Glutathione oxidation and PTPase inhibition by hydrogen peroxide in Caco-2 cell monolayer

R. K. Rao, L. Li, R. D. Baker, S. S. Baker, and A. Gupta

Department of Pediatrics, Medical University of South Carolina, Charleston, South Carolina 29425


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The role of H2O2 and protein thiol oxidation in oxidative stress-induced epithelial paracellular permeability was investigated in Caco-2 cell monolayers. Treatment with a H2O2 generating system (xanthine oxidase + xanthine) or H2O2 (20 µM) increased the paracellular permeability. Xanthine oxidase-induced permeability was potentiated by superoxide dismutase and prevented by catalase. H2O2-induced permeability was prevented by ferrous sulfate and potentiated by deferoxamine and 1,10-phenanthroline. GSH, N-acetyl-L-cysteine, dithiothreitol, mercaptosuccinate, and diethylmaleate inhibited H2O2-induced permeability, but it was potentiated by 1,3-bis(2-chloroethyl)-1-nitrosourea. H2O2 reduced cellular GSH and protein thiols and increased GSSG. H2O2-mediated reduction of GSH-to-GSSG ratio was prevented by ferrous sulfate, GSH, N-acetyl-L-cysteine, diethylmaleate, and mercaptosuccinate and potentiated by 1,10-phenanthroline and 1,3-bis(2-chloroethyl)-1-nitrosourea. Incubation of soluble fraction of cells with GSSG reduced protein tyrosine phosphatase (PTPase) activity, which was prevented by coincubation with GSH. PTPase activity was also lower in H2O2-treated cells. This study indicates that H2O2, but not O2-· or ·OH, increases paracellular permeability of Caco-2 cell monolayer by a mechanism that involves oxidation of GSH and inhibition of PTPases.

intestine; tight junction; protein tyrosine phosphatase; signal transduction; tyrosine kinase


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

REACTIVE OXYGEN SPECIES (ROS) play an important role in ischemic tissue injury and pathogenesis of a number of intestinal disorders, including inflammatory bowel disease and necrotizing enterocolitis (11, 22). Superoxide (O2-·) is generated by electron transport chain, NADPH oxidase, and xanthine oxidase activity (12). O2-· undergoes dismutation reaction to generate hydrogen peroxide (H2O2) (11); this reaction is catalyzed by superoxide dismutase (SOD). H2O2 is normally detoxified by antioxidant defense enzymes, such as catalase in peroxisomes and glutathione peroxidase (Gpx) in mitochondria and cytosol (11). However, H2O2 is also rapidly split to ·OH by Fenton reaction (13), which is catalyzed by transition metal ions Fe2+ and Cu2+. ·OH is considered more reactive than O2-· and H2O2. Although all known ROS were found to be cytotoxic, little is known about the specific role of each ROS in oxidative stress-induced tissue injury. It is suggested that O2-· plays an important role in inflammatory response (23), and anti-inflammatory action of SOD was seen in several animal models of induced inflammation as well as in clinical trials in humans (6, 20, 22). There is no evidence for the role of O2-· in lipid peroxidation; however, O2-· can reduce Fe3+ to Fe2+, which in turn may accelerate ·OH generation from H2O2 (Haber Weiss reaction) (10). Additionally, O2-· can react with nitric oxide to generate peroxynitrite (18), which appears to be more toxic than O2-·. Therefore, it is usually believed that the toxicity of O2-· or H2O2 is mediated by the conversion into ·OH. ·OH is the most reactive ROS, reacting with virtually all biological systems. This oxidant species is suggested to be responsible for membrane lipid peroxidation (8), mitochondrial energization (8), hyaluronic acid degradation (4), and DNA fragmentation (28).

