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Am J Physiol Lung Cell Mol Physiol 280: L779-L791, 2001;
1040-0605/01 $5.00
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Vol. 280, Issue 4, L779-L791, April 2001

Glutamine protects mitochondrial structure and function in oxygen toxicity

Shama Ahmad1, Carl W. White1, Ling-Yi Chang2, Barbara K. Schneider1, and Corrie B. Allen1

Departments of 2 Medicine and 1 Pediatrics, National Jewish Medical and Research Center, Denver, Colorado 80206


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Glutamine is an important mitochondrial substrate implicated in the protection of cells from oxidant injury, but the mechanisms of its action are incompletely understood. Human pulmonary epithelial-like (A549) cells were exposed to 95% O2 for 4 days in the absence and presence of glutamine. Cell proliferation in normoxia was dependent on glutamine, and glutamine deprivation markedly accelerated cell death in hyperoxia. Glutamine significantly increased cellular ATP levels in normoxia and prevented the loss of ATP in hyperoxia seen in glutamine-deprived cells. Mitochondrial membrane potential as assessed by flow cytometry with chloromethyltetramethylrosamine was increased by glutamine in hyperoxia-exposed A549 cells, and a glutamine dose-dependent increase in mitochondrial membrane potential was detected. Glutamine-supplemented, hyperoxia-exposed cells had a higher O2 consumption rate and GSH content. Electron and fluorescence microscopy revealed that, in hyperoxia, glutamine protected cellular structures, especially mitochondria, from damage. In hyperoxia, activity of the tricarboxylic acid cycle enzyme alpha -ketoglutarate dehydrogenase was partially protected by its indirect substrate, glutamine, indicating a mechanism of mitochondrial protection.

human; airway; epithelium; adenosine 5'-triphosphate; alpha -ketoglutarate dehydrogenase; mitochondrial membrane potential


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

HYPEROXIA CAUSES reactive oxygen species-mediated injury to lung cells that may contribute to the pathogenesis of various lung diseases (18, 23, 35, 45). In addition, elevated concentrations of O2 in cells in vitro cause inhibition of cellular proliferation and lung growth by effecting changes in mitochondrial metabolism and respiration and by causing DNA damage (10, 38). Hyperoxia rapidly inhibits aconitase, the initial enzymatic step in the tricarboxylic acid (TCA; Krebs) cycle both in cultured cells and in the lungs of rats (19) and newborn primates (35). Cellular respiration declines in parallel with the loss of aconitase activity in cells cultured in hyperoxia, and inhibitors of aconitase cause a similar decrease in respiration in cells cultured under normal O2 tensions (19). One mechanism by which cultured cells adapt to the stress of hyperoxic exposure is through increased glycolysis (4, 24). In vivo, the lungs of rats adapted to hyperoxia have increased total activity of hexokinase (3), the rate-limiting step in glycolysis in the rat lung (42). In addition, a novel isoform, hexokinase II, is expressed in the lungs of these rats, and there also is increased expression of hexokinase III, a nuclear isoform.

Although impairment of the aconitase step occurs rapidly during hyperoxic exposure, inhibition of subsequent steps in the TCA cycle pathway occurs later and less completely (24). The alternate substrate glutamine enters the TCA cycle subsequent to the aconitase step. Therefore, we hypothesized that increased utilization of glutamine could contribute to hyperoxic adaptation.

Mitochondria are a potential target of injury by oxygen radicals, and an alteration in mitochondrial membrane function is an important component of oxidative stress in cells (5, 52). Because the mitochondrial membrane potential (MMP) in situ is a measure of the energetic state of the cell as well as a sensitive indicator of mitochondrial function, we assessed the electrical potential across the inner mitochondrial membrane of air- and O2-exposed human pulmonary epithelial-like (A549) cells. This was done by the use of flow cytometry and the specific dye chloromethyltetramethylrosamine (CMTMRos; MitoTracker Orange, Molecular Probes, Eugene, OR). It was found that the inner MMP was profoundly affected by the presence or absence of glutamine, a mitochondrial substrate, in the growth medium of O2-exposed A549 cells. We also performed a series of experiments to evaluate the effect of glutamine supplementation on survival, growth, cell and organelle morphology, ATP content, respiration, alpha -ketoglutarate dehydrogenase activity, glutamine consumption, and cellular glutathione (GSH) content of these cells in hyperoxia. Our findings indicate that cells cultured in air in the absence of glutamine can survive, but not proliferate, and preserve mitochondrial integrity. In hyperoxia, on the other hand, these cells can neither proliferate nor survive, and their death is preceded by degeneration of their mitochondria. Paradoxically, cells provided with glutamine utilized the amino acid at a considerably increased rate in hyperoxia compared with cells exposed to normal O2 tensions.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cells and culture. The human epithelial-like lung carcinoma cell line A549 was obtained from the American Type Culture Collection (Manassas, VA). The cells were grown in 100-mm Falcon tissue culture dishes in 10 ml of F-12K growth medium (GIBCO BRL, Life Technologies, Grand Island, NY) containing 10% fetal calf serum, 100 U/ml of penicillin, 100 µg/ml of streptomycin, 20 mM glucose, and 2 mM glutamine incubated at 37°C under a humidified atmosphere of air containing 5% CO2. A549 cultures were routinely passaged by trypsinization and subcultured at an initial plating density of 0.5 million cells/plate. Hyperoxic exposures were maintained at Denver's atmospheric pressure (635 mmHg) and performed in a humidified airtight plastic incubator chamber (Billups Rothenberg, Del Mar, CA) gassed with 95% O2-5% CO2, and the cultures were incubated at 37°C. For experiments, custom-made F-12K medium (GIBCO BRL) without L-glutamine was used. It was supplemented with fresh L-glutamine for the studies involving the absence and presence of glutamine. Fresh medium was supplied daily during the hyperoxic exposures.

