Departments of Pathology and Pediatrics, University of Geneva Medical School, 1211 Geneva 4, Switzerland
Submitted 6 June 2003 ; accepted in final form 21 September 2003
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ABSTRACT |
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mice; apoptosis; mitochondria; type II epithelial cells; Bax
During oxygen exposure mitochondria actively participate in increasing the production of intracellular ROS (19). The mitochondrion plays a pivotal role in the regulation of cell death responses, since it maintains the cellular levels of ATP and is able to releases death-promoting factors, such as cytochrome c, when cells are exposed to proapoptotic signals. Although several death stimuli induce the mitochondrial release of cytochrome c, the molecular mechanisms of this process are not completely understood (16, 30). It has been shown that the activation and translocation from the cytosol to the mitochondrial membrane of the proapoptotic members of the Bcl-2 family, such as Bax, lead to the formation of channels through which cytochrome c is released without alteration of the morphology of mitochondria (3, 30). Alternatively, strong cell death signals or even Bax itself can affect directly the permeability of the mitochondrial membrane and induce the opening of the high-conductance channel permeability transition pore (PTP) (30, 47). Irreversible PTP opening induces large amplitude mitochondrial swelling, outer membrane break, and subsequent release of cytochrome c (24, 37). Once into the cytosol, cytochrome c is able to trigger the apoptotic signaling cascade by activating the caspase-9 and downstream effector caspases, in particular caspase-3, responsible for the nuclear fragmentation and cell death (23).
Morphological studies have described that mitochondria from hyperoxia-injured alveolar cells undergo structural changes such as swelling and crista disorganization (27, 32). Such alterations have been correlated with perturbation of the mitochondrial membrane permeability in vitro (1). However, a direct relationship between mitochondrial morphological changes, cytochrome c release, and apoptotic/necrotic signaling during hyperoxia in vivo has not been established yet.
In the present study, we explored the role of mitochondria-dependent cell death signaling in the pathogenesis of hyperoxia-induced lung injury. Exposure of mice to 100% oxygen induced the release of cytochrome c in high amounts from the mitochondria into the cytosol. This was accompanied by Bax translocation to the mitochondrial membrane. Besides the initiation of the mitochondrial apoptotic machinery, the active form of caspase-3 was not detectable. To investigate whether cytochrome c release was dependent on mitochondrial permeability transition, we injected mice with cyclosporin A (CsA), a specific blocker of PTP opening, before and during oxygen exposure (38, 48). CsA treatment significantly prevented cytochrome c release from mitochondria into the cytoplasm of alveolar cells. Electron microscopy (EM) analysis of alveolar type II cells, where mitochondria are abundant and easily recognizable, showed that mitochondria from CsA-treated mice were more electron dense and less swollen and had less disorganization of cristae compared with vehicle-treated mice. Concomitantly, CsA treatment ameliorated hyperoxia-induced lung damage, as shown by macroscopical lung injury score and lung weight. These results suggest that preventing from mitochondrial structure alteration cytochrome c release in alveolar epithelial cells can protect from hyperoxia-induced lung injury.
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MATERIALS AND METHODS |
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Hyperoxia exposure and in vivo treatment. Mice were placed in a sealed Plexiglas chamber and exposed to 100% O2 for 72 h, as described (5). Control mice were exposed to room air under the same conditions. CsA was injected intraperitoneally at the dose of 25 mg·kg-1·day-1 (Sandimmune, Novartis), diluted in vehicle, composed of 650 mg/ml cremaphor (Sigma-Aldrich, St. Louis, MO), 250 mg/ml ETOH, and 0.9% NaCl. The untreated group was injected with the vehicle only. This dosage regimen was shown to be effective in previous studies (33). The treatment with CsA or vehicle was started 3 days before oxygen exposure and continued during the time of air or 100% O2 exposure. After death, lung damage was macroscopically evaluated, according to the extension of hemorrhages, as described by Christofidou-Solomidou et al. (11), and a score ranging from 0 (no lesions) to 5 (complete hemorrhagic lung) was given by three experienced independent examiners. We determined pulmonary edema by measuring right wet lung weight, as described previously (5). The right lung was immediately frozen in liquid nitrogen and stored at -80°C until further analysis. The left lung was fixed and stored for histological analysis. Bronchoalveolar lavage (BAL) was performed as described (6). BAL fluid was recovered immediately on ice under negative hydrostatic pressure. After centrifugation, cells were counted, and protein concentration was determined.
