1Center for Cardiovascular Science, Albany Medical College, and 2Research Service, Stratton Veterans Affairs Medical Center, Albany, New York 1228
Submitted 16 April 2003 ; accepted in final form 5 June 2003
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ABSTRACT |
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antisense; edema; mRNA; permeability; transcription
NAD(P)H oxidase, recently explored in endothelium, is an enzyme that produces superoxide anion () via the reaction 2O2 + NAD(P)H
+ NADP+ + H+ (2123, 32). The NAD(P)H oxidase enzyme complex is composed of a membrane-bound flavocytochrome b558 and the cytosolic activating factors Rac-2, p40phox, p47phox, and p67phox (2123, 32). The membrane-bound flavocytochrome b558 consists of two heme redox centers and the subcomponents p22phox and gp91phox. The translocation of Rac-2, p40phox, p47phox, and p67phox to the flavocytochrome b558 is generally considered the signature event for activation of NAD(P)H oxidase and the subsequent generation of
(2123, 32). The activation of NAD(P)H oxidase in response to TNF is theorized to occur by an increase in PKC-mediated phosphorylation of p47phox, which causes its translocation to the peripheral membrane (2123, 32). This notion is supported by our previous work that indicates the TNF-induced increase in pulmonary endothelial permeability is dependent on PKC-
, a potential activator of NAD(P)H oxidase (8). However, the role of NAD(P)H oxidase-dependent generation of
in the TNF-induced pulmonary microvessel endothelial barrier dysfunction is not known. Therefore, we hypothesize that the increased endothelial permeability in response to TNF is mediated by NAD(P)H oxidase-dependent generation of
.
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MATERIALS AND METHODS |
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Bovine lung microvessel endothelial cells (BLMVEC) derived from fresh calf lungs were obtained at 4th passage (Vec Technologies, Rensselaer, NY). The preparations were identified by Vec Technologies as pure populations by 1) the characteristic "cobblestone" appearance as assessed by phase contrast microscopy, 2) the presence of factor VIII-related antigen (indirect immunofluorescence), 3) the uptake of acylated low-density lipoproteins, and 4) the absence of smooth muscle actin (indirect immunofluorescence).
For all studies, BLMVEC were cultured from 4 to 12 passages (2, 8) in medium containing Dulbecco's modified Eagle's Medium (DMEM; GIBCO-BRL, Grand Island, NY) supplemented with 20% fetal bovine serum (FBS; Hyclone Laboratories, Logan, UT), 15 µg/ml endothelial cell growth supplement (Upstate Biotechnology, Lake Placid, NY), and 1% nonessential amino acids (GIBCO-BRL) and maintained in 5% CO2 plus humidified air at 37°C. A confluent pulmonary microvessel endothelial monolayer (PMEM) was reached within two to three population doublings, which took 34 days.
Treatments
TNF treatment. Highly purified recombinant human TNF from Escherichia coli (Calbiochem-Novabiochem, La Jolla, CA) in a stock solution of 10 µg/ml was used. The endotoxin level was <0.1 ng/µg of TNF as determined by standard limulus assay (2, 8). We previously showed that boiling TNF for 0.75 h blocks the effect of TNF in our system (6, 7, 16, 18), which indicates no endotoxin contamination. A TNF dose of 50 ng/ml was used throughout this study because our previous studies indicated that this dose causes a PKC-- and ·NO-dependent increase in permeability of PMEM (2, 8).
SOD. SOD (Sigma, St. Louis, MO), an enzyme that catalyzes the dismutation of superoxide radicals to hydrogen peroxide (H2O2) and molecular oxygen (7, 18, 29), was used to verify the role of . PMEM were pretreated with SOD (100 U/ml) 0.25 h before its co-incubation with TNF. We previously showed that heat-inactivated SOD (autoclaving SOD for 0.75 h) blocked its protective effects, which verified the antioxidant specificity of SOD (7, 18, 27).
4,5-Dihydroxy-1,3-benzenedisulfonic acid. 4,5-Dihydroxy-1,3-benzenedisulfonic acid (Tiron, Sigma), a cell-permeable superoxide trap, was also used to verify -mediated alterations by TNF. We previously showed that Tiron prevents the TNF-induced latent decrease in ·NO and increase in peroxides in response to TNF (18). PMEM were pretreated with Tiron (1 mM) 0.25 h before its co-incubation with TNF.
Treatment medium. For all studies, incubation of PMEM with TNF and corresponding controls was performed with phenol-free DMEM (pf-DMEM, GIBCO-BRL) supplemented with 10% FBS to avoid the potential antioxidant effect of phenol as well as its spectrophotometric interference.
