Using oxidant-induced hyperpermeability of monolayers of intestinal (Caco-2) cells as a model for the pathophysiology of inflammatory bowel disease (IBD), we previously showed that oxidative injury to the F-actin cytoskeleton is necessary for the disruption of monolayer barrier integrity. We hypothesized that this cytoskeletal damage is caused by upregulation of an inducible nitric oxide (NO) synthase (iNOS)-driven pathway that overproduces reactive nitrogen metabolites (RNMs) such as NO and peroxynitrite (OONO−), which cause actin nitration and disassembly. Monolayers were exposed to H2O2 or to RNMs with and without pretreatment with antioxidants or iNOS inhibitors. H2O2concentrations that disassembled and/or disrupted the F-actin cytoskeleton and barrier integrity also caused rapid iNOS activation, NO overproduction, and actin nitration. Added OONO− mimicked H2O2; iNOS inhibitors and RNM scavengers were protective. Our results show that oxidant-induced F-actin and intestinal barrier disruption are caused by rapid iNOS upregulation that further increases oxidant levels; a similar positive feedback mechanism may underlie the episodic recurrence of the acute IBD attack. Confirming these mechanisms in vivo would provide a rationale for developing novel anti-RNM therapies for IBD.
- inflammatory bowel disease
- nitric oxide
- Caco-2 cells
- inducible nitric oxide synthase
under normal conditions, the gastrointestinal (GI) mucosa is a highly selective barrier that prevents the passage of toxic proinflammatory molecules from the gut lumen into the mucosa and the circulation (12, 20, 27). Under abnormal conditions, loss of GI barrier integrity can permit the penetration of normally excluded luminal substances (e.g., endotoxin and microbes) into or across the mucosa, which can lead to the initiation and/or perpetuation of inflammatory processes and mucosal injury. This injury and the ensuing loss of mucosal barrier integrity have been implicated in the pathophysiology of a wide range of inflammatory disorders including inflammatory bowel disease (IBD) (12, 16, 20, 27). The pathogenesis of mucosal barrier dysfunction in IBD remains poorly understood, but several studies, including ours (2, 6, 8), have shown that chronic gut inflammation is associated with high levels of reactive oxygen metabolites (ROMs) and that these oxidants appear to be involved in causing mucosal barrier dysfunction (2, 8, 26,30). However, the precise biochemical mechanisms have not been established.
While investigating these mechanisms with monolayers of intestinal cells as a model of barrier function, we (2, 5-8) demonstrated that oxidant-induced barrier disruption is dependent on the disruption of the cytoskeleton. For example, we observed that loss of barrier integrity required disruption of the polymerized (F)-actin cytoskeleton. The idea that cytoskeletal instability in general and actin filament disruption in particular could be major contributing factors to loss of barrier integrity is consistent with other studies in which we have shown the importance of cytoskeletal stability in GI healing in vivo (3, 4) as well as in vitro (2, 5–10a, 25). It is also consistent with the known roles of actin in cell function. Actin is one of the most abundant proteins in the eukaryotic cells, with the ability to polymerize into filaments of highly dynamic α-double helices (11, 38). In epithelial cells, such filaments constitute a dense cross-linked actin cortex located on the inner side of the plasma membrane. The actin cytoskeleton also contains filamentous stress fibers that traverse the cytosol and short fibers that extend into the lamellipodia in motile cells. This structural component is essential in maintaining normal cellular physiology, structure, locomotion, and support functions (2, 11, 29, 38, 39).
Our current investigation also derives from recent reports that elevated levels of peroxynitrite (ONOO−) may be an essential factor in tissue injury during IBD (23, 34, 35). Indeed, studies from our laboratory on ethanol-induced intestinal injury, as well as those from other laboratories, have shown that inducible nitric oxide (NO) synthase (iNOS) activation can lead to NO overproduction (8, 14, 36, 40) and that the injurious effects of NO overproduction appear to be mediated by ONOO−, the product of the reaction of NO with superoxide anions (O ·) (7, 18, 22, 34). Accordingly, the objectives of the current study were to determine1) whether disruption of the actin cytoskeleton and loss of barrier integrity after exposure to oxidants such as H2O2 are associated with elevated iNOS activity and with elevated levels of reactive nitrogen metabolites (RNMs) such as NO and ONOO−, 2) whether exposure of cells to these RNMs mimics the effects of oxidants, and 3) whether agents that scavenge these RNMs (antioxidants) or inhibit their formation (iNOS inhibitors) are protective.