Small-airway epithelial cells (SAECs) were purchased as frozen primary cultures from Clonetics (Walkersville, MD). They were cultured in 100-mm Falcon tissue culture dishes in 10 ml of the supplier's SAEC basal medium supplemented with gentamicin, amphotericin B, bovine pituitary extract, hydrocortisone, human epithelial cell growth factor, epinephrine, transferrin, insulin, retinoic acid, triiodothyronine, and bovine serum albumin as per the supplier's recommendations. Although human SAECs can be maintained in culture for five to six passages with the supplier's serum-free medium, all experiments presented here were performed with SAECs that had been passaged a maximum of four times to ensure no loss of phenotype. Cultures were split at 80-90% confluence by trypsin digestion and subcultured in the same SAEC medium.

Measurement of MMP. MMP was estimated by the uptake of a fixable dye, CMTMRos (MitoTracker Orange, Molecular Probes) according to the method of Macho et al. (30). Thirty minutes before the end of hyperoxic exposure, the medium overlying the cells was replaced with gas-conditioned (air- or O2-containing) medium containing 150 nM CMTMRos and returned to the incubator. At the end of the exposure, the medium over the cells was aspirated, and the cells were washed once with phosphate-buffered saline (PBS) and then harvested with a dye-containing trypsin-EDTA solution. The cells were pelleted at 200 g for 10 min, and the supernatant was removed. The cell pellet was resuspended in 1 ml of PBS and then fixed with 1 ml of 8% paraformaldehyde (Electron Microscopy Sciences, Fort Washington, PA) in PBS (pH 7.4). After incubation in the dark at room temperature on a shaker for 30 min, the cells were kept on ice and analyzed with an EPICS XL flow cytometer (Coulter, Hialeah, FL) operated by Coulter's System II software and incorporating an argon laser (488 nm, 15 mW) for excitation. MitoTracker Orange fluorescence was assessed in FL2 (575-nm band-pass filter). Carbonyl cyanide m-chlorophenylhydrozone (CCCP; Calbiochem, San Diego, CA) dissolved in dimethyl sulfoxide (DMSO) was added along with the dye MitoTracker Orange for MMP inhibition studies. List mode files were collected for each sample and transferred to a Macintosh G3 computer for subsequent analysis. Mean fluorescence intensity (MFI) of the cells was used as the primary index for comparison of MMP. Corrections for any changes in MFI due to forward light scatter (FS) were done by plotting MFI against FS. The slope (MFI/FS) of these distributions for each sample, calculated from the regression analysis and analysis of covariance (ANCOVA) with JMP software (SAS Institute, Cary, NC), was considered a better indicator of MMP.

Cellular O2 consumption. Cellular O2 consumption was measured in a custom-built six-place respirometer. Each chamber of this apparatus consisted of a glass water-jacketed cell (Gilson Medical Electronics) fitted with a Clark-style polarographic O2 electrode (model 5331, Yellow Springs Instruments, Yellow Springs, OH). The six chambers were fixed on a multiposition electromagnetic stir plate (Cole-Parmer, Vernon Hills, IL) placed within a tissue culture incubator maintained at 37°C. The six electrodes were connected to a chemical microsensor II (Diamond General Development, Ann Arbor, MI) through a 10-channel multiplexer. Channel selection and data collection were achieved by using LabVIEW software (National Instruments, Austin TX). Each electrode was preequilibrated with 1.4 ml of medium. Next, 1 million cells were added in a 100-µl volume, and the stopper was placed in the chamber. O2 concentration was measured for ~1 h, and the slopes representing O2 consumption were calculated. The medium O2 saturation values were calculated with the phenazine methosulfate-NADH method as described by Robinson and Cooper (40).

Electron microscopy. For electron-microscopic analysis, control and hyperoxic cells with and without glutamine were fixed in 2% glutaraldehyde in 0.085 M sodium cacodylate buffer, pH 7.4, containing 0.05% CaCl2. After being scraped, the fixed cells were suspended in fresh fixative and pelleted at 300 g for 10 min. After dehydration and embedding in resin, thin sections were cut with a Reichart Ultra Cut E microtome. The sections were collected on 0.4% Formvar-coated, 100-mesh circular grids (3-mm diameter) and stained with 2% uranyl acetate and 2% lead citrate for 15 min. The sections were examined for mitochondria with a Philips CM-10 electron microscope at 8 kV, and the images were photographed.

Fluorescence microscopy. A549 cells were seeded onto 22-mm glass coverslips in six-well plates at a density of 3 × 105 cells/well in both the presence and absence of glutamine. Hyperoxic exposures were performed as described in Cells and culture. At the end of exposure, dye (MitoTracker Orange, Molecular Probes) dissolved in DMSO and diluted with warm medium to a concentration of 150 nM was added. The cells were incubated with the dye for 30 min at 37°C under the appropriate conditions (21 or 95% O2). The coverslips with adherent cells were then rinsed with PBS and fixed in 4% paraformaldehyde for 10 min. After being rinsed, each coverslip was mounted upside down onto a glass slide with aqueous antifade mounting medium (ProLong, Molecular Probes) and allowed to dry overnight. An Olympus Vanox-T fluorescent microscope attached to a digital camera (Cooke, Auburn Hills, MI) was used to examine the fixed cells. Images were recorded with Slide Book 2.6.5.5 software (Intelligent Imaging Innovations, Denver, CO) on a Macintosh G3 computer.

Biochemical assays. Cellular growth was measured with the MTT assay (34) with the water-soluble tetrazolium salt 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (Sigma). Cells were plated in 96-well half-area tissue culture plates (Costar 3696), and the medium was replaced by 100 µl of a 1:1 serum- and phenol red-free DMEM-F-12 medium mixture. Fifty microliters of MTT (4 mg/ml) in the serum- and phenol red-free DMEM-F-12 medium mixture were added, and the plate was incubated for 4 h at 37°C. The purple formazan crystals thus formed were dissolved in 50 µl of DMSO, and the optical density of the wells on the plate was read at 540 nm with a plate reader.

Trypan blue exclusion was performed by adding 25 µl of 0.1% trypan blue solution to 100 µl of cells suspended in PBS, and the cells that excluded the dye were counted with a hemacytometer.

