Departments of Internal Medicine (Section of Gastroenterology and Nutrition), Pharmacology, and Molecular Physiology, Rush University School of Medicine, Chicago, Illinois 60612
Submitted 1 April 2003 ; accepted in final form 29 May 2003
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ABSTRACT |
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actin cytoskeleton; gut barrier; growth factors; oxidative stress; nitration and carbonylation; reactive nitrogen metabolites; phospholipase C isoform; inflammatory bowel disease; Caco-2 cells
An important discovery in the GI inflammation (IBD) field was the realization that a leaky and disrupted gut barrier can cause intestinal inflammation and that maintaining a normal mucosal epithelial barrier is required for intestinal health. For instance, intestinal barrier hyperpermeability that is induced by the injection of bacterial endotoxin into the mucosa of rodents can elicit an oxidative and inflammatory condition similar to IBD (60). Moreover, transgenic animals with a leaky gut barrier exhibit symptoms of intestinal inflammation (29). However, the pathophysiology of mucosal barrier disruption in IBD remains poorly understood. Nonetheless, several studies have shown that chronic gut inflammation in IBD is associated with excessive amounts of oxidants (e.g., H2O2) and that a high level of these oxidants appears to be a key contributor to mucosal injury (2, 10, 17, 18, 37, 39, 40, 43). Oxidant-induced disruption is of substantial clinical and biological value not only because oxidants are common in inflammation (e.g., they are elaborated by neutrophils that infiltrate the mucosa during inflammation) but also because they can lead to mucosal barrier dysfunction and, in turn, to the initiation and/or continuation of mucosal inflammation and injury (29-31, 38, 39, 60). Accordingly, understanding how gut barrier integrity can be protected against oxidative, proinflammatory conditions is of fundamental clinical and biological importance.
We have been investigating the mechanisms underlying oxidant-induced
mucosal injury and barrier disruption as well as protection against this
injury by growth factor pathways. Using monolayers of intestinal cells as a
well-established model of gut barrier integrity, we have shown that
cytoskeletal disassembly and disruption is a key event in oxidant injury and
that growth factors [EGF or transforming growth factor (TGF)-] appear
to prevent damage by stabilizing the cytoskeleton in large part through a
signaling pathway mediated by phospholipase C-
(PLC-
)
(1-3,
12,
18). The involvement in
protective mechanisms by PLC-
in the GI epithelium was a novel finding
(3,
12). We showed, using
wild-type Caco-2 intestinal cells, that EGF induces the membrane translocation
of the native PLC-
isoform and therefore considered it as a possible
contributor to EGF-mediated protection of the GI epithelial barrier. We then
noted that maintaining an intact cytoskeleton is required for protection of
intestinal barrier integrity by EGF apparently via PLC-
(3,
18). Despite the critical
importance of the
-isoform of PLC to intestinal barrier permeability,
the fundamental mechanism for PLC-
-mediated, EGF-induced protection of
monolayer barrier and actin cytoskeletal integrity remains elusive.
Inducible nitric oxide synthase (iNOS)-dependent processes are key in the underlying mechanism of oxidant-induced disruption of intestinal barrier integrity (9, 10). Indeed, overproduction and uncontrolled generation of iNOS-derived reactive nitrogen metabolites (e.g., NO, ONOO-) have been proposed to be an important factor in tissue damage during inflammation, including in IBD (17, 34, 37, 39, 40, 55). For example, we have shown that a number of these oxidative reactions, including cytoskeletal nitration and oxidation, also occur in intestinal mucosa from patients with IBD (17, 37) as well as in intestinal cell monolayers in culture (9, 10).
Accordingly, investigating the role of the -isoform of PLC in the
prevention of oxidative stress of iNOS-driven reactions in cells, we believe,
is both novel and significant because it is of substantial clinical and
biological importance to establish the idea that specific isoforms of PLC play
fundamental roles in endogenous protective mechanisms of cells against
oxidative stress to essential cellular structural proteins required for the
maintenance of GI integrity. Moreover, an improved understanding of
effectively suppressing (e.g., by PLC-
) the leakiness and disruption of
the intestinal barrier under conditions of oxidative stress should lead to the
development of new therapeutic modalities for inflammatory diseases of the GI
tract that are related to oxidative injury caused by hyperactivation of iNOS
and NO pathway.
In view of the above considerations, we tested the hypothesis that
PLC- not only prevents oxidant-induced iNOS upregulation and its
injurious consequences but also that it is key to EGF-mediated protection of
F-actin cytoskeleton and intestinal barrier integrity against the oxidative
stress of this upregulation. To this end, we utilized both pharmacological and
targeted molecular interventions employing several transfected intestinal cell
lines that we developed. In several clones the PLC-
isoform was
reliably overexpressed; in the other clones, PLC-
activity was severely
inhibited. Here, we report new mechanismsprevention of the oxidative
stress of iNOS upregulation and of cytoskeletal protein nitration and
oxidationby the
-isoform of PLC in cell monolayers. To our
knowledge, this is the first report that PLC-
can inhibit the dynamics
of iNOS-induced oxidative stress and cytoskeletal oxidation and disassembly in
cells.
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MATERIALS AND METHODS |
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Plasmids and stable transfection. The sense and dominant negative
plasmids of PLC- were constructed and then stably transfected by using
Lipofectin (Lipofectin reagent; GIBCO BRL) as we previously described
(3,
12). Expression was controlled
by SV40 early promoter present in pXf vector. The dominant negative
PLC-
1 fragment from the Z region (designated as PLCz) of human
PLC-
1, which covers the src homologous 2 and 3 (i.e., SH2 and
SH3) domains (amino acids 517-901), was isolated by RT-PCR and cloned into a
eukaryotic expression vector, pXf
(23,
57). Control conditions
included vector (pXf) alone. Multiple clones stably overexpressing PLC-
or lacking PLC-
activity were assessed by immunoblotting as well as
tested for PLC-
activity (see below). These cells were then plated on
Biocoat collagen I cell culture inserts (Becton Dickinson) and subsequently
used for experiments.
Experimental design. In the first series of experiments,
postconfluent monolayers of wild-type cells were preincubated with EGF (1-10
ng/ml) or isotonic saline for 10 min and then exposed to oxidant
(H2O2, 0-0.5 mM) or vehicle (saline) for 30 min. As we
previously showed, H2O2 at 0.5 mM disrupts actin
cytoskeleton and barrier integrity and upregulates iNOS
(2,
10,
18). EGF at 10 ng/ml (but not
1 ng/ml) prevents both actin and barrier disruption. These experiments were
then repeated using transfected cells. In all experiments, we assessed actin
cytoskeletal stability (cytoarchitecture, F-actin and G-actin
assembly/disassembly), barrier integrity, PLC- subcellular
distribution, PLC-
isoform activity, iNOS activity, NO levels, reactive
nitrogen metabolites (RNM) levels (e.g., ONOO-), oxidative stress
[dichlorofluorescein (DCF) fluorescence], actin nitration (nitrotyrosination),
and actin oxidation (carbonylation).
In the second series of experiments, cell monolayers that were stably
overexpressing PLC- were preincubated (10 min) with EGF (1 or 10 ng/ml)
or vehicle before exposure (30 min) to damaging concentrations of oxidant
(H2O2, 0.5 mM) or vehicle. Outcomes measured were as
described above.
In the third series of experiments, monolayers of dominant negative, namely
PLCz, transfected cells lacking PLC- activity were treated with high
(protective) doses of EGF and then oxidant. In corollary experiments, we
investigated the effects of PLC-
activation or inactivation on the
state of 1) actin nitration and oxidation, 2) actin assembly
and disassembly, and 3) stability of cytoarchitecture of the F-actin
cytoskeleton. Monomeric (G) and polymerized (F) fractions of actin were
isolated and then analyzed for outcomes (e.g., oxidation and nitration by
immunoblotting) (10,
18). Actin integrity was
assessed by 1) immunofluorescent labeling and fluorescence microscopy
to determine the percentage of cells with normal actin, 2) detailed
analysis by high-resolution laser scanning confocal microscopy (LSCM),
3) immunoblot analysis of G- and F-actin pools, and 4)
immunoblot analysis of oxidation and nitration of actin.
