Departments of Pediatrics and Environmental Medicine, The University of Rochester, Rochester, New York 14642
Received April 17, 2001; accepted June 20, 2001
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ABSTRACT |
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Key Words: cell cycle; DNA damage; oxidative stress; reactive oxygen species (ROS).
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INTRODUCTION |
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The lung is particularly affected by oxidative stress, because it functions to exchange oxygen with the environment and blood. Higher oxygen tensions used to reduce tissue hypoxia, as well as inhaled pollutants such as nitrogen dioxide and ozone, injure and kill pulmonary cells. Since hyperoxia produces oxidant-free radicals (Zweier et al., 1989), many investigators use hydrogen peroxide as an in vitro model for hyperoxia-induced lung injury. Even though both can damage DNA, it is unlikely that hydrogen peroxide mimics the toxic effects of oxygen for several reasons. Studies in Chinese hamster ovary cells revealed that hydrogen peroxide produces mutagenic single-strand DNA damage, whereas hyperoxia causes sister chromatid exchanges and other chromosome aberrations (Gille et al., 1989
). Iron chelators enhance survival of cells exposed to hydrogen peroxide, but not when exposed to hyperoxia (Gille et al., 1992
). Furthermore, hydrogen peroxide causes acute injury because it has a short half-life. In contrast, hyperoxia-induced stress may be continuously maintained until the cells are returned to a normoxic environment. Because hyperoxia is used clinically to reduce tissue hypoxia as well as provide a useful model for cancer or aging processes (Gille and Joenje, 1992
), it is important to understand how cells respond to its toxic effects. Although numerous studies in both rodents (Barazzone et al., 1998
; O'Reilly et al., 2000
) and a variety of cultured epithelial cell lines (Kazzaz et al., 1996
; Rancourt et al., 2001
, Rancourt et al., 1999
) have shown that oxygen, by >90%, kills cells by necrosis and not apoptosis, little is known about how cells prevent oxidant damage.
One of the earliest studies examining the cellular response to hyperoxia revealed that it inhibited proliferation of HeLa cells (Rueckert and Mueller, 1960). Clearly, preventing DNA replication during periods of genotoxic stress would be advantageous. Unfortunately, the mechanism by which cells cease proliferation during hyperoxia remained unclear until recently. Several studies in adult mice and cultured cell lines have now shown that >90% oxygen levels increase expression of the tumor suppressor protein p53 and its phosphorylation on serine 15 (Barazzone et al., 1998
; O'Reilly et al., 1998a
; Rancourt et al., 2001
; Shenberger and Dixon, 1999
). DNA damage activates a number of kinases that phosphorylate p53 at several sites, including serine 15. Phosphorylation at this site prevents mdm2 from binding p53 and targeting it for ubiquitin-mediated degradation (Shieh et al., 1997
). Hyperoxia also increases expression of the cyclin-dependent kinase inhibitor p21Cip1/WAF1/Sdi1 (p21; see O'Reilly et al., 1998b
; Rancourt et al., 2001
; Shenberger and Dixon, 1999
), which may be transcriptionally induced by p53 and various cytokines and steroids, or posttranscriptionally stabilized by ROS (Bellido et al., 1998
; Datto et al., 1995
; el-Deiry et al., 1993
). p21 inhibits cell proliferation by binding and inactivating G1 and S cyclin-dependent kinases (cdk; Luo et al., 1995
). It also binds proliferating cell nuclear antigen PCNA and blocks DNA polymerase activity. A role for p21 in the growth-arresting activities of hyperoxia was first described in SV40 transformed type-II epithelial cells. In that study, hyperoxia increased p21, which inhibited cyclin E/cdk2 kinase activity (Corroyer et al., 1996
). Active cyclin E/cdk2 kinase phosphorylates substrates that are required for DNA replication (Elledge, 1996
). Additional studies using a variety of p21-functional and deficient epithelial cell lines revealed that cells expressing p21 accumulated in G1 while cells lacking p21 exited G1 and accumulated in S phase (Rancourt et al., 2001
). Analogous studies revealed that hyperoxia inhibited cell proliferation in lungs of adult mice in vivo, but not in mice lacking p21 (O'Reilly et al., 2001
). Interestingly, the p21-deficient mice were markedly sensitive to hyperoxia, as shown by rapid necrotic death of parenchymal cells and an
50% reduction in mean survival time. This observation was consistent with other studies showing that p21 protects cells from the genotoxic effects of ionizing radiation, ultraviolet light, cisplatin, methylmethane sulfonate, and nitrogen mustard (Fan et al., 1997
; McDonald et al., 1996
; Sheikh et al., 1997
; Wang et al., 1997
). Because the mammalian lung contains nearly 40 distinct cell types, a simple cell-line model would be useful for clarifying how p21 protects cells from oxygen-induced toxicity.