Our recent study demonstrated that xanthine oxidase-mediated oxidative stress increases paracellular permeability in Caco-2 cell monolayer by a mechanism that involves protein tyrosine phosphorylation (25, 26). However, the specific ROS or the mechanism by which it increases protein tyrosine phosphorylation is not known. It is likely that oxidative stress modulates the activities of tyrosine kinases and protein tyrosine phosphatases (PTPases) to increase cellular protein tyrosine phosphorylation. Specific proteins such as occludin, ZO-1, E-cadherin, and alpha -catenin that construct and regulate tight junctions and adherens junctions are likely targets for tyrosine phosphorylation. Because tight junctions and adherens junctions are responsible for impedance of paracellular permeability of macromolecules, tyrosine phosphorylation of junctional proteins may alter their permeability. In the present study, we determined the role of H2O2 as the prominent ROS responsible for increasing paracellular permeability in Caco-2 cell monolayer and tested the role of Gpx-mediated oxidation of GSH and inhibition of PTPase activity in the mechanism of H2O2-induced permeability.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture. Caco-2 cells obtained from American Type Culture Collection (Rockville, MD) were maintained under standard cell culture conditions at 37°C in medium containing 20% (vol/vol) fetal bovine serum. Cells were grown on polycarbonate membranes in Transwells (6.5 mm; Costar, Cambridge, MA). Experiments were performed 12-14 days after seeding of the cells.

Treatment with oxidants, antioxidants, and other compounds. Confluent monolayers were bathed in PBS/BSA (Dulbecco's saline containing 1.2 mM CaCl2, 1 mM MgCl2, and 0.6% BSA). Oxidative stress was induced by administering (to both apical and basal compartments) 20 µM H2O2 or a combination of xanthine oxidase (20 mU/ml) and xanthine (0.25 mM) (XO+X). In certain experiments, XO+X was administered along with SOD (20 or 100 µg/ml) or catalase (10 U/ml). In other experiments, cell monolayers were pretreated with FeSO4 (0.1 mM), deferoxamine (1 mM), 1,10-phenanthroline (0.1 mM), GSH (1 mM), N-acetyl-L-cysteine (NAC; 0.1 mM), dithiothreitol (DTT; 0.5 mM), mercaptosuccinate (0.1-10 mM), diethylmaleate (DEM; 0.15 mM), (+)-alpha -tocopherol acid succinate (vitamin E; 1 mM), all trans-retinol acetate (vitamin A; 1 mM) and 1,3-bis(2-chloroethyl)-1-nitrosourea (BCNU; 0.3 mM) for 90 min before H2O2 administration. Cell monolayers were washed two times with PBS/BSA after the pretreatment. Control monolayers were incubated in PBS/BSA with or without pretreatment with different compounds.

Assay of H2O2. H2O2 level was measured as described previously by Pick and Mizel (24). Briefly, 100-µl aliquots of incubation medium containing XO+X or H2O2, with or without cell monolayers and with or without the presence of other compounds, were incubated with 100 µl of phenol red solution (40 U/ml of horseradish peroxidase and 1.16 mM phenol red in PBS) in 96-well plates at room temperature for 15 min. Reaction was terminated by adding 10 µl of 1 N NaOH. Plates were read at 610 nm in an automated plate reader (HTS 7000 Bio Assay Reader; Perkin Elmer, Norwalk, CT). A standard curve was constructed using 2-50 µM H2O2.

Measurement of transepithelial electrical resistance. Transepithelial electrical resistance (TER) was measured according to the method of Hidalgo et al. (16) using a Millicell-ERS electrical resistance system (Millipore, Bedford, MA) and calculated as Omega  · cm2 by multiplying it with the surface area of the monolayer (0.33 cm2). The resistance of the supporting membrane in Transwells (which is usually ~30 Omega  · cm2) was subtracted from all readings before calculations.

Unidirectional flux of mannitol. Cell monolayers in Transwells were incubated under different experimental conditions in the presence of 0.2 µCi/ml of D-[2-3H]mannitol (15 Ci/mmol; ICN Biomedicals, Costa Mesa, CA) in the basal well. At different times after oxidant administration, 100 µl each of apical and basal media was withdrawn and radioactivity was counted in a scintillation counter. The flux into the apical well was calculated as the percentage of total isotope administered into the basal well per hour per square centimeter of surface area.

