Mitogen-activated protein kinase pathway mediates
hyperoxia-induced apoptosis in cultured macrophage cells
Irina
Petrache1,
Mary E.
Choi2,3,
Leo E.
Otterbein4,
Beek Yoke
Chin4,
Lin L.
Mantell5,
Stuart
Horowitz5, and
Augustine M. K.
Choi2,6
6 Section of Pulmonary and
Critical Care Medicine and
3 Section of Nephrology,
Department of Internal Medicine, Yale University School of
Medicine, New Haven 06520;
2 Connecticut Veterans Affairs
HealthCare System, West Haven, Connecticut 06516;
5 Departments of Thoracic
Cardiovascular Surgery and Pediatrics, The CardioPulmonary Research
Institute, Winthrop-University Hospital, State University of New York
at Stony Brook School of Medicine, Mineola, New York 11501; and
1 Division of Pulmonary and
Critical Care Medicine and
4 Department of Environmental
Health Sciences, The Johns Hopkins Medical Institutions, Baltimore,
Maryland 21205
 |
ABSTRACT |
We have previously demonstrated that the lungs
of mice can exhibit increased programmed cell death or apoptosis after
hyperoxic exposure in vivo. In this report, we show that hyperoxic
exposure in vitro can also induce apoptosis in cultured murine
macrophage cells (RAW 264.7) as assessed by DNA-laddering, terminal
deoxynucleotidyltransferase dUTP nick end-labeling, and nucleosomal
assays. To further delineate the signaling pathway of hyperoxia-induced
apoptosis in RAW 264.7 macrophages, we first show that hyperoxia can
activate the mitogen-activated protein kinase (MAPK) pathway, the
extracellular signal-regulated kinases (ERKs) p42/p44, in a
time-dependent manner as assessed by increased phosphorylation of
ERK1/ERK2 by Western blot analyses. Neither the c-Jun
NH2-terminal
kinase/stress-activated protein kinase nor the p38 MAPK was activated
by hyperoxia in these cells. Chemical or genetic inhibition of the ERK
p42/p44 MAPK pathway by PD-98059, a selective inhibitor of MAPK kinase,
and dominant negative mutants of ERK, respectively, attenuated
hyperoxia-induced apoptosis as assessed by DNA laddering and
nucleosomal ELISAs. Taken together, our data suggest that hyperoxia can
induce apoptosis in cultured murine macrophages and that the MAPK
pathway mediates hyperoxia-induced apoptosis.
programmed cell death; oxygen; signal transduction; extracellular
signal-regulated kinase; cell death
 |
INTRODUCTION |
EXPOSURE TO HYPEROXIA, or supraphysiological
concentrations of O2, is
associated at the cellular level with an accumulation of reactive
oxygen species (ROS) such as superoxide, hydroxyl radicals, and
hydrogen peroxide
(H2O2),
with resultant damage of proteins, lipids, and DNA (10, 11). When the
oxidant insult is no longer compensated by the host's antioxidant
defense mechanisms, cell injury and death ensue (6, 8, 22, 26). The
lung in particular is a major target of oxidative injury in a variety of disease states including acute respiratory distress
syndrome, lung fibrosis, and transplant rejection (8, 11,
23). Moreover, the supplemental O2
administered to patients with such lung injury can in itself add to the
oxidative burden already present.
Experimentally, the exposure of animals to hyperoxia causes a form of
acute lung injury similar to that seen in human acute respiratory
distress syndrome, a pathological process characterized by edema and
influx of inflammatory cells into the lung where the cells release
toxic ROS capable of initiating or amplifying lung injury (6, 8, 23).
Recent studies by our laboratories (26, 29) have also suggested that a
major histological feature of hyperoxia-induced lung injury in vivo is
programmed cell death or apoptosis. Apoptosis is an active form of
cellular demise characterized by cell shrinkage, DNA fragmentation, and
nuclear condensation; unlike necrosis, apoptosis is not associated with
architectural distortion of the tissues and does not trigger an
inflammatory response (31). Kazzaz et al. (22) have demonstrated that
hyperoxia induces alveolar epithelial cell necrosis and not apoptosis
in human lung A549 epithelial cells, whereas oxidants such as
H2O2 trigger apoptosis of these cells. We show in this study that hyperoxia can induce apoptosis in vitro depending on the cell type. We
demonstrate in murine macrophages that hyperoxic toxicity might be
responsible for this phenomenon.
The signal transduction pathways of hyperoxia-induced cell necrosis, or
hyperoxia-induced apoptosis in particular, are not yet fully
elucidated. It is known that hyperoxia induces the activation of
transcription factors including activator protein-1 and nuclear factor-
B (23, 24). Both of these transcription factors have been
implicated in the signaling pathways in programmed cell death (16, 25,
27, 35). Moreover, extracellular stimuli such as growth factors and
cellular stresses including DNA damage, hyperosmolarity, oxidative
stress, and mechanical stress are known to activate cascades of protein
kinases that participate in signal transduction; of these,
mitogen-activated protein kinase (MAPK) family members play an
important role in cellular growth and differentiation (33, 36, 37).
Recent data (13, 15, 20) have demonstrated that the MAPK pathway plays
an important role in mediating apoptosis in various in vitro models.