Antisense Oligonucleotides
p22phox antisense. Expression of p22phox was inhibited with a p22phox antisense oligonucleotide complementary to bovine p22phox nucleotides 1,1651,189 (25). The antisense and a corresponding scrambled-nonsense oligonucleotide sequences were 5'-CCG-CGTGTC-GAG-GTC-CAT-GCA-GAC-C-3' and 5'-GAC-GAC-GTGACT GAC-CTG-GTG-CCG-C-3', respectively. Both oligonucleotides were second generation chimeras, consisting of phosphorothioated DNA (9 nucleotides) and 2'-O-methylated RNA (16 nucleotides) (Oligos Etc., Wilsonville, OR), to achieve improved specificity of binding to mRNA and nuclease resistance.
Transfection of oligonucleotides in PMEM. Transfection of oligonucleotides was accomplished with the cationic liposome carrier Lipofectin (GIBCO-BRL) by our standard technique (2, 8). Oligonucleotides from a 40-µM stock solution were added to serum-free DMEM containing Lipofectin (6.6 µl/ml) for a final concentration of 200 nM, then added to PMEM, and incubated at 37°C for 4.0 h. The Lipofectin/oligonucleotide-containing medium was removed before experimental treatment.
Detection of p47phox Protein by Immunoblot
Purification of endothelial luminal plasma membranes. To determine subcellular distribution of p47phox in response to TNF treatment, we isolated endothelial cell luminal plasma membranes as previously described (24, 31). After the 4.0-h stimulation with TNF, we rapidly cooled cells to 4°C by rinsing them with ice-cold MES-buffered saline (MBS, 20 mM, pH 6.0) to stabilize enzymatic activity and diffusion of molecules within the lipid bilayer of the plasma membrane. Subsequently, the cells were incubated with a positively charged colloidal silica solution for 10 min at 4°C. Subsequent cross-linking of the silica particles by incubation with polyacrylic acid (0.1%) serves to create a stable adherent silica pellicle that permits its purification by centrifugation to separate the silica-coated endothelial cell plasma membranes from the whole cell homogenate. Silica-coated endothelial cell monolayers were scraped and pooled in 1 ml of HEPES-buffered sucrose (HBS) containing a cocktail of protease inhibitors [0.25 M sucrose, 25 mM HEPES at pH 7.0, 10 µg/ml leupeptin, 10 µg/ml pepstatin A, 10 µg/ml o-phenanthroline, 10 µg/ml 4-(2-aminoethyl) benzenesulfonyl fluoride, and 50 µg/ml trans-epoxysuccinyl-L-leucinamido (4-guanidino) butane]. After sample homogenization, cell homogenates were mixed with 102% (wt/vol) Nycodenz (Life Sciences) with 20 mM KCl to make a 50% final solution. The Nycodenzhomogenate solution was layered over a continuous 5570% Nycodenz gradient containing 20 mM KCl and HBS and centrifuged in a Beckman SW 55 rotor at 15,000 rpm for 0.5 h at 4°C. The resulting plasma membrane pellet was resuspended in 0.5 ml of MBS.
Western analysis. Protein concentrations of the whole cell homogenate or purified luminal membrane fractions were determined by bicinchoninic acid analysis (Pierce). Equivalent amounts of protein from each sample were prepared and separated by SDS-PAGE (10% gels) followed by electrotransfer to nitrocellulose filters. Immunoblotting with antibodies against p47phox (Santa Cruz) served to detect p47phox protein present in each cell subfraction. Briefly, filters were incubated in blocking solution (Tris-buffered saline, 0.5% Tween 20, 5% nonfat dry milk) for 1.0 h, followed by a 2.0-h incubation in primary antibody diluted in blocking buffer. After extensive washing, membranes were incubated for 1.0 h with anti-rabbit horseradish peroxidase. Membranes were extensively washed, and p47phox signal was detected with enhanced chemiluminescence substrate (ECL, Amersham). Western blots were scanned and digitized (Molecular Dynamics, Sunnyvale, CA). Densitometric quantification of immunoblots with Imagequant software (Molecular Dynamics) enabled direct comparisons between control and TNF-stimulated cells. We normalized results by arbitrarily setting the densitometry of control cells to 1.0.
