Division of Digestive Diseases, Department of Internal Medicine, and Departments of Pharmacology and Molecular Physiology, Rush University Medical Center, Chicago, Illinois 60612
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ABSTRACT |
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Loss of intestinal
barrier integrity is associated with oxidative inflammatory GI
disorders including inflammatory bowel disease. Using monolayers of
human intestinal epithelial (Caco-2) cells, we recently
reported that epidermal growth factor (EGF) protects barrier
integrity against oxidants by stabilizing the microtubule cytoskeleton,
but the mechanism downstream of the EGF receptor (EGFR) is not
established. We hypothesized that phospholipase C (PLC)- is
required. Caco-2 monolayers were exposed to oxidant (H2O2) with or without pretreatment with EGF or
specific inhibitors of EGFR tyrosine kinase (AG-1478, tyrphostin 25) or
of PLC (L-108, U-73122). Other Caco-2 cells were stably transfected
with a dominant negative fragment for PLC-
(PLCz) to inhibit PLC-
activation. Doses of EGF that enhanced PLC activity also protected
monolayers against oxidant-induced tubulin disassembly, disruption of
the microtubule cytoskeleton, and barrier leakiness as assessed by radioimmunoassay, quantitative Western blots, high-resolution laser
confocal microscopy, and fluorometry, respectively.
Pretreatment with either type of inhibitor abolished EGF protection.
Transfected cells also lost EGF protection and showed reduced PLC-
phosphorylation and activity. We conclude that EGF protection requires
PLC-
signaling and that PLC-
may be a useful therapeutic target.
phospholipase Cz transfection; tubulin; epidermal growth factor; barrier integrity; Caco-2 cells
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INTRODUCTION |
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THE EPITHELIUM OF THE GASTROINTESTINAL (GI) mucosa is a highly selective permeability barrier that normally restricts the passage of harmful proinflammatory and toxic molecules into the GI mucosa or systemic circulation. Disruption of this barrier is thought to play a pivotal role in inflammatory diseases of the GI tract such as inflammatory bowel disease (IBD) (32). For example, an increase in intestinal mucosal permeability can allow greater intestinal penetration of toxic luminal antigens (endotoxin or other bacterial fragments and by-products, food by-products, and dietary additives), leading not only to initiation of intestinal inflammation and tissue damage but also to a vicious cycle that can perpetuate the inflammatory cascade and sustain the active IBD attack (32, 37). Loss of mucosal barrier integrity is also characteristic of necrotizing entercolitis and a variety of other GI disorders such as gastric mucosal injury induced by ethanol as well as several systemic disorders such as alcoholic liver disease (4, 32, 36, 37). Indeed, disrupting the epithelial barrier and allowing bacterial endotoxin into the mucosal layer in animal models can initiate an oxidative inflammatory condition similar to IBD (60). Furthermore, in mice that are genetically engineered to exhibit a leaky paracellular route, loss of mucosal barrier function can lead to intestinal inflammation (31).
Because of our group's interest in developing more effective treatment
regimens for oxidative inflammatory disorders of the GI tract, we have
been investigating mechanisms of protection of intestinal barrier
integrity by endogenous growth factors such as epidermal
growth factor (EGF). Increasing our understanding of these protective
mechanisms should identify potential targets for new therapeutic
agents. Using monolayers of human intestinal (Caco-2) cells as a model
of intestinal barrier function, we previously showed
(5-8) that EGF and transforming growth factor
(TGF)- protect against oxidant-induced disruption of monolayer
barrier integrity by protecting the microtubule cytoskeleton. We have also shown (4, 5-14) that cytoskeletal stability is
key in GI mucosal healing in vivo in rats as well as under in vitro
conditions in cell cultures. In a recent study, we reported (5,
6) that protection by EGF against oxidant-induced injury was
mediated by EGF receptor (EGFR) and its tyrosine kinase. However, the
immediate downstream signal transducer responsible for the protective
effects of EGF on the microtubule cytoskeleton and intestinal barrier function is not known.