MATERIALS AND METHODS
Intestinal Caco-2 cells (American Type Culture Collection, Manassas, VA) grown for barrier integrity work were split at a ratio of 1:2 and seeded at a density of 200,000 cells/cm2 in 0.4-μm collagen I cell culture inserts (0.3-cm2 growth surface; Biocoat, Becton Dickinson Labware, Bedford, MA), and experiments were performed at least 7 days after confluence. The utility, maintenance, and characterization of this cell line and the preparation of the monolayers of Caco-2 cells have been previously described (9, 17,39).
Determination of epithelial barrier function by fluorometry.
The barrier integrity of Caco-2 monolayers was determined by measuring the apical-to-basolateral flux of fluorescein sulfonic acid [(FSA); 200 μg/ml; 478 Da; Molecular Probes, Eugene, OR] as previously described (2, 6, 39). After the treatments, fluorescent signals from the samples were quantitated with a fluorescence multiplate reader (FL 600, Bio-Tek Instruments) with the excitation and emission spectra for FSA set as excitation = 485 nm and emission = 530 nm. Clearance (Cl) was calculated with the formula Cl (nl · h−1 · cm−2) =F ab/([FSA]a ×S), where F ab is the apical-to-basolateral flux of FSA in light units per hour, [FSA]a is the concentration at baseline in light units per nanoliter, and S is the surface area (0.3 cm2) (2, 8). Simultaneous controls were performed with each experiment.
Assay of NOS activity.
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). The conversion ofl-[3H]arginine (l-Arg; Amersham, Arlington Heights, IL) to l-[3H]citrulline was measured in the homogenates by scintillation counting as described previously (7, 8, 36). As we previously reported (7,8), experiments in the absence of NADPH and the presence of the NOS inhibitor N G-nitro-l-arginine (1 mM) were used to assess the extent ofl-[3H]citrulline formation that was independent of any NOS activity. Experiments in the presence of NADPH, without Ca2+ and with 5 mM EGTA, determined Ca2+-independent NOS (iNOS) activity. Experiments in the presence of NADPH and Ca2+ determined Ca2+-dependent NOS [constitutive NOS (cNOS)] activity. In selected experiments, we added the isoform-selective iNOS inhibitorl-N 6-(1-iminoethyl)lysine (l-NIL, 1 mM). Protein concentrations were determined by the Bradford method (13).
Western blot analysis of the level of iNOS protein.
After treatment, the cells were washed once with cold PBS, scraped in 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 gels. Subsequently, the proteins were transferred to nitrocellulose membranes and blocked in 3% BSA for 1 h, followed by several washes in Tris-buffered saline. The immunoblotted proteins were incubated for 2 h in Tween 20, Tris-buffered saline, 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) was used as a secondary antibody at 1:3,000 dilution. The membranes were visualized by enhanced chemiluminescence (Amersham) and autoradiography (7, 37).
Chemiluminescence analysis of NO concentration in cultures.
NO production was assessed by a novel chemiluminescence procedure (1, 7). Briefly, cells were homogenized by sonication, and the endogenous nitrate (NO ) and nitrite (NO ) and the metabolic degradation products of NO were then reduced to NO with the use of vanadium(III) (Sigma, St. Louis, MO) and HCl at 90°C before the measurement of NO concentration by chemiluminescence analysis. Chemiluminescence was measured with a Sievers NO 280 analyzer (NOA, Boulder, CO). NO (expressed in μM) was calculated by comparison with the chemiluminescence of a standard solution of NaNO2. The absolute NO values were reported as micromoles per 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 dichlorofluorescein (DCF) as previously described (8). The dependence of the assay on O · generation was shown by adding an active superoxide radical scavenger, superoxide dismutase (SOD, 300 U/ml), or an inactive superoxide radical scavenger, heat-inactivated SOD (iSOD). Briefly, monolayers grown in 96-well plates were preincubated with the membrane-permeant DCFD (10 μg/ml for 30 min) before the subsequent treatments. Fluorescent signals (i.e., DCF fluorescence) from the samples were quantitated with a fluorescence multiplate reader set at an excitation wavelength of 485 nm and an emission wavelength of 530 nm.