Assay of p22phox and p47phox by Immunofluorescence Cell Staining
BLMVEC (1 x 104/0.20 ml of culture medium) were plated on 18-mm coverslips inside a 35-mm culture dish, incubated at 37°C for 2 h to allow attachment, and then grown to confluence in an additional 2 ml of culture medium. The PMEM were treated as indicated, washed with Dulbecco's phosphate-buffered saline without ions (DPBS, GIBCO-BRL), fixed with 3.7% formaldehyde solution at room temperature (RT) for 20 min, and then permeabilized with 1% Triton X-100 (Sigma) in DPBS at RT for 5 min. The cells were washed with DPBS and then blocked in 10% normal goat serum (NGS, GIBCO-BRL) at RT for 1 h. PMEM were incubated with either a rabbit polyclonal anti-human p22phox peptide antibody or a rabbit polyclonal anti-bovine p47phox antibody (provided by Mark Quinn, Marsh Laboratory, Bozeman, MT), at a 1:1,000 dilution in 10% NGS at RT for 1 h then washed sufficiently. The secondary antibody, Alexa Fluor 488-labeled goat anti-rabbit IgG (Molecular Probes) was added at a 1:1,000 dilution in 10% NGS and incubated at RT for 1 h and then washed sufficiently. The nuclei were stained with 2.5 µg/ml propidium iodide (Molecular Probes) at RT for 5 min then washed 1x with DPBS. The coverslips were mounted on clean glass slides with Permafluor mounting media (Thermo Shandon, Pittsburgh, PA). The PMEM were visualized with a Spot RT color camera (Diagnostic Instruments, Sterling Heights, MI) mounted on an Olympus IX70 inverted microscope (Olympus America, Melville, NY) equipped for phase, light, and fluorescence detection. Images were captured at x100 magnification with an exposure time of 8 s and downloaded into Spot RT imaging software (Diagnostic Instruments).
The quantification strategy for the fluorescent images is as follows. PMEM were visualized and quantified with confocal microscopy with the Leica Confocal System TCS SP2 (Leica Microsystems, Exton, PA). There were four separate studies with six treatment groups per study. All fields were selected by random movement of the microscope stage to another area within an intact endothelial monolayer. Six entire fields per treatment group were analyzed with one image per field. We normalized all treatment groups for fluorescent intensity by initially adjusting the settings for noise, brightness, and contrast, as determined by the slide with the maximum fluorescence.
Detection of Oxidant Generation
The oxidation of 5 (and 6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate-acetyl ester (CM-H2DCFDA; Molecular Probes, Eugene, OR) or 6-carboxy-2',7'-dichlorodihydrofluorescein diacetate-di(acetoxymethyl ester) (C-H2DCFDA, Molecular Probes) was used to assess intracellular reactive oxygen species (ROS) (5, 9, 9a). Cleavage of the ester groups by intracellular esterases and oxidation by ROS results in intracellular dichlorofluorescein (DCF) derivatives, which are highly fluorescent. In addition, the oxidation of dihydroethidium (DHE, Molecular Probes) to ethidium was used to specifically assess . Upon oxidation, ethidium intercalates within the cell's DNA, becoming brightly fluorescent, and is considered a relatively specific measurement of
(5, 9, 9a).
Fluorescence detection of ROS in intact PMEM. DCF fluorescence was measured in a Wallac 1420 Victor2 multilabel counter (Wallac, Turku, Finland) using excitation and emission wavelengths of 490 and 535 nm, respectively. BLMVEC (1 x 104/0.20 ml of culture medium) were plated in 96-well microplates and grown to confluence. Before treatment, PMEM were washed 2x with Hanks' buffered saline solution (HBSS, GIBCO-BRL), loaded with 20 µM CM-H2DCFDA (in HBSS 100 µl/well), and incubated at 37°C for 0.5 h. PMEM were then washed 1x with HBSS and 1x with pf-DMEM + 10% FBS to remove free probe. Then, fresh pf-DMEM + 10% FBS (100 µl/well) was added, and a baseline fluorescence reading was taken before treatment. Fluorescence measurements were taken at 0.0, 0.5, 1.0, 2.0, 3.0, and 4.0 h during the treatments. Results are presented as percent change from baseline by the formula [(Ftexp - Ftbase)/Ftbase x 100], where Ftexp = fluorescence at any given time during the experiment in a given well and Ftbase = baseline fluorescence of the same well. The difference in DCF fluorescence generated in the presence and absence of SOD or Tiron was used as a further verification of -dependent pathways (7, 18, 27).