One likely candidate is phospholipase C (PLC)-. Studies in
epithelial cell models suggest that PLC-
signaling may be activated and phosphorylated by growth factor receptor tyrosine kinases via the
src homology 2 (SH2 or phosphotyrosine) domains (23, 43, 56). For example, PLC-
was proposed to be activated by the platelet-derived growth factor receptor and insulin-like growth factor-1 as well as by EGFR (19, 23, 39). It is also known that intestinal epithelial cells express the PLC-
isoform
(48). In the current investigation, we sought to determine
whether EGF-induced protection against oxidant injury to the
microtubule cytoskeleton and intestinal epithelial barrier integrity
occurs through activation of the PLC-
isoform. To this end, both
pharmacological as well as targeted molecular biological techniques
were used. The latter included the use of a dominant negative fragment
of PLC-
, namely PLCz, which contains SH2, SH3, and PLC-inhibitory
domains, and which is known to specifically inhibit PLC-
activation
but not other isoforms (23, 33, 56).
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MATERIALS AND METHODS |
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Cell culture.
Caco-2 cells, which were obtained from ATCC (Manassas, VA) at
passage 15, were chosen because they form monolayers that
morphologically resemble small intestinal cells, with defined apical
brush borders, junctional complexes, and a highly organized microtubule
network (11, 25, 44). Cells were maintained at 37°C in
complete DMEM in an atmosphere of 5% CO2 with 100%
relative humidity. Parental (wild type) cells and transfected cells
(see Stable transfection of dominant negative PLC-1 in Caco-2
cells) were split at a ratio of 1:6 on reaching confluence
and set up in either 6- or 24-well plates for experiments or in T-75
flasks for propagation. Cells grown for barrier function work were
split at a ratio of 1:2 and seeded at a density of 200,000 cells/cm2 into 0.4-µm BioCoat collagen I cell
culture inserts (0.3-cm2 growth surface; Becton Dickinson
Labware, Bedford, MA), and experiments were performed at least 7 days
postconfluence. The media were changed every 2 days. The utility and
characterization of this cell line has been previously reported
(11, 25, 44).
Stable transfection of dominant negative PLC-1 in Caco-2
cells.
The dominant negative PLC-
1 fragment from the Z region
(designated PLCz) was a generous gift from Dr. A. Wells (University of
Pittsburgh, Pittsburgh, PA). The Z region of human PLC-
1, which
covers the SH2 and SH3 domains (amino acids 517-901), was isolated
by RT-PCR and cloned into a eukaryotic expression vector, pXf
(23, 56). Expression was controlled by the SV40 early promoter present in pXf vector. Cultures of Caco-2 cells grown to
50-60% confluence were cotransfected with G418 resistance plasmid and PLCz (pXf/PLCz) with LIPOFECTIN (GIBCO BRL). Control conditions included vector (pXf) alone. Stable transfected cells were selected in
high-glucose DMEM-10% fetal bovine serum (FBS) supplemented with G418
(0.6 mg/ml). Stable expression of PLCz protein (~51 kDa) in these
cells was demonstrated by Western blot analysis with monoclonal
anti-PLC-
1. The PLCz-expressing cells were also tested for PLC
activity (see Experimental design). Clones stably expressing
PLCz were plated on Transwell cell culture inserts, allowed to form
confluent monolayers, and used subsequently for experiments. In
preliminary studies, we confirmed that PLCz expression did not injure
the Caco-2 cell monolayer barrier and that it did not affect the
expression levels of PLC-
.
Experimental design. In all experiments, barrier function, microtubule cytoskeletal integrity (intracellular architecture, tubulin assembly and disassembly), PLC activity, and PLCz protein expression were assessed. In the first series of experiments, postconfluent monolayers of wild-type Caco-2 cells were preincubated with EGF (1, 5, or 10 ng/ml) or isotonic saline for 10 min and then exposed to oxidant (0.5 mM H2O2) or vehicle (saline) for 30 min. These experiments were then repeated with cell monolayers stably expressing PLCz. Reagents, including inhibitors of EGFR-linked tyrosine kinase and of PLC, were applied on the apical side of monolayers unless otherwise indicated. The concentrations of oxidant or EGF used have been shown by our laboratory (5-8, 13, 14) to be effective at damaging and protecting, respectively, Caco-2 monolayers. Because our previous studies (6, 13) showed that the results were qualitatively similar regardless of whether apical or basolateral exposure of oxidants was used, all of the current studies used apical application.
