The role of p21CIP1/WAF1 in growth of epithelial cells exposed to hyperoxia

Raymond C. Rancourt1, Peter C. Keng2, Christopher E. Helt1, and Michael A. O'Reilly1,3

Departments of 1 Environmental Medicine, 2 Radiation Oncology, and 3 Pediatrics, University of Rochester, Rochester, New York 14642


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Previous studies have shown that hyperoxia inhibits proliferation and increases the expression of the tumor suppressor p53 and its downstream target, the cyclin-dependent kinase inhibitor p21CIP1/WAF1, which inhibits proliferation in the G1 phase of the cell cycle. To determine whether growth arrest was mediated through activation of the p21-dependent G1 checkpoint, the kinetics of cell cycle movement during exposure to 95% O2 were assessed in the Mv1Lu and A549 pulmonary adenocarcinoma cell lines. Cell counts, 5-bromo-2'-deoxyuridine incorporation, and cell cycle analyses revealed that growth arrest of both cell lines occurred in S phase, with A549 cells also showing evidence of a G1 arrest. Hyperoxia increased p21 in A549 but not in Mv1Lu cells, consistent with the activation of the p21-dependent G1 checkpoint. The ability of p21 to exert the G1 arrest was confirmed by showing that hyperoxia inhibited proliferation of HCT 116 colon carcinoma cells predominantly in G1, whereas an isogenic line lacking p21 arrested in S phase. The cell cycle arrest in S phase appears to be a p21-independent process caused by a gradual reduction in the rate of DNA strand elongation. Our data reveal that hyperoxia inhibits proliferation in G1 and S phase and demonstrate that p53 and p21 retain their ability to affect G1 checkpoint control during exposure to elevated O2 levels.

p53; oxidants; cell proliferation


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

HYPEROXIA REFERS TO AN ENVIRONMENT containing O2 levels higher than those found in the atmosphere at sea level. Although high O2 concentrations are used therapeutically for the treatment of a number of pulmonary and cardiovascular disorders, its continued use can also have harmful side effects. In vitro studies (5, 6, 10, 24) with a wide variety of cells have shown that continued exposure to hyperoxia inhibits proliferation and decreases survival. Boveris and Chance (4) laid the groundwork for understanding the biochemistry of O2 toxicity by demonstrating that H2O2 production by mitochondria increases during hyperoxia. Subsequent work has shown that cultured cells maintained in a high O2 atmosphere display elevated rates of O<UP><SUB>2</SUB><SUP>−</SUP></UP> formation in addition to H2O2 production (27). Although this increased flux of reactive oxygen species (ROS) during hyperoxic exposure is believed responsible for the physiological changes that result in decreased viability, the mechanism whereby cells growth arrest during these conditions has yet to be fully clarified.

Proliferation in eukaryotes occurs when a cell departs from the first stage of interphase (G1) and enters into a period of DNA synthesis termed S phase. After duplication of its DNA, the cell then enters G2 and M phases during which formation of the mitotic spindle and mitosis occur. In response to environmental conditions unfavorable for cell reproduction, replicative DNA synthesis and mitosis are restricted by the activation of cell cycle checkpoints at the G1/S and G2/M boundaries. The signal transduction pathway responsible for G1 checkpoint activation has been characterized predominantly by investigation of cell cycle delay in response to DNA-damaging agents. The p53 tumor suppressor is a principal regulator of the G1 checkpoint. p53 protein accumulates within 1-3 h after a DNA-damaging event by a posttranscriptional mechanism involving, in part, phosphorylation on serine residue 15 (17, 30). Increased levels of p53 regulate the expression of specific genes involved in growth regulation and apoptosis (12). One of the major targets of p53 involved in cell cycle control is the cyclin-dependent kinase (Cdk) inhibitor p21CIP1/WAF1 (hereafter termed "p21"). Elevated levels of p21 protein can prevent G1/S progression by binding and inhibiting the activities of the G1 and S phase kinases, cyclin E-Cdk2, cyclin D-Cdk4-6, and cyclin A-Cdk2 (9). The p21 protein has also been shown to interact by proliferating cell nuclear antigen (28), suggesting another route by which it may control proliferation at the G1/S boundary. Cells lacking either p53 or p21 fail to arrest in G1 in response to genomic damage (8, 21). Cultured cells have also been shown to undergo a p53-dependent G1 arrest when treated with drugs that deplete ribonucleotide pools (15). DNA replication by cells with insufficient ribonucleotide pools results in chromosomal instability. Thus the limitations on G1/S progression imposed by p53 serve not only to prevent replication of damaged DNA templates but also to prevent DNA damage from occurring during conditions that may bring about replication-associated damage.

There are two plausible theories, which are not mutually exclusive, that could explain how hyperoxia inhibits proliferation. The first is that damage to the cell replication machinery and/or energy processes has occurred in sufficient quantity, thereby making replication impossible. A number of studies (2, 25) have provided supportive evidence that the inactivation of mitochondrial enzymes, mitochondrial damage, and gradual respiratory failure occur in both intact lungs and cultured cells in response to hyperoxia. Because mitochondria supply much of the energy necessary for biosynthetic reactions within the cell, loss of ATP could impair the replication process. The second possibility is that hyperoxia could be activating G1 and/or G2 checkpoints, similar to what is observed after administration of genotoxic agents. Several studies have shown that hyperoxia increases p53 and p21 expression in vivo (16, 19, 20) and in vitro (7, 26). Corroyer et al. (7) were the first to suggest that regulatory molecules may act to limit G1/S progression during hyperoxia by demonstrating increased transforming growth factor-beta and p21 levels and lowered cyclin E-Cdk2 activity in O2-arrested SV40-immortalized rat lung type II cells. Although cell count measurements were used in that study to demonstrate growth inhibition, there was no evidence to indicate whether growth arrest was cell cycle phase specific. In a recent report by Schenberger and Dixon (26), smooth muscle cell cultures exposed to hyperoxia showed increased p53 and p21 levels. Although a decline in the number of cells with G1 phase DNA content and an increase in cells possessing S phase DNA content were observed in these cultures, there was no evidence to suggest that these cell cycle alterations were a functional consequence of p53 or p21 activity. Although Rancourt et al. (22) previously demonstrated that transforming growth factor-beta is not involved in hyperoxia-induced growth arrest of Mv1Lu (mink lung adenocarcinoma) cells, the question of whether it operates through a p53/p21 pathway remains untested.