Fluorescence detection of in PMEM lysate. BLMVEC (1 x 105 /2.0 ml of culture medium) were plated in six-well culture dishes and, upon reaching confluence, were treated as indicated. After treatment, PMEM were washed 1x with pf-DMEM + 2% FBS and incubated with 10 µM DHE in pf-DMEM + 2% FBS (1 ml/well) at 37°C for 0.5 h. PMEM were then washed on ice 2x with ice-cold DPBS to remove free probe and scraped with 1 ml of ice-cold DPBS into microtubes. The cells were centrifuged at 8,000 g for 10 min at 4°C, and the pellet was resuspended in 0.5 ml of ice-cold DPBS. The cell suspensions were sonicated on ice for 15 s, and 100 µl/well sonicate were added to Falcon 96-well black microplates (Becton Dickinson, Franklin Lakes, NJ) in quadruplicate. Probe-free cell sonicate was used for blanks. Ethidium fluorescence was measured in a Wallac 1420 Victor2 multilabel counter (Wallac) using excitation and emission wavelengths of 490 and 605 nm, respectively. Fluorescence was presented as percentage of control by the formula [Ftexp/Ftcontrol], where Ftexp = fluorescence at any time after treatment in a given lysate and Ftcontrol = fluorescence of the respective untreated control group.
Microscopic detection of ROS-generated fluorescence in intact PMEM. BLMVEC (1 x 105/2.0 ml of culture medium) were plated in 35-mm dishes and, upon reaching confluence, were treated as indicated. We performed detection of ROS by incubating the PMEM with 1.0 µM C-H2DCFDA diluted in pf-DMEM + 2% FBS at 37°C for 20 min. PMEM were then washed once in DPBS, fixed in a 3.7% formaldehyde solution for 20 min at RT, and then washed twice with DPBS. The PMEM fluorescence was visualized with a Spot RT color camera mounted on an Olympus IX70 inverted microscope. Images were captured at x20 magnification with an exposure time of 8 s and downloaded into Spot RT imaging software. Immunofluorescence was quantified by computerized densitometry with Sigma Scan Pro (SPSS Scientific Software, San Rafael, CA). There were four separate studies with six treatment groups per study. All fields were selected by random movement of the microscope stage to another area within an intact endothelial monolayer. Six entire fields per treatment group were analyzed with one image per field.
Assay of Endothelial Permeability
Nuclepore Track-Etch Polycarbonate Membranes (13-mm diameter, 0.8-µm pore size; Corning Costar, Cambridge, MA) were coated with gelatin (type B from bovine skin; Sigma) as previously described (2, 7), mounted on modified Boyden chemotaxis chambers (9-mm inner diameter; Adaps, Dedham, MA) with MF cement no. 1 (Millipore, Bedford, MA), and sterilized by ultraviolet light for 12.024.0 h. BLMVEC (1 x 105/0.50 ml of DMEM) were plated on the gelatinized membranes and allowed to reach confluence.
The experimental apparatus for the study of transendothelial transport in the absence of hydrostatic and oncotic pressure gradients has been described (2, 8, 26). In brief, the system consists of two compartments separated by a microporous polycarbonate membrane lined with the endothelial cell monolayer as described above. The upper, luminal compartment (700 µl) was suspended in the lower, abluminal compartment (25 ml). The abluminal compartment was stirred continuously for complete mixing. The entire system was kept in a water bath at a constant temperature of 37°C. The fluid height in both compartments was the same to eliminate convective flux.
Endothelial permeability was characterized by the clearance rate of Evans blue-labeled albumin as previously described (2, 6, 8). A buffer solution containing HBSS (GIBCO-BRL), 0.5% bovine serum albumin (Sigma), and 20 mM HEPES (Sigma) was used on both sides of the monolayer. The luminal compartment buffer was labeled with a final concentration of 0.057% Evans blue dye in a volume of 700 µl. The absorbance of free Evans blue dye in the luminal and abluminal compartments was always <1% of the total absorbance of Evans blue in the buffer. At the beginning of each study, a luminal compartment sample was diluted 1:100 to determine the initial absorbance of that compartment. Abluminal compartment samples (300 µl) were taken every 5 min for 1.0 h. The absorbance of the samples was measured in a SpectraMax Plus microplate spectrophotometer (Molecular Devices, Sunnyvale, CA) at 620 nm. The clearance rate of Evans blue-labeled albumin was determined by least-squares linear regression between 10 and 60 min for the control and experimental groups.
Assay of Cell Viability
Trypan blue exclusion. BLMVEC (1 x 105/2.0 ml of culture medium) were seeded in 35-mm dishes and grown until confluent. After respective drug treatments, PMEM were washed with DPBS and followed by treatment with 0.05% trypsin (0.2 ml) for 1.0 min at 37°C. The cells were resuspended in DPBS (1 ml), and an aliquot of cell suspension (50 µl) was combined with 0.08% trypan blue (50 µl) for 3 min. Cells were counted in 10 µl of the mixture with a hemocytometer. Cell viability was defined by the following formula: Cell viability = (cells excluding trypan blue/total cells) x 100 (10).
