First Department of Medicine, Tokyo Women's Medical University, Shinjuku-ku, Tokyo 162-8666, Japan
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Alveolar macrophages (AMs) may play a critical role
in cigarette smoke (CS)-related pulmonary diseases. This study was
designed to determine whether CS induces apoptosis of AMs. In
in vitro studies, mouse, rat, and human AMs and human blood
monocyte-derived macrophages cultured with aqueous whole CS extracts
underwent apoptosis that was detected by light and electron
microscopy and terminal deoxynucleotidyltransferase-mediated dUTP nick
end labeling. The gas phase of CSE did not cause apoptosis. The
CS-induced apoptosis was associated with increased oxidative
stress, Bax protein accumulation, mitochondrial dysfunction, and
mitochondrial cytochrome c release but was independent of
p53, Fas, and caspase activation. This apoptosis was inhibited
by antioxidants such as glutathione, ascorbic acid, and -tocopherol.
In in vivo studies where rats were exposed to the smoke from 10 cigarettes over 5 h in an exposure chamber, ~3% of AMs obtained
by bronchoalveolar lavage after 24 h showed apoptosis.
These results suggest that acute CS exposure is capable of inducing
apoptosis of AMs.
oxidative stress; Bax; mitochondria; p53; caspase
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
NUMEROUS STUDIES HAVE SHOWN that cigarette smoke (CS) directly modulates the metabolism and function of alveolar macrophages (AMs). For example, changes have been reported in morphology and ultrastructure (24), glucose oxidation rate (11), RNA and protein synthesis (13), phagocytic activity (7, 8), various enzymatic activities (27), protease release (26, 32), and cytokine production (37) of AMs.
CS is also known to be toxic to AMs. In in vitro studies, acute CS exposure has been shown to kill AMs (31, 35), although the type of cell death, i.e., necrosis or apoptosis, remains uncertain. The mechanism of CS toxicity to AMs may involve oxidative stress, an important mediator of cell death via both necrosis and apoptosis. Indeed, CS contains large quantities of reactive oxygen species (ROS) and ROS inducers (25), some of which can enter cells and may even reach the nucleus to cause oxidative DNA damage and cell death (30). The toxicity of CS to AMs may be relevant to a clinical observation that cigarette smokers often suffer from nonspecific respiratory infections that may enhance mortality in this population (7, 8).
Despite the acute toxicity of CS, AMs are known to be increased in smokers compared with nonsmokers (3). This is thought to be due to enhanced recruitment of blood monocytes into the lung. However, it is currently unknown whether the rate of AM death is different in smokers and nonsmokers. Smokers' AMs may become resistant to chronic and repetitive exposure to CS. Alternatively, AMs may be protected against the toxicity of CS in the microenvironment of the lung that contains large amounts of antioxidants. Further studies are needed to understand how CS induces AM death.
In the present study, we investigated whether acute exposure to CS induces apoptosis of AMs. To determine the direct effect of CS on AMs, we first conducted in vitro experiments in which AMs were exposed to aqueous CS extract (CSE). Next, we performed in vivo experiments in which rodents were acutely exposed to CS inhalation. From these experiments, we demonstrate that acute CS exposure induces apoptosis of AMs.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Experimental animals.
Ten-week-old male Sprague-Dawley rats and six-week-old male C57BL/6
wild-type, Fas-deficient lpr/lpr, and Fas
ligand-defective gld/gld mice were purchased from
SLC (Shizuoka, Japan). C57BL/6 p53-deficient [(/
)]
mice were obtained from Clea Japan (Tokyo, Japan). Animals were handled
in accordance with Institutional Animal Care and Use Committee
protocols approved by the animal facility of Tokyo Women's Medical
University (Tokyo, Japan). They were maintained under standard
conditions, with a dark period from 8 PM to 8 AM, and water and food
were provided ad libitum.
Preparations of macrophages. AMs were obtained by bronchoalveolar lavage (BAL) from rats, mice, and human healthy nonsmoking volunteers as previously described (5). Human blood monocyte-derived macrophages were prepared as previously described (19). Purity of the macrophage populations on May-Grünwald-Giemsa-stained cytospin slides was > 95%, and the cell viability as determined by trypan blue dye exclusion was > 98%.
Preparation of CSE solution. CSE was prepared as previously described (2). Commercial plain-ended cigarettes (Peace, Japan Tobacco, Tokyo, Japan) yielding 24 mg of tar and 2.4 mg of nicotine under a standard smoking regimen were used in this study. Mainstream smoke was generated from one cigarette by drawing consecutive puffs into a 20-ml plastic syringe, with a stopcock connected through one port to a glass vessel containing 10 ml of PBS (pH 7.4). A 20-ml puff drawn in 1 s was obtained at 10-s intervals, and each puff was held for 3 s and bubbled through PBS in 5 s. One cigarette yielded an average of 45 puffs by this procedure. To prepare gas-phase CSE, the smoke was drawn into the syringe through a 0.22-µm pore size filter (Milex-HA, Nihon Millipore, Tokyo, Japan) rated to remove the tar phase of CS. pH of the resultant solution was 7.4 in both whole and gas-phase CSE solutions. The CSE solutions were always prepared by the same person (K. Aoshiba) using exactly the same method and were used within 3 min after preparation.
