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Published ahead of print on August 12, 2004, doi:10.1165/rcmb.2004-0141OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 31, pp. 611-618, 2004
© 2004 American Thoracic Society
DOI: 10.1165/rcmb.2004-0141OC

Extracellular Signal–Regulated Kinase Activation Delays Hyperoxia-Induced Epithelial Cell Death in Conditions of Akt Downregulation

Son V. Truong, Martha M. Monick, Timur O. Yarovinsky, Linda S. Powers, Toru Nyunoya and Gary W. Hunninghake

Roy J. and Lucille A. Carver College of Medicine, University of Iowa, Iowa City; and Veterans Administration Medical Center, Iowa City, Iowa

Address correspondence to: Martha M. Monick, Division of Pulmonary, Critical Care, and Occupational Medicine; 100 EMRB, University of Iowa Roy J. and Lucille A. Carver College of Medicine, Iowa City, IA 52242. E-mail: martha-monick{at}uiowa.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Hyperoxia (fraction of inspired oxygen = 95%) induces death of lung epithelial cells. The duration of cell survival in the setting of hyperoxia depends on hyperoxia-induced activation of intracellular survival pathways. Two survival pathways with known effects on lung epithelial cells are the propidium iodide 3–kinase/Akt and extracellular signal–regulated kinase (ERK)/mitogen-activated protein (MAP) kinase pathways. We investigated the effect of hyperoxia on activity of both the Akt and ERK pathways in the A549 lung epithelial cell line. Hyperoxia-exposed cells show progressive loss of Akt activation and total Akt protein. Hyperoxia decreases Akt mRNA, consistent with the loss of total Akt. In addition, hyperoxia induces ERK activation. Inhibition of ERK with the MAP kinase kinase 1/2 inhibitor, U0126, shortens the survival time of cells in hyperoxia, suggesting that increased ERK activity partially compensates for the hyperoxia-induced Akt downregulation. Our findings show, for the first time, that hyperoxia has divergent effects on two survival pathways (Akt and ERK), and that ERK activity compensates for the loss of the Akt survival effects, delaying the death of hyperoxia-exposed lung epithelial cells.

Abbreviations: threshold cycle, Ct • extended chemiluminescence, ECL • extracellular signal–regulated kinase, ERK • ethidium homodimer, EthD-1 • fraction of inspired oxygen, FiO2 • hypoxanthine phosphoribosyltransferase, HPRT • mitogen-activated protein, MAP • MAP kinase kinase, MEK • nicotinamide adenine dinucleotide, NAD • poly(ADP-ribose) polymerase, PARP • propidium iodide, PI • phosphatidylinositol 3-kinase, PI3-K • reverse transcriptase–polymerase chain reaction, RT-PCR • tumor necrosis factor, TNF •


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Oxygen supplementation at supraphysiologic levels is necessary in patients with respiratory failure, especially in those with acute lung injury or in premature neonates with underdeveloped lungs (1, 2). Oxygen in these settings is a life-preserving supportive measure until the initial pathologic process that elicited the respiratory disease subsides. However, prolonged exposure to high oxygen concentration leads to cellular injury and death (1–6). Pulmonary oxygen toxicity was first described as early as 1897 (3), and toxicity was shown to induce alterations of the airway structures, lung tissue, and pulmonary vasculature. Alterations in airway structures include atelectasis, leukocyte, erythrocyte, and macrophage accumulation in the alveoli, and formation of membranes on the alveolar wall. Alterations in the pulmonary vasculature include hyperemia, conglomeration of blood cells in capillaries, and capillary proliferation. In many respects, the effects of hyperoxia resemble those typical of acute lung injury (3, 7).

It has been suggested that the toxic effect of oxygen is mediated by increased reactive oxygen intermediates, such as superoxide anion (O2–·), hydrogen peroxide (H2O2), and hydroxyl radical (OH·) (8). Hyperoxia predisposes cells to lipid peroxidation, protein oxidation, DNA damage, and depletion of cellular reducing agents (9–16). Injury to the respiratory epithelium results in impaired gas exchange due to distortion of the alveolar architecture (17, 18). Additionally, hyperoxia may result in increased susceptibility to infection from impaired macrophage function and disruption of the structural integrity of the alveolar epithelium (6, 19, 20).

Epithelial cells exposed to conditions of hyperoxia (95% fraction of inspired oxygen [FiO2]) exhibit morphologic changes characterized by cell flattening/stretching, cell cycle arrest, and death (6, 21–25). It has also been shown that other types of oxidative stress lead to Akt degradation via proteolysis mediated by caspases (26). However, the fate of Akt in conditions of hyperoxia remains poorly studied. Akt has been widely demonstrated to be a prosurvival kinase. It has been suggested that Akt protects against DNA strand breakage, detachment, and death (27–32). Animal studies have demonstrated that the intratracheal introduction of constitutively active Akt gene into murine lung by adenoviral vectors results in delayed death (32).