All study protocols were approved by the local ethical committee on animal experiments (Office Vétérinaire Cantonal of Geneva).
Light and electron microscopy. Left lungs were fixed intratracheally with 4% paraformaldehyde in phosphate buffer, with an hydrostatic pressure of 20 cmH2O, and embedded in paraffin. Sections (4 µm) were stained with hematoxylin and eosin for histological evaluation or processed for immunohistochemistry (IHC). For EM studies, lungs were fixed by the intratracheal instillation of 2% glutaraldehyde (in 0.1 M cacodylate buffer) and processed as described (6). Sections were embedded in Epon and then examined with a Philips CM10 (Philips, Zurich, Switzerland) 400 electron microscope at 70 kV.
IHC. Paraffin-embedded lung tissue sections (5 µm) were mounted on slides pretreated with 3-aminopropylxylane (Merck, Darmstadt, Germany), baked overnight at 55°C, deparaffinized, and rehydrated. Sections were cooked 3x 5 min in microwave to facilitate the access of the antibody. After cooling, samples were blocked with 5% bovine serum albumin in Tris-buffered solution and incubated overnight with an anti-cytochrome c monoclonal antibody (1:250-1:500, clone 7H8.2C12; BD Pharmingen, Franklin Lakes, NJ) or with an anti-cleaved-caspase-3 rabbit polyclonal antibody (1:100, 9661-S; Cell Signaling Technology, Beverly, MA). As secondary antibody, a biotinylated anti-mouse IgG, Fc-specific antibody (Jackson Laboratories, San Diego, CA) or a biotinylated anti-rabbit total Ig antibody (Santa Cruz Biotechnology, Santa Cruz, CA) was used and then labeled with streptavidinbiotin-alkaline phosphatase complex (A+B kit; Vector Laboratories, Burlingame, CA). The phosphatase reaction was revealed with the fast red substrate system (DaKo, Carpinteria, CA), and slides were counterstained with Hemalun. Normal mouse or rabbit IgG was used instead of the primary antibody as negative control for nonspecific binding. To quantify the mitochondrial content of cytochrome c, a score was established as follows: 1) the number of lung parenchymal positive cells was counted at x100 magnification in 25 fields for each lung section (2-3 different sections per animal, 7-8 mice in each group); 2) for each positive cell a range of positivity was given between 1 and 4, 1 corresponding to <25% of the cell surface covered by the granular pattern, 2 between 25 and 50%, 3 between 50 and 75%, and 4 >75%. The total score was expressed as the number of positive cells x the range of positivity. This value was divided by the number of positive cells to calculate a mean score per positive cell (mean ± SD). This evaluation was established independently by three different investigators.
Cytosolic and mitochondrial subcellular fractionations. Subcellular fractions were prepared from total lungs extracts, as described by Bossy-Wetzel and Green (9) with some modifications (22). Briefly, frozen lungs were minced into small pieces with a scalpel and suspended in 15 volumes of cold buffer (250 mM sucrose, 20 mM HEPES-KOH, pH 7.4, 10 mM KCl, 1.5 mM Na-EGTA, 1.5 mM Na-EDTA, 1.0 mM MgCl2, 1.0 mM DTT, and protease inhibitors). Tissue samples were then homogenized with a glass Dounce homogenizer and a tight Teflon pestle. Homogenates were first centrifuged (800 g for 10 min at 4°C) to eliminate nuclei and debris. The postnuclear supernatant was filtered through gauze and centrifuged at 9,500 g for 15 min at 4°C. Resulting pellets were designated as mitochondria-enriched heavy membrane fractions (HMF), according to Dubrez et al. (14), while supernatants were further ultracentrifuged at 100,000 g for 1 h, at 4°C, to obtain cytosolic fractions. To verify the integrity of mitochondrial membranes and exclude cytosol contamination by mitochondrial proteins, we blotted each fraction with a mouse monoclonal anti-cytochrome oxidase subunit IV (COX, A-6431; Molecular Probes, Eugene, OR) as a mitochondrial marker. A rabbit polyclonal antibody anti-actin (AL-20, a kind gift from the laboratory of G. Gabbiani) was used as a control for protein loading.