NaCl dilution potential. At the end of experimental treatments, monolayers were washed once with PBS/BSA and bathed in fresh PBS/BSA, 0.2 ml and 1.0 ml to apical and basal compartments, respectively. Transepithelial potential difference was recorded using Millicell-ERS. Twenty percent dilution in the apical compartment was developed by replacing 40 µl of apical medium with 40 µl of PBS/BSA in which NaCl was replaced with equiosmolar mannitol. The potential difference was recorded again, and the dilution potential was calculated from the difference between the initial potential difference and the potential difference recorded after 20% dilution.

Assay of GSH and protein thiols. Cell proteins were extracted in cold 5% 5-sulfosalicylic acid (5-SSA). Levels of GSH and GSSG in acid-soluble fractions were determined by enzymatic recycling assay using glutathione reductase (type IV; Sigma, St. Louis, MO) and 5,5'-dithio-bis-2-nitrobenzoic acid (DTNB; Sigma) as described previously (1). Total GSH was estimated by monitoring the rate of formation of chromophoric product 2-nitro-5-thiobenzoic acid at 412 nm in a Bio Assay Reader with the computer software HTSoft (Perkin Elmer).

For the measurement of GSSG, GSH in acid soluble fraction was first derivatized with 2-vinylpyridine in the presence of triethanolamine. Samples were then assayed for total GSH as described above. The amount of GSH or GSSG in samples was determined from a standard curve constructed by performing the recycling assay using 40-320 pmol of standard GSH. Because the level of GSSG in the cell was <1% of that of GSH, the level of GSH + GSSG was not significantly different from GSH alone.

The level of protein thiols was determined by measuring thiols in acid precipitates as described previously (5). Acid precipitates were washed with 5-SSA and dissolved in 0.5 M Tris · HCl, pH 7.6. Extracts were mixed with equal volumes of 0.2 mM DTNB. After 20 min at room temperature, the absorbency was measured at 420 nm. Data are expressed as nanomoles of thiol per milligram of protein, calculated on the basis of the GSH standard curve.

Preparation and treatment of plasma membrane and soluble fractions. Cell monolayers were washed twice with ice-cold PBS and once with lysis buffer F [PBS containing 10 mM beta -glycerophosphate, 2 µg/ml leupeptin, 10 µg/ml aprotinin, 10 µg/ml bestatin, 10 µg/ml pepstatin-A, 1 mM benzamidine, and 1 mM phenylmethylsulfonyl fluoride (PMSF)]. Cells were dispersed by homogenization in a glass/Teflon Dounce homogenizer with 50 strokes and lysed by sonication at 4°C for two strokes (5 s each) with a 30-s interval. Cell lysate was centrifuged first at 3,000 g for 10 min at 4°C to sediment the cell debris. Supernatant was further centrifuged at 30,000 g for 30 min at 4°C. Pellet was suspended in 500 µl of lysis buffer F. Aliquots of plasma membrane and soluble fractions were incubated at 37°C in the presence of varying concentrations of GSSG and/or GSH for 60 min before tyrosine kinase and PTPase assay. In certain experiments, plasma membrane and soluble fractions were isolated from cell monolayers treated with H2O2 for varying times. These fractions were directly assayed for PTPase activity.

PTPase assay. 32P-Raytide was first prepared by phosphorylating Raytide with [gamma -32P]ATP and c-Src (19). Soluble fraction of Caco-2 cell monolayers mixed in 60 µl of PTPase buffer (50 mM HEPES, pH 7.2, 60 mM NaCl, 60 mM KCl, 0.2 mM PMSF, 10 µg/ml aprotinin, 2 µg/ml leupeptin, and 10 µg/ml bestatin) containing 32P-Raytide (50,000 cpm). Assay mixture was incubated at 30°C for 10 min. Reaction was terminated by placing 50 µl of reaction mixture onto P81 filter paper disks (Whatman). Filter disks were washed with 0.5% phosphoric acid, and radioactivity was counted. For control, assay was conducted in the presence of 1 mM sodium orthovanadate. Activity was calculated as units (nmol phosphate hydrolyzed/hour from phosphopeptide substrate under assay conditions).

