Glucocorticoid-induced contractility and F-actin content of human lung fibroblasts in three-dimensional culture

Hiroyuki Miki, Tadashi Mio, Sonoko Nagai, Yuma Hoshino, Takeo Tsutsumi, Takeshi Mikuniya, and Takateru Izumi

Department of Respiratory Medicine, Graduate School of Medicine, Kyoto University, Kyoto 606-8507, Japan


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Fibroblast contractility plays a useful role in the wound healing process but contributes to architectural distortion in the lungs. Glucocorticoids (GCs) have been reported to reduce dermal fibroblast contractility, which may result in delaying wound healing, but the effects on lung fibroblasts are unknown. In this study, we examined how human lung fibroblast contractility is altered in the presence of GCs. Lung fibroblast cell lines (n = 5) were established from normal parts of surgically resected lung tissue. The effects of GCs on contractility were investigated with a type I collagen gel contraction assay. Filamentous actin (F-actin) content was detected by confocal microscopy and measured with a fluorescent phalloidin binding assay. GCs augmented fibroblast contraction in a concentration-dependent manner, with an approximate EC50 of 1.8 × 10-8 M, whereas other steroid derivatives had no effects. GC contractility needed de novo protein synthesis. The GC-induced increase in contractility was found to be consistent with an increase in F-actin content. In conclusion, lung fibroblast contractility was enhanced with GCs through an upregulation of lung fibroblast F-actin.

human lung fibroblast contractility; gel contraction assay; filamentous actin; fluorescent phalloidin binding assay


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

FIBROBLASTS ARE CONSIDERED to be important cells that can migrate into sites of inflammation in tissue healing (20, 22) and produce a variety of extracellular matrix proteins that contribute to the development of fibrosis with or without remodeling (9, 31). In the lung, fibroblast proliferation in excess of repair affects alveolar architecture, and fibroblast contractility could contribute to the remodeling process that occurs in pulmonary fibrosis (14, 33).

Tissue contraction is a part of normal wound healing. Tissues undergoing contraction can generate tension (1, 17). The generation of tension is believed to be a cell-mediated event; however, how cells generate the forces resulting in tension during tissue contraction is unclear. Fibroblasts have the potential to generate tension. They can exert tension on a flexible silicone rubber substratum (15) and can also generate tension when cultured within collagen gel (3, 12, 28). Actin filaments with associated myosin and actin binding proteins are present in tension-generating fibroblasts. Filamentous actin (F-actin) has been proposed to be organized in response to cell contraction (5, 38) and may participate in generating the forces responsible for continued development and maintenance of tension (12, 19, 23).

It has been reported that glucocorticoids (GCs) reduce dermal fibroblast-mediated collagen gel contraction (8) and that this reduction may play a role in the delay in the wound-healing process (13, 24). On the other hand, if fibroblast contractility can be reduced by GCs, a positive beneficial effect might be expected in patients with pulmonary fibrosis. Katzenstein and Fiorelli (21) and Nagai et al. (30) have recognized a heterogeneity of pulmonary fibrosis in patients with interstitial pneumonia (21, 30). Some patients with an acute or subacute type of pulmonary fibrosis show a favorable response to GCs, whereas GCs only show marginal effects in patients with a more chronic type of pulmonary fibrosis such as seen in the pathological pattern of usual interstitial pneumonia. Nagai and colleagues (26, 27) have previously reported that a number of factors can modulate fibroblast-mediated collagen gel contraction.

In this study, we examined how human lung fibroblast contractility is altered in the presence of GCs.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Study population. Five adult patients who underwent a thoracotomy for clinically relevant reasons (4 lung cancers and 1 benign lung tumor) were selected for this study. These patients included 3 men and 2 women with a mean age of 58.4 ± 4.6 yr. None of the patients were current smokers, and none were treated with GCs or immunosuppressants at the time of thoracotomy. No signs or symptoms that suggested infection were present at the time of study. Informed consent was obtained from each patient. The study was approved by the Ethical Committee of the Graduate School of Medicine, Kyoto University (Kyoto, Japan).