Fractionation and imunoblotting of PLC-. Cell monolayers
grown in 75-cm2 flasks were processed for the isolation of the
cytosolic, membrane, and cytoskeletal fractions
(7,
8). Protein content of the
various cell fractions was assessed by the Bradford method
(20). For immunoblotting,
samples (25 µg protein/lane) were added to a standard SDS buffer, boiled,
and then separated on 7.5% SDS-PAGE. The immunoblotted proteins were incubated
with the primary mouse monoclonal anti-PLC-
(Santa Cruz Biotechnology,
Santa Cruz, CA) at 1:2,000 dilution. A horseradish peroxidase (HRP)-conjugated
goat anti-mouse antibody (Molecular Probes, Eugene, OR) was used as a
secondary antibody at 1:4,000 dilution. Proteins were visualized by enhanced
chemiluminescence (ECL; Amersham, Arlington Heights, IL) and autoradiography
and subsequently analyzed. The identity of the PLC-
bands were
confirmed by 1) using a PLC-
blocking peptide in combination
with the anti-PLC-
antibody that prevents the appearance of the
corresponding "major" band in Western blots. 2)
Additionally, in the absence of the primary antibody to PLC-
, no
corresponding band for PLC-
was observed. 3) The PLC-
band ran at the expected molecular weight of 145 kDa as confirmed by a known
positive control for PLC-
(from rat brain lysates). 4)
Prestained molecular weight markers (Mr 34,900 and
205,000) were run in adjacent lanes. We also confirmed that overexpression of
PLC-
or dominant negative inhibition of PLC-
did not affect the
relative expression levels of other PLC isoforms and did not injure the Caco-2
cells.
Immunoprecipitation and PLC- activity.
Immunoprecipitated PLC-
was collected and processed for its ability to
form [3H]inositol phosphates
(12). Briefly, 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,
anti-protease cocktail (10 µg/ml), 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, the
lysates were incubated for 2 h at 4°C with monoclonal anti-PLC-
(1:1,000 dilution, in excess). The extracts were then incubated with protein
G-Sepharose for 1 h at 4°C. The immuno-complexes were collected by
centrifugation (2,500 g, 5 min) in microfuge tubes and washed three
times with immunoprecipitation buffer containing 5 mM Tris·HCl, pH 7.4,
and 0.2% Triton X-100. They were then washed one time with sample buffer (20
mM HEPES, pH 7.5) and resuspended in 20 µl of buffer and 5 µl of
reaction buffer (5 µCi/ml [3H]myoinositol) plus LiCl (10 mM,
which inhibits inositol phosphate hydrolysis) and subsequently incubated for 5
min at 30°C. Reactions were then stopped by the addition of 8 µl of
5x sample buffer, and the [3H]inositol phosphates (IP) were
recovered in the supernatant after centrifugation (16,000 g, 5 min).
The extracts were separated on Dowex formate ion-exchange minicolumns
(Bio-Rad, Hercules, CA). Radioactivity present (IP content) in samples was
quantified by scintillation counting with aqueous counting scintillant. Counts
for blanks were subtracted from the sample activity. Sample activity was also
corrected for protein concentration (Bradford method), and PLC-
activity was reported as picomoles per minute per milligram of protein.
Assay of NOS activity. Wild-type and transfected cells grown to confluence were removed by scraping and were centrifuged and homogenized on ice in a buffer containing 50 mM Tris·HCl, 0.1 mM EDTA, 0.1 mM EGTA, 12 mM 2-mercaptoethanol, and 1 mM phenylmethylsulfonyl fluoride, pH 7.4. Conversion of L-[3H]arginine (Amersham) to L-[3H]citrulline was measured in the cell homogenates by scintillation counting. Experiments in the presence of NADPH, without Ca+2 and with 5 mM EGTA, determined Ca2+-independent NOS (iNOS) activity (1, 4, 9, 10, 16).
Western blot of the level of iNOS. After treatments, the cells were washed once with cold PBS, scraped into 1 ml of cold PBS, and harvested in a standard anti-protease cocktail. For immunoblotting, samples (25 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 then separated on 7.5% SDS-PAGE. Subsequently, proteins were transferred to nitrocellulose membranes and then blocked in 3% BSA for 1 h, followed by several washes (Tris-buffered saline). The immunoblotted proteins were incubated for 2 h in Tris-buffered saline containing Tween 20 and 1% BSA with the primary antibody (mouse monoclonal anti-human iNOS, 1:3,000 dilution; Santa Cruz Biotechnology). An HRP-conjugated goat anti-mouse antibody (Molecular Probes) was used as a secondary antibody, at 1:3,000 dilution. Membranes were visualized by ECL and then autoradiographed (4, 9, 10, 16).
Chemiluminescence analysis of NO. NO production was assessed by a chemiluminescence procedure (4, 9, 10, 16). Briefly, cells were homogenized, and the endogenous nitrate (NO3 -) and nitrite (NO2 -), the metabolic degradation products of NO, were then reduced to NO by using vanadium (III) (Sigma, St. Louis, MO) and HCl at 90°C before measuring the NO concentration with a model 280 nitrix oxide analyzer (NOA) from Sievers (Boulder, CO). NO was expressed in micromolar concentration and calculated by comparison to the chemiluminescence of a standard solution of NaNO2. The absolute NO values were reported as the number of micromoles per 1 x 106 cells.
Determination of cell oxidative stress. Oxidative stress was assessed by measuring the conversion of a nonfluorescent compound, 2',7'-dichlorofluorescein diacetate (DCFD; Molecular Probes) into the fluorescent dye DCF (1, 2, 4, 10, 15). Monolayers grown in 96-well plates were preincubated with the membrane-permeable DCFD (10 µg/ml for 30 min) before the treatments. Subsequently, fluorescent signals (i.e., DCF fluorescence) from samples were quantitated using a fluorescence multiplate reader set at an excitation wavelength of 485 nm and an emission wavelength of 530 nm. DCF fluorescence was then expressed as a percentage of baseline oxidative stress. The dependence of the assay on reactive oxygen species (ROS) production (e.g.,.O2 - generation) was shown as we previously reported (1, 4, 9, 10) by adding either catalase, an active H2O2 oxidant scavenger, or SOD, an active superoxide radical scavenger, or, for control conditions, either an inactive H2O2 or inactive superoxide scavenger [heat-inactivated catalase or SOD (iSOD), respectively]. Similarly, we previously showed (1, 4, 9, 10) the dependence of this assay on RNM production (e.g., NO or ONOO- generation) by adding either an RNM scavenger (e.g., cysteine or urate) or an inhibitor of RNM biosynthesis [e.g., N6-(1-iminoethyl)-L-lysine (L-NIL)].
Immunofluorescent staining and high-resolution LSCM of actin
cytoskeleton. Cells from monolayers were fixed in cytoskeletal
stabilization buffer and then post-fixed in 95% ethanol at -20°C as we
previously described (10,
15,
18,
59). Cells were subsequently
processed for incubation with FITC-phalloidin (specific for F-actin staining;
Sigma), at 1:40 dilution for 1 h at 37°C. After staining, cells were
observed with an argon laser ( = 488 nm) using a x63
oil-immersion plan-apochromat objective, NA 1.4 (Zeiss). The cytoskeletal
elements were examined in a blinded fashion for their overall morphology,
orientation, and disruption (1,
2,
10,
11,
13,
14,
18). The identity of the
treatment groups for all slides was decoded only after examination was
complete.
Actin fractionation and quantitative Western immunoblotting of F- and G-actin. Polymerized (F) actin and monomeric (G) actin were isolated by using a especially developed series of extraction and ultracentrifugation steps as we described previously (10, 18). Fractionated F- and G-actin samples were then flash frozen in liquid N2 and stored at -70°C until immunoblotting. For immunoblotting, samples (5 µg protein/lane) were placed in a standard SDS sample buffer, boiled, and then subjected to PAGE on 7.5% gels. Standard (purified) actin loading controls (5 µg/lane) were also run concurrently with each run. To quantify the relative levels of actin, the optical density of the bands corresponding to immunolabeled actin were measured with a laser densitometer.