Tyrosine kinase assay. Aliquots of membrane fractions were incubated with a kinase buffer (50 mM imidazole, 250 mM NaCl, and 1 mM MnCl2, pH 7.4) containing 5 µg of poly(Glu,Tyr) and 5 µl of ATP mix (0.1 mM ATP, 72 mM MgCl2, 12 mM MnCl2, 0.6 mM vanadate, 12 mM p-nitrophenyl phosphate, and 0.6 mCi/ml [gamma -32P]ATP) in a total volume of 30 µl for 20 min at 25°C. Labeling of substrate was analyzed by placing 25 µl of assay mixture onto DE81 filter paper circles and washing with 5% TCA. Activity was expressed as micromoles phosphate incorporated to peptide per hour under assay conditions.

Chemicals. Cell culture media and related reagents were purchased from GIBCO BRL (Grand Island, NY). Xanthine oxidase, xanthine, GSH, GSSG, DTT, vitamin E, vitamin A, catalase, H2O2, deferoxamine mesylate, 1,10-phenanthroline, NAC, DEM, phenol red, mercaptosuccinate, horseradish peroxidase, poly(Glu,Tyr), ATP, and SOD were purchased from Sigma. BCNU was from Aldrich (Milwaukee, WI), and P81 and DE81 filter disks were from Whatman (Clifton, NJ). [gamma -32P]ATP was purchased from ICN Pharmaceuticals (Costa Mesa, CA). Raytide and pp60c-Src were from Oncogene Sciences (Cambridge, MA). All other chemicals were of analytical grade purchased either from Sigma or Fisher Scientific (Tustin, CA).

Statistics. Comparison between two groups was made by the Student's t-tests for grouped data or by analysis of variance and Fisher's post hoc test for comparisons of more than two groups. The significance in all tests was derived at the 95% or greater confidence level.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Absence of a role for O2-· in XO+X-induced increase in permeability. In the absence of cell monolayer, XO+X generated a maximum of 17.2 µM H2O2 by 15 min (Fig. 1A). In the presence of cell monolayer, XO+X generated 10.9 µM H2O2 in the basal well and 4.2 µM in the apical well at 15 min, followed by a drop to steady-state level of 3.0 µM and 1.3 µM at 60 min, respectively. XO+X treatment reduced the TER of Caco-2 cell monolayers in a time-dependent manner (Fig. 1B); nearly 80% decrease in TER was achieved at 3 h. XO+X-induced decrease in TER was associated with a reduction of dilution potential (Table 1) and an increase of mannitol flux (Fig. 1C).


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Fig. 1.   Time course of the effect of xanthine oxidase (20 mU/ml) and xanthine (0.25 mM) (XO+X) on paracellular permeability. A: H2O2 level achieved at varying times after XO+X. H2O2 in the incubation mixture in the absence of cell monolayer () and in the incubation mixture in the basal (black-triangle) and apical () wells in the presence of cell monolayer in Transwells were measured as described in MATERIALS AND METHODS. Results from a representative experiment are provided. B and C: transepithelial electrical resistance (TER; B) and mannitol flux (C) in Caco-2 cell monolayer incubated with () or without () XO+X. Values are means ± SE (n = 6). * P < 0.05 vs. corresponding values for monolayer incubated without XO+X.


                              
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Table 1.   Effect of superoxide dismutase and catalase on oxidative stress-induced increase in paracellular permeability

Coadministration of SOD (100 µg/ml) resulted in a significant potentiation of XO+X-induced changes in dilution potential and mannitol flux (Table 1). On the other hand, administration of catalase (10 U/ml) inhibited the effect of XO+X. SOD or catalase by themselves produced no significant effect on TER, dilution potential, or mannitol flux. SOD produced a slight but significant increase in H2O2 level, whereas the level was markedly reduced by catalase (Table 1).