Statistics
A one-way analysis of variance was used to compare values among the treatment groups. If significance was noted, a Bonferroni multiple-comparison test was used to determine significant differences between the groups. A t-test was used when appropriate. Each PMEM well and flask represent a single experiment. There were 510 samples (n) per group, in which each sample was from a separate study, unless noted otherwise. All data are reported as means ± SE. Significance was at P < 0.05.
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RESULTS |
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We previously showed that TNF, SOD, and Tiron do not affect cell viability (2, 6, 8, 10, 27). In the present study, treatment with p22phox oligonucleotides ± TNF (50 ng/ml) did not affect PMEM viability. The trypan blue exclusion was similar among the groups (i.e., control: 98.5 ± 0.5% vs. p22phox antisense: 97.5 ± 0.5% vs. p22phox antisense + TNF: 97.0 ± 0.5% vs. scrambled: 98.5 ± 0.5% vs. scrambled + TNF: 99.0 ± 0.5%).
TNF Induces an Increase in p22phox That Is Prevented by p22phox Antisense
In the representative study of Fig. 1, PMEM were incubated with TNF for 4.0 h both with and without pretreatment with either antisense oligonucleotides to p22phox or scrambled oligonucleotides and then assayed for p22phox fluorescence. We measured the p22phox protein to show that the antisense to p22phox can prevent the upregulation of p22phox protein during the response to TNF. In PMEM treated with TNF alone or scrambled oligonucleotides + TNF, there was an increase in p22phox fluorescence (green) compared with their respective controls; note the peripheral localization of p22phox fluorescence in these groups. In PMEM treated with anti-p22phox + TNF, there was no increase in p22phox fluorescence compared with anti-p22phox controls, nor was there notable peripheral localization of p22phox fluorescence in these groups. The nuclear morphology (red) was similar among all the groups. There was no significant fluorescence in the TNF group stained with only secondary antibody and without propidium iodide. In Fig. 2, quantification of the digitized images indicates that TNF induced an increase in total cellular p22phox fluorescence that was prevented by anti-p22phox, whereas the scrambled oligonucleotide had no effect. The data from Figs. 1 and 2 show that TNF induces an increase in p22phox protein. The increase in p22phox protein and the peripheral localization of the p22phox protein were specifically prevented by anti-p22phox.
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TNF Induces Activation of NAD(P)H Oxidase
PMEM were incubated with or without TNF for 4.0 h (Fig. 3). Total cell lysate and membrane fractions were assayed for p47phox protein by Western immunoblot. The intracellular distribution of p47phox protein was measured because increased localization of p47phox in the peripheral membrane is a signature event for NAD(P)H oxidase activation. The representative (n = 2) Western blot shows TNF increased the membrane p47phox/total p47phox ratios compared with the control group. We assayed the immunofluorescence of p47phox protein to further explore the NAD(P)H oxidase activation in response to TNF. In the representative micrographs of Fig. 3B, the control group shows faint p47phox fluorescence (green) diffusely distributed within the cell. In the TNF and scrambled + TNF groups, there are increases in intensity of p47phox fluorescence compared with the respective control and scrambled groups. In the anti-p22phox + TNF group, there is no increase in p47phox fluorescence compared with the anti-p22phox group. The nucleus (red) does not exhibit any significant change in morphology among the groups. PMEM treated with only secondary antibody reveals no significant fluorescence in any group. In Fig. 4, quantification of the digitized images indicates that TNF induced an increase in total cellular p47phox fluorescence that was prevented by anti-p22phox, whereas the scrambled oligonucleotide had no effect.
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The data of Figs. 3 and 4 demonstrate that TNF treatment causes increased localization of p47phox to the peripheral membrane and indicates, therefore, NAD(P)H oxidase activation in response to TNF. In addition, digital analysis of the images indicates that TNF induced an increase in p47phox protein that was specifically prevented by anti-p22phox.
p22phox Antisense Prevents the TNF-Induced Increase in Colocalization of p47phox with p22phox
Figures 5 and 6 show the apical-to-basal p22phox and p47phox fluorescence along the z-axis to further demonstrate the effect of TNF on the distribution of p22phox and p47phox protein. In Fig. 5A, the micrograph indicates there was greater p22phox fluorescence in the TNF group compared with the control group. Moreover, in the control group, the p22phox fluorescence was primarily between the cells, whereas the p22phox fluorescence was more diffuse in the TNF group. In Fig. 5B, the micrograph indicates there was greater p47phox fluorescence in the TNF group compared with the control group. Moreover, in the TNF group, there was greater p47phox fluorescence between the cells compared with the diffuse fluorescence in the control group. In Fig. 5C, the increase in p22phox and p47phox fluorescence was maximally localized to similar z-fractions in the TNF group, indicating increased localization of p47phox with p22phox in response to TNF.
