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Alveolar Wall Apoptosis Causes Lung Destruction and Emphysematous Changes

First Department of Medicine, Tokyo Women's Medical University, Tokyo, Japan

Address correspondence to: Kazutetsu Aoshiba, M.D., First Department of Medicine, Tokyo Women's Medical University, 8-1 Kawada-cho, Shinjuku-ku, Tokyo 162-8666, Japan. E-mail: kaoshiba{at}chi.twmu.ac.jp

	Abstract

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Pulmonary emphysema is characterized by alveolar wall destruction and airspace enlargement. Recent evidence indicates that epithelial or endothelial apoptosis may be involved in the pathogenesis of emphysema. Here, we describe the induction of emphysematous changes, including airspace enlargement, alveolar wall destruction, and enhanced lung distensibility, in mice receiving a single intratracheal injection of active caspase-3 and Chariot, a newly developed protein transfection reagent. Epithelial apoptosis and enhanced elastolytic activity (optimal at pH 5.5) in bronchoalveolar lavage were noted. Emphysematous changes were also generated in mice receiving an intratracheal injection of nodularin, a proapoptotic serine/threonine kinase inhibitor. This murine model provides direct evidence that confirms that alveolar wall apoptosis causes emphysematous changes. Furthermore, this simple technique for protein transfection of lung tissue can be used in a variety of future applications.

Abbreviations: bronchoalveolar lavage, BAL • Dulbecco's modified Eagle's medium, DMEM • dimethyl sulfoxide, DMSO • cell-extracellular matrix, ECM • fluorescein isothiocyanate, FITC • horseradish peroxidase, HRP • phosphate-buffered saline, PBS • single-stranded DNA, ssDNA • terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling, TUNEL

	Introduction

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Pulmonary emphysema is a pathologic condition in the lung characterized by enlargement of airspaces distal to the terminal bronchiole, the destruction of alveolar walls, and loss of the alveolar unit (1). Although emphysema is an important cause of morbidity and mortality in modern society (2), its pathogenesis remains enigmatic. The most prevailing hypothesis since the 1960s for the pathogenesis of emphysema has been the elastase/antielastase imbalance theory. According to this theory, chronic exposure to cigarette smoke leads to the invasion of inflammatory cells into the lung airspaces; the inflammatory cells subsequently release large quantities of elastases, leading to an elastase/antielastase imbalance with the destruction of lung tissue (3). However, this hypothesis does not fully explain why alveolar cells and alveolar wall structures are lost in emphysema. An alternative hypothesis presently being investigated posits the loss of alveolar wall structures through epithelial and endothelial apoptosis (4, 5). Recent clinical studies have shown elevated levels of apoptosis in alveolar walls of emphysematous lungs (6, 7). A recent study has also shown that the chronic treatment of rats with vascular endothelial growth factor receptor-2 inhibitor SU5416 causes endothelial apoptosis followed by airspace enlargement (4), although direct evidence linking apoptosis and emphysema has not been obtained.

Here, we describe the induction of emphysematous changes by alveolar wall apoptosis. When mice were given a single intratracheal injection of active caspase-3 and Chariot, a newly developed protein transfection reagent, alveolar epithelial apoptosis followed by alveolar wall destruction and airspace enlargement was observed. Similar lung lesions were also noted in mice receiving an intratracheal injection of nodularin, a proapoptotic serine/threonine kinase inhibitor. These results provide direct evidence that alveolar wall apoptosis causes emphysematous changes. In addition, Chariot-mediated protein transfection is a simple technique that could be used for a variety of future applications.