In vitro CS exposure of macrophages. For in vitro CS exposure, isolated macrophages (1 × 106 cells/ml) were suspended in serum-free DMEM containing 100 U/ml of penicillin and 100 µg/ml of streptomycin with and without 0.5-10 vol% of freshly prepared CSE solutions. The cells were then cultured in either 96-well culture plates (Beckon Dickinson, Lincoln Park, NJ) or 1.5-ml graduated microcentrifuge tubes (Assist, Tokyo, Japan) at 37°C in a 5% CO2 humidified atmosphere.
In vivo CS exposure. For in vivo CS exposure, three rats were placed in a plastic cage (27 × 27 × 18 cm) with a narrow orifice connected to a stopcock through which mainstream smoke puffs (45 puffs/cigarette, 20 ml/puff) were delivered. The CS was exhausted with four exhaust holes (1 cm) on the side panels. The animals were exposed to CS in the conscious state and breathed spontaneously in the exposure chamber. In brief, rats were exposed to CS from one cigarette for 20 min. Because the exhaust holes were narrow, the chamber was poorly ventilated so that the rats were expected to inspire high concentrations of CS. After the 20-min exposure to CS, the cage was ventilated by introducing fresh air for 10 min. A total of 10 cigarettes were smoked over 5 h at 30-min intervals. At the end of CS exposure, the rats were transferred to a new cage and allowed to inspire air. After 24 h, the rats were killed and BAL was performed.
Apoptosis assay by light and fluorescence microscopy. Cell samples were affixed to slides by cytospin (Cytospin 3, Shandon, Pittsburgh, PA) and air-dried. For light microscopy, cells were stained with May-Grünwald-Giemsa, and apoptotic cells were identified based on nuclear pyknosis or chromatin condensation together with cell shrinkage. For fluorescence microscopy, cells were fixed with 3% paraformaldehyde and stained with 10 µg/ml of acridine orange (Sigma). Apoptotic cells were identified by nuclear pyknosis or chromatin condensation. To identify macrophages in an apoptotic cell population, slides were first immunostained with anti-macrophage antibody (Ab) as described in Immunocytochemistry and then stained with Giemsa solution.
Terminal deoxynucleotidyltransferase-mediated dUTP nick end labeling assay. A terminal deoxynucleotidyltransferase (TdT)-mediated dUTP nick end labeling (TUNEL) assay was performed with an in situ apoptosis detection kit (Takara Biomedicals, Tokyo, Japan). With this kit, FITC-labeled nucleotides are incorporated at sites of DNA strand breaks by TdT, reacted with horseradish peroxidase-conjugated anti-FITC Ab, and visualized with a peroxidase-substrate reaction. Briefly, slides were fixed in 3% paraformaldehyde and rinsed in PBS. After inactivation of endogenous peroxidase activities in 0.3% H2O2 in methanol for 15 min, the slides were incubated with TdT buffer containing TdT- and FITC-labeled dUTP in a humid atmosphere at 37°C for 60 min. The slides were rinsed in PBS and then covered with mouse monoclonal anti-FITC Ab for 30 min at 37°C. The slides were then rinsed with PBS and stained with 3,3'-diaminobenzidine or 3-amino-9-ethylcarbazole reactions. The TUNEL assay was also performed without TdT as a negative control. Double staining by TUNEL and anti-macrophage Ab was performed as described in Immunocytochemistry.
Immunocytochemistry. Cells affixed to slides by cytospin were fixed with 3% paraformaldehyde, permeabilized in 0.5% Triton X in PBS for 5 min, treated with or without 0.3% H2O2 for 5 min, and washed three times with PBS. After nonspecific binding was blocked with PBS containing 3% BSA and 2% normal goat serum, the slides were incubated with rabbit polyclonal anti-Bax Ab (2 µg/ml; Ab-1, Calbiochem-Novabiochem, La Jolla, CA), mouse monoclonal anti-8-hydroxydeoxyguanosine (8-OHdG) Ab (5 µg/ml; Japan Institute for the Control of Aging, Shizuoka, Japan), or rabbit polyclonal anti-cytochrome c Ab (Santa Cruz Biotechnology, Santa Cruz, CA) for 1 h at room temperature. The primary Abs were captured with anti-IgG-conjugated biotin followed by incubation with a streptavidin-peroxidase complex. The immunoreactants were visualized by 3,3'-diaminobenzidine. Alternatively, the primary Abs were captured with anti-IgG-conjugated Alexa 488 (Molecular Probes, Eugene, OR), and the immunoreactants were observed under an epifluorescence microscope. For double staining with TUNEL and anti-rat macrophage Ab on cytospin slides, TUNEL-stained slides were immersed in 0.1 M glycine-HCl buffer (pH 2.2) for 2 h at room temperature to erase the prior antibody complexes, washed in PBS, and stained with monoclonal anti-rat macrophage Ab (3 µg/ml; ED2, BioSource International, Camarillo, CA) as described above.