Immunoblotting determination of protein actin oxidation and actin nitration. Oxidation and nitration of the actin cytoskeleton were assessed, respectively, by measuring protein carbonyl and nitrotyrosine formation (10, 18). To avoid unwanted oxidation of actin samples, all buffers contained 0.5 mM dithiothreitol (DTT) and 20 mM 4,5-dihydroxy-1,3-benzene sulfonic acid (Sigma). To determine the carbonyl content, samples were blotted to a polyvinylidene difluoride (PVDF) membrane, followed by successive incubations in 2 N HCl and 2,4-dinitrophenylhydrazine (DNPH; 100 µg/ml in 2 N HCl; Sigma) for 5 min each. Membranes were then washed three times in 2 N HCl and subsequently washed seven times in 100% methanol (5 min each), followed by blocking for 1 h in 5% BSA in 10x PBS-Tween 20 (PBS-T). Immunologic evaluation of carbonyl formation was performed for 1 h in 1% BSA/PBS-T buffer containing anti-DNPH (1:25,000 dilution; Molecular Probes). Membranes were then incubated with an HRP-conjugated secondary antibody (1:4,000 dilution, 1 h; Molecular Probes). To determine nitrotyrosine content, after the blocking step described above (i.e., BSA/PBS-T buffer), membranes were probed for nitrotyrosine by incubation with 2 µg/ml monoclonal anti-nitrotyrosine antibody for 1 h (Up-state Biotech, Lake Placid, NY), followed by the HRP-conjugated secondary antibody (as above). Processing and film exposure were as in a standard Western blot protocol. The relative levels of oxidized or nitrated actin were then quantified by measuring, with a laser densitometer, the optical density (OD) of the bands corresponding to anti-DNPH (carbonylation) or anti-nitrotyrosine (nitration) immunoreactivity. Immunoreactivity was reported as the carbonyl or nitrotyrosine formation (OD) in the treatment group compared with the maximally oxidized or nitrated actin standards, expressed as a percentage. Oxidized actin standards (5 µg/lane) were run concurrently with corresponding treatment groups.
Determination of barrier permeability by fluorometry. The status of the integrity of monolayer barrier function was confirmed by a widely used and validated technique that measures the apical-to-basolateral paracellular flux of fluorescent markers such as fluorescein sulfonic acid (FSA; 200 µg/ml, 0.478 kDa) as we and others have described previously (1, 2, 6-12, 18, 33, 36, 58). Briefly, fresh phenol-free DMEM (800 µl) was placed into the lower (basolateral) chamber, and phenol-free DMEM (300 µl) containing probe (FSA) was placed in the upper (apical) chamber. Aliquots (50 µl) were obtained from the upper and lower chambers at time 0 and at subsequent time points and transferred into clear 96-well plates (clear bottom; Costar, Cambridge, MA). Fluorescent signals from samples were quantitated using a Fluorescence multiplate reader (FL 600; BIO-TEK Instruments). The excitation and emission spectra for FSA were as follows: excitation = 485 nm, emission = 530 nm. Clearance (Cl) was calculated using the following formula: Cl (nl·h-1·cm-2) = Fab/([FSA]a x S), where Fab is the apical-to-basolateral flux of FSA (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 treatment group. Statistical analysis comparing treatment groups was performed using analysis of variance followed by Dunnett's multiple range test (27). Correlational analyses were done using 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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Stable overexpression of PLC- isoform protects against
oxidative damage to the cytoskeleton: inhibition of both actin nitration and
oxidation. Using both our wild-type and transfected cells, we measured
the "footprints" of RNM formation, nitrotyrosine moieties, under
conditions of oxidant challenge. We also simultaneously measured oxidation
footprints by assessing the carbonylation levels. This was done by
sequentially fractionating and purifying the 43-kDa actin molecule from cell
monolayers and subsequently immunoblotting these fractions. In wild-type cells
(those not overexpressing PLC-
), oxidant H2O2
alone resulted in a substantial levels of nitration and oxidation of the actin
cytoskeleton (Fig.
1A). In contrast, overexpression of PLC-
by itself
afforded protection against oxidant-induced actin nitration and actin
carbonylation compared with those in wild-type cells. Indeed, only cells
stably overexpressing PLC-
were protected against oxidant-induced
nitration and oxidation injuries. Protection did not require the presence of
the growth factor EGF in the cell culture media. Although 1 ng/ml EGF did not
afford significant protection against actin nitration or oxidation in
wild-type cells, this concentration did potentiate the protection observed in
cells overexpressing PLC-
. In wild-type cells, higher doses of EGF (10
ng/ml) were required for protection (Fig.
1A). Transfection of only the empty vector did not confer
protection against oxidation and nitration. For instance, the percentage of
actin that was nitrated was 0% for vector-transfected cells exposed to
vehicle, 0.73 ± 0.28% for vector-transfected cells exposed to
H2O2 alone, and 0.11 ± 0.5% for PLC-
sense-transfected cells incubated in H2O2. Similarly,
the percentage of actin that was carbonylated was 0% for vector-transfected
cells exposed to vehicle, 0.77 ± 0.25% for vector-transfected cells
exposed to H2O2 alone, and 0.09 ± 0.34% for
PLC-
sense-transfected cells incubated in H2O2.
These oxidative alterations did not appear to be caused by changes in the
ability of oxidants to cause oxidation/nitration because vector-transfected
cells and wild-type cells responded in a similar fashion to
H2O2, exhibiting comparable actin oxidation.
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Figure 1, B and
C, shows representative immunoblots of the alterations in
actin nitration and carbonylation. For instance, PLC- overexpression
substantially inhibited both actin nitration
(Fig. 1B) and
oxidation (Fig. 1C) as
shown by reduced band (lane) densities to a level close to that of controls,
indicating prevention of oxidative damage to the actin cytoskeleton in cells
overexpressing PLC-
. As above, only high (protective) doses of EGF
(e.g., 10 ng/ml) prevented actin oxidation and nitration in wild-type cells.
In contrast, oxidant caused the oxidation and nitration of actin in these
wild-type cells.
PLC--induced protection involves downregulation of
iNOS-driven reactions: inhibition of iNOS, NO, RNMs
(ONOO-), and oxidative stress. Because oxidants such
as H2O2 upregulate iNOS
(1,
15), we hypothesized that
inhibition of iNOS-driven pathways might be a key mechanism for
PLC-
-induced protection. To this end, multiple clones of intestinal
cells transfected with 1, 2, 3, or 5 µg of PLC-
sense cDNA showed
(Table 1) a dose-dependent
inhibition of iNOS upregulation (L-[3H]citrulline
formation) against oxidant (H2O2)-induced challenge. The
clone transfected with 3 µg of PLC-
sense provided the maximum
inhibition of iNOS upregulation against oxidative insult. Accordingly, we used
this stable (
3) clone in subsequent experiments.
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Figure 2A shows
that PLC- overexpression using the 3-µg sense-transfected clone,
which protects gut barrier integrity
(3), also caused a substantial
reduction in calcium-independent iNOS activity (
96% lower iNOS activity).
This is comparable to that of the controls, which displayed only low iNOS
activity. These measurements were done in lysates of both transfected and
nontransfected Caco-2 monolayers. In wild-type cells, this same dose of
H2O2 caused both hyperpermeability and increases in iNOS
activity. PLC-
-induced inhibition of iNOS upregulation did not require
EGF. However, a low EGF concentration, 1 ng/ml, which did not by itself afford
inhibition of iNOS in wild-type cells, potentiated PLC-
-induced iNOS
downregulation in transfected cells. Wild-type cells, which have native levels
of PLC-
, required a higher dose of EGF (10 ng/ml,
Fig. 2A). Transfection
of the empty vector alone did not confer protection against oxidant-induced
iNOS hyperactivation (iNOS activity was 0.48 ± 0.03
pmol·min-1·mg protein-1 for
vector-transfected cells exposed to vehicle, 5.95 ± 0.28
pmol·min-1·mg protein-1 for
vector-transfected cells exposed to H2O2 alone, and 0.65
± 0.23 pmol·min-1· mg protein-1 for
PLC-
sense-transfected cells incubated in
H2O2).
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Figure 2B depicts a
representative Western blot showing that H2O2
significantly increased iNOS protein levels in wild-type cells, whereas
transfected cells overexpressing PLC- exhibited only low, basal levels
of the iNOS protein. For example, the corresponding OD values were 857
± 78 for control, 4,518 ± 92 for 0.5 mM
H2O2, and 963 ± 106 for PLC-
sense-transfected cells incubated in H2O2. Transfection
of empty vector alone, similar to its lack of effects on iNOS activity and
actin oxidation, was ineffective in preventing iNOS protein upregulation (not
shown).