	Materials and Methods

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Protein Transfection Reagents
Chariot (Active Motif, Carlsbad, CA), a newly developed noncytotoxic protein transfection reagent, was used to transfer the proteins into the cells via a noncovalent complex based on a short amphipathic peptide carrier, Pep-1 (8). For in vivo experiments, 1 µl of Chariot was mixed with 20 µl of phosphate-buffered saline (PBS) containing either ß-galactosidase (2 µg = 20 unit; Active Motif) or recombinant human active caspase-3 (3.7 µg; MBL, Nagoya, Japan) with or without DEVD-CHO (5 µg; Biomol, Plymouth Meeting, PA). After incubation for 30 min at room temperature, the complex was diluted in PBS to a volume of 50 µl. For the in vitro experiments, 1 µl of Chariot diluted in 10 µl of 80% dimethyl sulfoxide (DMSO) was mixed with 10 µl of PBS containing active caspase-3 (1 µg). After incubation for 30 min at room temperature, the complex was diluted in serum-free Dulbecco's modified Eagle's medium (DMEM) to a volume of 100 µl.

Cell Culture
Primary rat type II alveolar epithelial cells were isolated as previously described (9).

Animal Treatment
The animal protocol was approved by the Animal Care and Use Committee of Tokyo Women's Medical University. Six-week-old male C57BL/6 mice were intratracheally given 50 µl of PBS solutions containing (i) PBS alone; (ii) active caspase-3 (3.7 µg); (iii) Chariot (1 µl); (iv) a complex of active caspase-3 and Chariot; (v) a complex of active caspase-3, DEVD-CHO (5 µg), and Chariot; (vi) a complex of Chariot and ß-galactosidase (2 µg); or (vii) nodularin (1 µg; Calbiochem, San Diego, CA). Animals were killed by terminal anesthesia at various time points, and then the hearts and lungs were excised en bloc.

Tissue Processing
The lungs were inflated and fixed by intratracheal instillation of 10% formalin at a constant pressure of 25 cm H₂O. The volume of the left lung was measured by water displacement, and the lung distensibility (lung volume divided by fixing pressure, µl/cm H₂O) was calculated. Bilateral lungs were then embedded in paraffin and sectioned sagittally (3 µm) for histologic analysis. For ß-galactosidase staining, the lungs were inflated by manual instillation with 50% O.C.T. compound, quick frozen, and sectioned (3 µm).

Bronchoalveolar Lavage
The lungs were lavaged with 1 ml of PBS through an intratracheal cannula. The cytocentrifuge preparations were stained with May-Grünwald-Giemsa or a modified papanicolaou solution (10), and the remaining slides were stored at -70°C until further use. The supernatant was concentrated 10-fold by ultrafiltration and stored at -70°C.

Lung Histology
The mean cord length (a measure of average alveolar size) was estimated using computer-assisted image analysis. Two sagittal paraffin sections from each lung were stained with hematoxylin and eosin, and digitized video images of the entire lung fields were captured using an Olympus BX60 microscope (Olympus Optical Co., Ltd., Tokyo, Japan) with an Olympus DP50 CCD camera. Fields containing nonalveolated structures, such as bronchovascular bundles, were discarded. The video output of the camera was sent to an Olympus imaging microscopic workstation (CUSL2G40; Olympus Optical Co., Ltd.) equipped with a computer running Microsoft Windows 98 and a computerized color image analysis software system (Win Roof Version 3.5; Mitani Corporation, Fukui, Japan). Each image was then subjected to operations with 400 horizontal grid lines, and the length of the lines overlying an air space was measured and averaged as the mean cord length. The destructive index (a measure of alveolar wall destruction) was estimated as described previously (11).

Immunostaining
Monoclonal anti-pancytokeratin (1:300 dilution; Sigma, St. Louis, MO), polyclonal anti-macrophages (1:200; Inter-Cell Technologies, Hopewell, NJ), or monoclonal anti–single-stranded DNA (ssDNA) (1:100; DAKO Japan, Tokyo, Japan) were used as the primary antibodies. The primary antibody was reacted with biotinylated anti-IgG and either streptavidin–alkaline phosphatase or streptavidin–horseradish peroxidase. Immunoreactants were visualized using a NBT/BCIP or diaminobenzidine-substrate solution. After immunostaining, some slides were stained by terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL).

