Department of Respiratory Medicine, Graduate School of Medicine, Kyoto University, Kyoto 606-8507, Japan
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ABSTRACT |
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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 × 108 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
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INTRODUCTION |
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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.
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MATERIALS AND METHODS |
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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-17 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 -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-
-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
-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 × 107 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.
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RESULTS |
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Effect of Dex on the gel contraction. Dex enhanced
fibroblast-mediated collagen gel contraction in a
concentration-dependent manner at concentrations ranging from
1010 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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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 × 108 (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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Effect of cycloheximide on contractility. When the gels were
incubated with 106 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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Immunostaining of human fibroblasts within collagen gel. The
five human fibroblast cell lines used in this study were all vimentin
positive, desmin negative, and -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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Quantification of F-actin content. Dex enhanced the F-actin
content in a concentration-dependent manner at concentrations ranging
from 1010 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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DISCUSSION |
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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 × 104 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- and platelet-derived growth factor are
candidate proteins that enhance contractility, and
-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.
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ACKNOWLEDGEMENTS |
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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.
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FOOTNOTES |
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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.
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