Cell cultures. Lung fibroblast cell lines were established from normal parts of surgically resected tissues as previously described (25). The lung tissue specimens were minced into small pieces, washed with Dulbecco's modified Eagle's medium (DMEM; GIBCO BRL, Life Technologies, Grand Island, NY), and plated in 100-mm tissue culture dishes (Iwaki Glass, Chiba, Japan). The lung tissues were then cultured with DMEM supplemented with 10% fetal bovine serum, 50 U/ml of penicillin, 50 µg/ml of streptomycin, and 0.25 µg/ml of Fungizone (GIBCO BRL) at 37°C in an atmosphere of 5% CO2. The medium was replaced twice a week. When the fibroblasts reached confluence, the cells were detached by brief trypsinization (0.05% trypsin and 0.53 mM EDTA · 4Na; GIBCO BRL) and subcultured at a 1:4 ratio of cell suspension to medium. The cells obtained from the fourth to seventh passages were used for the experiments.

We investigated the effect of GCs with a gel contraction assay, a viability assay, immunocytochemistry, and a phalloidin binding assay. The GCs tested included hydrocortisone, methylprednisolone, and dexamethasone (Dex). To examine the specificity of GC activity, we examined the effects of cholesterol, aldosterone, dehydroepiandrosterone, methyltestosterone, progesterone, and estradiol-17beta as controls. All of the steroids used in this study were commercially available (Sigma, St. Louis, MO), and the stock solutions were prepared by dissolving the steroids in absolute ethanol at concentrations 103 M higher than required in the experiments.

Gel contraction assay. Fibroblast contractility was assessed by measuring the changes in the surface area of type I collagen gels mediated by fibroblasts (2, 26, 37). The change in surface area (percent contraction) is expressed as the ratio of the change in surface gel area to the initial surface gel area. The type I collagen solution was prepared from rat tail tendons (10). The final gel constitution was type I collagen (0.75 mg/ml) and a fibroblast cell suspension (2 × 105 cells/ml) in HEPES-buffered DMEM (pH 7.4). Five hundred microliters of collagen gel were made in each well of the tissue culture plate (Iwaki Glass). Five hundred microliters of DMEM supplemented with 2 µg/ml of insulin and 20 µg/ml of transferrin (Sigma) were then overlaid. The fibroblasts suspended in type I collagen gels were cultured for 96 h. The gels were then incubated with various reagents for 48 h. Then the gels were separated from the tissue culture plates with a sterile scalpel and transferred to nontreated petri dishes (Iwaki Glass) containing 5 ml of prewarmed DMEM supplemented with reagents. The gel area was measured with an image scanner connected to a computer running a public domain planimetry application (National Institutes of Health Image version 1.61; available by anonymous FTP from zippy.nimh.nih.gov).

Viability assay. Viability of the fibroblasts was assessed with the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay (Sigma) (29). After culture of the fibroblasts in collagen gels with reagents, MTT was added to each well at a final concentration of 0.5 mg/ml. After incubation for 4 h at 37°C, the overlaid medium was removed, and formazan salt was dissolved by adding 0.5 ml of acid isopropyl alcohol. The aqueous phase was collected, and the optical density was determined with a Multiscan MCC/340 (reference wavelength 630 nm, test wavelength 570 nm; Labsystems, Helsinki, Finland).