Based on these observations, we hypothesized that hyperoxia activates
the MAPK pathway and that this activation, in turn, is important in
mediating hyperoxia-induced apoptosis. We demonstrate that hyperoxia
activates the extracellular signal-regulated kinase (ERK) p42/p44 MAPK
pathway preferentially and that the ERK MAPK pathway is necessary for
mediating hyperoxia-induced apoptosis in murine RAW 264.7 macrophage cells.
 |
EXPERIMENTAL PROCEDURES |
Cell culture and hyperoxic exposures.
Murine peritoneal macrophages (RAW 264.7 cells), alveolar macrophages
(MHS cells), and L929 fibroblasts were cultured in Dulbecco's modified
Eagle's medium and RPMI medium containing 10% fetal bovine serum
(FBS; GIBCO BRL, Life Technologies, Grand Island, NY), 100 U/ml of
penicillin, and 100 µg/ml of streptomycin. Rat alveolar
macrophages (NR 8383) and human pulmonary epithelial (A549) cells were
maintained in Ham's F-12 medium containing 10% FBS (GIBCO BRL). The
cultures were maintained at 37°C in a humidified atmosphere of 5%
CO2-95% air. The cells were
exposed to hyperoxia (95% O2-5%
CO2) in a tightly sealed modular
chamber (Billup-Rothberg, Del Mar, CA) at 37°C.
Cell transfections. RAW 264.7 cells
were transfected with a dominant negative mutant of the ERK MAPKs (9)
(provided by Dr. Andrew Larner, Food and Drug Administration, Bethesda,
MD) as previously described in our laboratory (22). Two
dominant negative mutant constructs were used: TEYE, where
Thr183 and
Tyr185 residues required for
phosphorylation were replaced with glutamic acid, and TAYE, where the
same residues were replaced with alanine and phenylalanine,
respectively (9). The wild-type (WT) ERK MAPK was obtained from Dr.
Larner (9). RAW 264.7 cells transfected with the neomycin gene or WT
ERK MAPK as previously described (22) were used as controls.
Genomic DNA isolation DNA-laddering
assay. Genomic DNA was isolated from cultured cells
with the Puregene DNA isolation kit (Gentra Systems, Minneapolis, MN).
Briefly, the cells were lysed after medium removal with lysis buffer
followed by a 1-h incubation with RNase A. The cell lysates were
precipitated for proteins and spun at 2,000 g for 15 min. The supernatant was
precipitated with isopropanol for isolation of DNA. After an alcohol
wash, DNA was hydrated and quantified. Equal amounts (20 µg) of DNA were electrophoresed on a 1.5% agarose gel (with incorporated ethidium
bromide) in 1× Tris-acetate buffer. The gel was then photographed
under ultraviolet (UV) luminescence.
Nucleosomal ELISA. Cell lysates were
isolated, and ELISAs were performed according to the
manufacturer's protocol (Calbiochem, San Diego, CA).
Terminal deoxynucleotidyltransferase dUTP nick
end-labeling method in cells. A two-step binding assay
was used to label the 3'-hydroxyl ends of the DNA breaks,
resulting in fluorescent labeling of the apoptotic cells. An APO-BRDU
kit (Phoenix Flow Systems, San Diego, CA) combines terminal
deoxynucleotidyltransferase to catalyze extensions of the
3'-hydroxyl ends and Br-dUTP to substitute for thymidine.
FITC-conjugated anti-5-bromo-2'-deoxyuridine results in the green
labeling. The cells were resuspended in RNase and propidium iodide to
hydrolyze the double-stranded RNA and to stoichiometrically label total
DNA, respectively. Apoptosis was correlated with cell cycle by
generating two independent fluorescent signals on the FACStarplus flow cytometer.
MAPK activation assays. The various
MAPK assays were performed according to the manufacturer's
instructions (New England Biolabs, Beverly, MA). Briefly, cells were
lysed in a buffer (20 mM Tris, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1%
Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM
-glycerophosphate, 1 mM
Na3VO4,
1 µg/ml of leupeptin, and 1 mM phenylmethylsulfonyl fluoride) and
sheared by passage through a 25-gauge needle. Protein concentrations of the cell lysates were determined by Coomassie blue dye-binding assay
(Bio-Rad, Hercules, CA). Total protein (200-µg) samples were
incubated with phospho-specific p42/44 MAPK antibody (1:50) overnight
on a rocker at 4°C. The p42/44 MAPK antibodies detect only the
Tyr204-phosphorylated forms of
ERK1 and ERK2 and thus select for the activated (phosphorylated) MAPK.
The samples were analyzed by 12% SDS-PAGE and electroblotted by
Western blot as per the manufacturer's protocol. After overnight
incubation with the primary antibody at 4°C, the membrane was
incubated for 1 h with a horseradish peroxidase-conjugated anti-rabbit
secondary antibody (1:2,000) at room temperature with gentle rocking.
The proteins were subsequently detected with LumiGLO (New England
Biolabs) and exposed to X-ray film. p38 MAPK activity was measured with
a PhosphoPlus p38 MAPK (Thr180/Tyr182)
antibody kit (New England Biolabs) according to the manufacturer's instructions. A PhosphoPlus stress-activated protein kinase
(SAPK)/c-Jun NH2-terminal kinase
(JNK;
Thr183/Tyr185)
antibody kit (New England Biolabs) was used to measure JNK
phosphorylation or activity. The MAPK kinase (MEK) inhibitor PD-98059
chemical was purchased from New England Biolabs.
O2 exposure of animals.
