Section of Gastroenterology and Nutrition, Department of Internal Medicine, Department of Pharmacology, and Department of Molecular Physiology, Rush University Medical Center, Chicago, Illinois 60612
Submitted 18 March 2004 ; accepted in final form 31 May 2004
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ABSTRACT |
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tubulin cytoskeleton; microtubules; oxidation/nitration; inducible nitric oxide synthase/peroxynitrite; inflammatory bowel disease; Caco-2 cells; gut barrier; nuclear factor-B/I
B
Although the pathophysiology of mucosal barrier disruption in IBD is not fully characterized, it was recently discovered that a leaky and hyperpermeable gut barrier can cause intestinal inflammation and that promoting a normal mucosal barrier function is essential for intestinal health. In animal models, intestinal barrier hyperpermeability induced by the injection of proinflammatory substances (e.g., peptidoglycan-polysaccharide, PG-PS) into the mucosa elicits oxidative and inflammatory conditions similar to those of IBD (54). Moreover, transgenic mice with a leaky gut exhibit symptoms of intestinal inflammation (27). It is also known that chronic gut inflammation in IBD is associated not only with high levels of oxidants (e.g., H2O2) but also with increased cytoskeletal instability, which together appear to be key contributors to mucosal injury (35, 3739). Indeed, upregulation of oxidants and the consequent oxidative stress and cytoskeletal disruption have been implicated in mucosal inflammation and damage in IBD (2, 8, 11, 12, 17, 18, 3539). Accordingly, understanding how gut barrier and cytoskeletal function are destabilized under oxidative, proinflammatory conditions is of substantial clinical and biological value.
During the past decade, we have been investigating injurious mechanisms such as oxidant-induced mucosal damage and barrier disruption not only to better understand endogenous defensive mechanisms (e.g., against oxidative disruption by H2O2) but also to devise a rational basis for the development of potentially more effective treatment regimes for IBD. Using monolayers of intestinal Caco-2 cells as a well-established model of gut cytoskeleton and barrier function, we previously reported several original findings that cytoskeletal oxidation and disassembly are required in oxidant-induced loss of barrier function (2, 4, 5, 7, 9) and that oxidants (H2O2, HOCl, and others) induce oxidative stress damage, in large part through the upregulation of inducible nitric oxide synthase (iNOS)-driven reactions (e.g., reactive nitrogen metabolites such as NO) (8, 14).
It is noteworthy that activation of NF-B (a key proinflammatory factor) is essential to the promotion of an oxidative and inflammatory response in gut disorders such as in IBD (ulcerative colitis and Crohn's disease) (11, 12, 43, 46, 48). Once activated, NF-
B appears to regulate several important processes involved in inflammatory response, including loss of barrier function (3, 15, 16). Indeed, we recently reported novel findings on the importance of NF-
B-dependent mechanisms in oxidant-induced barrier disruption (3, 11). We showed, using intestinal Caco-2 cells, that oxidants induce the nuclear translocation and activation of NF-
B and lead to disruption of monolayer barrier (permeability) function (3). Despite the importance of NF-
B to intestinal barrier integrity, the molecular mechanisms underlying NF-
B-mediated, oxidant-induced disruption of monolayer barrier and cytoskeletal function are poorly understood.
In light of the aforementioned considerations, we tested the hypothesis that not only is NF-B activation critical to oxidant-induced iNOS and NO upregulation but also that it is key to the injurious consequences of this upregulation, namely, microtubule cytoskeletal oxidation, nitration, disassembly and disarray, and monolayer dysfunction.
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MATERIALS AND METHODS |
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Plasmids and transfection.
The dominant negative mutant (superrepressor) of IB
was used as previously described (3, 42). This mutant contains double point mutations substituting key serines 32 and 36 with alanine residues, which stabilizes I
B
(and prevents the activation of NF-
B). The construct was cloned into a cytomegalovirus (CMV) expression vector to overexpress the dominant negative mutant. An expression vector containing CMV plasmid alone served as a control. Stable transfectants were determined by immunoblot analysis of cell fractions.
For transfection, cultures of cells grown to 5060% confluency were stably transfected with varying amounts (14 µg) of expression plasmid encoding a dominant negative mutant of IB
by using Lipofectin reagent (25 µl Lipofectamine/25-cm2 flask; GIBCO BRL) as we recently described (3, 42). Control conditions included vector alone. I
B
protein expression and lack of its degradation (i.e., stability) were verified by Western blot analysis of cellular cytosolic fractions. Multiple clones were subsequently plated and allowed to form confluent monolayers and were then used for experiments.
