critical role for PKC-
in
mediating MUC2 mucin secretion. Furthermore, the results provide an
important validation of the specificity of action of the
dominant-negative PKC-
, because the role of PKC-
in mediating
mucin secretion is strongly supported by independent evidence (12).
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DISCUSSION |
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Our results show that the expression of two mucin genes, MUC5AC and MUC2, is induced by the phorbol ester PMA. Both genes are located on chromosome 11p15.5 in a 400-kb region that also includes the genes for MUC5B and MUC6 (27). The four gene products have considerable sequence homology, particularly in their cysteine-rich carboxy-terminal regions, and their genes are thought to have evolved from a common ancestor. PMA had much less effect on the expression of MUC5B or MUC6, however, and may even decrease the expression of MUC5B, indicating that all genes in the cluster are unlikely to be controlled in the same manner by PMA. The two genes that do respond strongly are interesting, however, in that they are frequently coexpressed in goblet cells in the presence of duodenal and colonic inflammation (29). MUC2 is the dominant mucin in the normal intestine and is the major mucin expressed in colitis (34). However, when tissues are stained immunohistochemically for specific antigens, goblet cells that contain both MUC2 and MUC5AC are seen in areas of significant inflammation (29). Also, these two genes are coexpressed in many cells in areas of gastric metaplasia, an inflammatory response characterized by the appearance of cells with gastric phenotype in which the dominant mucin is MUC5AC (unpublished observations). Thus there is pathological evidence to suggest that intracellular stored amounts of these two gene products are regulated by inflammatory stimuli in intestinal cells.
PKC is a potentially important mediator of gene expression under these
conditions because it could be activated by a number of inflammatory
agents. The proinflammatory cytokine tumor necrosis factor-
(TNF-
) stimulates PKC activity, for example, in bovine bronchial
epithelial cells as part of the physiological response to wound healing
(39). In macrophages, PKC-
is activated by bacterial
lipopolysaccharide (LPS) and is the major isoform involved in the
LPS-induced secretion of TNF-
and other cytokines (30). PKC-
is
also induced by TNF-
to mediate inhibition of insulin signaling in
HEK-293 cells (13). As yet, the role of inflammatory agents in
regulating mucin gene expression is largely unexplored. TNF-
and LPS
have been shown to upregulate MUC2 in human airway epithelial cells
(20). TNF-
-mediated MUC2 gene expression was inhibited by calphostin
C and genistein, suggesting that signal transduction was dependent on
both PKC and tyrosine kinases.
In the present study, overexpression of a dominant-negative PKC-
mutant in HT29/A1 cells prevented the increase in MUC5AC and MUC2 mRNA
normally seen on exposure of these cells to phorbol ester. At the same
time, overexpression of wild-type PKC-
did not alter the increase in
mRNA levels for these two mucins when the cells were exposed to PMA,
indicating that endogenous PKC-
was sufficient for induction. These
results indicate that the inhibition of the PMA-induced increase in
MUC5AC and MUC2 mRNA levels by the dominant-negative PKC-
is not due
to overexpression of other domains of the PKC-
polypeptide.
The intestinal goblet cell secretes mucin by two processes, an
unregulated constitutive pathway that depends on the continuous movement of mucin granules from the Golgi to the apex of the cell and a
second regulated process that depends on the sudden release of mucin
from central storage granules (21). The first is directly related to
the rate of mucin synthesis and hence to mucin gene expression. The
second is independent of synthesis in the short term and is measured as
the increase in mucin output over baseline output that follows an
appropriate stimulus. We have now shown that PMA stimulates both
responses by activation of the same PKC isoform. In previous work we
demonstrated that PMA was able to stimulate mucin secretion
[subsequently identified as MUC2 secretion (22)] in T84
cells in a Ca2+-independent manner
(9). Later results pointed to mediation by the
Ca2+-independant isoform of PKC,
PKC-, because in three cell lines (LS-180, T84, and HT29/A1) the
secretory response to PMA correlated with the level and activation time
of PKC-
(12). Inhibition of PMA-regulated mucin secretion by a
dominant-negative PKC-
mutant in HT29/A1 cells, as demonstrated in
this study, provides the final confirmation that PKC-
mediates
PMA-dependent exocytosis of MUC2. PKC-
appears to mediate a variety
of secretory processes. For example, we have also shown in studies in
NIH/3T3 cells that PKC-
is involved in regulating the secretion of
glycosaminoglycans (18). Phorbol ester-dependent secretion of prolactin
was also increased by overproduction of PKC-
(2).
There are at least 12 closely related PKC isozymes that exhibit
distinct enzymological properties, differential tissue expression with
specific subcellular localization, and different modes of cellular
regulation (24). In HT29 and T84 cells, the major isoforms are PKC-,
PKC-
, and PKC-
(12), and of these only PKC-
and PKC-
are
activated by PMA. The role of PKC-
, which is abundantly expressed in
these cells, is not yet clear. However, it may be involved in
regulating cell differentiation, because it has been shown to modulate
growth and differentiation in the enterocyte colonic adenocarcinoma
cell line Caco-2 (1). The Caco-2 cell line differentiates into cells of
the enterocyte lineage, however, rather than into mucus-secreting
cells. Thus these findings may not be applicable to the HT29/A1 cells
used in the present study.
It is important to emphasize that our results do not discriminate
between an increase in transcription rate and mRNA stabilization. A
previous study (37) failed to determine conclusively whether phorbol
ester or forskolin acted to increase MUC2 transcription rates or
stabilize mRNA. Protein synthesis seemed to be required, suggesting
that activation of a rapidly degraded transcription factor such as the
activator protein-1 complex could be involved. Further studies now are
required to better define the mechanism(s) involved in mediating the
noted PKC--dependent changes in mucin gene expression.
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FOOTNOTES |
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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: G. Forstner, The Hospital for Sick Children, 555 Univ. Ave., Rm. 3423, Toronto, ON, Canada M5G 1X8 (E-mail: gforst{at}sickkids.on.ca).
Received 16 March 1999; accepted in final form 2 August 1999.
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