Eight-week-old pathogen-free male C57BL/6 mice were purchased from
Jackson Laboratories and allowed to acclimate on arrival for 7 days
before experimentation. The animals were fed rodent chow and water ad
libitum. The animals were exposed to hyperoxia (>99%
O2) at a flow rate of 12 l/min
in a 3.70-cubic foot Plexiglas exposure chamber. The mice were supplied
with rodent chow and water ad libitum during the exposure. At the start
of the exposure, the chamber was humidified for 10-15 min. These
experiments were carried out according to the animal protocol approved
by the Animal Care and Use Committee.
Lung tissue preparation. The lungs
were fixed by perfusion with 10% Formalin at 20 cmH2O pressure and embedded in
paraffin. Lung sections of 4-5 µm were mounted onto slides
pretreated with 3-aminopropylethoxysilane (Digene Diagnostics,
Beltsville, MD). The slides were baked for 30 min at 60°C and
washed twice in fresh xylene for 5 min each to remove the paraffin. The
slides were rehydrated though a series of graded alcohols and then
washed in distilled water for 3 min each.
Terminal deoxynucleotidyltransferase dUTP nick
end-labeling assay in tissue sections and
photomicrography. The terminal
deoxynucleotidyltransferase dUTP nick end-labeling (TUNEL) method was
used for the apoptosis assay of lung tissue sections as previously
described (26).
TUNEL reagents, including a rhodamine-conjugated anti-digoxigenin Fab
fragment, were obtained from Boehringer Mannheim (Indianapolis, IN).
The nonspecific signals or autofluorescence was excluded by subtracting
the signals generated from the TUNEL assay with signals generated by
all reagents except the terminal deoxynucleotidyltransferase in the
assay. Tissue sections were counterstained with 2 µg/ml of
4',6-diamidine-2-phenylindole dihydrochloride (Boehringer
Mannheim) for 10 min at room temperature. Photomicrographs were
recorded on 35-mm film with a Nikon Optiphot microscope and UFX camera system (Nikon, Melville, NY) and transferred onto a KodakPhotoCD. The
images were digitally adjusted for contrast with Adobe PhotoShop 3.0 (Adobe Systems, Mountain View, CA). The images were captured with a
charge-coupled device video camera. Uniform camera control settings
were used for image capture, and image threshholding was identical for
all images. The captured images were analyzed with the Image 1 system
(Universal Imaging, West Chester, PA) running on a personal computer.
At least 25 fields were analyzed from at least two animals at each time point.
Statistical analysis. Data are
expressed as means ± SE. Differences in measured variables between
the experimental and control groups were assessed with Student's
t-test. Statistical calculations were
performed on a Macintosh personal computer with the Statview II
statistical package (Abacus Concepts, Berkeley, CA). Significant difference was accepted at P < 0.05.
 |
RESULTS |
Hyperoxia induces apoptosis in macrophages in
vitro. To determine whether hyperoxia induces cell
death via programmed cell death (apoptosis), genomic DNA was isolated
from RAW 264.7 cells after hyperoxic (>95%
O2) treatment and analyzed for
DNA laddering. Figure
1A shows
marked DNA laddering at 24 h of hyperoxic treatment (lane 2) in contrast to cells
exposed to normoxia (lane 1). Agents such as endotoxin, ATP, or
H2O2,
which have been shown to induce apoptosis in other cell types, did not
induce apoptosis in RAW 264.7 cells (Fig.
1A). We also observed that
hyperoxic exposure induced apoptosis in another macrophage cell line,
MHS cells, as assessed by DNA-laddering assay (Fig.
1B) and nucleosomal assay (control,
0.05 nucleosomal U/mg protein; hyperoxia for 16 h, 0.39 nucleosomal
U/mg protein). To further complement our DNA-laddering assays and to
better quantify the effect of hyperoxia-induced apoptosis, identical
experiments were performed with the TUNEL and nucleosomal assays. Both
assays further demonstrate evidence of apoptosis in RAW 264.7 cells
after hyperoxia (Fig. 2).


View larger version (121K):
[in this window]
[in a new window]
|
Fig. 1.
Hyperoxia induces apoptosis in murine macrophage cells.
A: genomic DNA was isolated from RAW
264.7 cells exposed to hyperoxia for 24 h and fractionated with 1.5%
agarose gel electrophoresis as described in
EXPERIMENTAL PROCEDURES.
Lane 1, control normoxia;
lane 2, 95%
O2 for 24 h; lane
3, lipopolysaccharide (LPS; 10 µg/ml) for 24 h;
lane 4, 5 mM ATP for 6 h;
lane 5, 600 µM
H2O2
for 6 h. B: genomic DNA was isolated
from MHS cells exposed to hyperoxia and fractionated with 1.5% agarose
gel electrophoresis as described in EXPERIMENTAL
PROCEDURES. Lane 1,
control normoxia; lane 2, 95%
O2 for 4 h; lane
3, 95% O2 for 8 h; lane 4, 95%
O2 for 16 h.
|
|

View larger version (8K):
[in this window]
[in a new window]
|
Fig. 2.
A: terminal
deoxynucleotidyltransferase dUTP nick end-labeling (TUNEL) analysis of
RAW 264.7 cells exposed to hyperoxia. Cells were analyzed for positive
TUNEL staining as described in EXPERIMENTAL
PROCEDURES. 1, Control normoxia; 2, 95%
O2 for 24 h. Data are means ± SE of 3 independent experiments.
* P < 0.001 compared with
normoxic control cells. B: nucleosomal
ELISA of RAW 264.7 cells exposed to hyperoxia. Cells lysates were
isolated after 24 h of hyperoxic exposure, and ELISA was performed as
described in EXPERIMENTAL PROCEDURES.