For propidium iodide (PI) staining of nonviable cells, ~106 cells were suspended in 1 ml of PBS, and PI (2 µg/ml final concentration) was added. After incubation for 5 min on ice in the dark, flow cytometric analysis was performed.

The protein content of cells was measured with a DC protein assay kit (Bio-Rad Laboratories, Hercules, CA). For ATP analysis, the cells were harvested, an extract was prepared as previously described (4), and total cellular ATP content was estimated with a luciferase-luciferin kit (Analytical Luminescence Laboratory, Sparks, MD). The glutamine content of the medium was estimated by cation-exchange chromatography on a Beckman 6300 amino acid analyzer in a lithium citrate buffer (33).

For the measurement of alpha -ketoglutarate dehydrogenase activity, air- and O2-exposed cells were washed with PBS and harvested in 1.0 ml of ice-cold Tris · HCl buffer (25 mM Tris, pH 7.4, supplemented with 0.25 M sucrose, 2 mM EDTA, 10 mM K2HPO4, 5.0 mM MgCl2, 2.0 mM KCN, and 2 mM glutamine). After centrifugation at 12,000 g for 1 min, the pellet obtained was suspended in 200 µl of the above buffer and sonicated at 10% (setting 3) power three times in 10-s bursts with a model 50 sonic dismembranator (Fisher Scientific). alpha -Ketoglutarate dehydrogenase activity was measured essentially as described by Bergmeyer (9), at 340 nm and in the presence of 2 mM NAD+, 20 µM coenzyme A, 2 mM KCN, 200 µM thiamine pyrophosphate, and 2 mM alpha -ketoglutarate. The assay buffer contained 25 mM Tris · HCl, 0.25 M sucrose, 2 mM EDTA, 10 mM K2HPO4, and 5.0 mM MgCl2.

The total intracellular GSH was determined with the 5,5'-dithio-bis(nitrobenzoic acid)-glutathione reductase recycling assay (7, 13). A549 cells were harvested in PBS from different incubation conditions and transferred to microcentrifuge tubes. A volume of 200 µl of 2.5% sulfosalicylic acid with 0.2% Triton X-100 was added to the pellet and centrifuged. The supernatant was harvested, and 30 µl from each tube were transferred to a 96-well plate. After this step, 140 µl of 0.3 mM NADPH in stock buffer solution (125 mM sodium phosphate and 6.3 mM EDTA, pH 7.5) and 100 µl of glutathione reductase (1 U/ml; Sigma) were added to each well. Finally, the substrate 5,5'-dithio-bis(nitrobenzoic acid) (20 µl of a 6 mM solution; Sigma) was added to the reaction. The absorbance at 405 nm of each well was read with a microplate reader. GSH was quantified with a GSH standard curve.

Statistical analysis. All statistical calculations were performed with JMP software (SAS Institute, Cary, NC). Means were compared by one-way analysis of variance followed by two-tailed t-test for comparison between two groups and the Tukey-Kramer test for multiple comparisons. A P value of <0.05 was considered significant.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Effects of glutamine on cellular proliferation and survival in hyperoxia. The proliferation of cells was highly dependent on glutamine. Figure 1 shows the growth of A549 cells in the absence and presence of glutamine as assessed by the MTT assay. Cells deprived of glutamine did not grow at all during 4 days of culture. In 21% O2 and in the presence of glutamine (1-4 mM), the cells proliferated rapidly. Glutamine at a concentration of 1 mM appeared adequate, if not optimal, for the survival and proliferation of these cells.


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  		Fig. 1.   Effect of glutamine (Gln) supplementation on A549 cell growth in 21% O2. Cells were plated at a density of 5,000 cells/well in a 96-well tissue culture plate in the absence and presence of 1, 2, and 4 mM glutamine in F-12K medium supplemented with 10% FCS. Cell growth was assessed at each experimental point (24 h) with the MTT assay as described in METHODS. Fresh medium was supplied daily during the experiment. Values are means ± SE of each experiment performed in triplicate. * Significant difference from simultaneous control A549 cells without glutamine for the 3 overlapping symbols, P < 0.05 by t-test.

Cell survival in hyperoxia was profoundly affected by glutamine. The effect of glutamine on the survival of A549 cells in 21 and 95% O2, measured by counting trypan blue-excluding cells, is illustrated in Fig. 2. In hyperoxia, the glutamine-deprived cells showed a gradual decline in number, with only 25-30% of the cells remaining on the 4th day of exposure (0.128 × 106 ± 0.022 × 106 cells on the 1st day and 0.034 × 106 ± 0.01 × 106 cells on day 4). Glutamine-supplemented cells did not proliferate in the presence of elevated O2 tension but were able to survive for up to 4 days of exposure (0.150 × 106 ± 0.023 × 106 cells on the 1st day and 0.152 × 106 ± 0.018 × 106 cells on day 4 in 4 mM glutamine).


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  		Fig. 2.   Effect of glutamine on the viability of A549 cells in hyperoxia on days 1-4 of exposure. Cells were plated in a 24-well plate at a density of 10,000 cells/well in the absence and presence of glutamine and were exposed to 95% O2 (oxy) at 635 mmHg. Viable cells were examined by their ability to exclude trypan blue at each time interval (24 h) and compared with their respective 21% O2 controls. [Glutamine], glutamine concentration. Values are means ± SE of each experiment performed in triplicate; error bars not seen are within bars. * Significant difference from simultaneous control, air-exposed A549 cells with and without glutamine, P < 0.05 by t-test.

These results were further supported by PI staining to indicate the presence of nonviable cells (Fig. 3). These studies showed a significant (~70%) loss of viability of cells after the 4th day of hyperoxic exposure in the absence of glutamine. Glutamine supplementation prevented >50% of cell death at this time.