Western blot analysis. Samples were analyzed for protein concentration by Bio-Rad DC protein assay kit (ref. 500-0111; Bio-Rad Laboratories, Hercules, CA). To analyze the localization of cytochrome c, we electrophoresed 15 µg of total proteins from the cytosolic and mitochondrial fractions on a 12% SDS-polyacrylamide gel. We used 70 µg of protein for the analysis of Bax. All gels were blotted to nitrocellulose membranes (Amersham International, Amersham, UK), except for detection of COX, where samples were transferred to a polyvinylidene difluoride membrane (porablot; Macherey-Nagel, Düren, Germany). Membranes were blocked overnight in TBST buffer (0.2 M Tris, pH 7.6, 1.5 M NaCl, and 0.1% Tween 20) and 5% milk and incubated with the following antibodies: rabbit polyclonal anti-cytochrome c antibody (1:200 dilution, sc-7159; Santa Cruz Biotechnology), mouse monoclonal anti-COX antibody (1:500 dilution), rabbit polyclonal anti-actin antibody (1:2,000, AL-20; gift from G. Gabbiani's laboratory), rabbit polyclonal anti-Bax antibody (1:200 dilution, sc-493; Santa Cruz Biotechnology), and rabbit polyclonal anti-cleaved caspase-3 (1:1,000, 9661-S; Cell Signaling Technology). Horseradish peroxidase-conjugated anti-mouse and anti-rabbit (1:3,000 dilution, Bio-Rad Laboratories) were used as secondary antibodies. Bands were visualized with a chemiluminescent substrate (Amersham International). Scanning and quantification of signal intensity were performed on subsaturated films by with the Imagequant software.
Statistical analysis. For all parameters measured, the values for all animals in different groups were averaged, and the SD of the mean was calculated. The significance of differences between the values of the groups was determined with unpaired Student's t-test. Where appropriate, two-way ANOVA with multiple comparisons followed by unpaired t-test was used. Significance levels were set at P < 0.05.
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RESULTS |
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As a complementary approach, we also analyzed the cellular distribution of cytochrome c by WB. In total lung extracts, the content of cytochrome c was similar in air-breathing and oxygen-exposed mice (Fig. 2A). However, its intracellular distribution was significantly affected by hyperoxia. In air-breathing mice, cytochrome c was almost undetectable in lung cytosolic fractions, whereas it was present in mitochondria-enriched HMF (Fig. 2B). During oxygen exposure, cytochrome c appeared in the cytosol at 48 h, and the signal was even higher at 72 h (Fig. 2B). Quantification of four different experiments (n = 11 animals) showed that the cytosolic level of cytochrome c was significantly higher at 72 h of exposure compared with control lungs (P < 0.001; Fig. 2C, right). In some experiments, the release of cytochrome c into the cytosol was accompanied with a decrease, although not significant, of the signal in HMF fraction (Fig. 2C, left). These results demonstrate that hyperoxia induces the release of cytochrome c from mitochondria into cytosol of alveolar cells.
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Hyperoxia-induced release of cytochrome c is associated with Bax translocation to the mitochondria without caspase-3 activation. It has been described that during apoptosis Bax protein translocates from the cytosol into the mitochondria, inducing the release of cytochrome c (3). Therefore, we analyzed whether cytochrome c release was accompanied by Bax translocation. As shown by WB, Bax protein content increased significantly in HMF fractions of hyperoxia-exposed compared with control lungs (Fig. 3, A and B, left, P < 0.05), suggesting a translocation of the proapoptotic protein to mitochondria. However, the cytosolic content of Bax was not significantly reduced by oxygen exposure (Fig. 3, A and B, right).