1, Control normoxia; 2, 95% O2
for 24 h. Data are means ± SE of 3 independent
experiments. * P < 0.006 compared with normoxic control cells.
|
|
Hyperoxia induces apoptosis in mouse
lungs. Our laboratories (26, 29) have previously
reported that hyperoxia increases the total apoptotic index in the
lungs of both rats and mice. To examine whether lung alveolar
macrophages contributed to the increased total apoptotic index in the
lungs after hyperoxic exposure in vivo, lung sections were obtained
from mice exposed to hyperoxia and analyzed for apoptotic signals by an
in situ TUNEL assay, which labels the 3'-OH ends of DNA cut by
endonucleases that are activated during apoptosis (26). A marked
increase in TUNEL staining was observed in cells resembling macrophages
in the alveoli of lungs from mice exposed to hyperoxia compared with
mice exposed to normoxia alone (Fig. 3).

View larger version (38K):
[in this window]
[in a new window]
|
Fig. 3.
Apoptosis of alveolar macrophages and lung parenchyma. Normoxic
(A) and hyperoxic
(B) mouse lungs were isolated after
72 h of 100% O2 exposure and
fixed and embedded as described in EXPERIMENTAL
PROCEDURES. Lung tissue sections were processed for
TUNEL assay to identify apoptotic cells (TUNEL stain) and dual labeled
with 4',6-diamidine-2-phenylindole dihydrochloride (DAPI) to
visualize all nuclei in the field. Differential interference contrast
(DIC) image was digitally enhanced to emphasize surface topology.
Arrows, presumptive alveolar macrophage in lumen of alveolus;
arrowheads, epithelial cells of alveolar wall. Bar, 15 µm.
|
|
Hyperoxia activates the ERK p42/p44 MAPK signal
transduction pathway in murine macrophages.
Extracellular stimuli such as growth factors, cellular stresses
including DNA damage, hyperosmolarity, oxidative stress, and mechanical
stress are known to activate cascades of protein kinases that
participate in signal transduction. The MAPK pathway represents one
major signaling pathway of these cellular stressors (33, 36). We
hypothesized that hyperoxia activates the MAPK pathway in cultured
macrophages. To determine the ERK p42/p44 MAPK activity in response to
hyperoxia, we selectively immunoblotted cellular proteins isolated from
RAW 264.7 cells after hyperoxic treatment with a monoclonal
phospho-specific antibody to p42/p44 MAPK
(Thr202 and
Tyr204). Increased ERK p42/p44
MAPK activity was observed as early as 30 min, which returns to
baseline at 8-24 h of hyperoxic exposure (Fig.
4). In contrast, exposure of RAW 264.7 cells to hyperoxia did not result in activation of the JNK or p38 MAPK
pathway (Fig. 5).

View larger version (17K):
[in this window]
[in a new window]
|
Fig. 4.
Kinetics of extracellular signal-regulated kinase [ERK; p42/p44
mitogen-activated protein kinase (MAPK)] activity of RAW 264.7 cells exposed to hyperoxia. Lane 1,
control normoxia; lane 2, 30 min of
hyperoxia; lane 3, 1 h of hyperoxia;
lane 4, 2 h of hyperoxia;
lane 5, 4 h of hyperoxia;
lane 6, 8 h of hyperoxia;
lane 7, 24 h of hyperoxia. Blot is
representative of 4 independent experiments. Nos. on
right, molecular mass.
|
|

View larger version (62K):
[in this window]
[in a new window]
|
Fig. 5.
A: kinetics of stress-activated
protein kinase (SAPK)/c-Jun
NH2-terminal kinase (JNK) activity
of RAW 264.7 cells exposed to hyperoxia. Lane
1, control normoxia; lane
2, 30 min of hyperoxia; lane
3, 1 h of hyperoxia; lane
4, 2 h of hyperoxia; lane
5, 4 h of hyperoxia; lane
6, 8 h of hyperoxia; lane
7, 30 min of ultraviolet irradiation as positive
control. Blot is representative of 3 independent experiments.
B: kinetics of p38 MAPK activity of
RAW 264.7 cells exposed to hyperoxia. Lane
1, control normoxia; lane
2, 15 min of hyperoxia; lane
3, 30 min of hyperoxia; lane
4, 4 h of hyperoxia; lane
5, 4 h of 1 µg/ml of LPS as positive control. Blot is
representative of 3 independent experiments.
|
|
Requirement of ERK p42/p44 MAPK signaling pathway for
hyperoxia-induced apoptosis in macrophages. Based on
our observations that hyperoxia induces apoptosis and activates the ERK
p42/p44 MAPK pathway in RAW 264.7 cells, we hypothesized that the ERK p42/p44 MAPK pathway may mediate hyperoxia-induced apoptosis in RAW
264.7 cells. To test this hypothesis, our strategy was to inhibit ERK
MAPK genetically and chemically and then examine whether inhibition of
the ERK MAPK pathway modulates hyperoxia-induced apoptosis. First, RAW
264.7 cells were transfected with dominant negative mutants of ERK MAPK
(9) and then were exposed to hyperoxia. DNA laddering was detected in
the WT ERK MAPK RAW 264.7 cells after hyperoxic treatment but not in
the TEYE dominant negative mutant cells
(Thr183 and
Tyr185 residues required for
phosphorylation were replaced with glutamic acid), suggesting that the
ERK pathway plays an important role in mediating hyperoxia-induced
apoptosis (Fig. 6). Identical results were
obtained with the TAYE dominant negative mutant cells
(Thr183 and
Tyr185 residues were replaced with
alanine and phenylalanine, respectively; data not shown). To further
confirm the role of the ERK p42/p44 MAPK pathway in hyperoxia-induced
apoptosis, we treated RAW 264.7 cells with PD-98059, a selective
inhibitor of MEK (located upstream from ERK) before the hyperoxic
exposure and then examined the cells for DNA laddering. As seen in Fig.