Experimental design.
In the first series of experiments, postconfluence monolayers of wild-type cells were incubated with oxidant (H2O2, 00.5 mM) or vehicle (isotonic saline) for 30 min, and then outcomes were assessed over time (e.g., from 0 to 8 h). Outcomes measured are described below. As we previously showed, H2O2 at 0.5 mM disrupts microtubules and barrier integrity and upregulates iNOS in wild-type intestinal cells (2, 8, 14). These experiments were then repeated using monolayers composed of clones of transfected cells (i.e., IB
mutants). In all experiments, we assessed 1) microtubule cytoskeletal stability (cytoarchitecture, assembly), 2) polymerized (S2) and monomeric (S1) tubulin pools, 3) iNOS activity and protein, 4) NO levels, 5) reactive nitrogen metabolite (RNM) levels (e.g., ONOO), 6) oxidative stress [dichlorofluorescein (DCF) fluorescence], 7) tubulin nitration (nitrotyrosine formation), 8) tubulin oxidation (carbonylation), 9) I
B
distribution (cytosolic expression and instability), and 10) NF-
B p65 subunit activity (nuclear translocation and activity) and NF-
B p50 subunit activity (nuclear translocation and activity).
In the second series of experiments, we investigated the potential importance of the NF-B in oxidant-induced oxidative stress upregulation (e.g., NO overproduction) as well as cytoskeletal oxidation/nitration injury by using several pharmacological inhibitors (all 30-min preincubations). Monolayers of wild-type cells were preincubated with four different NF-
B/I
B
inhibitors and then incubated with or without oxidant or vehicle. These inhibitors included known inhibitors of the activation of NF-
B (1, 33), alone or in combination with H2O2: curcumin (20 µM), 1-pyrrolidinedithiocarbamate (PDTC; 20 µM), carbobenzyloxy-leu-leu-leucinol (MG-132; 10 µM), and lactacystin (10 µM). Controls were treated with vehicle. We confirmed that these doses of inhibitors were not toxic to cells. Outcomes measured were as described for the first series of experiments.
In a third series of experiments, we incubated monolayers of IB
dominant negative transfected cells (I
B
mutants) with oxidant or vehicle. In all experiments, I
B
levels and NF-
B activity were determined. Other outcomes measured were as described for the first series of experiments. In corollary experiments, we investigated the effects of NF-
B activation or inactivation on the state of 1) tubulin oxidation and tubulin nitration, 2) iNOS upregulation and NO overproduction, 3) tubulin assembly and disassembly, and 4) stability of the cytoarchitecture of the microtubule cytoskeleton. Monomeric (S1) and polymerized (S2) fractions of tubulin (the structural protein subunit of microtubules) were isolated and then analyzed for outcomes (e.g., oxidation and nitration by immunoblot analysis) (2, 14). Microtubule integrity was assessed by 1) immunofluorescent labeling and fluorescence microscopy to assess cells with normal microtubules, 2) detailed morphological analysis of cytoskeleton by ultrahigh-resolution laser scanning confocal microscopy, 3) immunoblot analyses of monomeric and polymerized tubulin pools, and 4) immunoblot analyses of oxidation and nitration of tubulin.
Assay of NOS activity. Wild-type and transfected cells grown to confluence were removed by scraping, 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, Arlington Heights, IL) to L-[3H]citrulline was measured in the cell homogenates by scintillation counting. Experiments in the presence of NADPH, without Ca2+ and with 5 mM EGTA, determined Ca2+-independent NOS (iNOS) activity (1, 6, 8).
Western blot analysis of iNOS level. After treatments, the cells were washed once with cold PBS, scraped into 1 ml of cold PBS, and harvested in a standard antiprotease cocktail. For immunoblot analysis, samples (20 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, TBS). The immunoblotted proteins were incubated for 2 h in Tween 20-TBS (TBST) and 1% BSA with the primary antibody (mouse monoclonal anti-human iNOS, at 1:3,000 dilution; Santa Cruz Biotechnology, Santa Cruz, CA). A horseradish peroxidase (HRP)-conjugated goat anti-mouse antibody (Molecular Probes, Eugene, OR) was used as a secondary antibody, at 1:3,000 dilution. Membranes were visualized by enhanced chemiluminescence (ECL) and then autoradiographed and processed for densitometry (8, 14).