The current study examines the toxic effects of hyperoxia on HCT116 colon carcinoma cells that express wild-types p53 and p21. An advantage of these cells over existing pulmonary cell lines is that clonal variants lacking p53 or p21 were created by homologous recombination (Bunz et al., 1998; Waldman et al., 1995
). This allows direct toxicologic comparisons between genetically identical cells without additional complications involving unknown genetic differences that may exist when studying nonhomologous cell lines. We now show that hyperoxia induces p53-dependent expression of p21, which prevents cells from exiting G1 as well as enhancing their survival. Our findings are consistent with the importance of limiting DNA replication during oxidant genotoxic stress and suggest that repair of oxidant DNA damage may be critical for survival. Since hyperoxia damages cells indirectly through production of toxic ROS, our findings with these colon cells may also have clinical significance for patients suffering from ROS-mediated inflammatory or ischemic bowel injuries.
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MATERIALS AND METHODS |
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Western-blot analysis.
Cells were harvested by trypsinization, followed by centrifugation at 300 x g for 6 min. The pellets were then washed with 1x phosphate-buffered saline (PBS, Life Technologies) and resuspended 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 phenylmethylsulfonyl fluoride. The lysate was cleared by centrifugation at maximum speed in a microcentrifuge and protein concentrations were determined by the Lowry Assay (DC Protein Assay, Bio-Rad, Hercules, CA). The lysates were boiled for five min in 3x Laemmli buffer (1x Laemmli contains 50 mM Tris pH 6.8, 1% ß-mercaptoethanol, 2% SDS, 0.1% bromophenol blue, and 10% glycerol). The extracted protein was separated by SDSpolyacrylamide gel electrophoresis and transferred to polyvinylidene difluoride membranes (PVDF, Millipore, Bedford MA). The membranes were blocked for 1 h at room temperature in phosphate or Tris-buffered saline (PBS or TBS) containing 5% nonfat dry milk plus 0.05% or 0.1% Tween-20 (PBST or TBST), respectively. TBST was used to wash membranes intended for phosphorylation-specific antisera. The membranes were then incubated in primary antibody p53 (1:1000, Oncogene Research Products, Cambridge, MA), p53 (ser-15; 1:1000, Cell Signaling Technology, Beverly, MA), or p21 (1:500, PharMingen, San Diego, CA), followed by 3 washes in PBS or TBS to remove nonspecific interactions. Membranes were then incubated in secondary antibody (p53 or p21:goat antimouse; Southern Biotechnology, Birmingham, AL or p53 (ser-15): goat antirabbit; Jackson Labs, West Grove, PA) for one h at room temperature. After washing the membranes with PBST or TBST, specific antibody interactions were visualized by chemiluminescence (Amersham, Arlington Heights, IL). As a loading control, membranes were also blotted for ß-actin (1:5000; Sigma, St. Louis, MO).
Growth curve and cell viability assays.
Cells were trypsinized, counted with a hemacytometer, and plated in triplicate in 60 mm culture dishes before exposing to room air or hyperoxia for 24, 48, 72, or 96 h. A hemacytometer was used to count cells and viability was determined by their ability to exclude 0.04% trypan blue dye. Colony-survival assays were initiated by exposing cells to room air or hyperoxia for 24 or 36 h. Cells were then trypsinized and counted using a hemacytometer. Serial dilutions were replated at 100 cells per 60 mm culture dish and allowed to grow in room air at 37°C for 13 days, at which time colonies were fixed in 15% methanol and stained with crystal violet (2.5 g/l). The total number of colonies produced on each plate was then recorded.
Flow cytometric cell cycle analysis.
HCT116 cells exposed to room air or hyperoxia were trypsinized, resuspended in their own media, and centrifuged at 300 x g for 6 min. Pellets were resuspended in 75% ethanol and stored at 4°C for a minimum of 24 h. Fixed cells were then centrifuged at 300 x g for 6 min. The pellets were resuspended in PBS containing 1 mg/ml RNase (Sigma, St. Louis, MO) and incubated at room temperature for 30 min, with intermittent vortexing. Cells were centrifuged again at 300 x g. The pellets were resuspended in PBS containing 20 µg/ml propidium iodide. Stained samples were filtered through a 37 µm mesh prior to analysis on an Epics Elite (Coulter Electronics) equipped with an argon laser excited at 488 nm. Relative DNA content per cell was determined by measuring fluorescence of the treated samples. Cell-phase percentages were determined by use of Multicycle by Phoenix Flow Systems software (Phoenix Flow Systems, San Diego, CA).
RNA extraction and analysis.