Immunofluorescent staining and high-resolution laser scanning confocal microscopy of the actin cytoskeleton.
Cells from monolayers were fixed in cytoskeleton stabilization buffer and then postfixed in 95% ethanol as previously described (2,9). Monolayers of 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 and then were mounted in Aquamount. The samples were examined by both standard fluorescence microscopy and ultra high-resolution laser scanning confocal microscopy (LSCM; Carl Zeiss). Cell monolayers on slides were analyzed in a blinded fashion using LSCM with a ×63 oil immersion plan-apochromat objective, NA 1.4 (Zeiss). An argon laser (wavelength = 488 nm) was used to examine FITC-labeled cells, and the cytoskeletal elements were examined for their overall morphology, orientation, and disruption as previously described (2, 9).
Actin fractionation and quantitative Western immunoblotting of F-actin and monomeric actin.
F-actin and monomeric (G)-actin were isolated as we previously described (2, 5). Briefly, cells were pelleted by centrifugation at low speed (700 rpm for 7 min at 4°C) and resuspended in actin stabilization-extraction buffer (0.1 M PIPES, pH 6.9, 30% glycerol, 5% DMSO, 1 mM MgSO4, 10 μg/ml of a standard anti-protease cocktail, 1 mM EGTA, and 1% Triton X-100) at room temperature for 20 min. F- and G-actin were separated after a series of ultracentrifugation and extraction steps. Fractionated actin samples were flash frozen in liquid N2 and stored at −70°C until immunoblotting was performed. For immunoblotting, samples (5 μg of protein) were placed in a standard SDS buffer, boiled for 5 min, and then subjected to PAGE (7.5% gel) (9). To quantify the relative levels of actin, the optical density (OD) of the bands corresponding to the immunoradiolabeled actin was measured with a laser densitometer (2).
Immunoblotting determination of actin oxidation and actin nitration.
Oxidation and nitration of the actin cytoskeleton were assessed, respectively, by measuring protein carbonyl and nitrotyrosine formation. Carbonylation and nitrotyrosination of actin were determined in a manner similar to the quantitative blotting of actin (2,8). To avoid the unwanted oxidation of the actin samples, all buffers contained 0.5 mM dithiothreitol and 20 mM 4,5-dihydroxy-1,3-benzenedisulfonic acid (Sigma). To determine the carbonyl content, samples were blotted to a polyvinylidene difluoride membrane and then subjected to successive 5-min incubations in 2 N HCl and 2,4-dinitrophenylhydrazine (DNPH; 100 μg/ml in 2 N HCl; Sigma). 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 10× 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). The membranes were then incubated with a HRP-conjugated secondary antibody (1:4,000 dilution; Molecular Probes) for 1 h. To determine nitrotyrosine content, after the blocking step listed above (i.e., BSA-PBS-T buffer), the membranes were probed for nitrotyrosine by incubation with 2 μg/ml of monoclonal anti-nitrotyrosine antibody for 1 h (Upstate Biotechnology, Lake Placid, NY) followed by the HRP-conjugated secondary antibody as described for carbonylation. The wash steps and film exposure were as in a standard Western protocol (8, 9). The relative levels of oxidized or nitrated actin were then quantified by measuring the OD of the bands corresponding to anti-DNPH or anti-nitrotyrosine immunoreactivity with a laser densitometer. Immunoreactivity was expressed as the percentage of carbonyl or nitrotyrosine formation (OD) in the treatment group compared with the maximally oxidized or nitrated tubulin standard. Oxidized or nitrated tubulin standards were run concurrently with the corresponding treatment groups.
Data are presented as means ± SE. All experiments were carried out with a sample size of at least 4–6 observations/group. Statistical analysis between or among groups was carried out with analysis of variance followed by Dunnett's multiple-range test (19). Correlational analyses were done with the Pearson test for parametric analysis or, when applicable, the Spearman test for nonparametric analysis. A P value < 0.05 was deemed to represent statistical significance.
Evidence that oxidant-induced leakiness of the intestinal barrier involves activation of iNOS.