NO is the product of the iNOS-catalyzed reaction.
Figure 3 shows NO levels both
in transfected monolayers and in wild-type monolayers exposed to
H2O2 as determined by sensitive chemiluminescence
analysis of cell lysates. PLC- overexpression markedly prevented
oxidant-induced NO overproduction (Fig.
3). In wild-type cells, as for actin oxidation and iNOS
upregulation, NO overproduction was inhibited only by high, protective doses
of EGF (e.g., 10 ng/ml). Transfection of vector alone did not confer
protection against NO overproduction (not shown).
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Table 1 also depicts the
results of NO analysis from multiple clones of transfected,
PLC--overexpressing intestinal cells showing a dose-dependent
inhibition of NO overproduction. As for iNOS suppression, the 3-µg stable
clone of PLC-
sense (
3) provided the highest protection against
NO overproduction.
Figure 4 shows the time
course for increases in iNOS protein, iNOS activity, and NO levels under
oxidative conditions and their prevention in transfected cells. PLC-
overexpression prevented the effects of H2O2 on all
three outcomes. Maximal fold increases under H2O2 alone
were
5.2 for iNOS protein,
12 for iNOS activity, and
12 for NO
levels; these increases were prevented by PLC-
overexpression.
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In parallel with the suppression of oxidant-induced affects, PLC-
overexpression inhibited oxidative stress as determined by a reduction in the
fluorescence of DCF (Fig. 5).
In wild-type cells, where H2O2 substantially increased
DCF fluorescence, oxidative stress was suppressed only by high, protective
doses (e.g., 10 ng/ml) of EGF. In the absence of oxidant, we observed
significantly lower but still substantial levels of oxidative stress [possibly
due to the normal generation of DCF reactive oxygen radicals (e.g.,
*O2-) by well-known cellular metabolic
processes such as the mitochondrial respiratory chain reactions
(1,
4,
9,
10)]. Transfection of the
empty vector alone did not suppress oxidative stress (not shown).
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Suppression of iNOS upregulation in transfected cells protects the
assembly of actin and the cytoarchitecture of F-actin cytoskeleton.
Because it is known that oxidants in this intestinal model disrupt the
cytoskeleton, we assessed the state of actin polymerization and its
intracellular architecture. PLC- overexpression confered protection to
the assembly of F-actin pool (Fig.
6) as well as the cytoarchitecture of actin cytoskeleton
(Fig. 7, a-c). For
example, to determine effects of PLC-
overexpression on the dynamic
alterations in the polymerization states of the F-actin, we performed
immunoblotting of actin cytoskeleton. To this end, the polymerized actin
fraction (F-actin, an index of actin stability) was isolated from monolayers.
Figure 6 shows that
PLC-
-overexpressing monolayers, which were exposed to oxidant,
exhibited a stable F-actin assembly, as indicated by an enhancement in this
polymerized actin fraction (i.e., increased band density). This state of
assembly is comparable to that of controls. In wild-type cells, in contrast,
oxidant decreased polymerized F-actin, indicating disassembly of actin
cytoskeleton. In these wild-type cells, only pretreatment with the higher
doses (10 ng/ml) of EGF resulted in a stable actin assembly. Indeed, confocal
microscopy corroborates this finding, showing that intestinal cells
overexpressing PLC-
had a smooth and normal architecture of the actin
cytoskeleton even after exposure to oxidant
(Fig. 7c). This
preserved appearance was indistinguishable from that of control (and
untreated) cells (Fig.
7a), which also showed an intact organization of the
actin cytoskeleton. In contrast, wild-type cells (not overexpressing
PLC-
) that are challenged with H2O2 exhibit
instability, fragmentation, and disruption of the actin cytoskeleton
(Fig. 7b). This
protection of both the assembly and cytoarchitecture of actin-based
cytoskeleton by PLC-
overexpression parallels the protective effects of
this overexpression against oxidant-induced iNOS and NO upregulation as well
as actin oxidation.
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Intracellular distribution and constitutive activation of the
overexpressed PLC- in transfected intestinal cells
correlates with several different indexes of iNOS and oxidative stress in
monolayers. Overexpressing the 145-kDa PLC-
in intestinal cells
led to its distribution into mostly the particulate fractions (particulate =
membrane + cytoskeletal fractions), with a much smaller distribution in the
cytosolic fractions (Fig.
8A), suggesting the constitutive activation of the
-isoform of PLC. In wild-type cells
(Fig. 8B), in
contrast, we found a mostly cytosolic distribution of PLC-
, with
smaller pools in the membrane and cytoskeletal (particulate) fractions,
suggesting inactivity of this isozyme.
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Table 2 is an analysis of
the intracellular distribution of the PLC- in various fractions of
either transfected or wild-type Caco-2 cell monolayers. Overexpressed
PLC-
isoform is "constitutively active" because achieving
this intracellular distribution did not require EGF or pharmacological
intervention. Pretreatment of these cells with EGF, however, enhanced the
fraction of PLC-
isoform in the membrane and cytoskeletal fractions,
reaching near total levels for PLC-
. On the other hand, in wild-type
cells PLC-
is found in a mostly cytosolic distribution (suggesting
inactivity), with smaller pools in the membrane and cytoskeletal (particulate)
fractions. Wild-type cells incubated with EGF also showed increased membrane
and cytoskeletal distribution of native PLC-
.
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Figure 9 shows the activity
levels of PLC- isoform (determined by in vitro assay) from
immunoprecipitated particulate cell fractions of Caco-2 cells, which were
stably transfected with PLC-
cDNA to overexpress this isoform. There
was a substantial increase in the activity levels of PLC-
isoform in
these transfected (vehicle exposed) cells, confirming findings in
Fig. 8 and
Table 2. EGF further activated
PLC-
in these transfected cells, reaching near maximal activation
levels for this isoform. Wild-type cells exposed to vehicle, in contrast,
showed basal activity levels for PLC-
in the particulate cell
fractions. In these wild-type cells, EGF further activated native PLC-
,
but at much lower levels compared with that of the transfected cells under
similar conditions.
|
Using data across all experimental conditions, we found significant inverse
correlations (e.g., r = -0.93, P < 0.05) between
PLC- levels (in vitro assay or optical density from the particulate
fraction) and iNOS downregulation, further suggesting that activation of
-isoform of PLC is key in protection against oxidant-induced iNOS
upregulation. Other robust correlations were seen when either NO
overproduction or oxidative stress (DCF fluorescence) was correlated with the
PLC-
levels (r = -0.90 or -0.89, respectively, P <
0.05 for each). When two other markers of oxidative stress, actin
carbonylation and actin nitration (RNM generation), were correlated with
PLC-
, additional robust correlations were observed (r = -0.95
or -0.94, respectively, P < 0.05 for each), further indicating
that activation of
-isoform is key in iNOS downregulation through
normalization of NO levels. Similarly, when markers of stability such as
either actin integrity or actin assembly were correlated with the PLC-
,
robust correlations were seen (r = 0.88 or 0.91, respectively,
P < 0.05 for each).
Stable dominant negative inhibition of PLC- by PLCz
fragment to inactivate native
-isoform and its prevention of
EGF-induced protection against oxidative stress of iNOS upregulation. The
above findings collectively indicate that PLC-
may play an essential
intracellular role in protection against oxidative stress of iNOS-driven
reactions. To independently investigate a possible role for PLC-
in
EGF-mediated protection against iNOS upregulation and consequent RNM driven
oxidative stress, we used stable dominant negative transfected PLCz clones of
Caco-2 cells, which we developed. To this end, cDNA encoding a PLCz dominant
negative fragment from the Z region of human PLC-
1 was utilized. Using
this dominant negative approach for PLC-
, we are capable of
substantially reducing the steady-state activity levels for native isoform by
99.3% (Fig. 9, 3-µg
clone). In these dominant negative PLCz cells, EGF could not increase the
native PLC-
isoform activity.
Table 1 further demonstrates
the dose-dependent effects of varying amounts (1, 2, 3, or 5 µg) of
PLC- dominant negative plasmid (i.e., PLCz) on suppression of both
EGF-induced iNOS downregulation and NO normalization in intestinal cells. The
cell clone that was stably transfected with 3 µg of PLCz dominant negative
plasmid resulted in maximum inability of EGF to prevent oxidant-induced iNOS
upregulation or NO overproduction. Thus this clone was utilized for other
inhibition experiments.