The extracellular signal–regulated kinase (ERK) 1/2 and mitogen-activated protein (MAP) kinase are activated by hyperoxia (10, 33, 34). However, there are conflicting results regarding the role of ERK in epithelial cells exposed to high concentration of oxygen. ERK activation has been shown to be both protective and detrimental to cells in hyperoxic condition. ERK activation has been demonstrated to protect against cellular injury through its protective effect on DNA strand breakage and antiapoptotic properties (35). On the other hand, ERK signaling has also been implicated in epithelial cell death in high oxygen tension (10, 36–38). Thus, the role of Akt and ERK signaling in response to oxidative stress from hyperoxia remains poorly studied.

In this study, we evaluated mechanisms of cell death in hyperoxia-exposed lung epithelial cells (A549 cells). The A549 cell line derived from explant carcinoma tissue has many of the features of lung epithelial cells. Some studies have found it to have characteristics of alveolar type II cells. In our study, A549 cells behaved very much like airway epithelial cells. Future studies will be needed to show definitively that the phenomena that we observed in the cell culture system also occur in the intact lung. We demonstrate for the first time that hyperoxia downregulates Akt activity, mRNA, and protein and that epithelial cell survival in conditions of low Akt depends on hyperoxia-induced ERK.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Reagents and Antibodies
Chemicals were obtained from Sigma (St. Louis, MO). Protease inhibitors were from Roche Applied Science (Indianapolis, IN). Nitrocellulose membrane, extended chemiluminescence (ECL), and ECL plus chemiluminescent substrates were from Amersham Biosciences (Arlington Heights, IL). Phosphorylation-specific antibodies for Akt and ERK, total Akt antibody, and cleaved poly(ADP-ribose) polymerase (PARP) antibody were from Cell Signaling (Beverly, MA). ERK2 antibody and horseradish peroxidase-conjugated goat anti-rabbit and goat anti-mouse developing antibodies were from Santa Cruz Biotechnology (Santa Cruz, CA). ß-Actin antibody was from Sigma. MAP kinase kinase (MEK)1/2 inhibitor (UO126) was from Calbiochem (Darmstadt, Germany). Fas ligand was obtained from Upstate Biotechnologies (Charlottesville, VA).

Cell Culture and Treatment Conditions
A549 cells, an adenocarcinoma cell line with properties of normal airway epithelial cells (39), obtained from American Type Culture Collection (Manassas, VA), were incubated in 5% CO2 at 37°C in a humidified incubator. The cells were cultured in Eagle's minimum essential medium (Invitrogen, Grand Island, NY) supplemented with 10% fetal bovine serum (Hy-Clone, Logan, UT) and 40 mg/ml gentamicin and subcultured every 5–7 d by harvesting in 0.025% trypsin (Invitrogen) no more than 20 times from stock originally designated at pass 70. Cells were cultured for 24 h in 35-mm tissue culture plates (Corning, Corning, NY) at 1.5 x 106 cells/ml and subsequently exposed to normoxia (21% FiO2, 5% CO2) or hyperoxia (95% FiO2, 5% CO2). The hyperoxia group was placed in a sealed humidified hyperoxia chamber (Reming Bioinstruments, Redfield, NY) with constant instillation of 95% O2 and 5% CO2. When indicated, the MEK1/2 inhibitor, UO126, was added 30 min before exposure to hyperoxia. In our cell death experiment, A549 cells were seeded at a starting density of 1 x 106 cells/ml instead of 1.5 x 106.

Cell Viability and Death Assays
Cell viability and death was measured by Trypan blue (Sigma) and by a commercially available kit, the LIVE/DEAD viability/cytotoxicity kit (Molecular Probes, Eugene, OR). For assessment of viability with Trypan blue, 1 ml of nonadherent cells in the supernatant was aspirated from the 35-mm tissue culture plate after treatment and centrifuged at 300 x g. The cells were resuspended in 100 µl of 1x Dulbecco's phosphate-buffered saline. 40 µl of suspended cells was counted in an electric particle counter (Coulter Electronics, Hialeah, FL). The remaining 60 µl of cells was added to an equal volume of 10% Trypan blue and counted using a hemacytometer for Trypan blue–stained (dead) and nonstained cells (live). Trypsin (0.025%) was added to the remaining adherent cells, centrifuged, resuspended, and counted using the electric particle counter and hemacytometer as above. The total number of cells was determined by adding the number of nonadherent and adherent cells. The fraction of live/dead nonadherent and adherent cells was determined by dividing the number of live cells per microscopic field of ~ 100 cells. Each slide was counted twice in two different microscopic fields for reproducibility. For assessment of viability using the LIVE/DEAD viability/cytotoxicity kit, A549 cells were stained with 1.0 µM calcein acetoxymethyl ester for 30 min. The same cells were subsequently stained with 1.5 µM ethidium homodimer (EthD-1) for 15 min and viability was measured using fluorescence microscopy. In some cases, membrane permeability to EthD-1 was used as a marker of cell viability. Two hundred cells from at least three different fields were counted for each measurement. Data are presented as % dead (number of EthD-1–expressing cells/total number of cells x 100).