7, RAW 264.7 cells when exposed to
hyperoxia alone (lane 3) exhibited
marked DNA laddering, whereas cells exposed to hyperoxia in the
presence of PD-98059 exhibited attenuation of DNA laddering
(lane 4). To better quantify the
effect of inhibition of ERK MAPK on hyperoxia-induced apoptosis, we
performed nucleosomal ELISA assays in TEYE dominant negative mutant
cells in the presence of hyperoxia. As shown in Fig.
8, a marked increase in nucleosomal units
(index of apoptosis) were observed in RAW 264.7 neo control cells after
hyperoxia (lane 2;
P < 0.03 compared with normoxic
control cells in lane 1). In contrast, no significant increase in nucleosomal units was observed in
TEYE dominant negative mutant cells after hyperoxia (Fig. 8, lane 4) compared with that after
normoxia (Fig. 8, lane 3;
P = not significant). Statistical
analysis of nucleosomal units (index of apoptosis) between RAW 264.7 neo control cells and TEYE dominant negative mutant cells after
hyperoxia demonstrated significant differences between these two groups
(P < 0.06). We observed similar attenuation of hyperoxia-induced nucleosomal units (index of apoptosis) in RAW 264.7 cells exposed to hyperoxia in the presence of the chemical
inhibitor PD-98059 (data not shown).

View larger version (58K):
[in this window]
[in a new window]
|
Fig. 6.
Effect of dominant negative mutants of ERK on hyperoxia-induced
apoptosis. Genomic DNA was isolated from cells exposed to hyperoxia for
24 h and then fractionated with 1.5% agarose gel electrophoresis as
described in EXPERIMENTAL PROCEDURES.
Left: lane
1, wild-type (WT) ERK MAPK RAW 264.7 cells in normoxia;
lane 2, WT ERK MAPK RAW 264.7 cells in
hyperoxia. Right:
lane 1, TEYE dominant negative mutant
RAW 264.7 cells in normoxia; lane 2,
TEYE dominant negative mutant RAW 264.7 cells in hyperoxia.
|
|

View larger version (105K):
[in this window]
[in a new window]
|
Fig. 7.
Effect of MAPK kinase (MEK) inhibitor on hyperoxia-induced apoptosis.
Genomic DNA was isolated from cells exposed to hyperoxia for 24 h in
absence and presence of PD-98059 (10 µM), a specific MEK inhibitor,
and then fractionated with 1.5% agarose gel electrophoresis as
described in EXPERIMENTAL PROCEDURES.
Lane 1, normoxia;
lane 2, PD-98059 alone;
lane 3, hyperoxia;
lane 4, PD-98059 plus hyperoxia.
|
|

View larger version (10K):
[in this window]
[in a new window]
|
Fig. 8.
Effect of dominant negative mutants of ERK on hyperoxia-induced
apoptosis by nucleosomal ELISA. Cells lysates were obtained at 24 h of
hyperoxic exposure, and nucleosomal ELISA was performed as described in
EXPERIMENTAL PROCEDURES. 1, RAW 264.7 neo control cells in normoxia; 2, RAW 264.7 neo control cells after 24 h in hyperoxia; 3, TEYE dominant negative mutant RAW 264.7 cells in
normoxia; 4, TEYE dominant negative mutant RAW 264.7 cells after 24 h
of hyperoxia. Data are means ± SE of 5 independent experiments.
* P < 0.03 compared with
control cell in normoxia; ** P < 0.006 compared with control cells after 24 h of hyperoxia.
|
|
 |
DISCUSSION |
It is well established that direct exposure of cultured cells to ROS
such as
H2O2,
superoxide-generating agents, and glutathione depletors induce cell
death via induction of programmed cell death or apoptosis in many model
systems. Although hyperoxia is a form of oxidant stress, being
associated with an accumulation of ROS and triggering common
antioxidant responses, recent data (22) suggest that hyperoxia to a
large extent induces cell death via cell necrosis rather than via
programmed cell death. For example, lung epithelial cells (A549) die by
cell necrosis when exposed to hyperoxia (22). Human bronchial
epithelial cells, rat alveolar macrophages (NR 8383 cells), and murine
fibroblasts (L929 cells) do not exhibit genomic DNA fragmentation
(laddering) when exposed to hyperoxia (data not shown). We show in this
study that murine macrophages (RAW 264.7 and MHS cells), in contrast to
rat macrophages (NR 8383 cells), have the capacity to undergo
programmed cell death or apoptosis after hyperoxic exposure, suggesting
species specificity of the hyperoxic insult. Moreover, the complexity of programmed cell death is further highlighted by recent observations (1, 19, 21, 34) demonstrating cell-type specificity in that hyperoxia
can cause apoptosis in cells such as human small-airway epithelial
cells, retinal capillary endothelial cells, cerebral endothelial cells,
and pheochromocytoma cells.
