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Published ahead of print on January 10, 2003, doi:10.1165/rcmb.2002-0112OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 28, pp. 705-712, 2003
© 2003 American Thoracic Society
DOI: 10.1165/rcmb.2002-0112OC

Reactive Nitrogen Species Block Cell Cycle Re-Entry through Sustained Production of Hydrogen Peroxide

Ziqiang Yuan, Harriet Schellekens, Loraine Warner, Yvonne Janssen-Heininger, Peter Burch and Nicholas H. Heintz

Department of Pathology, University of Vermont College of Medicine, Burlington, Vermont

Address correspondence to: Nicholas H. Heintz, Department of Pathology, University of Vermont College of Medicine, Burlington VT 05465. E-mail: nheintz{at}zoo.uvm.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Endogenous sources of reactive nitrogen species (RNS) act as second messengers in a variety of cell signaling events, whereas environmental sources of RNS like nitrogen dioxide (NO2) inhibit cell survival and growth through covalent modification of cellular macromolecules. To examine the effects of RNS on cell cycle progression, murine type II alveolar C10 cells arrested in G0 by serum deprivation were exposed to either NO2 or SIN-1, a generator of RNS, during cell cycle re-entry. In serum-stimulated cells, RNS did not prevent the immediate early gene response by AP-1, but rather blocked cyclin D1 gene expression, resulting cell cycle arrest at the boundary between G0 and G1. Dichlorofluorescin diacetate (DCF) fluorescence indicated that RNS induced sustained production of intracellular hydrogen peroxide (H2O2), which normally is produced only transiently in response to serum growth factors. Loading cells with catalase did not diminish the formation of 3-nitrotyrosine on the cell surface, but rather prevented enhanced DCF fluorescence and rescued cyclin D1 expression and S phase entry. These studies indicate environmental RNS interfere with cell cycle re-entry through an H2O2-dependent mechanism that influences expression of cyclin D1 and progression from G0 to the G1 phase of the cell cycle.

Abbreviations: dichlorofluorescin diacetate, DCF • Dulbecco's modified Eagle's medium, DMEM • epidermal growth factor, EGF • fetal bovine serum, FBS • mitogen-activated protein kinase, MAPK • reactive nitrogen species, RNS • superoxide dismutase, SOD


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Reactive oxygen and reactive nitrogen species (ROS/RNS) act as physiologic second messengers in many cell signaling processes, including immune responses, cell cycle progression, and apoptosis (1, 2). In contrast, environmental sources of ROS/RNS are known to contribute to the pathogenesis of many diseases and aging by damaging cellular macromolecules (3–5). As a model for environmental RNS we have used nitrogen dioxide (NO2), a byproduct of fossil fuel combustion that can accumulate to significant levels in both indoor and outdoor air (6). NO2 also is generated from metabolites of nitric oxide (NO) during inflammatory processes (7, 8). NO2 and other RNS interact and react with numerous macromolecules, including lipids, DNA, and proteins. For example, RNS is associated with the formation of 3-nitrotyrosine, a protein modification that may affect the function of epidermal growth factor receptor (9) and MnSOD (10).

In the lung, epithelial cells are the primary targets of inhaled NO2. Acute exposure to NO2 causes shedding of bronchiolar epithelium, inflammation, pulmonary edema, and type II alveolar lung epithelial cell proliferation (11, 12). Much of the toxicity of NO2, particularly in the lung, may be related to oxidation of lipids in cell membranes to produce nitrohydroperoxides, fatty acid epoxides, and fatty acid hydroperoxides (13).

By exposing cells to defined levels of pure NO2, we observed that cell density has a profound effect on apoptosis of pulmonary epithelial cells by RNS (14). At confluence, rat and murine alveolar type II cells were highly resistant to killing by NO2, whereas cells at low density were markedly susceptible to apoptosis by the same exposure regimen. Monolayer wounding experiments showed that cells repopulating the wounded area of a cell monolayer are preferentially sensitive to NO2 (14). In contrast, apoptosis by hydrogen peroxide (H2O2) was not dependent on cell density or exacerbated by monolayer wounding (14, 15).