For example, we have shown (Fig.
10) that stable dominant negative inhibition of native PLC-
activity substantially prevented the protection afforded by 10 ng/ml EGF
against iNOS upregulation. In wild-type (naive) cells, on the other hand, this
same concentration of EGF almost completely prevented oxidant-induced iNOS
upregulation. A very large percentage (
90%) of EGF-induced iNOS
downregulation is PLC-
dependent.
|
Analysis of both the NO levels and oxidative stress from these dominant
negative transfected cells additionally demonstrates that inactivation of
native PLC- isoform substantially attenuated both EGF's normalization
of NO levels (Fig.
11A) and downregulation of oxidative stress
(Fig. 11B, DCF
fluorescence). As for iNOS downregulation, a large percentage
(
90%) of EGF-induced NO normalization and DCF fluorescence downregulation
appears to be PLC-
dependent in intestinal monolayers.
|
Furthermore, immunoblotting analysis of the oxidative state of actin
(Fig. 12A) from these
same dominant negative clones further shows that stable inactivation of the
-isoform prevented EGF-induced protection against both actin nitration
and oxidation. PLC-
isoform inactivation by itself did not cause actin
oxidation. Finally, analysis of the state of actin assembly from these
dominant negative cells demonstrates (Fig.
12B) that inhibition of native PLC-
attenuated
protection against actin depolymerization by a high (protective) dose of EGF.
Here, EGF could no longer prevent oxidant-induced actin disassembly.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
These conclusions are based on several independent lines of findings.
Expression of PLC- mimics an EGF-like protection against
oxidant-induced iNOS upregulation, including downregulation of iNOS
activation, normalization of NO levels, reduction of RNM footprints, and
decreases in oxidative stress (DCF fluorescence). Moreover, activation of the
PLC-
suppresses the footprints of oxidative injury (i.e., RNM
formation) to the 43-kDa actin protein. These other protective effects include
decreases in the nitration (nitrotyrosination) of the actin molecule and
reduction of oxidation (carbonylation) of actin. In concert, PLC-
activation decreased the monomeric (G) actin and enhanced the stability of
polymerized (F) actin as well as preserved appearance of normal actin
cytoarchitecture. Additionally, a low, nonprotective concentration of EGF
potentiated all measures of PLC-
-mediated protection against oxidative
stress of iNOS upregulation. Furthermore, dominant negative PLC-
(i.e.,
PLCz mutant), which causes almost complete inactivation of native PLC-
,
substantially prevented EGF's protective ability to suppress iNOS
upregulation, actin instability, and F-actin disruption. EGF was also unable
to inhibit nitration and carbonylation of actin, normalize NO levels, or even
reduce DCF fluorescence in these PLCz mutant cells. Finally, PLC-
activation quantitatively correlated with decreases in all outcomes indicating
protection against oxidative stress.
Using both transfected and wild-type cells, we found correlations
1) between PLC- isoform activation and protection against
oxidant-induced iNOS upregulation (r = -0.93, P < 0.05)
as well as several other key outcomes. These others included 2)
protection against oxidant-induced NO overproduction and PLC-
activation (r = -0.90, P < 0.05), 3) actin
nitration (RNM footprint) and PLC-
activation (r = -0.94,
P < 0.05), and 4) oxidative stress (DCF fluorescence)
levels and PLC-
activation (r = -0.89, P < 0.05).
Similar correlation was also reached when 5) actin carbonylation
(oxidation) and PLC-
activation (r = -0.95, P <
0.05) are utilized. Furthermore, 6) protection against
oxidant-induced actin disassembly (decreased F-actin polymer pool) and
PLC-
activation (r = 0.91, P < 0.05) and
7) the percentage of normal F-actin cytoarchitecture and PLC-
activation (r = 0.88, P < 0.05) provide other supporting
correlations. The high strength as well as consistency of these correlations
further indicates that PLC-
isoform activation is essential to
protection against iNOS upregulation and consequent oxidative stress to the
assembly of F-actin cytoskeleton and integrity of intestinal barrier function.
In this view, activation of PLC-
leads to the normalization of NO
levels and subsequently protects actin cytoskeleton and barrier integrity
against oxidative injury induced by iNOS.
Other proteins can also be involved in maintaining the integrity of
permeability barrier in the GI epithelium. These include a large heterogeneous
family of proteins such as microtubules (- and
-tubulin),
occludin, ZO proteins (e.g., ZO-1, ZO-2, ZO-3), claudins (e.g., isoforms I and
V), and myosin (e.g., type II) as well as others such as E-cadherin,
connexin43,
-catenin, and adherin
(5,
30,
33,
36,
42). Among these proteins, we
choose to study actin because previous studies showed the critical role of
actin cytoskeleton, especially the so-called "apical ring of
actin," in modulation of barrier paracellular permeability in epithelial
cells such as Caco-2 monolayers (e.g., Refs.
6,
10,
18,
36,
58). Moreover, we have
consistently shown that actin stability is key to EGF-mediated protection of
intestinal barrier permeability
(6,
10,
18).
The new findings of this report, using targeted molecular interventions,
are consistent not only with our own previous studies but also with the
findings of other investigators. It is known that PLC- profoundly
affects cellular functions in nonepithelial cells as well as epithelial cells
(23,
26,
32,
53,
62). For example, migration of
intestinal cells that is stimulated by growth factor requires PLC-
activity (22,
41,
46,
49). Other studies have
implicated PLC-
in the pathway for remodeling of the components of the
cytoskeleton such as microtubules, profilin, and gelsolin
(3,
12,
19,
26,
35). Furthermore, PLC-
is the only epithelial PLC isoform with SH2 and SH3 domains that can be
activated by growth factor ligands
(12,
23,
49,
52). The Z region of human
PLC-
1, namely PLCz, which covers both the SH2 and SH3 domains (amino
acids 517-901), is known to specifically inhibit PLC-
1 activation and
not other PLC isoforms in epithelial cells
(12,
22,
23,
32,
49). Indeed, the PLCz mutant
utilized is specific for inhibition of PLC-
1 because previous studies
have shown that PLC-
1 is the only epithelial PLC isoform that contains
SH2 and SH3 domains and that is activated by EGF
(12,
23,
32,
44,
49,
52,
57). In our studies
PLCz-dominant mutant expression prevented the activation of PLC-
while
at the same time abrogating EGF protection against oxidative stress. Our new
findings on the 145-kDa
-isoform of PLC, we believe, suggest a unique
pathophysiological role among PLC isoforms in intestinal cell monolayers,
namely protection against oxidative stress of RNM upregulation and of
cytoskeletal oxidation.
PLC- hydrolyzes phosphatidylinositol 4,5-bisphosphate
(PIP2) to produce not only inositol 1,4,5-trisphosphate
(IP3) but also diacyl glycerol (DAG) in epithelial cells
(6,
8,
23,
49,
51,
57). DAG is one of the
best-characterized products of PLC-
-mediated reactions and that is
known to lead to downstream activation of serine/thereonine protein kinase C
(PKC) (8,
11,
12,
28,
47). This is consistent with
the fact that we showed, using the same intestinal model, that PKC signaling
is also required for EGF-mediated protection against oxidant-induced GI
barrier and cytoskeletal disruption
(6,
8,
11,
12,
16). That PKC is downstream
from PLC-
in the cell's protective cascade is also indicated by our
previous finding that PKC activators (OAG or TPA) can maintain both
cytoskeletal integrity and intestinal barrier function even in the presence of
PLC inhibition. Indeed, PKC has also been shown to be downstream of PLC in
other systems as well (47,
49,
51,
61). Also, other studies have
shown that a naturally occurring intracellular activator of PKC, namely DAG
(OAG used in our previous studies is a synthetic version of this compound),
modulates intestinal monolayer permeability in Caco-2 cells
(11,
42). Overall, it appears that
growth factor-induced protection is mediated by PLC-
and then PKC.