TUNEL
TUNEL was performed using the Takara In Situ Apoptosis Detection Kit (Takara Biomedicals, Tokyo, Japan), according to the manufacturer's instructions. With this kit, fluorescein isothiocyanate (FITC)-labeled nucleotides are incorporated at the sites of DNA strand breaks via TdT, reacted with horseradish peroxidase–conjugated anti-FITC antibody, and visualized by a diaminobenzidine-substrate reaction. The slides were counterstained with hematoxylin.

_ß-Galactosidase Staining
Frozen lung tissue sections were fixed in PBS containing 2% formaldehyde and 0.2% glutaraldehyde, rinsed with PBS, and stained with a solution containing 40 mM of potassium ferricyanide, 40 mM of potassium ferrocyanide, 20 mM of magnesium chloride, and 2 mg/ml of X-gal. Some slides were then immunostained for pan-cytokeratin antigen, as described below.

In Vitro Elastin Degradation
Elastin degradation was quantified by incubating insoluble FITC-conjugated elastin (3.3 mg/ml; Elastin Products, Owensville, MI) with human recombinant caspases-1, -2, -3, -6, -7, -8, -9, and -10 (20 unit/ml each; MBL), purified porcine pancreatic elastase (0.1, 0.5, 1 U/ml; Sigma), or 10-fold concentrates of bronchoalveolar lavage (BAL) fluid samples in either an acidic (40 mM sodium acetate, pH 5.5; 0.1% CHAPS; 10 mM DTT) or neutral (20 mM HEPES, pH 7.4; 4 mM CaCl₂; 50 mM KCl; 250 mM sucrose; 5 mM ZnCl₂; 0.1% CHAPS; 10 mM DTT) reaction buffer. After 24 h of incubation at 37°C, the aliquots were centrifuged, and the solubilized products were measured in a spectrofluorometer. Elastin degradation was also quantified using the EnzChek Elastase Assay kit (Molecular Probes, Eugene, OR) according to the manufacturer's instructions. This kit uses DQ elastin, a soluble elastin that has been labeled with quenched BODIPY FL dye (Molecular Probes); the nonfluorescent substrate produces highly fluorescent fragments upon digestion. Briefly, DQ elastin (25 µg/ml) was incubated with samples in the reaction buffer in the absence of DTT for 1 h at 37°C. After the incubation period, the digestion of DQ elastin was measured using a spectrofluorometer. Elastin degradation was also tested by incubating -elastin (40 µg; Elastin Products) with samples in the reaction buffer for 24 h at 37°C. Aliquots were then subjected to analysis by SDS-PAGE and silver staining, as described previously (12).

In Situ Elastin Zymography
Elastin degradation in BAL cells or primary rat type II epithelial cells was examined using in situ elastin zymography and DQ elastin. Cytocentrifuge slides of the BAL cells were overlaid with 250 µg/ml of DQ elastin in either an acidic or neutral reaction buffer as described above. In some experiments, the buffer included 10 µM of ZVAD-fmk (Bachem AG, Bubendorf, Switzerland), a broad caspase inhibitor. After 20 min of incubation at 37°C, the slides were fixed with 3% paraformaldehyde and counterstained with 5 µg/ml Hoechst33342 (Sigma). DQ elastin digestion and apoptosis were detected by epifluorescence microscopy. The degradation of elastin layers by type II epithelial cells was evaluated using a culture system. Primary rat type II epithelial cells were plated in DMEM containing 10% FCS on 8-chamber glass slides that were precoated with a mixture of 30 µg/ml of type I collagen (Vitrogen) and 250 µg/ml of DQ elastin. After 16 h of culture, apoptosis was induced by incubating the cells with 100 µl of serum-free DMEM containing Chariot and active caspase-3, prepared as described in the section on protein transfection reagents. After 2 h of apoptosis-induction period, the cells were fixed in 3% paraformaldehyde, counterstained with 5 µg/ml Hoechst33342, and observed by epifluorescence microscopy.