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  		Fig. 3.   Influence of glutamine on A549 cell survival. For the identification of nonviable cells by propidium iodide (PI) staining, A549 cells were plated at a density of 0.5 × 106 in 100-mm tissue culture plates with and without glutamine. They were exposed to 95% O2-5% CO2 for 4 days. Fresh medium was supplied daily, and the floating cells were pelleted and added back to the plates for analysis at 96 h. PI-stained cells (details in METHODS) were analyzed on an EPICS XL flow cytometer (see Estimation of MMP) with excitation at a wavelength of 488 nm (15-nm argon laser). PI fluorescence was assessed in FL3 (620-nm band-pass filter). Values are means ± SE of each experiment performed in triplicate; error bars not seen are within bars. Inset: bivariate plots of cell count vs. PI fluorescence. Dead cells are >2 decades brighter than live ones. a: PI staining of glutamine-deprived, hyperoxia-exposed cells. b: glutamine-supplemented, hyperoxia-exposed cells. c and d: respective 21% O2 controls. * Significant difference from simultaneous control, air-exposed A549 cells with and without glutamine, P < 0.05 by t-test. ** Significant difference from all other groups, P < 0.05 by Tukey-Kramer test.

Effect of glutamine on cellular ATP content, MMP, and respiration. Cellular ATP content of the glutamine-supplemented cells in 21% O2 normalized to cell protein was greater by 30-40% than that of the glutamine-deprived cells. Glutamine supplementation also prevented the loss of ATP observed in the unsupplemented cells after 4 days of hyperoxic exposure (Fig. 4). In addition, in the 95% O2-exposed cells, relatively less change in ATP content was observed to be related to the level of glutamine supplementation (1-4 mM) among the various groups of glutamine-supplemented cells. Nonetheless, there was a glutamine concentration-dependent increase in cell ATP content both in normoxia and in hyperoxia over the 0-2 mM glutamine concentration range. However, there was no further increase in cell ATP content at higher glutamine concentrations (4 mM) under either level of O2 exposure.


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  		Fig. 4.   Effect of glutamine on ATP levels in hyperoxia-exposed A549 cells. Cells were plated and exposed in the absence and presence of glutamine as described in METHODS. Values are means ± SE; n = 3 experiments; error bars not seen are within bars. * Significant difference from control A549 cells without glutamine, P < 0.05 by t-test. ** Significant difference from all other groups, P < 0.05 by Tukey-Kramer test.

Treatment of glutamine-supplemented A549 cells with 95% O2 for 4 days resulted in an increase in CMTMRos fluorescence (MFI), an indication of MMP (Fig. 5A). This increase was threefold in 1 and 2 mM glutamine-supplemented cells [52.96 ± 0.52 (SE) fluorescence intensity units without glutamine vs. 165 ± 0.40 fluorescence intensity units with 1 mM glutamine and 48.73 ± 1.6 vs. 163 ± 3.0 fluorescence intensity units with 2 mM glutamine] and more than threefold in 4 mM glutamine-supplemented cells (62.56 ± 1.1 vs. 229.3 ± 2.7 fluorescence intensity units). An increase in FS also was detected in the glutamine-supplemented cells after hyperoxic exposure (Fig. 5B). Changes in FS light intensity are believed to indicate changes in cellular and/or mitochondrial volume. We have observed that correction of MFI for FS considerably reduces the variability in the data produced by changing FS values. Therefore, we expressed estimated MMP as MFI and as the corrected MFI/FS slope value. MFI/FS slopes (Fig. 5C) were generated as described in METHODS. A dose-dependent increase in the slope was observed with glutamine treatment in the hyperoxia-exposed cells. Thus the group supplemented with the highest concentration (4 mM) of glutamine had the highest MMP as indicated by the slope values [MFI/FS 1.496 ± 0.001 (SE) vs. 3.7 ± 0.02 in O2]. A 50% decrease in MFI was measured in the glutamine-deprived cells on hyperoxic exposure for 4 days (95.46 ± 1.56 vs. 177.73 ± 1.35; Fig. 5A). Similarly, a consistent decrease was also seen in the MFI/FS slope values of these cells (1.93 ± 0.065 vs. 3.15 ± 0.04; Fig. 5C). The effect of CCCP, a potent protonophore, on the MFI, FS, and MFI/FS slope of glutamine-supplemented and -unsupplemented air- and O2-exposed cells also was examined (Fig. 6). Treatment of the glutamine-deprived A549 cells with 40 µM CCCP resulted in a 40% decrease in the MFI in 21% O2 (109.5 ± 0.4 vs. 171 ± 1.25) and a 27% decrease in the MFI in hyperoxia (64.067 ± 2.3 vs. 87.2 ± 1.1). Treatment of glutamine (2 mM)-supplemented cells with CCCP caused a 40% decrease in the MFI in 21% O2 (36.9 ± 0.68 vs. 49.7 ± 1.35) and a similar percent decrease in MFI (38%) in hyperoxia in these cells (109.53 ± 3.15 vs. 160.6 ± 2.69) relative to that in unsupplemented cells. Because CCCP-inhibitable MFI indicates a proton gradient-dependent MMP, these findings ensure that MFI and MFI/FS changes observed in glutamine-deprived cells signify a loss of MMP.


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  		Fig. 5.   Measurement of chloromethyltetramethylrosamine (MitoTracker Orange) fluorescence, an indicator of mitochondrial membrane potential, in human lung epithelial-like A549 cells exposed to 95% O2 (635 mmHg) for 4 days in absence and presence of glutamine. Cells were stained and examined by flow cytometry as described in METHODS. A: mean fluorescence intensity (MFI) of A549 cells exposed to 21 or 95% O2. B: forward scatter (FS). C: MFI/FS slope values. Values are means ± SE; n = 3 experiments; error bars not seen are within bars. * Significant difference from simultaneous control, air-exposed A549 cells with and without glutamine, P < 0.05 by t-test.