Because the transcriptional program in serum-stimulated fibroblasts mimics that of fibroblasts in wound healing (16), we examined the effects of NO2 and SIN-1, a generator of RNS, on cell cycle re-entry in serum-stimulated avelolar type II cells, a target cell for RNS in the lung. Here we show that generation of an irreversible cell cycle block by RNS in serum-stimulated lung epithelial cells is linked to the persistent production of intracellular H2O2, suggesting that RNS interfere with the regulation of mitogenic pathways that signal through H2O2.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell Culture and Synchronization
Murine type II alveolar C10 cells (17) were propagated in CMRL medium with 10% fetal bovine serum (FBS) with 100 U penicillin/100 µg streptomycin/ml. Cells were arrested in G0/G1 by incubation in medium containing 0.2% FBS for 72 h, and induced to re-enter the cell cycle by adding Dulbecco's modified Eagle's medium (DMEM) containing 10% FBS. Cell cycle progression was evaluated by flow cytometry (18).

Immunoblotting
Immunoblotting was performed with total cell lysates as described previously (19), using the enhanced chemiluminescence system (Amersham, Piscataway, NJ) for signal detection. Antibodies used were as follows: cyclin D1, H-295; PCNA, PC10 (Santa Cruz Biotechnology, Santa Cruz, CA); catalase, ab1877 (Abcam, Cambridge, UK).

Electrophoretic Gel Mobility Shift Assays
Gel mobility shift assays were performed with nuclear extracts and end-labeled double-stranded oligonucleotide probes for AP-1 (20, 21). Supershift assays were performed by adding 1 µl of antibody to the binding reaction after a 10-min incubation with probe.

Transfections and Reporter Gene Assays
Cells plated at 50% confluence in CMRL growth medium were transfected with cyclin D1 promoter–luciferase reporter plasmid pCD1–1745 (22) using LipofectAMINE PLUS Reagent (GIBCO-BRL, Grand Island, NY). After transfection, cells were arrested in G0 in CMRL medium with 0.2% FBS for 72 h, and then were stimulated to re-enter the cell cycle with DMEM medium containing 10% FBS, with or without exposure to RNS. Luciferase assays were performed with duplicate samples from at least two separate experiments using a kit (Promega Corp., Madison, WI): individual determinations from duplicate samples normally varied less than 3%. The statistical significance of differences between groups was assessed by ANOVA.

RNase Protection Assays
Total cellular RNA was isolated with TRIzol reagent (Life Technologies, Inc., Rockville, MD), precipitated with ethanol, and dissolved in RNase-free water. 32P-labeled RNA probes were synthesized from multiprobe template sets, hybridized to 2 µg of RNA sample, and treated with RNase and protease as described by the probe manufacturer (PharMingen, San Diego, CA). Samples were resolved on 8% denaturing sequencing gels, and the amount of radioactivity in specific bands was quantified with a phosphorimager.

Immunostaining
For immunostaining, cells were plated on glass coverslips (Corning, Corning, NY), synchronized, and treated with or without NO2 or SIN-1 as described in the text. Coverslips were rinsed twice in PBS, fixed for 5 min in 3.7% formaldehyde in PBS, rinsed in PBS, and then exposed to cold methanol (-20°C) for 3 min. After washing with PBS, samples were blocked in PBS with 1% BSA (PBA) for 30 min, incubated first with rabbit polyclonal anti-3-nitrotyrosine antibody (2 µg/ml; Upstate Biotechnology, Lake Placid, NY), and then FITC-conjugated anti-rabbit secondary antibody in PBA (Jackson Immunoresearch Laboratories, West Grove, PA). In control experiments, 3-nitrotyrosine staining was blocked by preabsorption of the primary antibody with 10-fold excess of nitrotyrosine or by chemical conversion of 3-nitro-tyrosine to aminotyrosine using sodium hydrosulfite (23). Confocal immunofluorescence and reflection image microscopy were performed with a Biorad confocal scanning laser microscopy. For statistical analysis, cells showing positive staining for 3-NT (defined as greater than 10 immunopositive foci per cell) were counted in five randomly selected fields from each experimental group.