Absence of a role for ·OH in H2O2-induced increase in permeability. Treatment with H2O2 (20 µM) significantly reduced TER and dilution potential of Caco-2 cell monolayer and increased mannitol permeability (Table 2). Pretreatment of cell monolayers with FeSO4 (0.1 mM) significantly inhibited H2O2-induced changes in TER, dilution potential, and mannitol flux (Table 2). On the other hand, pretreatment with deferoxamine (1 mM) or 1,10-phenanthroline (0.1 mM) significantly potentiated H2O2-induced changes in TER, dilution potential, and mannitol permeability. FeSO4, deferoxamine, and 1,10-phenanthroline by themselves produced no significant effect on permeability in the absence of H2O2. Vitamin E or vitamin A (·OH scavengers) showed no effect on permeability by themselves, nor did they influence H2O2-induced increase in permeability. In vitro incubation of H2O2 with FeSO4, but not 1,10-phenanthroline, vitamin E, or vitamin A, resulted in a dramatic reduction in H2O2 level (data not shown). However, the H2O2 level in the buffer incubating cell monolayers pretreated with FeSO4 was not significantly different from that in the buffer incubating cell monolayers without FeSO4 pretreatment (Table 2).

                              
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Table 2.   Effect of iron salts, iron chelators, and ·OH-scavengers on H2O2-induced paracellular permeability

Role of thiol oxidation in H2O2-induced increase in paracellular permeability. Pretreatment of cell monolayers with GSH (1.0 mM), NAC (0.1 mM), or DTT (0.5 mM) significantly inhibited H2O2-induced decrease in TER and increase in mannitol flux (Fig. 2). BCNU, an inhibitor of GSSG reductase, potentiated the effect of H2O2 on TER and mannitol flux. Interestingly, DEM, a sulfhydryl alkylator, prevents H2O2-induced changes in TER and mannitol flux. GSH, NAC, DTT, BCNU, or DEM by themselves produced no significant effect on TER or mannitol flux (data not shown). H2O2 levels in the buffer incubating cell monolayers pretreated with GSH, NAC, or DTT were not significantly different from that in the buffer incubating cell monolayers without pretreatment.


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Fig. 2.   Effect of thiol-modulating agents on H2O2-induced permeability. Cell monolayers were pretreated for 90 min with 1 mM GSH, 0.1 mM N-acetyl-L-cysteine (NAC), 0.5 mM dithiothreitol (DTT), 0.15 mM diethylmaleate (DEM), or 0.3 mM 1,3-bis(2-chloroethyl)-nitrosourea (BCNU) before administration of H2O2 (20 µM). Control cell monolayers received no H2O2 or thiol-modulating agents. TER (A) and mannitol flux (B) were measured at 120 min after H2O2 administration as described in MATERIALS AND METHODS. Values are means ± SE (n = 6). * P < 0.05 vs. corresponding values for control. # P < 0.05 vs. corresponding values for H2O2. In the absence of H2O2, GSH, NAC, DTT, DEM, or BCNU produced no significant effect on TER or mannitol flux.

Pretreatment of cell monolayers with mercaptosuccinate (0.1-10 mM), an inhibitor of Gpx, prevented H2O2-induced decrease in TER and the increase in mannitol flux in a concentration-related manner (Fig. 3). Aminotriazole (20 mM), an inhibitor of catalase, on the other hand, produced a slight but significant potentiation of the effect of H2O2 on TER and mannitol flux. Mercaptosuccinate or aminotriazole by themselves produced no significant effect on permeability of control cell monolayer (data not shown). H2O2 levels in the buffer incubating cell monolayers pretreated with mercaptosuccinate or aminotriazole were not significantly different from that in the buffer incubating cell monolayers without pretreatment.


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Fig. 3.   Effect of inhibitors of glutathione peroxidase (Gpx) and catalase on H2O2-induced permeability. Cell monolayers were pretreated for 90 min with 0.1 mM, 0.5 mM, 2 mM, or 10 mM mercaptosuccinate (MS, black-lozenge ), 20 mM aminotriazole (AT, triangle ), or buffer () before H2O2 administration. Control monolayers (open circle ) received no H2O2 or inhibitors. TER (A) was measured at varying times. Mannitol flux (B) was measured during 0-3 h time span. Values are means ± SE (n = 4 from 2 independent experiments). * P < 0.05 vs. control and # P < 0.05 vs. H2O2-treated monolayers.