In the present study, we have used both p21-functional and p21-deficient epithelial cells to show that hyperoxia inhibits proliferation in both G1 and S phases of the cell cycle. Our results indicate that the delay incurred at G1 phase during hyperoxia is mediated by the p53-dependent activities of p21. The cell cycle arrest that occurs in S phase appears to be a p21-independent process that is caused instead by a gradual reduction in the rate of nascent DNA strand elongation. Cell lines capable of p21-dependent G1 checkpoint activation during hyperoxia can attenuate this S phase trapping, however, by limiting the number of cells entering into S phase. Because hyperoxia did not significantly reduce ATP levels when growth arrest was observed, the mechanisms by which it attenuates S phase progression remain unclear. Our findings are consistent with previous studies (6, 7, 22, 26) and extend them by demonstrating that hyperoxia inhibits proliferation in several phases of the cell cycle.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture. Mv1Lu cells were obtained from Dr. Anita Roberts (National Cancer Institute, National Institutes of Health, Bethesda, MD). A549 and Saos-2 cells were purchased from the American Type Culture Collection (Manassas, VA). HCT 116 cells and HCT 116 cells lacking p21 were obtained from Dr. Bert Vogelstein (Johns Hopkins Oncology Center and the Program in Human Genetics and Molecular Biology, Baltimore, MD).

Mv1Lu, A549, and Saos-2 cells were incubated in 5% CO2 at 37°C in Dulbecco's modified Eagle's medium (high glucose) with 10% fetal bovine serum (FBS), 50 U/ml of penicillin, and 50 µg/ml of streptomycin (GIBCO BRL). HCT 116 cells were incubated in McCoy's medium with 10% FBS, 50 U/ml of penicillin, and 50 µg/ml of streptomycin.

Hyperoxic exposure. Cells were trypsinized, counted with an electric particle counter (Coulter Electronics, Hialeah, FL), and plated in 100-mm plastic dishes at a density of 5 × 105. After 24 h, the cultures were given 10 ml of fresh medium and placed in a Plexiglas box (Belco Glass, Vineland, NJ). The box was sealed and flooded with 95% O2-5% CO2 for 15 min at a flow rate of 5 l/min. O2 concentrations were monitored with a miniOXI analyzer from Catalyst Research (Owings Mills, MD).

Flow cytometric cell cycle analysis. Room air- and hyperoxia-treated cells were trypsinized, resuspended in their original medium, and centrifuged at 300 g. The cell pellet was washed in PBS and fixed in 10 ml of 75% ethanol for at least 24 h at -20°C. Fixed cells were centrifuged, washed once in PBS, and incubated in 1 ml of RNase (1 mg/ml) for 30 min. Cells were recentrifuged and resuspended in 0.6 ml of PBS containing 10 µg/ml of propidium iodide. Propidium iodide fluorescence (relative DNA content per cell) was measured with an Epics Profile (Coulter Electronics) equipped with an air-cooled argon laser excited at 488 nm. To test for the G1 checkpoint, cells were exposed to 6 Gy of 137Cs at a dose rate of 3.6 Gy/min. Colcemid (0.6 mg/ml; GIBCO BRL) was administered to cells 45 min before irradiation. Cells were harvested 3-24 h after irradiation and analyzed for DNA content by flow cytometry.

5-Bromo-2'-deoxyuridine-labeling experiments. Cells were exposed to hyperoxia for 0, 24, 48, and 72 h. Prewarmed 5-bromo-2'-deoxyuridine (BrdU; 10 µM)-containing medium was administered to these exposed cells for 1 h in a 5% CO2-containing atmosphere before harvest. The cells were then trypsinized, centrifuged, and stored in 75% ethanol for at least 24 h. On the day of analysis, the cells were treated with 3 ml of 2 N HCl in PBS containing 0.2 mg/ml of pepsin for 20 min at 37°C. After centrifugation, cells were resuspended in 1 ml of PBS containing 2% FBS. After removal of the PBS, cells were incubated in 10 ng of anti-BrdU-FITC (Boehringer Mannheim, Indianapolis, IN) for 45 min followed by RNase treatment and propidium iodide counterstaining. Measurements of red (DNA content) and green (BrdU content) fluorescence were determined by flow cytometry.

To determine rates of S phase progression, BrdU pulse-chase studies were performed. Cells were initially exposed to hyperoxia for 15 h. Cultures were then incubated with BrdU for 1 h in room air-5% CO2. After complete removal of the BrdU-containing medium, cultures were returned to hyperoxia with fresh medium and harvested at various time points. Room air-exposed cells were also treated with BrdU and harvested in parallel fashion for comparison. All samples were then fixed and analyzed for DNA and BrdU content as described in Flow cytometric cell cycle analysis and 5-Bromo-2'-deoxyuridine-labeling experiments. The movement of cells through S phase was quantified with the use of Cytology software (Coulter Electronics) as described by Begg et al. (3). A region was set around the BrdU-positive labeled (green fluorescent) cells, and their mean DNA content was calculated by measuring their red fluorescence. The movement of these S phase cells relative to the positions of G1 and G2 was then calculated by use of the formula RM = (FL - FG1)/ (FG2M - FG1), where RM is relative movement, FL is the mean DNA content of BrdU-labeled cells, FG1 is the mean DNA content of G1 cells, and FG2M is the mean DNA content of G2/M cells.