Chemiluminescence analysis of NO. NO production was assessed using a unique chemiluminescence procedure (8, 14). Briefly, cells were homogenized, and endogenous nitrate (NO3) and nitrite (NO2), the metabolic degradation products of NO, were then reduced to NO using vanadium (III) (Sigma, St. Louis, MO) and HCl at 90°C before NO concentration was measured using a Sievers NOA 280 analyzer (Sievers Instruments, Boulder, CO). NO was expressed as a micromolar concentration and was calculated by comparison to the chemiluminescence of a standard solution of NaNO2. The absolute NO values were reported as 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 a fluorescent dye, DCF (1, 6, 14). Briefly, monolayers grown in 96-well plates were preincubated with the membrane-permeable DCFD (10 µg/ml for 30 min) before the subsequent treatments were carried out. 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. As we previously showed (6, 8), the dependence of the DCF assay on oxidative stress of reactive oxygen metabolite (ROM) production (e.g., ·O2 generation) was demonstrated by adding the active superoxide radical scavenger superoxide dismutase (SOD) or, as a control condition, heat-inactivated SOD (iSOD). Similarly, we previously showed the usefulness of this assay for detecting other forms of oxidative stress such as 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., L-N6-(1-iminoethyl)lysine]. These studies thus indicated that oxidative stress seen under conditions of H2O2 challenge is most likely due to the generation of new ROM species (e.g., ·O2) as well as RNM species (e.g., NO).
Analysis of NF-B activation.
NF-
B (p65 and p50 subunit) activation was assessed by a unique ELISA-based procedure as we described previously (3, 15). Monolayers of wild-type and transfected cells grown in 25-cm2 flasks were processed for the isolation of cytosolic and nuclear fractions. Cell fractions were added to a 96-well plate to which oligonucleotides containing a consensus-binding site for NF-
B had been immobilized (Trans-Am; Active Motif, Carlsbad, CA). Specifically, the NF-
B activity test is based on a validated ELISA principle whereby NF-
B is captured by a double-stranded oligonucleotide probe containing the consensus-binding sequence for either NF-
B p65 or p50 subunit (45). Consequently, only the activated NF-
B is captured by the probe bound in microwell plates. The binding of NF-
B to its consensus sequence was then detected using a primary anti-NF-
B (p65 or p50) antibody (Santa Cruz Biotechnology), followed by a secondary antibody conjugated to HRP. The results were quantitated by a chromogenic reaction (45), which was then read for absorbance at 450 nm by using a multiplate reader (FL 600; BIO-TEK Instruments).
Western blot analysis of changes in NF-B subunit levels and nuclear translocation.
Cellular nuclear and cytosolic extracts from wild-type and transfected cells were prepared as described above. NF-
B nuclear distribution (translocation) was determined by comparing the levels of NF-
B protein (e.g., p65 subunit) expression in the cytosolic and nuclear extracts with the use of anti-p65 (or anti-p50) antibody on a nondenaturing gel (6%) (3, 15). Samples (20 µg protein/lane) were placed in a standard sample buffer, boiled, and then subjected to PAGE. Proteins were visualized by ECL (Amersham) and autoradiography. For comparison of different blots, standard (positive) loading controls (20 µg of HeLa cell extracts/lane) for NF-
B were included concurrently with each run. In addition, after the blots were stripped, actin (
43 kDa) immunoblotting was performed as an internal control for equal loading.
Electrophoretic mobility shift assays.
LightShift chemiluminescent EMSA kits (Pierce, Rockford, IL), which use a nonisotopic method to detect specific DNA-protein interactions, were utilized (51). Biotin-end-labeled DNA is incubated with nuclear extracts. This reaction is then subjected to gel electrophoresis on a nondenaturing (6%) polyacrylamide gel and transferred to a nylon membrane. The biotin-end-labeled DNA is detected using the streptavidin-HRP conjugate and LightShift chemiluminescent substrate. To this end, thawed nuclear extracts (610 µg) were prepared as described above and incubated with 1 ng of biotin-end-labeled NF-B-specific probes (provided in the kit) in a total volume of 25 µl in the presence of 10 mM Tris·HCl (pH 7.5), 80 mM NaCl, 1 mM EGTA, 1 mM DTT, 10% glycerol, and 1 µl of poly(dI-dC). DNA-nuclear protein complexes were separated by PAGE (6%, nondenaturing gels) and visualized by autoradiography. The specificity of binding interactions was further assessed by competition with an excess of unlabeled double-stranded oligonucleotide of the same identity.
Western blot analysis of IB
degradation and expression levels.