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To further investigate whether cytochrome c release was able to trigger caspase activation, we analyzed the expression of the cleaved (active) form of caspase-3, which is one of the major downstream effector caspases. As shown in Fig. 4, no positive signal was detected in the cytosol from air- and hyperoxia-exposed lung extracts, even when 150 µg of proteins were loaded. Similarly, no signal for cleaved caspase-3 was detected on hyperoxic lung sections by IHC (not shown).
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CsA treatment prevents hyperoxia-induced cytochrome c release and mitochondrial damage. Morphological studies have shown marked structural changes in mitochondria from hyperoxia-exposed alveolar cells (1, 27). Therefore, we hypothesized that hyperoxia could induce the opening of the mitochondrial PTP, leading to mitochondria swelling, membrane disruption, and secondary cytochrome c release. To verify this hypothesis, we explored whether CsA treatment of mice exposed to hyperoxia could prevent mitochondrial damage and cytochrome c release.
As analyzed by IHC, CsA treatment significantly prevented the loss of mitochondrial cytochrome c staining observed in vehicle-treated mice exposed to hyperoxia (compare Fig. 5, B and D vs. A and C). Indeed, the number of positive cells in 25 fields (not shown) and the mean of positive granules per cell were significantly higher in CsA-treated animals compared with vehicle-treated animals exposed to hyperoxia (mean score/cell: 2.0 ± 0.4 in CsA-treated vs. 1.6 ± 0.19 in vehicletreated group exposed to hyperoxia, P < 0.01, Fig. 5E). To note, in air-breathing mice, treatment with vehicle alone or CsA did not affect the strong intracellular staining pattern (not shown) that was observed in untreated animals (see Figs. 1, C and E, and 5E).
We then analyzed the mitochondrial morphology in lungs of vehicle- and CsA-treated animals by EM. In air-breathing mice, a high number of small, electron-dense mitochondria were easily recognized in alveolar type II cells (Fig. 6, A and D). Major mitochondrial alterations were observed during hyperoxia in these cells, according to previous morphological studies in hyperoxia-exposed rats (27). After 72 h of oxygen exposure, almost all mitochondria exhibited marked swelling, diminished matrix density, and disorganized cristae (Fig. 6, B and E). Mitochondria of alveolar cells from CsA-treated mice were in large part compact and more electron dense compared with those from vehicle-treated mice (Fig. 6, C and F, arrows). However, mitochondria with a large, swollen aspect and disrupted cristae were also found within the same cell (Fig. 6, C and F, arrowhead). Together, these observations suggest that CsA partially prevented, at least in a subset population of alveolar cells, hyperoxia-induced mitochondrial damage and consequent cytochrome c release.
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CsA treatment reduces hyperoxia-induced lung damage. Finally, we analyzed whether the effects on mitochondria observed with CsA treatment were associated with lung damage protection. Macroscopical examination of the lungs showed significantly lower extent of hemorrhages in CsA-treated compared with vehicle-treated mice (Fig. 7, compare C with B, and Table 1). Accordingly, the right lung weight and BAL protein content were significantly reduced in the CsA-treated group compared with the vehicle-treated group (Table 1). However, the number of cells recovered by BAL was similar in both groups (Table 1). Lung histology of hyperoxia-exposed CsA-treated mice showed some extent of alveolar septa thickening but less edema and hyaline membrane formation compared with vehicle-treated mice (Fig. 7, compare C vs. B). CsA treatment in itself affected neither the macroscopical and microscopical aspect (not shown) nor the number of cells recovered by BAL from air-exposed mice (Table 1).