Hydrogen Peroxide Assays
Extracellular H2O2 in medium from cell cultures exposed to NO2 or SIN-1 was measured as described (24). C10 cells were treated with NO2 and SIN-1 as before, except that DMEM without phenol red was used. To remove protein from media samples, 100 µl of 10 mg/ml BSA and 100 µl 100% trichloroacetic acid were added to 800 µl of medium, the samples were mixed and then centrifuged for 2 min in a microcentrifuge. To 800 µl of supernatant, 200 µl of 250 mM Fe(NH4)2SO4 and 100 µl 250 mM KSCN were added, the samples were mixed, and absorbance at 450 nm was measured within 10 min. A standard curve was generated by adding H2O2 to medium with 10% FBS and subjecting the samples to the same deproteination procedure.

For intracellular catalase activity, 1 mg/ml proteinase K (Life Technologies, Inc.) was added to medium for 30 min before harvest; cells then were collected by centrifugation, washed 3x in phosphate buffer (0.02 M KH2PO4, 0.03M Na2HPO4, pH 7.0) to remove proteinase K, and lysed in phosphate buffer containing 1% Triton X-100. Catalase activity was measured by determining the rate of decrease of absorbance of hydrogen peroxide at 240 nm (25).

Measurement of Intracellular Oxidation States
After exposure to test conditions, cells were washed twice with Hanks' balanced salt solution, and 10 µM 2',7'-dichlorodihydrofluorocein diacetate (DCF-DA; Molecular Probes, Eugene, OR) in phenol red-free DMEM with 10% FBS was added to the cultures at 37°C for 30 min. Cells then were trypsinized, collected by centrifugation, resuspended in Hanks' balanced salt solution, and DCF fluorescence was evaluated by flow cytometry (26, 27).

Reagents
3-morphoolinosydnonimine (SIN-1) and superoxide dismutase from bovine erythrocytes were obtained from Calbiochem (La Jolla, CA). Beef liver catalase was obtained from Boehringer Mannheim (Indianapolis, IN) and Worthington Biochemical Corp. (Freehold, NJ). Cell culture supplies were from GIBCO.

Statistics
Data were analyzed by ANOVA using the Student-Newman-Keuls procedure to adjust for multiple pairwise comparisons, with P values <= 0.05 considered as significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
RNS Induces a Cell Cycle Block
Monolayer wounding experiments showed pulmonary epithelial cells actively repopulating wounded areas are exquisitely sensitive to RNS (14). To model the effect of RNS on cell cycle re-entry, serum-starved C10 cells were exposed to pure NO2 gas during serum stimulation. Flow cytometry showed C10 cells arrest in G0/G1 after incubation in medium containing 0.2% FBS for 72 h (Figure 1A). Using a novel exposure system (14), replicate cultures were exposed for 4 h to either room air or 5 ppm NO2 in DMEM with 10% FBS as described (14). Approximately 600–800 µm nitrite accumulated in the medium under these exposure conditions (data not shown). After the initial exposure, cultures were incubated in fresh medium with 10% FBS under normal growth conditions for various periods of time. In room air, addition of medium with 10% FBS induced 80% of the cell population to enter the S phase by 12–16 h (Figure 1A). In contrast, nearly 90% of the C10 cells exposed to 5 ppm NO2 during the first 4 h of serum stimulation failed to enter the S phase (Figure 1B).



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  		Figure 1. RNS block cell cycle re-entry during the G1 phase of the cell cycle. Serum-starved C10 cells were induced to re-enter the cell cycle by the addition of fresh medium with 10% FBS in room air (A), 10% FBS with 5 ppm NO2 for 4 h (B) or 10% FBS plus 0.5 mM SIN-1 for 4 h (C). Cell cycle progression was evaluated by staining with propidium iodide and flow cytometry at 12 and 16 h after addition of medium with 10% FBS with or without RNS. (D) Serum-stimulated treated C10 cells were treated with 0.5 mM SIN-1 for various 3-h intervals during cell cycle re-entry to assess sensitivity to RNS during the G0 to G1 transition. The percentage of cells in S and G2/M at the 16-h time point was assessed by flow cytometry. (E) Dose response studies showed that 0.5–1.0 mM SIN-1 was sufficient to induce cell cycle arrest during serum stimulation. Asterisks indicate levels of cells in S and G2/M that are significantly different from control.