The level of I
B
expression in the cytosolic extracts as well as its degradation and/or instability (i.e., disappearance from the cytosolic fractions) was confirmed by anti-I
B
antibody (Santa Cruz Biotechnology) according to a Western blot protocol (10% gel) (3, 15). Briefly, samples (20 µg protein/lane) were added to a standard SDS buffer, boiled, and then separated on SDS-PAGE. As for NF-
B, proteins were visualized by ECL and subsequently autoradiographed and processed for densitometry. Standard (positive) loading controls (20 µg of HeLa cell extracts/lane) for I
B
were included in each run.
Immunofluorescent staining and high-resolution laser scanning confocal microscopy of microtubules.
Cells from monolayers were fixed in cytoskeletal stabilization buffer and then postfixed in 95% ethanol at 20°C as we previously described (117). Cells were then processed for incubation with a primary antibody, monoclonal mouse anti--tubulin (human reactive; Sigma) at 1:200 dilution for 1 h at 37°C and then incubated with a secondary antibody (FITC-conjugated goat anti-mouse; Sigma) at 1:50 dilution for 1 h at room temperature. Slides were washed three times in Dulbecco's PBS and subsequently mounted in aquamount. After staining, cells were observed with an argon laser (
= 488 nm) using a x63 oil-immersion Plan Apochromatic objective, NA 1.4 (Carl Zeiss, Oberkochen, Germany). Cells from desired areas of monolayers were processed using the image processing software on a Zeiss ultrahigh-resolution laser scanning confocal microscope. The cytoskeletal elements were examined in a blinded fashion for their overall morphology, orientation, and disruption as we described (117). The identity of the treatment groups for all slides was decoded only after examination was complete.
Microtubule (tubulin) fractionation and quantitative immunoblot analysis of tubulin assembly. Polymerized (S2) and monomeric (S1) fractions of tubulin were isolated using a series of extraction and ultracentrifugation steps as we described previously (2, 13). Briefly, cells were gently scraped and pelleted with centrifugation at low speed (700 rpm, 7 min, 4°C) and resuspended in microtubule stabilization-extraction buffer (0.1 M PIPES, pH 6.9, 30% glycerol, 5% DMSO, 1 mM MgSO4, 10 µg/ml antiprotease cocktail, 1 mM EGTA, and 1% Triton X-100) at room temperature for 20 min. Tubulin fractions were separated after a series of centrifugation and extraction steps. Specifically, cell lysates were centrifuged at 105,000 g for 45 min at 4°C, and the supernatant containing the soluble monomeric pool of tubulin (S1 fraction) was gently removed. The remaining pellet was then resuspended in 0.3 ml of Ca2+-containing depolymerization buffer (0.1 M PIPES, pH 6.9, 1 mM MgSO4, 10 µg/ml antiprotease cocktail, and 10 mM CaCl2) and incubated on ice for 60 min. Subsequently, samples were centrifuged at 48,000 g for 15 min at 4°C, and the supernatant (S2 fraction or cold/Ca2+-soluble fraction) was removed. To ensure complete removal of the S2 fraction, we treated the remaining pellet with the Ca2+-containing depolymerization buffer twice more by resuspension and centrifugation. The "microtubules" were recovered by separate incubation (at 37°C for 30 min) of the S1 and S2 fractions with the stabilizing agents taxol (10 µM) and GTP (1 mM) in microtubule stabilization buffer (MSB: 0.1 M PIPES, pH 6.9, 30% glycerol, 5% DMSO, 10 µg/ml antiprotease cocktail, 1 mM EGTA, 1 mM MgCl2, and 1 mM GTP) to promote polymerization of tubulin. Tubulin was then recovered by centrifugation and resuspended in MSB. Fractionated S1 and S2 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 for 5 min, and then subjected to PAGE on 7.5% gels. Procedures for Western blotting were performed as previously described (2, 13). The immunolabeled tubulin was then processed for autoradiography. Standard (purified) tubulin loading controls (5 µg/lane) were included concurrently with each run.
