Division of Digestive Diseases, Department of Internal Medicine, and Departments of Pharmacology and Molecular Biophysics and Physiology, Rush University Medical Center, Chicago, Illinois 60612
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ABSTRACT |
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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. H2O2
concentrations 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; G-actin; nitration; oxidation; disassembly; nitric oxide; peroxynitrite; Caco-2 cells; inducible nitric oxide synthase
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INTRODUCTION |
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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 -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
, 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.
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MATERIALS AND METHODS |
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Cell culture. 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 · h1 · cm
2) = Fab/([FSA]a × S), where Fab 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 of L-[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 NG-nitro-L-arginine (1 mM) were used to assess the extent of L-[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 inhibitor L-N6-(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
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
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.
Statistical analysis. 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.
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RESULTS |
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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.
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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
72%), which is similar to
L-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.
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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% by
L-NIL, 84% by urate, 81% by L-cysteine, and
88% by SOD. These effects on the actin cytoskeleton by
L-NIL and antioxidants paralleled their protective effects
on barrier function.
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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
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 as
L-cysteine, urate, and SOD significantly prevented barrier
disruption resulting from ONOO
mimetics (data not shown),
confirming our previous findings.
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DISCUSSION |
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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
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(1) |
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
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 of
L-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
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 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. 9A).
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 nor L-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.
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ACKNOWLEDGEMENTS |
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This work was supported in part by a grant from Rush University Medical Center and the American College of Gastroenterology.
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FOOTNOTES |
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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: ali_banan{at}rush.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 5 October 2000; accepted in final form 24 January 2001.
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