We first confirmed our earlier finding (2) that exposure of Caco-2 cell monolayers for 30 min to increasing concentrations of H2O2 causes hyperpermeability of the intestinal barrier in the monolayers in a dose-dependent manner. This is indicated by the increased clearance of FSA (Fig.1). We now show that preincubation (1 h) with a selective iNOS inhibitor (l-NIL, 1 mM) significantly attenuates (−72%) this effect. This inhibition was significantly lower (−50%) at the higher oxidant doses: FSA clearance (in nl · h−1 · cm−2) = 1,287 ± 72 for l-NIL + 5 mM H2O2 vs. 2,589 ± 89 for 5 mM H2O2 alone. A substrate for NOS,l-Arg (3 mM, 48-h exposure), by itself did not significantly affect permeability, but it did synergize with a nondamaging concentration of H2O2 (0.05 mM) to disrupt monolayer barrier integrity. Moreover, l-Arg potentiated the loss of monolayer barrier integrity in the presence of damaging H2O2 concentrations (0.5 mM H2O2 is shown). In both cases, potentiation was prevented by l-NIL.
Evidence that NO, oxidative stress, and ONOO− are also involved in oxidant-induced monolayer barrier dysfunction.
Pretreatment of monolayers with the NO and ONOO−scavengers urate (−66%) and l-cysteine (−74%) or the O · scavenger SOD (−72%), which is similar tol-NIL, significantly attenuated H2O2-induced monolayer hyperpermeability (Fig.2). Pretreatment with iSOD was not protective. iSOD by itself did not injure cells (clearance = 28 ± 9 vs. 22 ± 8 nl · h−1 · cm−2 for vehicle). The failure to elicit 100% protection by the aforementioned antioxidants was not due to technical problems because the addition of catalase, an H2O2 scavenger (1,000 U/ml), elicited 99% protection.
Figure 3 Ashows that doses of H2O2 that caused hyperpermeability also caused significant increases in Ca2+-independent, l-NIL-inhibitable NOS (i.e., iNOS) activity in lysates of Caco-2 monolayers compared with control monolayers, which displayed low iNOS activity levels. In contrast, neither oxidants nor l-NIL or their combination had any effect on Ca2+-dependent cNOS (oxidant, 0.31 ± 0.10;l-NIL, 0.33 ± 0.07; l-NIL plus oxidant, 0.30 ± 0.08; vehicle alone, 0.29 ± 0.12 pmol · min−1 · mg protein−1).
Figure 3 B depicts a representative Western blot showing that H2O2 significantly increased iNOS protein levels; control monolayers exhibited low basal levels of iNOS protein. The corresponding ODs were control, 888 ± 92; nondamaging concentrations (0.05 mM) of H2O2, 924 ± 81; and damaging concentrations (0.5 mM) of H2O2, 4,251 ± 107.
Figure 3 C shows NO overproduction in cell monolayers exposed to H2O2. NO overproduction was almost completely prevented by pretreating the monolayers withl-NIL. A nondamaging concentration of H2O2 (0.05 mM), one that did not significantly increase permeability, induced neither iNOS activity (0.40 ± 0.15 vs. 0.35 ± 0.04 pmol · min−1 · mg protein−1 for vehicle) nor NO overproduction (1.93 ± 0.42 vs. 1.72 ± 0.19 μmol/106 cells for vehicle).
Figure 4 shows the time course for increases in iNOS protein, iNOS activity, and NO levels. These effects of H2O2 were rapid; more than two-thirds of the changes occurred within the first 30 min. Maximal changes were 4.8-fold for iNOS protein, 10.1-fold for iNOS activity, and 10.1-fold for NO levels.
H2O2 also significantly increased the fluorescence of DCF (Fig. 5), a marker for oxidative stress. This increase was prevented by l-NIL, by NO and ONOO− scavengers (l-cysteine is shown), and by O · scavengers (SOD but not iSOD). These data suggest that iNOS activation and its reaction products contribute to the increase in oxidative stress in the cell. These data also confirm the generation of NO and O · after exposure to H2O2.
NO- and ONOO−-dependent mechanisms in the loss of F-actin cytoskeletal integrity.
Pretreatment of Caco-2 monolayers with l-NIL or with the above noted antioxidants protected the F-actin cytoskeleton against most of the H2O2-induced damage, an outcome that was quantitated with the use of LSCM and by calculating changes in the percentage of cells displaying normal actin (Fig.6). A 0.5 mM dose of H2O2 decreased the proportion of cells showing normal actin by 49%. The extent of this damage was inhibited 90% byl-NIL, 84% by urate, 81% by l-cysteine, and 88% by SOD. These effects on the actin cytoskeleton byl-NIL and antioxidants paralleled their protective effects on barrier function.
