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
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ABSTRACT |
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
-ketoglutarate dehydrogenase was partially protected by its indirect substrate, glutamine, indicating a mechanism of mitochondrial protection.
human; airway; epithelium; adenosine 5'-triphosphate;
-ketoglutarate dehydrogenase; mitochondrial membrane potential
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INTRODUCTION |
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,
-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.
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METHODS |
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
-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).
-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
-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.
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RESULTS |
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.
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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.
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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.
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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.
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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.
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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.
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Biochemical effects of glutamine deprivation.
Because glutamine, the substrate, is known to stabilize the enzyme
-ketoglutarate dehydrogenase, its activity was measured. A complete
loss of
-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
-ketoglutarate dehydrogenase activity compared with the
unsupplemented cells.

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Fig. 8.
Effect of glutamine supplementation on -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.
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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.
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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.
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.
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|
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 |
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
-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
-ketoglutarate dehydrogenase
activity. In addition, we demonstrate for the first time that glutamine
can protect
-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
-ketoglutarate dehydrogenase enzyme and decreased glutamine
utilization. In our model, however, complete loss of
-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
-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
-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
-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
-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
-ketoglutarate dehydrogenase. Our results indicate an increased resistance conferred by glutamine to the O2-sensitive
-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
-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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