It is noteworthy that we previously established (1, 4, 9, 10, 16) that 1) oxidant (e.g., H2O2, HOCl, ONOO-,or NO compounds)- or oxidative stressor (e.g., ethanol)-induced disruption of the cytoskeleton of intestinal cells and the consequent disruption of the permeability barrier of intestinal monolayers require rapid upregulation of an iNOS-driven reactive pathwaysincreased levels of RNMs such as NO and ONOO- (the latter from reaction of ·O2- + NO)which causes increased oxidative stress (DCF fluorescence) and damage through nitration and oxidation of cytoskeletal network. For example, H2O2 concentrations that caused actin damage and nitration and intestinal hyperpermeability also rapidly upregulated iNOS and increased RNMs and oxidative stress, including DCF fluorescence; several different exogenously added NO/RNM compounds [e.g., 3-morpholinosydnonimine (SIN-1), soluble N-ethylmaleimide-sensitive factor attachment protein (SNAP) + xanthine + xanthine oxidase, ONOO-] mimicked the effects of H2O2 (and potentiated the effects of low, nondamaging doses of H2O2); and either iNOS inhibitors (e.g., L-NIL) or several selective anti-oxidants (urate, L-cysteine, SOD) that scavenge NO/RNM or ·O2- anion prevented or substantially attenuated the injurious changes (e.g., DCF fluorescence) induced by H2O2 or RNM compounds (4, 9, 10).
2) Additionally, exposure of intestinal cells to oxidants, which models the oxidative stress that occurs during the active phase of IBD, can, surprisingly, further increase endogenous cellular synthesis of RNM compounds (such as NO and ONOO-) as well as ROS compounds (·O2-) (·O2- can, in turn, combine with the NO to generate other damaging reactive species such as ONOO-) (1, 4, 9, 10). Thus we demonstrated that oxidants (e.g., H2O2) per se appear to cause epithelial damage through the generation of reactive species (e.g., NO and RNMs), which then cause increases in oxidative stress (DCF) and nitration. 3) We also showed that growth factors (e.g., EGF) protect against oxidant-induced disruption through the underlying suppression of iNOS pathway upregulation and decreased levels of RNMs such as NO and ONOO-, which then reduce oxidative stress (16). 4) Finally, oxidants H2O2, HOCl, and ONOO- can all rapidly upregulate iNOS and NO in intestinal cells, because these cells appear to have a standing pool of iNOS mRNA and protein that is ready for immediate upregulation (4, 5, 10, 16). We found a similar mechanism of action for other oxidative stressors such as ethanol (1, 9).
Studies in endothelial cells as well as one in vivo study in rat gastric mucosal cells are consistent with our published findings of rapid iNOS changes (54, 56). In the study using isolated rat gastric mucosal cells (56), low basal levels of iNOS were noted in control (untreated) mucosa, whereas, after challenge with endotoxin, significant and rapid increases in iNOS activity were detected in the mucosal cells within 1 h, followed by peak levels at 2-4 h (almost identical to the rapid start and peak times for iNOS changes in our Caco-2 cells). Similarly, other studies showed detectable levels of iNOS activity as early as 60 min after H2O2 (0.1-1 mM) challenge (54). In yet another study in endothelial cells, a slight basal expression of iNOS protein (and iNOS mRNA as detected by RT-PCR) was shown in unstimulated cells (63).
Our findings are potentially relevant for developing new treatment
strategies for IBD. They suggest a novel antioxidative defensive mechanism
that might protect against oxidative stress of iNOS and NO upregulation and
prevent initiation, continuation, or manifestation of the IBD attack. The
potential therapeutic use of this antioxidative mechanism is consistent with
the current characterizations of the pathophysiology of the inflammation, in
general, and of IBD, in particular. iNOS is a key factor in the inflammatory
response triggered by an array of conditions promoting oxidative stress.
Specifically, upregulation of iNOS and the formation of RNMs (e.g., NO,
ONOO-) under conditions of oxidative stress appears to be essential
in the promotion of an inflammatory response in non-GI as well as GI models
(e.g., 34,
37,
40,
50,
55). Furthermore, we and
others have shown that upregulation of iNOS and RNMs is found in the
intestinal mucosa of patients with ulcerative colitis and Crohn's disease
(17,
37,
50,
55), in which high levels of
oxidants (e.g., H2O2) as well as loss of mucosal barrier
integrity have been reported
(17,
30,
31,
37,
39,
43). In these studies, tissue
nitration was associated with the inflamed human mucosa of IBD
(37,
50,
55) and was linked with the
upregulation of iNOS (37,
55). We further showed that
the amount of NO and RNM formation (i.e., nitration) correlated with the
degree of mucosal inflammation and disease severity score in IBD
(17,
37). Interestingly, some
nitrated tissue proteins have also been detected in vivo in non-GI models such
as the inflamed lung (34,
48). Thus iNOS and NO
upregulation appear to be critical to the pathogenesis of inflammatory
conditions such as IBD (17,
37). This upregulation is
though to be especially important in the transitions from the inactive to
active (flare up) phases of inflammation in IBD in which intestinal oxidants
and proinflammatory molecules periodically create a vicious cycle that can
lead to sustained iNOS upregulation, oxidative stress, hyperpermeability,
inflammation, and consequently to mucosal tissue damage. The protective
antioxidative effects of PLC-, such as those we have found in
intestinal cells, could play a pivotal role in preventing the establishment of
such a vicious inflammatory cycle.
A question that remains unanswered is how iNOS might be up- or
downregulated in intestinal cells. There are several mechanisms by which the
rapid iNOS modulation might occur. Regarding iNOS upregulation, we recently
showed that oxidant induces rapid iNOS upregulation (and its injurious
consequences such as nitration and DCF increases) through the underlying
activation of the (75-kDa)-isoform of PKC (PKC-
, a member of
the "novel" subfamily of PKC isoforms)
(4,
5,
10). Specifically, oxidant
H2O2 induces loss of epithelial barrier integrity by
nitrating and disassembling the cytoskeleton through the activation of
PKC-
isoform and consequent upregulation of iNOS-driven reactions
(including NO, ONOO-, DCF fluorescence, and cytoskeletal
nitration). Moreover, expression and activation of PKC-
is by itself
key for cellular injury induced by oxidative stress of iNOS-driven reactions,
indicating, once again, the critical role of this "injurious PKC
isoform" signal in iNOS upregulation under conditions of oxidative
stress.
Regarding iNOS downregulation, we recently showed that EGF can rapidly
suppress iNOS and NO upregulation through the downstream activation of the
(72-kDa)-isoform of PKC (PKC-
, a member of the
"atypical" subfamily of PKC isoforms)
(7,
16). Specifically, EGF
protects against oxidative disruption of the intestinal barrier integrity by
stabilizing the cytoskeleton, in large part, through the downstream activation
of PKC-
and consequent downregulation of iNOS. Consistent with these
findings, activation of PKC-
is by itself required for cellular
protection against oxidative stress of iNOS upregulation, further supporting a
key role for this "protective PKC isoform" in inhibition of iNOS
upregulation. These mechanisms appear to account for how iNOS is rapidly
modulated by oxidant and EGF in our intestinal model.
Nonetheless, there are other possible mechanisms for iNOS modulation as reported in non-GI models. For example, NOS contains consensus sequences for sites for protein phosphorylation (21, 25), especially tyrosine phosphorylation of calcium-independent NOS (i.e., iNOS), which has been shown in vitro and in endothelial cells following a variety of stimuli, and it was proposed that this mechanism could rapidly regulate the activity of NOS (21, 25). An alternative mechanism is the rapid assembly of the two known monomeric domains of iNOS into an active dimer, which is known to be required for NOS catalytic activity. Specifically, pools of inactive, monomeric iNOS would be available from a standing intracellular protein pool in unstimulated cells (like in Caco-2 cells). These monomers can be rapidly assembled by an appropriate stimulus into an active dimer (21). It remains to be seen whether there are additional molecular mechanisms underlying the iNOS modulation in GI epithelial cells.
In summary, our findings suggest that PLC- is responsible for a
substantial portion of the protection of the intestinal epithelium against
oxidant stress that is induced by iNOS upregulation and perhaps is key to
preventing amplification and perpetuation of an uncontrolled, oxidant-induced,
inflammatory cascade in IBD, one that can be ignited by free radicals and
other oxidants present in the GI tract. Our studies also suggest that
PLC-
is a potential therapeutic target for intervention against
inflammatory conditions in which oxidants are prevalent (e.g., IBD). For
instance, one therapeutic approach might be the exogenous delivery of a sense
vector for PLC-
isoform (targeted gene therapy) to the inflamed GI
mucosa in vivo. If this gene therapy approach is successful, one should be
able to protect and maintain epithelial integrity against oxidative stress
such as that occuring during incipient or rampant inflammation and
subsequently limit the initiation and progression of GI mucosal inflammation
and damage. PLC-
therapies might even synergize with the use of
currently employed antioxidants so that inflammatory processes are more
effectively attenuated through the manipulation of both the damaging and
protective intracellular pathways.