Statistical Analysis
Data are expressed as the means ± SEM. The differences were examined for significance using an ANOVA with Scheffe's procedure post hoc analysis or with the Student's two-tailed unpaired t test, as appropriate, using StatView software for the Macintosh (Abacus Concepts, Inc., Berkeley, CA).

	Results

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Protein Transfection of Alveolar Wall Tissue Using Chariot
ß-Galactosidase staining of lung sections from mice treated with Chariot and ß-galactosidase (a 119-kD marker protein) showed focal positive staining within the alveolar and bronchial epithelium (Figure 1). The majority (88.3 ± 2.3%, n = 3) of the ß-galactosidase–positive cells were positively immunostained for pan-cytokeratin antigen, a marker for epithelial cells. Alternatively, 3.25 ± 0.6% (n = 3) of pan-cytokeratin–positive alveolar epithelial cells were stained with ß-galactosidase. By contrast, very few of the ß-galactosidase–positive cells were positive for CD31 antigen, a marker for endothelial cells (data not shown). Alveolar macrophages were almost all negative for ß-galactosidase. These results show that Chariot can be used for protein transfection in the alveolar epithelium.

View larger version (134K): [in this window] [in a new window]   Figure 1. ß-Galactosidase staining of lung tissue from mice killed 2 h after intratracheal injection of Chariot and ß-galactosidase. (A) ß-Galactosidase–positive signals (green staining) are visible within the alveolar and bronchial wall (original magnification: x100). (B) The ß-galactosidase–positive cells are immunopositive for pan-cytokeratin antigen (brown staining; original magnification: x1,000). These photomicrographs are representative of the results obtained in three animals.

View larger version (99K): [in this window] [in a new window]   Figure 2. Active caspase-3 transfection induces alveolar wall destruction and emphysematous changes. (A) Hematoxylin and eosin–stained lung tissue sections (a–n) and Elastica and van Gieson–stained lung tissue sections (o and p) from mice killed 6 h after treatment with PBS (a, b, and o), active caspase-3 (3.7 µg) (c and d), Chariot (1 µl) (e and f), active caspase-3 and Chariot (g, h, m, and p), a mixture of Chariot, active caspase-3, and DEVD-CHO (5 µg) (i and j), or nodularin (2 µg) (k, l, and n) (original magnification: a, c, e, g, i, and k, x40; b, d, f, h, j, l, o, and p, x100; m and n, x1,000). Mice treated with either active caspase-3 and Chariot or nodularin exhibit alveolar enlargement (g, h, k, and l), the disappearance of alveolar cells and the loss of alveolar structures (m and n), and the loss of elastin fiber staining (p), all indicative of emphysema. These photomicrographs are representative of the results obtained for a minimum of four animals in each group. (B) Quantification of mean chord length (closed columns) and destructive index (open columns) in mice killed 6 h after treatment with PBS, active caspase-3, Chariot, Chariot and active caspase-3, a mixture of Chariot, active caspase-3, and DEVD-CHO, or nodularin. **P < 0.01 versus PBS control; P < 0.01 versus active caspase-3 and Chariot (n = 4 for each experimental group).

View larger version (39K): [in this window] [in a new window]   Figure 3. Hematoxylin and eosin–stained lung tissue sections from mice killed before (A) and at 2 h (B), 6 h (C), 1 d (D), 7 d (E), and 15 d (F) after treatment with active caspase-3 and Chariot (original magnification: x100). Airspace enlargement is evident as early as 2 h after treatment and persisted for at least 15 d. Similar lung histologies were observed on Day 3 (photographs not shown). Fibrosis or an increase in inflammatory cell infiltration was not noted at any of the time points. These photomicrographs are representative of the results obtained in a minimum of three animals for each time point.