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Figure 6, A and B, shows the comparisons of z-axis distribution for p47phox and p22phox fluorescence to assess the effect of the anti-p22phox on the response to TNF. Figure 6A indicates that in the anti-p22phox + TNF group, there was no change in localization of p47phox with p22phox compared with the anti-p22phox group. Figure 6B indicates that in the scrambled + TNF group, the localization of p47phox with p22phox increased compared with the scrambled group, a result similar to the TNF group. The results of Figs. 5 and 6 indicate that TNF induces an increase in localization of p47phox with p22phox in PMEM. Moreover, the data indicate that p22phox antisense specifically prevents the TNF-induced increase in localization of p47phox with p22phox.
p22phox Mediates Generation of ROS in Response to TNF
In Fig. 7, PMEM grown in 96-well microplates were assessed for the inhibited DCF fluorescence generated from CM-H2DCFDA during a 4-h incubation with TNF to indicate the temporal generation of ROS. In separate studies, PMEM were pretreated with SOD before TNF incubation to verify that the DCF fluorescence was dependent on the generation of . In the SOD-inhibited control group, there was no significant change in fluorescence during the 4.0-h study period. In the SOD-inhibited TNF group, there was a time-dependent increase in DCF fluorescence, which was significantly greater at 4.0 h than both its baseline value and the corresponding 4.0-h value in the SOD-inhibited control group. PMEM were also pretreated with the cell-permeable antioxidant Tiron for 0.25 h before TNF incubation to verify that the DCF fluorescence was indeed sensitive to changes in
. In the Tiron-inhibited control group, there was an increase in DCF fluorescence, which became significant at 4.0 h compared with baseline. In the Tiron-inhibited TNF group, there was an increase in DCF fluorescence that became significant at 2.0 and 4.0 h compared with baseline and the corresponding 4.0-h value in the Tiron-inhibited control and SOD-inhibited TNF groups. Thus Fig. 7 indicates that TNF causes the generation of ROS, which is inhibited by the anti-
agents SOD and Tiron.
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Figure 8A shows a representative micrograph, and Fig. 8B shows the mean pixels of the digitized images using DCF on fixed PMEM, to verify that the SOD-inhibited fluorescence was not specific to any variant of the DCF assay condition. In the TNF group, there was an increase in fluorescence at 4.0 h compared with the control group. In the SOD + TNF group, the fluorescence was the same as the SOD group and less than the TNF group. In addition, some PMEM were pretreated with antisense to p22phox or the scrambled oligonucleotide before the TNF incubation to show that the NAD(P)H oxidase system mediates the increase in fluorescence in response to the TNF. In the anti-p22phox + TNF group, the fluorescence was the same as the anti-p22phox group and less than the TNF group. The scrambled oligonucleotide had no effect on the TNF-induced increase in fluorescence. The data from Fig. 8 verify that the TNF-induced increase in ROS 1) is dependent on , 2) is consistent between different assays using DCF, and 3) is mediated by NAD(P)H oxidase.
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Figure 9 shows PMEM incubated with vehicle or TNF (50 ng/ml) for 4.0 h and assayed with the specific fluorescent probe DHE to verify
-dependent generation of ROS. In separate studies, PMEM were pretreated with SOD for 0.25 h before the TNF incubation to further verify that the DHE fluorescence was due to the generation of
. Also, some PMEM were pretreated with antisense to p22phox or the scrambled oligonucleotide before the TNF incubation to show that the NAD(P)H oxidase system mediates the increase in fluorescence in response to the TNF. In the TNF group, there was an increase in DHE fluorescence compared with the control group. In the SOD + TNF group, the DHE fluorescence was similar to the SOD group and less than in the TNF group. In the anti-p22phox + TNF group, the fluorescence was similar to the anti-p22phox group and less than in the TNF group. The scrambled oligonucleotide had no effect on the TNF-induced increase in fluorescence. The results shown in Fig. 9 confirm that TNF causes the p22phox-dependent generation of
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TNF-Induced Increase in Permeability Is Mediated by NAD(P)H Oxidase and
As shown in Fig. 10, the PMEM were incubated with vehicle or TNF (50 ng/ml) for 4.0 h. Some PMEM were pretreated with antisense to p22phox or the scrambled oligonucleotides for 4.0 h before TNF incubation, with the purpose of showing that the NAD(P)H oxidase system mediates the barrier dysfunction in response to the TNF. The permeability of Evans blue albumin was assayed and is shown in Fig. 10. Albumin flux was increased in the TNF and scrambled + TNF groups compared with the respective control and scrambled groups. In the anti-p22phox + TNF group, there was no increase in albumin flux compared with the control and anti-p22phox group. In separate studies, PMEM were incubated with Cu-Zn SOD (100 U/ml) throughout the TNF incubation to show that the increased permeability is due to the generation of in response to TNF. The albumin clearance rate increased in the TNF group; however, there was no increase in the albumin clearance rate in the Cu-Zn SOD-treated group. The data indicate that the activation of NAD(P)H oxidase, via
, mediates the pulmonary endothelial barrier dysfunction induced by TNF.