![]() |
DISCLOSURES |
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![]() |
ACKNOWLEDGMENTS |
---|
![]() |
FOOTNOTES |
---|
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.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
2. Banan A, Choudhary S, Zhang Y, Fields JZ, and Keshavarzian A. Oxidant-induced intestinal barrier disruption and its prevention by growth factors in a human colonic cell line: role of the microtubule cytoskeleton. Free Radic Biol Med 28: 727-738, 2000.[ISI][Medline]
3. Banan A,
Farhadi A, Fields JZ, Zhang L, and Keshavarzian A. The isoform of
phospholipase C (PLC-
) in EGF protection of intestinal F-actin
cytoskeletal assembly and barrier integrity (Abstract).
Gastroenterology 122,
Suppl 1: T863,
2002.
4. Banan A,
Farhadi A, Zhang L, Fields JZ, and Keshavarzian A. The -isoform of
protein kinase C causes inducible nitric oxide synthase and nitric oxide
up-regulation: key mechanism for oxidant-induced carbonylation, nitration, and
disassembly of cytoskeleton and disruption of the microtubule cytoskeleton and
hyperpermeability of barrier intestinal epithelia. J Pharmacol Exp
Ther 305:
482-494, 2003.
5. Banan A, Fields
JZ, Farhadi A, Talmadge DA, Zhang L, and Keshavarzian A. Activation of
-isoform of protein kinase C is required for oxidant-induced disruption
of both the microbule cytoskeleton and permeability barrier of intestinal
epithelia. J Pharmacol Exp Ther
303: 17-28,
2002.
6. Banan A, Fields
JZ, Talmage DA, Zhang L, Farhadi A, and Keshavarzian A. The 1
isoform of protein kinase C mediates the protective effects of EGF on the
dynamic assembly of the F-actin cytoskeleton and normalization of calcium
homeostasis in human colonic cells. J Pharmacol Exp
Ther 301:
852-866, 2002.
7. Banan A, Fields
JZ, Talmage DA, Zhang L, and Keshavarzian A. PKC- is required in EGF
protection of cytoskeleton and intestinal barrier integrity against oxidant
injury. Am J Physiol Gastrointest Liver Physiol
282: G794-G808,
2002.
8. Banan A, Fields
JZ, Talmage DA, Zhang Y, and Keshavarzian A. PKC-1 mediates EGF
protection of microtubules and barrier integrity of intestinal monolayers
against oxidants. Am J Physiol Gastrointest Liver
Physiol 281:
G833-G847, 2001.
9. Banan A, Fields
JZ, Zhang Y, and Keshavarzian A. Nitric oxide and its metabolites mediate
ethanol-induced microtubule disruption and intestinal barrier dysfunction.
J Pharmacol Exp Ther 294:
997-1008, 2000.
10. Banan A, Fields
JZ, Zhang Y, and Keshavarzian A. iNOS upregulation mediates
oxidant-induced disruption of F-actin and the permeability barrier of
intestinal monolayers. Am J Physiol Gastrointest Liver
Physiol 280:
G1234-G1246, 2001.
11. Banan A, Fields
JZ, Zhang Y, and Keshavarzian A. Key role of PKC and Ca2+ in
EGF protection of microtubules and intestinal barrier against oxidants.
Am J Physiol Gastrointest Liver Physiol
280: G828-G843,
2001.
12. Banan A, Fields
JZ, Zhang Y, and Keshavarzian A. Phospholipase C- inhibition
prevents EGF protection of intestinal cytoskeleton and barrier against
oxidants. Am J Physiol Gastrointest Liver Physiol
281: G412-G423,
2001.
13. Banan A,
McCormack SA, and Johnson LR. Polyamines are required for microtubule
formation during mucosal healing. Am J Physiol Gastrointest Liver
Physiol 274:
G879-G885, 1998.
14. Banan A, Smith
GS, Rickenberg C, Kokoska ER, and Miller TA. Protection against ethanol
injury by prostaglandin in a human intestinal cell line: role of microtubules.
Am J Physiol Gastrointest Liver Physiol
274: G111-G121,
1998.
15. Banan A, Wang
JY, McCormack SA, and Johnson LR. Relationship between polyamines, actin
distribution, and gastric healing in rats. Am J Physiol
Gastrointest Liver Physiol 271:
G893-G903, 1996.
16. Banan A, Zhang
L, Fields JZ, Talmage DA, and Keshavarzian A. PKC- prevents
oxidant-induced iNOS upregulation and protects the microtubules and intestinal
barrier integrity. Am J Physiol Gastrointest Liver
Physiol 283:
G909-G922, 2002.
17. Banan A, Zhang Y, Hutte R, and Keshavarzian A. Increased oxidation and nitration injury in intestinal mucosa of patients with inflammatory bowel disease (Abstract). Gastroenterology 118: 4266, 2000.
18. Banan A, Zhang
Y, Losurdo J, and Keshavarzian A. Disassembly of the F-actin in
oxidant-induced barrier dysfunction and its prevention by epidermal growth
factor and transforming growth factor- in a human colonic cell line.
Gut 46:
830-837, 2000.
19. Bar-Sagii D, Rotin D, Batzer A, Mandiyan V, and Schlessinger J. SH3 domains direct cellular localization of signaling molecules. Cell 74: 83-91, 1993.[ISI][Medline]
20. Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72: 224-254, 1976.
21. Bredt DS,
Ferris CD, and Synder SH. Nitric oxide synthase regulatory sites.
Phosphorylation by cyclic AMP-dependent protein kinase, protein kinase C, and
calcium/calmodulin protein kinase; identification of flavin and calmodulin
binding sites. J Biol Chem 267:
10976-10981, 1992.
22. Chen P, Gupta K, and Wells A. Cell movement elicited by epidermal growth factor receptor requires kinase and autophosphorylation but is separable from mitogenesis. J Cell Biol 124: 547-555, 1994.[Abstract]
23. Chen P, Xie H, and Wells A. Mitogenic signaling from the egf receptor is attenuated by a phospholipase C-gamma/protein kinase C feedback mechanism. Mol Biol Cell 7: 871-881, 1996.[Abstract]
25. Garcia-Cardena G, Fan R, Stern DF, Liu J, and Sessa WC.
Endothelial nitric oxide synthase is regulated by tyrosine phosphorylation and
interacts with caveolin-1. J Biol Chem
271: 27237-27240,
1996.
26. Goldschmidt-Clermont PH, Kim JW, Mcheskky LM, Rhee SG, and Pollart TD. Regulation of phospholipase C-gamma1 by profilin and tyrosine phosphorylation. Science 251: 1231-1233, 1996.
27. Harter JL. Critical values for Dunnett's new multiple range test. Biometrics 16: 671-685, 1960.[ISI]
28. Hartwig JH, Thelen M, Rosen A, Janmey PA, Nairn AC, and Aderem A. MARCKS is an actin filament crosslinking protein regulated by protein kinase C and calcium-calmodulin. Nature 356: 618-622, 1992.[ISI][Medline]
29. Hermiston ML and Gordon JI. Inflammatory bowel disease and adenomas in mice expressing a dominant negative N-cadherin. Science 270: 1203-1207, 1995.[Abstract]
30. Hollander D. The intestinal permeability barrier: a hypothesis as to its regulation and involvement in Crohn's disease. Scand J Gastroenterol 27: 721-726, 1992.[ISI][Medline]
31. Hollander D. Crohn's disease- a permeability disorder of the tight junction? Gut 26: 1621-1624, 1998.
32. Homma Y. and
Takenawa T. Inhibitory effect are src homology (SH) 2/SH3
fragments of phospholipase C-gamma on the catalytic activity of phospholipase
C isoforms. Identification of a novel phospholipase C inhibitor region.
J Biol Chem 267:
21844-21849, 1992.