Western blot analysis. The cells were harvested by scraping at 4°C in 50 mM Tris (pH 8.0), 120 mM NaCl, and 0.5% Nonidet P-40 supplemented with 2 µg/ml of aprotinin and 100 µg/ml of phenylmethylsulfonyl fluoride. The cell lysate was cleared by centrifugation, and the protein concentration was determined by the Bradford assay. The lysates were boiled in 3× Laemmli buffer [1× = 62.5 mM Tris (pH 6.8), 2% SDS, 10% glycerol, and 0.5% bromphenol blue]. Proteins were separated by size on SDS-polyacrylamide gels and transferred to nitrocellulose. The membranes were blocked overnight in 1× PBS containing 5% nonfat dry milk followed by incubation with the primary antibody, p53 (1:3,000; Santa Cruz Biotechnology, Santa Cruz, CA), anti-phospho-p53 (Ser 15; 1:2,000; New England Biolabs, Beverly, MA), or p21 (1:3,000; PharMingen, San Diego, CA). Nonspecific interactions were removed by a wash in Tris-buffered saline with 0.5% Tween 20 (TBS-T) for 1 h followed by incubation with goat anti-mouse secondary antibody (1:5,000; Southern Biotechnology, Birmingham, AL). The blots were washed again in TBS-T for 1 h, and the conjugates were visualized by chemiluminescence (Amersham, Arlington Heights, IL). The blots were reprobed with anti-beta -actin antibody (1:5,000; Sigma, St. Louis, MO) as a loading control.

p53 transfection studies. Saos-2 cells were plated at 500,000 cells/60-mm dish and allowed to grow for 20 h. Each plate was transfected with 2 µg of p53-luciferase and 0-2 µg of pFC-p53 expression plasmid (Stratagene, La Jolla, CA) with calcium phosphate precipitation. The p53-luciferase construct contained 15 copies of the p53 binding site within the p21 promoter in front of a minimal TATA box. The pFC-p53 plasmid expressed wild-type p53 under the control of the cytomegalovirus promoter. All samples were cotransfected with 0.1 µg of the Renilla luciferase-expressing plasmid pRL-SV40, which was used to normalize for transfection efficiency with the Dual-Luciferase reporter assay system (Promega, Madison, WI). Transfected cells were washed three times with PBS before exposure to room air or hyperoxia for 24 h, and then luciferase activity was determined.

Measurement of ATP. ATP measurements from cells were obtained with a luciferase-luciferin kit (Calbiochem, San Diego, CA). In triplicate, cells were exposed to room air or hyperoxia, harvested, and resuspended in Hanks' balanced salt solution at 50,000 cells/ml. Molar quantitation of ATP was determined from an ATP standard curve generated on the day of the analysis.

Statistical analysis. Values are means ± SD. Group means were compared by ANOVA with Fisher's procedure post hoc analysis with StatView software for Macintosh. P < 0.05 was considered significant.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Hyperoxia inhibits proliferation of Mv1Lu and A549 cells. Exponentially growing Mv1Lu and A549 cultures were either maintained under normal incubation conditions (95% room air-5% CO2; 37°C) or exposed to hyperoxia (95% O2-5% CO2; 37°C) for up to 72 h. To determine the effect on cell proliferation, cells were harvested every 24 h and counted by use of an electric particle counter (Fig. 1). As expected, cell numbers in Mv1Lu and A549 cultures exposed to room air increased daily up to 72 h. In contrast, hyperoxia inhibited the growth of Mv1Lu cells, as shown previously by Rancourt et al. (22), as well as the growth of A549 cells. Short-term measurement of cell viability by dye exclusion and terminal deoxynucleotidyltransferase-mediated dUTP nick end labeling (TUNEL) assay indicated that >90% of these cells remained viable during the course of 72 h of hyperoxia (13, 22) (data not shown).


View larger version (24K):
[in this window]
[in a new window]
 
Fig. 1.   Hyperoxia inhibited proliferation of Mv1Lu and A549 cells. Mv1Lu (A) and A549 (B) cells were cultured for 24, 48, or 72 h in room air (with 5% CO2) or hyperoxia (95% O2-5% CO2), harvested, and counted. Values are means ± SD of 3 independent experiments.

Cell cycle response to hyperoxia. To gain insight into the mechanisms involved in the inhibition of proliferation by hyperoxia, exponentially growing cells were exposed to room air or hyperoxia for 24, 48, or 72 h and analyzed for DNA content (Fig. 2A). DNA histograms depicted at the 0-h time point are representative of room air-exposed cultures in exponential growth. Mv1Lu cells exposed to hyperoxia began to demonstrate significant cell cycle alterations compared with those in room air-exposed control cells after 24 h when increases in the number of S phase cells and a decline in the fraction of cells in G1 phase were evident. Longer exposures caused further depletion of the G1 population so that by 72 h of exposure, >80% of Mv1Lu cells were in S phase. Hyperoxia also caused an increase in the percentage of A549 cells in S phase. As illustrated in Fig. 2B, however, the magnitude of this S phase accumulation and the associated decline in G1 cells was significantly less than that observed in Mv1Lu cells.