Given the fact that in vivo exposure to hyperoxia is associated with
significant apoptosis in the lungs of mice and rats (26, 29), it is
possible that the alveolar macrophage is the predominant cell
population that contributes to this effect (Fig. 3), although other
cell types may also be contributing to the increased apoptotic index of
mouse lungs exposed to hyperoxia. Rigorous studies are needed in the
future to accurately determine the type of cells undergoing apoptosis
both quantitatively and qualitatively.
During the apoptotic process, macrophages assume the role of
scavengers, "silently" engulfing adjacent apoptotic cells and possibly contributing to the resolution of acute pulmonary inflammation (7, 31). On the other hand, macrophages possess the ability to undergo
programmed cell death or apoptosis themselves as observed in this study
in response to hyperoxia and to other cellular stressors such as
asbestos, bleomycin, nitric oxide and peroxynitrite, endotoxin, and
interferon-
(2, 3, 14, 17, 30). The functional significance of
macrophage apoptosis to these stimuli, and hyperoxic injury in
particular, is not well established. Based on the known observations
that macrophages can undergo apoptosis to many cellular stimuli that
are potent activators of macrophages (3, 14, 17, 30), one can speculate
that macrophage cell death via apoptosis may represent a mechanism by
which the host attempts to minimize ongoing inflammatory process during
macrophage activation.
To date, there have been three main branches of the MAPK pathway
described, although there is considerable cross talk among them. The
ERK p42/p44 MAPK pathway is activated by cell growth and mitogenic
signals, DNA damage, or oxidative stress (33, 37, 38). SAPK/JNK and p38
protein kinase represent the two other branches of the MAPK pathway,
being activated by cellular stimuli such as cytokines, oxidant
stresses, and osmotic shock (13, 15, 28, 36). These three branches, ERK
p42/p44, JNK, and p38, of the MAPK pathway have been implicated in the
regulation of the signaling pathways leading to apoptosis depending on
the various models. In our model, with early activation of the ERK p42/p44 MAPK pathway relative to the unaffected JNK and p38 MAPK pathways, it appears that the ERK p42/p44 MAPK pathway mediates hyperoxia-induced apoptosis. Interruption of the ERK p42/p44 branch of
the MAPK signaling pathway, either genetically or chemically, blocked
hyperoxia-induced apoptosis. The importance of the ERK p42/p44 MAPK
pathway in mediating apoptosis has also been recently observed in other
models of oxidant-induced apoptosis including asbestosis, UV
irradiation, and tumor necrosis factor-
(12, 17, 18). However, the
complexity of the MAPK pathway in mediating apoptosis is further
highlighted by studies demonstrating the importance of either the JNK
or p38 branch of the MAPK pathway in regulating the signaling pathways
in the apoptotic process depending on cell type and cell stimulus. For
example, UV irradiation-induced apoptosis in small cell lung cancer
cells appears to be mediated by the JNK MAPK pathway (4, 5), whereas
aspirin-induced apoptosis in fibroblasts is mediated by the p38 MAPK
pathway (32). Additionally, both the JNK and p38 MAPK pathways have
been shown to mediate nerve growth factor removal-induced apoptosis in
PC-12 pheochromocytoma cells (38).
In summary, our study demonstrates that hyperoxia causes apoptosis in
cultured murine macrophages and that among the three branches of the
MAPK pathway, the ERK p42/p44 pathway appears to play an important role
in mediating hyperoxia-induced apoptosis. Murine macrophage cell lines
will serve as a useful cell culture model to investigate the signaling
pathway(s) upstream from the ERK p42/p44 MAPK pathway mediating
hyperoxia-induced apoptosis and to further understand the functional
significance of hyperoxia-induced apoptosis.
 |
ACKNOWLEDGEMENTS |
A. M. K. Choi was supported by National Heart, Lung, and Blood
Institute Grants R01-HL-55330 and R01-HL-60234; National Institute of
Allergy and Infectious Diseases Grant R01-AI-42365; and an American
Heart Association Established Investigator Award. S. Horowitz was
supported in part by Basic Research Grant 1-FY96-0752 from the March of
Dimes Birth Defects Foundation and grants from the National Institutes
of Health and Winthrop-University Hospital. L. L. Mantell was supported
in part by grants from the American Lung Association and the Stony
Wold-Herbert Fund. M. E. Choi was supported by National Institute of
Diabetes and Digestive and Kidney Diseases Grant DK-01298-09 and by a
Veterans Affairs Career Development Award.
 |
FOOTNOTES |
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: A. M. K. Choi,
Section of Pulmonary and Critical Care Medicine, Yale Univ. School of
Medicine, 333 Cedar St., LCI 105, New Haven, CT 06520 (E-mail:
augustine.choi{at}yale.edu).
Received 17 September 1998; accepted in final form 7 April 1999.
 |
REFERENCES |
1.
Alon, T.,
I. Hemo,
A. Itin,
J. Pe'er,
J. Stone,
and
E. Keshet.
Vascular endothelial growth factor acts as a survival factor for newly formed retinal vessels and has implications for retinopathy of prematurity.
Nat. Med.
1:
1024-1028,
1995[Medline].
2.
Bingisser, R.,
C. Stey,
M. Weller,
P. Groacurth,
E. Russi,
and
K. Frei.
Apoptosis in human alveolar macrophages is induced by endotoxin and is modulated by cytokines.
Am. J. Respir. Cell Mol. Biol.
15:
64-70,
1996[Abstract].
3.