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DISCUSSION |
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Mitochondria play a central role in regulating cell death, as they control calcium homeostasis and intracellular oxidants levels (18, 47), produce ATP, and release apoptogenic factors (30). In the present study we analyzed the mitochondria-dependent cell death pathway and in particular the intracellular distribution of cytochrome c in lungs of mice exposed to air or to hyperoxia. The granular staining seen by IHC in control lungs was highly suggestive of mitochondrial protein distribution, in agreement with previous descriptions (22). The staining was mainly evident in large alveolar cells located at septal junctions. Their localization and morphology were compatible with epithelial type II cells, which are known to have high mitochondria content. After 72 h of hyperoxia, the cytochrome c-specific staining of these cells lost its granular aspect, suggesting that cytochrome c spilled out of the mitochondria into the cytosol. By biochemical analysis, we confirmed that mitochondrial cytochrome c was released into the cytosol in a small amount after 48 h of oxygen exposure and in a significantly higher amount after 72 h. The presence of cytochrome c in cytosolic extracts during hyperoxia was not always correlated with a detectable decrease in its mitochondrial quantity. However, since cytochrome c is normally absent in the cytoplasm, it is far easier to detect its release in this compartment than its evasion from mitochondria where cytochrome c is very abundant. Indeed, equivalent amounts of proteins for cytosolic and mitochondrial extracts (15 µg) were loaded onto the gels, whereas in most reports higher amounts of cytosol were used compared with mitochondria [i.e., 30 µg of cytosol vs. 4 µg of mitochondrial extracts in (22)].
To decipher the mechanism leading to cytochrome c release, we analyzed the modulation of Bax protein in our experimental model. In vivo oxygen exposure has been associated with a significant increase in Bax lung mRNA, without any modifi-cation of the protein expression (7, 36). The cellular fractionation approach allowed us to determine that Bax was significantly increased in mitochondrial fractions after 72 h of hyperoxia, indicating the activation and translocation of this protein from the cytosol to the mitochondria. These results suggest a proapoptotic role for Bax in hyperoxia and are in agreement with a previous report showing that Bak/bax(-/-) embryonic fibroblasts were more resistant to cell death upon oxygen exposure (10). Surprisingly, the translocation of Bax was not evident at 48 h of oxygen exposure, as predicted by the assumption that Bax would induce cytochrome c release (30). Two different hypotheses could explain the asynchrony between Bax translocation and cytochrome c release: 1) the anti-Bax specific antibody used was not sensitive enough, precluding the detection of a small increase in mitochondria-associated Bax at 48 h; 2) Bax translocation may be restricted to some mitochondria, whereas others could have their membranes damaged directly by the high concentration of free radicals with subsequent release of cytochrome c independently of Bax translocation. Unfortunately, we could not reliably determine the subcellular localization of Bax by IHC and hence could not define whether it was translocated in the same cells as those in which cytochrome c was released.
A recent report demonstrated that the activation of the caspase-8/Bid signaling pathway was involved in the apoptotic response during hyperoxia, since gene disruption of Bid protected against cell death, both in vivo and in vitro (44). It has been proposed that, once activated, Bid can assist the conformational change of Bax into its active form leading to cytochrome c release (13, 44). However, Bax insertion to the mitochondrial membrane has been observed in Bid-null fibroblasts, suggesting an alternative Bid-independent pathway in Bax activation (39). It cannot be excluded that other BH3-only proteins, such as Bim or Bad, or other upstream molecules, might induce Bax translocation.