In the second series of experiments, we further explored the possible importance of PLC signaling in EGF-mediated protection. For these studies, cell monolayers that were stably expressing PLCz were preincubated (10 min) with a high dose of EGF (10 ng/ml) or vehicle before the subsequent exposure (30 min) to damaging concentrations of oxidant (0.5 mM H2O2 or vehicle). In these transfection experiments, expression of PLCz protein was determined by immunoblotting. Inhibition of PLC activity as well as the tyrosine phosphorylation of PLC-PLC activity in Caco-2 cells. PLC activity was assayed by the formation of myo-[3H]inositol phosphates in cell monolayers as previously described (48, 56). Cells were labeled in serum-free medium containing 5 µCi/ml of myo-[3H]inositol for 24 h and subsequently washed three times to remove unincorporated label. LiCl (10 mM), which inhibits inositol phosphate hydrolysis by inositol phosphatases, was added 15 min before the addition of EGF and/or other treatments. Cells were then washed thrice with ice-cold PBS and lysed with 10% TCA. Cell lysates were collected, and the myo-[3H]inositol phosphates were recovered in the supernatant after centrifugation (16,000 g for 5 min). The extracts were separated on Dowex formate ion-exchange mini-columns (Bio-Rad, Hercules, CA). The radioactivity present (inositol phosphate content) in samples was quantified by scintillation counting with aqueous counting scintillant. Counts for blanks were subtracted from the sample activity. PLC activity was reported as counts per minute (cpm) per 106 cells.
Western immunoblotting of PLCz protein expression.
Differentiated cell monolayers grown in 75-cm2 flasks
were scraped and ultrasonically homogenized in lysis buffer (150 mM
NaCl, 50 mM Tris · HCl, 1 mM EDTA, 1 mM EGTA, 1% Nonidet P-40,
0.1% sodium deoxycholate, 0.1% SDS, and 2 µg/ml each of aprotinin, pepstatin, leupeptin, and phenylmethylsulfonyl fluoride). The samples
were then centrifuged (100,000 g for 40 min at 4°C), and the supernatant was used for bulk protein determination as assessed by
the Bradford method (20). For immunoblotting, samples (100 µg protein/lane) were added to SDS buffer (250 mM Tris · HCl, pH 6.8, 2% glycerol, and 5% mercaptoethanol), boiled for 5 min, and
separated on 7.5% SDS-polyacrylamide gels (7, 56).
Subsequently, proteins were transferred to nitrocellulose membranes
(0.2-µm pore size), blocked in 3% BSA for 1 h, and washed
several times with Tris-buffered saline. The immunoblotted proteins
were incubated for 2 h in Tween 20, Tris-buffered saline, 1% BSA,
and the primary mouse monoclonal anti-PLC-1 (an antibody that
recognizes PLCz; Transduction Laboratories and Santa Cruz
Biotechnology) at a 1:2,000 dilution for 1 h at room temperature.
A horseradish peroxidase-conjugated goat anti-mouse antibody (Molecular
Probes, Eugene, OR) was used as a secondary antibody at 1:4,000
dilution. Proteins on membranes were visualized by enhanced
chemiluminescence (ECL, Amersham, Arlington Heights, IL) and
autoradiography. The identity of the PLCz band (~51 kDa) was
ascertained with the use of a monoclonal antibody that recognizes PLCz.
Additionally, in the absence of the primary antibody to PLCz, no
corresponding band for PLCz was observed. Furthermore, prestained
molecular weights (Mr 34,900 and 53,900) were
run in adjacent lanes.
Immunoprecipitation and Western blot analysis of EGFR and
PLC-1 phosphorylation.