View larger version (241K): [in this window] [in a new window]   Figure 4. Apoptosis in lung tissue and BAL cells. (A) Lung tissue sections stained with either TUNEL (a, c, and e) or anti-ssDNA immunostaining (b, d, and f), and counterstained with hematoxylin. The lungs of mice killed 2 h after treatment with active caspase-3 and Chariot (a and b) or nodularin (e and f) show positive staining (nuclear brown staining) for DNA strand breaks in the alveolar wall. The lungs of mice treated with either PBS, active caspase-3, Chariot (photographs not shown), or a mixture of active caspase-3, Chariot, and DEVD-CHO (b and d) exhibit negative staining for DNA strand breaks (original magnification: a–f, x200). TUNEL-stained slides (g) were stained for pan-cytokeratin antigen, an epithelial cell marker, and exactly the same portion on the slide was photographed (h). TUNEL-positive cells (nuclear brown staining; g) are positively immunostained for pan-cytokeratin (cytoplasmic blue staining; h) in lungs of mice treated with active caspase-3 and Chariot (original magnification: x400). These photomicrographs are representative of the results obtained in three animals for each group. (B) BAL cell preparations from mice 2 h after treatment with active caspase-3 and Chariot. (a and b) May-Grünwald-Giemsa staining showing apoptotic cells, some of which have cilia (arrows). (c, d, and e) Modified Papanicolaou staining showing apoptosis of type II cells. The type II inclusions are visualized as small cytoplasmic particles. (d) An apoptotic type II cell that is probably undergoing phagocytosis by an macrophage. (e) An apoptotic body with type II inclusions. (f) Double-staining with TUNEL and pan-cytokeratin antigen. Apoptotic cells with TUNEL-positive fragmented nuclei show specific staining for pan-cytokeratin (cytoplasmic blue staining). The white arrow shows a macrophage that is negative for both TUNEL and the pan-cytokeratin antigen. These photomicrograph are representative of the results obtained in three animals. Original magnification: x1,000.

Increased Solubilized Elastin Products and Elastolytic Activity in BAL Samples after Treatment with Active Caspase-3 and Chariot
Along with the loss of elastin layers within the alveolar walls, an increased level of solubilized elastin products was detected in BAL samples from mice treated with active caspase-3 and Chariot, as compared with control mice (Figure 5). BAL samples obtained from mice treated with active caspase-3 and Chariot showed enhanced elastolytic activity at pH 5.5, as measured using DQ elastin, an elastin labeled with quenched BODIPY FL dye (Molecular Probes), or insoluble FITC-conjugated elastin as a substrate (Figures 5A and 5B). BAL samples obtained 1 h after treatment had the highest level of elastolytic activity, and the elastolytic activity disappeared in samples obtained 6 h after treatment (Figure 5C). To localize the source of elastolytic activity, in situ elastin zymography was performed using the BAL cell preparations obtained from these mice. Significant levels of DQ elastin digestion were observed in apoptotic cells, but not in normal cells in the presence of acidic environment (pH 5.5) (Figures 6Ac and 6Ad). The digestion of DQ elastin in apoptotic cells was not inhibited by the broad caspase inhibitor ZVAD-fmk (Figure 6Ae), suggesting that proteases other than caspases may mediate the digestion of elastin in apoptotic cells. In contrast, significant levels of elastolytic activities were not detected in BAL cell and fluid samples at pH 7.4 (Figures 5A, 5B, 6Aa, and 6Ab). Because most of the apoptotic cells seen in the BAL cell preparations were epithelial cells, as described previously, epithelial cells undergoing apoptosis may express elastase(s) that require an acidic environment for their activity. The high autofluorescence of tissue collagens and elastin prevented the specific fluorescence of DQ elastin digestion from being observed in frozen lung tissue sections (photographs not shown).

View larger version (21K): [in this window] [in a new window]   Figure 5. Quantitative analysis of elastolytic activity in concentrated BAL fluid samples. BAL fluid was obtained from mice 1 h (A, B, and C), 2 h (C), and 6 h (C) after treatment with PBS, caspase-3, Chariot, or active caspase-3 and Chariot. The fluid samples were concentrated 10x and elastolytic activity was assessed using soluble DQ elastin (A) or insoluble FITC-conjugated elastin (B and C) as a substrate. *P < 0.05 versus PBS-treated mice; P < 0.05 versus pretreated mice (n = a minimum of three for each group at each time point).