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DISCUSSION |
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The movement of p40phox, 47phox, p67phox, and Rac from cytosol to membrane and their formation with membrane-bound p22phox and gp91phox are generally considered the events required for activation of the NAD(P)H oxidase (2123, 32). The translocation of p47phox into the cell membrane compartment is considered a signature for the activation of NAD(P)H oxidase (2123, 32). The present study indicates that TNF induced the activation of the NAD(P)H oxidase because the Western blot analysis showed translocation of p47phox to the peripheral luminal membrane. Moreover, the confocal microscopy showed increased localization of p47phox with p22phox in response to TNF. Finally, the immunofluorescent images of p22phox and p47phox indicate that the membrane-associated p22phox is more apparent than the membrane-associated p47phox. The compartmental regulation of p47phox and p22phox may be entirely different because p47phox normally resides in both cytoplasm and membrane, whereas p22phox primarily resides in membrane (2123, 32). Thus an overall increase in p47phox will be manifested as increases in both cytoplasm and membrane as opposed to the overall increase in p22phox, which will be manifested as an increase mostly in the membrane. The antisense to p22phox prevented the 1) peripheral localization of p22phox, 2) TNF-induced increase in p22phox, and 3) increased localization of p47phox with p22phox. The prevention of the increased localization of p47phox with p22phox by antisense to p22phox is consistent with the theory that membrane p22phox is essential for the integration of p47phox into the NAD(P)H oxidase complex (21). Our data agree with other studies indicating that TNF activates the NAD(P)H oxidase pathway in pulmonary endothelium (5, 9) and that NAD(P)H oxidase activity can be inhibited by antisense to p22phox (3, 25).
TNF caused generation of because the anti-
agents SOD and/or Tiron inhibited the TNF-induced increase in fluorescence of CM-H2DCFDA, C-H2DCFDA, and DHE. Moreover, the effect of the TNF was independent of our
assay conditions because the TNF-induced increase in
was detected with intact PMEM, fixed PMEM, and PMEM lysate. Finally, DHE is a more specific indicator of
than CMH2DCFDA and C-H2DCFDA (25). NAD(P)H oxidase mediated the TNF-induced increase in
because antisense to p22phox prevented the TNF-induced increase in fluorescence of C-H2DCFDA and DHE, and the scrambled oligonucleotide had no effect. Also, SOD prevented the TNF-induced increase in fluorescence of C-H2DCFDA and DHE, which is consistent with the notion that SOD is a "
sink" that binds the
generated by the membrane NAD(P)H oxidase because SOD is not cell permeable (27). Yet, our data do not rule out the fact that other pathways such as xanthine oxidase (33), eNOS (29), and mitochondrial enzymes (13) can mediate generation of ROS (e.g.,
, H2O2, ·OH) in response to TNF because Tiron, which is cell permeable, had a greater inhibitory effect than SOD on CM-H2DCFDA fluorescence, and CM-H2DCFDA may not be completely specific for ·O2 (27, 29).
The treatment with antisense to p22phox did not decrease constitutive p22phox protein, but it did prevent the TNF-induced increase in p22phox protein. We used a dose and time of antisense treatment that would only prevent the TNF-induced increase in p22phox protein to avoid parenthetic adaptive effects that may occur in response to decreases in an essential constitutive protein. Moreover, we wanted to study the role of the NAD(P)H oxidase only during the response to TNF. We used this approach in our previous study using anti-PKC- to deplete PKC-
(8). We previously showed that TNF caused a prolonged increase in PKC-
protein (8), which supported the endothelial barrier dysfunction in response to TNF, a result similar to our present conclusion about p22phox. De Keulenaer et al. (3) showed in smooth muscle that TNF causes an increase in p22phox mRNA. Interestingly, treatment with antisense to p22phox also prevented the TNF-induced increase in p47phox protein. The mechanism for the increase in p22phox and p47phox protein is not a focus of the present study; however, it has been shown that ROS mediate the activation of transcription factors (e.g., AP-1) and gene expression in response to TNF (5, 10, 30). In addition, a preliminary study (n = 2, data not shown) indicates that treatment of PMEM with SOD prevents the TNF-induced increase in p47phox protein, implicating
in the regulation of p47phox protein. Therefore, antisense to p22phox may have prevented the increase in p47phox protein by preventing the generation of
and the resultant downstream effect of ROS on gene and protein expression (5, 10, 30).