Immunoblotting determination of protein tubulin oxidation and tubulin nitration. Oxidation and nitration of the microtubule (tubulin) cytoskeleton were assessed, respectively, by measuring protein carbonyl and nitrotyrosine formation using a method we developed (2, 14). To prevent unwanted oxidation of tubulin samples, all buffers contained 0.5 mM DTT and 20 mM 4,5-dihydroxy-1,3-benzene sulfonic acid (Sigma). To determine the carbonyl content, we blotted samples to a polyvinylidene difluoride 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 containing 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 a HRP-conjugated secondary antibody (1:4,000 dilution, 1 h; Molecular Probes). To determine nitrotyrosine content, after the blocking step 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 (Upstate Biotechnology, Lake Placid, NY), followed by the HRP-conjugated secondary antibody (as above). Wash steps and film exposure were as described in a standard Western protocol (2, 14). The relative levels of oxidized or nitrated tubulin were then quantified by measuring, with a laser densitometer, the optical density (OD) of the bands corresponding to anti-DNPH or anti-nitrotyrosine immunoreactivity. Immunoreactivity was expressed as the percentage of carbonyl or nitrotyrosine formation (OD) in the treatment group compared with the maximally oxidized or nitrated tubulin standards, which also served as loading controls. Oxidized or nitrated tubulin standards (5 µg/lane) were run concurrently with corresponding treatment groups. To further verify equal loading of lanes (22), blots were routinely stained with 0.1% india ink in TBST buffer. Oxidized tubulin standard was prepared by utilizing purified tubulin (ICN, Costa Mesa, CA) that was subsequently carbonylated by exposure to 0.5 mM H2O2 and 1 mM FeSO4 (30 min at room temperature). Nitrated tubulin standard was prepared by reacting purified tubulin with 0.1 mM ONOO (30 min at 37°C). These oxidized standards were then precipitated with trichloroacetic acid (TCA) followed by the decanting of supernatant and were washed (3 times) with TCA to remove excess oxidizing agents.
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 (26). 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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Pharmacological inhibition of NF-B protects against oxidative damage to the cytoskeleton: Prevention of both tubulin nitration and oxidation.
Because NF-
B is key in oxidant-induced barrier disruption (3), we surmised that this disruption may be due to the activation of oxidant-activated pathways such as the one triggered by reactive metabolites. Using intestinal cells, we initially measured the "footprints" of RNM formation, namely, nitrotyrosine moieties under conditions of oxidant challenge with or without several pharmacological inhibitors of NF-
B. We also measured oxidation footprints by assessing carbonylation levels. These were performed by sequentially fractionating and purifying the 50-kDa tubulin molecule from cell monolayers and subsequently immunoblotting these fractions to assess the oxidation state of microtubule (tubulin-based) cytoskeleton. Oxidant (H2O2, 0.5 mM) alone resulted in substantial levels of nitration and oxidation of the tubulin cytoskeleton (Fig. 1A). In contrast, pharmacological inhibitors of NF-
B afforded protection against oxidant-induced tubulin nitration and tubulin carbonylation as demonstrated by decreased oxidation levels, which were comparable to those in control (vehicle treated) cells. For example, only cells pretreated with inhibitors of NF-
B activation (e.g., curcumin or PDTC) were protected against oxidant-induced nitration and oxidation injuries (95% less oxidation). Similarly, preincubation with inhibitors of the NF-
B modulator I
B
(e.g., lactacystin or MG-132) also led to protection against oxidation (by
9093%). In the absence of oxidants, these inhibitors by themselves did not affect tubulin (not shown).
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NF-B-induced oxidative injury involves upregulation of iNOS-driven processes: Increases in iNOS, NO, RNM (ONOO), and oxidative stress.
We next probed potential mechanisms through which NF-
B promotes oxidative stress to the cytoskeleton. Because oxidants such as H2O2 upregulate iNOS (8, 14), we hypothesized that activation of iNOS-driven pathways might be a novel mechanism for NF-
B-induced oxidative injury to the tubulin-based cytoskeleton.
To this end, the same four pharmacological inhibitors of NF-B also cause a substantial reduction in Ca2+-independent iNOS activity induced by oxidant (0.5 mM) challenge (
9698% lower iNOS activity) (Fig. 2A). This is comparable to that of the control, displaying only low (basal) iNOS activity. In contrast, oxidant by itself causes large increases in iNOS activity. Similarly, NO levels (a product of the iNOS-catalyzed reaction) in monolayers exposed to H2O2 were also increased (not shown). Inhibitors of NF-
B activity markedly prevented oxidant-induced NO overproduction. Indeed, as for both tubulin oxidation/nitration and iNOS upregulation, NO overproduction was suppressed by these inhibitors.
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In parallel with the prevention of oxidant-induced effects, NF-B inhibitors suppressed oxidative stress as determined by reduction in the fluorescence of DCF (Fig. 3). In cells where H2O2 substantially increased DCF fluorescence, oxidative stress was prevented only by inhibitors of NF-
B activity. In the absence of oxidant (i.e., vehicle), we observed significantly lower and basal levels of cellular oxidative stress [presumably 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, 6, 8, 14)]. Inhibitors by themselves did not affect basal oxidative stress (not shown).
