Hyperoxia inhibits proliferation of Mv1Lu epithelial cells independent of TGF-beta signaling

Raymond C. Rancourt1, Rhonda J. Staversky2, Peter C. Keng3, and Michael A. O'reilly1,2

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


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

High concentrations of O2 inhibit epithelial cell proliferation that resumes on recovery in room air. To determine whether growth arrest is mediated by transforming growth factor-beta (TGF-beta ), changes in cell proliferation during exposure to hyperoxia were assessed in the mink lung epithelial cell line Mv1Lu and the clonal variant R1B, which is deficient for the type I TGF-beta receptor. Mv1Lu cells treated with TGF-beta accumulated in the G1 phase of the cell cycle as determined by propidium iodide staining, whereas proliferation of R1B cells was unaffected by TGF-beta . In contrast, hyperoxia inhibited proliferation of both cell lines within 24 h of exposure through an accumulation in the S phase. Mv1Lu cells treated with TGF-beta and exposed to hyperoxia accumulated in the G1 phase, suggesting that TGF-beta can inhibit the S phase accumulation observed with hyperoxia alone. Cyclin A was detected in cultures exposed to room air or growth arrested by hyperoxia while decreasing in cells growth arrested in the G1 phase by TGF-beta . Finally, hyperoxia failed to activate a TGF-beta -dependent transcriptional reporter in both Mv1Lu and R1B cells. These findings reveal that simple growth arrest by hyperoxia involves a defect in S phase progression that is independent of TGF-beta signaling.

proliferation; transforming growth factor-beta


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

THERE HAS RECENTLY BEEN a greater appreciation for the role of reactive oxygen species (ROS) in regulating normal and abnormal disease processes. The lung is uniquely challenged by ROS because its large surface area is exposed to various environmental pollutants, inflammatory cells are recruited during infection, and supplemental O2 is used clinically to treat pulmonary disease. Studies (2, 7, 8) over the past 30 years in mice, rats, monkeys, and humans have described the effects of lethal (>90%) and sublethal (<85%) levels of O2 on the lung alveolus. The alveolus is composed of type I and II epithelial and microvascular endothelial cells. Endothelial and type I epithelial cells are rapidly killed when exposed to lethal levels of O2. Type II cells, which synthesize pulmonary surfactant, proliferate and differentiate into type I cells during recovery in room air (2, 30). Rapid proliferation of type II cells is crucial for both the survival and prevention of fibrosis (1). Similarly, sublethal levels of O2 kill endothelial cells. However, unlike during exposure to lethal levels of O2, type I epithelial cells survive and type II cells proliferate during the exposure (8). Because proliferation of type II cells plays a critical role in pulmonary repair processes, it is important to clarify molecular signals that regulate type II cell proliferation during and after oxidant stress.

Regulation of cell proliferation is complex and requires the interactions of intracellular and extracellular stimulatory and inhibitory signals that modulate activity of the cyclin-dependent kinases (Cdks). Cdk complexes are composed of a kinase subunit and a catalytic subunit termed cyclin because its expression cycles during the cell cycle (29). The G1 phase cyclins include cyclins D (D1, D2, and D3) and cyclin E, which bind different Cdks. Expression of cyclin A is observed in the S and early G2 phases and cyclin B in the G2/M phase. Active cyclin D and E complexes phosphorylate the retinoblastoma (Rb) gene product, resulting in release of the transcription factor E2F that increases transcription of S phase genes such as thymidine kinase. Growth inhibition is achieved through binding of small Cdk inhibitory proteins (CKIs) to cyclin-Cdk complexes, resulting in decreased kinase activity (29). Two families of CKIs have been identified and include the Cip/Kip family (p21, p27, and p57) and the Ink4 family (p15, p16, p18, and p19).