Transmission electron microscopy. Cells were prefixed with 1.5% glutaraldehyde in 0.2 M cacodylate buffer, postfixed in 1% osmium tetroxide, dehydrated through absolute alcohol, transferred to propylene oxide, and impregnated overnight in Epon resin (Cosmo-Bio, Tokyo, Japan). This preparation was finally embedded in Epon resin and polymerized at 50°C. The sections were cut with an LKB Ultratome Nova to a thickness of 60 nm and stained with uranyl acetate and lead citrate. The samples of AMs were observed with a transmission electron microscope (Hitachi H-7000) at a magnification of ×6,000.
Evaluation of oxidative stress in CS-exposed AMs. The level of 8-OHdG, which is a marker of oxidative DNA modification, was assessed by an immunocytochemical method with monoclonal anti-8-OHdG Ab (Japan Institute for the Control of Aging) as described in Immunocytochemistry. The cellular level of thiol antioxidants was determined with monochlorobimane (Molecular Probes), which passively diffuses across the plasma membrane into the cytoplasm where it forms blue fluorescent adducts with the reduced form of glutathione and other thiol-containing proteins (39). Cells (5 × 104) were incubated with 50 µM monochlorobimane in 96-well plates for 40 min at 37°C, and then the plates were read on a Cytofluor II multiplate fluorometer (Perseptive Biosystems, Framingham, MA) with excitation and emission wavelengths of 395 and 460 nm, respectively.
Immunoblot analysis. Cell lysates were solubilized in assay buffer (150 mM NaCl, 50 mM Tris-Cl, pH 7.4, 0.5% Nonidet P-40, 0.1% SDS, 10 µg/ml of leupeptin, 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml of aprotinin, and 1 mM sodium orthovanadate) and centrifuged at 10,000 g for 30 min at 4°C. The supernatant containing 50 µg of protein was fractionated by SDS-PAGE, transferred to polyvinylidene difluoride membranes, and probed with 1 µg/ml of Abs against Bcl-2, Bad (Transduction Laboratories, Lexington, KY), Bax, p53 (Calbiochem), and phosphorylated p53 at Ser392 (New England Biolabs, Beverly, MA). Primary Ab was detected by horseradish peroxidase-conjugated Ab (1:2,500), which, in turn, was visualized with enhanced chemiluminescence (SuperSignal, Pierce, Rockford, IL).
For analysis of poly(ADP)-ribose polymerase (PARP) degradation, cell lysates in 1× Laemmli sample buffer were separated by SDS-PAGE, transferred to polyvinylidene difluoride membranes, and probed with 1 µg/ml of anti-PARP Ab (Upstate Biotechnology).Evaluation of mitochondrial function. Functional mitochondria were labeled by incubating cells with 250 nM MitoTracker Red CMXRos (Molecular Probes) for 30 min. The cells were fixed with 3% paraformaldehyde in PBS and observed under an epifluorescence microscope. MitoTracker Red CMXRos is concentrated by active mitochondria, and its fluorescence is reduced and diffused during apoptosis-induced depolarization of mitochondrial inner membrane potential. Depolarization of mitochondrial inner membrane potential was also monitored with the mitochondrial membrane potential-sensitive dye 3,3'-dihexyloxacarbocyanine iodide [DiOC6(3); Molecular Probes]. Briefly, 105 cells were incubated with 40 nM DiOC6(3) in 96-well plates for 15 min at 37°C, and then the plates were read on a Cytofluor II multiplate fluorometer with excitation and emission wavelengths of 485 and 530 nm, respectively.
Assay for caspase-1- and caspase-3-like activities.
Caspase proteolytic activities were measured essentially as previously
described (20). Briefly, 106 cells were placed
in lysis buffer [20 mM Tris · HCl, pH 7.5, 2 mM EDTA, 0.1%
3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS), 10 µg/ml of aprotinin, 10 µg of leupeptin, 1 mM phenylmethylsulfonyl fluoride, and 3 mM dithiothreitol] and were lysed with two cycles of
freezing at 80°C and thawing at 4°C. After centrifugation for 10 min at 13,000 rpm, the supernatants were collected, diluted in 50 mM
Tris · HCl, pH 7.5, 0.1% CHAPS, 10% sucrose, and 10 mM dithiothreitol, and then incubated with the fluorogenic substrates Tyr-Val-Ala-Asp-7-amino-4-methylcoumarin (Peptide Institute, Osaka, Japan) or Asp-Glu-Val-Asp-7-amino-4-methylcoumarin (Peptide Institute) at a concentration of 50 µM for 60 min at 37°C. Release of
7-amino-4-methylcoumarin was measured with excitation at 360 nm and
emission at 460 nm with a spectrofluorometer.
Statistics. Results are presented as means ± SE. Comparisons were made by Student's t-test.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
CS induces AM apoptosis in vitro.