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Intracellular distribution and activation of NF-B subunit proteins (p50, p65) in intestinal cells parallel several different indexes of iNOS/NO and oxidative stress in monolayers.
Assessment of alterations in NF-
B subunit distribution in the nuclear fractions (Figs. 5) shows that NF-
B inhibitors suppressed oxidant-induced nuclear distribution of the NF-
B subunits p50 (Fig. 5A) and p65 (Fig. 5B) as shown by the corresponding reductions in band densities, which are comparable to those of controls. On the other hand, exposure to oxidant led to increased nuclear distribution of NF-
B subunits, paralleling findings on oxidant-induced activation of oxidative stress pathways.
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Using data across all experimental conditions, we found positive correlations (e.g., r = 0.92, P < 0.05) between NF-B activity (activity assay or OD from the nuclear fractions) and iNOS upregulation, further indicating that activation of NF-
B is key in oxidant-induced iNOS upregulation and its deleterious consequences (e.g., nitration). Other correlations were seen when either NO overproduction or oxidative stress (DCF fluorescence) was correlated with the NF-
B activity (r = 0.91 or 0.88, respectively, P < 0.05 for each). When two other markers of oxidative stress, tubulin carbonylation and tubulin nitration, were correlated with NF-
B, additional correlations were observed (r = 0.96 and 0.97, respectively, P < 0.05 for each), further indicating that activation of NF-
B is key in NO overproduction and cytoskeletal nitration through upregulation of iNOS enzyme. Similarly, when markers of stability such as either microtubule integrity or tubulin assembly (e.g., S2 pool) were correlated with the NF-
B, still other correlations were seen (r = 0.87 or 0.90, respectively, P < 0.05 for each).
Inhibition of NF-B activity by a dominant negative mutant (superrepressor) for I
B
after transfection of intestinal cells: Prevention of oxidative stress of iNOS and NO upregulation and cytoskeletal nitration/oxidation in I
B
mutant clones.
The aforementioned findings collectively indicate that NF-
B activation could play a novel role in promotion of the oxidative stress of NO upregulation and cytoskeletal nitration/oxidation (and disarray). To independently demonstrate that NF-
B contributes to oxidative injury to cytoskeleton and to further probe the underlying events, we used stable I
B
mutant (dominant negative) transfected clones of intestinal cells that we recently developed (in which I
B
is stabilized against degradation) (3). Because oxidants not only reduce the stability of I
B
(a 37-kDa endogenous modulator of NF-
B) but also increase monolayer dysfunction in intestinal Caco-2 cells, we surmised that reduced stability of I
B
could be a unique mechanism underlying NF-
B-induced upregulation of oxidative stress and cytoskeletal oxidation.
Initially, intestinal Caco-2 cells were stably transfected with plasmid DNA encoding both neomycin resistance (for selection) and varying amounts (1, 2, 3, or 4 µg) of the IB
mutant (3). Using this dominant negative approach, we were able to suppress NF-
B activation in Caco-2 clones as determined by an EMSA (a nonradioactive chemiluminescent gel shift), using biotin-end-labeled NF-
B-specific probes (Fig. 6, 3-µg mutant). Transfection of control vector alone (CMV plasmid), as might be expected, did not inhibit NF-
B activity. Indeed, NF-
B activity in the control vector clone, which was challenged with oxidant, was similar to that of wild-type cells under similar conditions. Utilizing a second technique involving a sensitive ELISA, we confirmed that in the I
B
mutant cells, oxidant could no longer increase the NF-
B activity in nuclear fractions of Caco-2 cells (Table 1). We observed similar effects by I
B
mutant transfection in a normal intestinal cell line, IEC-6 (Table 2).
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Furthermore, immunoblot analysis of the oxidative state of tubulin (Fig. 7A and Table 2) from the same IB
mutant clone shows that suppression of NF-
B activity prevented (protected against) oxidant-induced tubulin oxidation. In parallel, immunoblot analysis of the state of tubulin pool assembly from these I
B
mutant cells demonstrates (Fig. 7B, Caco-2 cells) that inhibition of NF-
B activity attenuated tubulin depolymerization induced by oxidant challenge. In these mutant clones, oxidant cannot cause tubulin disassembly as shown by enhanced polymerized (stable) tubulin to near control levels.