Isolation of Whole-Cell Protein Extracts
Whole-cell protein was obtained by lysing the cells on ice for 20 min in 200 µl of lysis buffer (0.05 M Tris, pH 7.4, 0.15 M NaCl, 1% Nonidet P-40, 1 protease minitab) (Roche Applied Science) per 10 ml and 1x phosphatase inhibitor mixture (catalog no. 524625, Calbiochem). The cell lysates were then sonicated for 20 s, incubated on ice for 30 min to complete lysis, and centrifuged at 15,000 x g for 10 min at 4°C to separate lysate from cellular debris. The protein isolated was quantified using a protein measurement kit (protein assay kit 500–0006, Bio-Rad). Cell lysates were stored at 70°C until analysis.

Western Blot Analysis
Western blot analysis for the presence of phosphorylated and nonphosphorylated proteins was performed on whole-cell protein lysates. Protein (30–80 µg) was mixed 1:1 with 2x sample buffer (20% glycerol, 4% sodium dodecyl sulfate, 10% mercaptoethanol, 0.05% bromphenol blue, and 1.25 M Tris, pH 6.8; all from Sigma), heated to 95°C for 5 min, and fractionated on a 10 or 12.5% sodium dodecyl sulfate–polyacrylamide gel run at 100 V for 90 min. Cell proteins were transferred to nitrocellulose filters (ECL) by semidry transfer (Bio-Rad) at 20 V for 45 min. Equal loading of protein on the blots was evaluated using Ponceau S, a staining solution designed specifically for proteins on nitrocellulose membranes (Sigma Chemical). The nitrocellulose filter was blocked with 5% milk in Tris-buffered saline with 0.1% Tween 20 for 1 h, washed, and then incubated with the primary antibody overnight. The blots were washed four times with Tris-buffered saline with Tween 20 and incubated for 1 h with horseradish peroxidase–conjugated anti-rabbit or anti-mouse immunoglobulin G antibody. Immunoreactive bands were developed using a chemiluminescent substrate, ECL or ECL plus Western blotting detection system (Amersham, Arlington Heights, IL). Autoradiographs were obtained by exposing Kodak Biomax MR films (Eastman Kodak, Rochester, NY) to the membranes for 10 s to 2 min.

Flow Cytometry for Apoptosis
Measurement of apoptosis was performed using two methods: DNA content analysis and annexin V/propidium iodide staining. A549 cells were grown to 80% confluency in 35-mm tissue culture dishes, incubated 24 h in minimal essential medium supplemented with 10% fetal calf serum, and cultured in conditions of normoxia (21% O2 and 5% CO2) or hyperoxia (95% O2 and 5% CO2) for 12, 24, 36, and 48 h. Cells were lifted off tissue culture plate with 0.025% trypsin and neutralized with equal volume of cell culture media. The cells were washed twice in phosphate-buffered saline and fixed in cold 70% ethanol until staining and analysis. For the DNA content analysis, cells were washed twice in phosphate-buffered saline, treated with 0.25 mg/ml RNase A (Roche Applied Science), and stained with 50 µg/ml propidium iodide (PI) at 4°C for 30 min in the dark and analyzed for DNA content. A total of 10,000 cells that satisfied a gate on forward and side scatter to eliminate aggregates and debris were evaluated with a FACScan flow cytometer (BD Biosciences). Data analysis was performed with ModFit LT (Verity Software House Inc., Topsham, ME). For the annexin V/propidium iodide staining, live A549 cell death was measured by staining with fluorescein isothiocyanate–labeled annexin V and propidium iodide. A total of 10,000 cells that satisfied a gate on forward and side scatter to eliminate aggregates and debris were evaluated using a FACScan flow cytometer (Becton Dickinson, Mountain View, CA).