After treatments, confluent cell monolayers were lysed by incubation
for 20 min in 500 µl of cold lysis buffer (20 mM Tris · HCl,
pH 7.4, 150 mM NaCl, 10 µg/ml of anti-protease cocktail, 10%
glycerol, 1 mM sodium orthovanadate, 5 mM NaF, and 1% Triton X-100).
The lysates were clarified by centrifugation at 14,000 g for
10 min at 4°C. For immunoprecipitation (48), the lysates were incubated for 90 min at 4°C with monoclonal anti-EGFR (1:1,000 dilution, in excess) or monoclonal anti-PLC-
1 antibody
(1:500 dilution, in excess). The extracts were then incubated
with protein A Sepharose for 1 h at 4°C. The immunocomplexes
were collected by centrifugation (2,500 g for 5 min) in a
microfuge tube and washed three times with immunoprecipitation buffer
containing 5 mM Tris · HCl, pH 7.4, and 0.2% Triton X-100. The
resultant pellets were resuspended in a standard SDS sample buffer and
boiled at 95°C for 5 min before separation by 7.5% PAGE. Prestained
molecular weights (Mr 34,900 and 205,000) were
also run. Gels were transferred to nitrocellulose membranes, blocked
with 1% BSA and 0.01% Tween 20 in PBS for blotting by monoclonal
anti-phosphotyrosine (1:5,000 dilution; Transduction Labs) and for
detection of immune complexes by horseradish peroxidase-conjugated
secondary antibody, incubated with chemiluminescence reagents, and autoradiographed.
Immunofluorescent staining and high-resolution LSCM of
microtubules.
Cell monolayers were fixed in cytoskeletal stabilization buffer and
then postfixed in 95% ethanol at 20°C as we previously described
(4, 5, 11). Cells were subsequently processed for
incubation with a primary antibody, monoclonal mouse anti-
-tubulin (IgG1, rat/human reactive; Sigma, St. Louis, MO) at a 1:200 dilution for 1 h at 37°C. Slides were washed three times in Dulbecco's PBS (D-PBS) and then incubated with a secondary antibody
(FITC-conjugated goat anti-mouse; Sigma) at a 1:50 dilution for 1 h at room temperature. Slides were washed thrice in D-PBS and once with
deionized H2O and subsequently mounted in Aquamount
(Fisher). All antibodies were diluted with D-PBS containing 0.1% BSA.
After staining, cells were observed with an argon laser (
= 488 nm) equipped with a ×63 oil immersion plan-apochromat objective (NA
1.4, Zeiss). Single cells and/or a clump of two to three cells from
desired areas of the monolayers were processed with the image
processing software on a Zeiss ultra high-resolution laser scanning
confocal microscope to create "neat black" areas surrounding the
cells. The cytoskeletal elements were examined in a blinded fashion for
their overall morphology, orientation, and disruption as we previously
described (5, 6, 11). Briefly, 200 cells/slide (culture
well) were examined in four different fields by LSCM, and the
percentage of cells displaying normal microtubules was determined. The
microtubule cytoskeleton in Caco-2 cells was considered not normal on
the basis of one or more of the following criteria: collapse,
fragmentation, kinking, or disruption of the microtubule organizer
center. Slides were examined in a blinded fashion to avoid bias. We
coded them in such a way that the examiner had no knowledge of the
experimental protocol. The slides were decoded only after examination
was complete.
Microtubule (tubulin) fractionation and quantitative
immunoblotting of tubulin.
Polymerized (S2) and monomeric (S1) fractions of tubulin were isolated
as we previously described (5, 11). Fractionated S1 and S2
samples were flash-frozen in liquid N2 and then stored at
70°C until being immunoblotted. For immunoblotting, samples (5 µg
protein/lane) were placed in a standard SDS sample buffer, boiled for 5 min, and then subjected to PAGE on 7.5% gels. Procedures for Western
blotting were performed as previously described (5, 6). To
quantify the relative levels of tubulin, the optical density of the
bands corresponding to immunoradiolabeled tubulin was measured with a
laser densitometer.
Determination of barrier function by fluorometry.