Immunocytochemistry. We identified cell strains as fibroblasts by immunodetection of alpha -smooth muscle actin, vimentin, and desmin (35). The cells were fixed on coverslips with 3% paraformaldehyde in phosphate-buffered saline (PBS) containing 2% sucrose and permeabilized in 20 mM HEPES buffer (300 mM sucrose, 0.5% Triton, 50 mM NaCl, and 3 mM MgCl2, pH 7.4), for 3 min at 0°C; aldehyde-induced fluorescence was quenched with 50 mM NH4Cl. After blocking nonspecific binding with 3% (vol/vol) normal serum, we incubated the cells with a monoclonal antibody for anti-alpha -smooth muscle actin (1:400 dilution), vimentin (1:200 dilution) or desmin (1:200 dilution; all from Sigma). Primary antibodies were visualized with rhodamine-labeled anti-mouse IgG or FITC-labeled anti-rabbit IgG (1:100 dilution; DAKO, Glostrup, Denmark). The stained cells were mounted in 90% glycerol in PBS containing 2.5% 1,4-diazabicycol[2.2.2]-octane (Aldrich, Milwaukee, WI). To quantify the number of fibroblasts in each experimental condition, we counted alpha -smooth muscle actin-stained cells and the total number of cells per coverslip stained with 2 µg/ml of propidium iodide in a minimum of three randomly chosen microscopic fields (total cells > 100) with a Zeiss Axiovert 135 inverted microscope and a Zeiss LSM 410 confocal attachment. Photomicrographs were taken with identical contrast and brightness settings.

Phalloidin binding assay. Phalloidin binds tightly to the actin subunits in filaments but not to the monomers so the amount of bound phalloidin reflects the amount of actin filaments (11). Therefore, F-actin can be measured with a fluorescent derivative of phalloidin (6, 18). In our experiments, fibroblasts (2 × 105 cells/ml) within collagen gels were fixed in 4% paraformaldehyde and the cell membranes were permeabilized with 0.2% Triton X-100, and then the cells were incubated for 30 min with rhodamine-labeled phalloidin at a concentration of 2 × 10-7 M. In control experiments, nonspecific binding was determined by incubating samples in rhodamine-phalloidin plus a 10-fold excess of unlabeled phalloidin (2 × 10-6 M; Sigma). After three washes in PBS, the bound phalloidin was extracted by adding 500 µl of 0.1 N NaOH and neutralized with 1.0 M Tris · HCl, pH 7.4. After centrifugation at 13,000 rpm at 4°C for 15 min, the intensity of the rhodamine fluorescence was measured (excitation at 540 nm, emission at 575 nm) with a fluorescence spectrophotometer (model F-3000, Hitachi, Tokyo, Japan). Data are expressed as the intensity of fluorescence after subtraction of nonspecific binding.

Statistics. The results are expressed as means ± SE of no less than three determinations. All of the results were confirmed by repeating the experiments on two separate occasions in triplicate. Analysis of variance was used to assess the differences of means between the treated and control groups, and post hoc analysis with Scheffé's test was used for comparison between any two groups. A P value of <0.05 was considered to be significant.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Effect of Dex on the gel contraction. Dex enhanced fibroblast-mediated collagen gel contraction in a concentration-dependent manner at concentrations ranging from 10-10 to 10-6 M, with an approximate EC50 of 1.8 × 10-8 M (Fig. 1). The maximal effect was observed at a concentration of 10-6 M (48.3% decrease in initial gel area). The gels treated with 10-6 M Dex showed a contraction occurring from 30 to 360 min after release (33.2% decrease in 30 min and 44.2% decrease in 60 min; Fig. 2). Dex augmented the contraction of all of the tested fibroblasts in a concentration-dependent manner at concentrations ranging from 10-9 to 10-7 M (P < 0.01; Fig. 3). No difference between cell lines was detected. The cell viability of fibroblasts treated with 10-5 or 10-4 M Dex as assessed by MTT were lower (82.5 ± 10.2% of control value with 10-5 M Dex and 63.3 ± 12.5% with 10-4 M Dex) than that of cells treated with 10-10 (99.8 ± 5.4%) to 10-6 M (98.6 ± 8.2%).


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Fig. 1.   Effect of dexamethasone (Dex) on fibroblast contractility. Fibroblasts in type I collagen gels were cultured with serum-free DMEM for 96 h to measure contraction. Cells were then incubated with Dex at concentrations ranging from 10-11 to 10-4 M. After incubation for 48 h, gels were released and transferred to tissue culture dishes containing culture medium with the same concentrations of Dex. Areas of gels were measured 1 h after release. Values are means ± SE of triplicate determinations.