Antimitogens such as the cytokine transforming growth factor-beta (TGF-beta ) inhibit cell cycle progression by altering the expression and activities of CKIs. TGF-beta first binds to the type II TGF-beta receptor (Tbeta R-II), which then recruits and activates the kinase domain of the type I TGF-beta receptor (Tbeta R-I) through phosphorylation (13). The activated receptor complex phosphorylates various Smad proteins that translocate to the nucleus and bind and transactivate downstream target genes. TGF-beta inhibits cell cycle progression at the G1/S phase boundary by increasing the expression of p15 or p21 (9, 26). Cells arrested by TGF-beta have reduced levels of cyclin A associated with increased levels of hypophosphorylated Rb gene product (27).

Molecular signals that regulate type II cell proliferation remain unknown, in part, because type II cells rapidly lose their differentiated characteristics when cultured in vitro (28). Although simian virus 40 (SV40) and other viral oncoproteins alter normal cell proliferation by rearranging cyclin and Cdk partners, several investigators have immortalized type II cells using SV40. The proliferative response of SV40-immortalized rat type II cells (SV40T-T2) exposed to hyperoxia has been extensively studied. SV40T-T2 cells rapidly growth arrest when exposed to hyperoxia and resume proliferation on recovery in room air (5). Hyperoxia increased mRNA levels of TGF-beta 1, Tbeta R-I, and Tbeta R-II (4, 6). Moreover, hyperoxia increased the expression of p21 that bound and inhibited the activity of cyclin E-Cdk 2 complexes. It was concluded that hyperoxia inhibited proliferation through TGF-beta signaling because increased cyclin E-Cdk2 activity was observed in cells exposed to both hyperoxia and neutralizing antibodies to TGF-beta . Although these studies suggested that hyperoxia inhibits proliferation through TGF-beta signaling, they never demonstrated that administration of neutralizing antibodies resulted in normal cell proliferation in the presence of hyperoxia. In addition, it remains to be determined where in the cell cycle hyperoxia inhibits proliferation.

The present study investigates the effects of hyperoxia on proliferation of the mink lung epithelial cell line Mv1Lu, which is markedly growth arrested by TGF-beta (31). This cell line was chosen because chemically induced mutant lines that are unresponsive to TGF-beta have also been identified (31). The R1B cell line is a member of the R class of mutant Mv1Lu cells that have lost expression of Tbeta R-I. In the present study, we tested the hypothesis that hyperoxia inhibits proliferation through TGF-beta signaling by analyzing proliferation of Mv1Lu and R1B cells exposed to hyperoxia. Our findings reveal that hyperoxia caused both cell lines to cease proliferation in the S phase of the cell cycle independent of TGF-beta signaling.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture. Mv1Lu (mink lung adenocarcinoma) cells were obtained from Dr. Anita Roberts (National Cancer Institute, National Institutes of Health, Bethesda, MD), and R1B (chemically induced mutant Mv1Lu) cells were obtained from Dr. Joan Massagué (Howard Hughes Medical Institute, Memorial Sloan-Kettering Cancer Center, New York, NY). The cells were incubated at 37°C in 5% CO2 in Dulbecco's modified Eagle's medium with 10% fetal bovine serum, 50 U/ml of penicillin, and 50 µg/ml of streptomycin (GIBCO BRL). The cells were maintained in tissue culture flasks and routinely passaged every 3 days.

For exposures to hyperoxia or TGF-beta , the cells were trypsinized, counted with a hemacytometer, and plated in 100-mm dishes at a density of 5 × 105 overnight. The medium was replenished the next morning, at which time the cells were treated with 5 ng/ml of porcine TGF-beta 1 obtained from R&D Systems (Minneapolis, MN) and/or exposed to hyperoxia 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). The cells were harvested at various times with 0.25% trypsin, counted with a hemacytometer, and stained for viability with 0.5% trypan blue or 10 µg/ml of propidium iodide.

Flow cytometry. Cells were trypsinized, resuspended in their original medium, and centrifuged at 300 g. The medium was removed, and the cells were fixed in 75% ethanol for 24 h. The cells were resuspended in 1 ml of RNase (1 mg/ml) for 30 min, centrifuged, and resuspended in 0.5 ml of propidium iodide (10 µg/ml). The samples were analyzed on an Epics Profile (Coulter Electronics, Hialeah, FL) set to collect 10,000 events. DNA histograms were analyzed, and the percentages of G1, S, and G2/M phase cells were determined according to the mathematical model of Fried et al. (11). Terminal deoxynucleotidyltransferase dUTP nick end-labeling (TUNEL) staining was performed with the Apo-BRDU kit obtained from Phoenix Flow Systems (San Diego, CA), and fluorescent-positive cells were measured by flow cytometry. As a positive control for TUNEL staining, the cells were exposed to 5 Gy of 137Cs at a dose rate of 3.7 Gy/min and recovered in room air for 24 h.