Barrier integrity was determined by measuring apical-to-basolateral
flux of a membrane-impermeable fluorescent marker, fluorescein sulfonic
acid (FSA; 200 µg/ml; 478 Da) as we previously described (4, 5,
7). Briefly, fresh phenol-free DMEM (800 µl) was placed in the
lower (basolateral) chamber, and phenol-free DMEM (300 µl) containing
FSA was placed in the upper (apical) chamber. Aliquots (50 µl) were
obtained from the upper and lower chambers at time 0 and at
various subsequent time points and transfered to clear 96-well plates
(clear bottom; Costar, Cambridge, MA). Fluorescent signals from the
samples were quantitated with a fluorescence multiplate reader (FL 600, Bio-Tek Instruments). The spectra for FSA were 485 nm for excitation
and 530 nm for emission. Clearance (Cl) was calculated with the
following formula: Cl
(nl · h1 · cm
2) = Fab/([FSA]a × S), where Fab is the
apical-to-basolateral flux of FSA (in light units/h),
[FSA]a is the concentration at baseline (light units/nl),
and S is the surface area (0.3 cm2).
Simultaneous controls were performed with each experiment.
Statistical analysis. Data are presented as means ± SE. All experiments were carried out with a sample size of at least six observations per group. Statistical analysis was performed with ANOVA followed by Dunnett's multiple-range test (29). Correlational analyses were done with the Pearson test for parametric analysis or, when applicable, the Spearman test for nonparametric analysis. P values <0.05 were deemed statistically significant.
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RESULTS |
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Inhibitors of EGFR tyrosine kinase and of PLC prevent EGF-induced
protection of intestinal barrier function and of microtubule
cytoskeletal integrity.
First, we found that pretreatment with growth factor protects monolayer
barrier integrity against oxidant-induced barrier hyperpermeability as
assessed by reduced FSA clearance (Fig.
1), confirming our previous
reports (6, 7). We also confirmed that this protection is
EGFR-mediated because monoclonal anti-EGFR antibody (anti-EGFR, 1 µg/ml) abolished EGF-induced protection (data not shown).
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EGF causes enhancement of PLC activity in monolayers.
To further show that PLC is involved in EGF protection, we investigated
whether EGF increases PLC activity in Caco-2 cells as assessed by
radioimmunoassay of cell extracts from cells labeled with
myo-[3H]inositol. EGF enhanced PLC activity in
a dose-dependent manner (Fig. 5). Doses
of EGF known to be protective (e.g., 10 ng/ml) significantly increased
PLC activity; nonprotective doses of EGF (e.g., 1 ng/ml) did not.
Figure 6 shows that pretreatment with the
PLC inhibitors U-73122 and L-108 (but not the inactive U-73343) almost
completely inhibited the ability of protective EGF to stimulate PLC
activity. Preincubation with EGFR tyrosine kinase inhibitors caused
similar effects. The drugs by themselves had no effect on basal PLC
activity.
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PLCz reduces EGF-induction of PLC activity and EGF-mediated
protection of cell monolayers.
To further demonstrate a key role for PLC in EGF-induced
protection, wild-type Caco-2 cells were stably transfected with cDNA encoding a PLCz dominant fragment from the Z region of human PLC-1 (i.e., pXf/PLCz). Figure 7 shows that
PLCz (~51 kDa) (Fig. 7B) was detected by immunoblotting of
the cell lysates from these transfected cells. Native PLC-
(Fig.
7A) is demarcated as a protein with an apparent molecular
weight of 145. Figure 7 also shows that the control vector (pXf) did
not introduce the PLCz mutant. Normal parental (wild type) cells
lacking PLCz are shown as an additional control.
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PLCz mutant prevents EGF induction of PLC-1 phosphorylation.
Finally, both EGFR and PLC-
1 were immunoprecipitated from both the
PLCz-transfected and wild-type cells (those not expressing PLCz) and
then subjected to Western immunoblotting to assess tyrosine phosphorylation (Fig. 11).
In parental Caco-2 cells, the addition of EGF caused an increase in
tyrsosine phosphorylation of EGFR (Fig. 11A) and PLC-
1
(Fig. 11B). Expression of PLCz prevented the tyrosine
phosphorylation of PLC-
1 by EGF, suggesting that PLCz mutant is an
effective inhibitor of EGF ligand-induced PLC-
1 phosphorylation
and/or activation.