View larger version (146K): [in this window] [in a new window]   Figure 6. In situ elastin zymogram. (A) BAL cells obtained from mice 2 h after treatment with active caspase-3 and Chariot were incubated with DQ elastin at pH 7.4 (a and b) or pH 5.5 (c, d, and e) and counterstained with Hoechst 33342. Apoptotic cells with condensed or fragmented nuclei (blue fluorescence) show diffuse green fluorescence throughout the cell at pH 5.5 (d) but not at pH 7.4 (b), indicating the specific degradation of DQ elastin. The degradation of DQ elastin in apoptotic cells is not inhibited in the presence of ZVAD-fmk (e). In contrast, nonapoptotic cells, presumably alveolar macrophages, show a minimum level of green fluorescence, with an occasional punctuated pattern (a and c). These photomicrographs are representative of the results obtained in three animals. Original magnification: x1,000. (B) Rat primary type II cells plated on the layer of DQ elastin in DMEM (pH 7.4) were treated with active caspase-3 and Chariot in 8% DMSO and counterstained with Hoechst 33342. Apoptotic cells (b) show diffuse green fluorescence, indicating the degradation of DQ elastin. In contrast, normal cells treated with 8% DMSO alone (a) do not exhibit green fluorescence. These photomicrographs are representative of the results obtained in three experiments.

View larger version (53K): [in this window] [in a new window]   Figure 7. The caspases are not elastases. Human recombinant caspases-1, 2, 3, 6, 7, 8, 9, 10 (20 U/ml each; MBL) or purified porcine pancreatic elastase (0.1, 0.5, 1 U/ml; Sigma) were incubated with insoluble FITC-conjugated elastin (A), DQ elastin (B), or k-elastin (C) at pH 7.4, or with DQ elastin (D) at pH 5.5. Elastin degradation was assessed by spectrofluorometry (A, B, and D) or SDS-PAGE followed by silver staining (C). (A, B, and D) **P < 0.01 versus the fluorescence intensity of control experiments (n = 4 for each experimental group). In C, k-elastin migrates as a high molecular weight species, a portion of which barely enters the separating compartment of the 20% gel, with the remainder trapped in the stacking compartment. When k-elastin was incubated with pancreatic elastase (1 U/ml), the substrate was extensively degraded without the accumulation of distinct identifiable peptide products (12).

	Discussion

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Although the pathogenesis of pulmonary emphysema is still unknown, the importance of apoptosis has recently come to light. Clinical studies, including our own, have shown higher levels of apoptosis in alveolar wall cells (endothelial or epithelial) of emphysematous lungs than in control lungs (6, 7; N. Yokohori, unpublished observations). A simple explanation for these clinical observations is that destruction of the basement membrane may cause the loss of cell–extracellular matrix (ECM) attachments and apoptosis of surrounding cells. However, a recent study has shown that chronic treatment of rats with the vascular endothelial growth factor receptor-2 inhibitor SU5416 causes endothelial apoptosis followed by airspace enlargement (4). Although this study strongly suggests the pathogenetic role of apoptosis in emphysema, it also generates questions, including whether the chronic vascular endothelial growth factor receptor-2 inhibition can cause various effects other than endothelial apoptosis (5). Another study has shown that liposomal delivery of the hypoxia-inducible factor 1–responsive gene, RTP801, to mouse lungs caused alveolar wall apoptosis, although whether emphysema occurred is unclear (14). We extended the framework of these observations using a novel approach that models the progression of emphysema, and we confirmed that apoptosis can lead to emphysematous changes. In our animal model, the distribution of emphysematous changes was focal as a result of intratracheal injection of the proapoptotic reagent, in contrast to a previous model of diffuse emphysema induced by apoptosis (4). Importantly, the induction of emphysematous changes in our model is very rapid, independent of inflammatory cells, and persistent.