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  		Fig. 6.   Effect of mitochondrial uncoupler carbonyl cyanide m-chlorophenylhydrozone (CCCP) on flow cytometric estimation of mitochondrial membrane potential by MitoTracker Orange fluorescence in A549 cells. Cells were treated with 40 µM CCCP for 30 min at 37°C, harvested, and examined by flow cytometry as described in METHODS. A: CCCP-inhibitable component of MFI (MFI - MFICCCP). B: CCCP-inhibitable components of MFI/FS (slope - slopeCCCP). * Significant difference from simultaneous control, air-exposed A549 cells with and without glutamine, P < 0.05 by t-test.

Glutamine had a sparing effect on respiration in hyperoxia. The effect of hyperoxic exposure (4 days) on cellular O2 consumption is shown in Fig. 7. Glutamine supplementation partially preserved O2 consumption in hyperoxia, whereas a total loss of respiration was observed in glutamine-unsupplemented cells. Glutamine supplementation also supported the O2 consumption of A549 cells grown in 21% O2 and increased it twofold compared with that in glutamine-deprived cells.


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  		Fig. 7.   Evaluation of respiratory capacity of A549 cells exposed to hyperoxia for 4 days in the absence and presence of glutamine. After hyperoxic exposure in various experimental conditions, the cells were harvested by trypsinization. Aliquots of these cells were studied in the respirometer fitted with a calibrated polarographic O2 electrode as described in METHODS. Values are means ± SE; n = 3 experiments; error bars not seen are within bars. * Significant difference from simultaneous control, air-exposed A549 cells with and without glutamine, P < 0.05 by t-test. ** Significant difference from all other groups, P < 0.05 by Tukey-Kramer test.

Biochemical effects of glutamine deprivation. Because glutamine, the substrate, is known to stabilize the enzyme alpha -ketoglutarate dehydrogenase, its activity was measured. A complete loss of alpha -ketoglutarate dehydrogenase activity was observed during cellular exposure to hyperoxia in the absence of glutamine (Fig. 8). However, glutamine supplementation partially protected against the loss of enzyme activity. In the presence of glutamine, the cells in 21% O2 had a threefold increase in alpha -ketoglutarate dehydrogenase activity compared with the unsupplemented cells.


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  		Fig. 8.   Effect of glutamine supplementation on alpha -ketoglutarate dehydrogenase activity of A549 cells exposed to hyperoxia for 4 days. Values are means ± SE; n = 3 experiments. * Significant difference from simultaneous control, air-exposed A549 cells in air with and without glutamine, P < 0.05 by t-test. ** Significant difference from all other groups, P < 0.05 by Tukey-Kramer test.

Glutamine consumption by 1 mM glutamine-supplemented hyperoxia-exposed cells was significantly higher than in normoxia-exposed control cells (Fig. 9), but the cellular utilization of glutamine did not increase with additional increases of glutamine in the growth medium.


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  		Fig. 9.   Effect of hyperoxic exposure on glutamine consumption of A549 cells. Cells were seeded at a density of 0.5 × 106 on 100-mm dishes with and without glutamine and exposed to 95 or 21% O2 at 635 mmHg for 4 days. At the end of the exposure, supernatant medium was collected and centrifuged at 1,000 g, and the glutamine content was determined. Values are means ± SE; n = 3 experiments; error bars not seen are within symbols. * Significant difference from simultaneous control, air-exposed A549 cells with and without glutamine, P < 0.05 by t-test.

Because glutamine is an important substrate for GSH synthesis and GSH is an important antioxidant in cellular tolerance to oxidative stress, total cellular GSH also was measured. Table 1 shows the effect of glutamine on the total GSH content of normoxia- and hyperoxia-exposed cells. The total GSH content of hyperoxia-exposed glutamine-unsupplemented cells decreased by 42% compared with that in similar cells in normoxia (2.8 ± 0.4 vs. 4.8 ± 0.4 nmol/107 cells). Glutamine-supplemented cells maintained their GSH content in hyperoxia.

                              
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  		Table 1.   Effect of glutamine on total GSH content

Effects of glutamine deprivation on cellular and mitochondrial morphology. The effect of glutamine supplementation on the cellular morphology of A549 cells in normoxia was studied with electron microscopy (Fig. 10). Sections of A549 cells grown in normoxia (21% O2-5% CO2) revealed that in the absence of glutamine, the cells appeared thin, with spreading into monolayers (Fig. 10d). They had dense but normal-appearing mitochondria (Fig. 11a). There were no apparent ultrastructural abnormalities present except that the cell membranes had only small indentations with virtually no microvilli. With 1 mM glutamine supplementation (Fig. 10e), the cells appeared larger than those lacking glutamine, were cuboidal in shape, showed spreading into monolayers, and had numerous surface microvilli, suggesting enhanced metabolic activity. Mitochondria were abundant in these cells, and they appeared larger and had a less dense matrix than those in glutamine-unsupplemented cells. With 4 mM glutamine supplementation (Fig. 10f), the cells also appeared thicker and had a cuboidal shape. These cells possessed numerous surface microvilli. Glutamine-supplemented A549 cells had abundant Golgi apparatus, endoplasmic reticulum, ribosomes, and fibrillar bundles (Fig. 10, e and f), again suggesting a greater state of differentiation.


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  		Fig. 10.   Electron micrographs of A549 cells cultured in the absence (d) and presence of 1 (e) and 4 (f) mM glutamine in 21% O2 and of A549 cells exposed to hyperoxia in the absence (a) and presence of 1 (b) and 4 (c) mM glutamine. a: thin long arrow, lysosomes; thick short arrow, cellular debris in cells exposed to hyperoxia in the absence of glutamine. d: thick short arrow, lack of microvilli on the flattened surface of a cell deprived of glutamine in 21% O2, whereas microvilli are abundant (see space between 2 cells in e) in cells grown with glutamine supplementation at that gas tension. Original magnification, ×6,000.



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  		Fig. 11.   Electron micrograph of mitochondria in A549 cells exposed to hyperoxia in absence (b) and presence (c-h) of glutamine. a: dense round mitochondria of 21% O2-exposed A549 cells without glutamine supplementation. Original magnification, ×47,500.