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Fig. 2.   Time-dependent contraction of collagen gel. Fibroblasts within gels were cultured with 1 µM Dex (), 1 nM Dex (), or medium alone (open circle ) for 48 h before release. Gels were released and transferred to dishes containing medium with the same stimuli. Areas of gels were measured at indicated times from 0 to 720 min after release. Values are means ± SE of triplicate determinations.



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Fig. 3.   Contractility of 5 human fibroblast cell lines. Fibroblast cell lines were cultured for 96 h and incubated with Dex in concentrations ranging from 10-9 to 10-7 M for 48 h or with medium alone (control). Gels were then released and transferred to dishes containing the same concentrations of Dex. Areas of gels were measured 1 h after release. Each symbol represents a different cell line from a different subject. Values are means of area of triplicate determinations.

Effect of other steroid derivatives or cholesterol. Dex, hydrocortisone, and methylprednisolone enhanced the contraction in a concentration-dependent manner, with EC50 values of ~1.8 × 10-8 (Dex), 2.5 × 10-8 (hydrocortisone), and 2.4 × 10-8 M (methylprednisolone; Fig. 4). There were no significant differences in the enhancement among Dex, hydrocortisone, and methylprednisolone. In contrast, no other steroid derivatives had any effect on contraction. MTT assay revealed that these steroids and their derivatives did not affect cell viability at the concentrations and incubation times used in these experiments (data not shown).


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Fig. 4.   Effects of various steroid derivatives on gel contraction. Fibroblasts embedded in collagen gels were cultured for 96 h and then incubated with Dex (), methylprednisolone (black-triangle), or hydrocortisone () as glucocorticoids (GCs) or with cholesterol (black-diamond ), aldosterone (), dehydroepiandrosterone (star ), methyltestosterone (triangle ), progesterone (open circle ), or estradiol-17beta (down-triangle) as other steroid derivatives. After incubation for 48 h, gels were released and transferred to dishes containing the same concentrations of reagents. Control gels were incubated with medium alone. Areas of gels were measured 1 h after release. Values are means of area of triplicate determinations.

Effect of cycloheximide on contractility. When the gels were incubated with 10-6 M Dex for 0, 1, 8, 24, 48, and 72 h before release, the contractions were 53.4 ± 4.7% decrease in initial gel area at 72 h, 48.7 ± 5.3% at 48 h, 45.8 ± 4.4% at 24 h, 30.8 ± 9.5% at 8 h, 1.5 ± 6.4% at 1 h, and 0.2 ± 1.2% at 0 h. The augmentative effect of Dex was observed only after a minimum of 8 h of incubation (P < 0.01 at 72, 48, 24, and 8 h), whereas no effect was observed with 1- and 0-h incubations. To assess whether the effect of GCs was mediated by protein synthesis, the effect of cycloheximide was investigated. Lysophosphatidic acid, which is known to enhance fibroblast contractility without de novo protein synthesis (7, 32), was used as a control. The increase in fibroblast contractility with Dex was abolished with 0.1 mM cycloheximide (P < 0.01; Fig. 5), whereas the increase with lysophosphatidic acid was not. Cycloheximide and lysophosphatidic acid did not affect cell viability at the concentration and incubation times used in these experiments (cycloheximide, 99.3 ± 10.5% of the control value; lysophosphatidic acid, 101.5 ± 8.7%). The effect of cycloheximide was reversible. After the cells were incubated with 0.1 mM cycloheximide for 8 h, the cells were washed with medium, then incubated for 48 h with and without 10-6 M GC. The contractility was enhanced with the GC after the removal of cycloheximide (35.8 ± 8.9% decrease in initial gel area with 10-6 M Dex; 2.7 ± 10.3% without Dex; 1.8 ± 4.4% with medium alone).