GSH level in H2O2-treated Caco-2 cell monolayer was significantly lower than that in control cell monolayer (Fig. 4A). GSSG level, on the other hand, was significantly higher in H2O2-treated cells (Fig. 4B). Pretreatment with FeSO4 and mercaptosuccinate prevented H2O2-mediated changes in GSH and GSSG levels, whereas 1,10-phenanthroline and BCNU potentiated this effect of H2O2. Similar to GSH levels, GSH-to-GSSG ratio (Fig. 5A) and the levels of protein thiols (Fig. 5B) were also lower in H2O2-treated cells, which was prevented by FeSO4 and mercaptosuccinate and potentiated by 1,10-phenanthroline and BCNU. DEM markedly reduced cellular GSH levels (Fig. 4A) but prevented H2O2-mediated increase in GSSG (Fig. 4B) and decrease in protein thiols (Fig. 5B). DEM reduced GSH levels in control cell monolayers without altering the levels of GSSG or protein thiols (data not shown). Treatment with GSH or NAC markedly increased cellular GSH level (Fig. 4A) and protein thiols (Fig. 5B); H2O2-mediated GSSG formation was significantly increased (Fig. 4B).


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Fig. 4.   Effect of H2O2 on GSH and GSSG levels. Cell monolayers were pretreated with buffer, FeSO4 (0.1 mM), 1,10-phenanthroline (0.1 mM), mercaptosuccinate (2 mM), BCNU (0.3 mM), DEM (0.15 mM), GSH (1 mM), or NAC (0.1 mM) before administration of H2O2. After 3-h incubation, cell monolayers were homogenized in 5% 5-sulfosalicylic acid. GSH (A) and GSSG (B) levels were determined in acid-soluble extracts as described in MATERIALS AND METHODS. Values are means ± SE (n = 4). * P < 0.05 vs. control and # P < 0.05 vs. H2O2.



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Fig. 5.   Effect of H2O2 on GSH-to-GSSG ratio and protein thiols. Cell monolayers were treated with various compounds, and the levels of GSH and GSSG were measured as described for Fig. 4. GSH-to-GSSG ratio (A) was calculated from GSH and GSSG values in Fig. 4. Protein thiol (B) was measured in acid precipitates from the experiments described in Fig. 4. Values are means ± SE (n = 4). * P < 0.05 vs. control and # P < 0.05 vs. H2O2.

Inhibition of PTPase activity. In vitro incubation of soluble fraction of Caco-2 cell monolayer with GSSG (1-100 nM) resulted in a concentration-related inhibition of PTPase activity (Fig. 6A). Inhibition of PTPase activity by 30 nM GSSG was prevented by coadministration of GSH (30-300 nM) in a concentration-related manner. Incubation with either GSH or GSSG produced no significant effect on tyrosine kinase activity associated with the plasma membrane fraction (Fig. 6B). In vivo incubation of cell monolayers with H2O2 also resulted in a partial decrease in PTPase activity in both plasma membrane and soluble fractions (Fig. 7). Maximal inhibition was achieved by 20 min of XO+X treatment.


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Fig. 6.   Effect of GSSG on tyrosine kinase and PTPase activities in membrane and soluble fractions in vitro. Soluble and plasma membrane fractions were isolated from untreated cell monolayers and incubated with varying concentrations of GSSG (1-100 nM) with or without GSH (30-300 nM) at 37°C for 60 min. PTPase activity (A) in soluble fraction and tyrosine kinase activity (B) in membrane fractions were measured as described in MATERIALS AND METHODS. Values are means ± SE (n = 4 from 2 independent experiments). * P < 0.05 vs. untreated cell monolayers and # P < 0.05 vs. cell monolayer treated with 30 nM GSSG.