The cells also were examined for hyperoxia-mediated morphological changes in both the absence and presence of glutamine. Hyperoxia (95% O2-5% CO2)-exposed cells grown in glutamine-free medium (Fig. 10a) were irregular or spherical in shape. There was extensive cellular disintegration and injury, complete loss of cell-to-cell contacts, and cellular aggregation rather than spreading into monolayers. Considerable cellular debris was visible, and the remaining intact cells showed considerable and extensive lysosome and vacuole formation. Intact mitochondria were very rarely detected in these cells (Fig. 11b). Cells exposed to hyperoxia and supplemented with glutamine (1 mM; Fig. 10b) exhibited less necrosis, less aggregation, more spreading into a monolayer, and fewer vacuoles and lysosomes than cells lacking glutamine. There was a spectrum of mitochondrial morphologies observed in hyperoxia-exposed glutamine-supplemented cells (Fig. 11, c-h), with some appearing relatively normal, some appearing distended and empty, and some with very few cristae apparent. Additional morphological abnormalities such as the appearance of "mitochondria within mitochondria" also were detected in hyperoxia-exposed cells (Fig. 11e).

With 4 mM glutamine supplementation (Fig. 10c), the cells also maintained normal spreading and appeared less necrotic than those lacking glutamine. Few vacuoles and lysosomes were detected, and the mitochondria appeared similar to those in 1 mM glutamine-supplemented cells exposed to hyperoxia. In general, hyperoxia-exposed cells supplemented with 1 or 4 mM glutamine had a similar appearance.

To further confirm the changes in mitochondrial size and function on glutamine supplementation and hyperoxic exposure, A549 cells were stained with the potential-sensitive dye CMTMRos and observed under a fluorescence microscope. Mitochondria in normoxia-exposed control cells with or without glutamine showed thin filamentous structures ("spaghetti"; Fig. 12, a and b). In the presence of glutamine and hyperoxia, the mitochondria appeared more round in shape and densely packed around the nucleus ("meatballs"). Hyperoxic exposure of glutamine-unsupplemented cells resulted in a complete loss of the filamentous structure. In glutamine-supplemented cells, the mitochondria were predominantly enlarged and granular, suggesting the formation of megamitochondria. The latter were confirmed by electron microscopy (see above).


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  		Fig. 12.   Fluorescence micrographs of A549 cells stained with chloromethyltetramethylrosamine. Cells were cultured in the absence (a) and presence (b) of 1 mM glutamine and exposed to 21% O2. Additional cells were exposed to hyperoxia for 4 days without (c) and with (d) 1 mM glutamine. Other experimental details are given in METHODS.

Biochemical effects of glutamine deprivation in primary human SAECs. A549 cells are tumor-derived lung epithelial cells with type II pneumocyte characteristics. The metabolism of transformed or tumor-derived mammalian cell lines is significantly different from the primary mammalian cell lines. However, glucose and glutamine are usually the growth-limiting nutrients in all mammalian cells. A recent study (11) on glutamine dependence and utilization by normal and tumor-derived breast cell lines revealed that the origin of cell lines is not a determinant of glutamine metabolism and that both primary and tumor cells exhibit similar glutamine dependence and utilization rates. To determine whether a similar protective role could be attributed to glutamine in a primary cell line, we studied SAECs in hyperoxia with and without glutamine supplementation (Fig. 13).


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  		Fig. 13.   Effect of glutamine on the ATP content (A), respiration (B), and survival (C) of small-airway epithelial cells in 21 (air) and 95% O2. Small-airway epithelial cells were cultured in absence and presence (6 mM) of glutamine and exposed to 95% O2 for 4 days. ATP content, O2 consumption, and cell survival were measured with procedures identical to those used for A549 cells. * Significant difference from 21% O2-exposed control cells. ** Significant difference from glutamine-supplemented, 95% O2-exposed cells.

About a 30-40% loss in the total cellular ATP content (Fig. 13A) was observed in SAECs exposed to hyperoxia in the absence of glutamine compared with those that were supplemented with 6 mM glutamine. These cells (glutamine-deprived O2-exposed SAECs) also had decreased O2 consumption rates (Fig. 13B) compared with the glutamine-supplemented cells, but the decline in respiration was not as severe as in the case of glutamine-deprived hyperoxia-exposed A549 cells. This was accompanied by a parallel decline in the growth of these cells in 21% O2 in the absence of glutamine (0.37 × 106 ± 0.01 × 106 cells compared with 0.50 × 106 ± 0.04 × 106 cells). Hyperoxic exposure of these cells resulted in growth arrest and extensive cell death, with only 0.09 × 106 ± 0.01 × 106 cells remaining in the absence of glutamine. Glutamine supplementation was associated with a greater absolute number of surviving cells in hyperoxia, to 0.14 × 106 ± 0.01 × 106 cells. In comparing glutamine-deprived and glutamine-supplemented cells exposed to hyperoxia, cellular ATP contents were significantly lower in glutamine-deprived cells. Differences in cell respiration and surviving cell numbers were marginally, but not statistically, significant.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Our present study demonstrated that glutamine deprivation greatly accelerates the toxicity of O2 to human pulmonary epithelial-like (A549) cells. Enhanced injury to mitochondria was evidenced by severe ultrastructural damage, decreased MMP, decreased cellular ATP content, and complete loss of cell respiration. Two biochemical mechanisms that may have contributed to these effects were identified: 1) a decline in cell GSH content and 2) a complete inhibition of alpha -ketoglutarate dehydrogenase activity in the TCA cycle. The essential nature of glutamine in oxidative stress was further supported when it was found that cell glutamine consumption was doubled during exposure to hyperoxia even in the presence of a marked decline in alpha -ketoglutarate dehydrogenase activity. In addition, we demonstrate for the first time that glutamine can protect alpha -ketoglutarate dehydrogenase from inactivation under oxidative stress (hyperoxia).