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Fig. 5.   Effect of cycloheximide on Dex-induced gel contraction. Fibroblasts embedded in collagen gels were cultured for 136 h and then incubated with Dex, lysophosphatidic acid (LPA), or medium alone [Dex(-)LPA(-)] in presence (hatched columns) and absence (open columns) of cycloheximide (0.1 mM) for 8 h before release. Areas of gels were measured after 1 h of incubation. Values are means ± SE of triplicate determinations.

Immunostaining of human fibroblasts within collagen gel. The five human fibroblast cell lines used in this study were all vimentin positive, desmin negative, and alpha -smooth muscle actin positive. The positive staining was found consistently in >70% of cells over 20 passages with our culture conditions. Treatment with 10-6 M Dex for 48 h induced an increase in the thickness of actin bundles (Fig. 6).


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Fig. 6.   Immunochemical staining with anti-alpha -smooth muscle actin antibody. Fibroblasts in type I collagen gels were cultured in serum-free DMEM for 96 h. Cells were then incubated with (A) and without (B) 1 µM Dex for 48 h. Then fibroblasts were fixed in 4% paraformaldehyde, and cell membranes were permeabilized with 0.2% Triton X-100 and immunostained with anti-alpha -smooth muscle actin antibody followed by FITC conjugate as a second antibody. Fluorescence intensity of actin bundles was seen in cytoplasm. These observations were made on a minimum of 3 randomly chosen microscopic fields (>100 total cells) on 3 separate occasions. Micrographs are representative sections. Bars, 20 µm.

Quantification of F-actin content. Dex enhanced the F-actin content in a concentration-dependent manner at concentrations ranging from 10-10 to 10-7 M, with an approximate EC50 of 8.4 × 10-7 M (Fig. 7A). This Dex-induced enhancement of F-actin content was also time dependent (Fig. 7B). The relative fluorescence intensity was 40.2 ± 3.6 at 48 h, 38.5 ± 2.8 at 24 h, 33.8 ± 1.7 at 8 h, and 21.9 ± 3.0 at 1 h of incubation with Dex and 20.2 ± 2.5 without incubation. The enhancement of F-actin content was observed at 48, 24, and 8 h of incubation but not within 1 h or without Dex.



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Fig. 7.   Effect of Dex on F-actin content. A: fibroblasts in type I collagen gels were cultured with serum-free DMEM for 96 h. Cells were then incubated with Dex at indicated concentrations for 48 h. Control gels were incubated with medium alone. Cells were then fixed and permeabilized, and binding of phalloidin to F-actin was detected as described in MATERIALS AND METHODS. B: fibroblasts were cultured in serum-free DMEM for 96 h and then incubated with 10-6 M Dex for indicated times (total incubation time was 144 h). Cells were then fixed and permeabilized, and binding of phalloidin to F-actin was detected. Values are means ± SE of triplicate determinations.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We demonstrated that five cell lines of human lung fibroblasts obtained from surgically resected specimens showed contractility in three-dimensional type I collagen gel cultures. The contractility and F-actin content in fibroblasts were augmented in the presence of GCs but not in the presence of other steroid derivatives. The increase in contractility was shown to be both concentration and time dependent. These phenomena were dependent on de novo protein synthesis because GC-induced enhancement in contractility required at least 8 h of incubation and cycloheximide abolished the augmentation.

Coulomb et al. (8) reported that the contractility of human dermal fibroblasts in collagen gel was reduced by GCs, suggesting that dermal fibroblasts may respond in different ways from lung fibroblasts as evidenced by our studies. However, we noticed two methodological differences between the study of Coulomb et al. and our present study: 1) in their report, they used Dex at concentrations ranging from 2 × 10-4 to 2 × 10-3 M and hydrocortisone at concentrations ranging from 2 × 10-4 to 1.5 × 10-3 M. In our experiments, the viability of fibroblasts within collagen gels decreased with treatment with GCs at concentrations > 10-5 M (2). In their study, culture medium supplemented with serum was used for the gel contraction assay, whereas in our study, the fibroblasts were cultured under serum-free conditions. Because serum factors can augment fibroblast-mediated gel contraction (37), their culture conditions may have made it difficult to detect the effect of GCs. Different culture conditions in the gel contraction assay may bring about different results.