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DISCUSSION |
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Identifying and characterizing the intracellular mechanisms
underlying the ability of EGF to confer protection to the intestinal epithelial barrier is important because loss of GI barrier integrity is
considered to be a key mechanism in the pathogenesis of IBD (32). Because oxidative tissue damage is also implicated
in the pathophysiology of IBD (37, 42), we have been
investigating mechanisms by which oxidants disrupt the intestinal
barrier and growth factors protect it. Using monolayers of intestinal
cells as a model of barrier function, our previous reports (5,
6) demonstrated that damage to the microtubule cytoskeleton is
required for oxidant-induced damage to the intestinal barrier and that pretreatment with growth factors prevents this damage. We further showed that the EGFR and PKC activation mediate the ability of EGF to
protect both microtubule cytoskeletal organization and intestinal
barrier integrity (6-8). Continuing this probe into protective pathways, our current investigation demonstrates that PLC- is a critical downstream signal from the EGFR and is required for EGF-induced protection. To the best of our knowledge, this is the
first demonstration of this mechanism for the defense and protection of
intestinal epithelial barrier integrity.
Several independent lines of evidence support this conclusion. First, preincubation of oxidant-exposed monolayers with EGF not only increased PLC activity but also concomitantly prevented oxidant-induced intestinal monolayer hyperpermeability. In concert, EGF prevented oxidant-induced decreases in tubulin assembly (S2) and the percentage of cells with intact microtubule architecture and increases in tubulin disassembly (S1). Second, these protective effects of EGF were significantly attenuated by two different classes of inhibitors, those that inhibit EGFR tyrosine kinase and those that inhibit PLC, whereas the noninhibitory analogs of these same agents did not prevent protection by EGF.
Third, transfected cells in which PLC-1 activity was severely
attenuated by PLCz expression were no longer protected by EGF against
oxidant damage to tubulin assembly, microtubules, and barrier
integrity. These transfection experiments were done, in part, because
pharmacological inhibitors of PLC, namely U-73122 and L-108, are
specific to PLC but not to any one PLC isoform. Moreover, PLCz, the
dominant negative fragment of PLC-
1 obtained from the Z region of
PLC-
1, is known to contain the SH2-SH2-SH3 domains necessary for
activation and/or phosphorylation of PLC-
1 by EGFR (23, 33,
56). The concordance of our findings resulting from the use of
both pharmacological inhibition and molecular targeting supports a
central role for PLC-
in protection by EGF.
The findings of other investigators are also consistent with the
conclusions of our study. It is known that PLC-1 profoundly affects
similar cellular functions in epithelial cells as well as in
nonepithelial cells (23, 24, 27, 30, 33, 38, 53, 54, 57,
62). Several recent studies, for example, showed that migration
of intestinal cells that is stimulated by growth factor receptors
(i.e., a motogenic pathway) requires PLC-
1 activity (15, 21,
45, 48, 55). Additionally, PLC-
1 has been implicated in the
pathway for EGF-mediated remodeling of the actin component of the
cytoskeleton (16, 27, 34) and of EGF-induced regulation of
profilin and gelsolin (34). Furthermore, several recent
studies have suggested that PLC-
1 is the only epithelial PLC isoform
with SH2 and SH3 domains that may be activated by EGF ligands
(23, 48, 51).
PLC-1 hydrolyzes PIP2 to produce not only
inositol trisphosphate but also diacylglycerol (DAG) in epithelial
cells (23, 48, 56). DAG is one of the best-characterized
products of PLC-
-mediated reactions and is known to activate
serine/threonine protein kinase C (PKC) (3, 7,
46). This is consistent with the fact that we have
recently shown, with the use of pharmacological and targeted molecular
inhibition (i.e., antisense), that PKC, especially the PKC-
1
isoform, is an essential mediator of the EGF-enhanced signaling that
leads to the maintenance of intestinal barrier permeability (7,
8). Moreover, PKC activators (OAG or
12-O-tetradecanoylphorbol 13-acetate) can maintain both the cytoskeletal integrity and intestinal barrier function in the presence
of PLC inhibitors, suggesting the integral role of PKC activation
downstream of PLC-
1 in protection. Indeed, PKC activity has also
been shown to be PLC dependent in other systems (46, 61).