Glutamine, an important precursor of peptides, proteins, and nucleotides, also serves a critical role in cellular metabolic pathways (16, 36, 54). Acute glutamine deprivation is associated with DNA damage as well as elevated expression of growth-arrest genes (1, 21). Such deprivation often leads to a severe decline in cell viability (43). Glutamine is a highly labile constituent of cell culture medium, and yet most media are prepackaged to contain glutamine, thus severely limiting shelf life. Various investigators may or may not use fresh medium and/or add supplemental glutamine. The findings of the present study indicate that the effects of oxidative stress on cells may be profoundly affected depending on the actual concentrations of glutamine present during such exposures.

Glutamine deprivation significantly affected the growth of A549 cells, and hyperoxic exposure of these cells caused a markedly exaggerated injury. Another study (25) has also revealed the glutamine requirement of pulmonary endothelial cells and A549 cells for growth. In this study, glutamine supplementation increased cell proliferation in normoxia (21% O2), and 1 mM glutamine was adequate to support growth. Additionally, glutamine supplementation not only supported the growth of A549 cells but protected them from hyperoxia-mediated injury and death. In the experiments reported here, more than two-thirds of the cell death observed could have been prevented by glutamine supplementation.

Lung tissue and cellular injury, growth arrest, and accumulation of cells in the G1/G2 and S phases of the cell cycle commonly occur on hyperoxic exposures (12, 22, 39, 47). In this study, several important differences were found in the structure of A549 cells on hyperoxic exposure in the absence and presence of glutamine. Glutamine supplementation improved the mitochondrial viability such that, in hyperoxia, enlarged mitochondria, a spectrum of abnormal mitochondria, and even a few normal-looking mitochondria were observed. In the absence of glutamine, however, very few mitochondrial structures could be identified after hyperoxic exposure. Enlarged mitochondria with ruptured membranes have been found to be a feature of cells from lungs, hearts, and brains from hyperoxia-exposed animals (8, 37, 41). Enlarged mitochondria also frequently occur in hyperoxia-adapted HeLa cells, Chinese hamster ovary cells (24, 50), and other cells stressed by oxidants. Although the enlargement of mitochondria likely represents injury, it also may indicate adaptation to compensate during oxidative stress by increasing the ratio of surface area to volume within mitochondria.

Lung cells exposed to hyperoxia can generate free radicals like superoxide anion, hydroxyl, and alkyl radicals via mitochondrial electron transport (18). Mitochondrial DNA, metabolism, and function are highly susceptible to injury by these species. Such mitochondrial injury can contribute to the pathogenesis of necrotic and apoptotic cell death (17, 28, 38). Whether enhancement of cell survival in this study is related to energy homeostasis and protection of mitochondrial function was investigated by measuring the MMP and the total ATP content of hyperoxia-exposed glutamine-supplemented and -unsupplemented cells. In our studies of MMP, glutamine-deprived 21% O2-exposed control cells actually had higher MFI values than glutamine-supplemented normoxia-exposed cells. This could be due to hyperpolarization of the mitochondrial membrane caused by mitochondrial ATP accumulation in these metabolically inactive, growth-retarded cells, which may also have diminished ATP turnover. In addition, there was a remarkable increase in MFI observed in hyperoxia-exposed glutamine-supplemented cells (1-4 mM; Fig. 5). There were two possible causes for this increase in fluorescence. This could be due to either an increase in the MMP or an increase in the apparent MMP due to an increase in the total number of mitochondria per cell (49). Indeed, electron microscopy revealed morphological changes in mitochondria that were prominent in hyperoxia both with and without glutamine supplementation.

To help distinguish the changes in the MMP from changes due to potential alterations in background fluorescence, such as those that may result from altered mitochondrial volume, cells were treated with 40 µM CCCP, a potent protonophore, and stained with CMTMRos. In the presence of CCCP, the cells showed a considerable reduction in fluorescence that was dependent on the proton gradient (Fig. 6). On the other hand, there was considerable background fluorescence that persisted. This indicates that there is some considerable equilibration of the dye CMTMRos at the loading concentration (150 nM) within the mitochondrial matrix space and/or cytoplasm even with no potential across the inner mitochondrial membrane. This level of background fluorescence may result from nonspecific dye binding by mitochondrial or other cellular components. Therefore, that portion of total fluorescence that can be dynamically inhibited by the uncoupler CCCP is the portion that represents MMP. The difference in the mean MFI/FS slope values of control and hyperoxia-exposed cells with and without CCCP was calculated. The difference for the normoxic, glutamine-deprived cells was 1.06 and for hyperoxia-exposed, glutamine-deprived cells was 0.38. This indicates a significant decrease in H+-dependent MMP in the glutamine-deprived cells in hyperoxia relative to normoxia.

Fluorescence and electron microscopy further supported the protection of mitochondrial function. Mitochondria in glutamine-supplemented cells stained with mitochondria-specific CMTMRos are spread out rather than densely packed around the nucleus as in glutamine-deprived cells. Glutamine-supplemented cells in hyperoxia stained with CMTMRos and observed by fluorescence microscopy revealed the presence of large mitochondria, again confirming the swollen and enlarged mitochondria observed by electron microscopy. Electron-microscopic examination of hyperoxia-exposed glutamine-deprived cells revealed few detectable mitochondria in these cells. The positive CMTMRos staining of these cells (Fig. 12c) reflects the background fluorescence that is also evident in the flow cytometry results (Fig. 5). Background fluorescence, which is retained even after treatment with high concentrations of uncouplers, was reported in other studies (29, 49, 53) on flow cytometry with other mitochondrial dyes like rhodamine 123 and 5,5',6,6-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1).