Actin filaments with associated myosin and actin binding proteins are present in tension-generating fibroblasts. F-actin has been proposed to be organized in response to cell contraction (5, 38) and may participate in generating the forces responsible for continued development and maintenance of tension (12, 19, 23). However, there is no definite understanding of the precise regulatory mechanisms in terms of F-actin production. On the basis of our results, >8 h of culture were needed for demonstrating the effect of GCs on either fibroblast contractility or F-actin content. This suggests that a complex interplay may be required to induce a GC-induced contractility. Transforming growth factor-beta and platelet-derived growth factor are candidate proteins that enhance contractility, and beta -stimulants and prostaglandins are other candidates that suppress a contraction (26). GCs may modulate these proteins in terms of production or function. In addition, we propose the mechanism by which GC modulates actin-related proteins based on our result that F-actin content related to GC-induced contractility.

In our experimental conditions, there may be a possibility that a subset of fibroblasts that augments fibroblast contractility by GCs preferentially proliferated (4, 16, 34, 36). Five cell lines derived from five different adult lungs, however, showed similar contraction profiles in the presence of GCs. Thus further studies with cell lines obtained from patients with pulmonary fibrosis would be required to reveal potentially different profiles of contractility with and without GCs. Because hydrocortisone is a naturally occurring GC and methylprednisolone and Dex are both frequently used as therapeutic drugs for the treatment of various inflammatory lung injuries and fibrosis, we used all three of these GCs to compare the effects on contractility. In this study, we used physiological concentrations of GCs as anti-inflammatory drugs, although the anti-inflammatory activities differ markedly among these three GCs.

During wound healing, fibroblast-mediated contraction may favor repair processes at injured sites (20, 22). With regard to pulmonary fibrosis, exaggerated repair processes involving fibroblast proliferation relates lung dysfunction and architectural distortion. Theoretically, therefore, augmentation of fibroblast contractility by GCs may augment physiological dysfunction. It is not yet known whether GCs can modulate lung fibroblast contractility in cells obtained from disease sites. It will be important to compare fibroblasts from healthy subjects, patients with immature fibrosis of the type found in bronchiolitis obliterans organizing pneumonia, and patients with the mature fibrosis found in usual interstitial pneumonia (30).

In conclusion, human lung fibroblasts showed contractility in three-dimensional type I collagen gel culture that differed from dermal fibroblasts. This contractility was enhanced in the presence of GCs but not in the presence of other steroids. This contractility was induced through de novo protein synthesis and was consistent with increased F-actin content in lung fibroblasts.


    ACKNOWLEDGEMENTS

We thank Dr. Roland M. duBois for reviewing the manuscript. We thank all of the staff of the surgical department of Kyoto University (Kyoto, Japan) and Dr. Seiichi Matsunobe of Social Insurance Shiga Hospital (Otsu, Shiga Prefecture, Japan) for kindly providing the lung specimens. We also thank Fumiko Tanioka and Machiko Yamada for technical assistance in the experimental works and Simon Johnson for checking linguistic problems.


    FOOTNOTES

This study was supported by Grant-in-Aid 08670661 from the Ministry of Education of Japan and the Smoking Research Foundation in Japan.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Address for reprint requests and other correspondence: S. Nagai, Department of Respiratory Medicine, Graduate School of Medicine, Kyoto University, 53 Shogoin, Kawahara-cho, Sakyo-ku, Kyoto 606-8507, Japan (E-mail: nagai{at}kuhp.kyoto-u.ac.jp).

Received 30 November 1998; accepted in final form 18 August 1999.


    REFERENCES
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Am J Physiol Lung Cell Mol Physiol 278(1):L13-L18
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