Real-Time Reverse Transcriptase–Polymerase Chain Reaction for Akt
Total RNA was isolated using the Absolutely RNA reverse transcriptase–polymerase chain reaction (RT-PCR) Miniprep Kit (Stratagene, La Jolla, CA) following manufacturer's instructions. RNA quantification was performed using the RiboGreen kit (Molecular Probes). A 1 µg sample of total RNA was reverse-transcribed to cDNA using the RETROscript RT-PCR kit (Ambion, Austin, TX). In a 0.2-ml PCR tube (Bio-Rad), 2 µl of cDNA (10% cDNA synthesis reaction volume) was added to 48 µl of PCR reaction mixture containing 160 µM each of deoxyribonucleotide triphosphate (dNTP) (Invitrogen), 3.0 mM MgCl2 (Invitrogen), 1:10,000 SYBR Green I DNA dye (Molecular Probes), 0.2 µM of each sense and antisense primers (IDT, Coralville, IA), and 2.5 units of platinum TaqDNA polymerase (Invitrogen). Amplification was then performed in an iCycler iQ fluorescence thermocycler (Bio-Rad) as follows: 5 min at 95°C followed by 45 cycles of 20 s at 95°C, 20 s at 60°C, and 20 s at 72°C. Fluorescence data were captured during the 72°C dwell time. Specificity of the amplification was confirmed using melting curve analysis. In addition, 5 µl aliquots of the end products of PCR amplification were separated in 1.5% agarose gel in Tris-borate–ethylenediaminetetraacetic acid buffer, stained with ethidium bromide, and visualized using GelDoc 2000 gel documentation system (Bio-Rad). The remaining PCR products were purified with QIAquick PCR purification kit (QIAGEN, Valencia, CA) and sequenced at the University of Iowa DNA facility. Data for real-time RT-PCR were collected and recorded by iCycler iQ software (Bio-Rad) and expressed as a function of threshold cycle (Ct), the cycle at which the fluorescent intensity in a given reaction tube rises above background (calculated as 10 times the mean SD of fluorescence in all of the wells over the baseline cycles). Specific primer sets used were as follows (5' to 3'): Akt sense, CCTCAAGAATG ATGGCACCT; Akt antisense, TGAGTTGTCACTGGGTGAGC; hypoxanthine phosphoribosyltransferase (HPRT) sense, CCTCATG GACTGATTATGGAC; and HPRT antisense, CAGATTCAACTTGCGCTCATC. Primers were selected based on nucleotide sequences downloaded from the National Center for Biotechnology Information data bank and designed with software by Steve Rozen and Helen J. Skaletsky (1998 Primer3; code available at www.genome.wi.mit.edu/genome_software/other/primer3.html).

Quantitation of Akt Gene Expression
Quantitation of Akt mRNA expression was calculated as follows: for each sample assayed, the Ct for reactions amplifying Akt and the HPRT housekeeping gene were determined. The Akt Ct for each sample was corrected by subtracting the Ct for HPRT (Ct). Akt mRNA abundance, relative to HPRT mRNA abundance, was calculated by the formula 2(–{Delta}Ct). Validity of this approach was confirmed by using serial 10-fold dilutions of template for Akt and HPRT. Using the 10-fold dilutions, the amplification efficiencies for Akt and HPRT were found to be identical.

Statistical Analysis
The numbers of cells in each condition at various time points in adherent or nonadherent conditions were compared using unpaired (two-tailed) t tests. Values in figures are mean ± SEM. Values of P < 0.05 were considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Exposure to Hyperoxia Leads to Lung Epithelial Cell Death
In the present study, we investigated the mechanism of epithelial cell injury induced by hyperoxia. To do so, we utilized an in vitro system using A549 (transformed alveolar type II epithelial) cells exposed to normoxia and hyperoxia for varying times. A549 cells were cultured in 35-mm tissue culture plates at a starting cell density of 1.5 x 106/ml. After progressive exposure to a normoxic (21% O2) or hyperoxic (95% O2) environment at 0, 12, 24, and 48 h, the total numbers of live and dead cells were determined by evaluating plasma membrane integrity with Trypan blue exclusion or EthD-1 staining (LIVE/DEAD viability/cytotoxicity kit; Molecular Probes). As shown in Figure 1, after 48 h of culture, ~ 15% of the cells die in normoxia compared with 55% of the cells dying in hyperoxia. The same result was obtained by staining the normoxia and hyperoxia-exposed cells with EthD-1/calcein and quantifying the numbers of live and dead cells by direct fluorescence microscopy (data not shown). These data confirm the findings of other studies that have shown that prolonged exposure to a hyperoxic environment is lethal to lung epithelial cells (6, 12, 40). It is unclear whether hyperoxia-induced cell death is by necrosis or apoptosis (7, 41–43). We next evaluated the amount of apoptosis in our model.



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  		Figure 1. (A) Hyperoxia induces significant cell death in epithelial cells. A549 cells were cultured in conditions of normoxia (squares; 21% O2, 5% CO2) and hyperoxia (diamonds; 95% O2, 5% CO2). At 0, 12, 24, and 48 h, the percent of total cell death (adherent and nonadherent) was determined using 10% Trypan blue staining. Experiments were done in triplicate. (B) Hyperoxia induces low levels of apoptosis. A549 cells were cultured in conditions of normoxia (21% O2, 5% CO2) and hyperoxia (95% O2, 5% CO2). At 0, 12, 24, 36, and 48 h, percent of apoptotic cells was assessed by DNA analysis for subdiploid cells. This graph represents the DNA analysis data from the normoxia and hyperoxia treatment groups at 48 h. Experiments were done in triplicate.

 
Hyperoxia Induces Very Low Levels of Apoptosis
Under light and fluorescence microscopy, epithelial cell morphologic changes (cell flattening and stretching, membrane fragmentation, and loss of cytoplasm) were observed in hyperoxia (95% O2). The nuclei, however, were "normal" in appearance. These changes are consistent with necrotic cell death. This led us to hypothesize that lung epithelial cells die predominantly of necrosis.