View larger version (39K):
[in this window]
[in a new window]
 
Fig. 2.   Cell cycle analysis of Mv1Lu cells and A549 cells. A: flow cytometric profiles for Mv1Lu and A549 cells exposed to room air (room air-5% CO2) and 24, 48, or 72 h of hyperoxia (95% O2-5% CO2). Values are means ± SD of 3 independent experiments. B: mean average change of Mv1Lu and A549 cells in G1, S, and G2/M phases over the course of 72 h of hyperoxia.

To examine the impact of hyperoxia on DNA synthesis, cells exposed to room air or 24-72 h of hyperoxia were incubated with BrdU for 1 h and analyzed for DNA content and BrdU uptake by flow cytometry (Fig. 3). Compared with the room air-treated control cells, BrdU incorporation was diminished after 24 h of hyperoxia in both Mv1Lu and A549 cells. Cells from cultures exposed to hyperoxia for 48 h or longer showed a further reduction in BrdU fluorescence intensity, which approached the threshold level of unlabeled negative control populations. Thus cultures exposed to hyperoxia lost the capacity to proliferate despite overall increases in S phase cell number.


View larger version (54K):
[in this window]
[in a new window]
 
Fig. 3.   Effect of hyperoxia on 5-bromo-2'-deoxyuridine (BrdU) incorporation. Bivariate distributions of BrdU vs. DNA content from Mv1Lu and A549 cells maintained in room air (0 h) or exposed to hyperoxia for 24, 48, or 72 h are shown. All cultures were labeled with 10 µM BrdU for 1 h and harvested, and incorporated BrdU was identified immunologically with fluorescent secondary antibodies. Insets, cells exhibiting fluorescence content greater than the level detected in unlabeled control cultures. Data represent a typical experiment (n = 3 experiments).

Rates of DNA elongation are reduced during hyperoxia. BrdU pulse-chase studies were performed to determine whether cells exposed to hyperoxia were capable of elongating DNA strands. The purpose of this experiment was to determine the rate at which cells progress through S phase towards the G2 phase of the cell cycle. Mv1Lu and A549 cells were initially exposed to 15 h of hyperoxia, pulsed with BrdU for 1 h in room air, returned to hyperoxia, and harvested at times between 0 and 15 h. Cells not exposed to hyperoxia were also treated with BrdU and harvested in a parallel fashion for purposes of comparison. We chose to label cells after 15 h of exposure because pilot studies revealed that it was difficult to obtain labeled populations if they were exposed to hyperoxia for longer than this period of time (see Fig. 3 as an example of BrdU-labeled cells after 24 h of hyperoxia). Figure 4 displays a representative experiment that was performed on the Mv1Lu cell line. The BrdU-labeled S phase fractions from room air-exposed cells progressed steadily toward the G2 phase of the cell cycle. More than 60% of the labeled cells had divided and returned to G1 phase after 7 h. By 9 h, 95% of the cells had completed mitosis. In contrast, the rates of progression through S phase for cells exposed to hyperoxia were considerably slower. After 7 h, most of the labeled cells were still in S phase, with <1% having undergone a round of cell division. After 9 h, only 6% of the labeled fraction had undergone mitosis. Using the formula developed by Begg et al. (3), we calculated that S phase transit for Mv1Lu cells exposed to room air was 7 h and increased by an additional 8.3 h when the cells were exposed to hyperoxia (Table 1). Similar studies revealed that hyperoxia increased the duration of S phase in A549 cells by an additional 3.5 h.


View larger version (43K):
[in this window]
[in a new window]
 
Fig. 4.   Effect of hyperoxia on DNA synthesis. Bivariate distributions of BrdU incorporation vs. DNA content for Mv1Lu cells exposed to room air (top) or preexposed to hyperoxia for 15 h (bottom) are shown. After the initial exposure, cultures were labeled with 10 µM BrdU for 1 h and then returned to room air or hyperoxia. Cultures were harvested after the 1-h pulse and after 3, 7, and 9 h of chase. Insets, cells exhibiting fluorescence content greater than the level detected in unlabeled control cultures. Data are representative of a typical experiment (n = 3 experiments for Mv1Lu cells).


                              
View this table:
[in this window]
[in a new window]
 
Table 1.   Effect of hyperoxia on S phase transit time

Determination of p53 and p21 expression during hyperoxia. Previous studies (7, 16, 19, 20, 26) have demonstrated that p53 and p21, the key regulatory proteins in controlling G1/S progression, are increased in both cell lines and mouse lungs in response to hyperoxia. To determine whether these gene products contribute to the cell cycle alterations that occur in Mv1Lu and A549 cultures during hyperoxia, changes in their expression were determined during exposure. A low constitutive level of p53 was detected in Mv1Lu and A549 cells exposed to room air (Fig. 5A). A marked induction of p53 expression was observed in both cell lines after exposure to hyperoxia for 24 or 48 h. The same sets of extracts were also used for detection of p21. p21 was expressed only at a very low level in A549 cells exposed to room air. However, it was easily detected after exposure to hyperoxia for 24 h and increased further after 48 h. In contrast, p21 was never detected in Mv1Lu cells despite the use of three different commercially available antibodies.


View larger version (32K):
[in this window]
[in a new window]
 
Fig. 5.   Hyperoxia activates p53 transcription and induction of p21CIP1/WAF1 (p21). A: Mv1Lu and A549 cells were exposed to room air (0 h) or hyperoxia for 24 and 48 h. Cells were harvested, and the expression of p53, anti-phospho-p53 (Ser 15), and p21 were detected by Western blotting. The expression of beta -actin was used to confirm that comparable levels of protein were present in all samples. The blots represent a typical experiment (n = 5 experiments). B: Saos-2 cells were transfected with p53-luciferase alone or with increasing amounts of the p53 expression vector pFC-p53 and exposed to room air (RA) or hyperoxia (O2). Values are multiples of increase ± SD of relative luciferase activity compared with room air-exposed cultures; n = 3 experiments. Hyperoxia significantly increased luciferase activity, *P < 0.05.