 
The effect of SIN-1, a generator of RNS (28), on cell cycle re-entry was also examined. Dose response experiments indicated that 4 h of exposure to 0.5–0.65 mM SIN-1 during serum stimulation was sufficient to induce arrest in G1 without inducing DNA fragmentation or other signs of apoptosis by 16 h (Figures 1C and 1E, and data not shown). As for NO2, cells exposed to SIN-1 during the first 3–4 h of serum-stimulation were most sensitive to cell cycle arrest. Although exposures for 3-h intervals later in G1 showed reduced numbers of cells in S and G2/M after 16 h (Figure 1D), time course flow cytometry and cell growth experiments showed that exposure to RNS at later intervals in G1 only delayed cell cycle progression (data not shown). In contrast, the block imposed during the initial 4 h of serum stimulation by RNS was permanent, and could not be reversed by incubation in fresh growth medium alone (see below). We then examined cell cycle markers to determine the point of RNS-induced arrest in G1.

RNS Block Cell Cycle Progression at the G0 to G1 Boundary
In serum-stimulated cells, NO2 and SIN-1 blocked phosphorylation of pRB, activation of E2F, accumulation of PCNA, and expression of cyclin D1 (data not shown, but see below), suggesting that arrest occurred upstream of the G1 restriction point at the G0 to G1 transition. Transition from G0 to G1 in serum-stimulated cells is accompanied by a cascade of cell signaling events that connect activation of growth factor receptors to cyclin D1 gene expression. The immediate early gene response to serum activates transcription largely by modification of pre-existing factors, such as phosphorylation of c-Fos and c-Jun family members to form active AP-1 (reviewed in Ref. 29). Transcriptional control of the cyclin D1 gene has been linked directly to AP-1 complexes, with JunB acting as a negative regulator of cyclin D1 promoter activity in G2/M (30), and c-Jun and the combination of c-Fos/FosB acting as obligate positive activators in G1 (30–32). Gel mobility shift experiments showed AP-1 DNA-binding activity in cell extracts accumulated at normal levels in serum-stimulated C10 cells treated with SIN-1 (Figure 2A). Moreover, analysis of the constituents of AP-1 DNA-binding complexes in extracts from cells treated with or without RNS using antibody supershift experiments revealed no quantitative alteration in the constituents in complexes bound to a consensus AP-1 probe (Figure 2B). Indeed, no obvious differences in the components of AP-1 DNA-binding complexes was observed in extracts from cells treated with SIN-1 from 30 min to 8 h (Figure 2D and data not shown). Upon cell cycle re-entry, two waves of AP-1–dependent reporter gene activity were observed, the first at 2 h and a second just before S phase (Figure 2C). In cells treated with SIN-1, the second wave of AP-1–dependent reporter gene transcription that occurred at 8 h as C10 cells approached S phase was greatly reduced (Figure 2C).



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  		Figure 2. Effect of RNS on AP-1 during cell cycle re-entry. Serum-starved C10 were incubated with medium containing 10% FBS or 10% FBS with 0.5 mM SIN-1 as before. (A) To assess AP-1 DNA-binding activity, cell extracts were prepared at the indicated time points and analyzed using electrophoretic mobility shift assays with a consensus AP-1 site (TGACTCA) as probe. (B) Supershift assays with the indicated antibodies were used to examine the constituents of AP-1 complexes in extracts from cells at the 8-h time point. Identical results were observed for supershift assays performed with extracts harvested at the 0.5-, 1-, 2-, 4-, and 6-h time points (not shown). (C) AP-1–dependent gene expression was evaluated by exposing a C10 cell line with a stably integrated c-Jun luciferase reported construct to serum or serum with SIN-1 during cell cycle re-entry. Cell extracts were normalized for protein concentration and luciferase activity is expressed as relative light units (RLU) ± standard error of the mean. Note that despite high levels of AP-1 DNA binding activity with apparently identical subunits at 8 h (A and B), C10 cells treated with SIN-1 do not execute a second wave of AP-1–dependent expression as cells approach S phase (C). Solid line with open squares, FBS; dotted line with filled diamonds, FBS + SIN-1. Asterisks indicate levels of luciferase activity in control samples that are statistically different from FBS with SIN-1.