The membranes of necrotic and late apoptotic cells are permeable to annexin V, and these cells cannot be distinguished with certainty. Thus, double staining (annexin V with PI) is required to estimate the number of early apoptotic cells (annexin V–positive, PI-negative). When we stained live A549 cells with annexin V and analyzed the cells by flow cytometry, we observed < 5% cells stained with annexin V only (apoptotic cells) in both normoxia and hyperoxia. This staining could be attributed to ether low levels of apoptosis in both groups or to the membrane damage that occurs during cell harvesting for flow cytometry analysis (44). More than 40% cells in the hyperoxia group (48 h) were stained with both annexin V and PI. It is important to note that a positive control group for apoptosis (tumor necrosis factor [TNF]-{alpha} plus actinomycin D) consistently produced more than 30% cells that stained with annexin V only. Based on these observations, we concluded that annexin V staining was not sufficient to determine the mode of cell death in hyperoxia, and preferred to analyze DNA content, morphology of the nuclei, and sensitivity to caspase inhibition.

DNA content analysis was done to measure the proportion of epithelial cells that die from apoptosis. To assess apoptosis in hyperoxia-exposed cells, A549 cells were cultured in normoxia (21% O2) and hyperoxia (95% O2) at 12, 24, 36, and 48 h, and the percent of apoptotic cells was measured by flow cytometry (DNA content analysis). As shown in Figure 1B, the most significant difference occured after 48 h of exposure to hyperoxia. There were 2.73% subdiploid cells in the normoxic group and 7.26% subdiploid cells in the hyperoxic group. Of note, the cell death attributable to apoptosis was significantly lower than the total cell death as measured by Trypan blue exclusion (15% in normoxia and 55% in hyperoxia). The same groups were analyzed using PI and annexin V, measures of membrane permeability, and phosphatidylserine translocation across the plasma membrane. Flow cytometry of both PI- and annexin V–stained cells demonstrated identical results to those of the DNA content analysis: the majority of the cells die by necrosis (data not shown). These experiments, as an aggregate, strongly suggest that hyperoxia induces epithelial cell death, mainly by necrosis. These data are consistent with the observations of Wang and colleagues (43).

Hyperoxia-Exposed Cells Demonstrate Nuclear Morphology Consistent with Necrosis and Not Apoptosis
To confirm the nonapoptotic mechanism of hyperoxia-induced death, A549 cells were exposed to either hyperoxia, Fas ligand or a combination of TNF-{alpha} and actinomycin D for 48 h. Cells were then stained for nuclear morphology using EthD-1 and images acquired by fluorescence microscopy (Figure 2A). Both Fas ligand and TNF-{alpha}/actinomycin D induced nuclear condensation consistent with apoptosis. In contrast, the hyperoxia-exposed cells demonstrated no nuclear condensation. These data confirm the conclusion of the PI staining shown in Figure 1: hyperoxia kills epithelial cells by nonapoptotic methods.



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  		Figure 2. Hyperoxia results in a nonapoptotic cell death. (A) Hyperoxia results in cell death without nuclear condensation typical for apoptosis. A549 cells were cultured in hyperoxia for 48 h and stained with EthD-1 (red nuclear staining for dead cells) and calcein AM (green cytoplasmic staining for live cells). A549 cells were also treated with Fas ligand (0.01 ng/µl) (24 h) or a combination of TNF-{alpha} (1 ng/ml) and actinomycin D (2.5 ug/ml) (16 h) to show typical apoptotic nuclei for positive control. Scale bar = 20 µm. (B) Caspase inhibition prevents hyperoxia-induced PARP cleavage. A549 cells were cultured in conditions of hyperoxia (95% O2, 5% CO2) at 0 (control) and 48 h. A total of 50 µM of ZVAD was added to cells 30 min before exposure to hyperoxia where indicated. Fas ligand treatment groups served as positive controls. 0.01 ng/µl of Fas ligand was added to cells in the positive control group for 12 h. Whole-cell lysates were harvested for Western blot analysis for the 89 kD cleaved PARP. ß Actin immunoreactive bands are used to demonstrate equal loading. Experiments were done in triplicate. (B) Caspase inhibition does not prevent hyperoxia-induced epithelial cell death. A549 cells were cultured in conditions of hyperoxia (95% O2, 5% CO2) at 0 (control) and 48 h. A total of 50 µM of ZVAD was added to cells 30 min before exposure to hyperoxia in the Hyp + ZVAD group. Percent of total cell death (adherent and nonadherent) in hyperoxia and hyperoxia + ZVAD groups was determined by 10% Trypan blue staining. Experiments were done in triplicate.