Western blot analysis. The cells were harvested at 4°C by scraping in 50 mM Tris, pH 7.4, 150 mM sodium chloride, 2 mM EDTA, 25 mM sodium fluoride, 25 mM beta -glycerol phosphate, 0.1 mM sodium vanadate, 1 mM phenylmethylsulfonyl fluoride, 0.2% Triton X-100, 0.3% Nonidet P-40, 10 µg/ml of leupeptin, 10 µg/ml of pepstatin, and 10 µg/ml of aprotinin. The cell lysates were cleared by centrifugation, and protein concentrations were determined with a modified Lowry assay (Bio-Rad, Hercules, CA) with bovine serum albumin as a standard. The lysates were boiled in 2× Laemmli buffer (1× is 62.5 mM Tris, pH 6.8, 2% SDS, 10% glycerol, 0.025% bromphenol blue, and 5% beta -mercaptoethanol). Proteins (10 µg/ml) were separated by size on polyacrylamide-SDS gels and transferred to nitrocellulose. The membranes were blocked in PBS containing 5% nonfat dry milk overnight at 4°C before incubation with an anti-cyclin A antibody (Santa Cruz Biotechnology, Santa Cruz, CA) at a 1:1,000 dilution for 1 h at room temperature. Nonspecific interactions were removed by washing in PBS containing 0.05% Tween 20 before the blots were incubated in goat anti-rabbit peroxidase-conjugated secondary antibody at 1:5,000 (Jackson ImmunoResearch Laboratories, West Grove, PA). The blots were extensively washed again, and the conjugates were visualized with chemiluminescence (Amersham, Arlington Heights, IL) by exposure to Kodak Bio-Max film. The blots were reblotted with anti-beta -actin antibody (Sigma, St. Louis MO) at a 1:5,000 dilution as a loading control.

TGF-beta -inducible luciferase reporter assays. The cells were transfected with the TGF-beta inducible reporter p3TP-Lux with calcium phosphate as previously described (21). The p3TP-Lux reporter contains three 12-O-tetradecanoylphorbol 13-acetate (TPA) response elements (TRE) from the human collagenase promoter and the TGF-beta responsive element from the plasminogen activator inhibitor-1 promoter ligated upstream to the adenovirus E4 minimal promoter (31). Transfection efficiencies were normalized with the pRL-TK vector (Promega, Madison, WI) that expresses the Renilla luciferase gene. Luciferase assays were performed with the dual-luciferase reporter assay system (Promega) and measured with a luminometer from Tropix (Bedford, MA).

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


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Effects of TGF-beta on cell cycle distribution of Mv1Lu and R1B cells. Mv1Lu and R1B cells were incubated for 24 h in the absence and presence of 5 ng/ml of TGF-beta 1 and harvested, and genomic DNA was stained with propidium iodide. Flow cytometric analysis revealed that asynchronous cultures of Mv1Lu cells cultured in room air contained cells in all phases of the cell cycle. Approximately 55% of the cells were in the G1 [diploid (2n) of DNA where n is haploid] phase, 30% in the S phase, and 15% in the G2 (4n) phase (Table 1). Mv1Lu cells treated with TGF-beta accumulated in the G1 phase of the cell cycle, with a significant decrease in the percentage of cells in the S and G2 phases. In fact, nearly 90% of TGF-beta -treated cells accumulated in the G1 phase. Although asynchronous cultures of R1B cells had a similar cell cycle distribution as Mv1Lu cells when cultured in room air, they were unaffected by exposure to TGF-beta (Table 1). These findings confirm a previous study (31) that demonstrated that the Mv1Lu clonal variant R1B cell line is unresponsive to TGF-beta .