Immunofluorescent staining, as used in the experiments shown in Fig.7, revealed that pretreatment withl-NIL before H2O2 protected the actin cytoskeleton against injury. This was shown by a normal smooth and continuous pattern of the actin “ring” at the areas of cell-cell contact (Fig. 7 C), which was comparable in appearance to the control monolayers (Fig. 7 A) and quite different from the injured (i.e., disrupted, fragmented, condensed, and beaded) appearance of actin in cells exposed to oxidants (Fig.7 B).
Quantitative Western immunoblotting analysis (Fig.8 A) demonstrated that 0.5 mM H2O2 decreased the fraction of stable (polymerized) F-actin by 31% and increased the fraction of monomeric G-actin. Pretreatment with l-NIL or antioxidants attenuated this effect (l-NIL, −82%; urate, −94%; l-cysteine, −88%; SOD, −82%).
A representative Western blot gel (Fig. 8 B) demonstrated that preincubation with l-NIL or antioxidants enhanced the autoradiographic band density for F-actin extracted from the monolayers to the control level, independently confirming enhanced assembly (and stabilization) of actin.
To measure the “footprints” of ONOO− formation, namely nitrotyrosine and carbonyl moieties, the F-actin cytoskeleton of the cell monolayers was fractionated and isolated. We then used a sensitive quantitative immunoblot (Fig.9 A). H2O2 resulted in the nitration and oxidation of the F-actin cytoskeleton. For instance, the fraction of F-actin that was nitrated by 0.5 mM H2O2 was equal to 0.65 ± 0.03%, and this was similar to the fraction of F-actin that was carbonylated (0.70 ± 0.015%, ratios normalized to a nitrated or oxidized actin standard run concurrently). Figure9 B shows that pretreatment of cell monolayers with the iNOS inhibitor l-NIL or with any of our antioxidants significantly protected against the nitration and carbonylation of the F-actin filaments (nitration shown in Fig. 9 B:l-NIL, −93%; urate, −85%; l-cysteine, −98%; SOD, −73%). There was a significant (P < 0.05) positive correlation between 1) F-actin disruption and2) F-actin nitration (r = 0.98) or oxidation (r = 0.96). These data are consistent with the protective effects of these same antioxidants against barrier dysfunction.
ONOO− compounds mimic the ability of H2O2 to cause F-actin cytoskeletal instability and barrier disruption.
If ONOO− mediates oxidant-induced damage, then chemically authentic ONOO− or compounds capable of generating ONOO− should mimic H2O2 and cause similar disruption of the F-actin cytoskeleton and monolayer barrier, and these effects should be inhibited by the same antioxidants that inhibit the disruption that is caused by H2O2. First, we found that ONOO− and ONOO−generators [e.g., 3-morpholinosydnonimine (SIN-1), a NO and O · donor, andS-nitroso-N-acetylpenicillamine, a NO donor] in combination with xanthine plus xanthine oxidase (an O · donor) significantly and dose dependently disrupted the monolayer barrier (data not shown), which confirmed our previous findings (7) that these ONOO−compounds increase FSA clearance. ONOO− generator systems were then added to the cell culture media at a pH of 7.4. To promote the stability of authentic added ONOO− in solution, ONOO− (180 mM stock in 0.3 M NaOH) was added to the cell culture media to a final pH of 7.6. Pilot studies confirmed that there were no adverse effects of a pH of 7.6 on the cytoskeleton or on monolayer barrier function. We also found that antioxidants such asl-cysteine, urate, and SOD significantly prevented barrier disruption resulting from ONOO− mimetics (data not shown), confirming our previous findings.
In the present study, these same ONOO− mimetics also depolymerized F-actin filaments as shown by increased G-actin and reduced F-actin (Fig. 10 A). In contrast, antioxidants (urate, l-cysteine, SOD) almost completely prevented both the actin oxidation and nitration (Table1) and the actin depolymerization (Fig.10 B) that result from exposure of monolayers to ONOO− compounds. LSCM (Fig.11) showed that ONOO−elicited actin disruption, aggregation, and kinking (Fig.11 B). Pretreatment of the monolayers with the antioxidantl-cysteine (Fig. 11 C) protected the F-actin cytoskeleton against disruption induced by ONOO−. The antioxidant-treated group (C) was indistinguishable from the control group (Fig. 11 A).