 
Caspase Inhibition Prevents PARP Cleavage in Hyperoxia-Exposed Cells without Affecting Cell Death
To further study the relative contribution of apoptosis versus necrosis, we compared the effect of caspase inhibition on PARP cleavage (dependent on caspase activity) and cell death. A549 cells were subjected to hyperoxic conditions (95% O2) for 48 h with and without the polycaspase inhibitor, fmk-ZVAD. Whole-cell lysates were obtained for Western blot analysis and cell viability was analyzed by Trypan blue exclusion. Using an antibody specific for the 89 kD PARP cleavage product, we observed induction of PARP cleavage in hyperoxia-exposed cells that was blocked by the caspase inhibitor (Figure 2B). These data are consistent with our observations that a small fraction of the cells die via apoptosis (see above). As a control, we also treated the lung epithelial cells with Fas ligand to obtain caspase-dependent signaling. Figure 2B shows that PARP cleavage by Fas ligand was prevented by the caspase inhibitor, fmk-ZVAD. Compared with the effect of caspase inhibition on PARP cleavage, when we analyzed lung epithelial cells exposed to the same conditions, there was little effect of caspase inhibition on hyperoxia-induced cell death (Figure 2C). There was no significant difference in survival between the hyperoxia-exposed group and the hyperoxia group exposed to the caspase inhibitor (~ 70% cell death after hyperoxia and fmk-ZVAD exposure compared with 65% cell death after hyperoxia alone). ZVAD was effective in blocking caspase-mediated PARP cleavage but not in altering cell death after hyperoxia. These data are consistent with the low amounts of apoptosis occurring in hyperoxia-exposed epithelial cells.

Hyperoxia Downregulates Active Akt and Total Akt
We have recently demonstrated that the PI3-kinase/Akt and ERK pathways cooperate in maintaining viability of lung epithelial cells (45). We next wanted to look at the contribution of these two pathways to cell survival in hyperoxia-exposed lung epithelial cells. A549 cells were exposed to normoxia (21% O2) and hyperoxia (95% O2) and, at 24, 36, and 48 h, whole-cell lysates were obtained for Western blot analysis for active (phosphorylated on serine 473) and total Akt. The data demonstrate that by 48 h there is a significant decline in the level of both phosphorylated Akt and total Akt (Figure 3A). Very little cell death occurred before 36 h of exposure to hyperoxia. Though we cannot entirely rule out the idea that the decrease in Akt is simply a reflection of decreased viability of the cells, the equalization of protein levels and consistent ß actin staining suggest that the decreased Akt is an active process. Further studies will be needed to prove this definitively. The loss of Akt is consistent with an important role for ERK activity in prolonging survival of hyperoxia-exposed epithelial cells.



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  		Figure 3. (A) Hyperoxia decreases Akt amounts and activity. A549 cells were cultured in conditions of normoxia (21% O2, 5% CO2; open bars) and hyperoxia (95% O2, 5% CO2; shaded bars) at 0 (control), 24, 36, and 48 h. Whole-cell lysates were harvested for Western blot analysis for phospho-Akt and total Akt. ß Actin immunoreactive bands are used to demonstrate equal loading. Experiments were done in triplicate. Densitometry was determined by comparing experimental samples to the zero-time control (densitometry of experimental band/densitometry of control band). Significance was determined using Student's t test, and compared normoxia to hyperoxia at equivalent time points (P < 0.01). (B) Hyperoxia decreases Akt mRNA. A549 cells were cultured in conditions of normoxia (21% O2, 5% CO2) and hyperoxia (95% O2, 5% CO2) at 0 (control), 36, and 48 h. Left: RNA gel of Akt mRNA after RT-PCR amplification at 0 (control), 36, and 48 h. Right: bar graph showing the level of Akt mRNA (expressed as Akt mRNA abundance relative to HPRT mRNA abundance) in normoxia and hyperoxia at 48 h using RT-PCR. Experiments were done in triplicate.

 
Hyperoxia Downregulates Akt mRNA
Hyperoxia has been shown previously to decrease Akt activity (46, 47). In this study, the decrease in Akt was due to caspase-mediated cleavage of the protein. Our study shows that pretreatment with ZVAD reverses caspase-mediated PARP cleavage. However, we were not able to show that ZVAD inhibited the decrease in Akt activity or protein (data not shown). Our data demonstrate the novel finding that total Akt levels are decreased as well. To more completely study the decrease in total Akt, we evaluated Akt mRNA levels in normoxia- versus hyperoxia-exposed cells. A549 cells were cultured in normoxia (21% O2) and hyperoxia (95% O2) at 24, 36, and 48 h. Total RNA was isolated at varying time points, and quantitative real-time RT-PCR was performed. In Figure 3B, we show both DNA products from an Akt-specific RT-PCR and a graph from three separate experiments using real-time RT-PCR to quantitate Akt mRNA levels. Both data show the same result: hyperoxia decreases Akt mRNA levels. These data suggest that one-way hyperoxia decreases total Akt by downregulation of Akt mRNA.