33. Hurani MA, Noach AB, Blom-Roosemalen CM, DeBoer AG, Nagelkerke JF, and Breimer DD. Permeability enhancement in Caco-2 cell monolayers by sodium salicylate and sodium taurodihydrosulfate: assessment of effect-reversibility and imaging of transepithelial transport routes by laser confocal microscopy. J Pharmacol Exp Ther 267: 942-950, 1993.[Abstract]
34. Ischiropoulos H, Al-Mehdi HA, and Fisher AB. Reactive species
in ischemic rat lung injury: contribution of peroxynitrite. Am J
Physiol Lung Cell Mol Physiol 269:
L158-L164, 1995.
35. Janmey PA, Lamb
J, Allen PG, and Matsudaira PT. Phosphoinositide-binding peptides derived
from the sequences of gelsolin and villin. J Biol Chem
267: 11818-11823,
1992.
36. Kennedy M, Denenberg AG, Szabo C, and Salzman AL. Poly(ADP-ribose) synthetase activation mediates increased permeability induced by peroxynitrite in Caco-2BBe cells. Gastroenterology 114: 510-518, 1998.[ISI][Medline]
37. Keshavarzian A,
Banan A, Farhadi A, Kommandori S, Zhang L, and Fields JZ. Increases in
free radicals and cytoskeletal protein oxidation in the colon of patients with
inflammatory bowel disease. Gut
52: 720-728,
2003.
38. Keshavarzian A, Holmes EW, Patel M, Iber F, and Pethkar S. Leaky gut in alcoholic cirrhosis: a possible mechanism for alcohol induced liver damage. Am J Gastroenterol 94: 200-207, 1999.[ISI][Medline]
39. Keshavarzian A, Sedghi S, Kanofsky J, List T, Robinson C, Ibrahim C, and Winship D. Excessive production of reactive oxygen metabolites by inflamed colon: analysis by chemiluminescence probe. Gastroenterology 103: 177-185, 1992.[ISI][Medline]
40. Kimura H,
Hokari R, Miura S, Shigematsu T, Hirokawa M, Akiba Y, Kurose I, Higuchi H,
Fujimori H, Tsuzuki Y, Serizawa H, and Ishii H. Increased expression of an
inducible isoform of nitric oxide synthase and the formation of peroxynitrite
in colonic mucosa of patients with active ulcerative colitis.
Gut 42:
180-187, 1998.
41. Kinashi T,
Escobeda JA, Williams LT, Takatsu K, and Springer TA. Receptor tyrosine
kinase stimulates cell-matrix adhesion by phosphatidylinositol 3 kinase and
phospholipase C-1 pathways. Blood
86: 2086-2090,
1995.
42. Lindmark T,
Kimura Y, and Artursson P. Absorption enhancement through intracellular
regulation of tight junction permeability by medium-chain fatty acids in
Caco-2 cells. J Pharmacol Exp Ther
284: 362-369,
1998.
43. McKenizie SJ,
Baker MS, Buffington GD, and Doe WF. Evidence for oxidant-induced injury
to epithelial cells during inflammatory bowel disease. J Clin
Invest 98:
136-141, 1996.
44. Meisenhelder J, Suh PG, Rhee G, and Hunter T. Phospholipase C-gamma is a substrate for the PDGF and EGF receptor protein tyrosine kinase in vivo and in vitro. Cell 57: 1109-1122, 1989.[ISI][Medline]
45. Meunier VM, Bourrie Y, Berger Y, and Fabre G. The human intestinal epithelial cell line Caco-2: pharmacological and pharmacokinetics applications. Cell Biol Toxicol 11: 187-194, 1995.[ISI][Medline]
46. Nishimura R, Li W, and Kashishian A, Mondino A, Zhou M, Cooper J, and Schlessinger J. Two signaling molecules share a phosphotyrosine-containing binding site in PDGF receptor. Mol Cell Biol 13: 6889-6896, 1993.[Abstract]
47. Nishizuka Y. Intracellular signaling by hydrolysis of phospholipids and activation of protein kinase C. The role of protein kinase C in cell surface signal transduction and tumor promotion. Science 258: 607-614, 1992.[ISI][Medline]
48. Phelps DT,
Ferro TJ, Higgins PJ, Shankar R, Parker DM, and Johnson A. TNF-
induces peroxynitrite-mediated depletion of lung endothelial glutathione via
protein kinase C. Am J Physiol Lung Cell Mol Physiol
269: L551-L559,
1995.
49. Polk DB. Epidermal growth factor receptor-stimulated intestinal epithelial cell migration requires phospholipase C activity. Gastroenterology 114: 493-502, 1998.[ISI][Medline]
50. Ramchilewitz D, Stamler JS, Bachwich D, Karmeli F, Ackerman Z, and Podolsky DK. Enhanced colonic nitric oxide generation and nitric oxide synthase activity in ulcerative colitis and Crohn's disease. Gut 36: 718-723, 1995.[Abstract]
51. Reynolds NJ, Talwar HS, Baldassare JJ, Henderson PA, Elder JT, Voorhees JJ, and Fisher GJ. Differential induction of phosphotidylcholine hydrolysis, diacylglycerol formation and protein kinase C activation by EGF and TGF-alpha in normal human skin fibroblasts and keratinocytes. Biochem J 294: 535-544, 1993.[ISI][Medline]
52. Rhee SG and
Chio KD. Regulation of inositol phospholipidspecific phospholipase C
isozymes. J Biol Chem 267:
12393-12396, 1992.
53. Rotin D, Margolis B, Mohammadi M, Daly RJ, Daum G, Li N, Fischer EH, Burgess WH, Ulrich A, and Schlessinger J. SH2 domains prevent tyrosine dephosphorylation of EGF-R: identification of Tyr992 as the high affinity binding site for SH2 domains of PLC-gamma. EMBO J 11: 559-567, 1992.[Abstract]
54. Shimizu S, Nomoto M, Naito S, Yamamoto T, and Momose K. Stimulation of nitric oxide synthase during oxidative endothelial cell injury. Biochem Pharmacol 55: 77-83, 1998.[ISI][Medline]
55. Singer II, Kawka DW, Scott S, Weidner JR, Mumford RA, Riehl TE, and Stenson WF. Expression of inducible nitric oxide synthase and nitrotyrosine in colonic epithelium in inflammatory bowel disease. Gastroenterology 111: 871-885, 1996.[ISI][Medline]
56. Tripp MA and Tepperman BL. Role of calcium in nitric oxide-mediated injury to gastric mucosal cells. Gastroenterology 111: 65-72, 1996.[ISI][Medline]
57. Turner T, Epps-Fung MV, Kassis J, and Wells A. Molecular inhibitor of phospholipase C-gamma signaling abrogates DU-145 prostate tumor cell invasion. Clin Cancer Res 3: 2275-2282, 1997.[Abstract]
58. Unno N, Menconi
MJ, Smith M, and Fink MP. Hyperperme-ability of intestinal epithelial
monolayers induced by NO: effect of low extracellular pH. Am J
Physiol Gastrointest Liver Physiol 272:
G923-G934, 1997.
59. Wall RL, Albrecht T, Thompson WC, James O, and Carney DH. Thrombin and phorbol myristate acetate stimulate cytoskeletal polymerization in quiescent cells: a potential link to mitogenesis. Cell Motil Cytoskeleton 23: 265-278, 1992.[ISI][Medline]
60. Yamada T, Sarto RB, Marshall S, Special RD, and Grisham MB. Mucosal injury and inflammation in a model of chronic granulomatous colitis in rats. Gastroenterology 104: 759-771, 1993.[ISI][Medline]
61. Yang LJ, Rhees
SG, and Williamson JR. Growth factor-induced activation and translocation
of phospholipase C-1 to the cytoskeleton in rat hepatocytes.
J Biol Chem 269:
7156-7162, 1994.
62. Yeo E-J,
Kalzlasukas A, and Exton JH. Activation of PLC-gamma is necessary for
stimulation of Phosholipase by PDGF. J Biol Chem
269: 27823-27826,
1994.
63. Zadeh MS, Kolb JP, Geromin D, D'Anna R, Boulmerka A, Marconi A, Dugas B, Marsac C, and D'Alessio P. Regulation of ICAM-1/CD54 expression on human endothelial cells by hydrogen peroxide involves inducible NO synthase. J Leukoc Biol 67: 327-334, 2000.[Abstract]
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