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Fig. 7.   Effect of XO+X on PTPase activity in vivo. Cell monolayers were incubated with XO+X for varying times, and plasma membrane and soluble fractions were isolated. PTPase activity was measured as described in MATERIALS AND METHODS. Values are means ± SE (n = 4). * P < 0.05 vs. corresponding values at 0 min.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

This study shows that H2O2, but not O2-· or ·OH, increases paracellular permeability in Caco-2 cell monolayer and that H2O2-induced increase in paracellular permeability is mediated by Gpx-dependent oxidation of GSH and GSSG-mediated inhibition of PTPase activity. We previously demonstrated that XO+X increases Caco-2 epithelial paracellular permeability (26), and the results of the present study provide evidence of the identity of the oxidant species responsible for this effect. Although it is relatively nonreactive, O2-· has been shown to induce a few biological effects and appears to play an important role in inflammation (7, 23). It may directly affect a biological system or induce its effect indirectly after conversion into peroxynitrite. The present study shows that SOD, the enzyme that catalyzes the conversion of O2-· to H2O2, failed to inhibit the XO+X-induced increase in paracellular permeability, suggesting that O2-· may not play a role in increasing the permeability. Catalase, however, markedly reduced H2O2 level and inhibited the XO+X-induced increase in permeability. The observation that H2O2 at 20 µM concentration (similar to that generated by XO+X) increases paracellular permeability to an extent similar to the XO+X-induced permeability supports the view that O2-· is not involved.

Although it is well established that ·OH is the major oxidant species involved in cell injury, our present study demonstrates that ·OH does not play a role in increasing the paracellular permeability in Caco-2 cell monolayer. FeSO4 inhibited H2O2-induced paracellular permeability, suggesting a rapid conversion of H2O2 to ·OH by Fe2+. In contrast, pretreatment of cell monolayer with deferoxamine or 1,10-phenanthroline (Fe chelators) resulted in a significant potentiation of H2O2-induced increase in paracellular permeability. The prevention of H2O2-induced increase in permeability by Fe2+ and potentiation by Fe chelators clearly demonstrate that ·OH does not increase paracellular permeability in Caco-2 cell monolayer. This conclusion of a lack of a role for ·OH in increasing paracellular permeability was further supported by the observation that vitamin E and vitamin A, ·OH scavengers, do not prevent H2O2-induced permeability. Therefore, H2O2 itself is responsible for increasing the paracellular permeability in Caco-2 cell monolayer.

Although it is usually thought that most or all of the toxicity of O2-· and H2O2 involves their conversion to ·OH, very little is known about the specific role of H2O2 itself in tissue injury. H2O2 at low micromolar levels is poorly reactive with biological systems (12). However, at higher concentration, H2O2 can inactivate glyceraldehyde-3-phosphate dehydrogenase, a glycolytic enzyme (21). Ginsburg et al. (9) showed in kidney epithelial cells that injury caused by xanthine oxidase in combination with a bacterial toxin was prevented by H2O2 scavengers but was unaffected by SOD or deferoxamine, suggesting that H2O2 was the specific oxidant species that was responsible for cell injury in their model. Another study showed that hepatocyte injury by H2O2 generated by glucose oxidase was not affected by N,N'-diphenyl-p-phenylenediamine, a scavenger of lipid peroxides, suggesting that lipid peroxidation (usually caused by ·OH) played no role in this cell injury; the effects of Fe2+ or Fe chelators were not tested. ·OH seems to play an important role in H2O2-induced cell injury in cultured gastric mucosal cells (17) and hepatocytes (27). Therefore, H2O2 and ·OH may play distinct roles in cell injury depending on the cell type, doses of oxidant, and duration of treatment.

The results of our present study also suggest that the mechanism of H2O2-induced increase in paracellular permeability may involve oxidation of GSH and protein thiols. H2O2-induced paracellular permeability was significantly inhibited by pretreatment of cell monolayers with thiol compounds, such as GSH, NAC, or DTT. Treatment with thiol compounds also showed increased levels of GSH and protein thiols in Caco-2 cell monolayer. This observation suggests that an elevation of intracellular thiols may protect the epithelium from H2O2, possibly by protecting the cellular protein thiols from oxidation. Direct evidence of H2O2-induced GSH oxidation in increasing paracellular permeability was provided by the effect of mercaptosuccinate, a Gpx inhibitor (3). Pretreatment of cell monolayer with mercaptosuccinate resulted in a concentration-related inhibition of H2O2-induced increase in paracellular permeability, suggesting that Gpx activity may be required in this effect of H2O2. On the other hand, treatment with BCNU, an inhibitor of GSSG reductase, potentiated H2O2-induced increase in permeability. In contrast to Gpx inhibitor, catalase inhibitor potentiated the effect of H2O2 on permeability. Inhibition of catalase may increase intracellular H2O2 level and exacerbate the effect on permeability. Inhibition of Gpx activity may also increase intracellular H2O2 level; however, this H2O2 cannot oxidize GSH to GSSG in the absence of Gpx activity. These findings support our suggestion that GSH oxidation and GSSG accumulation are crucial for H2O2-induced increase in permeability.