Hyperoxia Activates ERK MAP Kinase
In addition to phosphatidylinositol 3-kinase (PI3-K), the ERK signaling cascade has been shown to be important in promoting cell survival in response to oxidative stress (10, 33, 34). To further evaluate the role of ERK in our system, lung epithelial cells were exposed to normoxia (21% O2) and hyperoxia (95% O2) and, at 24, 36, and 48 h, whole-cell lysates were obtained for analysis of ERK activity (Western analysis for phosphorylated ERK). Figure 4 demonstrates that hyperoxia induces extended ERK activity starting at 36 h and continuing through 48 h (a time point associated with significant cell death). The blot suggests a possible decrease in the normoxic phosphorylated ERK, over multiple blots (see densitometry graph, Figure 4) this change is not significant. The decrease is probably due to cell cycle changes in the serum-dependent proliferating A549 cells and, if anything, only serves to enhance the hyperoxia-linked increases in phosphorylated ERK. The increase in ERK activity coincides with the onset of loss of Akt and cell death.



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  		Figure 4. Hyperoxia activates ERK. A549 cells were cultured in conditions of normoxia (white bars; 21% O2, 5% CO2) and hyperoxia (gray bars; 95% O2, 5% CO2) at 0, 24, 36, and 48 h. Whole-cell lysates were harvested for Western blot analysis for phosphorylated (active) ERK. Total ERK immunoreactive bands are used to demonstrate equal loading. Experiments were done in triplicate. Densitometry was determined by comparing experimental samples to the zero-time control (densitometry of experimental band/densitometry of control band). Significance was determined using Student's t test and compared normoxia to hyperoxia at equivalent time points (P < 0.01).

 
Erk Activity Delays Hyperoxia-Induced Death of Lung Epithelial Cells
Because ERK activity increased at time points that coincided with the onset of cell death, we wished to determine if ERK activity was responsible for or delayed cell death with hyperoxia. To evaluate this question, we used UO126, an inhibitor of the ERK upstream kinase, MEK1. We have previously shown that in A549 cells, at the doses used, that U0126 blocks ERK activity while having no effect on Akt activity (45). A549 cells were pretreated with UO126 for 30 min and exposed to normoxia (21% O2) or hyperoxia (95% O2) for 24 and 48 h. Cell viability was assessed by EthD-1/calcein staining as described previously here. Figure 5 demonstrates that blocking ERK activity leads to significantly earlier cell death. The increased death was not due to an increase in apoptosis (data not shown). The increase in cell death with ERK inhibition begins at 24 h, rises to ~ 60% at 36 h, and is at 100% by 48 h. ERK inhibition in normoxia has a deleterious effect on cell survival (as does PI3 kinase inhibition) (45). This supports the important role these two pathways play in epithelial cell homeostasis. In hyperoxia, with decreasing Akt protein, ERK activity becomes an important source of survival activity in A549 cells. This study provides strong evidence that ERK activation promotes extended epithelial cell survival and decreased cell death by necrosis in a hyperoxic environment.



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  		Figure 5. ERK activity delays hyperoxia-induced epithelial cell death. A549 cells were cultured in conditions of normoxia (21% O2, 5% CO2) and hyperoxia (95% O2, 5% CO2) at 0 (control), 24 h, 36, and 48 h. 10 µM of UO126 was added to cells 30 min prior to exposure to hyperoxia where indicated. Percent of total live cells (adherent and nonadherent) in the normoxia/hyperoxia without UO126 and normoxia/hyperoxia with UO126 groups was determined using EthD-1 uptake as described in MATERIALS AND METHODS. Experiments were done in triplicate. Purple diamonds, normoxia; blue squares, normoxia + UO126; red triangles, hyperoxia; yellow squares, hyperoxia + UO126.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this study, we evaluated the role of two cell survival pathways in lung epithelial cells exposed to hyperoxia. We found that under hyperoxic conditions, epithelial cells undergo significant cell death, but with low levels of apoptosis. Consistent with this finding, caspase inhibition did not significantly decrease cell death. This suggests that the major mechanism of hyperoxia-induced cell death is caspase-independent and due to necrosis. When we evaluated Akt and ERK activity in hyperoxia-exposed cells, we found that hyperoxia had a discordant effect on the two pathways. Akt activity, total protein, and mRNA were decreased after hyperoxia. In contrast, hyperoxia induced significant and extended ERK activation. Inhibition of ERK resulted in earlier and more pronounced cell death. Taken together, these data suggest that ERK activation delays hyperoxia-induced necrotic cell death in the absence of Akt protein.

Interpretation of these results should take into account the fact that the experiments were all done in a proliferating lung epithelial cell line (A549). These cells do not entirely replicate the conditions found in polarized, nonreplicating airway epithelial cells. However, we have shown previously that A549 cells mimic primary airway cells, especially as regards signaling pathway activity (45, 48). For that reason, we believe that the data presented in this article have a high probability of replicating those gathered on the effects of hyperoxia on primary airway epithelial cells.

Our DNA analysis showed minimal apoptosis in A549 cells exposed to hyperoxia. In addition, we observed no increase in Fas mRNA after treatment with hyperoxia (data not shown). However, cells exposed to hyperoxia showed some increase in PARP cleavage that was reversed with caspase inhibition. A recent review by Bouchard and colleagues proposed that excessive PARP activation causes depletion of intracellular nicotinamide adenine dinucleotide (NAD) pools, resulting in disruption of the NAD/NAD phosphate balance. This process depletes cellular adenosine triphosphate and leads to necrotic cell death from disruption of all energy-dependent cellular processes (49). This remains a possible mechanism for the hyperoxia-induced epithelial cell death in our system.