To determine the direct effect of CS on AM apoptosis, we added
aqueous CSE to culture medium of in vitro AMs (38), which is a simple, reproducible, and widely used method. Treatment of rat AMs with CSE-induced apoptosis as demonstrated by light
microscopy of May-Grünwald- Giemsa-stained cells (Fig.
1, A and B),
fluorescence microscopy of acridine orange-stained cells (Fig. 1,
C and D), TUNEL (Fig. 1, E and
F), and transmission electron microscopy (Fig. 1,
G and H). AMs exposed to CS clearly showed
morphological hallmarks of apoptosis such as cellular
shrinkage, cell surface smoothing, nuclear compaction, and chromatin
condensation at the periphery of the nuclear envelope (Fig. 1,
B and F). However, they rarely showed nuclear
fragmentation, another feature of apoptosis that was observed
in AMs exposed to diethyl maleate (5 × 105 M),
tumor necrosis factor-
(1 µg/ml) plus cycloheximide (1 × 10
4 M), or bleomycin (100 µg/ml; data not shown),
suggesting that AMs exposed to CS undergo a different mechanism of
apoptosis. As shown in Fig. 2,
apoptosis by CSE occurred in a dose- and time-dependent manner,
with 93% of AMs showing apoptosis after 24-h exposure of AMs
to 10 vol% of CSE solution. Gas-phase CSE, which was generated by
removing the particle (tar) component from whole smoke, induced apoptosis much less than whole CSE (8.5 ± 0.6 vs.
72.5 ± 2.1% apoptosis after 24-h exposure to gas-phase
CSE and whole CSE, respectively), suggesting that some of the tar
component is required for induction of apoptosis.
Apoptosis of AMs was not induced by nicotine (10
5
to 10
8 M), a major tar component of CS (data not shown).
|
|
|
Oxidative stress mediates CS-induced AM apoptosis.
Because both CS, particularly its tar component, and CSE solutions
contain ROS (25, 30), we evaluated the role of oxidative stress in CS-induced apoptosis of AMs. As shown in Fig.
4, A and B,
exposure of rat AMs to CSE increased the level of 8-OHdG, a marker of
oxidative DNA modification, and decreased the level of thiol
antioxidants such as glutathione, indicating that CS imposes oxidative
stress on AMs. Because ROS in the respiratory tract are effectively
eliminated by many antioxidants in respiratory tract lining fluids
(RTLFs) (4), we examined whether CS-induced AM
apoptosis is inhibited by antioxidants. As shown in Fig.
4C, CS-induced AM apoptosis was significantly
inhibited by N-acetylcysteine (100 µM) and the RTLF
antioxidants glutathione (100 µM), ascorbic acid (100 µM), and
-tocopherol (0.7 µM) at physiological concentrations (4).
|
CS promotes accumulation of Bax protein.
The Bcl-2 protein family is a major class of intracellular regulators
of apoptosis (1). Some, such as Bcl-2 and
Bcl-XL, suppress apoptosis, whereas others such as
Bax, Bad, and Bid promote it (1). The balance between
proapoptotic and antiapoptotic members determines the fate of
various types of cells. Therefore, we evaluated the effect of CS on the
cellular levels of Bcl-2, Bax, and Bad proteins. CS promoted the
accumulation of Bax protein in rat AMs (Fig.
5) but had no effect on the levels of
Bcl-2 or Bad (Fig. 5A). These results suggest that CS
promotes the accumulation of Bax protein by AMs.
|
CS-induced AM apoptosis is independent of p53 and Fas.
We then examined the mechanism of CS-induced apoptosis in a
mouse model lacking the p53 gene. Because p53 has been demonstrated to
induce Bax gene expression and ROS-dependent apoptosis
(23), we asked whether CS-induced apoptosis is
dependent on p53. We found that CS induced both Bax protein
accumulation and apoptosis in AMs obtained from
p53(/
) mice (Fig. 6,
A-D). In addition, immunoblot analysis
showed that CS did not promote p53 protein accumulation and
phosphorylation by rat AMs (data not shown). These results suggest that
CS-induced AM apoptosis is independent of p53. We also asked
whether Fas-Fas ligand interactions, which mediate
apoptosis in various types of cells including macrophages, are
required for CS-induced apoptosis of AMs. CS induced
apoptosis in AMs obtained from Fas ligand-defective mice and
those from Fas-deficient mice (Fig. 6, E-H),
suggesting that CS-induced apoptosis is also independent of
Fas-Fas ligand interactions.
|
Mitochondrial dysfunction and cytochrome c release during
CS-induced apoptosis.
Because Bax protein can trigger mitochondria to release
caspase-activating proteins including cytochrome c
(6), probably accelerating the opening of the
voltage-dependent anion channel, we evaluated the effect of CS on
mitochondrial function and cytochrome c release. MitoTracker
Red CMXRos, a fluorescent dye that accumulates in functional
mitochondria, was used to evaluate mitochondrial function (Fig.