 
Effects of RNS on Cell Cycle–Regulated Gene Expression
Results with AP-1 reporter genes suggested that the immediate early gene response was not impaired by RNS during the early phases of the G0 to G1 transition. RNase protection assays (RPAs) were used to examine mRNA levels for cyclins and various AP-1 family members, including the c-jun and fra1 genes that are regulated in part by AP-1 (Figure 3). SIN-1 did not prevent expression of AP-1 family members after serum stimulation, although mRNA for JunB and Fra1 persisted at elevated levels for longer periods of time in the presence of SIN-1 (compare Figure 3A to 3B). Under the same conditions, however, expression of cyclin D1 mRNA, which begins to accumulate by 4 h after serum stimulation (Figure 3C), was completely blocked by RNS (Figure 3D). Thus, as suggested by immunoblotting for cyclin D1 and other cell cycle markers, RNS blocked C10 cells during serum stimulation in a narrow window between the immediate early gene response and transcriptional activation of cyclin D1, or at the transition from G0 to G1.



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  		Figure 3. Analysis of mRNA levels for AP-1 and cyclin family members during cell cycle arrest by SIN-1. Total RNA was prepared from duplicate samples at the indicated time points from serum-starved C10 cells treated with 10% FBS (A and C) or 10% FBS and SIN-1 (B and D). After the 4-h exposure to SIN-1, cells were washed and incubated in fresh medium with 10% FBS as before. RNA levels for the indicated genes were determined with RNase protection assays (see supplemental data) and signals were quantified with a phosphorimager. Cyclin D1 mRNA levels did not increase in cells in treated with SIN-1, showing the block in cyclin D1 expression by SIN-1 occurs at the transcriptional level, and placing the block at the G0 to G1 transition.

 
Catalase Rescues Cell Cycle Progression from Inhibition by RNS
The chemistry of RNS is complex, as RNS may interact with ROS to generate a variety of reactive species that attack a plethora of cellular targets (reviewed in Refs. 33, 34). Moreover, spontaneous decomposition of SIN-1 to form NO and superoxide anion, which then react to form ONOO-, is affected by pH, medium components such as HEPES buffer, and other factors (35, 36). To determine how NO2 and SIN-1 induce arrest in early G1, antioxidant enzymes were tested for the ability to rescue cell cycle progression. Superoxide dismutase (SOD) partially relieved the cell cycle block by SIN-1 (Figure 4A), suggesting the enzyme coverted superoxide anion to H2O2 before it could react with NO to form ONOO- (35). However, SOD did not prevent cell cycle arrest by NO2 (Figure 4A). In contrast, addition of 1,000 U of beef liver catalase per milliliter to the culture medium completely restored S phase entry after treatment with either SIN-1 or NO2 (Figure 4A). Heat-inactivated catalase at any concentration was unable to restore cell cycle progression (data not shown).



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  		Figure 4. Catalase, but not SOD, prevents cell cycle arrest by RNS. (A) Serum-starved C10 cells were exposed to NO2 or SIN-1 for the first 4 h of serum stimulation with or without addition of SOD or catalase to the culture medium as indicated. At 16 h, cell cycle progression was measured by quantifying the number of cells in S phase and G2/M by flow cytometry. Catalase, but not SOD, rescued cell cycle progression in cells treated either with NO2 or SIN-1. Single asterisks indicate levels of cells in S and G2/M that are significantly different from control; double asterisks indicate levels of cells in S and G2/M that are significantly different from all other groups. (B) Catalase restores cyclin D1 promoter activity in serum-stimulated cells treated with SIN-1. C10 cells were transfected with a cyclin D1 luciferase reporter plasmid, synchronized in G0 by serum deprivation, and treated with medium containing 10% FBS, 10% FBS with SIN-1, or 10% FBS with SIN-1 and SOD or catalase as in (A). At 4 h, cell lysates were prepared and assayed for luciferase activity. Activity was normalized to protein concentration and is expressed as relative light units (RLU) + SEM. Single asterisk for the FBS group indicates a statistically significant difference from the G0 cell lysate. Double asterisks indicate statistically significant difference from all other groups. (C) Catalase restores expression of Cyclin D1. Serum-starved C10 cells were treated with control medium, medium with SIN-1, or medium with SIN-1 plus catalase as in A and B. Cell extracts were prepared at the indicated times and probed for the expression of cyclin D1 by immunoblotting. (D) Catalase does not prevent the formation of 3-nitrotyrosine (3-NT) residues on cellular proteins. Immunofluorescence microscopy with an antibody to 3-NT showed that exposure to 5 ppm NO2 for 4 h in the presence of catalase (NO2 + Cat) did not cause a statistically significant difference in the level of 3-nitrotyrosine residues on the cell surface in response to NO2. The 3-NT signal is shown in green; cell nuclei were stained with propidium iodide (red).