Hyperoxia induces p53 phosphorylation and transcriptional activities. Phosphorylation of p53 on serine-15 resulted in stabilization of p53 protein and increased the ability to bind and transcriptionally activate target genes such as p21 (17, 30). The anti-phospho-p53 (Ser 15) antibody was used to examine the phosphorylation of p53 on serine residue 15 during hyperoxia. Phosphorylated p53 was not readily detected in cultures of Mv1Lu and A549 cells exposed to room air (Fig. 5A). Marked increases in phosphorylated p53 were detected in both cell lines after 24 and 48 h of hyperoxia. Thus hyperoxia-induced accumulation of p53 coincides temporally with phosphorylation of serine-15.

To determine whether increased levels of p53 were capable of transactivating p21, we assessed whether hyperoxia could activate a p53-luciferase reporter that contained tandem copies of the p53 binding site within the p21 promoter linked to a minimal TATA box. Preliminary studies found that this construct expressed a very high basal level of reporter activity when transfected into Mv1Lu or A549 cells that were exposed to room air (data not shown). This finding was consistent with a study (23) demonstrating that transfected DNAs can activate the p53-dependent damage response. A closer examination of the literature revealed that p53 transactivation assays have been assessed in either cell lines lacking p53, where low levels of p53 are provided through cotransfection, or cell lines in which the p53-luciferase gene is integrated stably within the genome. We therefore assessed whether hyperoxia could activate p53-dependent transactivation in the p53-deficient human osteosarcoma cell line Saos-2. Minimal luciferase activity was detected in Saos-2 cells exposed to room air or hyperoxia (Fig. 5B). A modest induction was obtained when the cells were cotransfected with 20-2,000 ng of p53 expression vector and exposed to room air. In contrast, luciferase activity increased 100- to 500-fold in these cells on exposure to hyperoxia. Thus hyperoxia increases posttranscriptional phosphorylation and transcriptional activity of p53.

G1 checkpoint activation in response to gamma radiation treatment. Our data are consistent with the concept that hyperoxia increases p53 in part through phosphorylation, which can transcriptionally increase p21. Increased levels of p21 would in turn inhibit proliferation by checkpoint activation at the G1/S transition. However, hyperoxia increased p53 and p21 in A549 cells, whereas only p53 was increased in Mv1Lu cells. Because hyperoxia caused more A549 cells compared with Mv1Lu cells to accumulate in G1, we hypothesized that the p53 and p21-dependent checkpoint activation was defective in Mv1Lu cells. This hypothesis was tested by assessing the G1 checkpoint response in Mv1Lu and A549 cells after exposure to ionizing radiation. Colcemid was administered to cells 45 min before irradiation to block all cells in mitosis from dividing. Blocking cells in M phase with colcemid prevented second-cycle G1 cells from contaminating the DNA histograms used in investigating the G1 delay response. DNA histograms for A549 and Mv1Lu cells treated with colcemid alone or colcemid followed by a single dose of 6 Gy of radiation are presented in Fig. 6. Cells administered colcemid moved through the cell cycle until encountering the colcemid block in G2/M phase. In response to radiation exposure, however, A549 cells experienced a significant delay in their ability to move out of G1 and into S phase. The higher proportion of G1 cells in the irradiated group compared with the nonirradiated group at all sampling times was an indication that radiation can invoke a G1 delay in this cell type. Irradiated and nonirradiated Mv1Lu cells, however, revealed no difference in the number of cells in G1 phase under simultaneous experimental conditions. Ionizing radiation also increased p53 and p21 in A549 cells but only p53 in Mv1Lu cells (data not shown). Thus ionizing radiation was able to elicit a G1 checkpoint in A549 but not in Mv1Lu cells, consistent with the findings after exposure to hyperoxia.


View larger version (23K):
[in this window]
[in a new window]
 
Fig. 6.   G1 phase arrest after gamma radiation. DNA histograms of A549 (A) and Mv1Lu (B) cells treated with colcemid with and without exposure to 6 Gy of ionizing radiation are shown. Cells were harvested at the times indicated, and DNA content was measured by flow cytometry. Figure represents a typical experiment (n = 2) comparing cell number and DNA content.

Effect of hyperoxia on proliferation of HCT 116 cells. A fundamental difficulty encountered when working with transformed cell types of different origins is the likelihood of unrecognized phenotypic differences that could confound experimental comparisons. To confirm that hyperoxia activates the p21-dependent G1 checkpoint, we assessed its effects on proliferation in the human colorectal cell line HCT 116 p21(+/+), which expresses wild-type p21, and an isogenic clone, HCT 116 p21(-/-), in which the p21 was knocked out by homologous recombination (29). The parental HCT 116 p21(+/+) cells showed only slight decreases in the fractions of cells in G1 and G2 phases, with a modest increase in S phase during the course of 72 h of hyperoxia (Fig. 7). In contrast, HCT 116 p21(-/-) cells showed a sharp decline in G1 cells and a large increase in S phase cell number after 24 h of hyperoxia. Western blotting revealed that hyperoxia increased p21 in HCT 116 p21(+/+) cells, and it was not detected in HCT 116 p21(-/-) cells (data not shown). The failure of this latter cell type to retain cells in G1 phase indicates that p21 is responsible for the G1 checkpoint during hyperoxia.


