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PROTEIN AND VESICLE TRAFFICKING, CYTOSKELETON
in EGF protection of epithelial barrier against iNOS upregulation and F-actin nitration and disassembly
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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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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appears to be essential for monolayer integrity. We thus hypothesized that
PLC- activation is essential in EGF protection against iNOS
upregulation and the consequent cytoskeletal oxidation and disarray and
monolayer disruption. Intestinal cells were transfected to stably overexpress
PLC- or to inhibit its activation and were then pretreated with EGF
± oxidant (H2O2). Wild-type (WT) intestinal cells
were treated similarly. Relative to WT monolayers exposed to oxidant,
pretreatment with EGF protected monolayers by: increasing native PLC-
activity; decreasing six iNOS-related variables (iNOS activity/protein, NO
levels, oxidative stress, actin oxidation/nitration); increasing stable
F-actin; maintaining actin stability; and enhancing barrier integrity.
Relative to WT cells exposed to oxidant, transfected monolayers overexpressing
PLC- (+2.3-fold) were protected, as indicated by decreases in all
measures of iNOS-driven pathway and enhanced actin and barrier integrity.
Overexpression-induced inhibition of iNOS was potentiated by low doses of EGF.
Stable inhibition of PLC- prevented all measures of EGF protection
against iNOS upregulation. We conclude that 1) EGF protects against
oxidative stress disruption of intestinal barrier by stabilizing F-Actin,
largely through the activation of PLC- and downregulation of iNOS
pathway; 2) activation of PLC- is by itself essential for
cellular protection against oxidative stress of iNOS; and 3) the
ability to suppress iNOS-driven reactions and cytoskeletal oxidation and
disassembly is a novel mechanism not previously attributed to the PLC family
of isoforms. 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 mechanisms—prevention of the oxidative
stress of iNOS upregulation and of cytoskeletal protein nitration and
oxidation—by 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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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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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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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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sense stably overexpress the (145
kDa)-isoform of phospholipase C (
2.3-fold compared with wild-type cells)
and that this overexpression protects monolayer barrier integrity against
exposure to oxidant challenge. Because PLC- protects against
oxidant-induced disruption, we surmised that this protection may be due to the
inhibition of oxidant-activated pathways such as the one triggered by reactive
metabolites. In the current investigation, utilizing both pharmacological and
molecular biological interventions, we studied the underlying mechanism by
which PLC- protects.
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.
|
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.
|
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).
|
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.
|
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).
|
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.
|
|
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.
|
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-.
|
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 |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
-isoform of PLC is
required for EGF-mediated protection against oxidant-induced iNOS upregulation
and the consequent oxidative stress injury to the integrity of F-actin
cytoskeleton and the intestinal epithelial barrier. A second conclusion is
that PLC- by itself is key in cellular protection against stress of
iNOS-driven reactions. The underlying mechanism for this protective effect of
PLC- isoform appears to be the suppression of both nitration and
oxidation stress injury to the 43-kDa subunit components of the F-actin
network and consequent stabilization of actin assembly and
cytoarchitecture.
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 grow