 
To assess the effects of catalase on cyclin D1 promoter activity, cells were transfected with a cyclin D1 reporter gene, synchronized in G0/G1 by serum deprivation, and treated with or without SIN-1 during serum stimulation as before. These experiments showed catalase, but not SOD, rescued activation of the cyclin D1 promoter in the presence of RNS (Figure 4B). Immunoblotting confirmed that catalase restored expression of cyclin D1 (Figure 4C), although both expression of cyclin D1 and S phase entry were slightly delayed by RNS under these exposure conditions.

To ascertain if catalase prevented access of NO2 to the cell surface, C10 cells were stained for the presence of 3-nitrotyrosine (3-NT) adducts, a marker of RNS exposure (34), after exposure in the presence and absence of catalase (Figure 4D). Statistical analysis showed no significant difference in the number of 3-NT–positive cells or degree of 3-NT staining in the presence or absence of catalase. Finally, incubating C10 cells with catalase in starvation medium for 2 h before serum stimulation also protected cells from cell cycle arrest by either NO2 or SIN-1, whereas addition of catalase after exposure did not rescue cell cycle progression (data not shown).

Catalase Prevents the Accumulation of Intracellular H2O2 by RNS
Because the enzymatic function of catalase is to detoxify H2O2, rescue of cell cycle progression by catalase indicated that the arrest induced by RNS was linked to production of H2O2. Because NO2 is not a substrate for catalase, we attempted to identify the source of H2O2 leading to cell cycle arrest. Using a ferrous thiocyanide assay (24), the level of H2O2 in medium from cell cultures exposed to NO2 was measured over time. Although reactive oxidants accumulated in medium from cultures exposed to NO2 or SIN-1 for 4 h, catalase had no effect on the levels of reactive species in either culture medium (Figure 5A), suggesting the formation of hydroperoxides or other reactive species (13). In control experiments, catalase was able to eliminate H2O2 from culture medium (Figure 5A). Measurement of intracellular levels of catalase activity indicated that C10 cells, like several other cell types (37), adsorbed catalase from medium containing either 0.2% or 10% FBS, resulting in a statistically significant increase (2- to 4-fold) in intracellular catalase activity (Figure 5B).



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  		Figure 5. Accumulation of intracellular hydrogen peroxide is required for arrest in G0 by RNS. (A) Culture medium from control or RNS-exposed cells was assayed for ROS. In control experiments, catalase removed greater than 95% of 200 µM H2O2 that was added to culture medium (Media), but did not alter the levels of ROS in medium from cells exposed to either 5 ppm NO2 or 0.5 mM SIN-1 for 4 h, suggesting the formation of hydroperoxides. Asterisk indicates statistically significant from matched control. (B) Intracellular levels of catalase activity increased after addition of beef liver catalase to culture media. Serum-starved or serum-stimulated C10 cells were incubated with 1,000 U beef liver catalase/ml medium for 4 h. Cells were treated with proteinase K for 30 min, washed, and cell extracts were assayed for catalase activity. In culture medium alone, addition of proteinase K for 15 min was sufficient to completely inactivate catalase activity (not shown). Asterisk indicates statistically significant difference from matched control. (C) C10 cells were exposed to medium for 4 h with 1,000 (lanes 3 and 5) or 3,000 (lanes 1, 4, and 6) U/ml beef liver catalase. To assay for cellular uptake of catalase, proteinase K was added to medium for the final 30 min (lanes 3 and 4) or the first 30 min (lanes 5 and 6) of the incubation, and cell extracts then were analyzed for the presence of catalase (arrow) by immunoblotting. Extract from control cells not treated with catalase is in lane 2. Immunoreactive material that migrates at less than 60 kD (bracket) likely represents degradation products. (D) Serum-starved C10 cells were treated with medium containing 10% FBS, 10% FBS with SIN-1 or 5 ppm NO2, or 10% FBS with SIN-1 or 5 ppm NO2 with 1,000 U/ml catalase. After 4 h, cells were examined for intracellular oxidative stress by DCF fluorescence and flow cytometry. Catalase reduced DCF fluorescence to control levels in cells treated with either NO2 or SIN-1.