View larger version (26K):
[in this window]
[in a new window]
 
Fig. 7.   Cell cycle analysis of HCT 116 cells. A: flow cytometric profiles for parental HCT 116 [p21(+/+)] and isogenic HCT 116-deficient [p21(-/-)] cells exposed to room air (0 h) or hyperoxia for 24, 48, and 72 h. B: mean average percent of cells in G1, S, and G2/M phases at each time point (n = 3 experiments).

ATP levels during exposure to hyperoxia. Although hyperoxia inhibits proliferation in G1 phase through induction of p21, it remains unclear how cells accumulate in S phase. Because S phase checkpoints have yet to be fully defined, we hypothesized that loss of respiratory activity and ATP production could account for the inability of cells to complete S phase. A previous study (1) has shown that ATP levels increase in cells exposed to hyperoxia and then decline as glucose becomes depleted. As shown in Fig. 8, ATP levels in both Mv1Lu and A549 cells increased modestly over the first 48 h of hyperoxia and then declined to room air levels. We conclude that a decline in ATP levels is not likely to account for failure to complete S phase because ATP levels did not decline below room air-exposed levels when S phase growth arrest was observed.


View larger version (20K):
[in this window]
[in a new window]
 
Fig. 8.   ATP levels in Mv1Lu and A549 cells exposed to hyperoxia. Cells were exposed to room air (0 h) or hyperoxia for 16, 24, 48, and 72 h. Cells were harvested and counted, and ATP levels were determined. Data are means ± SD; n = 3 experiments.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Exposure to high atmospheric concentrations of O2 increases the intracellular formation of ROS, which are, in turn, believed to impinge on a wide array of biochemical processes. Some of the ROS-mediated events that have been implicated as possible mechanisms for hyperoxia-induced growth arrest include oxidation of DNA and lipids, chromosomal anomalies, mitochondrial dysfunction, and/or changes in gene expression of cell cycle regulatory proteins. The large number of variables involved, however, has hampered efforts at providing a strong correlation between any one particular phenomenon and the growth arrest process observed during hyperoxia exposure. The objective of this study was to use flow cytometric techniques to test the hypothesis that the block to proliferation imposed by hyperoxia occurs in a cell cycle phase-specific manner. Our findings indicate that hyperoxia inhibits proliferation of epithelial cells through p21-dependent G1 checkpoint activation and a failure to complete S phase.

One of the more prominent cell cycle alterations that occurred in epithelial cells during continued exposure was growth arrest during G1 and in S phase. Throughout the course of these experiments, however, A549 cells had a significantly lower percentage of cells accumulating in S phase then did Mv1Lu cells. The inability to detect any p21 induction in Mv1Lu cells suggested that this cell type may lack a functional G1 delay, a deficiency that, in turn, contributes to its greater degree of S phase arrest during hyperoxia. Support for this argument is that Mv1Lu cells, unlike A549 cells, do not incur any G1 delay after exposure to a single dose of 6 Gy of ionizing radiation. The ability of p21 to exert a G1 checkpoint during hyperoxia was also confirmed with HCT 116 colon carcinoma cells, which express functional p21, and an isogenic line that lacked p21. Our findings corroborate the previous work of Corroyer et al. (7), which demonstrated that hyperoxia increased p21, which blocked cyclin E-cdk2 activity in immortalized rat type II epithelial cells. These authors concluded that the p21-dependent block of cyclin E-cdk 2 activity could be a mechanism by which hyperoxia inhibited proliferation. Using p21-competent and -deficient cell lines, we have now confirmed that p21 is responsible for the G1 growth-arresting activities of hyperoxia.

It is worth discussing that cells with intact p53 and p21 pathways still experience a progressive decline in G1 cell numbers during hyperoxia. Possible reasons for this occurrence may relate to the transient nature of the G1 delay (14), which cannot be sustained over the protracted time period that hyperoxia is administered to cells. Weinert and Lydall (32) have also shown that cells may succumb to growth-stimulating factors in cultured media and make attempts at checkpoint override during prolonged delay (32). In hyperoxia, the decline in the G1 fraction of cells that are released from the G1 delay is also exacerbated by the failure of cells to complete S phase. As a result of failure to complete S phase, cells do not enter mitosis, and the G1 fraction of these cultures is not replenished.

Our findings also reveal that exposure of Mv1Lu and A549 cultures to hyperoxia leads to the accumulation of cells in S phase. Although G1 and G2 cell cycle checkpoints in response to DNA damage have been studied extensively, the only evidence supporting the existence of an S phase checkpoint comes from a study with yeast (31) in which hydroxyurea-induced nucleotide depletion was shown to elicit an arrest in early S phase (31). In this study, the authors isolated several mutant strains (rad53; mec2) that fail to arrest in S phase when exposed to hydroxyurea. This observation supports the concept that certain stress-responsive pathways capable of slowing or halting transcriptional activity during S phase may exist in all eukaryotes. Currently, there is no direct evidence to support or refute the idea that a novel S phase checkpoint is activated during hyperoxia. Our current results do, however, suggest that defects in DNA synthesis caused by hyperoxic toxicity could be responsible for trapping cells in S phase. The BrdU pulse-chase studies indicated that Mv1Lu and A549 cells in S phase exposed to hyperoxia do not abruptly stop DNA synthesis but, instead, exhibit a lowered rate of transit through S phase. Despite their lowered rates of progression, BrdU-positive cells in S phase did eventually complete mitosis, as evidenced by the appearance of BrdU-positive cells in G1 phase at later time points. Unlike G1 and G2 checkpoints, which enact a temporary block toward cell cycle progression, the constraints on S phase movement caused by hyperoxia at early times are not yet severe enough to cause complete arrest. However, continuous exposure to hyperoxia extends the time required to traverse S phase until proliferation ceases.