The mechanism for the activation of NAD(P)H oxidase in response to TNF was not investigated in the present study. We previously indicated that both PKC and/or PKC- activation occurs in response to TNF and mediates the 1) latent
-mediated decrease in ·NO (18), 2) increase in ONOO- (27), and 3) increase in glutathione oxidation (27). Frey et al. (9) showed that PKC-
regulates TNF-induced activation of NAD(P)H oxidase in endothelial cells. In response to TNF, the PKC-mediated phosphorylation of the NAD(P)H oxidase components such as p47phox can induce the translocation of p47phox to the peripheral membrane and the formation of the NAD(P)H oxidase enzyme complex (3, 9). Thus probable downstream targets for PKC activation are pathways leading to generation of ROS via NAD(P)H oxidase. Finally, other mechanisms for NAD(P)H activation in response to TNF can include activation of the mitogen-activated protein kinase pathways (11, 25).
The present results show that TNF caused an increase in protein permeability, which is consistent with our previous studies (2, 6, 8). The compartmentalization of , via NAD(P)H oxidase, within the cell peripheral membrane is a primary event responsible for the barrier dysfunction in response to TNF, because p22phox antisense and SOD completely blocked the increase in permeability. In separate ongoing studies of ours (n = 3), antisense to p47phox also prevented the TNF-induced increase in permeability (control: 0.274 ± 0.021
anti-p47phox + TNF 0.320 ± 0.013 µl/min; control: 0.274 ± 0.021 << TNF: 0.392 ± 0.018, scrambled p47phox + TNF: 0.379 ± 0.031 µl/min; P < 0.05). Our previous work indicates that TNF-induced barrier dysfunction is mediated by PKC-
(8), eNOS (2), and ·NO (6); in addition, our present study finds that NAD(P)H oxidase and
mediate barrier dysfunction. Moreover, our previous studies (6, 27) and the literature (29) indicate a role for ONOO- in pulmonary endothelial injury, which is a paradigm that integrates the mediators ·NO and
. ·NO and
can react with each other at a diffusion-limited rate to form the potent reactive oxygen/nitrogen molecule ONOO- (29). ONOO- is known to cause endothelial injury and to increase endothelial permeability (29). The role of ONOO- in TNF-induced endothelial injury is now under intense investigation in our laboratory.
It is known that reactive nitrogen (e.g., ·NO, ONOO-) and oxygen (, H2O2, and ·OH) species mediate alterations in the cytoskeleton and extracellular matrix that may ultimately have an impact on the barrier dysfunction in response to TNF. There are pathways that link reactive nitrogen and oxygen species with the cytoskeleton and permeability, such as direct oxidation and nitration of cytoskeletal protein (e.g., actin, tubulin; 1, 3, 4, 6), modulation of kinase pathways (e.g.., PKC, ERK, Src, and Rho; 11, 19, 25, 36), reorganization of cytoskeletal proteins (e.g., actin; 12, 28), and increases in protease activity (e.g., metalloproteases; 26). The interpretation of the z-axis images of p22phox and p47phox integrated with the increase in p47phox in the luminal membrane suggests that the TNF-induced increase in localization of p47phox with p22phox occurs, at least in part, at the apical membrane within the boundaries of the cell-cell interface. The cell-cell interface is an area in which cell-cell adhesion occurs. Actin, in association with zonula occludens protein ZO-1, claudins, occludins, and zonula adherence proteins catenins and cadherins, is a primary component of the mechanism for cell-cell adhesion (11, 19, 25, 36). Therefore, the results support the notion that a potential mechanism for the
-induced increase in permeability is the generation of ONOO- followed by alteration of cytoskeletal targets such as nitration of actin, resulting in alterations in cell adhesion and cell shape and increases in paracellular permeability (6, 12, 27).
In summary, our study indicates for the first time that TNF induces increased pulmonary microvascular endothelial permeability, which is dependent on NAD(P)H oxidase-mediated generation of . The development of strategies that target ongoing NAD(P)H oxidase activity may provide therapy for TNF-mediated acute lung injury.
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ACKNOWLEDGMENTS |
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This work was supported by National Heart, Lung, and Blood Institute Grants RO1-HL-59901-02 (A. Johnson) and RO1-HL-66301-01 (V. Rizzo).
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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