Cells grown on culture dishes were lysed by adding 1 ml of 4 M guanidine isothiocyanate, 0.5% N-laurylsarcosine, 20 mM sodium citrate, and 0.1 M ß-mercaptoethanol followed by scraping the dish surface. RNA was extracted from the resulting solution using acid phenol and phase-lock gel columns (5 Prime-3 Prime, Boulder, Colorado) and resuspended in diethylpyrocarbonate-treated water. The amount of RNA in an aqueous solution was determined by absorbance at 260 nm. RNase protection assays were performed with human cell cycle-2 multi-probe template kit, according to the manufacturer's instructions (PharMingen, San Diego, CA). Riboprobe synthesis reaction was incubated for 60 min at room temperature, followed by an additional 30 min at 37°C in the presence of 2 units of RNase-free DNase. Riboprobes were extracted with phenol/chloroform and precipitated with ethanol and ammonium acetate. Probes were resuspended in 50 µl of hybridization buffer (400 mM NaCl, 40 mM PIPES, pH 6.7, 1 mM EDTA, pH 8.0, and 80% formamide) and diluted to 3.37 x 105 cpm/µl before incubating with 5 µg denatured total RNA at 56°C for 16 h. Hybridized probes were digested with RNase buffer for 45 min at 30°C. Samples were incubated with proteinase K and yeast tRNA before extracting with phenol/chloroform and precipitating with ethanol. Protected products were separated on a 6% acrylamide/8 M urea sequencing gel, dried, and visualized by exposure on PhosphorImager screens.
Statistical analysis.
Values are expressed as means ± SD. Group means were compared by ANOVA with Fisher's procedure post hoc analysis, using Excel software for the Macintosh with p < 0.05 considered as significant.
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RESULTS |
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p21 Prevents Exit from G1 during Hyperoxia
The effect of hyperoxia on cell proliferation was determined by counting cells with a hemacytometer (Fig. 3). Cultures of parental HCT116 cells or those lacking p53 or p21 showed a linear increase in cell number over time when exposed to room air. In contrast, cell numbers did not increase in cultures exposed to hyperoxia. The inhibitory effects of hyperoxia were observed in the parental cells and in both the p53 and p21-deficient lines after 48 h of exposure.
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DISCUSSION |
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One of the important observations in the current study was that p53 was necessary for the induction of p21. Previous studies exposing p53-deficient mice to 92% oxygen revealed that p21 induction was dependent upon p53. This is consistent with our recent finding that hyperoxia activates a p53-dependent transcriptional reporter in cultured cell lines (Rancourt et al., 2001). On the other hand, we also showed that modest levels of p21 were still induced in p53-deficient mice who were exposed to 100% FiO2 in vivo, while maximal expression was detected in p53-wild-type mice. The latter finding is consistent with studies showing that hyperoxia increases p21 in SV40-transformed pulmonary epithelial cells that presumably contain nonfunctional p53, due to their expression of SV40 (Corroyer et al., 1996
). One explanation for these differences is that p21 is regulated by multiple signal transduction pathways. For example, oxidative stress can increase the dithiol-reducing protein thioredoxin, which potentiates redox factor (Ref)-1-dependent regulation of p53 and p21 (Ueno et al., 1999
). p53-independent changes in p21 mRNA stability have been reported in cells treated with diethylmaleate, a compound that oxidizes cells by depleting them of reduced glutathione (Esposito et al., 1997
). The cytokines transforming growth factor (TGF)-ß and interleukin (IL)-6 increase p53-independent transcription of p21 in keratinocytes and osteoblast cells, respectively (Bellido et al., 1998
; Datto et al., 1995
). Interestingly, the expression of both cytokines increases in mouse lungs exposed to hyperoxia (Johnston et al., 1998
; O'Reilly et al., 1997
). Additional response elements have been identified in the p21 promoter, which mediate regulation by phorbol esters, steroids, cisplatin, epidermal growth factor, and UV radiation (Gartel and Tyner, 1999
). Although multiple pathways exist to regulate p21, all of the available evidence suggests that p53 participates in the induction of p21 by hyperoxia, because it is absolutely required in the HCT116 cell line model and is required for maximal induction in vivo.