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Finally, analysis of oxidative stress (Fig. 8,DCF fluorescence in Caco-2 cells) from these IB
mutant clones additionally demonstrates that inhibition of NF-
B activity substantially attenuated oxidants' upregulation of oxidative stress. As for iNOS and NO upregulation, a large percentage (
9598%) of oxidant-induced effects on DCF fluorescence upregulation appears to be NF-
B dependent.
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DISCUSSION |
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These conclusions are based on several independent lines of evidence. First, activation of NF-B induces an oxidant-like injury, including oxidant-induced iNOS upregulation. NF-
B activation evokes an oxidative cascade of alterations, including hyperactivation of iNOS enzyme, overgeneration of NO, increases in RNM, and promotion of oxidative stress (DCF fluorescence). This oxidative injury appears to require activation of the NF-
B. Second, NF-
B induces footprints of oxidative damage to tubulin (50 kDa) protein subunits of the microtubule network. Activation of NF-
B (e.g., p65 subunit) promotes the nitration (nitrotyrosination) of tubulin and increases the oxidation (carbonylation) of tubulin (and as we previously showed, destabilizes the appearance of the microtubule cytoskeleton; Ref. 3). Third, dominant negative mutation of I
B
, which causes stabilization of I
B
and inactivation of NF-
B, substantially interfered with oxidant-induced increases in iNOS upregulation (by
96%) and with nitration and carbonylation of tubulin (as well as instability of microtubules). Oxidant also did not overproduce NO or increase DCF fluorescence in these dominant negative clones. Indeed, the I
B
mutants (e.g., 3-µg clones) induce almost complete protection against oxidation and/or nitration. Fourth, pretreatment of intestinal monolayers with pharmacological inhibitors of NF-
B suppresses NF-
B activity and evokes protective cascade of alterations that are further consistent with the proposed mechanism. Not surprisingly, the effects of I
B
mutant clones against oxidation are selectively mimicked by the I
B
inhibitors (e.g., lactacystin) as well as NF-
B suppressors (e.g., curcumin). These various inhibitors of NF-
B thus substantially prevented cascade of oxidant stress injury, including cytoskeletal nitration. The concordance of our findings utilizing both pharmacological inhibition and molecular targeting further supports a key role for NF-
B in these injurious oxidative stress processes for the epithelial cytoskeletal dysfunction.
Finally, NF-B activation quantitatively correlates with increases in all parameters indicating oxidative stress, which further supports our conclusions. Using both transfected clones and wild type cells, we found correlations (P < 0.05) between NF-
B activation and microtubule oxidation (r = 0.96) and several other outcomes of oxidative stress and cytoskeletal oxidation; for example, tubulin nitration and NF-
B activation (r = 0.97), tubulin oxidation and NF-
B activation (r = 0.96), and tubulin disassembly (increase in S1 monomeric pool) and NF-
B activation (r = 0.90). Importantly, other consistent correlations are also reached when oxidant-induced iNOS upregulation and NF-
B activation (r = 0.92), NO overgeneration and NF-
B activation (r = 0.91), or DCF fluorescence levels and NF-
B activation (r = 0.88) are utilized. The high strength of these various correlations, which explains 8595% of the variance, indicates that NF-
B activation is required for the injury induced by iNOS upregulation and consequent oxidative stress (e.g., nitration, carbonylation) to the tubulin cytoskeleton and intestinal epithelial monolayer. In this view, activation of NF-
B leads to the overproduction of NO (a RNM) and instability of the cytoskeleton following oxidative stress of iNOS activation. Our previous and current studies on NF-
B are consistent with a unique model for cytoskeletal and monolayer dysfunction under proinflammatory conditions of oxidant stress in which
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Our findings utilizing both pharmacology and targeted molecular approaches are further consistent with known properties of NF-B. NF-
B is typically composed of two subunits (p50 and p65), and its activation is tightly regulated by an endogenous cytoplasmic inhibitor, I
B
, which complexes with NF-
B and traps it in the cytoplasm in an inactive form (3, 15, 33, 42). NF-
B is a crucial proinflammatory factor in the immune response induced by a wide variety of other agents including cAMP, phorbol esters, free radicals, and cytokines (e.g., TNF-
, IL-1, IL-6) as well as viral transactivators (19, 21, 24, 40). For example, cytokine-triggered nuclear distribution and activation of NF-
B has been shown to be key in the promotion of inflammatory processes in both non-GI (e.g., peritoneal macrophages) (40, 46, 52) and GI models (e.g., intestinal cells) (3, 33, 42). Also, bacterial lipopolysaccharide (LPS) induces instability of I