Also, two recent studies (2, 40) proposed that a naturally occurring intracellular activator of PKC, namely DAG (OAG used in our
recent studies is a synthetic version of this compound), modulates
intestinal monolayer permeability in Caco-2 cells. Overall, it appears
that growth factor-induced protection is mediated by PLC-
1 and then
PKC-
1. In our studies, specific inhibition of PLC-
1 was required
to demonstrate a causal relationship between this enzyme and
EGF-mediated protection.
First, two different specific inhibitors of PLC activity, U-73122 (and its control condition, the inactive analog U-73343) and L-108, were used to determine whether inhibition of PLC would prevent protection. These drugs had no deleterious side effects on the "basal" rate of the parameters under consideration (e.g., microtubules, barrier, PLC). Numerous previous studies have used an identical pharmacological approach to answer similar kinds of questions in epithelial cells, including studies in colonic cells (23, 43, 48, 62, 63). For example, using identical PLC inhibitors, Polk (48) demonstrated the importance of PLC in EGF-induced motility of intestinal epithelial cells.
Second, pharmacological inhibitors of PLC do not necessarily prove that
EGFR/PLC- signaling is critical to EGF-induced protection. To more
convincingly identify PLC-
as a required downstream signal for EGF
protection, we used a previously validated and widely used molecular
approach, PLCz mutant (22, 23, 33, 56). PLCz, which covers
both the SH2 and SH3 domains (amino acids 517-901), is known to
specifically inhibit PLC-
activation and/or phosphorylation and not
other PLC isoforms in epithelial cells (22, 23, 33, 56).
The PLCz mutant is specific for inhibition of PLC-
because extensive
previous studies from several independent labs have shown that PLC-
1
is the only epithelial PLC isoform that contains SH2 and SH3 domains
and that is activated by EGF (23, 33, 43, 48, 51, 56).
Indeed, in our own studies, PLCz dominant mutant expression prevented
the tyrosine phosphorylation (activation) of PLC-
1 (Fig. 11), while
at the same time, it abrogated activation of PLC (Fig. 8) by EGF.
Third, we note that although PLCz dominant negative mutant exerts its
inhibitory effect by binding to receptor phosphotyrosine motifs (i.e.,
SH2 motifs), it is possible that other, non-PLC- SH2-mediated
interactions may also be interrupted. However, this scenario seems
unlikely because 1) the U-73122 and L-108 (pharmacological) data point to a phospholipase (PLC) being the target of action and
2) the PLCz mutant data specifically indicate an SH2
domain-containing molecule as a requirement for EGF-induced protection.
PLC-
is the only candidate that, once again, fulfills both requirements.
Several other reports, including our own, suggest that our proposed
mechanism for EGF protection is both generalizable and relevant to IBD.
We observed protective effects by EGF when hypochlorous acid (HOCl) or
peroxynitrite (ONOO) were the oxidative agents used to
damage microtubules and the monolayer barrier (5-7).
A key role for these oxidants, including the oxidant
H2O2, in IBD is likely because the natural
course of IBD involves recurrent episodes of an inactive phase (where there are no neutrophils) followed by acute flare-up, as characterized by mucosal infiltration of large numbers of leukocytes including neutrophils (37, 42). These leukocytes are capable of
producing large quantities of reactive oxygen metabolites (e.g.,
H2O2 and HOCl) and reactive nitrogen
metabolites (e.g., ONOO
), reactive species that create a
vicious cycle and sustain an inflammatory cascade and tissue damage.