7A). Rat AMs not exposed to
CSE displayed a bright and punctuate MitoTracker Red CMXRos staining
pattern. By contrast, when treated with 5 vol% CSE for 12 h, AMs
showed diffuse MitoTracker Red CMXRos staining, suggesting that the
mitochondrial inner membrane potential had decayed. A reduction in
mitochondrial inner membrane potential in CSE-exposed AMs was also
shown by a decline in the fluorescence level of
DiOC6(3), which reflects depolarization of the
mitochondrial membrane (12) (Fig. 7B).
Immunocytochemistry demonstrated cytochrome c release from
mitochondria to the cytoplasm on exposure to CSE (Fig. 7C).
These results suggest that mitochondrial dysfunction and cytochrome
c release occur during CS-induced apoptosis.
|
CS-induced AM apoptosis is independent of caspase
activation.
Although caspases are key mediators of apoptosis, recent
evidence has shown that Bax induces apoptosis by either a
caspase-dependent or caspase-independent mechanism (1, 18, 22,
36). Therefore, we examined whether CS-induced AM
apoptosis is dependent on caspase activation. First, we tested
several cell membrane-permeable peptide inhibitors of caspases for
their ability to prevent CS-induced apoptosis. As shown in Fig.
8A, CS-induced apoptosis of rat
AMs was not inhibited in the presence of Boc-D-FMK (general caspase inhibitor; Enzyme System Products, Livermore, CA),
Tyr-Val-Ala-Asp-chloromethylketone (inhibitor of caspase-1 and -4;
Bachem), or Asp-Glu-Val-Asp-aldehyde (inhibitor of caspase-3, -6, -7, -8, and -10; BIOMOL Research, Plymouth Meeting, PA) at a concentration
of 50 µM. We had previously confirmed that these inhibitors at a
concentration of 50 µM had no toxicity to rat AMs by a colorimetric
MTT assay (data not shown). Second, CS did not promote caspase-1-like
or caspase-3-like proteolytic activities (Fig. 8B). Third,
immunoblot analysis demonstrated that PARP, a nuclear protein cleaved
by multiple caspases to an 85-kDa inactive form, was not degraded
during CS-induced apoptosis of AMs (Fig. 8C). Taken
together, these results suggest that CS-induced apoptosis of
AMs is independent of any known major caspases.
|
Induction of AM apoptosis by CS in vivo.
To determine whether CS also induces AM apoptosis in vivo, rats
were exposed to the smoke from 10 cigarettes over 5 h at 30-min intervals in an exposure chamber. BAL fluid cells were obtained 24 h after CS exposure, and apoptosis was evaluated by light
microscopy of May-Grünwald-Giemsa-stained cells (Fig.
9, A and B) and
Giemsa-stained cells (Fig. 9, E and F),
transmission electron microscopy (Fig. 9, C and
D), and TUNEL (Fig. 9, G and H). BAL
fluid cells recovered from CS-exposed rats contained a small population
of apoptotic cells (Fig. 9, B, D,
F, and H). These apoptotic cells were
positively stained with anti-macrophage Ab (Fig. 9, F and
H), indicating that some AMs underwent apoptosis in
CS-exposed rats. However, the incidence of AM apoptosis in
CS-exposed rats (3.2%) was much lower than expected based on the
results of in vitro studies (Fig. 2). This may have been due to the
long 24-h interval between CS exposure and BAL fluid collection.
Alternatively, because antioxidants inhibited CS-induced AM
apoptosis in vitro (Fig. 4C), antioxidants present
in the respiratory tract may have prevented the effect of CS inhalation
on AM apoptosis.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
AMs represent the chief detoxifying mechanism for inhaled materials. Some inhaled toxic materials such as silica, asbestos, and other particulates are known to induce apoptosis of AMs (9, 14). The present study demonstrated that acute exposure to CS also induces apoptosis of AMs in vitro and, to some extent, in vivo. This apoptosis was associated with increased oxidative stress, Bax protein accumulation, mitochondrial dysfunction, and mitochondrial cytochrome c release but was independent of p53, Fas, and caspase activation. Importantly, the CS-induced apoptosis of AMs was inhibited by several antioxidants known to be present in the respiratory tract.