B
, resulting in distribution of "free" NF-
B into the nucleus, which, in turn, activates an inflammatory response (19). Consistent with our current and previous findings, H2O2 has been shown to activate NF-
B (through destabilization of I
B
) in several cellular models. For instance, activation of NF-
B induced by H2O2 or TNF-
has been shown in endothelium (20). Similarly, NF-
B activation induced by oxidant has been reported in colonic smooth muscles (49) as well as in Jurkat T-cells (23). Moreover, recent studies from our laboratory, which were based on both pharmacological and molecular biological approaches, showed that the 78-kDa "classical"
1 isoform of protein kinase C (PKC-
1) and the 72-kDa "atypical"
isoform of PKC (PKC-
) are both required for suppression of NF-
B activation in colonocytes (15, 16). We showed that both PKC-
1 and PKC-
isoforms are key for growth factor (EGF, TGF-
)-induced protection of the intestinal epithelium (4, 5, 15, 16). Nevertheless, altogether, our studies on damage support a new model showing that a fundamental mechanism in the cascade of events that underlies disruption of the GI epithelial monolayer and cytoskeletal integrity under oxidant stress involves an injurious cascade of events that is likely initiated by free radicals. In this injurious cascade, oxidants induce I
B
instability (degradation) and then activate NF-
B, a crucial inflammatory mediator. We have now expanded on previous studies and shown that oxidants cause oxidative stress injury, especially nitrotyrosination and carbonylation, to the cytoskeletal subunit assembly through the activation of the NF-
B, which then leads to NO overproduction and its injurious oxidation consequences. Thus it appears that activating NF-
B will have distinct injurious, oxidative stress effects on the intestinal epithelial cytoskeleton, including nitration. Overall, our findings, we believe, indicate a new function for NF-
B under pathophysiological conditions of oxidant challenge, namely, promotion of the stress of NO overproduction and consequent cytoskeletal nitration and carbonylation in intestinal epithelial monolayers. We have thus discovered a novel biological mechanism for cytoskeletal and cellular oxidation.
Our findings are potentially relevant for developing new treatment modalities for inflammation, in general, and IBD, in particular. The manifestations of IBD, including ulcerative colitis and Crohn's disease, wax and wane between active (symptomatic) phases of disease, when oxidant stress is high, and inactive (asymptomatic) phases, when oxidative stress is minimal. Our series of findings suggests a unique oxidative stress mechanism that could, if it occurred in vivo, promote oxidative injury and initiate or perpetuate manifestation of the IBD attacks. This concept is consistent with recent characterizations of the pathophysiology of IBD and of the proinflammatory nature of NF-B (12, 21, 48). NF-
B activation is a critical event in the inflammatory response induced by an array of conditions, especially free radicals (3, 11, 15, 19, 20, 24, 33). NF-
B activation (as indicated by increased p65 nuclear levels) occurs in the inflamed mucosa of patients with IBD (ulcerative colitis or Crohn's disease) (11, 12, 43, 46, 48), in which excessive concentrations of oxidants (H2O2) as well as hyperpermeability of mucosal barrier function have been found (17, 28, 32, 35, 37, 39). We have found that the amount of oxidant stress and NF-
B activation closely parallel the degree of mucosal inflammation in patients with IBD (11, 12, 17, 35). Equally important, we recently found that the degree of gut mucosal cytoskeletal oxidation (and instability) correlates with the degree of inflammation and disease severity score in IBD patients (35). Thus induction of NF-
B appears to be key to the perpetuation of the active, symptomatic phase of IBD, in which intestinal oxidant stress creates a vicious cycle of inflammation dependent on sustained NF-
B activation, oxidant production, cytoskeletal instability, and, ultimately, tissue injury. The oxidative stress injury to key cytoskeletal structures mediated by NF-
B, as we have shown here in intestinal cells, could be pivotal in developing more effective strategies to suppress the continuation of inflammatory processes and structural damage in IBD mucosa.
In summary, our findings utilizing several independent approaches, including targeted molecular interventions, support the novel idea that NF-B is responsible for a substantial portion of nitration/oxidation disruption of the intestinal mucosal epithelial cytoskeleton following NO upregulation. NF-
B, perhaps, is also crucial to promoting amplification and establishment of an uncontrolled, oxidant-initiated inflammatory stress cascade that causes structural (cytoskeletal) tissue damage in IBD, one that can be ignited by free radicals and other oxidants present in the GI tract under pathophysiological conditions.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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