Because prominent injury to mitochondria is also sustained during hyperoxia, Gille et al. (11) have suggested that the production of metabolic O2 radicals may not cause genomic injury by direct interaction with DNA but by interfering with the cell energy status of the cell. Attempts to proliferate with low ATP levels or unbalanced deoxynucleotide triphosphate pools would not only compromise the replication process but would also lead to breaks in DNA. Hyperoxia-induced genomic instability by indirect means is an attractive argument, considering that exposure to hyperoxia has been shown to have little or no mutagenic potential in a number of studies (e.g., Ref. 11). It is therefore tempting to speculate that the reductions in S phase transit that occur during early hyperoxia exposure and the arrest of cells in S phase at later duration are the result of a nucleotide precursor shortage arising from a compromised energy status. In fact, short-term exposure of A549 cells to hyperoxia resulted in a modest increase in ATP levels that decreased when cell death was detected (1). Decreased ATP levels and death could be attenuated by the addition of glucose to the medium. Although the current study found that hyperoxia increased ATP levels in Mv1Lu and A549 cells, they did not decrease below room air levels, even though cells had arrested in S phase. The ability to maintain ATP levels in the current study compared with a previous study (1) was likely due to the higher level of glucose in our culture medium (4,500 vs. 1,800 mg/l). Nevertheless, the observation that cells accumulated in S phase without losing ATP is not consistent with the concept that S phase growth arrest by hyperoxia is due to ATP depletion. Interestingly, the proliferation of Mv1Lu and A549 cells was also inhibited by the herbicide paraquat, which generates O2 free radicals (data not shown). However, growth arrest was not associated with the same marked accumulation of cells in S phase that is observed when the cells are exposed to hyperoxia. Thus the mechanism by which hyperoxia and associated O2 free radicals exert S phase growth arrest is complex and requires further study.

The G2 checkpoint delay is a universal response that occurs in eukaryotic cells in response to a DNA damage event (32). In contrast to G1 delays, which are of short duration, delays at the G2 position can last up to 12 h depending on cell type and the extent of damage (18). Also, the damage threshold for activating growth arrest appears to be higher for the G1 checkpoint than for the G2 checkpoint (32). Mv1Lu and A549 cells undergo a pronounced G2 delay in response to ionizing radiation (Rancourt, unpublished observations). Neither cell type, however, showed any significant increase in the proportion of cells in G2 phase during the 72-h course of hyperoxia exposure. Although speculative, if hyperoxia is damaging DNA, the failure to observe cells in G2 may be due to the arrest of cells in S phase. Lowering O2 concentrations may attenuate the constraints that cause S phase arrest and thereby unmask the G2 delay. This is consistent with a recent study (26) in which smooth muscle cells exposed to 40% O2 exhibited slightly larger G2 phase accumulations then when exposed to 80% O2.

An important observation of the present study is that hyperoxia-associated growth arrest has several components, including a p21-dependent G1 delay and an S phase arrest. These findings extend earlier observations of increased p53 and p21 expression during hyperoxia exposure by demonstrating that these molecules retain their ability to affect G1 checkpoint control during such conditions. Growth arrest in S phase, however, is not controlled by p21 but results instead from a decrease in the rate of DNA elongation. Although the mechanism causing this latter form of arrest is not clear, it may be related to cytotoxicity or the activation of a novel S phase-specific checkpoint.


    ACKNOWLEDGEMENTS

This work was funded by National Institutes of Health (NIH) Grants CA-73725 (to P. C. Keng) and HL-58774 (to M. A. O'Reilly). R. C. Rancourt and C. E. Helt were supported by NIH Training Grant ES-07026.


    FOOTNOTES

Address for reprint requests and other correspondence: M. A. O'Reilly, Dept. of Pediatrics, Box 777, Children's Hospital at Strong, Univ. of Rochester, 601 Elmwood Ave., Rochester, NY 14642 (E-mail: michael_oreilly{at}urmc.rochester.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 19 May 2000; accepted in final form 19 October 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Allen, CB, and White CW. Glucose modulates cell death due to normobaric hyperoxia by maintaining cellular ATP. Am J Physiol Lung Cell Mol Physiol 274: L159-L164, 1998[Abstract/Free Full Text].

2.   Bassett, DJP, Elbon CL, and Reichenbaugh SS. Respiratory activity of lung mitochondria isolated from oxygen-exposed rats. Am J Physiol Lung Cell Mol Physiol 263: L439-L445, 1992[Abstract/Free Full Text].

3.   Begg, AC, McNally NJ, Shrieve DC, and Karcher HA. A method to measure the duration of DNA synthesis and the potential doubling time from a single sample. Cytometry 6: 620-626, 1985[ISI][Medline].

4.   Boveris, A, and Chance B. The mitochondrial generation of hydrogen peroxide. General properties and effect of hyperbaric oxygen. Biochem J 134: 707-716, 1973[ISI][Medline].

5.   Caldwell, J. Effects of high partial pressures of oxygen on fungi and bacteria. Nature 206: 321-323, 1965[ISI][Medline].

6.   Clement, A, Edeas M, Chadelat K, and Brody JS. Inhibition of lung epithelial cell proliferation by hyperoxia. J Clin Invest 90: 1812-1818, 1992[ISI][Medline].

7.   Corroyer, S, Maitre B, Cazals V, and Clement A. Altered regulation of G1 cyclins in oxidant induced growth arrest of lung alveolar epithelial cells. J Biol Chem 271: 25117-25125, 1996[Abstract/Free Full Text].

8.   Deng, C, Zhang P, Harper JW, Elledge SJ, and Leder P. Mice lacking p21CIP1/Waf1 undergo normal development, but are defective in G1 checkpoint control. Cell 82: 675-684, 1995[ISI][Medline].