Table 2 shows that ONOO− and HOCl, like H2O2, upregulated both iNOS activity and NO levels (compare Fig. 3, A and C, with Table 2). Indeed, iNOS upregulation occurs after the addition of other agents that are injurious to the intestinal barrier, such as ethanol.
Together, our present findings support our main conclusion that the H2O2-induced disruption of the F-actin cytoskeleton of Caco-2 cells and the consequent disruption of the permeability barrier of Caco-2 monolayers require activation of an iNOS-driven pathway and increased levels of RNMs such as NO and ONOO−, which appear to mediate this damage through nitration and oxidation of a 43-kDa actin molecule. A second conclusion, and a novel finding, is that exposure of these intestinal cells to oxidants, a process that models the oxidative stress that occurs during the acute IBD attack, can, surprisingly, further increase cellular synthesis of RNMs and ROMs. A third conclusion, also novel, is that oxidants such as H2O2, HOCl, and ONOO− can rapidly upregulate iNOS enzyme activity and NO levels.
Our primary conclusion is relevant to IBD. It extends our previous investigation into the role of oxidants in the pathophysiological mechanisms of this disease. Although the primary etiology of IBD is multifactorial, gut leakiness and diffusion of intraluminal proinflammatory antigens (e.g., bacterial products) are considered to be reasonable initial steps for subsequent intestinal inflammation. Similarly, in our monolayer model, oxidants induced barrier hyperpermeability. We (2) previously traced the in vitro cause of this monolayer hyperpermeability back to disruption of the functioning and architectural integrity of actin filaments. We now show that iNOS upregulation and oxidation and nitration of the subunit components of the actin cytoskeleton are a major part of the mechanism for the disruption of the F-actin filaments. In this view, the connection between iNOS and actin nitration is through the reaction Equation 1an idea that is supported by both the existing literature (21, 31, 34) and our finding here that DCF fluorescence is quenched by both O · scavengers (SOD) and by NO and ONOO− scavengers.
Two factors further enhance the validity of this primary conclusion. First, the conclusion is supported in the present study by three independent lines of investigation: 1) H2O2 concentrations that cause actin damage and intestinal hyperpermeability simultaneously activate iNOS and increase RNMs and oxidative stress; 2) three different exogenously added RNMs mimicked all the effects of H2O2; and 3) both iNOS inhibitors and antioxidants that scavenge RNMs prevented or substantially attenuated the injurious changes that resulted from exposure either to H2O2 or to RNM compounds. Second, we found robust positive correlations between increases in RNMs, increases in oxidative stress, RNM-associated nitration and disruption of the F-actin cytoskeleton, and increases in monolayer permeability. Correlation analysis showed a significant (P < 0.05) and robust correlation between NO and nitrotyrosine levels (r = 0.97) and between nitrotyrosine levels and either actin disruption (r = 0.98) or actin disassembly (r = 0.94). Similar to nitration, oxidation, as measured by either carbonyl levels or DCF fluorescence, predicted actin disruption (r values of 0.92 and 0.96, respectively). There was also a significant positive correlation (r = 0.95) between H2O2-induced barrier disruption (FSA) and actin disassembly. Finally, the two markers for actin integrity (i.e., percentage of polymerization and percentage of normal actin) strongly correlated with each other (r = 0.96). Although it is possible that the increase in ONOO− levels observed after exposure of the cells to H2O2 derived from sources other than iNOS and its reaction product NO, our data suggest that this would likely be a relatively minor source because the iNOS inhibitor l-NIL nearly completely abolished oxidant-induced nitration injury to the actin cytoskeleton and maintained barrier integrity. Overall, the biochemical cause of injury to the actin network appears to be the nitration and oxidation of its 43-kDa protein subunits.