Brune, B.,
C. Golkel,
and
A. von Knethen.
Cytokine and low-level nitric oxide prestimulation block p53 accumulation and apoptosis of RAW 264.7 macrophages.
Biochem. Biophys. Res. Commun.
229:
396-401,
1996[Medline].
4.
Butterfield, L.,
B. Storey,
L. Maas,
and
L. E. Heasley.
c-Jun NH2-terminal kinase regulation of the apoptotic response of small cell lung cancer cells to ultraviolet radiation.
J. Biol. Chem.
272:
10110-10116,
1997[Abstract/Free Full Text].
5.
Chen, Y. R.,
X. P. Wang,
D. Templeton,
R. J. Davis,
and
T. H. Tan.
The role of c-Jun N-terminal kinase (JNK) in apoptosis induced by ultraviolet C and gamma radiaton: duration of JNK activation may determine cell death and proliferation.
J. Biol. Chem.
271:
31929-31936,
1996[Abstract/Free Full Text].
6.
Clerch, L. B.,
and
D. Massaro.
Tolerance of rats to hyperoxia: lung antioxidant enzyme gene expression.
J. Clin. Invest.
91:
499-508,
1993[Medline].
7.
Cox, G.,
J. Crossley,
and
Z. Xing.
Macrophage engulfment of apoptotic neutrophils contributes to the resolution of acute pulmonary inflammation in vivo.
Am. J. Respir. Cell Mol. Biol.
12:
232-237,
1995[Abstract].
8.
Crapo, J. D.,
B. E. Barry,
H. A. Foscue,
and
J. Shelburne.
Structural and biochemical changes in rat lungs occurring during exposure to lethal and adaptive doses of oxygen.
Am. Rev. Respir. Dis.
122:
123-143,
1980[Medline].
9.
David, M.,
E. Petricoin,
C. Benjamin,
R. Pine,
M. J. Weber,
and
A. C. Larner.
Requirement for MAP kinase (ERK2) activity in interferon alpha- and interferon beta-stimulated gene expression through STAT proteins.
Science
269:
1721-1723,
1995[Medline].
10.
Freeman, B. A.,
and
J. D. Crapo.
Hyperoxia increases oxygen radical production in rat lungs and lung mitochondria.
J. Biol. Chem.
256:
10986-10992,
1981[Free Full Text].
11.
Fridovich, I.
The biology of oxygen radicals.
Science
201:
875-880,
1978[Medline].
12.
Gardner, A. M.,
and
G. L. Johnson.
Fibroblast growth factor-2 suppression of tumor necrosis factor alpha-mediated apoptosis requires Ras and the activation of mitogen-activated protein kinase.
J. Biol. Chem.
271:
14560-14566,
1996[Abstract/Free Full Text].
13.
Goillot, E. J.,
J. Raingeaud,
A. Ranger,
R. I. Tepper,
R. J. Davis,
E. Harlow,
and
I. Sanchez.
Mitogen-activated protein kinase-mediated Fas apoptotic signaling pathway.
Proc. Natl. Acad. Sci. USA
94:
3302-3307,
1997[Abstract/Free Full Text].
14.
Hamilton, R. F.,
L. Li,
T. B. Felder,
and
A. Holian.
Bleomycin induces apoptosis in human alveolar macrophages.
Am. J. Physiol.
269 (Lung Cell. Mol. Physiol. 13):
L318-L325,
1995[Abstract/Free Full Text].
15.
Ichijo, H.,
E. Nishida,
K. Irie,
P. ten Dijke,
M. Saitoh,
T. Moriguchi,
M. Takagi,
K. Matsumoto,
K. Miyazono,
and
Y. Gotoh.
Induction of apoptosis by ASK1, a mammalian MAPKKK that activates SAPK/JNK and p38 signaling pathways.
Science
275:
90-94,
1997[Abstract/Free Full Text].
16.
Ishikawa, Y.,
T. Yokoo,
and
M. Kitamura.
c-Jun/AP-1, but not NF-kappa B, is a mediator for oxidant-initiated apoptosis in glomerular mesangial cells.
Biochem. Biophys. Res. Commun.
240:
496-501,
1997[Medline].
17.
Jimenez, L. A.,
C. Zanella,
H. Fung,
Y. M. Janssen,
P. Vacek,
C. Charland,
J. Goldberg,
and
B. T. Mossman.
Role of extracellular signal-regulated protein kinases in apoptosis by asbestos and H2O2.
Am. J. Physiol.
273 (Lung Cell. Mol. Physiol. 17):
L1029-L1035,
1997[Abstract/Free Full Text].
18.
Johnson, N. L.,
A. M. Gardner,
K. M. Diener,
C. A. Lange-Carter,
J. Gleavy,
M. B. Jarpe,
A. Minden,
M. Karin,
L. I. Zon,
and
G. L. Johnson.
Signal transduction pathways regulated by mitogen-activated/extracellular response kinase kinase kinase induce cell death.
J. Biol. Chem.
271:
3229-3237,
1996[Abstract/Free Full Text].
19.
Jyonouchi, H.,
S. Sun,
T. Abiru,
S. Chareancholvanich,
and
D. H. Ingbar.
The effects of hyperoxic injury and antioxidant vitamins on death and proliferation of human small airway epithelial cells.
Am. J. Respir. Cell Mol. Biol.
19:
426-436,
1998[Abstract/Free Full Text].
20.
Karin, M.,
Z. Liu,
and
E. Zandi.
AP-1 function and regulation.
Curr. Opin. Cell Biol.
9:
240-246,
1997[Medline].