                              
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Table 1.   Effects of TGF-beta and hyperoxia on cell cycle distribution of Mv1Lu and R1B cells

Hyperoxia inhibits Mv1Lu and R1B proliferation. Subconfluent cultures of Mv1Lu and R1B cells were exposed to room air or hyperoxia to determine whether hyperoxia inhibited their proliferation. The cells were harvested every 24 h and counted with a hemacytometer. Mv1Lu and R1B cultures had an increasing number of cells over time when incubated under normoxic conditions (Fig. 1). In contrast, the cell number did not increase in cultures exposed to hyperoxia. The effects of hyperoxia on total cell number were distinguishable within the first 24 h of exposure.


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Fig. 1.   Hyperoxia inhibits proliferation of Mv1Lu (A) and R1B (B) cells. Cells were cultured for indicated times in room air with 5% CO2 or hyperoxia (95% O2-5% CO2), harvested, and counted. Values are means ± SD of 3 cultures for each time point. Hyperoxia significantly inhibited total cell number of both cell lines by 24 h of exposure, P < 0.05 by ANOVA.

Changes in total cell number in the presence of hyperoxia could be due to decreased cell proliferation or increased cell death. Although Mv1Lu and R1B cells remain attached to their plates for the first 72 h of exposure, trypan blue dye exclusion was used to quantitatively measure cell viability. More than 90% of the cells exposed to hyperoxia for up to 72 h continued to possess good membrane integrity based on their ability to exclude dye (Fig. 2A). Similarly, R1B cells maintained membrane integrity over this time period (data not shown). Although membrane integrity remains a reasonable method to determine the viability of necrotic cells, it does not adequately detect apoptotic cells (25). TUNEL staining was used to assess apoptosis in Mv1Lu cells exposed to hyperoxia. Minimal TUNEL-positive cells were observed in cultures exposed to room air or hyperoxia (Fig. 2B). Hyperoxia also does not induce DNA laddering in Mv1Lu cells (data not shown). As a positive control for apoptotic cell death, Mv1Lu cells were exposed to 5 Gy of ionizing radiation. In contrast to the effects of hyperoxia, nearly 90% of Mv1Lu cells exposed to radiation became TUNEL positive. TUNEL staining was not measured in R1B cells because this finding and a previous study (16) have shown that hyperoxia kills cells in vitro through necrosis. Collectively, our findings suggest that the effects of hyperoxia on cell number were not due to increased cell death.


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Fig. 2.   Effect of hyperoxia on cell viability. Mv1Lu cells were exposed to room air or hyperoxia for indicated times. Cells were harvested, stained with trypan blue, and counted with a hemacytometer. A: values are means ± SD of cells from triplicate cultures excluding trypan blue. B: percentage of terminal deoxynucleotidyltransferase dUTP nick end labeled (TUNEL)-positive cells was assessed in Mv1Lu cells exposed to room air or hyperoxia for indicated times. Hatched bar, percentage of TUNEL-positive cells after exposure to 5 Gy of ionizing radiation.

Mv1Lu and R1B cells were exposed to room air or hyperoxia for 24 h and analyzed by flow cytometry to determine whether hyperoxia altered their cell cycle distribution. Asynchronous cultures of Mv1Lu and R1B cells cultured in room air had cells in all phases of the cell cycle (Fig. 3). In contrast, cultures of both cell lines exposed to hyperoxia for 24 h had a marked increase in the percentage of cells in the S phase associated with a decrease in the percentage of cells in the G1 phase. In fact, the percentage of cells in the S phase nearly doubled after 24 h of exposure (Table 1). Continued exposure to hyperoxia for 48 and 72 h resulted in a continued increase in the percentage of cells in the S phase (Fig. 4).


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Fig. 3.   Flow cytometric analysis of cells cultured in normoxia or hyperoxia. Mv1Lu and R1B cells were exposed to room air (room air-5% CO2) or hyperoxia (95% O2-5% CO2) for 24 h. Cells were harvested, stained with propidium iodide, and analyzed by flow cytometry. This is representative of a typical histogram comparing cell number and DNA content (2n vs. 4n, where n is haploid).