Cytochrome c release into the cytosol has been associated with caspase signaling activation and apoptotic cell death in several human diseases, such as heart failure, neurodegenerative diseases, and traumatic brain injury (35, 40, 41). We were unable to detect any active caspase-3 during hyperoxia, supporting our previous results, where no significant increase in caspase activity was measured in total lung extracts. In addition, the administration of specific caspase inhibitors (ZVAD) did not prevent mouse lung injury (7). These data are in contrast with a recent report from Zhang et al. (49) in which activated caspase-3 was detected by IHC in lung epithelial and bronchial cells of mice exposed to hyperoxia. This discordance could account for the different sensitivity of the antibodies and the different technical approaches used in either study. Nevertheless, these results leave open the question whether, besides the possible activation of caspase-3 in alveolar cells, hyperoxia-induced lung injury is dependent on the classical caspase-3-mediated apoptotic cascade. Indeed, a clear-cut answer might be provided by studying the survival of caspase-3(-/-) mice in hyperoxia (29). Moreover, several in vitro results indicate that the signaling pathways used to initiate cell death are not predicting the final outcome of cell necrosis or apoptosis. For instance, activation of caspase-9 upon cytochrome c release and phenotypic features of apoptosis have been shown in cultured Rat-1 cells and endothelial cells after 40-72 h of oxygen exposure (10, 25). Conversely, A549 cells exposed to 100% oxygen, although presenting caspase-8 and caspase-9 activation, did not undergo apoptotic cell death but presented morphological features of necrosis (26, 44). Our previous results based on EM analysis suggest that most of the apoptotic features occurred in endothelial cells, although some alveolar cells presented overlapping features of apoptosis and necrosis (7). This indicates that hyperoxia-induced epithelial cell death might involve the initiation of a caspase-mediated, mitochondria-dependent apoptotic pathway, despite a final outcome of cellular necrosis (44).
CsA treatment was previously described to be effective in protecting lungs from hyperoxia-induced injury (33). However, no mechanism was proposed for its beneficial effect. Accordingly, our study shows that CsA attenuated lung damage, as evaluated by different injury markers such as macroscopical injury score, lung weight, and BAL protein content. Importantly, such a protective effect of CsA correlated with a preserved morphological aspect of mitochondria in type II cells and with a decrease of cytochrome c release, as demonstrated by IHC. These results are in agreement with previous reports where CsA prevented irreversible damage to mitochondria (21) and cytochrome c release in several pathological conditions (34, 46). It is interesting to note that alveolar type II cells from CsA-treated mice presented a mixed population of still electron-dense and swollen mitochondria, suggesting that the number of functional mitochondria and the level of ATP are crucial for cell survival and lung damage.
Morphological mitochondria alterations under hyperoxia have been mainly studied in type II and type II-like epithelial cells (1, 12, 27). These cells have been shown to play a critical role in the maintenance of the alveolar space, since they are more resistant to oxidative stress compared with other alveolar cells and are the precursors of type I cells (31). Adaptation to hyperoxia and survival after lung injury may depend on the capacity of alveolar progenitor cells to proliferate and reestablish the integrity of the alveolar epithelium. For this reason, type II cells have been targeted for the overexpression of protecting molecules, to prevent hyperoxia-induced lung injury (43). For instance, overexpression of mitochondrial antioxidant enzyme Mn-superoxide dismutase in surfactant protein (SP)C-expressing cells conferred protection from oxygen-induced lung damage in mice and correlated with prevention of mitochondrial injury and preservation of ATP content in those cells (45). Accordingly, our results emphasize the importance of epithelial type II cells in maintaining critical epithelial function.
However, it cannot be excluded that CsA would also prevent cytochrome c release and mitochondrial damage in endothelial and epithelial type I cells. Alternatively, type II cells could indirectly influence the sensitivity to oxygen of endothelial and epithelial type I cells. For example, a deficiency of SP-B, a protein specifically synthesized by type II cells, sensitized mice to hyperoxia. Increased lung permeability and protein content in BAL were observed in these mice, suggesting that epithelial type II cells affect the physiological function of endothelial and epithelial type I cells (42). Additional evidence of cross talk between type II cells and endothelial cells has been provided by in vitro studies, where A549 and primary type II cells were able to modulate transendothelial migration of leukocytes under specific stimuli (15, 28). It is likely that these cells may also contribute to changes of endothelial function in vivo and that preserving the integrity of type II cells might exert a protective effect on alveolo-capillary barrier.
In conclusion, this report indicates that CsA, by preventing mitochondrial alterations and cytochrome c release in type II cells, confers protection against hyperoxia-mediated lung injury.
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ACKNOWLEDGMENTS |
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GRANTS
This work was supported by Fonds National de la Recherche Scientifique Grant 3200-067865.02 and by the Wölfermann-Naügele, Novartis, and Lancardis Foundations.
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FOOTNOTES |
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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.
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REFERENCES |
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