Treatment with H2O2 reduced the level of GSH and protein thiols. Decrease in GSH was accompanied by an increase in GSSG level. The amount of increase in GSSG did not account for the amount of decrease in GSH. However, decrease in protein thiols did account for most of GSH decrease. This observation suggests that GSSG may rapidly react with protein thiols to form mixed disulfides. This effect of H2O2 on GSH oxidation was prevented by FeSO4 and potentiated by 1,10-phenanthroline. Mercaptosuccinate prevented H2O2-induced oxidation of GSH and protein thiols, whereas BCNU potentiated this effect of H2O2. Additionally, pretreatment with DEM (a sulfhydryl alkylator) inhibited H2O2-induced decrease in TER. DEM produced no significant effect on TER in the absence of H2O2, but it markedly reduced cellular GSH levels. However, DEM prevented H2O2-induced generation of GSSG and depletion of protein thiols. These results suggest that generation of GSSG, rather than GSH depletion, is important in H2O2-induced permeability. Reduced level of GSH by alkylation may prevent Gpx-mediated metabolism of H2O2 and reduce GSSG accumulation.

In a previous study we demonstrated that H2O2-induced increase in paracellular permeability in Caco-2 cell monolayer is mediated by protein tyrosine phosphorylation (26). The present study demonstrates that H2O2 treatment results in a partial decrease in PTPase activity in Caco-2 cell monolayers. The inhibition of PTPase activity was rapid, with maximal inhibition achieved by 20 min, which corresponds well with the peak level of H2O2 generated by XO+X. Interestingly, the PTPase activity in soluble fractions of Caco-2 cells was also inhibited by in vitro incubation with GSSG, suggesting that GSSG may directly interact with PTPases. Inhibition of PTPases may contribute to the protein tyrosine phosphorylation induced by H2O2. These results indicate that elevation of intracellular GSSG is required for H2O2-induced increase in protein tyrosine phosphorylation and paracellular permeability. As outlined in Fig. 8, the level of GSSG attained may be regulated by the ratio of the activities of Gpx and GSSG reductase. The activity of Gpx in the Caco-2 cell is 35-fold greater than the activity of GSSG reductase (2). Elevated GSSG caused by Gpx-dependent H2O2 metabolism may mediate protein thiol oxidation in the cell. The protein thiol oxidation by H2O2 may result in inhibition of PTPases and/or activation of protein tyrosine kinases. This view is supported by previous observations that H2O2 can activate tyrosine kinase (14) and inhibit PTPase (15). The inhibition of PTPases may involve an oxidation of cysteine thiol of signature motif sequence at the active site of PTPases (29). Protection by GSH and other thiol compounds may have been caused by the prevention of GSSG-mediated protein thiol oxidation. In summary, this study shows that H2O2, but not O2-· or ·OH, increases the paracellular permeability in Caco-2 cell monolayer by a mechanism that may involve protein thiol oxidation and inhibition of PTPases.


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Fig. 8.   Proposed mechanism for H2O2-induced increases in protein tyrosine phosphorylation and paracellular permeability in Caco-2 cell monolayer.


    ACKNOWLEDGEMENTS

This study was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant R01-DK-55532-01 (to R. K. Rao).


    FOOTNOTES

Address for reprint requests and other correspondence: R. K. Rao, Dept. of Pediatrics, Medical Univ. of South Carolina, 158 Rutledge Ave., Charleston, SC 29403.

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. §1734 solely to indicate this fact.

Received 24 June 1999; accepted in final form 6 March 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Am J Physiol Gastrointest Liver Physiol 279(2):G332-G340
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