In contrast to our animal model, human emphysema is a chronic progressive disease. However, many chronic progressive diseases occur as a result of repeating the acute process of the disease. This is exactly true for emphysema, which is believed to occur as a result of repeated acute lung injury, presumably caused by an excess of elastases. Moreover, clinical findings of increased alveolar wall apoptosis in human emphysema (6, 7) indicate that apoptosis occurs repeatedly in human emphysema, and also that repeated apoptosis may lead to the development of emphysema.

Airspace enlargement in our animal model of emphysema was evident for at least 15 d after treatment with active caspase-3 and Chariot (Figure 3F). However, the increase in mean chord length was reduced by 60% from 6 h to 15 d, suggesting that induction of emphysema by alveolar wall apoptosis may be partially reversible. Future studies are needed to address this issue.

The traditional hypothesis for the pathogenesis of emphysema is that cigarette smoke results in the accumulation of inflammatory cells that release proteases. These proteases disrupt ECM, including elastin, and this is followed by the loss of cell–ECM attachments, leading to apoptosis and the loss of alveolar units (5). Our study reinforces an alternative hypothesis whereby apoptosis is the primary event that leads to the loss of ECM components and subsequent loss of alveolar units (5). In emphysematous lungs, epithelial apoptosis could be induced by various stimuli, including oxidative stress, proteases, and infiltrating cytotoxic CD8+ T cells (15–17). Regardless of which of these stimuli is active, subsequent epithelial apoptosis is likely to result in the loss of alveolar units.

In the present study, enhanced elastolytic activity and increased solubilized elastin products in BAL samples suggest that our model may operate along the lines of the elastase/antielastase imbalance theory (3). The noninflammatory aspect of our model further suggests that the proteases released by apoptotic epithelial cells may be sufficient to cause elastin destruction. This notion is supported by our in vitro findings, which showed the destruction of elastin layer beneath type II cells as they were undergoing apoptosis.

We did not investigate which protease(s) mediate the elastin destruction in the present study. However, the optimal acidic pH of the protease(s) makes lysosomal cathepsins the logical candidates. Many studies have implicated lysosomal cathepsins in the pericellular degradation of the ECM, including elastin (18–21), as well as in intracellular apoptotic processes (18, 19, 22). Furthermore, alveolar epithelial cells constitutively express enzymatically active, mature forms of cathepsins B, K, and L (23), some of which (i.e., cathepsins K and L) have strong elastolytic activities (24, 25). Despite this evidence favoring the role of cathepsins, BAL elastolytic activity was inhibited by only 30% in the presence of the broad cathepsin inhibitor E64 (K. Aoshiba, unpublished observations). Thus, we could not determine unequivocally that elastase(s) mediated apoptosis-induced emphysema. We speculate that multiple classes of elastases, including cathepsins and unknown caspases, may be involved in the destruction of the ECM.

Our animal model of emphysema did not exhibit alveolar and airway inflammation, such as that seen in human emphysema. However, this does not mean that emphysema occurs without inflammation. The potential roles of the neutrophils, macrophages, and T cells that are traditionally associated with the pathogenesis of emphysema should not be excluded. These cells may initiate lung damage and ECM destruction through mechanisms involving oxidants, proteases, and cell-mediated cytotoxicity, thereby leading to alveolar cell apoptosis, the promotion of ECM destruction, and further apoptosis. Thus, emphysema may be enhanced by repeated induction of apoptosis or by recruitment of inflammatory cells. In conclusion, the present study provides strong evidence for the ability of apoptosis to cause emphysematous changes, suggesting that apoptosis may play a part in the mechanism of emphysema.

	Acknowledgments

 
This work was supported by Grant-in Aid for Scientific Research #12670580 from the Ministry of Education, Science, and Culture, Japan, and by the Respiratory Failure Research Group in Japan. The authors are very grateful to Masayuki Shino and Yoshimi Sugimura for their excellent technical assistance.

Received in original form June 18, 2002

Received in final form October 24, 2002

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