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Fig. 4.   Transforming growth factor (TGF)-beta inhibits S phase entry of Mv1Lu cells exposed to hyperoxia. Mv1Lu cells were treated in absence and presence of 5 ng/ml of TGF-beta 1 and exposed to hyperoxia (O2) for indicated times. Cells were harvested, stained with propidium iodide, and analyzed by flow cytometry. This is representative of a typical histogram comparing cell number and DNA content (2n vs. 4n).

Differential effects on the cell cycle by TGF-beta and hyperoxia. Asynchronous cultures of Mv1Lu cells were then exposed to hyperoxia in the absence and presence of 5 ng/ml of TGF-beta 1 to determine whether TGF-beta could maintain a G1 phase growth arrest in the presence of hyperoxia. Changes in the percentage of Mv1Lu cells in the S phase were time dependent as noticed by the continual increase in the number of cells with a DNA content of >2n and <4n (Fig. 4, left). Approximately 80% of the cells were in the S phase based on propidium iodide staining after 72 h of hyperoxia. In contrast, cells exposed to hyperoxia and treated with TGF-beta accumulated in the G1 phase and remained predominantly in the G1 phase even after 72 h of hyperoxia (Fig. 4, right). Although the percentage of cells retained in the G1 phase with TGF-beta treatment decreased from 90% after 24 h to ~70% after 72 h, it was still significantly greater than the percentage of cells exposed to hyperoxia alone. TGF-beta inhibited S phase entry of cells exposed to hyperoxia for 3 days when the medium was replenished every 24 h as well as when the cells were treated once at the beginning of the exposure (data not shown).

The expression of cyclin A was also determined as further evidence that TGF-beta and hyperoxia inhibit proliferation through distinct mechanisms. Cyclin A is expressed by cells in the S and early G2 phases and is decreased in Mv1Lu cells treated with TGF-beta (27, 29). Mv1Lu cells were exposed to room air, TGF-beta , hyperoxia, or TGF-beta and hyperoxia for 24 h. Cyclin A was readily detected in asynchronous cultures growing under normoxic conditions and in cultures growth arrested by hyperoxia (Fig. 5). In contrast, cyclin A abundance was decreased to nearly undetectable levels in cultures exposed to TGF-beta or TGF-beta and hyperoxia. Thus cyclin A was detected in asynchronous cultures exposed to room air or arrested by hyperoxia but not in cultures arrested by TGF-beta .


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Fig. 5.   Expression of cyclin A. Mv1Lu cells were exposed to room air (RA), TGF-beta , O2, or TGF-beta +O2 for 24 h. Cells were harvested, and expression of cyclin A was detected by Western blotting. Expression of beta -actin was used to confirm that equal amounts of protein were loaded in each lane.

Effects of hyperoxia on a TGF-beta -dependent transcriptional reporter. These observations demonstrate that TGF-beta inhibits proliferation in the G1 phase, whereas hyperoxia causes cells to accumulate in the S phase. Furthermore, they suggest that TGF-beta signaling can overcome the effects of hyperoxia because cells treated with TGF-beta and hyperoxia accumulate in the G1, not in the S, phase. To confirm that TGF-beta and hyperoxia signal through distinct pathways, the TGF-beta -responsive luciferase reporter gene p3TP-Lux was used to measure TGF-beta -dependent transcriptional responses (31). Mv1Lu cells were transfected with this plasmid and exposed to room air, TGF-beta , hyperoxia, or TGF-beta and hyperoxia for 24 h. Room air-exposed cells had minimal luciferase activity that was markedly induced by treatment with TGF-beta (Fig. 6). Cells exposed to hyperoxia alone also had minimal luciferase activity that was not significantly different from that in room air-exposed cells. In contrast, cells exposed to both TGF-beta and hyperoxia had induced luciferase activity, consistent with the ability of TGF-beta to signal even in O2-exposed cultures. As a control for reporter specificity, R1B cells were transfected and exposed to room air, TGF-beta , hyperoxia, or TGF-beta and hyperoxia. Minimal luciferase activity was detected in room air-exposed cultures and was not induced by TGF-beta or hyperoxia (Fig. 6).