It might be argued that the antioxidants l-cysteine and urate are not specific to RNMs and might inhibit the synthesis of ONOO− (see eq. 1) by scavenging O · as SOD does. Although this could conceivably occur, there would still be overwhelming evidence that NO and ONOO− are involved in oxidant-induced damage in our model. First, ONOO− mimetics (authentic ONOO− or ONOO−-generating systems) mimic H2O2. Second, nitration as measured by nitrotyrosine formation on actin protein occurs in Caco-2 cells after the addition of oxidants (H2O2, ONOO−). Third, a selective iNOS inhibitor (l-NIL) prevents nitration. Fourth, our own previous experiments (7) involving the addition ofl-cysteine to a test tube containing authentic ONOO− (in which l-cysteine completely scavenged the ONOO−) and other literature (15, 22,32-34) clearly indicate that l-cysteine and urate are capable of scavenging RNMs. Moreover, they must be doing so here because urate and l-cysteine prevented both the F-actin nitration and the F-actin depolymerization that was caused when we directly added authentic ONOO− to our monolayers. Fifth, the free sulfhydryl groups of l-cysteine react with ONOO− with a rate constant (5,900 M−1s−1) that is over 1,000-fold greater than its rate constants for reaction with other oxygen species such as H2O2 or O · (∼4 M−1s−1)(32), making it more likely that l-cysteine is effectively scavenging ONOO− at a greater rate than O ·.
Our primary conclusion is consistent with recent studies showing that O · reacts rapidly with NO to generate ONOO− in vivo (21, 31). These in vivo studies have led to the proposal that the formation of ONOO−radical is key in the pathogenesis of IBD and a variety of other inflammatory GI and systemic disorders (23, 28, 37), but the target proteins were unclear. For example, tissue nitration, which was detected by immunofluorescent staining of nitrotyrosine, has been associated with the inflamed human mucosa in IBD (28, 37) and was linked with the upregulation of iNOS (37). Some nitrated tissue proteins (e.g., SOD and glutathione) have been detected in vivo in non-GI models such as the inflamed lung (23,32). Moreover, it appears that ONOO−-induced tissue nitration (nitrotyrosination) involves the addition of nitro groups to the ortho position of tyrosine residues (22). We also previously showed (7) that ethanol induces tubulin nitration and oxidation in vitro in monolayers of human intestinal cells. The current study suggests that actin molecules and the F-actin cytoskeleton are key target proteins of oxidant-induced nitration. On the basis of our findings with ethanol (7, 8), it seems likely that tubulin and the microtubule cytoskeleton are also key target proteins. These conjectures are further supported by previous studies (2, 6) in which we showed that phalloidin and taxol, agents that prevent oxidative damage to the F-actin and the microtubule cytoskeletons, respectively, also prevent barrier disruption.
Interestingly, cytoskeletal injury does not need to affect all cells in the monolayer to elicit intestinal leakiness. The data presented in Fig. 6 indicate that significant damage occurs when only ∼50% of cells in the monolayer no longer show a normal actin cytoskeleton. This is accomplished under oxidative conditions in which there is a 65% increase in actin nitration (Fig. 9 A).
Finally, our primary conclusion is consistent with our most recent in vivo studies (48) in which immunoblotting analysis of the mucosal pinch biopsy specimens of inflamed intestinal tissues from IBD patients showed increased tissue levels of NO and nitrotyrosination of actin.
Our second conclusion is also relevant to IBD and suggests a novel positive feedback mechanism that could, if it occurred in vivo, very likely overwhelm endogenous antioxidant defenses and either initiate or sustain the acute IBD attack. This positive feedback is seen in the ability of three separate oxidants (H2O2, HOCl, and ONOO−) to upregulate iNOS, which then synthesizes NO and ONOO−. Although the reasons behind the presence of such a positive feedback mechanism in our model are unclear (oxidants may be activating cellular stress responses), the existence of this mechanism is consistent with the current characterization of the pathophysiology of IBD (26,28). This is especially true for the transition 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 leads to sustained inflammation and tissue damage. In particular, the natural course of IBD involves recurrent episodes of the inactive phase (in which there are no neutrophils) followed by acute exacerbation (flare-up) that is characterized by mucosal infiltration of large numbers of leukocytes including neutrophils. These plasma cells are capable of producing large quantities of ROMs (e.g., H2O2 and HOCl) and RNMs (e.g., ONOO−), reactive species that create a vicious cycle and sustain an inflammatory cascade. A positive feedback loop, such as the one we observed in Caco-2 cells, could play a key role in establishing such a vicious cycle.