The findings of the present study implicate oxidative stress as a
mechanism of CS-induced apoptosis. CS is a rich source of ROS
and ROS inducers, and ROS, particularly at low levels, can induce
apoptosis in some types of cells. However, the composition of
ROS differs between different components of CS. The tar component of CS
contains large quantities of stable and cell membrane-permeable radicals such as hydroquinones that redox cycle to form
O
Our findings also suggest that AM apoptosis by CS is associated with Bax protein accumulation but not with caspase activation. Although caspases are key mediators of apoptosis, recent evidence (18, 21, 22, 36) indicates that some apoptosis may occur without caspase involvement. For example, infection of AMs by Chlamydia psittaci has recently been reported to induce apoptosis by a caspase-independent mechanism (21). Furthermore, Bax protein has been shown to induce both cytochrome c release and apoptosis-like cell death without caspase activation (16, 36). In this respect, recent evidence suggests that Bax and Bax-like proteins mediate caspase-independent death via channel-forming activity (1), which could promote mitochondrial permeability or puncture the mitochondrial outer membrane (8). In addition, CS may directly inhibit caspase functions through its oxidative activity because the caspases are cysteine-dependent enzymes that are sensitive to the redox status of the cells (10). This concept is supported by a recent report (29) documenting that CS directly inhibits caspase-3 activity. Taken together, these lines of evidence favor our conclusion that CS induces apoptosis of AMs through a mechanism other than caspase activation. However, we cannot exclude the possibility that CS-induced apoptosis is mediated by an as yet uncharacterized caspase or by other proteases. The mechanisms of apoptosis other than caspase activation may include activation of apoptosis-inducing factor, a newly identified mitochondrial protein that translocates into the nuclei and degrades DNA in response to apoptotic stimuli (15). However, this mechanism is unlikely involved in CS-induced AM apoptosis because an immunocytochemical study of CS-exposed AMs could not detect nuclear translocation of apoptosis-inducing factor from the mitochondria (data not shown).
The morphological features of CS-induced apoptosis resemble those of apoptosis previously reported to occur by a caspase-independent mechanism (22, 36). Caspase-independent apoptosis, some of which involves Bax, is associated with chromatin condensation, cell shrinkage, and mitochondrial dysfunction but not with chromatin fragmentation (22, 36). These changes are similar to those in CS-induced AM apoptosis found in this study. Both internucleosomal DNA fragmentation and chromatin fragmentation are known to require caspase activation. However, it has been reported that chromatin condensation during apoptosis does not require DNA fragmentation (28). In this context, agarose gel electrophoresis of DNA isolated from CS-exposed AMs in the present study could not detect any significant level of internucleosomal DNA fragmentation (data not shown).
Our finding of CS-induced AM apoptosis seems to be inconsistent with clinical observations in chronic cigarette smokers. The number of AMs in the lung is much greater in smokers than in nonsmokers (3). Indeed, macrophage alveolitis and respiratory bronchiolitis are thought to be the early changes in cigarette smokers, which, in susceptible individuals, probably lead to the alveolar wall destruction seen in pulmonary emphysema. However, if CS induces AM apoptosis in vivo, the number of AMs in the lung would decrease.
A simple explanation for the increased number of AMs in smokers is that
the rate of monocyte recruitment from the blood into the lung exceeds
the rate of AM death by apoptosis. Alternatively, there may be
a mechanism in place to limit CS-mediated apoptosis of AMs in
smokers' lungs. First, AMs exposed to chronic CS in smokers' lungs
may become resistant to apoptosis. A very recent study
(34) showed that expression of Bcl-XL, an
antiapoptotic protein, was increased in AMs from smokers compared
with those from nonsmokers. Second, antioxidants present in the
respiratory tract may prevent apoptosis of AMs by CS
inhalation. The respiratory tract is covered with RTLFs that contain
various antioxidants acting as a defensive shield against inhaled ROS
and CS (4). In this context, the results of our in vitro
studies show that glutathione, ascorbic acid, and -tocopherol, which
are considered to be important antioxidants in the respiratory tract
(4), inhibited CS-induced apoptosis of AMs.
Chronic smoke exposure in humans and hamsters has also been shown to
increase antioxidant enzyme activities in AMs (17). This
increased activity may serve as a mechanism to limit CS-mediated
apoptosis in AM in smokers' lungs. Although the life span of
AMs in the normal lung is estimated to be ~80 days (33),
studies are needed to determine the survival of AMs within smokers' lungs.
The CS exposure experiments in vitro and in vivo in the present study have several limitations that may make them difficult to extrapolate to the clinical situation. First, the in vitro exposure to CSE does not simulate in vivo CS exposure. Although exposure to CSE is a standard procedure, its relevancy to the in vivo state remains unclear. Second, the CS exposure in vivo in the present study seems to be very intense as evidenced by the occurrence of some degree of alveolar bleeding. This is never observed in human smokers. Third, because mice are obligatory nasal breathers, some toxic products that would be normally inhaled by humans may have been deposited in the nasal passage of the mice. Fourth, the CS exposure experiments in this study are a model of acute smoking and cannot be extrapolated to what occurs in smokers who inhale CS chronically and intermittently. Although these limitations must be taken into account when interpreting the results of this study in relation to cigarette smokers, the results demonstrate that CS has the ability to induce apoptosis of AMs.
![]() |
ACKNOWLEDGEMENTS |
---|
We are very grateful to Masayuki Shino and Yoshimi Sugimura for excellent technical assistance.
![]() |
FOOTNOTES |
---|
This work was supported by Grant-in Aid for Scientific Research 12670580 from the Ministry of Education, Science, and Culture, Japan.
Address for reprint requests and other correspondence: K. Aoshiba, First Dept. of Medicine, Tokyo Women's Medical Univ., 8-1 Kawada-cho, Shinjuku-ku, Tokyo 162-8666, Japan (E-mail: kaoshiba{at}chi.twmu.ac.jp).