The current study confirmed previous observations that lethal levels of oxygen inhibit cell proliferation in both animal models (Evans et al., 1969) and cultured cell lines (Rueckert and Mueller, 1960
). More recent findings revealed that hyperoxia inhibits proliferation in G1 phase of the cell cycle through induction of p21 (Corroyer et al., 1996
; Rancourt et al., 2001
). Cells that fail to induce p21 exit G1 where they arrest predominantly in S phase. As expected, the percent of p21-deficient HCT116 cells in G1 phase of the cell cycle decreased when they were exposed to hyperoxia compared to the parental cells that expressed p21. This decrease in the G1 population correlated with an increase in the percentage of cells in S phase. Although hyperoxia also decreased the percent of p53-deficient HCT116 cells in G1, the percent of cells in S phase remained constant while the G2/M population increased significantly. It remains unclear why a greater percentage of p53-deficient cells were able to successfully complete S phase, because the mechanism by which hyperoxia causes cells to accumulate in S phase remains unknown. Nonetheless, failure to induce p21 during hyperoxia is associated with a significant percentage of cells exiting the G1 compartment. Cells that vacate G1 enter S phase, where they either accumulate or successfully progress through and accumulate in G2.
Another important observation in the current study is that p21 protects cells from the toxic effects of oxygen exposure. This finding is consistent with our recent observation that p21 enhanced survival of adult mice exposed to 100% FiO2 (O'Reilly et al., 2001). The mean survival of p21-deficient mice was 72 h compared to 120 h for p21-wild-type mice. Ultrastructural analyses of their lungs revealed rapid necrosis of alveolar endothelial and of type I epithelial cells that lead to pulmonary edema. The same cell types are injured and killed in p21-wild-type mice after longer periods of exposure. These observations suggest that p21 protects cells from hyperoxia by delaying necrosis, rather than by changing the population of cells that are sensitive to hyperoxia. The current study extends these findings by showing that p21 also delayed death of HCT116 cells. To date, there is no evidence that hyperoxia kills cells in vitro by apoptosis (Kazzaz et al., 1996
; Rancourt et al., 1999
) or in vivo as shown by ultrastructural findings (O'Reilly et al., 2000
, 2001
). ROS may cause more damage to p21-deficient cells because they are trapped in S phase where their DNA is actively replicating and not protected by histones. In contrast, the modest increase in survival of the p53-deficient compared to p21-deficient cells may be attributed to the larger percentage of p53-deficient cells that successfully complete S phase and arrest in G2/M. Cells that arrest in G2 are also more resistant to genotoxins, presumably because they can repair damaged DNA before cytokinesis.
p21 may protect cells from oxygen-induced toxicity by preventing them from replicating their damaged DNA before it has been repaired. The observation that, cultures that express p21 retain a greater percentage of cells in G1 compared to cells that lack p21, supports this hypothesis. Alternatively, p21 may participate in DNA repair activities through its ability to bind PCNA that interacts with FEN-1 and DNA polymerase. p21 may function in nucleotide excision repair because p21-deficient HCT116 cells have a reduced capacity to repair DNA damaged by UV or cisplatin (McDonald et al., 1996). Similar results were found when cells were challenged with nitrogen mustard, which also damages DNA (Fan et al., 1997
). p21 also enhanced protection and repair from UV-induced DNA damage in DLD1 colorectal carcinoma cells (Sheikh et al., 1997
). In contrast, p21-deficient mouse embryo fibroblasts exhibited minimal repair capacity when damaged by UV (Smith et al., 2000
). Even though the biochemical evidence supports the argument that p21 participates in DNA repair processes, this issue remains to be clarified especially in the context of oxidative DNA damage.
Another important finding in this study was that hyperoxia only affected the expression of p21 mRNA. As shown in Figure 2, hyperoxia did not markedly alter mRNA levels of p27 or p57, both of which share structural homology to p21. Previous studies using rat type-II epithelial cells showed that hyperoxia increased p27 mRNA but not p27 protein (Corroyer et al., 1996
). The current study also showed that hyperoxia did not alter mRNA of the INK4 proteins p15, p16, p18, or p19. The INK4 proteins prevent S-phase progression by binding cdk4 and preventing association with its catalytic partner, cyclin D. Finally, hyperoxia did not alter expression of Rb or its related members p107 or p130. Interestingly, loss of p53 or p21 did not effect the basal expression of the Kip or INK4 genes or when the cells were exposed to hyperoxia. Thus, the effect of hyperoxia on mRNA levels of G1 kinase inhibitors is specific for p21 and is dependent upon functional p53.
In summary, the current study demonstrates that hyperoxia induces p53-dependent expression of p21 that prevented cells from exiting G1 and enhanced their survival. These findings are consistent with an in vivo mouse model of oxygen-induced lung injury that showed p21 enhances survival. Both models may now be exploited to understand how hyperoxia activates the p53 suppressor pathway and how its downstream targets, such as p21, modify the genotoxic effects of oxygen. A better understanding of how cells respond to the toxic effects of hyperoxia can provide insight into how to block the damaging effects of ROS associated with supplemental oxygen exposure, reperfusion injury, cancer, and the aging process.
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ACKNOWLEDGMENTS |
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NOTES |
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