9.   Dulic, V, Kaufmann WK, Wilson SJ, Lees E, Harper JW, and Elledge SJ. p53-dependent inhibition of cyclin-dependent kinase activities in human fibroblasts during radiation-induced G1 arrest. Cell 76: 1013-1023, 1994[ISI][Medline].

10.   Gille, JJ, and Joenje H. Cell culture models for oxidative stress: superoxide and hydrogen peroxide vs. normobaric hyperoxia. Mutat Res 275: 405-414, 1992[ISI][Medline].

11.   Gille, JJ, van Berkel CG, and Joenje H. Mutagenicity of metabolic oxygen radicals in mammalian cell cultures. Carcinogenesis 15: 2695-2699, 1994[Abstract].

12.   Gottlieb, TM, and Oren M. p53 in growth control and neoplasia. Biochim Biophys Acta 1287: 77-102, 1996[ISI][Medline].

13.   Kazzaz, JA, Xu J, Palaia TA, Mantell L, Fein MA, and Horowitz S. Cellular oxygen toxicity: oxidant injury without apoptosis. J Biol Chem 271: 15182-15186, 1996[Abstract/Free Full Text].

14.   Leeper, DB, Schneiderman MH, and Dewey WC. Radiation induced cycle delay in synchronized Chinese hamster cells: comparison between DNA synthesis and division. Radiat Res 53: 322-328, 1973.

15.   Linke, SP, Clarkin KC, DiLeonardo A, Tsou A, and Wahl GM. A reversible, p53-dependent G0/G1 cell cycle arrest induced by ribonucleotide depletion in the absence of detectable DNA damage. Genes Dev 10: 934-947, 1996[Abstract].

16.   McGrath, SA. Induction of p21 WAF1/CIP1 during hyperoxia. Am J Respir Cell Mol Biol 18: 179-187, 1998[Abstract/Free Full Text].

17.   Meek, DW. Posttranslational modification of p53. Semin Cancer Biol 5: 203-210, 1994[ISI][Medline].

18.   Nagasawa, H, Keng P, Harley R, Dahlberg W, and Little JB. Relationship between gamma-ray-induced G2/M delay and cellular radiosensitivity. Int J Radiat Biol 66: 373-379, 1994[ISI][Medline].

19.   O'Reilly, MA, Staversky RJ, Stripp BR, and Finkelstein JN. Exposure to oxygen induces p53 expression in mouse lung epithelium. Am J Respir Cell Mol Biol 18: 43-50, 1998[Abstract/Free Full Text].

20.   O'Reilly, MA, Staversky RJ, Watkins RH, and Maniscalco WM. Accumulation of p21(Cip1/WAF1) during hyperoxic lung injury in mice. Am J Respir Cell Mol Biol 19: 777-785, 1998[Abstract/Free Full Text].

21.   Polyak, K, Waldman T, He TC, Kinzler KW, and Vogelstein B. Genetic determinants of p53-induced apoptosis and growth arrest. Genes Dev 10: 1945-1952, 1996[Abstract].

22.   Rancourt, RC, Keng PC, and O'Reilly MA. Hyperoxia inhibits proliferation of Mv1Lu epithelial cells independent of TGF-beta signaling. Am J Physiol Lung Cell Mol Physiol 277: L1172-L1178, 1999[Abstract/Free Full Text].

23.   Renzig, J, and Lane DP. p53-dependent growth arrest following calcium phosphate-mediated transfection of murine fibroblasts. Oncogene 10: 1865-1868, 1995[ISI][Medline].

24.   Rueckert, RR, and Mueller CJ. Effect of oxygen tension on HeLa cell growth. Cancer Res 20: 944-949, 1960[ISI].

25.   Schoonen, WGEJ, Wanamarta AH, van der Klei-van Moorsel JM, Jakobs C, and Joenje H. Hyperoxia-induced clonogenic killing of HeLa cells is associated with respiratory failure and selective inactivation of Krebs cycle enzymes. Mutat Res 237: 173-181, 1990[ISI][Medline].

26.   Shenberger, JS, and Dixon PS. Oxygen induces S-phase growth arrest and increases p53 and p21WAF1/CIP1 expression in human bronchial smooth-muscle cells. Am J Respir Cell Mol Biol 21: 395-402, 1999[Abstract/Free Full Text].

27.   Turrens, JF, Freeman B, Levitt JG, and Crapo JD. The effect of hyperoxia on superoxide production by lung submitochondrial particles. Arch Biochem Biophys 217: 401-410, 1982[ISI][Medline].

28.   Waga, S, Hannon GJ, Beach D, and Stillman B. The p21 inhibitor of cyclin dependent kinases controls DNA replication by interaction with PCNA. Nature 364: 574-578, 1994[ISI].

29.   Waldman, T, Kinzler KW, and Vogelstein B. p21 is necessary for the p53-mediated G1 arrest in human cancer cells. Cancer Res 55: 5187-5190, 1995[Abstract].

30.   Wang, Y, and Eckhart W. Phosphorylation sites in the amino-terminal region of mouse p53. Proc Natl Acad Sci USA 89: 4231-4235, 1992[Abstract].

31.   Weinert, TA, Kiser GL, and Hartwell LH. Mitotic checkpoint genes in budding yeast and the dependence of mitosis on DNA replication and repair. Genes Dev 8: 652-655, 1994[Abstract].

32.   Weinert, TA, and Lydall D. Cell cycle checkpoints, genetic instability and cancer. Semin Cancer Biol 4: 129-140, 1993[ISI][Medline].


Am J Physiol Lung Cell Mol Physiol 280(4):L617-L626
1040-0605/01 $5.00 Copyright © 2001 the American Physiological Society