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 12 June 2001; accepted in final form 30 July 2001.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Adams, JM,
and
Cory S.
The Bcl-2 protein family: arbiters of cell survival.
Science
281:
1322-1326,
1998
2.
Aoshiba, K,
Nagai A,
and
Konno K.
Nicotine prevents a reduction in neutrophil filterability induced by cigarette smoke exposure.
Am J Respir Crit Care Med
150:
1101-1107,
1994[Abstract].
3.
Costabel, U,
and
Guzman J.
Effect of smoking on bronchoalveolar lavage constituents.
Eur Respir J
5:
776-779,
1992[ISI][Medline].
4.
Cross, CE,
Van Der Vliet A,
Eiserich JP,
and
Wong J.
Oxidative stress and antioxidants in respiratory tract lung fluids.
In: Oxygen, Gene Expression, and Cellular Function, edited by Clerch LB,
and Massaro DJ.. New York: Dekker, 1997, vol. 105, p. 367-398, 1997. (Lung Biol Health Dis Ser)
5.
Foster, PS,
Hogan SP,
Ramsay AJ,
Matthaei KI,
and
Young IG.
Interleukin 5 deficiency abolishes eosinophilia, airway hyperreactivity, and lung damage in a mouse asthma model.
J Exp Med
183:
195-201,
1996[Abstract].
6.
Green, DR,
and
Reed JC.
Mitochondria and apoptosis.
Science
281:
1309-1312,
1998
7.
Green, GM.
Cigarette smoke: protection of alveolar macrophages by glutathione and cysteine.
Science
162:
810-811,
1968[ISI][Medline].
8.
Green, GM,
and
Carolin D.
The depressant effect of cigarette smoke on the in vitro antibacterial activity of alveolar macrophages.
N Engl J Med
276:
421-427,
1967[ISI][Medline].
9.
Hamilton, RF,
Li L,
Lyer R,
and
Holian A.
Asbestos induces apoptosis in human alveolar macrophages.
Am J Physiol Lung Cell Mol Physiol
271:
L813-L819,
1996
10.
Hampton, MB,
Fadeel B,
and
Orrenius S.
Redox regulation of the caspases during apoptosis.
Ann NY Acad Sci
854:
328-335,
1998
11.
Harris, JO,
Swenson EW,
and
Johnson JE.
Human alveolar macrophages: comparison of phagocytic ability, glucose utilization, and ultrastructure in smokers and nonsmokers.
J Clin Invest
49:
2086-2096,
1970[ISI][Medline].
12.
Haugland, RP.
Handbook of Fluorescent Probes and Research Chemicals (6th ed.). Eugene, OR: Molecular Probes, 1996.
13.
Holt, PG,
and
Keast D.
The effect of tobacco smoke on protein synthesis in macrophages.
Proc Soc Exp Biol Med
142:
1243-1247,
1973.
14.
Iyer, R,
Hamilton RF,
Li L,
and
Holian A.
Silica-induced apoptosis mediated via scavenger receptor in human alveolar macrophages.
Toxicol Appl Pharmacol
141:
84-92,
1996[ISI][Medline].
15.
Joza, N,
Susin SA,
Daugas E,
Stanford WL,
Cho SK,
Li CYJ,
Sasaki T,
Elia AJ,
Cheng MM,
Ravagnan L,
Ferri KF,
Zamzami N,
Wakeham A,
Hakem R,
Yoshida H,
Kong Y,
Mark TW,
Zuniga-Pflücker JC,
Kroemer G,
and
Penninger JM.
Essential role of the mitochondrial apoptosis-inducing factor in programmed cell death.
Nature
410:
549-554,
2001[ISI][Medline].
16.
McCarthy, NJ,
Whyte MKB,
Gilbert CS,
and
Evan GI.
Inhibition of Ced-3/ICE-related proteases does not prevent cell death induced by oncogene DNA damage, or the Bcl-2 homologue Bak.
J Cell Biol
136:
215-227,
1997
17.
McCusker, K,
and
Hoidal J.
Selective increase of antioxidant enzyme activity in the alveolar macrophages from cigarette smokers and smoke-exposed hamsters.
Am Rev Respir Dis
141:
678-682,
1990[ISI][Medline].
18.
Miller, TM,
Moulder KL,
Knudson CM,
Creedon DJ,
Deshmukh M,
Korsmeyer SJ,
and
Johnson EM, Jr.
Bax deletion further orders the cell death pathway in cerebellar granule cells and suggests a caspase-independent pathway to cell death.
J Cell Biol
139:
205-217,
1997
19.
Oddo, M,
Renno T,
Attinger A,
Bakker T,
MacDonald HR,
and
Meylan PRA
Fas ligand-induced apoptosis of infected human macrophages reduces the viability of intracellular Mycobacterium tuberculosis.
J Immunol
160:
5448-5454,
1998
20.
Ohta, T,
Kinoshita T,
Naito M,
Nozaki T,
Masutani M,
Tsuruo T,
and
Miyajima A.