21.
Katoh, S.,
Y. Mitsui,
K. Kitani,
and
T. Suzuki.
The rescuing effect of nerve growth factor is the result of up-regulation of bcl-2 in hyperoxia-induced apoptosis of a subclone of pheochromocytoma cells, PC12H.
Neurosci. Lett.
232:
71-74,
1997[Medline].
22.
Kazzaz, J. A.,
J. Xu,
T. A. Palaia,
L. Mantell,
A. M. Fein,
and
S. Horowitz.
Cellular oxygen toxicity. Oxidant injury without apoptosis.
J. Biol. Chem.
271:
15182-15186,
1996[Abstract/Free Full Text].
23.
Lee, P. J.,
J. Alam,
S. L. Sylvester,
N. Inamdar,
L. Otterbein,
and
A. M. K. Choi.
Regulation of heme oxygenase-1 expression in vivo and in vitro in hyperoxic lung injury.
Am. J. Respir. Cell Mol. Biol.
14:
556-569,
1996[Abstract].
24.
Li, Y. C.,
W. X. Zhang,
L. L. Mantell,
J. A. Kazzaz,
A. M. Fein,
and
S. Horowitz.
Nuclear factor-kappa-b is activated by hyperoxia but does not protect from cell death.
J. Biol. Chem.
272:
20646-20649,
1997[Abstract/Free Full Text].
25.
Liebermann, D. A.,
B. Gregory,
and
B. Hoffman.
AP-1 (Fos/Jun) transcription factors in hematopoietic differentiation and apoptosis.
Int. J. Oncol.
12:
685-700,
1998[Medline].
26.
Mantell, L. L.,
J. A. Kazzaz,
J. Xu,
T. A. Palaia,
B. Piedboeuf,
S. Hall,
G. C. Rhodes,
G. Niu,
A. F. Fein,
and
S. Horowitz.
Unscheduled apoptosis during inflammatory lung injury.
Cell Death Differ.
4:
600-607,
1997.
27.
Mayo, M. W.,
C. Y. Wang,
P. C. Cogswell,
K. S. Rogers-Graham,
S. W. Lowe,
C. J. Der,
and
A. S. Baldwin, Jr.
Requirement of NF-kappaB activation to suppress p53-independent apoptosis induced by oncogenic Ras.
Science
278:
1812-1815,
1997[Abstract/Free Full Text].
28.
Moriguchi, T.,
F. Toyoshima,
N. Masuyama,
H. Hanafusa,
Y. Gotoh,
and
E. Nishida.
A novel SAPK/JNK kinase, MKK7, stimulated by TNF alpha and cellular stresses.
EMBO J.
16:
7045-7053,
1997[Abstract/Free Full Text].
29.
Otterbein, L.,
B. Y. Chin,
L. L. Mantell,
L. Stansberry,
S. Horowitz,
and
A. M. K. Choi.
Pulmonary apoptosis in aged and oxygen-tolerant rats exposed to hyperoxia.
Am. J. Physiol.
275 (Lung Cell. Mol. Physiol. 19):
L14-L20,
1998[Abstract/Free Full Text].
30.
Sandoval, M.,
X. J. Zhang,
X. Liu,
E. E. Mannick,
D. A. Clark,
and
M. J. Miller.
Peroxynitrite-induced apoptosis in T84 and RAW 264.7 cells: attenuation by L-ascorbic acid.
Free Radic. Biol. Med.
22:
489-495,
1997[Medline].
31.
Savill, J.
Apoptosis in resolution of inflammation.
J. Leukoc. Biol.
61:
375-380,
1997[Abstract].
32.
Schwenger, P.,
P. Bellosta,
I. Vietor,
C. Basilico,
E. Y. Skolnik,
and
J. Vilcek.
Sodium salicylate induces apoptosis via p38 mitogen-activated protein kinase but inhibits tumor necrosis factor-induced c-Jun N-terminal kinase/stress-activated protein kinase activation.
Proc. Natl. Acad. Sci. USA
94:
2869-2873,
1997[Abstract/Free Full Text].
33.
Seger, R.,
and
E. G. Krebs.
The MAPK signaling cascade.
FASEB J.
9:
726-735,
1995[Abstract/Free Full Text].
34.
Shaikh, A. Y.,
J. Xu,
Y. J. Wu,
L. He,
and
C. Y. Hsu.
Melatonin protects bovine cerebral endothelial cells from hyperoxia-induced DNA damage and death.
Neurosci. Lett.
229:
193-197,
1997[Medline].
35.
Sonenshein, G. E.
Rel/NF-kappa B transcription factors and the control of apoptosis.
Semin. Cancer Biol.
8:
113-119,
1997[Medline].
36.
Su, B.,
and
M. Karin.
Mitogen-activated protein kinase cascades and regulation of gene expression.
Curr. Opin. Immunol.
8:
402-411,
1996[Medline].
37.
Woodgett, J. R.,
J. Avruch,
and
J. Kyriakis.
The stress activated protein kinase pathway.
Cancer Surv.
27:
127-138,
1996[Medline].
38.
Xia, Z.,
M. Dickens,
J. Raingeaud,
R. J. Davis,
and
M. E. Greenberg.
Opposing effects of ERK and JNK-p38 MAP kinases on apoptosis.
Science
270:
1326-1331,
1995[Abstract].
Am J Physiol Lung Cell Mol Physiol 277(3):L589-L595
0002-9513/99 $5.00
Copyright © 1999 the American Physiological Society