Additionally, in recent studies (5, 6, 13), we noted
similar protective effects for TGF-
, a structurally similar growth
factor synthesized by the intestinal mucosa that acts through the same
EGFR. In fact, no report has convincingly disassociated the biological
activities of EGF and TGF-
in any cell population, including GI
epithelium (28, 47). Indeed, it is likely that both growth
factors play a role in protection. Moreover, salivary EGF and EGF
contained within secretions of the Brunner's glands and exocrine
pancreas are the most important sources of gut EGF, and they play a
major role in protection of both the small and large intestine
(18, 47, 49, 50). For example, a previous study showed the
presence of EGF-like immunoreactivity in a novel cell lineage derived
from intestinal stem cells in the inflamed intestinal mucosa, such as
occurs in IBD and peptic ulcers (58). EGF also prevents
injury to the intestinal epithelium in animal models of IBD
(trinitrobenzenesulfonic acid) (18, 49) as well
as in vitro (Clostridium difficile toxin model of epithelial
damage to human colonocytes) (52).
MAPK inhibitors did not prevent EGF-induced protection in our intestinal model of barrier function (data not shown). It is therefore likely that MAPKs are not involved directly in EGF-mediated protection. Not surprisingly, MAPKs regulate many deleterious (not protective) pathways in cells such as programmed cell death, oxidative stress, and the inflammatory response (e.g., cytokine production, immune cell degranulation, and oxidant stress by "extracellular signal-regulated kinase"-type MAPK) as well as mitogenesis and differentiation responses (e.g., by p38 MAPK) (1, 26, 35, 41). For example, MAPK can be activated by oxidants and is a key cellular response to oxidant stress (an upstream target of MAPK, p21 ras, is a target of reactive oxygen species) (1). Furthermore, although it is known that EGF can signal MAPK, different cells respond differently to EGF (1, 17, 26, 59). For instance, activation of EGF/EGFR does not necessarily cause MAPK signaling in lung epithelial cells (i.e., A549 cells, an adenocarcinoma epithelial cell line similar to our Caco-2 colonic cancer cells) (26) as well as in several other cell types (23). Consistent with the above studies, several previous studies showed that MAPK is not always involved in EGF-induced effects (e.g., cell motility response) (23, 59).
In light of our new findings, it appears that an EGFR/PLC signaling
pathway is responsible for a significant fraction of the normal
protection of GI mucosal epithelium and perhaps is the key to
preventing amplification and perpetuation of an uncontrolled oxidant-induced inflammatory cascade that can be ignited by free radicals and other oxidants that are ever-present in the GI tract. Our
studies not only describe pathophysiological mechanisms, they also
suggest possible ways to develop novel therapeutic strategies. In
particular, our studies indicate that PLC-1 is a potential therapeutic target for pharmacological and perhaps genetic
interventions against a wide variety of oxidative inflammatory
conditions of the GI tract including IBD. For instance, one therapeutic
approach ("targeted gene therapy") might be the exogenous delivery
of a sense vector for the PLC-
1 isoform to the inflamed GI mucosa in
vivo. If either the pharmacological or the gene therapy approach is
successful, one should be able to protect and maintain epithelial integrity against oxidative stress, such as that which occurs during
incipient or rampant inflammation, and subsequently limit the
initiation and progression of GI mucosal inflammation and damage. Also,
such therapies might synergize with the use of antioxidants so that
inflammatory processes are attenuated through the manipulation of both
the damaging and protective intracellular pathways. In summary, our
findings demonstrate that PLC-
1 is a key mediator in protection of
the GI mucosal epithelium by growth factors.
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ACKNOWLEDGEMENTS |
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We thank Dr. A. Wells at the Pittsburgh University Medical Center for his generous help in providing the PLCz vector.
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FOOTNOTES |
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This work was supported, in part, by a grant from the Department of Internal Medicine at Rush University Medical Center and by a grant from the American College of Gastroenterology.
This work was presented, in part, at the annual meeting of the American Gastroenterological Association, May 2001.
Address for reprint requests and other correspondence: A. Banan, Rush Univ. Medical Center, Div. of Digestive Diseases, 1725 W. Harrison, Suite 206, Chicago, IL 60612 (E-mail: ali_banan{at}rush.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 7 March 2001; accepted in final form 23 April 2001.
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