Requirement of the caspase-3/CPP32 protease cascade for apoptotic death following cytokine deprivation in hematopoietic cells.
J Biol Chem
272:
23111-23116,
1997
21.
Ojcius, DM,
Souque P,
Perfettini JL,
and
Dautry-Varsat A.
Apoptosis of epithelial cells and macrophages due to infection with the obligate intracellular pathogen Chlamydia psittaci.
J Immunol
161:
4220-4226,
1998
22.
Okuno, S,
Shimizu S,
Ito T,
Nomura M,
Hamada E,
Tsujimoto Y,
and
Matsuda H.
Bcl-2 prevents caspase-independent cell death.
J Biol Chem
273:
34272-34277,
1998
23.
Polyak, K,
Xia Y,
Zweier JL,
Kinzler KW,
and
Vogelstein B.
A model for p53-induced apoptosis.
Nature
389:
300-305,
1997[ISI][Medline].
24.
Pratt, SA,
Smith MH,
Ladman AJ,
and
Finley TN.
The ultrastructure of alveolar macrophages from human cigarette smokers and nonsmokers.
Lab Invest
24:
331-338,
1971[ISI][Medline].
25.
Pryor, WA,
and
Stone K.
Oxidants in cigarette smoke: radicals, hydrogen peroxide, peroxynitrate, and peroxynitrite.
Ann NY Acad Sci
686:
12-28,
1993[ISI][Medline].
26.
Rodriguez, RJ,
White RR,
Senior RM,
and
Levine EA.
Elastase release from human alveolar macrophages: comparison between smokers and non-smokers.
Science
198:
313-314,
1977[ISI][Medline].
27.
Roque, AL,
and
Pickren JW.
Enzymatic changes in fluorescent alveolar macrophages of the lungs of cigarette smokers.
Acta Cytol
12:
420-429,
1968[ISI][Medline].
28.
Sakahira, H,
Enari M,
Ohsawa Y,
Uchiyama Y,
and
Nagata S.
Apoptotic nuclear morphologic changes without DNA fragmentation.
Curr Biol
9:
543-560,
1999[ISI][Medline].
29.
Sarafian, TA,
Roth MD,
and
Tashkin DP.
Inhibition of caspase-3 activity by marijuana smoke (Abstract).
Am J Respir Crit Care Med
159:
A619,
1999[ISI].
30.
Stone, K,
Bermúdez E,
and
Pryor WA.
Aqueous extracts of cigarette tar containing the tar free radical cause DNA nicks in mammalian cells.
Environ Health Perspect
102, Suppl10:
173-178,
1994[ISI][Medline].
31.
Stuart, RS,
Higgins WH,
and
Brown PW.
In vitro toxicity of tobacco smoke solutions to rabbit alveolar macrophages.
Arch Environ Health
33:
135-140,
1978[ISI][Medline].
32.
Takahashi, H,
Ishidoh K,
Muno D,
Ohwada A,
Nukiwa T,
Kominami E,
and
Kira S.
Cathepsin L activity is increased in alveolar macrophages and bronchoalveolar lavage fluid of smokers.
Am Rev Respir Dis
147:
1562-1568,
1993[ISI][Medline].
33.
Thomas, ED,
Ramberg RE,
Sale GE,
Sparkes RS,
and
Golde DW.
Direct evidence for a bone marrow origin of the alveolar macrophages in man.
Science
192:
1016-1018,
1976[ISI][Medline].
34.
Tomita, K,
Caramori G,
Lim S,
Chung KF,
Shimizu E,
Barnes PJ,
and
Adcock IM.
Enhanced expression of Bcl-XL in alveolar macrophages from smokers (Abstract).
Am J Respir Crit Care Med
163:
A188,
2001.
35.
Voisin, C,
Aerts C,
Fournier E,
and
Firlik M.
Acute effects of tobacco smoke on alveolar macrophages cultured in gas phase.
Eur J Respir Dis
66, Suppl139:
76-81,
1985[ISI].
36.
Xian, J,
Chao DT,
and
Korsmeyer SJ.
BAX-induced cell death may not require interleukin 1-converting enzyme-like proteases.
Proc Natl Acad Sci USA
93:
14559-14563,
1996
37.
Yamaguchi, E,
Okazaki N,
Itoh A,
Abe S,
Kawakami Y,
and
Okuyama H.
Interleukin 1 production by alveolar macrophages is decreased in smokers.
Am Rev Respir Dis
140:
397-402,
1989[ISI][Medline].
38.
York, GK,
Arth C,
Stumbo JA,
Cross CE,
and
Mustafa MG.
Pulmonary macrophages respiration as affected by cigarette smoke and tobacco extract.
Arch Environ Health
27:
96-98,
1973[ISI][Medline].
39.
Young, PR,
ConnorsWhite AL,
and
Dzido GA.
Kinetic analysis of the intracellular conjugation of monochlorobimane by IC-21 murine macrophage glutathione-S-transferase.
Biochim Biophys Acta
1201:
461-465,
1994[ISI][Medline].