Glutamine is a major energy source for mammalian cells in culture, and glutamine supplementation significantly increased the ATP levels in normoxic A549 cells (Fig. 4). However, during hyperoxic exposure, the ATP content in unsupplemented cells was significantly decreased relative to that in the glutamine-supplemented groups. The higher ATP content in glutamine-supplemented cells suggested that it can be used as an oxidizable substrate for ATP synthesis in cells under hyperoxic stress. This ATP could then be utilized for homeostatic, protective, and repair mechanisms. Schoonen et al. (45) have reported depletion of ATP on hyperoxic exposure of HeLa cells, which, in those cells, is associated with a complete inactivation of alpha -ketoglutarate dehydrogenase enzyme and decreased glutamine utilization. In our model, however, complete loss of alpha -ketoglutarate dehydrogenase activity was only observed in hyperoxia-exposed glutamine-deprived A549 cells, and glutamine supplementation of A549 cells caused a partial protection against the loss of alpha -ketoglutarate dehydrogenase activity during hyperoxic exposure (Fig. 8). Such exposure was also associated with an increased utilization of glutamine (Fig. 9) and an associated enhancement in ATP content. Because these cells are growth arrested due to hyperoxic exposure and may have decreased ATP turnover, some preservation of ATP may be understood. Here it is important to note that the ATP content is reported as a function of total protein. Therefore, changes due to any cellular hypertrophy or atrophy resulting from hyperoxic exposure are taken into account.

Studies in A549 cells previously done in our laboratory (19, 20) have shown that hyperoxia results in a rapid inactivation of aconitase that is accompanied by a parallel loss in both basal respiration and respiratory capacity. In addition, progressive respiratory failure has been reported in Chinese hamster ovary cells exposed to normobaric hyperoxia for 3 days (44). Hence the question of whether glutamine, an alternate indirect substrate that enters the TCA cycle subsequent to the aconitase step, modulates cellular O2 consumption in hyperoxia-exposed A549 cells was addressed. Glutamine supplementation partially preserved O2 consumption, whereas a total impairment of respiration was observed in glutamine-unsupplemented cells in hyperoxia. Glutamine supplementation supported the O2 consumption of A549 cells grown in air and increased it twofold compared with that in the glutamine-deprived cells. This finding shows that these cells are only partially dependent on glutamine for respiration during life in normoxic environments. However, they become completely dependent on glutamine for respiration during prolonged exposure to hyperoxia. This finding further confirms the essential nature of glutamine in oxidative stress.

The significant preservation of respiration in glutamine-supplemented cells supports the importance of the protection of alpha -ketoglutarate dehydrogenase activity that we observed in these cells. The nature of this protection in hyperoxia is unknown. We speculate that the active site could be directly protected by glutamine or that protection by glutamine was indirect, possibly provided by the increased GSH afforded by glutamine supplementation. The latter seems likely because others (38, 44) have reported inactivation of at least three -SH group-containing flavoprotein complexes, i.e., the NADH dehydrogenase, succinate dehydrogenase, and alpha -ketoglutarate dehydrogenase complexes on hyperoxic exposure of cells. Joenje et al. (24) adapted HeLa cells over several months for tolerance to 80% O2. These cells exhibited a similar O2 consumption to 20% O2-adapted control cells. In addition, antioxidant enzyme activities in these cells were not different before and after adaptation. On the other hand, Chinese hamster ovary cells resistant to 99% O2-1% CO2 had two- to fourfold increased activities of superoxide dismutase, catalase, and glutathione peroxidase (51). The alpha -ketoglutarate dehydrogenase activity of these cells was found to be resistant to hyperoxic damage, and they had an enhanced capacity to respire and survive under hyperoxic conditions (46). Hence the increased tolerance was postulated to be largely attributed to a genetically determined increased resistance of O2-sensitive cellular targets like alpha -ketoglutarate dehydrogenase. Our results indicate an increased resistance conferred by glutamine to the O2-sensitive alpha -ketoglutarate dehydrogenase.

Hyperoxia-exposed A549 cells had a twofold higher rate of glutamine consumption compared with normoxic cells (Fig. 9). Increased glutamic acid uptake in parallel with increased GSH levels has been reported in hyperoxia-exposed bovine pulmonary endothelial cells (14, 15). GSH and glutamine, as a precursor of GSH, are known to play important roles in the protection against oxidative stress caused by hyperoxia (6, 26). In this previous study (15), a 50% decrease in cellular GSH level was found on exposure of glutamine-deprived cells to hyperoxia. In our study, glutamine supplementation prevented hyperoxic depletion of GSH levels in A549 cells. Therefore, protection of mitochondrial function and increased cell viability in A549 cells could be attributed, at least in part, to the restored GSH levels. Decreased cell GSH contents have been associated with disruption of MMP and hence mitochondrial function during oxidative stress (17, 31, 53).

In this study, enhanced cell survival in hyperoxia with glutamine supplementation could be attributed to the protection of mitochondrial structure and function, increased ATP content, O2 consumption, and protection of alpha -ketoglutarate dehydrogenase and cellular GSH. Studies with human SAECs indicated that glutamine deprivation can cause similar abnormalities in bioenergetics in primary and tumor-derived epithelial cells. Survival of A549 cells in air and the absence of glutamine could also be due to endogenous glutamine synthetase activity (48). Glutamine synthetase is highly susceptible to oxidant stress (2), and through this mechanism, oxidant-stressed cells grown in the absence of glutamine could be further deprived of glutamine. A recent report (27) also suggests that inflammatory necrotic cell death occurring as a result of oxidant stress in pulmonary endothelial cells could be converted to noninflammatory apoptosis by glutamine supplementation through increased ATP levels. Moreover, glutamine supplementation in patients during advanced disease states is a subject of recent medical interest (32), and further studies in this area may be useful.


    ACKNOWLEDGEMENTS

We thank Stephanie Park for preparing the manuscript, Tim Pattison for help in performing the electron microscopy, and Dr. Steve Goodman for glutamine analysis.


    FOOTNOTES

This work was supported by National Heart, Lung, and Blood Institute Grants HL-57144, HL-52732, and HL-56263.

S. Ahmad was a recipient of the Robert J. Suslow Fellowship in Environmental Lung Disease funded by National Jewish Medical and Research Center (Denver, CO).

Address for reprint requests and other correspondence: C. W. White, 1400 Jackson St., Rm. J101, Denver, CO 80206 (E-mail: whitec{at}njc.org).

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

Received 8 May 2000; accepted in final form 20 October 2000.


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