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Fig. 6.   Hyperoxia does not induce TGF-beta -dependent transcription. A: Mv1Lu and R1B cells were transfected with TGF-beta -regulatable luciferase reporter p3TP-Lux containing 3 12-O-tetradecanoylphorbol 13-acetate response elements [(TRE)3] and plasminogen activator inhibitor (PAI)-1 promoter. Arrows, orientation of each element relative to transcription initiation site. B: cells were exposed to RA, TGF-beta , O2, or TGF-beta +O2 for 24 h, and luciferase activity was determined. Values are means ± SD of luciferase activity from triplicate transfections. * Luciferase activity was significantly increased in Mv1Lu cells treated with TGF-beta , P < 0.05.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The present study extends previous observations on the growth-arresting activities of hyperoxia in other cell lines by showing that hyperoxia caused Mv1Lu and R1B cells to accumulate in the S phase of the cell cycle. These changes were independent of TGF-beta signaling because hyperoxia inhibited proliferation of the TGF-beta -unresponsive cell line R1B. Mv1Lu cells exposed to room air or hyperoxia expressed cyclin A, whereas cells treated with TGF-beta accumulated in the G1 phase and had reduced levels of cyclin A. Moreover, Mv1Lu cells exposed to both TGF-beta and hyperoxia accumulated in the G1 phase, with decreased expression of cyclin A, suggesting that TGF-beta signaling can overcome the growth-arresting activities of simple hyperoxia. This was confirmed by demonstrating functional activity of a TGF-beta -dependent transcriptional reporter in Mv1Lu cells cultured in room air or hyperoxia. However, hyperoxia by itself was unable to activate TGF-beta -dependent transcription. Collectively, these observations suggest that TGF-beta does not participate in the growth-arresting activities of hyperoxia. However, TGF-beta may modify the cellular response to hyperoxia by preventing S phase entry.

Previous studies with rat SV40T-T2 cells demonstrated that their proliferation was inhibited by hyperoxia. Although growth arrest was associated with increased mRNA expression of histone and thymidine kinase mRNAs, they were not efficiently translated, which could account for the failure of these cells to proliferate (5). In addition, hyperoxia increased mRNA levels of TGF-beta 1, Tbeta R-I, and Tbeta R-II (4, 6). Hyperoxia also increased p21, which inhibited the kinase activity of cyclin E-Cdk2 complexes (6). The authors concluded that TGF-beta participates in mediating the growth-arresting activities of hyperoxia because SV40T-T2 cells cultured with a neutralizing antibody to TGF-beta had a modest increase in cyclin E-Cdk2 activity. Unfortunately, these studies never measured cell cycle progression or determined whether addition of the neutralizing TGF-beta antibody resulted in an increase in total cell number. Based on the present findings, it is possible that blocking TGF-beta activity resulted in more cells exiting the G1 phase and entering the S and/or G2/M phases where they arrested. Alternatively, hyperoxia may inhibit cell proliferation by different mechanisms that are cell-type dependent. For example, the effects of hyperoxia on SV40T-T2 cells may be unique to these cells because SV40 and other DNA tumor proteins are known to rearrange cyclin and Cdk partners (32). In contrast, there is no evidence that the Mv1Lu cells, which are derived from the fetal mink lung, express viral genes (14).

The present study reveals that Mv1Lu cells exposed to hyperoxia accumulate predominantly in the S phase of the cell cycle. Although G1 and G2 cell cycle checkpoints in response to DNA damage have been studied extensively, less is known about the existence of an S phase checkpoint. Cells exposed to ionizing radiation growth arrest in the G1 phase through the DNA damage-dependent accumulation of the tumor suppressor p53, which transcriptionally increases p21 (10, 17, 20). In addition to blocking S phase entry, p21 also binds proliferating cell nuclear antigen, which participates in both DNA replication and repair (18). Additional studies in yeast have identified DNA polymerase-epsilon as a DNA damage sensor that may integrate DNA replication and repair (19). Ionizing radiation also induces a G2 phase checkpoint that is controlled by RAD53 and other related kinases (29). In contrast, ultraviolet B radiation, which creates DNA adducts, prolonged G1 and S phase progression in neonatal rat keratinocytes (24). Similarly, alkylating agents slow S phase progression in Saccharomyces cerevisiae (23). Because DNA replication was dependent on the MEC1 and RAD53 genes, the authors concluded that alkylating agents inhibited proliferation through activation of a checkpoint control rather than through failure to bypass DNA lesions.