 
Immunoblotting of cell lysates from cultures exposed to catalase and then treated with proteinase K was used to determine if C10 cells import catalase from culture medium. In the absence of added catalase, immunoblotting of extracts from cells treated with proteinase K showed no 60 kD band for catalase (Figure 5D, lane 2). Extracts from control cultures treated with catalase, but not proteinase K, clearly showed the presence of catalase (Figure 5D, lane 1). When proteinase K was added along with catalase to culture medium, catalase was completely digested by 30 min (Figure 5C, compare lane 1 with lanes 5 and 6). In contrast, when proteinase K was added for the last 30 min of a 4-h incubation with catalase, a dose-dependent increase in catalase was observed in C10 cell lysates (Figure 5C, lanes 3 and 4), suggesting catalase was imported into C10 cells, thereby protecting the enzyme from protease digestion. These results agree well with catalase activity assays (Figure 5B). We assume that minor bands observed below the catalase signal at 60 kD represent degradation products (bracket, Figure 5C).

Transient production of intracellular H2O2 is required for growth factor signaling leading to mitogenesis (37–39). A cell-permeable fluorescent dye (DCF-DA) that is indirectly sensitive to oxidation by H2O2 was used assess the levels of intracellular oxidative stress after exposure to either NO2 or SIN-1 for 4 h. After exposure, cells were washed and loaded with DCF-DA for 30 min, trypsinized, and examined by flow cytometry. C10 cells had increased intracellular DCF fluorescence 4 h after serum stimulation when treated with either NO2 or SIN-1 (Figure 5D), long after transient increases in DCF fluorescence due to growth factor stimulation had diminished in control cells (data not shown). Although the fraction of cells displaying enhanced DCF fluorescence was lower in cultures treated with NO2 ({bsim} 44%) as compared with cultures treated with SIN-1 ({bsim} 76%), addition of catalase to the culture medium completely prevented sustained DCF fluorescence in both instances (Figure 5D). Thus, sustained intracellular DCF fluorescence was associated with cell cycle arrest at the G0 to G1 boundary, and restoration of cell cycle progression by catalase was correlated with elimination of enhanced levels of DCF fluorescence.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Mammalian cells mount a variety of responses to counter the effects of H2O2 and other external sources of oxidative stress, including expression of antioxidant enzymes such as glutathione S-transferases, peroxidases, and SODs. Extracellular H2O2 also influences growth factor receptor signaling and activation of mitogen-activated protein kinase (MAPK), leading to changes in cell survival and apoptosis (reviewed in Refs. 1, 4, 40). Recent studies also have shown that transient pulses of intracellular hydrogen peroxide play a critical role in growth factor signaling through the epidermal growth factor (EGF) and platelet-derived growth factor receptors leading to mitogenesis (37–39). Our studies suggest there may be a relationship between exposure to extracellular sources of RNS, deregulation of intracellular H2O2 levels, and cell cycle progression. Because cell cycle arrest in response to NO2 or SIN-1 can be rescued by catalase, and catalase acts through dampening intracellular levels of H2O2, RNS must induce changes in specific H2O2-dependent signaling pathways that govern growth control.

Understanding the relationship between intra- and extracellular sources of oxidants in growth control has been complicated by the ability of extracellular H2O2 and other ROS to penetrate the cell membrane. For example, SIN-1 has been used as a generating system in studies on the effects of RNS on cell growth (41, 42), but these studies are complicated by the fact that SIN-1 generates H2O2 in addition to RNS. In contrast, the use of pure NO2 gas in studies here clearly show that cell cycle responses to extracellular sources of RNS can be mediated via pathways that are responsive to intracellular levels of H2O2.

Our results suggest a model in which perturbation of signaling pathways by RNS leads to sustained production of intracellular H2O2, the downstream effects of which can be bypassed by catalase (Figure 6). Upon addition of serum, tyrosine kinase activity of growth factor receptors is stimulated, leading to the assembly of signaling complexes that activate MAPK and other downstream kinase cascades. In serum-stimulated cells, these events occur within minutes. Activation of growth factor receptors (and integrins) in turn activates Ras and Rac, which stimulate membrane-bound NADPH oxidase complexes to produce superoxide anion, which is rapidly converted to H2O2 by SOD (reviewed in Ref. 1). Mitogenic signaling in response to platelet-derived growth factor, EGF, and activated HER-2/neu is known to be a redox-sensitive process that requires the production of intracellular H2O2 (37–39), which acts as a second messenger on unknown targets. Interestingly, Rac regulates both ROS generation in response to growth factors (43) and translation of cyclin D1 mRNA in response to integrin signaling (44), providing a potential node for integrating signals that culminate in cyclin D1 expression.