Our third conclusion is that increases in the level or activity of iNOS can occur rapidly. This conclusion is supported by parallel increases in three separate variables: iNOS protein levels, iNOS enzyme activity, and NO levels. The findings of recent studies in endothelial cells, as well as one in vivo study in rat gastric mucosal cells, are also consistent with our finding of rapid iNOS upregulation (41-43). In the study that used isolated rat gastric mucosal cells, low basal levels of iNOS were noted in control (untreated) mucosa (41), whereas after challenge with endotoxin, significant increases in iNOS activity were detected in the mucosal cells within 1 h, followed by peak levels at 2–4 h. Similarly, other studies on endothelial cells showed detectable levels of iNOS activity as early as 60–90 min after H2O2 (0.1–1 mM) challenge (42,43). 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 (44). Furthermore, it seems unlikely that oxidant-induced increases in cNOS activity are occurring and confounding our finding that Ca2+-independent iNOS is upregulated, because we found that neither oxidants norl-NIL affect Ca2+-dependent cNOS activity.
A question that remains to be answered is how iNOS might be activated so rapidly. We now suggest three mechanisms by which the rapid iNOS upregulation might occur: protein synthesis starting from a preexisting mRNA pool; upregulation of inactive iNOS enzyme molecules by any of several well-known cellular mechanisms such as phosphorylation-dephosphorylation; and iNOS dimerization. The first mechanism requires a basal constitutive level of iNOS mRNA in unstimulated cells, as was suggested by a recent study in endothelial cells (44). In general, protein expression (i.e., transcription) from a “standing pool of mRNA” does not require more than 30–40 min.
A second proposed mechanism is the phosphorylation of the iNOS enzyme as a means of rapidly regulating its enzymatic activity. NOS contains consensus sequences for sites of protein phosphorylation (45,46); tyrosine phosphorylation of Ca2+-independent NOS has been shown in vitro and in endothelial cells after a variety of stimuli, and it was proposed that this mechanism could rapidly regulate the activity of NOS (45, 46).
A third 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. These monomers can be rapidly assembled by an appropriate stimulus into an active dimer (47). Future studies will be needed to investigate which of these mechanisms is operative in our model and in intestinal cells in general.
Although ONOO− is relatively short-lived at physiological pH (7.4), we believe that our methods, results, and conclusions drawn from the ONOO− systems that we have employed are valid for several reasons. 1) Our findings are consistent with several reports from the literature that indicate that ONOO− or its footprints have been detected in vivo in inflamed mucosal tissue from patients with IBD as well as from patients with other systemic inflammatory disorders such as in the lung (23, 26, 28, 35, 37). For example, ONOO− has been detected in the inflamed colonic mucosa by immunofluorescence analysis (28). Thus, although ONOO− is short-lived, its footprints have been detectable in vivo under a variety of pathophysiological conditions and, apparently, enough ONOO− is around for a long enough time for this to occur.2) ONOO− generator systems (SIN-1 and SNAP-xanthine-xanthine oxidase were used in our studies) caused nitration damage to actin and to the permeability barrier of our monolayers at pH 7.4. Both of these models are known to spontaneously generate ONOO− in vitro (25, 34). Moreover, the extent of the damage was equivalent to damage induced by oxidants (H2O2 or authentic ONOO− at pH 7.6). 3) Added authentic ONOO− caused a degree of damage at pH 7.6 similar to that caused by ONOO−generators at pH 7.4. At pH 7.6, ONOO− decreased over 30 min from 100% of added ONOO− to ∼38% as our laboratory previously reported (7). Thus a substantial fraction of the added ONOO− remains in contact with our monolayers and could very well lead to injury.
In summary, our studies to date indicate that the disruption of intestinal barrier function induced by oxidants is caused by iNOS upregulation, RNMs, and oxidative injury to the actin cytoskeleton and the microtubule cytoskeleton. If this mechanism can be demonstrated in vivo, then our in vitro findings would suggest intracellular mechanisms (e.g., iNOS inhibitors and ONOO− scavengers) that might serve as targets for the development of novel therapies for IBD.
This work was supported in part by a grant from Rush University Medical Center and the American College of Gastroenterology.
Portions of this work were presented in abstract form at the annual meeting of the American Gastroenterological Association in San Diego, CA, 2000.
Address for reprint requests and other correspondence: A. Banan, Rush Univ. Medical Ctr., Div. of Digestive Diseases, 1725 W. Harrison, Ste. 206, Chicago, IL 60612 (E-mail:).
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- Copyright © 2001 the American Physiological Society