Checkpoints were defined as places in the cell cycle where progression was dependent on completion of the previous phases (12). Failure to complete the previous phase resulted in a transient growth arrest during which the cells were able to conclude incomplete biochemical processes. Although hyperoxia alone is not cytotoxic, it is converted to genotoxic ROS that would be predicted to elicit the classic G1 phase checkpoint involving p53 and p21 or the G2 phase checkpoint involving RAD53 (20, 29). The present finding that cells exposed to hyperoxia fail to progress through the S phase is consistent with an inability to appropriately replicate DNA. Further studies are needed to determine whether this is due to a global shutdown in cellular function or a novel DNA damage checkpoint.

The present study found that TGF-beta could maintain the G1 phase checkpoint in cells exposed to hyperoxia, thereby preventing the entry and subsequent arrest in the S phase caused by hyperoxia alone. It is widely believed that the cells arrested in the G1 or G2 phase will have a greater capacity to survive genotoxic stress compared with cells in the S phase when DNA replication is occurring (29). However, yeast cells arrested in the S phase with the alkylating agent methylmethane sulfonate have enhanced survival when challenged with ionizing radiation (23). The authors suggest that methylmethane sulfonate may protect against radiation-induced DNA damage because DNA repair events are coupled with replication. This hypothesis is consistent with the observation that DNA polymerase-epsilon and p21 coordinate DNA repair and replication (18, 19). The concept that resistance to genotoxins may be coupled to the cell cycle is also consistent with a recent study (15) where confluent (growth-arrested) cultures of small-airway epithelial cells were more resistant to hyperoxic injury than subconfluent (proliferating) cultures. Although these studies use different cell types and DNA-damaging agents, they suggest that cells in different phases of the cell cycle are not equally injured by genotoxins. Thus cells arrested in the G1 phase by TGF-beta may have an altered ability to survive exposure to hyperoxia than cells arrested in the S and G2 phases by hyperoxia alone. Because TGF-beta decreases antioxidant enzyme expression and enhances the cytotoxicity of hydrogen peroxide in A549 cells, future experiments must take into account the multitude of biological activities associated with this cytokine in addition to its ability to simply inhibit proliferation (3).

In summary, the findings in this study reveal that hyperoxia induced Mv1Lu and R1B cells to accumulate in the S phase independent of TGF-beta signaling. TGF-beta induced a G1 phase growth arrest that blocked entry into and subsequent arrest in the S phase caused by hyperoxia. These changes in cell cycle progression were not associated with decreased cell viability. It is worth noting that TGF-beta expression increased within 1-3 h of exposure to hyperoxia in pulmonary epithelial cells of adult mice (22). Although the role that TGF-beta plays in hyperoxic lung injury remains to be determined, this finding is interesting because it suggests that TGF-beta may prevent S phase entry of pulmonary cells exposed to hyperoxia in vivo.


    ACKNOWLEDGEMENTS

We thank Anita Roberts for the Mv1Lu cell line and Joan Massagué for the R1B cell line and p3TP-Lux plasmid.


    FOOTNOTES

This work was funded by the American Lung Association; American Heart Association Beginning Grant-in-Aid 9860004T; and National Heart, Blood, and Lung Institute Grant HL-58774 (to M. A. O'Reilly). Additional support was provided by National Cancer Institute Grants CA-73725 and CA-11198 (to P. C. Keng).

R. C. Rancourt was supported by National Institute of Environmental Health Sciences Training Grant ES-07026.

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

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

Received 28 October 1998; accepted in final form 3 August 1999.


    REFERENCES
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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Am J Physiol Lung Cell Mol Physiol 277(6):L1172-L1178
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