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  		Figure 6. A model for cell cycle arrest in response to serum and RNS. We propose that RNS interferes with cell signaling pathways in response to serum, leading to sustained production of intracellular H2O2. Catalase does not affect the accumulation of reactive species in the cell culture medium or reduce the formation of 3-NT residues on cellular proteins, indicating that it acts downstream of the immediate early gene response by detoxifying intracellular H2O2. Cyclin D1 transcription is restored by catalase, suggesting activation of this promoter is sensitive to redox regulation at the transition between G0 and G1.

 
Under normal circumstances, like other signaling events (45), redox-dependent signaling events leading to mitogenesis must be transient and decay over time. Here RNS induced prolonged production of intracellular H2O2 in cells responding to serum. Analysis of AP-1–dependent gene activation showed that RNS does not perturb the initial phases of the immediate early gene response, but rather cause arrest at the G0 to G1 transition just upstream of cyclin D1 expression. In other contexts, sustained activation of ERK mediated by signaling through integrins and growth factor receptors has been shown to be critical for activation of cyclin D1 expression in adherent cells (43, 44). Kinase assays and immunoblotting for phospho-ERK have shown that ERK is activated in the presence of RNS, and remains phosphorylated under conditions of catalase rescue, suggesting that failure to signal through ERK is not responsible for failure to induce cyclin D1 transcription (data not shown).

The potential targets of RNS that might promote cell cycle arrest through prolonged redox-dependent signaling include growth factor receptors, membrane lipids, MAPK kinases, phosphatases, the proteosome, and other macromolecules (reviewed in Ref. 1, 4, 5, 40). For example, peroxynitrite, a potent nitrating agent, induces covalent dimerization of EGF receptors, a process that is enhanced by EGF (9). Hence, crosslinking of EGFR and other receptors by RNS in the presence of serum ligands could result prolonged signaling, leading to sustained, rather than transient, production of H2O2. Although there are many potential sources of intracellular H2O2, preliminary results show apocynin, but not cyclosporin A, is able to prevent irreversible growth inhibition (Yuan and coworkers), suggesting NADPH oxidase complexes activated in response to growth factors may be one source of the elevated intracellular H2O2 detected under our experimental conditions. This possibility is supported by the observation that overexpression of Nox1 causes loss of growth control and neoplastic transformation (46), phenotypes that are mediated increased production of intracellular H2O2 and completely reversed by the expression of catalase (47).

By using activation of cyclin D1 transcription as a marker of entry into G1, the sensitivity of specific signaling factors to redox-dependent processes that mediate cell cycle progression may be delineated. Our present studies indicate that RNS do not prevent the accumulation of sequence-specific transcription factors such as CREB, AP-1, nuclear factor-{kappa}B, or TCF that regulate cyclin D1 transcription, but actually prolong the time that these factors remain activated. In this regard, the discordance between AP-1 binding activity and AP-1 reporter gene activity reported here is instructive (Figure 2). Chromatin binding experiments show that trafficking of AP-1 subunits on and off chromatin after serum stimulation is altered by RNS (data not shown), raising the possibility that sustained production of intracellular H2O2 interferes with other steps in activation of cyclin D1 transcription, such as chromatin remodeling or recruitment of coactivators. Because cycling cells do not pass through G0, our model for examining the G0 to G1 transition in serum-stimulated cells should prove to be useful for identifying specific steps in cell cycle re-entry that are subject to regulation by intracellular H2O2.


   Acknowledgments
 
The authors thank S. Tighe and J. Koh for assistance with flow cytometry, R. Pestell for the cyclin D1 reporter construct, R. Persinger and B. Mossman for discussions, P. Vacek for statistical analysis, and D. Hemenway for assistance with the NO2 exposure platform. This work was supported by a grant from NIEHS (ES09673).

Received in original form July 10, 2002

Received in final form December 6, 2002


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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