Akt activation has been reported to protect cells from various stress stimuli (50). Akt's functions include maintaining homeostasis, regulating cell growth, and inhibiting cell death (28, 32). Inactivation of Akt function or loss of Akt protein can lead to homeostatic dysregulation, cell cycle arrest, and cell death. Akt's protective effect is mediated by phosphorylation of a number of downstream substrates, including members of the B cell leukemia (Bcl-2) family, Bcl-2-associated death promoter (BAD) (30, 31), transcription factors of the forkhead family (51, 52), nitric oxide synthase (53), and caspase-9 (54). A study of Lu and colleagues demonstrated that intratracheal instillation of constitutively active Akt using an adenovirus vector (myr-Adeno-Akt) protects against lung injury and prolongs survival in mice (32). This is consistent with our finding that Akt levels and activity are significantly decreased as cell viability decreases. A decrease in Akt activity by caspase-mediated cleavage of the Akt protein has been previously shown (46, 47). In our study, we were not able to reverse hyperoxia-mediated Akt protein loss with ZVAD. However, we were able to show that hyperoxia significantly decreases the level of Akt mRNA production. This finding implies that total Akt protein loss, under our experimental conditions, is not due to caspase-mediated protein cleavage, but may be due to Akt mRNA downregulation. To our knowledge, this is a novel mechanism of hyperoxia-induced Akt protein loss.

Both ERK and Akt have been demonstrated to inhibit apoptosis by phosphorylating and inactivating caspase-9 (55). This suggests that ERK activity may protect cells from the proapoptotic effects of loss of Akt. The ERK signaling pathway has been implicated in regulating cell growth, differentiation (56, 57), and survival (35). Although ERK has been known to be a prosurvival mediator, its role in hyperoxia remains elusive. Prior studies have shown that hyperoxia activates ERK, but there is conflicting evidence on whether ERK activation is protective or detrimental to cells in this environment. Our study shows that ERK is protective against hyperoxia-mediated epithelial cell injury. Inhibition of ERK activation caused significantly earlier and higher levels of cell death in response to hyperoxia.

Our finding that ERK is protective against hyperoxia-induced cell injury and death is in accord with several prior studies. Jones and colleagues found that hyperoxia generates reactive oxygen species, which, in turn, activate the MEK1/2 pathway and, subsequently, induces early growth response gene–1. They proposed that early growth response gene–1, which is known to be activated by various forms of stress, and may be involved in cell signaling pathways, confers protection in hyperoxia (34). Buckley and colleagues demonstrated that rat epithelial cells cultured on laminin are protective against DNA strand breakage and apoptosis induced by hyperoxia. They speculated that hyperoxia stimulates signaling from the basement membrane, which, in turn, activates the ERK1/2 pathway and, subsequently, leads to protection against cell death (35). Additionally, Allan and colleagues have shown that ERK phosphorylates caspase-9 at threonine 124 and subsequently protect HeLa cells from caspase-3–mediated apoptosis (55).

One study that conflicts with our data is the study by Zhang and colleagues, demonstrating that, in a mouse epithelial cell line and in whole animal studies, hyperoxia increases reactive oxygen species leading to ERK activation, increased cytochrome C release, and apoptosis (10). The reasons for these discordant data are unclear. It is potentially a species difference, as their experiments are in a murine system and our experiments and others are in human lung epithelial cells. Additionally, in the study by Zhang and colleagues, only ERK activity at very early time points (up to 1 h) was evaluated, as opposed to the extended activation seen in our studies. A number of recent studies in other systems have investigated the concept that extended ERK activity has very different outcomes than transient ERK activity (58, 59), suggesting that the extended ERK activation in the human cells prevents the apoptosis seen in murine cells where there is only a transient ERK activation. As an aggregate, our study shows that hyperoxia primarily causes necrotic cell death in epithelial cells. Hyperoxia-induced epithelial cell death is associated with decreased Akt activity, protein, and mRNA. Further, ERK activity contributes to delayed cell death in hyperoxia-exposed airway epithelial cells.


   Acknowledgments
 
This manuscript was supported by a Vererans Administration Merit Review grant, National Institutes of Health (NIH): HL-60316 and NIH HL-077431, and RR00059 from the General Clinical Research Centers Program, National Center for Research Resources, NIH.


   Footnotes
 
Conflict of Interest Statement: S.V.T. has no declared conflicts of interest; M.M.M. has no declared conflicts of interest; T.O.Y. has no declared conflicts of interest; L.S.P. has no declared conflicts of interest; T.N. has no declared conflicts of interest; and G.W.H. has no declared conflicts of interest.

Received in original form April 29, 2004

Received in final form August 4, 2004


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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